Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review

Merkle, Sybille; Kleeberg, Kim Karen; Fritsche, Jan

doi:10.3390/chromatography2030293

Open AccessReview

Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review

by

Sybille Merkle

^1,†,

Kim Karen Kleeberg

^2,† and

Jan Fritsche

^1,*

¹

Faculty Life Sciences, Department Nutrition & Home Economics/Food Science, Hamburg University of Applied Sciences, Ulmenliet 20, 21033, Hamburg, Germany

²

ODOCON Dr. Kim Kleeberg & Dr. Jan Streese-Kleeberg GbR, Amtsstraße 7, 22143, Hamburg, Germany

^*

Author to whom correspondence should be addressed.

^†

These authors contributed equally to this work.

Chromatography 2015, 2(3), 293-381; https://doi.org/10.3390/chromatography2030293

Submission received: 1 April 2015 / Revised: 10 June 2015 / Accepted: 11 June 2015 / Published: 30 June 2015

(This article belongs to the Special Issue Solid Phase Micro-Extraction)

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Abstract

:

Solid-phase microextraction (SPME) is a simple, sensitive, rapid and solvent-free technique for the extraction of analytes from gaseous, liquid and solid samples and takes a leading position among microextraction methods. Application of SPME in sample preparation has been increasing continuously over the last decade. It is most often used as an automatized fiber injection system coupled to chromatographic separation modules for the extraction of volatile and semivolatile organic compounds and also allows for the trace analysis of compounds in complex matrices. Since SPME was first introduced in the early 1990s, several modifications have been made to adapt the procedure to specific application requirements. More robust fiber assemblies and coatings with higher extraction efficiencies, selectivity and stability have been commercialized. Automation and on-line coupling to analytical instruments have been achieved in many applications and new derivatization strategies as well as improved calibration procedures have been developed to overcome existing limitations regarding quantitation. Furthermore, devices using tubes, needles or tips for extraction instead of a fiber have been designed. In the field of food analysis, SPME has been most often applied to fruit/vegetables, fats/oils, wine, meat products, dairy and beverages whereas environmental applications focus on the analysis of air, water, soil and sediment samples.

Keywords:

SPME; food analysis; environmental analysis; volatile compounds; aroma; off-flavor; GC; LC; SBSE

Graphical Abstract

1. Introduction

The quality of trace analysis in food and environmental samples mainly depends on the selected sample treatment [1]. There is an increasing demand for strategies minimizing the analyte-matrix interferences, especially in case of volatile compounds occurring at low concentrations in complex matrices. These conditions require the reduction of final volumes of analytes to furnish higher concentrations of analytes at low quantification limits [2]. Moreover, there is a demand for environmentally sustainable procedures that apply reduced volumes of organic solvents [2]. Another aspect of analytical method development is automation [3]. There are numerous advantages of automation, e.g., lower time consumption, increase of simplicity, lower probability of sample contamination and higher repeatability. These are just a few reasons for the widespread application of solid phase microextration (SPME) techniques [3].

SPME is a relatively recent and easy to automate technique for the extraction of analytes from gaseous, liquid, and solid matrices. The simple, rapid and solvent-free technique was introduced in the early 1990s by Pawliszyn and coworkers [4,5,6]. The SPME process is composed of two basic steps: (i) partitioning of analytes between the extraction phase and the sample matrix and (ii) desorption of concentrated extracts into an analytical instrument [7]. Due to the combination of sampling, extraction, pre-concentration and sample introduction into an analytical instrument in one single step, SPME has gained popularity in many fields of application in recent years, especially in routine laboratories and industrial applications [8,9,10]. Areas of application have been steadily growing, including food packaging and environmental uses.

Automatised fiber injection systems hyphenated with gas chromatographic (GC) and gas chromatography-mass spectrometry (GC-MS) separation modules are the most popular instruments combined with SPME in use [11]. These instrumental assemblies have been successfully applied to a wide variety of compounds, especially for the extraction of volatile and semivolatile organic compounds from complex sample matrices [12]. Due to the inaccessibility of weakly volatile or thermally labile compounds to GC or GC-MS, SPME was also developed for direct coupling to high-performance liquid chromatography (HPLC) and liquid chromatography-mass spectrometry (LC-MS) [12].

A number of reviews on SPME have already been published, dealing with food analysis [12,13,14,15], environmental analysis [16,17,18,19,20,21], calibration methods [22], SPME coatings [11,23,24,25,26], drug analysis [12,27], the analysis of specific compounds [28,29] and specific SPME techniques [30]. In addition, general overviews of SPME techniques and contemporary developments are provided by others [2,6,17,31].

This review provides a comprehensive overview on latest trends of SPME method development including new techniques and new devices, as well as essential parameters in SPME processes. The latest development of fiber coating development and the application of nanotechnology in SPME technology are also covered extensively by the present review. Moreover, this review reports for the first time simultaneously on two important life sciences SPME application areas—food and environmental applications published over the last decade (2005–2015).

2. SPME Techniques

The underlying principle of the SPME methodology is based on the partitioning of analytes between a coated fiber and a sample. The reusable fiber is coated with a stationary phase, which can be a liquid polymer, a solid sorbent, or a combination of both [9]. A mass transfer begins after exposure to the vapour phase above a solution (HS-SPME) or direct immersion in the solution (DI-SPME). This process is driven by the second law of thermodynamics. Thereby the chemical potential of each compound should be equal. Once equilibrium is reached, the extracted compounds on the fiber are desorbed and inserted into the injection port of a chromatograph. The release of the analytes is accomplished in GC by thermal desorption and in HPLC by dissolution and subsequent injection of the elution solvent [32]. Both methods lead to detection of the substances by the analytical instrument detector. The application of SPME-LC has lagged behind SPME-GC, which might be due to the small number of commercially available SPME sorbents for LC applications, the lack of commercially available interfaces, long equilibration times, and the lack of automation [2].

The aim of SPME is to reach the equilibrium between the sample matrix and the coating of the SPME device as rapidly as possible [33]. Further exposure of the fiber after equilibrium has been reached does not increase the concentration of extracted compounds. For that reason, sample extraction and pre-concentration could be carried out in one single step [34].

SPME has several advantages over traditional extraction methods. It is not just a rapid, simple and solvent-free method, but it is also sensitive, provides linear results for a wide range of concentrations and analytes [2]. Notwithstanding the low concentrations of analytes, quantitative or semi-quantitative data are provided and losses that can occur during the extraction, concentration and clean-up steps of traditional sample procedures are mostly avoidable [2].

One of the main drawbacks of SPME techniques is the limited number of commercially available stationary phases (fiber materials) only roughly covering the scale of polarity of target analytes. In particular, the extraction of polar analytes from samples with a polar matrix poses a problem [35,36,37,38,39]. Other challenges are the relatively low recommended operating temperature (240–280 °C), the instability and swelling in organic solvents, breakage of the fiber, stripping of coatings, bending of the needle, and the cost as well as the limited lifetime of the fiber [2,36]. Furthermore, sample carry-over may occur [40] and high molecular weight compounds cannot be analyzed in combination with GC [36]. In certain cases, low extraction efficiencies are reported, in particular in case of very volatile, polar, or thermally unstable analytes [2]. The early theoretical and practical aspects of SPME have undergone continuous technical development over the years.

SPME techniques cover a wide range of sampling techniques, including field, in situ and air sampling. The two different implementations fiber SPME and in-tube SPME are frequently used [9]. Detailed SPME theory has been explained in a number of reviews [6,15,41,42].

2.1. Fiber Solid-Phase Microextraction

“Fiber”-SPME devices have been developed and currently represent the widespread SPME technique which comprises a fiber holder and a fiber assembly [41]. The fiber assembly consists of a 1- to 2-cm long retractable SPME fiber [41] and a built-in coated fiber that looks like a modified syringe [6,33]. After the sample is placed in a vial sealed with a septum-type cap, the SPME needle is pierced through the septum and the fiber is extended into the vial. The analytes partition between the sample matrix and the extraction phase until equilibrium of concentrations is reached. The maximum sensitivity is achieved and a proportional relationship is obtained between the amount of the extracted analyte by the SPME fiber and its initial concentration in the sample [43,44].

