The analytical pyrolysis technique hyphenated to gas chromatography–mass spectrometry (GC–MS) has extended the range of possible tools for the characterization of synthetic polymers and copolymers. Pyrolysis involves thermal fragmentation of the analytical sample at temperatures of 500–1400 °C. In the presence of an inert gas, reproducible decomposition products characteristic for the original polymer or copolymer sample are formed. The pyrolysis products are chromatographically separated using a fused-silica capillary column and are subsequently identified by interpretation of the obtained mass spectra or by using mass spectra libraries. The analytical technique eliminates the need for pretreatment by performing analyses directly on the solid or liquid polymer sample. In this article, application examples of analytical pyrolysis hyphenated to GC–MS for the identification of different polymeric materials in the plastic and automotive industry, dentistry, and occupational safety are demonstrated. For the first time, results of identification of commercial light-curing dental filling material and a car wrapping foil by pyrolysis–GC–MS are presented.

Structural analysis and the study of degradation properties are important to understand and improve performance characteristics of synthetic polymers and copolymers in many industrial applications. Traditional analytical techniques used for characterization of polymers and copolymers such as thermal analysis and Fourier transform infrared (FT–IR) spectroscopy have limitations or are not sufficiently sensitive (1). Pyrolysis techniques hyphenated to gas chromatography–mass spectrometry (GC–MS) have extended the range of possible tools for the characterization of synthetic polymers and copolymers. Under controlled conditions, at elevated temperatures (500–1400 °C) in the presence of an inert gas, reproducible decomposition products characteristic for the original polymer or copolymer sample are formed. The pyrolysis products are chromatographically separated using a fused-silica capillary column and subsequently identified by interpretation of the obtained mass spectra or by using mass spectra libraries (such as the National Institute of Standards and Technology [NIST] or Wiley). Pyrolysis methods eliminate the need for pretreatment by performing analyses directly on the solid polymer or copolymer sample (1). (Please note that this article was presented at the XVII European Conference on Analytical Chemistry, which was held in Warsaw, Poland, on August 25–29, 2013).  

Most of the thermal degradation results from free radical reactions initiated by bond breaking and depends on the relative strengths of the bonds that hold the molecules together. A large molecule will break apart and rearrange in a characteristic way (2–4). If the energy transfer to the sample is controlled by temperature, heating rate, and time, the fragmentation pattern is reproducible and characteristic for the original polymer or copolymer. Another sample of the same composition, heated at the same rate to the same temperature for the same period of time, will produce the same decomposition products. Therefore, the essential requirements of the apparatus in analytical pyrolysis are reproducibility of the final pyrolysis temperature, rapid temperature rise, and accurate temperature control. Depending on the heating mechanism, pyrolysis systems have been classified into two groups: the continuous-mode pyrolyzer (furnace pyrolyzer) and pulse-mode pyrolyzer (flash pyrolyzer, such as the heated filament, Curie-point, and laser pyrolyzer). The pyrolysis unit is directly connected to the injector port of a gas chromatograph. A flow of an inert carrier gas, such as helium, flushes the pyrolyzates into the fused-silica capillary column. Figure 1 shows the schematic view of the furnace pyrolyzer used in our investigation. The detection technique of the separated compounds is typically MS, but other GC detectors have also been used depending on the intentions of the analysis (1,4). 

 Figure 1: Schematic view of the furnace pyrolyzer used in this study.			

The applications of analytical pyrolysis–GC–MS range from research and development of new materials, quality control, characterization and competitor product evaluation, medicine, biology and biotechnology, geology, airspace, and environmental analysis to forensic purposes or conservation and restoration of cultural heritage. These applications cover analysis and identification of polymers, copolymers, and additives in components of automobiles, tires, packaging materials, textile fibers, coatings, half-finished products for electronics, paints or varnishes, lacquers, leather, paper or wood products, food, pharmaceuticals, surfactants, and fragrances.  

Our earlier publications (1,5–12) presented the analysis and identification of degradation products of commercially available synthetic polymers and copolymers by using analytical pyrolysis hyphenated to gas chromatography with flame ionization detection (GC–FID) and GC–MS. In this work, new examples of applications of this analytical technique for the identification of different polymeric materials are demonstrated. 

Plastic particles from industrial filter fins, a car wrapping foil, unknown fibers, and commercial light-curing dental filling material were used in the investigation. 

Instrumentation and Analytical Conditions

Approximately 100–200 µg of solid sample was cut out with a scalpel and inserted without any further preparation into the bore of the pyrolysis solids-injector and then placed with the plunger on the quartz wool of the quartz tube of the furnace pyrolyzer Pyrojector II (SGE Analytical Science). Three spots on each sample were analyzed in duplicate. The pyrolyzer was operated at a constant temperature of 550, 600, 700, or 900 °C. The pressure of helium carrier gas at the inlet to the furnace was 95 kPa.  

Pyrolysis–GC–MS measurements were made using two apparatus. In the first apparatus (1), the pyrolyzer was connected to a Trace 2000 gas chromatograph (ThermoQuest, CE Instruments) with a quadrupole mass spectrometer Voyager (ThermoQuest, Finnigan, MassLab Group) operated in electron ionization (EI) mode. A 60 m × 0.25 mm, 0.25-µm Elite-5ms fused-silica GC capillary column (PerkinElmer Instruments) was used. The GC conditions were as follows: programmed column temperature: 60 °C for 1 min, then 60–100 °C at 2.5 °C/min, 100–280 °C at 10 °C/min (20-min hold at 280 °C). The temperature of the split–splitless injector was 250 °C and the split flow was 50 cm

/min. Helium, grade 5.0 (Westfalen AG), was used as a carrier gas. The helium programmed pressure was 70 kPa for 1 min, then 70–110 kPa at 1 kPa/min (hold at 110 kPa to the end of analysis) was used. The transfer line temperature was 280 °C. The MS EI ion source temperature was kept at 250 °C. The ionization occurred with a kinetic energy of the impacting electrons of 70 eV. The current emission of the rhenium filament was 150 µA. The MS detector voltage was 350 V. Mass spectra and reconstructed chromatograms (total ion current [TIC]) were obtained by automatic scanning in the mass range 

35–450 u. Pyrolysis–GC–MS data were processed with the Xcalibur software (ThermoQuest) and the NIST 05 mass spectra library. 

In the second apparatus (2), the pyrolyzer was connected to a 7890A gas chromatograph with a series 5975C quadrupole mass spectrometer (Agilent Technologies Inc.) operated in EI mode. A 59 m × 0.25 mm, 0.25-µm 

 DB-5ms fused-silica GC capillary column (J&W Scientific) was used. Helium, grade 5.0 (Westfalen AG), was used as a carrier gas. The GC conditions were as follows: programmed column temperature: 60 °C for 1 min, then 60–280 °C at 7 °C/min (hold at 280 °C to the end of analysis); programmed helium pressure: 122.2 kPa for 1 min, then 122.2–212.9 kPa at 7 kPa/min (hold at 212.9 kPa to the end of analysis). Second set of GC conditions: programmed column temperature: 75 °C for 1 min, then 75–280 °C at 7 °C/min (hold at 280 °C to the end of analysis); programmed helium pressure: 122.2 kPa for 1 min, then 122.2–212.9 kPa at 7 kPa/min (hold at 212.9 kPa to the end of analysis). 

