Strategies for coupling solid-phase microextraction with mass spectrometry

Trends in Analytical Chemistry 55 (2014) 55–67

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Review

Strategies for coupling solid-phase microextraction with mass spectrometry Jiewei Deng a,1, Yunyun Yang b,1, Xiaowei Wang c, Tiangang Luan a,⇑ a

MOE Key Laboratory of Aquatic Product Safety, School of Life Sciences, Sun Yat-Sen University, Guangzhou 510275, China Guangdong Provincial Public Laboratory of Analysis and Testing Technology, China National Analytical Center Guangzhou, Guangzhou 510070, China c School of Marine Sciences, Sun Yat-Sen University, Guangzhou 510275, China b

a r t i c l e

i n f o

Keywords: Ambient mass spectrometry Atmospheric pressure ionization mass spectrometry Coupling Electron-impact mass spectrometry Inductively-coupled plasma mass spectrometry Interface Laser-desorption/ionization mass spectrometry Mass spectrometry Sample pretreatment Solid-phase microextraction

a b s t r a c t Solid-phase microextraction (SPME) has experienced significant development since its introduction as a sample-pretreatment technique in the early 1990s. SPME is suitable for interfacing with chromatography and mass spectrometry (MS), but progress in coupling with chromatography has exceeded that with MS. In the past two decades, efforts have been made to couple SPME and MS with different applications in various research fields. Based on these previous studies, this review article summarizes historical developments, principles and operation, practical applications, and recent trends in SPME coupled with five types of MS: (1) electron-impact MS, (2) inductively-coupled plasma MS, (3) laser-desorption/ionization MS, (4) atmospheric-pressure ionization MS and (5) ambient MS (AMS). We particularly emphasize efforts on SPME coupled with AMS. Ó 2014 Elsevier Ltd. All rights reserved.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SPME coupled with EI-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Coupling SPME with EI-MS using the GC-injection port as interface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Fiber introduction MS (FIMS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SPME coupled with ICP-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Coupling of SPME to ICP-MS with thermal desorption interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Coupling SPME to ICP-MS with a solvent-desorption interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SPME coupled with LDI-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. SPME coupled with vacuum LDI-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. SPME coupled with AP-LDI-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SPME coupled with API-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Coupling SPME with API-MS using an SPME desorption chamber as interface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Coupling SPME to API-MS using in-tube SPME as interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SPME coupled with AMS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. SPME coupled with solid-substrate ESI-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. SPME coupled with desorption ESI MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. SPME coupled with direct analysis in real time MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4. SPME coupled with desorption corona beam ionization MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.5. SPME coupled with low-temperature plasma (LTP) probe MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Future perspective and new opportunities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

⇑ Corresponding author. Tel.: +86 20 84112958; Fax: +86 20 84112199. 1

E-mail address: [email protected] (T. Luan). Equal contributions by these authors.

0165-9936/$ - see front matter Ó 2014 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.trac.2013.12.004

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1. Introduction Solid-phase microextraction (SPME), a solvent-free samplepretreatment technique that integrated sampling, isolation and enrichment of analytes into one step, was developed and implemented in analytical practice in the early 1990s by Professor Janusz Pawliszyn of University of Waterloo (Ontario, Canada) [1–3]. SPME provides significant advantages of simplicity, speed, sensitivity and ease of operation, when compared with conventional extraction techniques, such as liquid–liquid extraction and Soxhlet extraction. In an SPME-based analytical method, a fused-silica fiber coated with polymer is introduced directly into the sample or the headspace (HS) above the sample for enrichment and concentration of analytes, and then transferred to an analytical instrument for thermal/solvent/energy desorption and analysis of analytes [1–6]. The most commonly used analytical approach for desorption, separation, and detection of the analytes enriched on SPME fibers is chromatography. The combination of SPME with different chromatographic techniques has been investigated thoroughly by many research groups since the invention of the SPME technique [1,2,7–14]. SPME coupled with gas chromatography (GC) has been achieved by employing a GC-injection port as interface, in which analytes are released by thermal desorption and carried into GC column for separation and analysis [1,2]. Interfacing of SPME to liquid chromatography (LC) has been developed by applying an SPME desorption chamber as interface for solvent desorption of analytes and then introduction into LC column [10], and auto-

mated SPME-LC coupling has been successfully using in-tube SPME as interface [11]. The coupling of SPME with capillary electrophoresis (CE) was successfully realized by applying an adapter for connecting the SPME fiber and the capillary column [12], and in-tube SPME has also been introduced as another interface for on-line coupling of SPME and CE [13]. SPME is also capable of interfacing with mass spectrometry (MS) directly for detection of analytes without employing chromatographic separation. It is well known that MS allows high sensitivity and excellent specificity for rapid determination of compounds from complex matrices, and is a powerful tool for confirmation of targeted components, elucidation of non-targeted analytes, and identification of unknown compounds. However, direct introduction of complex samples into the mass spectrometer usually results in low sensitivity, high matrix effect, and rapid contamination of the instrument. By coupling SPME to MS, extraction, clean up, enrichment and detection of analytes are integrated into one single step. Thus, higher sensitivity and lower matrix-suppression effects can be achieved, and are extremely beneficial for detection of trace compounds from complex matrices. Moreover, the analytical time is greatly reduced because it eliminates the need for chromatographic separation. The feasibility of SPME coupled with MS has been investigated since the late 1990s, but its progress is not nearly as good as that of SPME coupled with chromatographic techniques. In most cases, the interface of SPME to MS is complicated, requiring a series of modifications of the mass spectrometer. In the past two decades, sub-

Fig. 1. Logical and methodological categorization of the schemes involved in solid-phase microextraction coupled with mass spectrometry.

J. Deng et al. / Trends in Analytical Chemistry 55 (2014) 55–67

stantial progress has been made for coupling SPME with MS, with different applications in various research fields. Using different strategies, SPME has been successfully coupled with five types of MS: (1) electron impact (EI)-MS; (2) inductively-coupled plasma (ICP)-MS; (3) laser-desorption/ionization (LDI)-MS; (4) atmospheric-pressure ionization (API)-MS; and, (5) ambient MS (AMS) (Fig. 1). To date, numerous reviews on interfacing SPME with chromatography have been published [15–17]. However, the principles,

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the developments and the applications for coupling SPME with MS have not yet been summarized systematically. In this review article, we discuss publications to date on SPME coupled with MS with different designs and combinations. For each coupled technique, we describe, the history of development, the underlying principles of operation, the possible desorption and ionization processes, and the representative applications. We highlight SPME coupled with AMS, including a brief discussion of future perspectives and new opportunities of this coupled technique.

