Solid-phase microextraction (SPME) was originally introduced as a solvent-free sample preparation technique ideally suited for analysis of fragrance compounds and environmental pollutants requiring gas chromatographic (GC) analysis. Introduction by thermal desorption into a standard GC injector helped to make this technique widely used. Its applicability for additional compound classes including pesticides, drugs and endogenous chemicals requiring liquid chromatographic (LC) analysis was demonstrated later, where the desorption of extracted analyte into a minimal volume of solvent compatible for injection into LC instrumentation was employed. Recently, new SPME sorbent phases have been introduced to further broaden the range of applications of the technology, and facilitate both on-site and 

 sampling and sample preparation from dynamic living systems. This article briefly introduces the theory relevant to such applications and provides examples of its use.

Solid-phase microextraction (SPME) is an extraction and pre-concentration technology developed to address the need for rapid sample preparation procedures both in the laboratory and in the field (1,2). In the past two decades it has found widespread use as a sample preparation tool for investigations using gas chromatography (GC) and liquid chromatography (LC). As a result of its simple design and operation, it has also enabled automation and high-throughput sample preparation for both types of separation.  

The basic concept of the technology in its current commercial configuration is a sorbent-coated rod that is put into contact with a sample (gaseous, liquid, semi-solid) or the headspace of liquids or solids. The sorbent is selected to have a good affinity with the analyte of interest in the sample. After a pre-defined exposure time, sufficient analyte will have moved from the sample to the sorbent to permit quantitative analysis. The amount extracted is proportional to the original concentration of analyte in the sample, which permits the easy determination of sample concentration (3).  

Although up till now SPME devices have been used mainly in laboratory applications, recent research has been directed towards field monitoring, particularly for clinical, environmental and industrial hygiene applications. A new line of biocompatible SPME devices has been specifically developed to better exploit these opportunities, particularly for 

 analysis (4). These devices use a C18 porous silica sorbent (similar to particles typically used in HPLC columns or as SPE sorbents) in a proprietary biocompatible binder [45 μm thickness, 15 mm length of the coating] (Supelco, Bellefonte, Pennsylvania, USA).

The binder used is a non-swelling polymer which resists fouling by biological matrix components (5). The solid support used for these probes is a flexible metal alloy (0.008" dia), and the coatings are housed inside a 22-gauge hypodermic needle with a seal incorporated to prevent sample loss during sampling. Because of the C18 extraction phase, they perform absorptive extraction.  

The devices may be employed for single use and are ideally suited for application, either 

to sample directly from whole blood or plasma in sample vials sealed with hole caps and septa, or through an injection bulb on an intravenous catheter for 

studies. Devices without the attached hypodermic needle are also available where sealing is not critical (for example, tissue sampling or sampling from open vials). The operating principles of such devices are analogous to those of the original devices. The modifications are made for greater convenience in given applications. 

An important advantage of SPME, particularly for on-site sampling, is the possibility of performing analyses without pre-defining a specific sample size. The reason for this may be understood from a brief review of SPME theory. The amount of an analyte extracted from a sample by SPME at equilibrium is given in Equation 1 (6):  

 is the initial sample concentration of the analyte, 

 is the amount of analyte extracted by the device, 

 is the analyte distribution constant between the sorbent and sample matrix. However, when sample size is large relative to the fibre capacity, the amount extracted is insignificant relative to the amount present in the sample (that is, conditions of negligible depletion). Specifically this occurs when 

. In this case Equation 1 reduces to Equation 2. This renders the amount of analyte extracted by SPME independent of the sample volume.  

This allows for the application of SPME to on-site analysis and eliminates the need to remove a representative sample from the system under study to perform the analysis. From a bioanalytical perspective, this permits the use of SPME to directly sample blood or tissue of animals

, without having to withdraw an appropriate biofluid/tissue sample. Several applications for 

SPME analysis where this benefit has been exploited in animals have appeared in recent years. 

A determination of the magnitude of sample volume required to satisfy this condition may be made from calibration data using Equation 1. Where mass extracted is plotted against sample concentration, the fibre constant (

) may be derived from the slope of the resulting linear regression curve.  

Selection of the extraction time should also be given some consideration. While full equilibrium is practical for gas sample extractions, where diffusion coefficients in the sample are high, it is commonly impractical for extractions from liquid samples. This is particularly true for 

analysis, where a very short (pre-equilibrium) extraction time is usually preferred. Very short extraction times typically fall in the kinetic region of the adsorption time uptake profile. 

extraction calibration because in this region uptake is controlled by diffusion across the boundary layer rather than the kinetics and/or thermodynamics of sorbent interactions. In such cases, calibration of mass uptake relative to sample concentration may be performed by monitoring the loss of an internal standard pre-loaded onto the sorbent.  

Where uptake kinetics are controlled by the boundary layer diffusion, there is a symmetry between analyte uptake and standard loss. Although it is almost always valid, it is prudent to verify that the relationship holds for a particular experiment. In our experience, this symmetry only breaks down in a few rare cases where the kinetics of analyte interaction with the sorbent surface are very slow and the boundary layer very thin. In the majority of cases, loss of pre-loaded internal standard from the sorbent may be used to control for variable extraction time and sample convection conditions, as well as providing for calibration of sample concentration.  

 SPME studies were applied to study the pharmacokinetics (PK) of various drugs in directly from the veins of animals (7–12). Specifically SPME devices based on polypyrrole and polyethylene glycol coatings were used for direct extraction of diazepam and its metabolites form the flowing blood of beagle dogs (13). LC coupled to tandem mass spectrometry (LC–MS–MS) was used for analysis. Two calibration strategies, external and kinetic calibration, were used to correlate the extracted amount of analytes with the 

 concentrations. The probes were exposed to the flowing blood for 2 min for equilibrium extraction and 30 s for kinetic calibration. The fast microextraction technique caused minimal disturbances to the drug/protein binding equilibria so additional information on the free concentration could be obtained. These studies exhibited the quantitative capability of SPME for 

Although rats are extensively used for traditional PK studies in drug discovery, there is an increased interest in the use of mice because of the availability of various strains of gene knockout mice and indications that mouse data may be more useful than rat data for allometric scaling of PK parameters from animals to humans (9). Traditional PK studies in mice rely on the sacrifice of multiple animals at each time point of interest, which introduces inter-animal variation and dosing inaccuracies into the PK data. This disadvantage has been addressed by performing studies where all samples are acquired from a single animal by serial blood sample withdrawal, but to date such studies have been difficult to execute in mice because a limited circulating blood volume (< 2 mL/25 g mouse) and difficulties in the handling and sample preparation of the small blood sample volumes.  