The temperature of the split–splitless injector was 250 °C and the split ratio was 50:1. The transfer line temperature was 280 °C. The MS EI ion source temperature was kept at 230 °C. The ionization occurred with a kinetic energy of the impacting electrons of 70 eV. The quadrupole temperature was 150 °C. Mass spectra and reconstructed chromatograms (total ion current) were obtained by automatic scanning in the mass range 

 35–750 u. Pyrolysis–GC–MS data were processed with the ChemStation software (Agilent Technologies) and the NIST 05 mass spectra library. 

Figure 2: PyrolysisâGCâMS chromatogram of plastic particles from industrial filter fins at 700 Â°C obtained with apparatus 1. Fused-silica GC capillary column: 60 m Ã 0.25 mm, 0.25-Âµm df Elite-5ms. GC conditions: programmed column temperature: 60 Â°C for 1 min, then 60â100 Â°C at 2.5 Â°C/min and then 100â280 Â°C at 10 Â°C/min (20 min hold at 280 Â°C); splitâsplitless injector temperature: 250 Â°C; split flow: 50 cm3/min; helium programmed pressure: 70 kPa for 1 min, then 70â110 kPa at 1 kPa/min (hold at 110 kPa to the end of analysis). For peak identification, see Table I.			

Pyrolysis–GC–MS of Plastic Particles from Industrial Filter Fins

A sample of plastic particles from industrial filter fins was pyrolyzed at 700 °C to identify its composition. Figure 2 shows the obtained pyrolysis–GC–MS chromatogram of the sample. Based on the decomposition products summarized in Table I, the plastic particles were identified as a mixture of poly(hexamethylene adipamide) (nylon 6-6) and polypropylene glycol with a small amount of styrene–butadiene-rubber (SBR). The peaks of propylene and acetone indicate the presence of polypropylene glycol. The main decomposition product of nylon 6-6 is cyclopentanone (retention time [

] = 10.51 min). Other peaks in Figure 2, like butene/1,3-butadiene (

 = 14.39 min) are typical pyrolysis products of SBR (1,2,5,6,18). The small peak of 4-isopropylphenol (

 = 28.15 min) may be a clue to the presence of polycarbonate or bisphenol A epoxy resin (5,6). All of the pyrolysis products and the materials identified from pyrolysis products in filter fins are summarized in Table I. 

Table I: Pyrolysis products and identified materials in plastic particles from industrial filter fins 			

Pyrolysis–GC–MS of a Car Wrapping Foil

The next object of identification was a car wrapping foil pyrolyzed at 600 °C. Figure 3 shows the obtained pyrolysis–GC–MS chromatogram of the car wrapping foil. Based on the decomposition products summarized in Table II, the plastic material was identified as a mixture of flexible poly(vinyl chloride) (PVC) with bis(2-ethylhexyl) phthalate (BEHP) plasticizer and poly(hexamethylene adipamide) (nylon 6-6). The chromatogram in Figure 3 shows the typical pyrolysis products of PVC, like hydrogen chloride (

 = 25.93 min). This is the result of the formation of double bonds by the elimination of hydrogen chloride from the poly(vinyl chloride) macromolecules, followed by the breaking of the carbon chain with or without cyclization reaction (2). The detected cyclopentanone (

 = 10.46 min) is generally known as characteristic pyrolysis product of nylon 6-6 (2,3,6). Methyl methacrylate (

 = 8.20 min) identified in pyrolyzate is formed from poly(methyl methacrylate) (6) and most likely comes from the adhesive film. Thus, the identified 3,3-diphenylacrylonitrile (

 = 36.69 min) may be from the adhesive layer of the foil. 

Figure 3: PyrolysisâGCâMS chromatogram of a car wrapping material at 600 Â°C obtained with apparatus 1. Fused-silica GC capillary column 60 m Ã 0.25 mm, 0.25-Âµm df Elite-5ms. GC conditions: programmed column temperature: 60 Â°C for 1 min, then 60â100 Â°C at 2.5 Â°C/min and then 100â280 Â°C at 10 Â°C/min (20 min hold at 280 Â°C); splitâsplitless injector temperature: 250 Â°C; split flow: 50 cm3/min; helium programmed pressure: 70 kPa for 1 min, then 70â110 kPa at 1 kPa/min (hold at 110 kPa to the end of analysis). For peak identification, see Table II.			

The thermal decomposition of the plasticizer bis(2-ethylhexyl) phthalate identified in car wrapping foil leads to the formation at 600 °C of 2-ethyl-1-hexene (

= 28.63 min) (1,7). In the car wrapping material, the rest of the tetrahydrofuran solvent (

 = 6.99 min) was also detected. Table II shows the identified ingredients of the pyrolyzed car wrapping foil. 

Table II: Pyrolysis products and identified materials in car wrapping foil 			

A sample of unknown plastic fibers was pyrolyzed at 700 °C and 900 °C, respectively, to identify its composition. Figure 4 shows the pyrolysis–GC–MS chromatogram of the sample pyrolyzed at 900 °C. Based on the decomposition products summarized in Table III, the fibers were identified as polyaramid [poly(

-phenylene terephthalamide)] (Figure 5). The main identified degradation products of polyaramid at 900 °C are benzene (

 = 11.81 min). Currently, polyaramid fibers have only been characterized in a few publications using thermal analysis (thermogravimetry, derivative thermogravimetry, and differential thermal analysis), infrared spectroscopy techniques (13–15), and pyrolysis–GC–MS (2,16–19). 

Figure 4: PyrolysisâGCâMS chromatogram of polyaramid fibers at 900 Â°C obtained with apparatus 2. Fused-silica GC capillary column: 59 m Ã 0.25 mm, 0.25-Âµm df DB-5ms. GC conditions: programmed column temperature: 75 Â°C for 1 min, then 75â280 Â°C at 7 Â°C/min (hold to the end of analysis); programmed pressure of helium carrier gas: 122.2 kPa for 1 min, then 122.2â212.9 kPa at 7 kPa/min (hold at 212.9 kPa to the end of analysis). For peak identification, see Table III.			

Polyaramid fibers are a class of heat-resistant, strong synthetic fibers. They are used in aerospace and military applications for ballistic-rated body armor, fabric, ballistic composites, and fire fighters protective clothing as well as in bicycle tires and as an asbestos substitute. 