Fig. 2. (a) The main steps involved in FIMS analysis. (i) SPME fiber ready for insertion into the mass spectrometer; (ii) SPME fiber exposed between the two EI filaments; (iii) analyte desorption from the SPME fiber and ionization by EI (70 eV electron beam) and transmission to the MS analyzer. (Reprinted from [27] with permission of Elsevier). (b) The interface directly coupling SPME to a miniature ion-trap mass spectrometer using the FIMS technique (Reproduced from [28] with permission from The Royal Society of Chemistry).

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2. SPME coupled with EI-MS 2.1. Coupling SPME with EI-MS using the GC-injection port as interface SPME coupled with EI-MS using the GC-injection port as interface is simple and easy to realize, as the only modification of a commercial instrument is to replace the analytical column of a GC-MS system with a transfer line. This coupling allows analytes enriched on the SPME fiber for thermal desorption in the GC-injection port, and then transferred to the EI-MS for detection. The first application of this coupled technique was reported by Marsili in 1999 [18]. The author utilized a 75-lm Carboxen (CAR)/ polydimethylsiloxane (PDMS) fiber to extract the volatiles from milk samples. After extraction, the fiber was inserted into a GCinjection port for thermal desorption, and the desorbed analytes were then transferred through a 1.0 m  0.25 mm uncoated, fused-silica column into the ion source for MS analysis without employing any chromatographic separation. Principal-component analysis was then applied to the acquired mass spectra, providing rapid differentiation of control reduced-fat milk (2% butterfat content) samples from reduced-fat milk samples affected by light, heat, copper, and microbial contamination. Similar research was done by Pérès et al. in 2001 [19] for rapid analysis and characterization of the volatile fraction of cheeses. A 75-lm CAR/PDMS fiber was applied for extraction, and then thermally desorbed via a GC-injection port. The gas-state analytes were delivered to EI-MS via a 1 m  0.1 mm deactivated silica capillary column heated to 210°C for analysis. Because interfacing SPME to EI-MS via a GC-injection port requires little modification of existing equipment, it is easy for many laboratories to achieve and to manipulate this coupling. Thus, it is still being used to date. For example, Mildner-Szkudlarz and Jelen´ developed an SPME-EI-MS method for adulteration of olive oil with hazelnut oil in 2008, using a 15 m  0.2 lm uncoated fused-silica column as transfer line [20]. An SPME-EI-MS method for identification of the botanical origin of raw spirits produced from rye, potato, and corn based on volatile compounds was conducted by Jelen´ et al. in 2010 [21], using a 5 m  0.2 mm fused-silica column without phase coating for transmission of the analytes. Non-separative HS-SPME-EI-MS approaches via GC-injection port and a 6.70-m deactivated fused-silica column were demonstrated by Bicchi et al. in 2011 [22], and Liberto et al. in 2013 [23], respectively, in view of their applications to on-line monitoring of the coffee roasting process. In spite of that, the advantages of such coupling are not obvious, with the exception of reduced analytical times in comparison with conventional SPME coupled with GC-MS. By eliminating of the chromatographic separation step, qualitative and quantitative analysis of a complex mixture based on only 70 eV EI mass spectra is difficult. These limitations restrict widespread use of this technique. 2.2. Fiber introduction MS (FIMS) FIMS is a successful technique for direct introduction of SPME fiber into the vacuum ion-source region of EI-MS and simultaneous thermal desorption and electron ionization of analytes for MS analysis. Fig. 2a shows the major steps involved in FIMS analysis. First, an SPME fiber, which contains the analytes previously extracted from the sample, is introduced into the vacuum region of an EI mass spectrometer via a silicone septum interface. The fiber is then exposed to the two filaments, heated and bombarded with a 70 eV electron beam. The rapid heating causes rapid, efficient desorption of the analytes, which are in turn ionized in the gas phase via the electron beam. The resulting ions are rapidly transmitted to the mass analyzer for detection.