2.2. In-tube Solid-Phase Microextraction

The more recent in-tube (IT) SPME was developed by Eisert and Pawliszyn [45] for application with HPLC or liquid chromatography (LC)-MS because fiber SPME could barely withstand aggressive HPLC solvent conditions [6,45]. In-tube SPME overcomes fiber-related drawbacks such as fragility, low sorption capacity and bleeding from thick-film coatings. It uses an open tubular fused-silica capillary in which the extraction phase is either an inner surface coating or a sorbent bed. Consequently, in-tube SPME devices exhibit considerably higher mechanical stability than fiber SPME devices. Another reason for the development of in-tube SPME was the lack of automation in the use of SPME coupled to HPLC [2]. By means of automation it is possible to perform extraction, desorption and injection simultaneously [6,12]. The advantages of automated systems compared to manual techniques are shorter total analysis times as well as higher accuracy and precision [2]. A drawback of in-tube SPME is the tendency of the capillary to clog up. This can be avoided by working with samples without interfering phases like particles or macromolecules [2]. Moreover, the enrichment factor is reduced compared to fiber SPME [2]. Kataoka [30] and Globig and Weickhardt [46] provided an overview of automated sample preparation using in-tube SPME and applications of this technique in environmental, clinical and food analysis, mainly for the determination of polar and thermolabile compounds.

In-tube SPME can be categorized into methods using extraction coatings where the coating is used as an internal extraction phase immobilized in the capillary wall and methods using extraction fillings where the extraction phase is a sorbent packing, as demonstrated in Figure 1 [47].

Figure 1. “Classical” solid-phase microextraction (SPME) fiber compared with in-tube fibers, adapted from Nerín et al. (2009) [2], reprinted with permission of Springer Verlag.

The latter techniques are particularly similar to the previously developed solid-phase extraction (SPE) and use different sorbent types. Both in-tube SPME techniques can be easily applied with existing autosampler systems. Several in-tube devices with extraction fillings have been developed, firstly in 2004 by Kubinek et al. [48], and are now commercially available, for example as in-tube extraction (ITEX) [2].

There are two modes of extraction in in-tube SPME. In the static mode, the analytes are transferred to the stationary phase by mere diffusion, while in the dynamic mode, the sample is actively drawn through the tube by repeated draw/eject cycles [49]. When the equilibrium is virtually reached, the extracted analytes are desorbed from the capillary either by direct transfer to the liquid chromatographic column or by a stream of the mobile phase. In case of a stronger attachment of the analytes to the capillary, desorption is achieved by a static desorption solvent which is subsequently injected into the chromatographic system [50]. The desorbed analytes are carried through the column and separated and detected by a mass selective detector [9].

Even though in-tube SPME was originally developed for HPLC applications, it can also be used with other instrumental equipment, for example capillary electrophoresis (CE) or GC [30]. On-line coupling to GC can be achieved by open-tubular trapping (OTT), which is mainly applied to HS samples [30]. In OTT, desorption of the analytes is carried out with a small amount of solvent or by thermal desorption. However, OTT is characterized by a complex instrumental setup and unfavorable sampling conditions, such as high pressure drop from long traps and limited flow rates [2].

In-tube SPME techniques have been developed further and modifications like wire-in-tube and fiber-in-tube techniques have emerged [51]. The wire-in-tube technique contains an additional stainless steel wire inserted into the extraction capillary. Thereby the surface of the coating material stay more or less the same, but the internal volume of the capillary is significantly reduced. A change in the phase ratio takes place which leads to preconcentration of the sample. This results in a more effective extraction [51]. In the fiber-in-tube technique, the extraction phase comprises several hundreds of delicate filaments of polymeric material packed longitudinally into a short capillary. This composition leads to enhanced pre-concentration of analytes [9].

2.3. Cooled Coated Fiber Device

Apart from in-tube SPME, procedures using a cooled fiber and other methods that do not necessarily rely on the use of a fiber have emerged. These new developments will be explained in the following section.

A recent development in the application of HS-SPME is an internally cooled coated fiber device (CCF) or cold fiber HS-SPME device. The main reason for developing this method was the improved release of analytes from the interfering phases in complex matrices [52,53].

For desorption of analytes from solid particles it is often useful to increase the extraction temperature. However absorption of analytes by the fiber coating is an exothermic process, therefore this process leads to a decrease of the partition coefficients [2]. The aim is to accelerate the mass transfer process and simultaneously increase the distribution constants of analytes [2]. For that purpose, an internally cooled coated fiber device was developed heating the sample matrix at the same time as the fiber coating is cooled down. Especially for matrices with high viscosity or for volatiles with low partition coefficients CCF is useful [2]. This technology was successfully employed for the first time to extract analytes from various environmental matrices [52,53,54]. CCF was also successfully applied to food analysis, such as extracting volatiles and semivolatiles from tropical fruit pulps by Carasek and Pawliszyn (2006). They stated that the cold fiber was the most suitable fiber for the purpose of extracting volatile compounds from the five fruit pulps studied [55]. Additionally, it was reported that cold-fiber HS-SPME offers more sensitivity and higher sample throughput than conventional HS-SPME [56]. One drawback, however, of the increased fiber capacity through this cooled coated fiber is the loss of selectivity [2]. Not only the analytes but also the interferences are extracted exhaustively onto the coating [6]. CCF has been miniaturized and automated first by Chen and Pawliszyn (2006) [53] and is nowadays routinely used (see Figure 2) [2].

Figure 2. Internally cooled SPME device (A) and its automation (B) according to Chen and Pawliszyn (2006) [53], reprinted with permission of the American Chemical Society.

2.4. Non-fiber SPME Techniques

Non-fiber SPME techniques developed over the last two decades can be divided into static methods using sample stirring and dynamic techniques relying on the flow-through of the sample. The former include stir-bar sorptive extraction (SBSE) and thin-film microextraction (TFME). The latter can be classified as in-needle and in-tip SPME [21,47]. Figure 3 provides an overview of these new techniques.

Figure 3. Overview of SPME techniques–techniques marked in grey are more recent developments (SBSE: stir-bar sorptive extraction; TFME: thin-film microextraction; SPDE: solid-phase dynamic extraction; FNME: fiber-packed needle microextraction; MEPS: microextraction by packed syringe).

In stir-bar sorptive extraction (SBSE), a magnetic stir bar coated with polydimethylsiloxane (PDMS) is stirred in or positioned above an aqueous sample [21,50]. After extraction, the stir bar is removed, which is usually done manually with tweezers. In case of a SBSE-GC coupling, desorption of the analytes is induced by inserting the bar into the heated GC injection port or by placing it in a small vial and back-extracting with a few microliters of an appropriate liquid solvent. In case of LC, the mobile phase can be added directly to the stir bar. Compared to fiber SPME, SBSE has a higher concentration capacity as the PDMS and the layer is 50 to 250 times thicker [21,50]. However, SBSE is not yet as widely accepted as fiber SPME due to the difficulty of achieving full automation and the limited number of commercially available coatings [21,50]. Dual-phase twisters using different carbon-based adsorbents in addition to PDMS are being developed at the moment as well as other phases covering a wide range of polarities and containing polymers like poly(methacrylic acid stearyl ester-ethylene dimethacrylate) and polypyrrole [21,50].

In thin-film microextraction (TFME), a flat film with a high surface area-to-volume ratio is used as the extraction phase [57,58]. This higher surface-to-volume ratio as well as an increased volume of extraction phase result in an enhanced sensitivity compared to conventional SPME without sacrificing the sampling time.The first step in TFME is the conditioning of the blades. Subsequently, the blades are exposed to the sample to extract the analytes [57,58]. A 96-well plate is placed on an orbital shaker for agitation. After washing with an appropriate solution to remove interfering substances, the blades can be transferred to a second 96-well plate containing the desorption solvent which is eventually injected into the chromatograph. A robotic workstation providing full automation of all the TFME steps including conditioning, extraction, washing and desorption may be used [57,58]. Moreover, coupling to liquid and gas chromatographic systems is possible [57,58]. TFME can be applied to gaseous, liquid and solid samples and is used in different sampling formats. In combination with liquid chromatography, it can be easily performed in a 96-well plate thus allowing for simultaneous analyses. The first TFME application used 96 pieces of small cut SPE disk membranes mounted on a stainless steel wire for support [57,58]. Nowadays, the extraction phase is coated onto a blade-shaped substrate and PDMS is the most common sorbent [57,58].