Table III: Pyrolysis products of polyaramid fibers 			

Identification of Commercial Light-Curing Dental Filling Material

A number of dental filling materials are presently available for tooth restorations. The four main groups of these materials, which dentists have used for about 35 years, are the conventional glass-ionomer cements, resin-based composites, resin-modified glass-ionomer cements, and polyacid-modified resinous composites (20). Light-curing glass-ionomer cements contain polyacrylic acid, chemically or photo-curing monomers (multifunctional methacrylates, like triethylene glycol dimethacrylate or 2-hydroxyethyl methacrylate), an ion-leaching glass, and additives (initiators, inhibitors, stabilizers, and others) (20). Resin-modified glass-ionomer cements are now widely used in dentistry as direct filling materials, liners, bases, luting cements, and fissure sealants (21). These materials mainly consist of polymer matrix and glass-ionomer parts. The polymer matrix is based on a monomer system and different multifunctional methacrylates with additives (21). Methacrylic monomers, such as bisphenol A glycidyl methacrylate (Bis-GMA), urethane dimethacrylate (UDMA), triethylene glycol dimethacrylate (TEGDMA), and 2-hydroxyethyl methacrylate (HEMA), are the main components of resin-based dental filling materials. The presence of additives such as initiators, activators, inhibitors, and plasticizers in uncured dental material mixture is necessary (21). 

Figure 5: Chemical structure of poly(p-phenylene terephthalamide) (polyaramid).			

Figure 6 shows the total ion current pyrolysis–GC–MS chromatogram of commercial light-curing dental filling material pyrolyzed at 550 °C. The pyrolysis products identified by using mass spectra library NIST 05 are summarized in Table IV. The carbon dioxide (

 = 6.85 min) identified in pyrolyzate is formed from polyacrylic acid (2,18). The identified substances 2-hydroxyethyl methacrylate (HEMA) (

 = 13.65 min), ethylene glycol dimethacrylate (EGDMA) (

 = 19.48 min), and triethylene glycol dimethacrylate (TEDMA) (

 = 28.72 min) are known as standard composites of dental filling materials (1). Other compounds in Table IV, such as bisphenol A (

 = 33.10 min) or bisphenol A diglycidyl ether (

 = 42.42 min), are probably formed by thermal degradation of bisphenol A diglycidyl mono- or dimethacrylates. The presence of the additives, such as the antioxidant 2,6-bis(1,1-dimethylethyl)-4-methylphenol (BHT) (

 = 23.17 min) or the UV-absorber drometrizol (

 = 31.95 min) was also confirmed. The triphenylantimony (

 = 34.55 min) identified in pyrolyzate is used as catalyst in the UV-induced polymerization (1). 

Figure 6: PyrolysisâGCâMS chromatogram of commercial light-curing dental filling material at 550 Â°C obtained with apparatus 2. Fused-silica GC capillary column: 59 m Ã 0.25 mm, 0.25-Âµm df DB-5ms. GC conditions: programmed column temperature: 60 Â°C for 1 min, then 60â280 Â°C at 7 Â°C/min (hold at 280 Â°C to the end of analysis); programmed helium pressure: 122.2 kPa for 1 min, then 122.2â212.9 kPa at 7 kPa/min (hold at 212.9 kPa to the end of analysis). For peak identification, see Table IV.			

Analytical pyrolysis–GC–MS has been proven as a valuable technique for the analysis and identification of organic polymeric materials in the plastic and rubber industry. For the first time pyrolysis–GC–MS was used for the identification of commercial light-curing dental filling material and for the identification of a car wrapping foil. This technique allows the direct analysis of very small sample amounts (5–200 µg) without the need for time-consuming sample preparation. 

Table IV: Pyrolysis products of commercial light-curing dental filling material 			

Advanced Gas Chromatography – Progress in Agricultural, Biomedical and Industrial Applications

, M.A. Mohd, Ed. (InTech, Rijeka, Croatia, 2012), pp. 343–362.  

Analytical Pyrolysis of Synthetic Organic Polymers

 (Elsevier, Amsterdam, The Netherlands, 2005). 

 (CRC Press, Boca Raton, Florida, 2007). 

(4) K.L. Sobeih, M. Baron, and J. Gonzales-Rodrigues, 

(6) P. Kusch, G. Knupp, and A. Morrisson, in  

 R.K. Bregg, Ed. (Nova Science Publishers, New York, New York, 2005), pp. 141–191. 

(8) P. Kusch, V. Obst, D. Schroeder-Obst, W. Fink, G. Knupp, and J. Steinhaus, 

(10) P. Kusch, V. Obst, D. Schroeder-Obst, G. Knupp, and W. Fink, 

(12) P. Kusch, W. Fink, D. Schroeder-Obst, and V. Obst, 

Articles

Peer-Reviewed Articles

This pyrolysis–GC–MS method enables direct analysis of solid or liquid polymers without sample pretreatment, as illustrated here for various materials, including a dental filling material and a car wrapping foil.

Application of Pyrolysis–Gas Chromatography–Mass Spectrometry for the Identification of Polymeric Materials

In this study, the procedure for analyzing amphetamines and synthetic cathinones (also known as bath salts drugs) in hair samples using a mixed-mode solid-phase extraction (SPE) is described. Samples of hair were digested with a dilute solution of base (containing internal standards), neutralized, and diluted with an aqueous phosphate buffer (pH 6). Each sample was applied to a conditioned SPE column, after which the sorbent was rinsed with deionized water, acetic acid, and methanol. After drying, the analytes were eluted and collected from the SPE column with 3 mL of an elution solvent consisting of methylene chloride–isopropanol–ammonium hydroxide. To the eluate, 200 µL of mobile phase was added and the samples were evaporated to the mobile phase for analysis by liquid chromatography–tandem mass spectrometry (LC–MS-MS). Chromatography was performed in gradient mode using a C18 column and a mobile phase consisting of acetonitrile and 0.1% aqueous formic acid. The total run time for each analysis was under 5 min.

Amphetamine (α-methylphen-ethylamine) (Figure 1) belongs to a class of compounds known as 

(1). This class of drugs includes the illicit drugs methamphetamine, methylenedioxyamphetamine (MDA), and methylenedioxymethamphetamine (MDMA), as well as ephedrine and pseudoephedrine, which can be found in over-the-counter medications. Amphetamine is administered as a prescription medication (for example, Adderall [Shire Pharmaceuticals]) for treating medical issues such as narcolepsy, obesity, or hypotension (2), whereas methamphetamine, MDA, and MDMA are considered controlled substances — that is, pharmaceuticals with little or no medical use.  

Synthetic cathinones are derived structurally from the parent compound (Figure 2) and have become noticeable in the scientific literature in recent times because of the fatalities arising from administration (3–6). The drugs are commonly referred to as "bath salts" because they were originally packaged with names such as "Ivory Wave" and marketed as "not for human consumption" or "research chemicals." These drugs are now scheduled in the same way as other controlled substances. 