FIMS was first demonstrated by Meurer et al. in 2002 [24]. Using this technique, they successfully detected and quantified low-ppb to ppt levels of volatile organic compounds (VOCs) and semi-VOCs (SVOCs), such as carbon tetrachloride, benzene, toluene, xylenes, c-terpinene, diisoamyl ether, chlorobenzene, and polycyclic aromatic hydrocarbon, from aqueous solutions, by applying a 100-lm PDMS fiber for HS or direct-immersion extraction. FIMS is also applicable for quantitative determination of trace weak polar or moderately polar analytes. Silva et al. [25] developed a FIMS method using selective ion monitoring to detect and to quantify dimethylphthalate, diethylphthalate and dipropylphthalate in mineral water. The results showed that FIMS with a 65lm PDMS/divinylbenzene (DVB) fiber allowed simple extraction and rapid MS detection of the investigated phthalates with good linearity and precision. The limits of detection (LODs) were 3.6– 5.1 lg/L, which were smaller than the US Environmental Protection Agency (EPA) regulation limit of 6 lg/L for di-2-ethylhexyl phthalate (a similar contaminant) in drinking water. Another FIMS method was also developed by Silva et al. [26] for determination of two organochlorine (i.e. chlorothalonil and aendosulfan) and three organophosphorus (i.e. fenthion, malathion and methyl parathion) pesticides in herbal infusions of Passiflora L. The authors developed an SPME fiber coated with a composite of PDMS and poly(vinyl alcohol). Simple HS extraction (25 min) and fast FIMS detection (less than 40 s) provided good linearity (correlation coefficients of 0.991–0.999) for each compound with concentrations of 10–140 ng/mL. In addition, good accuracy and precision, and low LODs (0.3–3.9 ng/mL) were also obtained. With regard to polar compounds, FIMS also shows considerable capability for detection even without derivatization of analytes. Eberlin et al. [27] described a FIMS method for direct analysis of five chlorophenols in water without pre-separation and derivatization, and the result showed that FIMS was faster and simpler than current EPA methods with comparable sensitivity, and was particularly suitable for on-site analysis. Another valuable use of FIMS was direct coupling of SPME to a portable miniature ion-trap (IT) mass spectrometer (Version 7), which was reported by Riter et al. in 2003 [28]. The vacuum interface was constructed of a 25 mm Quick Flange (QF25) blank stub with a 1/16-inch o.d. stainless-steel tube fitted through 1/8-inch Swagelok fittings held in place by 1/8–1/16-inch polytetrafluoroethylene reducing ferrules. At the high-pressure end of the Swagelok fittings, a GC septum allowed direct introduction of an SPME needle and fiber assembly into the vacuum system of the mass spectrometer. Thermal desorption of the analytes in the coated fiber was achieved with a built-in Nichrome heater, followed by electron ionization and cylindrical IT analysis (Fig. 2b). They applied this system to analyze VOCs in air with HS extraction using a 100-lm PDMS fiber, with LODs in the low-ppb range. In addition, this system has been successfully applied to quantitation of toluene in benzene, toluene, xylene mixtures in water and gasoline. In comparison with the coupled technique using GC-injection port as interface for coupling SPME to EI–MS, FIMS shows higher sensitivity and better reproducibility because the desorption and ionization of analytes take place in the ion source region of mass spectrometer simultaneously, allowing rapid and efficient transmission of analytes to the mass analyzer for detection. However, accurate positioning of the SPME fiber placed and exposed between the two filaments is extremely critical, and requires the SPME holder and the MS inlet to be tightly controlled. Another disadvantage of FIMS, which is similar to that using a GC-injection port as interface, is that qualitative and quantitative analysis become unsatisfactory, when based on 70 eV EI mass spectra only, as the mixture becomes increasingly complex. This limitation can be minimized by increasing the selectivity by using softer ioniza-

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Fig. 3. (a) The thermal-desorption interface for SPME analyte introduction into ICP-MS. (Reproduced from [29] with permission from The Royal Society of Chemistry). (b) On-line capillary microextraction (CME)-ICP-MS with PNIPA-coated fiber-in-tube capillary as extraction device. (Reprinted from [34] with permission of Elsevier).

tion techniques, such as chemical ionization, with high-resolution MS and MS/MS.

3. SPME coupled with ICP-MS 3.1. Coupling of SPME to ICP-MS with thermal desorption interface Direct coupling of SPME with ICP-MS was first described by Mester et al. in 2000 [29]. They utilized a thermal-desorptiongas-introduction interface to introduce analytes enriched on SPME fiber into ICP-MS. The thermal-desorption interface consisted of a heated, glass-lined, splitless type of GC injector, placing directly at the base of the torch to minimize the length of transfer line (Fig. 3a). This arrangement provided fast desorption and high sample-introduction efficiency, allowing sample introduction of volatile metal species into ICP-MS directly. Using this coupled device, they successfully detected methylmercury in biological-tissue samples via direct immersion or HS extraction using a 65-lm PDMS/DVB fiber. Subsequently, Mester et al. [30] used the same device for the non-selective determination of arsenic, selenium, antimony and tin species amenable to hydride generation. The LODs for As, Se, Sn and Sb hydrides using a 75-lm PDMS/CAR fiber were 70 pg/ mL, 5300 pg/mL, 8 pg/mL and 310 pg/mL, respectively. Tributyltin in aqueous samples was also successfully detected by Mester and co-workers, with an LOD of 5.8 pg/mL by using a 65-lm PDMS/ DVB fiber for HS extraction [31]. Guo et al. [32] developed an SPME coupled with an ICP-time-offlight (TOF)-MS method for quantitative determination of trace germanium. They compared chloride-generation and hydride-generation HS-SPME techniques. LODs of 20 pg/mL and 92 pg/mL and the precisions of 18% (n = 11) and 9.7% (n = 11) were achieved for chloride generation and hydride generation, respectively. The gen-

erated germanium chloride and hydride species were identified as GeCl4 and GeH4. Chloride generation doubled sensitivity compared to hydride generation, while the LODs for continuous hydride generation were 20-fold better than reported atomic fluorescence data. A method based on SPME-ICP-MS for speciation of volatile organo-selenium compounds was presented by Dietz et al. in 2003 [33]. They used an in-house desorption unit for thermal desorption of analytes enriched on an SPME fiber, and then detection by different detectors, such as ICP-MS and atomic absorption spectroscopy (AAS). Dimethylselenium and dimethyldiselenium could be separated and quantified in less than 2 min, achieving LODs of 0.7 mg/L and 0.9 mg/L, respectively, when ICP-MS detection was used. In addition, they applied the method to quantify organo-selenium in garlic samples.

3.2. Coupling SPME to ICP-MS with a solvent-desorption interface Coupling of SPME to ICP-MS using in-tube SPME as a solventdesorption interface was reported by Zheng and Hu in 2011 [34]. The poly(N-isopropylacrylamide) (PNIPA) gel was prepared and applied as a polymer coating for fiber-in-tube capillary microextraction of trace Co, Ni and Cd. The fiber-in-tube capillary was connected to the sample-introduction loop of the six-port injection valve, and the sample solution was allowed to pass through the capillary. Afterwards, the analytes retained on the capillary were eluted on-line with 50 lL of 0.1 mol/L HNO3 for desorption and further ICP-MS detection (Fig. 3b). The PNIPA coating showed high extraction efficiency, high adsorption capacity, good chemical stability and good regeneration capability, with LODs of 0.45 ng/L, 4.6 ng/L and 6.9 ng/L for Co, Ni and Cd, respectively. The method was applied to the determination of Co, Ni and Cd in human serum and urine with satisfactory results.

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Fig. 4. (a) The SPME-SGALDI method. (Reprinted from [38] with permission of Wiley-VCH). (b) The SPME-MALDI-QqTOF-MS system. (1. Laser source; 2. Focusing lens; 3. Fiber holder; 4. SPME/MALDI fiber; 5. QqMS; and- 6. TOF MS) (Reproduced from [40] with permission from The Royal Society of Chemistry). (c) The SPME/AP MALDI configuration (the target plate held an array of 16 SPME extraction fibers) for high-throughput determination of biological samples. (Reprinted from [42] with permission of American Chemical Society).