In case of coupling TFME to a gas chromatograph, desorption may also be achieved by direct thin-film injection inside a thermal desorption unit adapted to the special requirements of the thin-film with its particular geometry. The thermal desorption unit contains a large volume inlet and a programmed temperature vaporizer. The latter cryotraps the desorbed compounds. Once the thermal desorption process is completed, the cryotrap is heated and the trapped substances are transferred to the GC column [57,58]. In contrast to solvent desorption, direct thin-film injection allows for a complete introduction of the extracted analytes into the chromatograph [57,58]. As a thermal desorption unit is not always available and as solvent-desorption can be carried out in full automation, the latter is often used in practice [57,58].

In-needle SPME methods use a needle instead of a tube for extraction. In 2001, Koziel et al. [59] developed a needle trap (NT) device with quartz wool in order to trap particulate matter and aerosols in air. Wang et al. [60] used this development as a basis for the design of a sorbent-packed NT device for the analysis of VOCs in gaseous samples. The NT device was particularly robust because the quartz wool was protected by the needle. In 2006, Saito et al. [61] developed an in-needle extraction device for the analysis of VOCs using a copolymer of methacrylic acid and ethylene glycol dimethacrylate, which made the procedure more convenient [50,62,63,64]. This needle showed high extraction performance and thermal stability and was therefore suitable for typical GC applications. Also, the extracted analytes were very stable and could be analyzed even after several days of storage at room temperature [50,62,63,64].

Today, in-needle SPME methods can be classified as solid-phase dynamic extraction (SPDE), microextraction by packed syringe (MEPS) and fiber-packed needle microextraction (FNME). In SPDE, the analytes derived from a liquid or HS sample are concentrated onto a film of PDMS and 10% activated carbon coated on the inner wall of a stainless steel needle of a 2.5-mL gas-tight syringe by repeatedly moving the plunger of the syringe up and down [50,62,63,64]. Recovery of the trapped substances takes place by heat desorption directly into the GC injection port. Full automation is possible [50,62,63,64]. The first commercially available SPDE devices were developed in 2000 by Chromtech (Idstein, Germany) [50,62,63,64]. Lipinski [64] was the first who applied SPDE for the analysis of pesticides in water [50,62,63,64]. A significant advantage of SPDE over fiber SPME is the larger coating volume, which is approximately four to five times the volume of a 100-µm fiber and causes an increased in concentration capacity. Moreover, the extraction time is shorter, the repeatability better and the mechanical stability of the SPDE device higher [50,62,63,64]. On the other hand, carry-over may be an issue in SPDE as the analytes tend to remain on the inner surface of the needle after heat desorption [50,62,63,64].

Microextraction by packed syringe (MEPS) is a miniaturized form of the classical SPE where sample extraction takes place in a packed bed [50,62,63,64]. MEPS can be on-line coupled to liquid and gas chromatographic systems and is easily performed in automation. In contrast to SPE, the solid packing material is inserted directly into a needle coupled with a syringe and not into a separate column [50,62,63,64]. Furthermore, the sample preparation time as well as the volume of the sample and the organic solvents are considerably reduced and the MEPS sorbent can be used up to 100 times [50,62,63,64]. Sorbent materials include reversed phase (C2, C8 and C18), normal phase (silica), restricted access material (RAM) and molecular imprinted polymers (MIPs) [50,62,63,64]. After the packed bed has been conditioned, the sample solution is pumped several times through the syringe. The sorbent is then washed and the analytes are eluted either with an organic solvent which is injected into the chromatograph afterwards or with the LC mobile phase directly into the chromatographic injector [50,62,63,64]. Fiber-packed needle microextraction (FNME) is an alternative technique using fiber instead of particle materials inside the needle [50,62,63,64].

In-tip SPME differs from MEPS in that the solid packing material is inserted into pipette tips rather than a needle. As extraction is generally done off-line, only parts of the sample are injected into the chromatograph resulting in a lower sensitivity compared to on-line MEPS [50]. On the other hand, a major benefit of in-tip SPME is that several samples can be handled in parallel. Automation is possible by commercially available systems using 96-well extraction plates and a rosdbot [50]. Sorbents used for in-tip SPME comprise silica and methacrylate monoliths. In contrast to fiber and in-tube SPME, in-tip SPME and MEPS provide quantitative recoveries [50]. On the other hand, these techniques exhibit only a moderate enrichment capacity and they have a tendency towards carry-over [50].

3. SPME Process

This section will provide an overview of the different available extraction and desorption techniques for SPME applications. The variety of these techniques is shown in Figure 4. Furthermore it will provide recent information on commercially available fiber coatings as well as current developments in fiber coating procedures. Finally, aspects of quantitation techniques applicable for SPME will be described.

Figure 4. Extraction and desorption techniques for SPME applications.

3.1. Extraction and Desorption Techniques

The extraction process of fiber SPME can be conducted in three common ways: direct or immersion extraction (DI), headspace (HS) and membrane protection extraction (see Figure 5 and Figure 6). In Direct SPME the fiber is directly immersed in a gaseous or liquid sample. In case of a liquid sample certain agitation is required to reduce the extraction time. For volatile compounds in gaseous samples, the natural occurring air flow is often sufficient to reach the equilibrium [65].

In HS-SPME the fiber is exposed to the vapor phase above the liquid or solid sample [9]. The advantages are the protection of the fiber from damaging substances of the sample matrix and a possible adjustment of pH conditions. Only for very volatile compounds a significant difference between DI-SPME and HS-SPME techniques was observed [66]. The extraction kinetics are governed by Henry’s law [67]. If the Henry’s constant of a given substance is high, then the concentration of the compounds in the headspace is high, too. Under these conditions a rapid extraction from the headspace takes place. In case of aqueous sample matrices, volatile and non-polar compounds are extracted faster than semi-volatiles and polar volatiles [65]. Moreover, increasing sample temperature and agitation efficacy may decrease the extraction time [65].

Figure 5. SPME procedure for total-immersion and headspace sampling according to Nerín et al. (2009) [2], reprinted with permission of Springer-Verlag.

Figure 6. SPME procedure for membrane protection sampling according to Basheer and Lee (2004) [68], reprinted with permission of Elsevier B.V. (HFM: hollow fiber membrane).

In static headspace sampling, diffusion occurs between the fiber and the sample without any interference, whereas dynamic headspace sampling involves air movement devices like air sampling pumps that serve to move the headspace air [69]. To collect and pre-concentrate the headspace gas, the headspace air is transferred into another chamber where the SPME filament or another extraction trap is present [70].

An advantage of static headspace sampling in comparison with dynamic headspace sampling is that static headspace sampling does not require careful calibration processes and expensive air sampling pumps [71]. However, differences in temperature and pressure affect the efficiency of the static sampling process as well as wall effects from sampling containers that also play a role [69]. In summary static headspace sampling shows advantages of simplicity, sensitivity, selectivity and ease of automation over dynamic headspace sampling techniques [72].

The third extraction technique is the membrane protection extraction. Extraction which is typically applied to the analysis of samples containing interfering compounds such as proteins, humic acids and fatty material [68]. It is important to prevent the extraction of these high molecular weight compounds which potentially complicate, or even inhibit, sample analysis [68]. The extraction is conducted using a membrane which is selectively permeable for analytes of interest [68]. Membran protection extraction is slow in comparison with other extraction techniques, but effectively extracts compounds with low volatility [65].

All extraction procedures require an optimization of performing parameters in order to achieve good reproducibility, resulting from a compromise between sensitivity and extraction time. Method optimization parameters include agitation conditions, extraction time, extraction technique, temperature, headspace and sample volume vial shape, condition of fiber coating, depth of the SPME fiber inside the vial, geometry of the fiber, and pH conditions [33,73].

After extraction, target analytes are introduced into a suitable instrument for detection [74]. Desorption can be proceed in two different ways; static mode, by dipping the fiber into the mobile phase or solvent for a specified period, and dynamic mode, where the analytes are desorbed into a flowing mobile phase [65]. It is important to identify desorption solutions that completely remove the analytes from the fiber, eliminating carry over, but that do not damage the sorbent material or degrade the analytes of interest [74].