The popularity of amphetamines is because of their euphoria effect and ease of synthesis. Their use or abuse is generally verified by the analysis of biological samples, such as urine, blood, oral fluid, or hair. Of these samples, hair is a biological matrix that has been used as an alternative to urine or blood for drug testing because it allows noninvasive sampling and can document the use of the drugs over a longer period of time than blood or urine (7). In this study, amphetamine in the form of Adderall was determined in the hair of a subject along with several other amphetamines and a range of synthetic cathinones. 

In terms of metabolism in the human system, amphetamine is inactivated during this process, undergoing deamination to form phenylacetone, which is converted to benzoic acid and excreted in a conjugated form (2). A small amount of the parent is oxidized to norephedrine, which is also metabolized to the parahydroxylated forms of this compound, all of which is pharmacologically active and are thought to contribute to the effects of the drug (8,9). The therapeutic, toxic, and fatal concentrations of amphetamines in samples such as blood and urine are well documented (2,10), but  not so much for the synthetic cathinones because of the recent nature of their abuse. Most of the published studies on hair analysis have been performed in the area of work place drug testing and drugs or driving cases (11,12), not postmortem studies. 

Gas chromatography coupled to mass spectrometry (GC–MS) has been reported as a technique for quantifying amphetamines in hair (13). GC–MS analysis of the amphetamines requires derivatization using compounds such as heptafluorobutyric anhydride (HFAA) or pentafluoropropionic anhydride (PFAA) (14,15). Because amphetamine exists as a 

 isomeric pair, some laboratories have used chiral modification to separate the isomers in samples such as hair (16). The ratio of the isomeric forms may indicate whether or not the amphetamine has been taken legally. GC–MS analysis of the cathinones used similar fluoroacyl derivatives (3). Liquid chromatography coupled to tandem mass spectrometry (LC–MS-MS) is gaining popularity for analysis in this matrix (17,18). The use of solid-phase extraction (SPE) described in this article uses the LC mobile phase as a keeper solvent for amphetamine, reducing its volatility. In previous methods the addition of methanolic hydrochloric acid or a solvent such as dimethylformamide has been reported, and the solvent was evaporated to dryness (19). The addition of the mobile phase presents the LC–MS-MS with a more amenable analytical solvent. SPE has been reported in the analysis of hair samples previously (20–22), but not using this type of keeper solution format. 

, butylone, ethylone, flephedrone, mephedrone, mephedrone-

, methylone, methedrone, methcathinone (4-MEC), methylenedioxypyravalerone (MDPV), and pyravalerone  were obtained from Lipomed as 1-mg/mL methanolic solutions. Acetonitrile, acetic acid (glacial), concentrated ammonium hydroxide solution (32% by volume), formic acid, isopropanol, methanol, and methylene chloride were obtained from Fisher Scientific. The SPE columns (CSDAU206) were obtained from UCT Inc. Deionized (DI) water was laboratory grade, and it was generated in the Massachusetts State Police Crime Laboratory (MSPCL). All chemicals were of ACS grade. 

Acetic acid was prepared as a 0.1 M solution by diluting glacial acetic acid (5.8 mL to 500 mL) and then increasing the volume to 1 L by adding DI water and mixing well. Formic acid was prepared as a 0.1% (v/v) solution by adding 1 mL of the acid to 900 mL of DI water and diluting it to 1 L (mobile-phase A). Acetonitrile containing 0.1% formic acid (v/v) was prepared by adding 1 mL of formic acid to 900 mL of acetonitrile and diluting to 1 L (mobile-phase B). Aqueous sodium hydroxide was prepared as a 0.1 M solution by adding 4 g of the solid to 500 mL of DI water and dissolving before diluting to 1 L. Phosphate buffer (pH 6, 0.1 M) was purchased from Fisher Scientific as a ready-to-use solution. 

Analysis was performed using an API 3200 Q-Trap instrument supplied by Applied Biosystems. The chromatographic system consisted of a Shimadzu CBM 20 A controller, two Shimadzu LC 20 AD pumps (including a degasser), a Shimadzu SIL 20 AC autosampler, and a Shimadzu CTO AC oven (set at 10 °C). The instrument was fitted with a 50 mm × 2 mm, 5-µm 

 Cadenza-C18 column from Imtakt USA (formerly Silvertone Sciences) that was attached to a 5 mm × 2 mm Cadenza-C18 guard column obtained from the same supplier. The LC column oven was maintained at 40 °C throughout the analyses. The injection volume was 10 µL. The mobile phase was delivered at a flow rate of 0.5 mL/min. The mobile-phase gradient program started at 5% mobile-phase B, rose to 90% B in 4.0 min, and then returned to 5.0% B. The instrument was readied for reinjection after 5.1 min. 

The mass spectrometry was performed using positive multiple reaction monitoring (MRM). The mass spectrometer conditions for each of the amphetamines and synthetic cathinones are shown in Table I. Tandem MS was performed using the following conditions: curtain gas setting, 15; collision gas setting, medium; ion spray voltage setting, 5000 V; temperature setting, 650 °C; ion source gas 1 setting, 50; ion source gas 2 setting, 50. The tandem mass spectrometer conditions are shown in Table I. The analytical data were collected using Analyst software version 1.5.2 supplied by Applied Biosystems.  

Table I: Tandem mass spectrometry conditions			

A solution of amphetamines and synthetic cathinones (amphetamine, methamphetamine, MDA, MDMA, butylone, ethylone, flephedrone, mephedrone, methylone, methedrone, methcathinone [4-MEC], methylenedioxypyravalerone [MDPV], and pyravalerone) was prepared at a concentration of 1 µg/mL by the dilution of 10 µL of stock solution with acetonitrile to 10 mL in a volumetric flask. A 1-µg/mL solution of the internal standards (amphetamine-

) was prepared by the diluting 100 µL of the stock solution (100 µg/mL) to 10 mL with acetonitrile in a volumetric flask. The choice of internal standard was based on the fact that deuterated analogues of amphetamines and synthetic cathinones would not only chromatograph in a very similar mode to the drugs themselves, but also would be extracted via SPE as efficiently as amphetamine and the synthetic cathinones and would not likely be observed in the samples under analysis. 

Calibrators were prepared by the addition of 1.0, 2.5, 5.0, 10.0, 25.0, 50.0, and 100.0 ng of amphetamines and synthetic cathinones into 10-mg samples of drug-free hair samples. The hair samples had been previously decontaminated by washing two times with methylene chloride (10 mL) and two times with DI water (10 mL) before being air dried for 24 h. Each wash step was analyzed for the presence of drugs and found to be negative. Next, 50 ng of the internal standard was added to these samples. Control samples were prepared by the addition of 2 ng and 8 ng of amphetamine and synthetic cathinones to drug-free hair samples in addition to 50 ng of the internal standard. All determinations were performed in duplicate. A negative control sample was prepared by the addition of only the internal standard (50 ng) to a sample of drug-free hair samples (10 mg). Calibrators, control samples, and test samples were treated in an identical mode with regard to sample extraction. 