4. SPME coupled with LDI-MS 4.1. SPME coupled with vacuum LDI-MS Since its introduction in the mid-1980s, LDI has become a powerful technique for the analysis of biomolecules, such as proteins, peptides and oligonucleotides, by MS. Developed in 1988, matrix-assisted laser desorption/ionization (MALDI)-MS, which uses an ultraviolet-absorbing matrix as medium to transmit laser energy for desorption and ionization of analytes, is the most representative and widely-used LDI-MS technique [35,36]. LDI-MS can work with vacuum or atmospheric pressure (AP) conditions. The combined use of SPME with vacuum LDI-MS was first reported by Chen and Sun in 2002 [37]. They used a polycrystalline graphite (pencil lead) as the SPME fiber to combine with surface-assisted LDI (SALDI)-MS for detection of non-ionic alkylphenol-ethoxylate surfactants in water. The analytical procedure involved immersing the fiber in the sample solution for extraction, removing the absorbed analytes by scraping carbon powder from the graphite surface, mixing the powder with a SALDI liquid (15% sucrose/glycerol in methanol), and subjecting the sample to LDI-MS analysis. The carbon power served as the medium to transfer the UV-laser energy into the liquid to assist and to enhance LDI. As a result, an LOD of low ng/L was achieved for the analysis of nonyl phenyl polyethylene glycol ether. Special attention needed to be paid to the scratching procedure to ensure that the carbon powder obtained contained the trace analytes from the surface of the graphite fiber. Teng and Chen [38] demonstrated a strategy using a sol–gel derivative as both SPME coating and matrix material to assist LDI

without the addition of matrix, which was termed sol–gel-assisted LDI (SGALDI)-MS. In brief, an optical fiber was first coated with a thin layer of sol–gel 2,5-dihydroxybenzoic acid (DHB) derivative. Then, the fiber was immersed in the aqueous solutions to extract trace benzo[a]pyrene. Following extraction, the whole fiber was fixed on a Parafilm membrane (previously attached to a MALDI sample plate) using a transparent tape on both ends of the fiber. Subsequently, the fiber was subjected to LDI and detected by MS (Fig. 4a). A similar analytical method was also described by Perera et al. [39], with DHB-bonded silica particles coated on the fiber being used as both the SPME extraction phase and the MALDI substrate for extraction and LDI analysis of peptides and proteins. However, these procedures were laborious, since the SPME fiber was manually fixed to the plate, and only one side of the analytes on the SPME fiber could be introduced into the mass spectrometer. In general, SPME coupled with vacuum LDI-MS is easy to realize without any requirement for additional instrumentation design. However, such approaches are operated off-line, and the experimental procedures are somewhat laborious, since the SPME fibers must be manually scratched or fixed on the plate. All these drawbacks might be limitations to achieving rapid, high-throughput analysis. 4.2. SPME coupled with AP-LDI-MS SPME coupled with AP-MALDI-MS was initially reported by Pawliszyn’s group in 2002 [40]. They used a fused-silica optical fiber to produce an SPME/MALDI fiber to extract and to ionize analytes for direct MS analysis. One end of the optical fiber was silanized with 3-aminopropyltriethoxysilane for the extraction of

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analytes, and the other end was fixed into a connector ferrule with epoxy and then carefully polished to allow the maximum laser transmission through the fiber. Laser energy was transferred through the end of the optical fiber to desorb and to ionize analytes for subsequent analysis by ion-mobility spectrometry (IMS) or quadrupole time-of-flight (QTOF)-MS. A mixture of encephalin and substance P (a neuropeptide) solution in a-cyano-4-hydroxy cinnaminic acid as the matrix were investigated by using QTOFMS for detection. Samples were loaded by depositing a small amount of solution onto the end of the fiber and letting it dry. A prototype QStar QTOF-MS system was used for the experiment by simply removing the IonSpray source to expose the sampling orifice in ambient pressure conditions. The end of the SPME/MALDI fiber was fitted into a stainless-steel tube, which was supported on an insulator. The tube was positioned so that the optical fiber was aligned with the sampling orifice. An optimum laser power (about 100 lJ per pulse) was applied to the fiber for desorption and ionization of analytes for MS analysis (Fig. 4b). On-line coupling of SPME and LDI-MS was achieved by applying this approach, and good-quality mass spectra were obtained for a 500 fmol/lL solution. However, the sensitivity of the method developed was relatively poor due to MALDI being worked at AP conditions, and no signal was detected from a solution of 50 fmol/lL. It has been a challenge to use MALDI for the analysis of lowmass analytes, because of the presence of matrix-related ions in the low-mass range. Efforts have been made to overcome these limitations, and several matrix-free approaches have been developed. The matrix-free techniques are usually described as surface-enhanced LDI (SELDI). Direct coupling of SPME to SELDI was first developed by Wang et al. in 2004 [41] for determination of leucine encephalin. A polypyrrole SPME coating was employed as both the extraction phase and the SELDI surface to enhance LDI of analytes. Since the matrix was not required, the mass spectra obtained were free of matrix interferences, so that direct analysis and quantification of small peptides without peak interferences from the matrix showed interesting advantages. A high-performance SPME-AP-MALDI system for high-throughput sampling and determination of peptides was also reported by Pawliszyn’s group in 2005 [42]. This SPME-AP-MALDI system showed potential for high-throughput extraction from biological samples, with a multiplexed SPME plate capable of simultaneous extraction from 16 different wells on a multi-well plate without the need for extensive sample preparation (Fig. 4c). The AP-MALDI source stage was repositioned by removal of shims so that the tips of the SPME fibers could be placed approximately 2 mm from the inlet of the laminar flow chamber. The laser beam was focused directly onto the front side of the fiber surface at a 28° angle for desorption and ionization. As a result, sub-femtomole sensitivity for peptide standards and protein digests and run-to-run reproducibility of 13–31% were obtained. SPME coupled with LDI-MS has emerged as a promising technique, because it integrates significant advantages of both techniques, such as simplicity, speed, high sensitivity, high ion transmission and high spectral acquisition rates. More importantly, it extends the applications of SPME to polar biomacromolecules, with great potential in biochemical analysis, pharmaceutical research, and clinical diagnostics. 5. SPME coupled with API-MS 5.1. Coupling SPME with API-MS using an SPME desorption chamber as interface