In the case of a GC coupling the analytes are released by thermal desorption, for which the SPME fiber is inserted into the GC inlet and heated to temperatures that increase analyte volatility sufficient for their release. The analytes are physically released from the sorbent through heating whereby the volatility of the target compounds is increased. The carrier gas flow rate and the injector temperature are critical factors for effective thermal desorption [75]. High desorption temperatures effect rapid transfer of the target analytes from the injector to the chromatographic column, but may reduce the stability of the sorbent and lead to bleeding of the polymeric material [73]. Therefore the thermal stability of the fiber coating determines the upper desorption temperature limit [74].

Combined with an HPLC interface desorption is performed by solvent extraction in the desorption chamber [12], namely liquid desorption. Liquid desorption is conducted using a small volume of suitable solvents to transfer the target analytes to the analytical instrumentation [74]. Desorption of the analytes can also be conducted using a polar organic solvent, such as methanol or acetonitrile [65]. This approach is often used in combination with liquid chromatography (LC) [65]. For thermally labile compounds liquid desorption has the advantage of not requiring elevated temperatures compared to thermal desorption [74]. Liquid desorption is the combination of SPME with HPLC where desorption is achieved using an appropriate extraction solvent in a desorption chamber. Individually designed desorption interfaces have been reported [51,76]. There is a commercially available device that enables desorption of all analytes directly into the LC injector [65].

Another form of desorption is conducted by means of laser techniques. SPME in combination with matrix-assisted laser desorption/ionization mass spectrometry was reported [74,77].

3.2. Derivatization

Under some conditions target analytes cannot be efficiently extracted or determined by the applied method due to the physicochemical properties of the analytes or the composition of the matrix. In these cases derivatization may modify the analytes and improve their extraction, desorption, or chromatography relative to matrix interferences. Derivatization might also improve the separation performance of analytes with poor chromatographic behavior [78]. Desired requirements for derivatization reactions are a fast and quantitatively formation at room temperature as well as a high stability and solubility in the applied solvent or phase [74].

There are different derivatization methods applicable for SPME (see Figure 7). The main difference of the various derivatisation approaches is the point of time when the derivatization reactions take place. Firstly, the derivatization reaction can be conducted by directly adding the derivatizing agent to the sample, named direct or pre-extraction derivatization. Furthermore, the reaction can take place during the extraction process (simultaneous derivatization and extraction) or immediately after that (derivatization following extraction or post-extraction derivatization). Under these conditions, the derivatization is performed by exposing the enriched SPME fiber to the derivatization agent solution after the extraction [74]. Another option is the post-extraction derivatization, which take place in the injection port of a gas chromatograph through thermal conversion (derivatization in GC Injector Port (see Figure 7).

Figure 7. Classification of derivatization techniques in SPME, adapted from Pan and Pawliszyn (1997) [79].

In direct derivatization the chemical structure of the analyte is modified. Afterwards the common extraction and desorption procedures are performed. The obtained derivatives show better performance characteristics than the original analytes by respective extraction phases [79]. Direct derivatization is a simple derivatization technique and is frequently applied and easy to implement [74].

Another option for derivatization takes place simultaneously with the extraction process or after the extraction process. Due to the combination of extraction and derivatization this procedure is quite convenient to apply and the time needed for the analysis can be significantly reduced [74]. Barriers of this procedure are the compatibility of the derivatizing agent with the extraction phase and with the applied extraction method [74]. However, optimization of the composition of the derivatizing agent is frequently required [74]. Therefore, post-extraction derivatization is preferred when extraction and derivatization cannot be conducted simultaneously or the partition coefficients of the derivative agents are lower than those of the analytes [74].

Injection port derivatization is a post-extraction derivatization procedure, for which analytes containingpolar functional groups such as –OH, –SH, –NH, and –COOH are thermally converted into derivatives.

Derivatization in food analysis for example has been applied for analysis of chlorophenols and chloroanisoles in wine [80] or formaldehyde in fish products [81]. Interesting fields of application of derivatization in environmental samples are, for example, the analysis of formaldehyde and other carbonyl compounds in indoor air [82], organometallic compounds (mercury, lead, tin) in river and seawater [83] or organotin compounds in sediment [84]. Further applications of derivatization in the fields of food [80,81,85,86] and environment [82,83,84,87,88,89,90,91,92,93,94,95,96] are provided in Section 3 and Section 4, respectively. Recently, commonly used derivatization agents were reviewed by Quintana and Rodriguez [97] and Stalikas and Fiamegos [78].

The main drawback of derivatization, however is the formation of artefacts or increased matrix interferences [74]. Furthermore, derivatization reactions favors the generation of waste and therefore, it is recommended to avoid them where possible [98].

3.3. Salt-addition

The addition of salting-out reagents, usually sodium chloride or sodium sulphate, increases the ionic strength of the solution. The extraction efficiency is improved by decreasing the solubility of analytes, thus increasing the amount of analyte sorbed on the fiber. However it should be realized that this effect depends on the particular analyte and salt concentration in the sample [99].

3.4. Fiber Coatings

The choice of the fiber coating mainly depends on the nature of the analytes. Before a sample is analyzed, the fiber should be cleaned in order to remove contaminants that might increase background noise in the chromatogram [65]. Frequently used commercially available fiber coatings for food and environmental analysis are polydimethylsiloxane (PDMS), polyacylate (PA), Carboxen (CAR; a carbon molecular sieve), divinylbenzene (DVB), and Carbowax (CW; polyethylene glycol). These fibers are available in various coating combinations, blends or copolymers, film thicknesses, and fiber assemblies. The variety is constantly growing increasing the number of applications [11]. The most widespread examples of absorption or liquid fiber coatings are PDMS and PA. Mixed coatings in which the primary extraction phase is a porous solid are also available [100]. CAR/PDMS and DVB/CAR/PDMS fibers provide the best extraction efficiencies for a wide range of analytes with different polarities and molecular weights [65]. PDMS has the ability to withstand high temperatures up to 300 °C and shows great stability [65]. PDMS fibers are better suited to the analysis of nonpolar analytes, whereas PA fibers are more useful to extract polar analytes [65]. DVB is also a polar porous solid coating and therefore efficient in extracting polar compounds such as disulfides and trisulfides [101]. Bipolar compounds like alcohols, aldehydes, ketones, ethers and carboxylic acids are often most efficiently extracted by fibers consisting of a combination of e.g., non-polar material (PDMS) and a polar material (DVB) [65]. Another appropriate choice to extract analytes with different polarities are coating mixtures consisting of CARB/PDMS or DVB/CARB/PDMS [65]. However, a disadvantage of this coating material is the displacement effect of analytes with a lower affinity to the coating [102]. Apart from commercially-available sorbents, different new coating procedures have been applied to expand coating types in the commercially applicable and reproducible SPME devices. These new approaches also address biocompatibility, on-site compatibility, selectivity and sensitivity limitations [103,104]. One can differentiate coating procedures based on the sorbent type into dipping and physical agglutinating methods, sol-gel technology, chemical grafting, electrochemical methods, electrospinning, liquid-phase deposition and the hydrothermal method [24]. Further new coatings are polypyrrole polymer coatings [105], molecularly imprinted polymers (MIPs) [106] and immunoaffinity systems [107].

The following section gives a brief overview of the variety of coating procedures. More detailed information, including advantages and disadvantages of various coatings, is available in other recent technology reviews [11,24].

Table 1 lists the different coating procedures, sorbent materials and examples of applications.

Physical coating processes were the first to be developed and constitute the most convenient procedures [11]. They are compatible with nearly all kinds of sorbent materials. Application of new coatings such as carbon nanomaterials (CNMs), ordered mesoporous materials, ionic liquids (ILs) and polymeric ionic liquids (PILs) is very simple. These coatings can be used for fused silica fibers as well as for metal wires. The Dipping process is conducted by placing a fiber in a concentrated organic solvent solution of the material to be deposited for a short time. Crosslinking of the deposited material can take place after evaporation of the solvent by drying. In case of a metal wire, support material can be attached using a sticky paste layer of the sorbent (epoxy or similar) [11,24].