To assess the performance of the procedure, calibration curves were constructed twice daily over five consecutive days using the spiked controls, and from these data intraday and interday values were obtained. 

A 1-mL volume of 0.1 M aqueous sodium hydroxide solution was added to each sample (calibrator, control, and test) in a clean glass tube (75 mm × 12 mm) that was then capped. The tubes and contents were digested for 30 min at room temperature. Then, 4 mL of 0.1 M phosphate buffer (pH 6) was added to the solution, which was mixed and centrifuged at 3000 rpm for 10 min before it was applied to the SPE column. 

SPE columns were conditioned by the sequential addition of 3 mL of methanol, 3 mL of DI water, and 1 mL of 0.1 M phosphate buffer (pH 6). Each liquid was allowed to percolate through the sorbent using gravity without allowing the sorbent to dry out between steps. 

Following the passage of the methanol, DI water, and 0.1 M phosphate buffer (pH 6) through the SPE columns, each diluted sample (that is, calibrator, control, and case item) was loaded on to an individually marked SPE tube and allowed to pass through the sorbent using gravitational flow. The columns were then washed with sequential additions of 3 mL of DI water, 1 mL of 0.1 M acetic acid, and 3 mL of methanol. The SPE columns were then dried by applying a vacuum to the SPE manifold at 15 in. of mercury pressure using an electric vacuum pump. 

The analytes were eluted from the SPE columns by the addition of 3 mL of a 78:20:2 methylene chloride–isopropanol–ammonium hydroxide solution. This solution was prepared daily by adding 2 mL of concentrated ammonium hydroxide solution to 20 mL of isopropanol and mixing well. Then, 78 mL of methylene chloride was added to this solution and the resultant solution was transferred to a clean screw-top bottle for use. A screw-top bottle ensures that the basicity of the solution remains high by eliminating any loss of ammonia from the bottle. The elution solvent was allowed to flow through the SPE sorbent with the aid of gravity and collected in separate glass tubes (75 mm × 12 mm). Glass tubes were chosen because they are standard laboratory materials within this toxicology laboratory. 

Following elution, 200 µL of a solution containing 95% of mobile-phase A and 5% of mobile-phase B was added to each sample tube. The tubes were vortex mixed for approximately 1 min before the eluates were evaporated to the mobile phase using a gentle stream of nitrogen at 35 °C. After further vortex mixing, the samples were transferred to an autosampler vial (2 mL) containing a low-volume insert (250 µL) and the vial was capped for analysis. 

Studies of the matrix effects were performed according to a previously published procedure (23). In this part of the study, aliquots of amphetamine and synthetic cathinones (covering the linear range) were introduced into 200 µL of a solution containing 95% of mobile-phase A and 5% mobile-phase B. Each elution solution was evaporated to remove the organic component until only the mobile phase remained and then they were analyzed by LC–MS-MS (analysis A). Concurrently, a set of hair samples were subjected to the SPE process; after elution of the analytes from the SPE columns, the elution solvent was spiked with amphetamine, and 200 µL of mobile phase (95:5 A–B) was added, and then the solution was evaporated to the mobile phase (analysis B). A second set of hair samples was spiked with amphetamine and synthetic cathinones and processed via the SPE method. After elution, 200 µL of mobile phase was added and after vortex mixing, the solution was evaporated to the mobile phase (analysis C). The data (peak areas) for analyses A, B, and C were collected by the data analysis software. By comparing the peak areas of analysis B with those of A, an assessment of matrix effects was made. The comparison of peak areas for C with B provided data for the recoveries. 

Amphetamine and synthetic cathinone solution (concentration: 50 ng/mg) was infused into the tandem mass spectrometer using the on-board syringe pump (controlled by the data analysis software) via a Hamilton syringe (model 1001TLL, 1 mL volume, supplied by Fisher Scientific) at a flow rate of 5 µL/min. At the same time as the amphetamine solution was flowing into the mass spectrometer, a 10-µL aliquot of the SPE-extracted hair matrix (samples of hair confirmed to contain no drug material) was injected using the autosampler syringe on the Shimadzu liquid chromatograph. The liquid chromatograph and mass spectrometer were arranged so that samples from the liquid chromatograph were mixed into the flow of amphetamine and synthetic cathinones via a three-port tee section before the total flow entered the mass spectrometer. Any suppression effects on the amphetamine could be monitored at the MRM step for the noted drugs. 

When analyzing samples of hair extracts via SPE and LC–MS-MS it is essential to ensure that the interfering effects of other drug compounds can be eliminated. In this procedure, samples of hair extracts were spiked with 49 drugs at a concentration equivalent to 100 ng/mg of hair sample: (bupropion, lidocaine, methadone, amitriptyline, nortriptyline, thioridazine, trazodone, mesoridazine, meperidine, diphenhydramine, phenyltoloxamine, imipramine, desipramine, benztropine, trimethoprim, diltiazem, haloperidol, strychnine, morphine, codeine, 6-acetylmorphine, oxycodone, oxymorphone, hydrocodone, noroxycodone, hydromorphone, diazepam, nordiazepam, oxazepam, temazepam, alprazolam, α-hydroxyalprazolam, lorazepam, triazolam, α-hydroxytriazolam, flunitrazepam, 7-amino-flunitrazepam, chlordiazepoxide, midazolam, α-hydroxymidazolam, flurazepam, desalkyl-flurazepam, cocaine, ecgonine methyl ester, ecgonine ethyl ester, benzoylecgonine, cocaethylene, clonazepam, and 7-amino-clonazepam) and extracted according to the SPE method. It was observed that the interfering effect of these compounds was not found to be significant.

The mean recovery of amphetamine and synthetic cathinones from drug-free hair samples was determined to be 95% (±2%). This is an excellent indicator for the efficiency of the extraction procedure of amphetamines and synthetic cathinones using hair as a matrix. This procedure was performed twice daily over a period of five days. 

The results of the analysis of the spiked control samples of hair (2 ng/mg and 8 ng/mg, respectively) are shown in Table II. Analysis of the control samples was performed at the same time as the calibration curves were constructed — that is, over a period of five days. Control samples were prepared by adding the amphetamine and synthetic cathinone solution to the hair sample (10 mg) in the digestion mixture and treating as per the test samples. 

Table II: Precision results for control hair samples shown as percent recovery			

Intraday and interday variation for the analysis of amphetamines and synthetic cathinones was found to be less than 7% and less than 10%, respectively. Ion suppression studies revealed that suppression of the monitored ions was less than 5%. This method was found to be linear (

 > 0.995) over the dynamic range 0.1–10 ng/mg. 