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technique was first reported by Möder et al. in 1997 [43] for analysis of acylcarnitines in aqueous solutions, urine and blood plasma samples. They used a 70-lL SPME desorption chamber, described in detail by Chen and Pawliszyn in 1995 [10], as the interface for coupling SPME with electrospray ionization (ESI)-MS directly. The loop of the six-port injection valve was replaced by a threeway tee, the so-called desorption chamber. The valve was first switched to ‘‘load’’ position and the chamber was filled with desorption solution. Then, an SPME fiber loaded with analytes was carefully inserted through a ferrule and a Teflon tube into the tee for solvent desorption. After switching the valve to the ‘‘inject’’ position, the desorption solution was pumped into the ESI nebulizer for MS analysis. By applying the same coupling design, Ceglarek et al. [44] developed SPME coupled with ESI-MS for the qualitative and quantitative determination of linear alkylbenzene sulfonates (LASs) in wastewater samples. A 50-lm Carbowax/templated resin-coated fiber was directly immersed in influent and effluent samples of a sewage-treatment plant for extraction. The extracted LASs were desorbed for 15 min with a solvent of isopropanol/methanol (1:1) in an SPME desorption chamber, and then transferred for analysis by ESI-MS. Linear ranges of the external calibration were 0.5–100 ng/mL, with LODs of 0.5 ng/mL for each LAS homologue. McCooeye et al. [45] had an analogous experimental set-up for fast screening amphetamine, methamphetamine and their methylenedioxy derivatives in urine. An SPME desorption chamber was assembled in-house from a tee union. An SPME fiber was inserted through a graphite-filled vespel ferrule into the desorption area and secured in the upper end of the tee union by tightening the nut on the tee union. The sample was dynamically desorbed from the fiber by a continuous flow of 1.5 lL/min of buffer solution delivered by an HPLC pump, or statically desorbed by stopping the HPLC pump, and placing the fiber in the desorption chamber for 3 min. Desorption solution was then carried to the ESI needle via a silanized fused-silica capillary, with detection by high-field asymmetric waveform IMS (FAIMS)-MS. The LODs of the examined drugs in human urine were in the range 7.5–200 ng/mL. Another application was reported by van Hout et al. [46], with a method based on SPME coupled with AP chemical-ionization (APCI)-MS and an SPME desorption chamber as interface determining lidocaine in a sample of urine. The SPME procedure was performed by using a 100-lm PDMS fiber for direct immersion extraction under non-equilibrium conditions, permitting an analysis time within 10 min per sample, with 5 min of sorption and 4 min of static desorption. The LOD was 0.4 ng/mL for lidocaine, and the intra-day and inter-day reproducibility were within 14% over a concentration range of 2–45 ng/mL. A further investigation was performed by van Hout et al. [47], who applied non-equilibrium SPME at elevated temperatures and direct coupling to APCI-MS, with a modified desorption chamber allowing thermostatting as interface, for analysis of lidocaine in urine. Elevated temperatures during sorption (65°C) and desorption (55°C) had considerable influence on the extraction speed. Only 1 min sorption and 1 min desorption were performed, followed by MS detection, resulting in a total analysis time of 3 min. The determination of lidocaine in urine had acceptable reproducibilities, with relative standard deviations (RSDs) less than 10%, and a limit of quantification (LOQ) of about 1 ng/mL was obtained. The short sorption and desorption times ensured rapid analysis, showing interesting potential for high-throughput analysis of biological samples. 5.2. Coupling SPME to API-MS using in-tube SPME as interface

Interfacing SPME to API-MS using an SPME desorption chamber as interface for solvent desorption of analytes was the subject of the earliest research into a technique coupling SPME with MS. This

In-tube SPME has also been successfully coupled with API-MS, as first realized by Mester et al. in 1999 [48]. Trimethyl lead and

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Table 1 Typical applications of solid-phase microextraction (SPME) coupled with ambient mass spectrometry (AMS) SPME coating

Analyte

Extraction mode

Graphite RAM PDMS/DVB PDMS/DVB

Surfactants (Triton X-100) Peptides, proteins CWAs CWAs

Direct immersion Direct immersion Headspace Headspace

CAR/DVB CAR/DVB C18 C18/SCX DVB/CAR/ PDMS

CWAs Anabolic steroids Illicit drugs and pain medications Carbamazepine, triclosan Volatile fingerprint

PAN over C18 PAN C18 PAN PDMS Graphite

Diazepam

Headspace Direct immersion Direct immersion Direct immersion Headspace/direct immersion Direct immersion

Cocaine, methadone Pesticides Volatile organic compounds

Direct immersion Direct immersion Direct immersion

Sample matrix

AMS technique

Ref.

Aqueous solution Body fluids Spiked carpet Spiked office carpet, fabrics, photocopy paper, swabs Spiked swab, office furniture fabric, cardboard Urine Urine Wastewater Beer

DEP Nanospray DESI DESI

[57] [58] [63] [64]

DESI DESI DESI DESI DART

[65] [66] [67] [71] [73]

Whole blood

DART

[74]

Urine Water Standard solutions

DART DCBI LTP probe

[75] [77] [79]

CWA, Chemical-warfare agent; PDMS, Polydimethylsiloxane; RAM, Restricted access material; DVB, Divinylbenzene; CAR, Carboxen; SCX, strong cation exchanger; PAN, Polyacrylonitrile; DEP, Direct electrospray probe; DESI, Desorption electrospray ionization ; DART, Direct analysis in real time; DCBI, Desorption corona beam ionization; LTP, Low-temperature plasma.

Fig. 5. SPME coupled with ambient mass spectrometry (AMS) with ionization techniques of (a) solid-substrate ESI-MS; (b) DESI-MS); (c) DART-MS; and, (d) DCBI-MS.

triethyl lead in water samples were extracted using an open-tube capillary with porous DVB polymer coated inside. The capillary was connected to an HPLC system in place of the sample-introduction loop. Using an auto-injector with the six-port valve in the ‘load’ position, the capillary was washed several times with the sample solution by applying ‘‘draw from the sample-eject into the sample’’ extraction cycles. The cycles were repeated until equilibrium was reached. After the extraction step, the valve was switched to the ‘inject’ position and the mobile phase was passed

through the extraction capillary for desorption of analytes and sent into the ESI nebulizer for MS analysis.