An alternative to physical coating procedures is the sol-gel technology developed by Chong and co-workers (1997) [108]. Sol-gel process is based on the building of an inorganic network consisting of a colloidal suspension (sol) and gelation of this sol which forms a network in a continuous liquid phase (gel). The extraction phase can be chemically bonded to the silica thus producing highly cross-linked phase networks [2]. The precursors for the synthesis of the colloids consist of a metal or metalloid element (alkoxysilanes tetramethoxysilane or tetraethoxysilane (TMOS/TEOS) surrounded by various reactive ligands [109]. The sol-gel fibers therefore exhibit higher thermal stability, hydrolytic stability towards organic solvents as well as high and low pH solutions, and higher surface areas. Molecularly selective coatings can be prepared by using substances with specific functionalities such as crown ethers or ß-cyclodextrin derivatives [2]. Kumar et al. (2008) published a review on the development of sol-gel methods including various applications (e.g., PAHs, aromatic amines, phenols and pesticides) in environmental, pharmaceutical and food analysis [2,25]. A large number of new extraction phases with unique features have been introduced by application of the sol-gel technology. The most important advancements are summarized in Table 1. As there is no standardized sol-gel procedure, this new technology lacks inter-laboratory reproducibility [11]. Further disadvantages of the sol-gel technology are the large number of optimization factors and the fragility of the fused silica fiber used mostly as a support. Due to these limitations of the sol-gel procedures, simple chemical grafting has been developed. In this technique, the surface of the sorbents and supports are chemically modified in order to allow them to react with each other. Possible sorbents applicable to both metal wires and fused silica fibers are nanomaterials, antibodies, MIPs, ILs and PILs [24].

Electrochemical procedures are able to generate extraction films of variable thickness on an unbreakable metal support at a low cost and with a simple setup. Coatings show high thermal stability, have a long shelf life and some may provide higher capability to sorb polar compounds. Electrochemical procedures may be divided into electrodeposition, anodization and electrophoretic deposition (EPD) [110]. Electrodeposition can be used to create coatings with porous structure and refers to the deposition of a metallic or conductive polymer (CP) coating onto a base material by the electrochemical reduction of metal ions or electropolymerization of CPs from an electrolyte. This technique has been applied to the coating of metal supports with CPs, CPs nanocomposites, PIL composites and metal oxides [24]. Anodized metal wires were first developed by Djozan and co-workers in 2001 [111]. They anodized aluminum wires by direct current in sulfuric acid to obtain a porous layer of aluminum oxide on the aluminum surface [110]. Electrophoretic deposition (EPD) is a direct particle assembly method that is based on particle coagulation rather than an electrochemical reaction. Charged nanoparticles from a solution are deposited onto a substrate using an electric field [24]. Single-walled carbon nanotube (SWCNT) [112,113,114] and multi-walled carbon nanotube (MWCNT) [115] coatings have been prepared on the surfaces of platinum and stainless steel wires, respectively. In EPD, adsorption takes place in the spaces between the nanotubes and amorphous carbon in a three-dimensional network of nanotubes [24].

In electrospinning, a solution of a high molecular weight polymer with high viscosity is drawn into nanofibers by repulsive electrostatic forces. Zewe et al. (2010) were the first to prepare nanostructured polymeric based SPME coatings by electrospinning. They used a polymeric negative photoresist, SU-8 2100, which was converted to carbon by pyrolysis. The resultant SPME devices were used to extract both nonpolar and polar compounds [116].

Lin et al. (2008) employed liquid phase deposition (LPD) to produce a nanomaterial-based in-tube SPME coating with increased extraction efficiency [117]. Due to the chemical bonding between the substrate and the coating, LPD offers highly stable nanomaterial thin film coatings. It has been used to deposit thin films of SiO₂, TiO₂, SnO₂, ZrO₂ and the three dimensional transition metal oxides V, Cr, Mn, Fe, Co, Ni, Cu, Zn, In (individually or combined) [24].

The hydrothermal growth technique has been used to produce ZnO nanoparticles on the surface of fused silica fiber and stainless steel wire [118,119]. At first, the procedure consisted of two steps, seeding and growing [24]. A simplification to a single step was realized by Alizadeh et al. (2011) [118]. It is possible to coat 50 fibers at once. The hydrothermal method is used to produce organic framework (MOF) SPME coatings which offer high thermal and mechanical stability and exceptionally large surface areas [24].

In their review on procedures for the preparation of SPME coatings, Aziz-Zanjani and Mehdinia (2014) conclude that this variety of coatings forms a good basis for the development of more designable structures. Durable and stable SPME devices can be produced by metal supports and chemical bonding between support and sorbent. However, poor selectivity and limited applicability to samples with complex matrices remain problematic [24].

Table 1. Coating procedures and sorbent materials used [24].

**Table 1.** Coating procedures and sorbent materials used [24].
Dipping and physical agglutinating methods	Sol-gel technology	Chemical grafting	Electrochemical methods	Electro-spinning	Liquid-phase deposition	Hydro-thermal methods
Carbon nanomaterials [26,120] Ordered mesoporous materials [121,122] Ionic liquids and polymeric ionic liquids [123,124]	Functionalized or polymer-functionalized carbon nanomaterials [125,126,127] Ionic liquid-mediated SPME coating [128,129] Sol-gel derived polymeric ionic liquid-based SPME coatings [130,131] Sol-gel ordered mesoporous silica SPME coating [132,133] Sol-gel coating on metal wires [134,135] Sol-gel molecularly imprinted polymer coatings [113,136] Aptamer Sol-gel SPME [137]	Nanomaterials [138,139] Immunoaffinity SPME [140] Molecularly imprinted polymers [141,142] Substrate-bonded ionic liquid coatings [143] Substrate-bonded polymeric ionic liquid coatings [144]	Electrodeposition Electropolymerized conductive polymers [88,145,146,147] Electropolymerized conductive polymers nanocomposite [148,149] Conductive polymer-ionic liquid composites [150,151] Metal oxides [152,153] Anodized metal wires Metal oxides [154,155] Electrophoretic deposition Carbon nanotubes (CNTs) [112,113,114,115]	Electrospun epoxide polymer Carbon nanofiber-based SPME [116,156,157]	SiO₂, TiO₂, SnO₂, ZrO2 [117] Three dimensional transition metal oxides (V, Cr, Mn, Fe, Co, Ni, Cu, Zn, In, individually or combined) [158]	ZnO nanoparticles Organic frameworks (MOFs) [118,119]

3.5. Quantitation

Contrary to traditional sample preparation methods such as liquid-liquid extraction, solid-phase extraction or Soxleth, SPME is a non-exhaustive technique where only a fraction of the target analyte is extracted from the sample [22]. Consequently SPME must be calibrated carefully for quantitative analysis.

Quantitation of analytes in complex food and environmental matrices is currently one of the major challenges of SPME procedures. Nevertheless, several international standards applying the SPME methodology have recently been implemented, proving that SPME has been accepted as a state-of-the-art analytical technique at least in some fields of application (e.g., environmental). Firstly, the International Standard ISO 27108 [159] “Water quality–Determination of selected plant treatment agents and biocide products–Method using solid-phase microextraction (SPME) followed by gas chromatography-mass spectrometry (GC-MS)” has been established. The described procedure is based on the German Standard DIN 38407-34 [160]. Another standard based on DIN 38407-41 [161] has been published as a Draft International Standard in 2014: “Water quality – Determination of volatile organic compounds in water–Method using headspace solid-phase microextraction (HS-SPME) followed by gas chromatography-mass spectrometry (GC-MS) (ISO/DIS 17943) [162]. In addition, two other standards applying SPME methods have been published: the Austrian OENORM A 1117 [163] (Determination of volatile compounds in cellulose based materials by SPME) and the American ASTM D 7363a [164] (Standard test method for determination of parent and alkyl polycyclic aromatics in sediment pore water using solid-phase microextraction and gas chromatography/mass spectrometry in selected ion monitoring mode).