Limit of Detection and Limit of Quantification

 (LOD) of a particular method can be defined as the level at which the signal-to-noise ratio for the particular analyte is greater than or equal to 3:1. The 

 (LOQ) for the method is the level at which the signal-to-noise ratio for a particular analyte is greater than or equal to 10:1. In this study, LOD values were determined empirically by analyzing extracted samples of drug-free hair fortified with amphetamines and synthetic cathinones by LC–MS-MS according to the SPE method. This was performed until the lowest level at which each of the respective analytes just failed the signal-to-noise ratio of 3:1. This was observed as 0.05 ng/mg. In terms of LOQ, samples of drug-free hair samples were spiked with amphetamines and synthetic cathinones at concentrations below 10 ng/mg and extracted according to the SPE procedure until the analytes just failed a signal-to-noise ratio of 10:1; this value was found to be 0.1 ng/mg. This is better than the recommendations by the Society of Hair Testing (SoHT) of 0.2 ng/mg (24). Representative chromatograms at LOQ and genuine hair sample are shown in Figures 3 and 4. 

Figure 3: Chromatogram of hair analysis containing amphetamines and synthetic cathinones at the LOQ (0.1 ng/mg). See Table I for compound identification.			

Because hair is a solid matrix, the sample requires digestion to produce a liquid that is able to flow through the SPE column. As the digest is basic, buffering to pH 6 permits both efficient flow and optimal sorbing of the drugs onto the SPE sorbent. In using a mixed-mode (C8 and strong cation-exchange chemistries), the sample can be cleaned up via aqueous acid and methanol washes, which leaves the drugs in a much cleaner state than when they were originally applied to the SPE column. This effect is noted in the low matrix effects and ion suppression values.  

Figure 4: Chromatogram of genuine hair demonstrating the presence of amphetamine only (upper trace). Lower trace indicates internal standards. See Table I for compound identification.			

In this new methodology, the SPE eluates are not evaporated to dryness as in typical SPE procedures, but are evaporated to a solution of mobile phase. Without the addition of methanolic hydrochloric acid to reduce their volatility, these drugs are known to be lost during this evaporation step. Using the mobile phase as a keeper solvent permits the volatile amphetamines and synthetic cathinones to be kept in solution and results in high recovery values by eliminating the loss during the evaporation. 

In this methodology, LC–MS-MS has been successfully applied to the extraction and analysis of amphetamine and synthetic cathinones rather than GC–MS in which a multistep derivatization procedure (that is, reaction with a fluoroacyl reagent such as heptafluorobutyric anhydride), evaporation, and reconstitution in a volatile solvent is required not only to quantify, but also to confirm the identity of the amphetamine or synthetic cathinone. By using LC–MS-MS with specific MRM values, amphetamine and synthetic cathinones can be targeted, confirmed, and quantified in hair samples without the use of derivatization. Coupling this procedure with a quick LC method offers analysts the ability of determine concentrations of the drug within a short turnaround time. 

Concentrations of Amphetamines and Synthetic Cathinones in Genuine Hair Sample 

An efficient method for forensic analysis of amphetamines and synthetic cathinones - the illicit drugs often called "bathsalts" - in hair samples

Analysis of Amphetamines and Synthetic Cathinones in Hair Samples Using LC–Tandem Mass Spectrometry and Solid-Phase Extraction

Daniel W. Armstrong to Receive ACS Award in Separation Science and Technology 2014

Daniel W. Armstrong, a professor of chemistry and biochemistry at the University of Texas Arlington in Arlington, Texas, is to receive the 2014 American Chemical Society Award in Separations Science and Technology. This is the second time this year that the ACS has recognized the work of Armstrong — in August last year he was selected to join the ACS Class of Fellows 2013. 

The award, sponsored by Waters Corporation, recognizes "the development of novel applications with major impacts and/or the practical implementations of modern advancements in the field of separation science and technology." It will be presented to Armstrong on March 17 at the ACS national conference in Dallas, Texas. 

Armstrong is thought of as a leader in his field and is considered to be the "father" of pseudo-phase separations — a type of chromatography that lowers costs, volatility, and toxicity while providing higher selectivity than other analytical methods. His commercialized innovations are applied across the drug development, environmental analysis, and petrochemical industries. 

In his career, Armstrong has developed more than 30 different chromatography columns — a gas chromatography column that he developed is now part of the Rosetta mission of the European Space Agency exploring the composition of comets in space. 

Purdue Research Foundation Invests in Two Start-up Companies

The Purdue Research Foundation (West Lafayette, Indiana) has awarded $150,000 in investment to two companies that have licensed Purdue University Innovations: Telos Discovery Systems Inc., and bioVidria Inc. The investment is from the Emerging Innovations Fund, which was set up in 2008 to accelerate the commercialization of early stage technologies in the Purdue community. 

The company bioVidria Inc., received $75,000. It is based on the work of Mary J. Wirth, the W. Brooks Fortune Distinguished Professor of Analytical Chemistry in the Department of Chemistry. The company focuses on developing new materials for chromatography, specifically protein analysis, for the pharmaceutical and agricultural industries. 

Telos Discovery Systems Inc., also received $75,000. The company is a boutique equipment manufacturer that supports basic and discovery medical research. The company is based on the work of Joseph Garner, a former Purdue associate professor in the Department of Animal Sciences. 

Elizabeth Hart-Wells, assistant vice president and director of the Purdue Office of Technology Commercialization said: "The fund makes seed investments and early-stage investments in the form of convertible debt and/or warrants, with a goal of accelerating the commercialization of early-stage innovations."

Departments

Daniel W. Armstrong, a professor of chemistry and biochemistry at the University of Texas Arlington in Arlington, Texas, is to receive the 2014 American Chemical Society Award in Separations Science and Technology.

Peaks of Interest

If scaling isocratic separations is so simple, why is gradient scaling so confusing?

In last month's "LC Troubleshooting" installment (1), we looked at how to scale isocratic separations when the column size or packing particle size is changed. The process is quite simple. First, find a column with approximately the same plate number, then adjust the flow rate to give an acceptable pressure. The most common problems that result from mistakes in this scaling process give somewhat lower resolution than is expected or higher pressures. With gradients, unexpected consequences may occur from the changes that may be relatively unimportant in isocratic methods. In this month's discussion, we turn our attention to the proper scaling of gradient methods. 

Last month (1) we looked at the fundamental resolution equation for isocratic conditions: 

 is the column plate number, α is the separation factor, and 

 is the retention factor. A similar equation can be stated for gradient separations: 

* is the effective plate number under gradient conditions, α* is the gradient separation factor, and 

* is the gradient retention factor. As with isocratic conditions, we must be careful to keep from changing the chemistry of the system by keeping the same brand and series of column packing, the same mobile phase, and the same column temperature. We'll see below that we have some additional factors to be careful of with gradients. If we keep these things constant, 

* values for two adjacent peaks) should remain constant. If α* is unchanged, we will obtain the same resolution if we keep the same column plate number. 