6. SPME coupled with AMS AMS [49] {also named ambient desorption ionization MS [50] or ambient ionization MS [51]} comprises useful techniques for the analysis of samples under ordinary ambient and open-air condi-

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tions [51]. It allows direct, rapid, real-time and high-throughput analyses with minimal, or no, sample pretreatment and without chromatographic separation [49–54]. Most of the ambient ionization techniques are based on the mechanisms of ESI or APCI [50]. According to the differences on ionization and analytical processes, ambient ionization techniques can be classified into three main analytical strategies: (1) direct ionization; (2) direct desorption/ionization; and, (3) two-step ionization [51]. In the past two decades, efforts have been made to develop SPME coupled with AMS techniques for highly-sensitive, highthroughput and automated analysis, and the typical applications are summarized in Table 1. 6.1. SPME coupled with solid-substrate ESI-MS Solid-substrate ESI [55], which uses non-capillary solid materials for loading sample and ionization of analytes directly in a high electric field, is one of the simplest ambient ionization techniques based on the ESI mechanism with the analytical strategy of direct ionization. Various materials, including wick element, copper wire, metal needle [i.e. probe ESI (PESI)], optical fiber wired with a metal coil [i.e. direct electrospray probe (DEP)], surface-modified glass rod, nanostructured tungsten oxide, paper (i.e. paper spray), toothpick (i.e. wooden-tip ESI) and biological tissues, have been employed as solid substrates for sampling and direct ionization of analytes in complex matrices [55,56]. It is interesting that the SPME fiber can be used as a solid substrate, by applying some spray solvent and a high voltage, for direct ionization of the analytes enriched on the fiber. An electric field applied to the analyte solution on a pointed end leads to formation of Taylor cone and production of droplets, which are then sprayed to generate charged ions for MS analysis (Fig. 5a). The first SPME-solid-substrate ESI study was reported by Shiea’s group in 1999 [57]. They used an SPME fiber as the electrospray probe to construct an SPME-DEP assembly, by circling the copper wire on a graphite fiber for 12 turns. The SPME-DEP assembly was exposed to water for extraction of trace amounts of Triton X-100 (a surfactant). Then, the loading SPME-DEP assembly was mounted on an acrylic plate held on a XYZ-translation stage, 2 lL of desorption solution (50% methanol/water with 1% acetic

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acid) was deposited onto the copper coil, and a high voltage was applied to produce an electrospray on the tip of the coil, and the mass spectra of the analytes were obtained. Another SPME-solid-substrate ESI study was the coupling of SPME to nanospray, which was reported by Pawliszyn’s group in 2005 [58]. In their study, two biocompatible restricted access materials containing octadecyl silica (C18) extraction centers or strong cation-exchange properties were used as SPME fiber coatings to extract drugs and peptides from biofluids and tryptic digests. The loaded fiber was inserted into a commercial nanospray tip for solvent desorption. The nanospray tip then served as a solid substrate for direct MS analysis. The results demonstrated that SPME coupled with nanospray was a desirable approach for analysis of peptides from an aqueous solution, with an LOD down to 50 fmol/mL. SPME coupled with solid-substrate ESI is rather simple and straightforward with high sensitivity, but only a few publications are found in the literature, but this should improve in the future.

6.2. SPME coupled with desorption ESI MS Desorption ESI (DESI), a very successful ambient ionization technique based on the ESI mechanism with the analytical strategy of direct desorption/ionization, was developed by Cooks and coworkers from Purdue University in 2004 [59]. DESI is extremely suitable for detection of analytes on surfaces [59] and possesses the capability of imaging [60–62]. Thus, this technique is a desirable approach for direct desorption and ionization of analytes concentrated on an SPME fiber. The SPME-DESI-MS technique involves a pneumatically-assisted electrospray to impinge charged droplets and ions of solvent onto the SPME coating surface for desorption and generation of gaseous ions of analytes, and then introduced into a mass analyzer for detection (Fig. 5b). Accurate positioning of the SPME probe in the spray plume of DESI is extremely critical. Otherwise, it may cause a loss in sensitivity and contribute to the error in the measurements. The first application of SPME coupling to DESI-MS was reported by D’Agostino et al. in 2006 [63]. A 65-lm PDMS/DVB fiber was exposed to HS above the office-carpet samples contaminated with triethyl phosphate and the chemical-warfare agents (CWAs) of sarin and soman for extraction, and then the fiber was directly analyzed by DESI-MS and DESI-MS/MS. High-resolution DESI-MS/MS spectra were obtained for all three spiked compounds at the lg/g

Fig. 6. The experimental set-up of TFME-DESI-MS for high-throughput analysis (1. Home-built DESI source mounted on an Orbitrap Discovery hybrid instrument; 2. Electrosonic sprayer; 3. Secured onto a rotating stage; 4. which is mounted on a 3D moving stage; 5–7. Screws adjusting in the x-, y-, and z- directions; 8. Sample table, movable in the x-, y-, and z- directions; 9. Gas supply; 10. High-voltage supply; 11. Solvent supply; 12. Atmospheric inlet of mass spectrometer; 13. Inlet capillary; and, 14. TFME blades taped onto glass slides). (Reproduced from [71] with permission from The Royal Society of Chemistry).