A specialized sampling interface was developed to permit monitoring of small rodents (mice and rats) (11, 12, 14) and the blood draw-free SPME technique was applied for 

 pharmacokinetic studies drugs directly in the bloodstream (8). The SPME probes were inserted into the surgically-implanted interface through which blood was allowed to circulate. The main advantages of single rodent PK studies include: (1) a reduction in animal use which is particularly important when using rare or expensive strains of mice; (2) a reduction of dosing errors in the data; (3) a decrease in the amount of compound needed to perform the study, which is particularly advantageous in early drug discovery where the quantity of test compound is limited; (4) elimination of the effects of anesthesia from the PK data; (5) the ability to study inter-animal variation; and (6) the opportunity to perform multiple and/or simultaneous pharmacokinetics/pharmacodynamics investigations in the same mice (10–12, 14). The PK data obtained from serial SPME sampling was equivalent to the data obtained using terminal sampling from multiple animals, indicating the validity of this approach (12).  

SPME technique with conventional SPME fibres was successfully used for the detection of toluene levels in the brains of conscious, free-moving mice (15, 16). In order to measure the pharmacokinetics of inhaled toluene in the brains of mice, a SPME fibre was inserted into the hippocampus (CA1) through a cannula fixed onto the animal. BALB/c mice were exposed to toluene for 30 min. The pharmacokinetics of toluene in the brain of mice exposed to 50  ppm toluene showed that the toluene level decreased rapidly after the exposure, and returned to control levels after 60 min. The technique does not require lethal animal testing, and has the potential to become a useful tool for studies on the field of neurotoxicology (16).  

We have also recently studied the use of the new SPME fibres for monitoring brain function (17). Probes with a short (4 mm) sorbent length were prepared and used to monitor selected neurotransmitter levels in response to electrical stimulation of the brain. The SPME probes were positioned in the target brain region (hippocampus) through a surgically implanted microdialysis cannula, similar to the positioning of a microdialysis probe. The neurotransmitters studied were: serotonin, dopamine, γ-aminobutyric acid and glutamic acid and these were detected by LC–MS–MS. No significant changes were observed for the studied neurotransmitters before and during electrical stimulation, except for serotonin, which showed an approxiamte 2–3× (~ 50 pg/mL–400 pg/mL) increase during deep brain stimulation (DBS). Relative standard deviations (RSD) for serotonin at basal concentrations ranged from 3–20%. The results were equivalent in the target brain region in both the left and right hemispheres of all four rats studied. The SPME method addresses some of the limitations of microdialysis for these analyses, such as better sensitivity for a given sampling time, the unrestricted movement of the animals during sampling as a result of the lack of peripheral tubing, as well as the elimination of the significant signal suppression as a result of matrix components of the dialysate in microdialysis.  

The increase in serotonin after DBS in the present study was similar in magnitude to previous results obtained by Hamani et al. using microdialysis (18). This suggests that SPME as a tissue sampling tool may be a valid alternative approach to measure neurotransmitter changes after particular interventions. 

Over the past five years, we have applied SPME to the 

 determination of pharmaceuticals in fish tissue using direct muscle sampling (19–22). A significant advantage of SPME fibres is their ability to extract a variety of trace contaminants from fish tissues without lethal sampling. This means it is possible to monitor the presence of a target analyte in the same fish over several days. We have also evaluated the use of the devices for analysis of fish fillets after lethal sampling. In either case, sampling involves using a hypodermic needle to create a small track in the tissue followed by inserting the SPME sampler so that the sorbent is in direct contact with the target tissue for a few minutes.  

This is followed by withdrawing the needle and measuring the chemicals removed by the sorbent after transport to the laboratory. By pre-loading internal standard onto the fibre we are able to determine the concentrations of the chemicals in the tissue based on the loss of standard from the fibre during sampling. We have demonstrated repeatedly that the results obtained have the same accuracy as other more conventional methods of analysis (12, 7, 23). The system is much simpler to use than conventional analysis, thus field technicians do not require advanced training in chemical analysis. Laboratory-based steps are also greatly simplified. The method has been demonstrated for the analysis of pharmaceuticals to low to moderate ng/g levels in greenside darter, fathead minnow and rainbow trout (24). We have also demonstrated a method to calculate bioaccumulation factors for different species and pollutants (19, 20, 25).  

Currently there is an increased interest in the development of global or untargeted metabolomic profiling methods which can capture hundreds or thousands of metabolites in a single liquid chromatography–mass spectrometry (LC–MS) run. The choice of sampling, sample collection and sample preparation strategies plays an important role in the quality of metabolomics data during these studies (26). Issues with incomplete metabolism quenching, ionization suppression and metabolite instability have been well-documented and can adversely impact data interpretation. 

 solid-phase microextraction (SPME) is an attractive sampling method for global metabolomics studies by LC–MS.  

The potential benefits of SPME for metabolomics studies, such as the capture of short-lived and unstable metabolites and good sample throughput, have been demonstrated for the detection of biotransformation intermediates and both global and targeted metabolomics studies using GC–MS primarily in headspace mode (27–33). However, to date SPME in direct extraction mode in combination with LC–MS analysis has been used in the main for the analysis of a small number of 

 selected target compounds (23, 34). The main difficulty with applying these existing workflows to envisioned global metabolomics studies in complex biological matrices, such as blood or tissue, is the lack of coatings that can simultaneously extract both polar and nonpolar species using direct extraction mode. In fact, only four SPME coatings have been available commercially until now: poly(dimethylsiloxane) (PDMS), poly(dimethylsiloxane/divinylbenzene) (PDMS/DVB), polyacrylate (PA) and Carbowax-templated resin (CW-TPR). They exhibit poor extraction efficiency toward polar analytes, can suffer from the fouling of the extraction phase upon exposure to biological samples and are too expensive, which makes the use of multiple fibres during large-scale metabolomics studies cost prohibitive.  