Essential to developing quantitative SPME methods is the knowledge of fundamental principles governing the mass transfer of analytes in multiphase systems, such as thermodynamics and mass-transfer kinetics [5,43,44]. The current available SPME calibration methods can be categorized into traditional calibration methods including external standard, internal standard and standard addition as well as newer methods such as equilibrium extraction, exhaustive extraction and diffusion-based calibration (see Table 2). Ouyang and Pawliszyn [18,22] published a review giving more detailed information about the mentioned individual methods.

3.5.1. Traditional Calibration Methods

Traditional calibration methods include external standards (calibration curve), internal standard or standard addition method.

SPME calibration can be performed either by equilibrium or pre-equilibrium calibration, respectively. In equilibrium calibration a partitioning equilibrium between the sample matrix and extraction phase is reached. The convection conditions do not affect the amount of extracted analytes due to the present partitioning equilibrium [22]. In pre-equilibrium calibration the amount of extracted analytes is related to the extraction time [22]. The convection/agitation thereby is constant. Applying equilibrium calibration for quantitation purpose is more sensitive and does not depend on the time. However, if the equilibrium extraction is too long, the pre-equilibrium extraction is preferred [22].

The internal standard method is widely applied for quantitation in SPME methods. It has been applied for the quantitative analysis of environmental [165,166] and food [167,168] samples. For heterogeneous samples the standard addition calibration method is preferred [169,170]. Using an internal or a surrogate standard (preferably mass-labeled internal standards) compensates for matrix effects and instrumental or sample preparation variability [171,172,173,174,175]. In addition, external standards are widely used [22,176,177,178,179,180].

Table 2. SPME calibration methods [22].

**Table 2.** SPME calibration methods [22].
Calibration method		Advantages	Disadvantages	Applications
Traditional	External standard Standard addition Internal standard	No extensive sample preparation Correction of sample matrix effects Compensation of matrix effects and losses of analytes during sample preparation and irreproducibility in parameters (injection in GC/LC)	Need for availability of blank sample matrices Need for stable sampling procedure and chromatographic conditions Extensive sample preparation and analysis Limited availability of suitable internal standards High cost and limited availability of isotope-labelled standards	[176,177,178] [169,170] [165,166,167,168,171,172]
Equilibrium extraction		Possibility to calculate concentration of analytes by amount of extracted analytes Independence of amount of extracted analytes of sample volume	Need for knowledge about distribution coefficients	[181,182]
Exhaustive extraction		Possibility to calculate concentration of analytes by amound of extracted analytes and sample volume	Suitable only for small sample volumes and large distribution coefficients or need for special devices	[56,183]
Diffusion-based	Fick’s first law of diffusion Interface model and cross-flow mode Kinetic calibration with standard Standard-free kinetic calibration	Suitability for TWA sampling Independency of sampling rate of face velocity Minimizing of competitive effects for solid coating through high sampling rate and short time Suitability for on-site sampling Suitability for TWA sampling No need for standard loading Possibility to calculate concentrations of all extracted analytes in sample	Sorbent should be zero sink for target analytes Very low sample rate for water sampling Need for controlled or determined flow velocity of sampling matrix Application limited to linear sampling regime Need for determination of standard loading Need for stable sampling conditions Unsuitability for long-term monitoring	[184,185] [186,187] [188,189] [190]

3.5.2. Equilibrium Extraction

The equilibrium extraction method is a widely applied method for quantitation [181,182]. A small part of the extraction phase is exposed to the sample until equilibrium is reached. The concentration of the extracted analyte in the extraction phase is not dependent on the sample volume but corresponds directly to its concentration in the sample [22]. Therefore in practice, there is no need to collect a defined sample prior to analysis. The fiber can be exposed directly to the sample. The analytical process can be accelerated by removing the sampling step. Errors related to analyte losses through the decomposition or adsorption on the sampling container walls will be prevented. In that case the concentration of the target analytes can be determined. However, the distribution coefficients of the analytes between the fiber coating and the sample matrix have to be determined by experimentations. The concentration then can be determined by the amount of the analytes on the fiber under extraction equilibrium.

This calibration method was successfully applied for on-site air [181] or water sampling [182]. For air sampling the extraction can be performed in static and dynamic model [22]. In dynamic model, extraction rates from air-samples can be increased significantly with an air pump [181].

3.5.3. Exhaustive Extraction

Exhaustive extraction calibration can be done by using an internally cooled fiber device [52]. The higher analyte recoveries achieved using internally cooled fiber extraction result in greater confidence in quantitation and reporting of analyte concentrations (see Section 2.3, Figure 2). Almost quantitative extraction of the analytes was reported [22].

3.5.4. Diffusion-based Calibration

The diffusion coefficient is essential for description of kinetic processes of SPME [22]. Recently various diffusion-based calibration methods have been developed [22]. They were developed based on Fick’s first law of diffusion, the interface model, the cross-flow model and the kinetic process of absorption/adsorption and desorption [22]. Main applications are in on-site sampling, comprising grab sampling and long-term monitoring [22].

Fick’s first law is applicable to calibration when diffusion paths are well-defined [22]. The main applications are air and water sampling. Fiber-retracted SPME devices are used in which the analyte molecules access the fiber coating only by means of diffusion through the static air/water gap between the needle opening and the fiber coating. In air, the diffusion of the molecules is fast and the length of the diffusion path can be adjusted [191]. These kinds of samplers were applied for time-weighted average (TWA) sampling of several analytes in air [56,184]. For water sampling, a commercially SPME fiber assembly was developed and used for monitoring polycyclic aromatic hydrocarbons (PAHs) in water [185].

In the case of a poorly defined or unknown diffusion path for on-site sampling (when the SPME fiber is inserted directly into the air or water sample , the interface model and cross-flow model can be used for calibration. However, these models are limited to the linear sampling regime and a constant convection of air/water [22]. Applying these calibration methods requires a controlled velocity of air/water [22]. The Interface model was developed by Koziel et al. [186] and applied for sampling of volatile organic compounds (VOC) in air. The cross-flow model was developed by Chen et al. [187].

Kinetic calibration is based on a diffusion-controlled mass transfer model proposed by Ai [43,44]. The underlying assumption of this dynamic model is that there is a linearly proportional relationship between the adsorbed analyte and its initial concentration in the sample matrix. Based on this model two calibration methods of SPME, the kinetic calibration with standard or in-fiber standardization technique [192] and the standard-free kinetic calibration [190] were proposed. The applicability of this technique for TWA water sampling was established by both theoretical derivations and field trials [188,189]. Furthermore this method has become more convenient and applicable through the single point calibration [193]. Kinetic calibration with standards in the extraction phase can be applied to grab sampling as well as long-term monitoring [22]. Due to the loss of the standard during sampling this calibration method may not work in some fast sampling situations. For fast on-site and in vivo analysis, a standard-free kinetic calibration method was developed [190]. With this calibration method, all analyes can be directly calibrated with only two samplings and can be quantified without bearing in mind of reaching equilibrium in the system. The method was validated for identification of PAHs and BTEX in a standard aqueous solution and a standard gas flow-through system [190]. Compared with the previous calibration methods for rapid on-site analysis by SPME this method does not require a standard to calibrate the extraction. The total amount of the extracted analytes can be quantified without bearing in mind of reaching equilibrium in the system [190].

4. Applications in Food Analysis

The research of aroma-active compounds in foods was developed simultaneously with the introduction of the first gas chromatographs in the early 1960s [194]. In the headspace of foodstuffs the occurrence of around 10,000 compounds was estimated [195]. When considering the choice of suitable methods for food analysis, it is essential to realize that food is a complex, heterogeneous mixture composed of a multitude of different compounds. The analysis of volatile target analytes requires the extraction from the food matrix. It is nearly impossible to analyse food samples without sample pretreatment even though advanced techniques of separation and identification have been made available. Therefore, sample extraction, removal of interferences and the pre-concentration of analytes are mandatory procedures [196].

Developing reliable and sensitive SPME methods for food analysis requires the selection and optimization of parameters like the fiber coating, sampling technique, agitation conditions, sample volume and extraction and desorption conditions [197]. Simultaneous variation of several control variables is possible by applying multivariate test designs reducing the experimental effort for SPME optimization [198]. Multivariate test designs such as full factorial design, Doehlert matrix, 2⁴ experimental design and Design of Experiments are valid multivariate test designs for SPME method optimization [55,56,199].