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level. Continued investigations of the CWAs using SPME-DESI-MS were described by D’Agostino et al. [64,65]. Tabun and a number of related organophosphorus compounds spiked in office media, including flooring, wall surfaces, office fabrics, and paper products, were all successfully detected by SPMEDESI-MS methods [64,65]. Sulfur mustard, a compound that has not been successfully analyzed by electrospray MS in the past, was also successfully analyzed for the first time using SPMEDESI-MS [64]. The experimental results demonstrate that SPME coupled with DESI-MS can sample and analyze CWAs from contaminated media with minimal sample-handling requirements, with potential application within the forensic, defense and homeland security communities. Rapid screening of anabolic steroids in raw urine samples using SPME coupled with reactive DESI-MS was implemented by Huang et al. in 2007 [66]. The spray solvents used in DESI-MS contained hydroxylamine for heterogeneous reactions of hydroxylamine with the carbonyl group of the steroids, providing significant improvements on ionization efficiency of these steroids in undiluted raw urine when compared to conventional DESI. The use of SPME has greatly improved the concentration LODs, and low concentration levels of ketosteroids in raw urine relevant to screening for sports doping (20 ng/mL) can be reached by using a CAR/DVB fiber for preconcentration and direct analysis. Another SPME-DESI-MS method for fast extraction and quantification of a multitude of illicit drugs and pain medications in urine was presented by Kennedy et al. in 2010 [67]. They developed specially-designed SPME fibers coated uniformly with silica-C18 stationary phase for direct immersion extraction of seven drugs from unprocessed raw urine samples, and then removed, rinsed, and analyzed them directly by DESI-MS(MS/MS). The investigated drugs were all successfully quantitatively determined, using an isotopically-labeled internal-standard calibration. Generally, it would be necessary to use an internal standard in an AMS method, because the absolute intensity of ions varies greatly due to the differences in desorption and ionization efficiency [68]. For quantitative analysis, an isotope-labeled internal standard is especially recommended for improvement of reproducibility by compensating for signal fluctuation and matrix effect. 19 urine samples containing drugs from actual use were analyzed, and the experimental results showed good agreements on all the samples screened between the SPME-DESI-MS/MS method and the respective screening and confirmation method (conventional GC-MS and solvent desorption of the SPME fibers followed by LC-MS/MS) for each analyte. Special attention should be paid to the non-exhaustive desorption appearance in this SPME-DESI-MS method, in which a ‘‘wicking’’ effect may occur (the signal intensities remain relatively constant after successive analyses), whereby the outermost molecules in the boundary layer are desorbed, and the molecules embedded within the stationary phase migrate to the boundary layer and are desorbed in the next scan. In such a case, isotope-labeled internal standards play an important role in quantitative analysis for calibrating and compensating for the desorption and ionization efficiency and the instrumental responses. As suggested by this study, even though the desorption of the analytes by DESI is not exhaustive, sufficient sensitivity can still be obtained for this application, and the developed method shows great potential for screening and monitoring the levels of drug molecules in biological samples. Thin-film microextraction (TFME) [69,70], which uses a membrane rather than a fiber to carry the extraction phase, is a variant of SPME. TFME takes advantages of the large surface area of porous sorbents, and the robustness and the easy handling of organic adsorbents. Recently, TFME was directly coupled with DESI-MS for determination of pharmaceuticals and personal-care product

components in wastewater samples, as reported by Strittmatter et al. in 2012 [71]. They used porous, mixed mode C18/strong cation-exchanger (SCE) sorbent TFME blades for microextraction. The coated TFME blades were cut from combs with 12 blades each. After extraction, the TFME blades were air dried and then mounted onto a computer-controlled two-dimensional moving-stage system. A home-built DESI source, of which the electron-spray emitter was mounted on a rotating stage and secured onto a three-dimensional manual moving stage, was used for high-throughput MS analysis (Fig. 6). Using a high-resolution Orbitrap mass spectrometer, detection of analytes [targeted (i.e. carbamazepine and triclosan) and non-targeted (i.e. beta-blockers, non-steroidal antiinflammatory drugs, and UV filters)] was accomplished in wastewater matrices. The TFME-DESI-MS technique, which offers the advantages of cost effectiveness, short analysis times, possibility of miniaturization, and high-throughput capabilities, is suitable for high-throughput, and potentially on-site, screening of trace organic chemicals. The successful coupling of SPME and DESI-MS extends both techniques, being an excellent approach for rapid screening of a larger number of samples. Moreover, this coupled technique works at ambient and open-air conditions, in which automation and high-throughput analysis can be readily achieved. However, because DESI is based on the ESI mechanism, SPME coupled with DESI-MS, in both single-fiber and high-throughput formats, is suitable for determination of polar analytes, especially by using the biocompatible materials, such as C18 and C18/SCE. When using the GC-type coatings, low desorption efficiency of non-polar analytes by polar spray solvent of the DESI source may occur and result in cross contamination or potential loss of non-polar analytes. 6.3. SPME coupled with direct analysis in real time MS Direct analysis in real time (DART) is a well-known APCI-based ambient ionization technique with the analytical strategy of twostep ionization, as proposed by Cody and co-workers in 2005 [72]. An electrical discharge is applied to a gas (typically nitrogen or helium) to form a plasma of excited-state species, which are carried by the heated gas stream towards the sample for desorption and ionization of the small molecules from the sample surface. An SPME fiber or thin film can be held in the sampling module of the DART source, and the pre-concentrated analytes are thermally desorbed by the heated gas and ionized by the plasma, then directly introduced into the mass spectrometer for analysis (Fig. 5c). The first SPME-DART-MS method was investigated by Cajka et al. in 2010 [73]. A 50/30-lm DVB/CAR/PDMS fiber was employed for extraction of beer with both HS and direct-immersion modes. Then, the fiber was desorbed by DART and introduced directly for TOF-MS analysis to obtain chemical fingerprints. The mass spectra fingerprints of HS-SPME obtained (most m/z values under 200 Da) significantly differed from those obtained by direct immersion SPME (m/z values up to 500 Da). This appearance could be explained by HS sampling favoring the extraction of highly volatile analytes, while direct immersion was advantageous for less volatile analytes in the complex matrix. The experimental results revealed that DART-TOF-MS employing the direct immersion SPME sampling approach had the possibility of sorption/ desorption/ionization of relatively polar compounds, which were not volatile enough for HS-SPME sampling and conventional GCMS analysis, so representing a promising tool for obtaining chemical profiles for rapid authentication of food commodities. It is worth mentioning that, in such coupling, the original form of the SPME fiber needs to be fixed manually in the module of DART source, so TFME was developed to cater to this coupling system. TFME coupled with DART-MS was first demonstrated by Mirnaghi and Pawliszyn in 2012 [74], using a reusable UV-dried poly-