We have recently reported on the use of SPME in direct extraction mode for untargeted metabolite profiling studies in combination with LC–MS using the new generation biocompatible SPME probes (35). We have shown that mixed-mode, phenylboronic acid and polystyrene-divinylbenzene coatings are particularly suitable for this type of application. We have demonstrated for the first time that it is possible to extract hundreds of endogenous metabolites with a single coated fibre, whereas previously all existing reports of direct immersion SPME deal with targeted analysis of a few 

studies on human biological fluids, SPME can serve as a complementary technology to the widely used solvent precipitation approaches. It has been shown to provide improved quantitative information on polar species because of a reduction in ionization suppression, complementary metabolite coverage and information on the biologically relevant free concentration. In contrast to ultrafiltration, SPME reduces ionization suppression and improves coverage of hydrophobic species. These are typically highly bound and may not be observed by ultrafiltration either as a result of adsorption to the membrane or a high degree of protein binding. Further work is currently underway to exploit the 

 sampling capability of SPME and demonstrate that SPME can more accurately capture the metabolome at the time of sampling, in particular for the unstable metabolites and/or metabolites with fast turnover rates. 

SPME offers important advantages over conventional methods, such as simplified sample clean-up, fast stabilization of unstable analytes, elimination of enzymatic degradation after extraction and reduced ion suppression for mass spectrometry analyses. Furthermore, because both sampling and sample clean-up are combined into one step, the number of sample preparation steps, which can potentially result in loss of analyte or accidental contamination, is minimized (9). A recent article in 

 details all of the steps involved in performing 

 SPME for intravenous drug and metabolite monitoring (36).  

 is a research associate in the group of Professor Pawliszyn with experience in analytical device and method development, sorbent design and solid-phase sample preparation. She has authored over 40 scientific papers in biochemical toxicology and bioanalytical method development. Her primary research interests are in the areas of 

sample preparation and new bioanalytical methods development. 

 research programme is the design of highly automated and integrated instrumentation for the isolation of analytes from complex matrices and the subsequent separation, identification and determination of these species. Currently, his research is focused on the elimination of organic solvents from the sample preparation step to facilitate on-site monitoring and 

analysis. He is also exploring the application of computational and modelling techniques to enhance performance of sample preparation, as well as the development and application of imaging detection combined with microseparation approaches. 

(4) D. Vuckovic, R. Shirey, Y. Chen, L. Sidisky, C. Aurand, K. Stenerson and J. Pawliszyn, 

(5) L. Musteata, M. Musteata and J. Pawliszyn, 

Solid phase microextraction, theory and practice

, J. Pawliszyn, Ed.; Wiley-WCH, Inc.: USA (1997).  

(7) H.L. Lord, R.P. Grant, M. Walles, B. Incledon, B. Fahie and J.B. Pawliszyn, 

(8) F.M. Musteata, M.L. Musteata and J. Pawliszyn, 

(9) X. Zhang, A. Es-Haghi, F.M. Musteata, G. Ouyang and J. Pawliszyn, 

(10) A. Es-Haghi, X. Zhang, F.M. Musteata, H. Bagheri and J. Pawliszyn, 

(11) F.M. Musteata, I. de Lannoy, B. Gien and J. Pawliszyn, 

(12) D. Vuckovic, I. de Lannoy, B. Gien, Y. Yang, M.F. Musteata and J. Pawliszyn, 

(13) F.M. Musteata, M.L. Musteata and J. Pawliszyn, 

(14) D. Vuckovic, X. Zhang, E. Cudjoe and J. Pawliszyn, 

(15) T.T. Win-Shwe, D. Mitsushima, D. Nakajima, S. Ahmed, S. Yamamoto, S. Tsukahara, M. Kakeyama, S. Goto and H. Fujimaki, 

(16) D. Nakajima, T.T. Win-Shwe, M. Kakeyama, H. Fujimaki and S. Goto, 

Articles

The use of in vivo SPME offers important advantages over conventional methods according to the authors, including simplified sample clean-up, fast stabilization of unstable analytes, elimination of enzymatic degradation after extraction and reduced ion suppression for mass spectrometry analyses.

Introduction to SPME for In Vivo Analysis

In this instalment we review gas chromatography (GC) instruments and accessories that were newly presented at Pittcon 2012 or were introduced to the marketplace in the preceding year.

The 2012 Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy (Pittcon) was held 11–15 March, in Orlando, Florida, USA, at the Orange County Convention Centre. The 63rd annual occurrence of this venerable analytical chemists' symposium was only slightly less attended than the conference's 2010 southern sojourn at the same location, with 15754 registrants. I'm comparing the attendance to that at the previous Orlando venue rather than Atlanta from 2011 because the attendance always has been somewhat lower when the conference heads south. The amount of exhibitor booth space also declined by a similar margin, driven by many companies' decisions to attend with a smaller footprint, although the total number of exhibiting companies only dropped by about 2%. 

Significantly absent this year were Agilent and PerkinElmer, which are both engaging in post-conference seminar tours and on-line product displays. Both companies exhibited at Analytica in Münich, Germany, 17–20 April, 2012. Several companies opted for nearby hotel suites to meet key customers at rather than hosting an in-exhibition booth. To an aisle wanderer such as myself, the overall impression was one of reduced acreage accompanied by a larger Technology Park area and, although I did not measure them, perhaps slightly wider corridors. Smaller though it may have been, the exhibit floor experience was almost relaxed, with more time to visit each exhibitor of interest, unlike some earlier years where I often had to choose between visiting a booth or attending a session. 