SPME has been widely applied to the sampling and analysis of food matrices. Table 3 provides a comprehensive overview of SPME applications divided into different analyte classes; aroma, off-flavor (food products), off-Flavor (food packaging migrants) and volatile toxic compounds. Figure 8 shows the numbers of papers dealing with applications of SPME to the different types of analytes based on a ScienceDirect literature research for years 2005 to 2015. Hundreds of papers reporting SPME applications in the field of food analysis have been published in recent years (see Table 3). The vast majority of which referred to aroma analysis (see Figure 8). The applications of SPME to different food matrices are shown in Figure 9. The majority of applications refer to the food classes fruits/vegetables, fats/oils, wine, meat, dairy and (non-) alcoholic-beverages. The results are comparable to those published by Jeleń et al. [13] derived from a Web of Knowledge search in food aroma analysis between 2006 and 2011.

The microextraction technology in greatest use second to SPME (91%) was SBSE (5%), followed by SDME and LLME. The advantage of SBSE is a particularly high sensitivity for semi volatiles. However, SBSE suffered for a long time due to only one available coating (PDMS). Due to the lack of automated versions of SDME and LLME, and also due to their lower popularity in the scientific community (compared to SPME and SBSE) their use in aroma analysis is yet not enough explored. The diversity of matrices for which microextraction methods other than SPME are used proves that these methods await broader applications in aroma analysis [13].

Figure 8. Application of SPME to different types of analytes. Number of papers based on ScienceDirect search for years 2005–2015.

Figure 9. Application of SPME to different food matrices. Number of papers based on ScienceDirect search for years 2005–2015.

Table 3. SPME applications in food analysis.