The column plate number cannot be measured easily under gradient conditions, so we measure it under isocratic conditions. Because the plate number is a characteristic of the column, the use of isocratic conditions is not a problem. We use the same approach we used with isocratic conditions to select a column with an equivalent plate number so that we maintain the same resolution with the scaled method. We saw that we could obtain approximately the same plate number if we kept the column length-to-particle-diameter ratio constant within a range of +50% to -25%. Thus, we can determine the desired column length from 

 are the particle diameters of the original and new column, respectively. So far, nothing is different between the isocratic and gradient scaling process. 

With isocratic separation, we saw that the flow rate had no influence on the retention factor, 

, and only a minor influence on the plate number, 

. So, although we often start the isocratic scaling process by keeping the linear velocity of the mobile phase the same, from a practical standpoint the flow rate is adjusted to maintain an acceptable pressure. With gradients, however, a change in the flow rate without some other compensating change can generate problems. This is because 

 is the flow rate (milliliters per minute), 

 is the column volume (milliliters), ΔΦ is the gradient range (for example, 5–95% = 0.9), and 

 is a constant related to the molecular weight and is characteristic of each analyte. For the present discussion on scaling separations, we will not be changing the gradient range or the analytes, so ΔΦ and 

 will be constant and can be dropped for a simpler relationship: 

With isocratic separations, the separation factor, α, is an expression of the peak spacing of two peaks with 

As long as we do not change the chemistry of the system (mobile phase composition or column chemistry) or the column temperature, α will remain constant, as will the peak spacing. Isocratic 

 is changed by nonchemical or temperature means, 

 stays constant. This is why we can change the column length, the column diameter, or the flow rate and not affect 

 is constant, α will also remain constant. On the other hand, if we do something that changes 

, we expect that α will change because it is rare for a variable that changes 

 in exactly the same manner for all the other compounds in the separation. Thus, a change in the mobile phase or column temperature usually results in a change in peak spacing — in fact, we count on this effect as a tool for moving peaks relative to each other during method development. 

A similar requirement of keeping α* constant holds for gradient separations, where α* is expressed as follows: 

As with isocratic separation, chemical or temperature changes are not likely to change 

 in the same way, so these variables must be kept constant with gradients too. But with gradients, the added complication is that the variables on the right-hand side of equation 5 must also be kept constant, or they should be changed in a manner that allows α* to stay constant. Next, let's consider how this requirement influences the scaling of gradient methods.  

We'll start with a simple linear gradient of 5–95% acetonitrile over 20 min at 1 mL/min with a 150 mm × 4.6 mm, 5-μm particle diameter (

) column. Let's say that we want to get the same separation under ultrahigh-pressure liquid chromatography (UHPLC) conditions on a 1.8-μm 

 column with an internal diameter of 2.1 mm. 

The first step is to find a column with approximately the same plate number. We can use equation 3 to help us. The new column length (

) should be (150 mm × 1.8 μm)/5 μm = 54 mm long. Because 54-mm columns aren't available, we can use any length between 41 mm (-25%) and 81 mm (+50%) and be within our -25% to +50% guidelines. I would probably choose a 50 mm × 2.1 mm column in the present case. 

As with isocratic separations, with gradients it usually is a good idea to start with a flow rate that gives us the same linear velocity. We use the same technique we discussed last month: 

 is the column internal diameter for columns 1 and 2. In the present case, the new flow rate should be 1 mL/min × (2.1 mm/4.6 mm)

  = 0.208 mL/min; we'll round this to 0.2 mL/min for convenience. So, our new method will run on a 50 mm × 2.1 mm, 1.8-μm 

Let's check to see if we are done. Because we want 

* to remain constant, we can restate equation 5 as follows: 

We can calculate each half of equation 10 for the proposed conditions and see how close we are. For the column volume, we'll use the approximation of (

), because these are the only two variables that change between the columns. For the original conditions, (20 × 1)/(150 × 4.6

) = 0.00630, and for the proposed conditions, (20 × 0.2)/(50 × 2.1

) = 0.0181. The value for the new conditions is approximately three times too large. Because we've already defined the column size and the flow rate, the only variable we can adjust to fix this problem is the gradient time. If we reduce the gradient time to 7 min, the calculated value drops to 0.0635, which is close enough. So the gradient equivalent to the original conditions is a 7-min gradient at 0.2 mL/min on a 50 mm × 2.1 mm, 1.8-μm 

 column. This means that the switch from the high performance liquid chromatography (HPLC) to the UHPLC column reduces the run time by approximately threefold (20 min to 7 min). 

But if we're running under UHPLC conditions, we can operate at higher pressures and because we're using sub-2-μm particles, flow rate will not affect 

, so a higher flow rate may help to further speed the separation. We use the same equation to calculate the system pressure as we did last month for isocratic conditions: 

 are the original and new pressures, respectively; all the other variables were defined above. Let's assume that the original method generated a 2000 psi (140 bar) back pressure. For the proposed conditions, 

  × (0.2/1) ≈ 4935 psi. For UHPLC, we probably can tolerate twice this pressure, and the easiest way to do this is to double the flow rate so that a new pressure of ~9875 psi would be expected. Wait! We've changed the flow rate, so equation 5 tells us that this will change 

*, and we may find a change in peak spacing — not something we want. So we need to make another compensating change using equation 5 as a guideline. If we double 

* will stay constant. Now our method would be a 3.5-min gradient at 0.4 mL/min on a 50 mm × 2.1 mm, 1.8-μm 

 column. This would reduce the original run time by approximately sixfold (20 min to 3.5 min). 

The example above used a simple, one-segment linear gradient. Let's add a bit of complication and see how a two-segment gradient is handled. We'll use a 5–50% gradient in 10 min followed by a 50–95% gradient in 5 min, the other conditions (column and flow rate) are the same as before. Let's scale this to the same 2.1-mm i.d., 1.8-μm 

 column. The same column, flow rate, and pressure scaling results will be obtained: scale to a 50 mm × 2.1 mm column operated at 0.2 mL/min resulting in a back pressure of ~4935 psi. 