The conference organizers' emphasis on the technical programme was evident in the consistent number of poster sessions that were mounted in two large areas of the exhibition floor, as well as in the continuing high count of oral presentations, symposia and workshops. Short-course attendance was up 16% from last year.  

Among the conferees, 36% listed liquid chromatography (LC) and 34% gas chromatography (GC) among their selected scientific specialties; these were the top two categories at the conference. Spectroscopic specialties accounted for the majority of the remaining mentions; ion chromatography was cited by 11%, and other chromatography techniques were cited by 7%. 

Pittcon 2013 will be held in Philadelphia for the first time, 17–21 March, at the Pennsylvania Convention Centre. The conference is scheduled to return to Chicago (2014, 2017), New Orleans (2015) and Atlanta (2016), before returning to Orlando in 2018. 

This instalment is our annual review of GC instrumentation and accessories shown at this year's Pittcon or introduced during the previous year. For a review of new GC and LC chromatography columns and related accessories, please see Ron Majors' "Column Watch" in the April and May 2012 issues of 

The information presented here is based on manufacturers' replies to questionnaires, as well as on additional information from manufacturers' press releases, websites and product literature, and not upon my actual use or experience. During Pittcon, I took time to stroll around the convention aisles and see some of the new products firsthand, as well as discover a number of items that weren't covered by the questionnaires. Every effort has been made to collect accurate information, but because of the preliminary nature of some of the material 

 cannot be responsible for errors or omissions. This column cannot be considered a complete record of all new GC products introduced this year at Pittcon or elsewhere because not all manufacturers chose to respond to the questionnaire, nor is all of the submitted information necessarily included here because of the limited available space and the editors' judgment as to its suitability.  

Another new crop of GC instruments appeared in 2012, which demonstrates once again the ongoing resilience of the technique and also emphasizes a continuing shift toward mass-selective detection since what used to be top-end capabilities have become more affordable. Nearly half of the newly introduced instruments are tightly associated with a mass spectrometric (MS) detector. Most were more about the MS detector than the GC itself. See Table 2 for the details of each new product.  

Shimazdu's GCMS-QP2010 SE system is an upgraded version of the company's QP2010 S system that features high-performance ion optics and a quadrupole mass filter along with lab-friendly energy- and time-saving features. While not seen at Pittcon, PerkinElmer introduced the Clarus SQ 8 GC/Mass Spectrometer system with high sensitivity, tool-free source maintenance and a rapid scanning speed earlier this year. Bruker Daltonics highlighted its high-end Scion TQ GC–MS-MS system with 180° optics that enables multiple ion-selective scan modes for resolution of complex samples and matrices. This triple-quadrupole GC–MS system, introduced in July 2011, won the 2012 Pittcon Editors' Silver Award. All of these new GC–MS systems operate in tandem with the companies' respective existing gas chromatograph offerings. 

From Torion, the Tridion-9 GC-TMS system is a gas chromatograph–toroidal mass spectrometer system that is aimed at the rapid screening of chemicals such as environmental volatile organic compounds (VOCs) and semivolatile organic compounds (SVOCs), explosives, chemical threats and hazardous chemicals. The unit is billed as a "Person-Portable" GC–MS system that can easily be carried to remote sites. It uses a conventional 100-μm i.d. capillary column bundled with a low-thermal-mass resistive heater and helium carrier gas. Sample injection is accomplished via the company's Custodion solid-phase microextraction (SPME) syringe. 

Thermo Fisher Scientific came to Pittcon with the 1300-Series GC system, a completely new frame. With a unique system of "instant-connect" modular interchangeable inlet and detector options, the 1300-Series system allows end users to quickly exchange or replace either or both inlets or detectors as pluggable modules. Each module includes pneumatic and electronic connections that are engaged by putting them in place and tightening a couple of screws. 

PAC introduced the AC Analytical Controls CNS SIMDIS for Crude Oil analyser. The system simultaneously determines the boiling range distribution of carbon, nitrogen and sulphur in crude oils and final products, using simulated distillation and a combination of flame ionization detection (FID) and chemiluminescence detection. 

A unique portable GC instrument was on display at the 2012 Pittcon. The Frog-4000 system from Defiant Technologies is a miniaturized GC system with photoionization detection (PID). Based on micromachined preconcentrator and GC designs, the Frog-4000 system targets field analysis of VOCs in water and soil.  

This year saw strong growth in new automated headspace GC (HSGC) samplers. Together with Agilent's 7697A headspace sampler that was introduced at Pittcon 2011, this burgeoning of HSGC is indicative of the ongoing demand for volatiles analysis in solid, semisolid and liquid matrices such as for pharmaceutical, food or flavour products. Table 3 lists the salient details.  

Table 3: Sampling systems and accessories.			

The new offerings bring a wide range of sizes and capabilities. The SHS-40 headspace sampler from Bruker Daltonics is a multivial equilibrium headspace autosampler for the company's 400-series GC systems as well as the Scion Series GC–MS system. It accommodates up to 40 vials and supports a range of modes and capabilities. The Master SHS static headspace sampling system from Dani is designed to work with the company's Master GC system, but it also interfaces with most other GC systems. The Master SHS system has a 120-vial capacity and a range of options such as a bar-code reader. The Versa automated headspace vial sampler from Teldyne-Tekmar is a compact device that holds up to 20 vials and works with most GC systems as well. 

A related application-specific headspace sampler, EST Analytical's Centurion RSK50, performs US EPA RSK 175 analysis for dissolved methane, ethane and ethylene in water, including sample preparation of the requisite 40-mL volatile organics analysis (VOA) vial and sampling of the evolved headspace gases onto nearly any model of GC system. 

Two new cannister sampling devices were on-hand. Markes International introduced its CIA Advantage canister sampling system in 2011, after previewing it at last year's Pittcon. The cryogen-free sampler handles multiple cannisters and is fully compliant with US EPA TO-15. From Entech, the 7200 Automated Preconcentrater uses a three-stage trapping procedure to preconcentrate gaseous samples from a variety of sources, add internal standards or spikes and like the Markes unit it works with most GC systems. 