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acrylonitrile (PAN)-over C18-PAN TFME for extraction and direct analysis of diazepam in whole blood. The addition of PAN on the C18-PAN coating provided a protective layer on the outer surface of the coating, so improving reusability and reproducibility. In addition, the strongly polar nitrile groups present in the PAN layer were perfectly suited to participate in hydrogen binding with water molecules, forming a hydrated layer that inhibited interaction of blood cells and proteins with the coating surface. To prevent complete blocking of the mesh holes with the coating and to ensure transmission of the heated gas through the coating for efficient desorption and ionization of analytes, the thickness of the coated mesh should be very thin. Moreover, 10 tiny holes were made on the spotting area of the coated mesh using a small needle, so keeping the holes for efficient transmission of the heated gas through the coating. 5 lL of blood sample was deposited on the meshes for 5 min of extraction, and then the coated meshes were rinsed for 5 s with Nanopure water. Afterwards, the dried SPMEcoated mesh was positioned in the transmission module of DART source. Quantitative analysis was performed with an isotope-labeled internal standard and the result showed good linearity and recovery, with an LOD and an LOQ of 0.3 lg/mL and 1 lg/mL, respectively. When compared with the bare mesh, the matrix effect was greatly reduced using the coated mesh, resulting in much cleaner mass spectra. This biocompatible coating was reusable and had a reproducible extraction efficiency for analysis of diazepam from whole blood, and the coupling method attracted interest in the field of clinical and biological analysis. Another TFME-DART-MS application was investigated by Rodriguez-Lafuente et al. in 2013 [75]. They applied a C18-PAN thin-film for direct immersion extraction of cocaine and methadone in urine samples, and then direct analysis by DART-MS/MS. Results showed that TFME improved the detectability, with signal-to-blank ratios of 5 for cocaine and 13 for methadone, respectively, at a concentration of 0.5 ng/mL in human urine. Extraction with TFME provided efficient preconcentration of the analytes, thus increasing the sensitivity of the whole procedure. In addition, by using the coated mesh, the contamination of the ion source by salt residues from the urine samples could be avoided, providing longer intervals between maintenance for MS analysis. 6.4. SPME coupled with desorption corona beam ionization MS Desorption corona beam ionization (DCBI), an APCI-based ionization technique with the analytical strategy of direct desorption/ ionization, was introduced by Ding et al. in 2010 [76]. A high direct current (DC) voltage of 1–5 kV is applied to a metal tube to induce corona discharge from helium atoms. A visible corona-discharging beam is observed when a high DC voltage is applied to the source at a low current (mA). When an SPME substrate is introduced to the DCBI source, the analytes are desorbed by the visible heated corona-discharging beam, and ionized in the gas phase via the reactive species (ions or metastable helium atoms) embedded in the gas stream (Fig. 5d). Li et al. [77] used a PDMS thin film coupled with the DCBI technique for detection of pesticide compounds. The PDMS substrate was dipped in water for microextraction of five pesticides (acephate, isoprocarb, dimethoate, dichlorvos, and dicofol) and then transferred to the DCBI source for desorption and ionization. The LODs of the pesticides were 1 lg/L, The results indicated that the DCBI technique coupled with PDMS sampling is an excellent method for the analysis of organic pesticides in solution. 6.5. SPME coupled with low-temperature plasma (LTP) probe MS The LTP probe is another APCI-based ionization technique with the analytical strategy of direct desorption/ionization, developed

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by Cooks and co-workers in 2008 [78]. LTP is based on the dielectric barrier-discharge mechanism, applying an alternating current (AC) electric field to the discharge gas (He, Ar, N2, or ambient air) to form the plasma through the use of a specially-designed electrode configuration. The sampling plasma probe can operate at low temperature, and interacts directly with the sample being analyzed, desorbing and ionizing surface molecules for MS analysis. SPME coupled with the LTP probe was developed by Almasian et al. in 2010 [79]. In their study, the graphite-based electrode was capable of serving as a sample adsorbent for preconcentration of organic compounds on the electrode surface, and direct desorption/ionization in the LTP source followed by MS detection. Specifically, selected VOCs were preconcentrated on the graphite-based electrode surface via a home-made HS-SPME vial. After adsorption of the samples, the graphite electrodes were transferred into the LTP source for sample desorption and ionization. Because this method enables low-temperature operation without the need for solvents, it exhibits remarkable advantages for those samples that are sensitive to heat or solvents. 7. Future perspective and new opportunities SPME coupled with AMS will be a hot research topic in the next several years, even if its application is still limited. From previous research and the characteristics of both SPME and AMS techniques, we can summarize that coupling SPME to AMS will have the following merits: (1) combining isolation, enrichment, and analysis of analytes into one step under ordinary ambient and open-air conditions with minimal sample pretreatment and without chromatographic separation, possessing the ability of rapid screening and monitoring with low matrix effects, and providing the possibility of automation and high-throughput; (2) capability of on-line monitoring in real time, in situ, in vivo, non-destructive, and reactive analysis, providing chemical information at the molecular level; (3) high sensitivity and excellent specificity for the detection of trace analytes from samples with a complex matrix composition; (4) possessing the ability to confirm targeted components, to elucidate non-targeted components, and to identify unknown compounds when using a high-resolution hybrid mass analyzer (e.g., QTOF, IT time-of-flight (IT-TOF), and linear IT Orbitrap (LTQ-Orbitrap)) for MS analysis; and, (5) providing acceptable linearity, reasonable repeatability and reproducibility for quantitative/semi-quantitative analysis of the targeted compounds with an internal standard for MS analysis. The rapid development of ambient ionization techniques in the early twenty-first century gives opportunities for innovation of SPME coupled with AMS, with promising prospects in environmental, food, forensic, biological, pharmaceutical analysis, clinical, high-throughput, and in vivo applications. Both SPME [69,70,80] and AMS [54,81,82] are powerful tools for in vivo analysis, and the combined application of these two techniques will give a new insights in in vivo studies in the future. Acknowledgements Financial support from National Natural Science Foundation of China (NSFC, Nos. 21277177, 21307167), the Scientific and Technological Project of Guangdong Province (No. 2011B060100005), the South China Sea Branch of State Oceanic Administration and the

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Administration of Ocean and Fisheries of Guangdong Province is gratefully acknowledged.

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