When it comes to scaling the remaining gradient conditions, we proceed in the same manner as with a single-segment gradient, but we need to treat each segment as a separate gradient. Thus, we have two gradients to consider: 5–50%/10 min and 50–95%/5 min. Because we're not changing the gradient range, we can scale the gradient times so that equation 10 is balanced. For the first segment, the original conditions give (10 × 1)/(150 × 4.6

) = 0.00315 and the proposed conditions give (10 × 0.2)/(50 × 2.1

) = 0.00907, approximately three times too large. A change in the gradient time to 3.5 min gives a value of 0.00317. For the second segment, our original gives (5 × 1)/(150 × 4.6

) = 0.00158. We would expect the same gradient time ratio adjustment to hold for the second segment as for the first, so we can use it in our trial calculation. So we'll use (3.5/10) × 5 min = 1.75 min, rounded to 1.7 min for convenience: (1.7 × 0.2)/(50 × 2.1

) = 0.00154, which is close enough. Now our new method will be 5–50% in 3.5 min followed by 50–95% in 1.7 min at 0.2 mL/min on a 50 mm × 2.1 mm, 1.8-μm 

If we want to increase the flow rate to accommodate the higher pressure capability of a UHPLC system, we use the same technique as discussed above with equation 11, but remember to adjust the gradient time of each segment by a factor of two using equation 5 if you change the flow rate by a factor of two. 

Don't Forget to Adjust the Injection Volume

One thing I didn't mention in last month's discussion of isocratic scaling is the potential need to adjust the injection volume when the column size is changed. The same rules hold for both isocratic and gradient conditions. As a general rule, the injection volume should be adjusted in proportion to the column volume. We can use the following relationship as a guideline: 

 are the injection volumes of the original and new conditions, respectively. If our original method used a 25-μL injection, the new method should use an injection of (25 × 50 × 2.1

) = 1.75 μL, which we can round to 2 μL. 

Often the injection volume for the original method may not have been optimized, so although equation 12 gives the proper scaling, I suggest using an empirical test to determine the injection volume of the new method. I would try the calculated volume plus additional injections at twice and half the recommended volume. In the present case, I would try injections of 1, 2, and 4 (or 5) μL and observe the chromatogram. If there is no unacceptable degradation of the chromatogram, especially in the first part of the run, it is likely that the injection volume can be tolerated. I like to have a safety margin in my conditions, so the injection volume should not be so large that it is right on the edge of failure. This can be achieved by allowing a safety margin of 50–100%. For example, if in the above example, 5 μL still looked OK, try 8 or 10 μL. If these injection volumes are OK, 5 μL is safe. 

As you can see from the discussion above, when the number of segments in the gradient increases, the work in calculating new conditions increases because each gradient segment must be scaled individually. What happens when there is a curved (convex or concave) gradient? I recommend strongly against using this type of gradient, because they are hard to reproduce from one instrument to another, and very difficult to scale. I don't know of any way to scale such gradients, because they are essentially composed of an infinite number of very small segments with different slopes. So, if you have an existing curved gradient that needs to be scaled, use the single-segment rules and hope that the new gradient works. 

Those of you who are familiar with gradients will notice that I ignored the instrument dwell volume (gradient delay volume) in this discussion. The new dwell volume requirements can be calculated if the dwell time (the time to wash out the dwell volume) is considered as a gradient segment. The complication is that the dwell volume is seldom adjustable. Usually when you are moving a method from a conventional column to a small-volume or UHPLC column, you also will be moving from a conventional LC system to a UHPLC or a newer LC system, either of which is likely to have a reduced dwell volume. Dwell volume differences are most likely to affect the peak spacing for peaks eluted early in the gradient, so watch for such changes when the method is scaled. You may need to adjust the initial conditions to compensate for such changes. Dwell volume, of course, is not important in isocratic methods. 

Finally, as with the isocratic examples last month, gradient calculations are tedious. You can simplify the process by using one of the on-line calculators to do the work for you. Some of these also calculate scaling of the injection volume and take dwell volume into account. Search the internet for "HPLC method transfer calculator" and you'll find several to choose from. I suggest that you try several and settle on the one that seems like the easiest for you.  

"LC Troubleshooting" Editor John Dolan has been writing "LC Troubleshooting" for 

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

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

LC Troubleshooting

LC Method Scaling, Part II: Gradient Separations

In this installment, guest authors and capillary electrophoresis (CE) experts Breadmore and Sänger-van de Griend bring a practical perspective to enhancing the sensitivity of CE. They compare stacking and sweeping injection methods and provide practical suggestions about how to enhance sensitivity for charged and neutral compounds. 

One of the most frequently cited concerns in capillary electrophoresis (CE) is that the concentration limits of detection are inferior to what can be achieved with liquid chromatography (LC) (1). Extensive research has been carried out to overcome this limitation and now there are many different ways to increase sensitivity by controlling the way the sample is injected, how it is prepared, and how the targets are separated. After all, the concentration in the sample vial does not need to be the same as the concentration needed for detection if we can concentrate on-line. Early researchers in the field diluted samples in the electrolyte (parallel to dissolving or diluting in eluent in LC); however, doing so dilutes the target analytes, further reducing sensitivity. A better approach is to use the sample straight, which means that the composition of the sample plug is different from the composition of the electrolyte. Although intuitively this may not seem like a smart idea, depending on the exact analyte and conditions there may be a significant difference in their migration behavior in the sample zone compared to their migration behavior in the electrolyte. Very early researchers realized that this approach could enhance sensitivity. A brief search of the literature yields many excellent research papers and reviews (1–5), with different kinds of concentration mechanisms occurring because of different analyte, matrix, and electrolyte properties. Table I gives an overview of the most frequently used concentration techniques; as can be seen there are quite a number of them. It is worthwhile noting that these are the most common single concentration approaches used. It is possible to combine multiple mechanisms to produce even more powerful approaches; however, these are more complex. Most reviews discuss the various techniques from the mechanistic point of view. Here, we'd like to consider these options from a practical point of view and ask the question: I have XYZ in my sample; how do I improve my sensitivity? After all, in real life one usually cannot pick all conditions and sample matrices to suit a theoretically useful technique; instead, one has to work from the situation at hand. 

Table I: In-capillary sample concentration in CE			

First of all, the fundamental premise of concentration in CE comes back to creating a different velocity of the analytes of interest in the sample zone compared to their velocity in the background electrolyte (BGE) (6). There are many different techniques based on a change in velocity. Each technique has its own abbreviation, but the only difference may be a change in direction of the electroosmotic flow (EOF). These techniques fall into two groups: stacking techniques and sweeping techniques. If the velocity in the sample zone is higher than in the BGE (Figure 1b), the technique is generally called 

Stacking and sweeping injection methods are compared and practical suggestions are provided about how to enhance sensitivity for charged and neutral compounds.

LCGC North America-03-01-2014

Application of Pyrolysis–Gas Chromatography–Mass Spectrometry for the Identification of Polymeric Materials

Analysis of Amphetamines and Synthetic Cathinones in Hair Samples Using LC–Tandem Mass Spectrometry and Solid-Phase Extraction

Peaks of Interest

LC Method Scaling, Part II: Gradient Separations

In-Capillary Sample Concentration in CE

Mixed-Mode HPLC Separations: What, Why, and How

The Storage and Use of Gases for Gas Chromatography

Vol 32 No 3 LCGC North America March 2014 Regular Issue PDF