Two new liquid GC autosampler systems were shown this year. HTA s.r.l. displayed its model HT3100A autoinjector, HT3000A autosampler and HT3200A top-class autosampler. Together, the three models deliver a wide range of capabilities to almost any GC system. From Thermo Fisher Scientific, the TriPlus RSH autosampler is a robotic 

 axis sample handling system and autoinjector with all of the flexibility and versatility that comes with this type of device. One TriPlus RSH autosampler can address up to two GC systems with sample preparation, derivatization and micro-volume injection among many sampling options. 

Finally, Gerstel introduced two accessories for its robotic Multi-Purpose Sampler (MPS). The mVAP MultiPosition Evaporation Station concentrates up to six samples at user-defined temperatures and vacuum levels. Solvent exchange, solid-phase extraction and filtration are a few of the possible sample preparation techniques. Gerstel's mVORX vortex and shaker accessory for its MPS system agitates up to eight sample vials in the horizontal plane to eliminate vial-cap contamination and over-mixing of microsized samples. 

I would like to thank the manufacturers and distributors that kindly furnished the requested information before, during and after Pittcon 2012, allowing a timely report on new product introductions. For those manufacturers who did not receive a preconference questionnaire this year and would like to receive one and be considered for early inclusion into Pittcon 2013 coverage, please send the name of the primary company contact, the mailing address, fax number and e-mail address to Laura Bush, Editorial Director, 

, Advanstar Communications, 485F US 1 South, Iselin, New Jersey 08830, USA, Attn: 2013 New GC Products. 

 is senior research scientist at BPL Global, Ltd., Hillsboro, Oregon, USA, and a member of

 editorial advisory board. Direct correspondence about this column should go to 

 editor, Alasdair Matheson, 4A Bridgegate Pavilion, Chester Business Park, Wrexham Road, Chester, CH4 9QH, UK, or email 

GC Connections

In this instalment John V. Hinshaw reviews gas chromatography instruments and accessories that were newly presented at Pittcon 2012 or were introduced to the marketplace in the preceding year.

New Gas Chromatography Products 2012

Hyaluronic acid (HA) is an ubiquitous, very high molar mass polysaccharide that has been of particular importance in opthalmic surgery. HA acts as a molecular "shock-absorber" and stabilizer for cells and its visco-elastic properties are valuable for separating tissue and maintaining shape. It is a critical component in tissue lubrication and is believed to play a leading role in wound repair. Finally, HA's property of non-pyrogenicity makes it an ideal sheath for implants, whose presence might cause the body to suffer an immune response.			

Among the many benefits high molecular weight HA hold are											

the maintenance of tissue space for surgery

HA's therapeutic effectiveness depends critically on molecular weight: the higher the molecular weight, the longer its benefit. However, because of its visco-elastic properties, standards-based GPC analysis is inappropriate for characterizing HA. There exist no standards identical to HA and the desirability of altering experimental conditions renders conventional GPC/SEC impractical. 

 Figure 1: From the molar mass versus time plot, subtle differences can be seen among the samples.			

Combining a DAWN with HPLC separation, however, provides an ideal platform for absolute characterization, since the lightscattering measurements do not depend on pump speed, polymer standards or molecular conformation. 

Figure 2: The differential molar mass distributions calculated by ASTRA immediately confirm the large differences among the three HA samples.			

A DAWN was connected to a GPC/SEC line (100 mM NaN03 buffer, TSK-Gel G6000PW column, Optilab DSP refractometer, Waters 510 pump) and generated the data required to determine not only absolute molar mass, but also molecular size, for a variety of HA products. 

Figure 1 shows the molar mass versus time (with the 90° light-scattering chromatograms in the background) for the three samples, ranging from approximately 2 million to less than 200K Daltons. Figure 2 illustrates how profoundly different the samples are by revealing their differential molar mass distributions. These indicate that the samples will behave in different ways when used medically, depending on the content of their high molecular weight HA. 

Santa Barbara, California 93117, USA,				

				tel. +1 805 681 9009, fax +1 805 681 0123,			

Advertisement Feature

Hyaluronic Acid (Polysaccharides)

Phenol formaldehyde resins are synthesized by a step-growth polymerization. GPC/SEC with RI and UV detection is often used to characterize resins and to quantify the amount of different oligomeric species. Additional information can be obtained if an ESI-MS-spectrometer is added in-inline to the GPC/SEC system. This technique combines the separation ability of GPC/SEC with the sensitivity and specificity of detection from MS and allows the identification of oligomeric species and gives information about the degree of CH

GPC/SEC analysis was performed on a PSS SECcurity 1200 System consisting of: 									

an autosampler with variable injection volume

Figure 1: RI trace oligomeric phenol formaldehyed resin, degree of polymerization identified from corresponding mass spectrum.						

			Columns:  PSS SDV, 5 μm, 8 × 300 mm; 50 Å, 100 Å,    1000 Å, precolumn  

Software:  PSS WinGPC Unity 7.4, Thermo Fisher   Excalibur 2.07 

Figure 2: Mass spectrum for n = 1 showing the 3 different species (CH2 âOH substitution).			

Figure 1 shows the elugram of an oligomeric phenol formaldehyde resin, separated into three different peaks with 1, 2 and 3 repeat units. The mass spectra are then measured for each species. Figure 2 shows as an example the mass spectrum for peak n = 1 while Table 1 summarizes the masses possible in theory and identified (green) or not detected (grey). This combined approach can be used for molar masses up to approx. 2000 Da, higher degrees of polymerization have also been investigated.  

Table 1: Expected molar masses for different degrees of polymerization (n) and for CH2OH units at the aromatic ring. Colour code: grey: possible in theory but not found, green: identified in MS spectrum.			

In der Dalheimer Wiese 5, D-55120 Mainz, Germany  

tel. +49 6131 96239 50 fax +49 6131 96239 11  

Polymer Standards Service GmbH Application Note

Absolute Molar Masses for Phenol Formaldehyde Resins with GPC/SEC-ESI

What happens when the same method gives different retention times on different instruments?

This month's instalment comes from a question submitted by a regular reader (Justin Chow), which developed into a joint troubleshooting project. Before we get into the question, let me remind and encourage you to send me questions about problems you are having (see contact information at the end of this article). I'm happy to give you my input, and who knows your problem may form the basis of one of the "

" discussions, as is the case this month.  

The problem concerns the differences in retention time that were observed when the same method was run on three different brands of liquid chromatographs, which we'll refer to as system A, system B and system C. As is often the case, the method was one that was transferred from another laboratory, so complete details about the logic behind the method are not available. The method comprises an isocratic segment for 62 min at 95% A and 5% B followed by a 23-min gradient segment of 5–95% B. Finally, the mobile phase is stepped back to the initial conditions and allowed to equilibrate for the next injection. Both mobile-phase A and B contain buffer and organic solvent, with more organic in B. Note that both buffers A and B are prepared using volumetric glassware to eliminate variations from graduated cylinders. The column is a 150 mm × 3.0 mm, 2.6-μm particle diameter (

), C18 column operated at 0.6 mL/min. As part of the method transfer process, the same method was used to compare the chromatography on the three LC systems, all of which are four-solvent, low-pressure mixing systems. The dwell volumes for all systems are approximately the same. The main peak gave different retention times on each system: 49 min (system A), 55 min (system B) and 61 min (system C). Although the variability between systems A and B (49 vs. 55 min) is within the range observed by workers in other laboratories, the retention time for system C (61 min) is excessively long. Furthermore, latereluted peaks that normally appeared in the isocratic segment now came off the column during the gradient segment and were not adequately separated. The problem is, of course, why are the retention differences observed, how can the problem source be verified and how can it be fixed? This issue also serves to illustrate how we can use some simple calculations to evaluate the probable cause of the problem.  

It is slightly unusual to have such a long isocratic portion in the method, but it works for the present application and is part of a validated method that is best not modified, if possible. However, it should be noted that the peak properties will suffer from such large retention times. The column plate number, 

 are the retention time and baseline peak width, respectively. We can estimate the plate number for a column when real samples are run as  

 is the column length (in millimetres) and 

 is the particle diameter (in micrometres). So for our 150 mm × 3.0 mm, 2.6-μm 

 = 1.5 min or 0.9 mL at 0.6 mL/min flow rate. A peak eluted at 10 min would have one-fifth this width and consequently five times the height — not a small change in response. Therefore, the long retention time means that method sensitivity (peak height) may be poor. 

Another check we can make on the quality of the chromatography is to consider the retention factor, 

. For the best isocratic methods, we like 2 ≤ 

 ≤ 10, but in some cases all the peaks won't fit within this range so we accept 1 ≤ 

 ≤ 20. When the retention range exceeds this, a gradient method is usually preferred. Recall that 

 is the column dead time. The column volume can be estimated as  

 is the column volume (in millilitres) and

 is the column internal diameter (in millimetres). Our 150 mm ×  3.0 mm column will have a volume of ≈ 0.675 mL, and a flow rate of 0.6 mL/min, 

 = (50  – 1.125)/1.125 = 43; this is much larger than the maximum desired 

 ≈ 20, so a gradient method probably would be a better choice. But we have to work with the current method to separate some very closely related compounds, so let's try how to figure out what's wrong. 

Because the same method is used on all three instruments, one possible cause of the problem is that the systems are not producing the same blend of mobile phase. We can estimate how much the mobile phase will change retention by taking advantage of the relationship between mobile phase strength and retention:  

 is the retention time in 100% water and Φ is the %B as a decimal (Φ = 0.01 = 1% B). 

 is a constant for a given compound and can be estimated as 

, where MW is the molecular weight. If we assume a 400 Da compound, we can use 

 = 5 for our present calculations. Let's consider two cases, first the 49 min (

 ≈ 47.9, system B) "normal" variation and the unexpected 52 min (

 ≈ 45.2, average for systems A and B) vs. 61 min (

For the first case, log 42.6 = 1.629 and log 47.9 = 1.680, so Δ log 

 = (1.680 – 1.629) = 0.05 (note that I've rounded the numbers for convenience, so if you try to reproduce these calculations, don't round during the process). Divide by 

 and we get ΔΦ = 0.010. This suggests that systems A and B differ by 1% B, which is marginally within the normal calibration range of ±1% B (see discussion below). For the second case, log 45.2 = 1.655 and log 53.2 = 1.726, so Δ log 

 = (1.726 – 1.655) = 0.070, so ΔΦ = 0.070/5 or 1.4% B. In other words, we can account for the difference between the longest retention time (61 min, system C) and the average retention from systems A and B (52 min) by a difference of 0.4% B in the mobile phase composition. This small variation is probably within normal error, but it is possible that system C may be out of calibration. Next, we need to check the calibration of the mixing system for system C. 

A check of the calibration of the mobile-phase mixing system is part of system performance qualification (1), which we recommend performing every 6–12 months on each LC system in your laboratory. The mixing can be tested quite easily with the gradient step-test. This test is performed as follows:  

Remove the column and replace it with a piece of capillary tubing; ≈ 1 m of 0.125-mm (0.005-in.) i.d. tubing, such as the red PEEK (polyetheretherketone) tubing, which works well. This has an insignificant volume (≈ 12 μL) and generates enough back pressure that the pump works well. 

Replace the A-solvent with water and the B-solvent with water containing 0.1% acetone. If you want to check the C- and D-solvents, put the C-inlet tubing in the A reservoir and D in B. 

Set the flow rate to 2 mL/min and the UV detector wavelength to 265 nm, where acetone absorbs strongly. 

Programme and run a series of isocratic steps, 4 min each. Steps of 0, 10, 20, 30, 40, 50, 60, 70, 80, 90 and 100% B work well.  

The test should be run for each LC system and each combination of solvents that might be used for a method. Currently we're only concerned about the A- and B-solvents, but C and D should be tested as well. 

The results for a test of system C are shown in Figure 1. Measure the difference between the 0% and 100% steps and use this to determine the height of each intermediate step. In Figure 1, you can see that most of the steps vary a little from the target values (even 10% increments), but the worst deviation is the 20% step, which comes out 0.6% low at 19.4%. Pump specifications typically are ± 0.5–1%, so ± 1% is a reasonable acceptance limit for this test. Everything looks OK for system C from the step test.  

Figure 1: Gradient step-test results for system C using 10% steps from 0% B to 100% B; actual composition shown for each step. 

For cases in which subtle solvent proportioning errors are suspected, such as in the present example, it may make sense to make finer step sizes around the suspect region — 0–10% in 1% steps in this case. The results are shown in Figure 2. The absolute %B obtained is shown above each step and the actual step size shown to the right of each step. Here, as in the 10% steps of Figure 1, the absolute value of each of the steps is within 1% of the set-point, so the instrument passes specifications in the 0–10% region (note that from Figure 1, we know that this is 0–9.8%; Figure 2 reflects this). Furthermore, the anticipated 0.4% deviation at 5% B, determined from the earlier calculations, is not observed. The 5% step in Figure 2 measures 4.8%, only 0.2% from the ideal. However, the step sizes in Figure 2 are a bit odd. Note in Figure 1 that all the steps except 0 and 100% are low by 0.1–0.6%, with a general trend of larger deviations at lower %B-values. But in Figure 2, the step 1–2 and 2–3 intervals are 0.8% and the step 0–1 and 3–4 intervals are 1.2%. This inconsistency in step sizes was the only difference that could be linked to system C, so the proportioning valve assembly was replaced as a precautionary measure. System C with the new proportioning valve gave a retention time of 57.7 min for the main peak, which was still slightly longer than the other laboratories were finding. However, the poorly separated peaks that were eluted during the early part of the gradient segment are now eluted in the isocratic separation and have adequate separation. At this point, the problem was considered solved and the method was put into routine use on system C.  

Figure 2: Gradient step-test results as in Figure 1, but 1% steps from 0â10% B (actually 0â9.8% B based on the results of Figure 1). Actual composition shown above each step; step size to the right. 

The above problem highlights the challenge of obtaining sufficient precision and accuracy with on-line mixing to obtain consistent retention times for strongly retained peaks. A much simpler solution to the problem of instrument-to-instrument differences would be to use hand-mixed mobile phase for the isocratic portion. That is, hand mix a mobile phase of 95% A and 5% B and put this in the A-reservoir. Separately, hand mix a mobile phase of 5% A and 95% B and put this in the B-reservoir. Now, run the programme for 62 min at 0% reservoir-B and then programme a gradient to 100% reservoir-B in 23 min. This suggestion was tested and the retention time of the main peak came out much later. A further minor adjustment of the initial mobile-phase composition (by adding a little more of the B-solvent for the initial isocratic hold) should bring the retention time back to the target value.  

We've looked at a problem of instrument-to-instrument differences in retention time and speculated that the most likely cause was a difference in mobile-phase proportioning between the systems. We used simple back-of-the-envelope calculations based on normal chromatographic behaviour to estimate the magnitude of mobile-phase composition errors. After we knew what to look for in terms of the scale of the problem, we used the gradient step-test as a means of checking instrument calibration. Although the test results showed compositional differences that would most likely be unimportant for methods with shorter retention times, a replacement of the mobile-phase proportioning valve corrected the problem. If you would like to see more examples of problems related to proportioning errors, consult reference 3. 

is an associate scientist at Gilead Sciences, Inc., in Foster City, California, USA. He specializes in HPLC method development, separation of closely related compounds, impurity elucidation using LC–MS and instrument troubleshooting. 

 is vice president of LC Resources, Walnut Creek, California, USA. He is also a member of 

 editorial advisory board. Direct correspondence about this column should go to "LC Troubleshooting", 

4A Bridgegate Pavilion, Chester Business Park, Wrexham Road, Chester, CH4 9QH, UK, or e-mail the editor, Alasdair Matheson, at 

 (Wiley-Interscience, Hoboken, New Jersey, 2007), section 5.5.3. 

LC Troubleshooting

Retention Time Changes

In this instalment of "Sample Preparation Perspectives," Norwegian authors from the University of Oslo describe the practical aspects of hollow fibre liquid-phase microextraction in the three-phase mode (HF

LPME). The guest authors highlight important practical issues related to the supported liquid membrane, the hollow fibre and the extraction itself. They also discuss practical work with electromembrane extraction (EME), which is related to HF

LPME but uses an electrical potential as the driving force for the extraction.

This month's instalment describes practical aspects of hollow-fibre liquidphase microextraction in the three-phase mode (HF

 LPME is a microscale sample preparation technique (1) where target analytes are extracted from an aqueous sample through a supported liquid membrane (SLM) that is immobilized in the pores of a porous polymeric material and into a volume of acceptor solution (typically, 10–30 µL). In this context, the porous polymeric material is a hollow fibre. Here, we highlight important practical issues related to the SLM, the hollow fibre and the extraction itself because these issues are important for successful HF

LPME. We also discuss practical work with electromembrane extraction (EME), which is related to the HF

LPME device but uses an electrical potential as the driving force for the extraction (2). 

LPME can be used for extraction of basic or acidic analytes from aqueous samples. Figure 1 illustrates a setup for HF

LPME. The sample is contained in a sample vial and the pH is adjusted in the sample before extraction to keep the analytes in their uncharged state. For basic analytes, the sample is made alkaline, and for acidic analytes, the sample is acidified. A small piece of a porous hollow fibre, typically made of polypropylene, is closed in one end and dipped in an organic solvent immiscible with water. In a few seconds, this organic solvent is immobilized in the pores in the wall of the hollow fibre by capillary forces, forming an SLM. 

 Figure 1: Illustration of a typical hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME) setup.			

A 10–30 µL volume of aqueous acceptor solution is then injected into the lumen of the hollow fibre. For basic analytes, the acceptor solution is acidic, whereas it is alkaline for acidic analytes. The hollow fibre is finally placed into the sample and the whole assembly (sample vial and hollow fibre) is agitated for typically 15–45 min. During this time, analyte molecules are extracted in their uncharged state from the sample into the SLM, and further into the acceptor solution. In the acceptor solution, the analyte molecules become ionized, which prevents them from re-entering the SLM. After extraction, the acceptor solution is collected and analysed directly by high performance liquid chromatography (HPLC), capillary electrophoresis (CE), mass spectrometry (MS) or other related analytical techniques.  

Direct compatibility with HPLC, CE and MS

Low solvent consumption (10–30 µL of solvent for each extraction).

Advantages, as well as limitations, of HF

LPME have been discussed substantially in the literature and several reviews discussed a broad range of applications (4–11). 

LPME are typically made of polypropylene (4). Most work published in the literature has been accomplished with a polypropylene hollow fibre from Membrana (Wuppertal, Germany) termed "Q3/2" that has an internal diameter of 600 µm, a wall thickness of 200 µm and a pore diameter of 0.2 µm (4). The hollow fibre is connected to a pre-cut pipette tip or a medical syringe needle. The pipette tip or syringe needle serves as a guide tube to facilitate the injection and withdrawal of the acceptor solution, as illustrated in Figure 2. The fibre can be arranged either in the loop configuration with connections in both ends [Figure 2(b)] or in the rod configuration with a connection in one end and the other end closed [Figure 2(a)]. To close the hollow fibre, mechanical pressure with a pair of pincers can be used — no heat or glue is required. In our laboratory, we use pipette tips for the connections. In this case, we carefully heat the connection between the tip and the fibre with a soldering iron to prevent disruption. 

Figure 2: Different configurations for hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME): (a) rod configuration, (b) loop configuration and (c) hollow fiber attached directly to the needle of a microsyringe.			

Alternatively, the hollow fibre can also be attached directly to the needle of a microsyringe for easy injection and withdrawal of the acceptor solution, as illustrated in Figure 2(c) (12). Alternative fibre dimensions can also be used, but generally the thickness of the wall of the fibre should not exceed 200–300 µm because the extraction speed and the recovery are dependent on the thickness of the SLM. Fibres with internal diameters less than 600 µm have been reported to speed up HF

LPME with small volumes of acceptor solution for high enrichment (3), and hollow fibres with internal diameters larger than 600 µm have been used for easier injection and withdrawal of the acceptor solution (13,14). 

LPME, the SLM is an intermediate extraction medium. Analytes should be extracted rapidly and efficiently into the SLM, but transport out of the SLM and into the acceptor phase should also be efficient to avoid substantial trapping of analytes in the SLM itself. Substantial trapping in the SLM is undesirable because it reduces the extraction recovery. In other words, finding the optimum SLM solvent for the application is an important step. 

Table 1: Boiling point, evaporation, water solubility, and dissolution data for various solvents for hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME).			

LPME, dihexyl ether and 1-octanol have been the most popular SLM solvents (4). As seen in Table 1, these two solvents have a high boiling point (> 195 °C), and when the hollow fibre is dipped in the solvents, little or almost no evaporation of the SLM is observed during 2 min of air exposure (Table 1). This observation is important because the hollow fibre with the immobilized SLM is normally exposed for a short time (< 2 min) to open air before it is placed in the sample. After the hollow fibre with the SLM is inserted in the sample in a capped vial, evaporation is no longer an issue because the SLM is protected by the aqueous sample and the system is closed. Volatile solvents such as toluene and 1-chloropentane may be difficult to use in HF

LPME because they evaporate quickly and give an unstable SLM (see Table 1). In general, it is recommended not to use solvents with a boiling point below190–200 °C for HF

In addition to the volatility of the solvent, the water solubility is also important. For dihexyl ether, the water solubility is low (< 110 µg/mL), therefore, SLMs made from this solvent are very stable during extraction. With 1-octanol, the water solubility is higher (1200 µg/mL) and this solvent may leak into the sample (and the acceptor solution) in significant amounts. As seen in Table 1, about 11% of a 1-octanol SLM may theoretically leak into 1 mL of sample based on the water solubility, resulting in a significant reduction of the SLM. This level of SLM leakage has been verified experimentally in our laboratory by analysing the sample solution after HF

LPME with gas chromatography–mass spectrometry (GC–MS) to determine traces of 1-octanol. This loss may be even greater if the sample volume is increased or if surfactants and other emulsifying agents are present in the sample. In general, it is preferred not to use solvents exceeding 200–400 µg/mL in terms of water solubility. In addition to dihexyl ether, solvents such as 1-decanol, dodecyl acetate, 2-nitrophenyl octyl ether and 1-nonanol fulfill the criteria discussed above and have been used in HF

LPME studies (Table 1) (4). Some new solvents that have also been tested recently in our laboratory for HF

LPME are included in Table 1 and may be interesting SLM candidates for the future.  

How Much Does the SLM Affect the Extraction?

Table 2 (basic drugs as model analytes) and Table 3 (acidic drugs as model analytes) illustrate recent examples from our laboratory in which the extraction recovery in HF

This month's "Sample Prepartion Perspectives" describes practical aspects of hollow-fibre liquid-phase microextraction in the three phase mode (HF3LPME) and also electromembrane extraction (EME), which uses an electrical potential as the driving force for the extraction.

LCGC Asia Pacific-06-01-2012

Introduction to SPME for In Vivo Analysis

New Gas Chromatography Products 2012

Hyaluronic Acid (Polysaccharides)

Absolute Molar Masses for Phenol Formaldehyde Resins with GPC/SEC-ESI

Retention Time Changes

Hollow-Fibre Liquid-Phase Microextraction in the Three-Phase Mode — Practical Considerations

Vol 15 No 2 LCGC Asia Pacific June 2012 Regular Issue PDF