**Table 3.** SPME applications in food analysis.
		Extraction conditions						Desorption conditions
Food Sample	Analyte	Technique	Fiber	Time [min]	Temp. [°C]	Salt addition	Deriv.	Time [min]	Temp. [°C]	Detection	LOD	Recovery	Ref.
Aroma
Dairy products
Butter	diacetyl	-	PDMS-DVB	5	37	-	-	1	250	GC-MS-MS	0.0078 ppm	-	[200]
European PDO hard cheeses	VVC	HS-SPME	DVB-CAR-PDMS	60	45	-	-	10	260	GC-MS / GC-FID	-	-	[201]
PDO Cheese, Oscypek	VVC	HS-SPME	CAR-PDMS	15	50	-	-	5	260	GC-MS	-	-	[202]
Van Herby Cheeses	esters, ketones, alde-hydes, acids, alcohols, hydrocarbons, terpenes	HS-SPME	DVB-CAR-PDMS	30	40	-	-	5	250	GC-MS / GC-FID	-	-	[203]
Milk, cheese and whey powder	VVC	HS-SPME	DVB-CAR-PDMS	30	40	4 g NaCl	-	5	250	GC-MS	-	-	[204]
Meat and meat products
Roasted pork of mini-pig	VVC	DI-SPME	CAR-PDMS	60	80	0.22 g salt	-	4	280	GC-MS / GO-O / GC-FID	-	-	[205]
Traditional smoke-cured bacon (CSCB)	alkane, aldehydes, ketones, alcohols, thioethers, thiols, furans, phenols	-	CAR-PDMS / DVB-CAR-PDMS	30	60	-	-	5	280	GC-MS	-	-	[206]
Minced beef	VVC	HS-SPME	DVB-CAR-PDMS	30	40	25% NaCl	-	1	250	GC-MS	-	-	[207]
Cooked, cured pork ham	VVC	HS-SPME / SBSE	CAR-PDB-DVB / 0.5mm PDMS phase thickness stir bars	30 / 90	40 / RT	-	-	10	250 / 30	GC-MS	-	-	[208]
Slow fermented sausages	dimethyl trisulfide, 3-methyl thiophene, 2,3-butanedione, 2-nonanoneacetic acid	HS-SPME	CAR-PDMS	180	37	-	-	5	240	GC-MS / GC-O	-	-	[209]
Cooked, fermented sausage	VVC	SPME	CAR-PDMS	40	47	-	-	10	250	GC-MS	-	-	[210]
Cooked beef	VVC	HS-SPME	DVB–CAR–PDMS	25	40	6% NaCl	-	3	250	GC-MS	-	-	[211]
Juice and alcoholic beverages
Whisky	fatty acid ethyl esters, higher alcohols, fatty acids, carbonyl com-pounds, monoterpenols, C13 norisoprenoids, volatile phenols	HS-SPME	CAR-PDMS	60	40	30% NaCl	-	6	220	GC-MS	-	-	[212]
Banana Terra spirit	3-methylbutan-1-ol, 3-methylbutan-1-ol acetate, 2-phenylethylacetate, phenylethyl alcohol	HS-SPME	PDMS-CAR-DVB	25	60	-	-	2	240	GC-MS / GC-O	-	-	[213]
Chinese Laobaigan liquor	VVC	HS-SPME	DVB-CAR-PDMS / CAR-PDMS	40	60	3 g NaCl	-	5	250	GC-MS	-	-	[214]
Ice wine	aroma compounds	HS-SPME	DVB-CAR-PDMS	5	45	1 g NaCl	-	2	260	GC-TOF-MS	trans-OL: 0.015 g/mL; cis-OL: 0.01 g/mL	-	[215,216]
Wine	2-methyl-3-furanthiol, 4-mercapto-4-methyl-2-pentanone, 3-mercaptohexanol, 2-furanmethane-thiol, 3-mercaptohexyl acetate	HS-SPME	PDMS–DVB	10	55	-	on-fiber deri-vatization	2	250	GC-NCI-MS	0.03–0.8 fg	-	[85]
Spanish white wines	VVC	HS-SPME	CAR-PDMS	60	46	-	-	2	280	GC-MS / GC-FID	0.1–900 ng/mL	97–110%	[217]
Cherry wines	VVC	HS-SPME	DVB-CAR-PDMS	45	50	2 g NaCl	-	5	230	GC-MS	0.03–7.27 µg/L	60.7–125.6%	[218]
China gingko wine	VVC	HS-SPME	CAR-PDMS	45	50	saturated NaCl solution	-	4	250	GC-MS	-	-	[219]
Black raspberry wines	VVC	HS-SPME	PDMS-DVB	30	60	-	-	5	230	GC-MS	-	-	[220]
Beer	alcohols, esters, organic acids, aldehydes, ketones, terpenes, sulfur com-pounds, amines, phenols	HS-SPME	TMSPMA-OH-TSO prepared by sol-gel technology	30	40	2 g NaCl	-	5	300	GC	0.01–35.2 µg/L	92.8–105.8%	[221]
Yellow passion fruit juice	VVC	HS-SPME	PDMS-DVB	20	50	17% NaCl	-	6	250	GC-MS	-	-	[222]
Grape juice	aroma compounds	HS-SBSE	PDMS	120	RT	-	-	5	-50	GC-MS	-	28.4%	[223]
Orange beverage emulsion	VVC	HS-SPME	CAR-PDMS	15	45	15% NaCl	-	8	250	GC-MS	0.06–2.27 mg/L	88.3–121.7%	[224]
Coffee	aroma compounds	HS-SPME	PDMS-DVB	20	60	-	-	1	270	GC-MS	-	-	[225]
Coffee	VVC	HS-SPME	PDMS	5	30	-	-	-	220	SAW	-	-	[226]
Coffee	VVC	HS-SPME	DVB-CAR-PDMS	30	60	-	-	5	230	GC	-	-	[227]
Coffee	furans, methoxyphenols, pyrazines, and ketones	HS-SPME	poly [VC₁₆Im][NTf₂] with 50% [VBIm₂C₁₂]2[Ntf₂]	30	RT	-	-	5	175	GC-MS / GC-FID	-	-	[123]
Fruits and vegetables
Various apricot varieties	ethyl acetate, hexyl acetate, limonene, b-cyclocitral, c-decalactone, 6-methyl-5-hepten-2-one, linalool, b-ionone, menthone and (E)-hexen-2-al	HS-SPME	CAR-PDMS	20	40	-	-	4	250	GC-MS / GC-O	-	-	[228]
Apricot varieties	linalool, a-terpineol, b-ionone and c-decalactone	HS-SPME	CAR-PDMS	20	40	saturated NaCl solution	-	4	250	GC-MS	-	-	[229]
Apricot varieties	aldehydes, alcohols, acetates, esters, terpenes and acids	HS-SPME	CAR-PDMS	30	40	-	-	2	250	GC-MS	-	-	[230]
Jackfruit	ethyl isovalerate, 3-methylbutyl acetate, 1-butanol, propyl isovalerate, isobutyl isovalerate, 2-methyl-butanol, butyl isovalerate	HS-SPME	DVB-CAR-PDMS	30	10	-	-	5	250	GC-TOF-MS	-	-	[231]
Cooked peaches	VVC	HS-SPME	DVB-CAR-PDMS	20	40	-	-	10	270	GC-MS	-	-	[232]
Monstera deliciosa fruit	VVC	HS-SPME	PDMS-DVB	60	40	15% NaCl	-	6	250	GC–qMS	-	-	[233]
Pineapple fruit	VVC	HS-SPME	PDMS-DVB	30	40	-	-	2	240	GC×GC-qMS	-	-	[234]
Sweet cherry cultivars	VVC	HS-SPME	CAR-PDMS	20	45	0.2 g NaCl	-	10	260	GC-MS	-	-	[235]
Air-dried raisins	free and glycosidically bound volatile compounds	HS-SPME	CAR-PDMS-DVB	40	60	1.3 g NaCl	-	8	-	GC-MS	-	-	[236]
Table grapes	alcohols, carbonyls, C6 compounds, terpenoids, esters	HS-SPME	PDMS-DVB	30	40	2 g CaCl₂, 20 g NaCl	-	3	220	GC-MS	-	-	[237]
Tomato	VVC	HS-SPME	DVB-CAR-PDMS	15	50	saturated CaCl₂ solution	-	10	250	GC-MS	-	-	[238]
Miscellaneous
Thistle honey	VVC	HS-SPME	DVB-CAR-PDMS	40	60	30% NaCl	-	2	250	GC-MS	-	-	[239]
Croatian lime tree, fir honey-dew, sage honey	VVC	HS-SPME	DVB-CAR-PDMS	20	40	0.5 g an-hydrous Na₂SO₄	-	3	250	GC-MS	-	-	[240]
Honey	VVC	HS-SPME	PDMS-DVB	40	50	-	-	2	250	GC-QTOF-MS	-	-	[241]
Extra virgin olive oils	VVC	HS-SPME	DVB-CAR-PDMS	10	40	-	-	5	260	GC-FID	-	-	[242]
Extra virgin olive oils	VVC	HS-SPME	DVB-CAR-PDMS	360	30	-	-	4	270	GC-MS	-	-	[243]
Virgin olive oil	VVC	HS-SPME	DVB-CAR-PDMS	40	40	-	-	5	300	GC-MS	0.1–2.54 mg/kg	-	[244]
Black and white rice bran	terpenoid flavor odorants	HS-SPME	PDMS	30	100	-	-	0.2	250	GCxGC-MS	-	-	[245]
Italian rice cultivars	VVC	HS-SPME	DVB-CAR-PDMS	60	60	-	-	5	250	GC-MS	-	-	[246]
Palm sugar	N-heterocyclic and O-heterocyclic compounds	HS-SPME	DVB-CAR-PDMS	10	50	-	-	5	240	GC-MS	-	-	[247]
Almond cultivars	VVC	HS-SPME	DVB-CAR-PDMS	60	60	-	-	10	270	GC-MS	-	-	[248]
Saffron	VVC	HS-SPME	PDMS	20	36	-	-	0.4	250	GC-MS	-	-	[249]
Garlic	VVC	HS-SPME	CAR-PDMS	30	30	-	-	3	220	GC-MS	-	-	[250]
Atlantic shellfish species	VVC	HS-SPME	CAR-PDMS	30	80	10 mL saturated NaCl solution	-	10	260	GC-MS	0.12–1.19 ppb	59.3–119.6%	[251]
Off-Flavor
Food products
Beer and beverage	sulfur compounds	HS-SDME / DI-SPME / HS-SDME	PDMS	5	25	2.0 g NaCl / 20% NaCl solution	-	5	250	GC-FPD	0.5 ng/mL for DPrDS -208.1 ng/mL	beer: 85.5–106.9%; bevera-ges: 95.2–110.8%	[252]
Beer	esters and vicinal diketones	HS-SPME	DVB-CAR-PDMS	30	60	3.0 g NaCl	-	-	260	GC-MS	-	-	[253]
Wine	volatile sulfur compouds	HS-SPME	CAR-PDMS	20	35	-	-	7	300	GC-pFPD	0.5 µg/L	0–100%	[254]
Chardonnay and Pinot gris wines	2-aminoacetophenone	DI-SPME	DVB-CAR-PDMS	30	30	-	-	-	250	GC-MS	-	70–80%	[255]
Orange juice	guaiacol and halogenated phenol	HS-SPME	DVB-CAR-PDMS	30	40	-	-	5	220	GC-MS / GC-O	-	-	[256]
Water and apple juice	geosmin	HS-SPME	PDMS synthesized as coated fiber by sol-gel technology	25	40	37% NaCl	-	4	250	GC-MS	1–1.000 ng/L	95–102%	[257]
Coffee beverage	volatile compounds produced by fungi	HS-SPME	DVB-CAR	30	65	-	-	0.7	270	GC-MS	-	-	[258]
Rapeseed oil	hexanal, 2,4-heptadienal, 2-heptenal and 1-pentene-3-ol	HS-SPME	CAR-DVB-PDMS	35	50	-	-	-	150	MS	-	-	[259]
Conventional and high-oleic sunflower oil	hexanal, (E)-2-heptenal, (E)-2-decenal, (E,E)-2,4-nonadienal	HS-SPME	CAR-DVB-PDMS	60 / 90 / 120	40 / 60 / 80	-	-	5	270	MS	0.4–4.3 mg/L	-	[260]
Conventional and high-oleic rapeseed oil	octanal, 3-octanone, propanal, (E,E)-2,4-hexa-dienal, (E)-2-heptenal	HS-SPME	CAR-DVB-PDMS	90	40	-	-	5	270	MS	3.7–816.5 µg/L	-	[261]
Various frying oils	(E,E)-2,4-decadienal, heptanal, (E,E)-2,4-heptadienal, (E)-2-decenal	HS-SPME	CAR-DVB-PDMS	90	40	-	-	5	270	MS	0.03–47.2 µg/L	-	[262]
Almond oils	hexanal, (E)-2-heptenal, (E)-2-octenal, nonanal, (E)-2-nonenal, (E,E)-2,4-nonadienal, (E,E)-2,4-decadienal	HS-SPME	DVB-CAR-PDMS	60	60	-	-	10	270	GC-MS	-	-	[263]
Butter	hexanal	HS-SPME	CAR-PDMS	180	4	-	-	5	250	GC-MS	-	97.37%	[264]
Soymilk	aldehydes, alcohols, ketones, aromatic compounds, esters, furans	HS-SPME	CAR-PDMS	20	40	-	-	3	300	GC-MS	-	-	[265]
Fresh chilled pasteurised milk	microbially induced changes in volatile constituents	HS-SPME	CAR-PDMS	30	40	-	-	2	240	GC-MS / PTR-MS	-	-	[266]
Full fat bovine milk	volatile compounds (pentanal, pentanol, hexanal) produced by photooxidation	HS-SPME	CAR-PDMS	30	50	-	-	0.02	250	GC-MS	-	-	[267]
Chicken breast	sulfides methanethiol, dimethyl disulfide, di-methyl trisulfide, ethanol, 1- and 2-butanol, 1-butanol isomers, free fatty acids	HS-SPME	PDMS	15	50	-	-	3	200	GC-MS-FASST	-	-	[268]
Whiting	trimethylamine, 3-methyl-butanal, 2-methyl-butanal, 3-hydroxy-2-butanone, 3-methyl-1-butanol, 2-methyl-1-butanol	HS-SPME	CAR-PDMS	40	50	saturated NaCl solution	-	0.16	250	GC-MS	-	-	[269]
Rainbow trout	geosmin	HS-SPME	DVB-PDMS	20	65	3.0 g NaCl	-	3	270	GC-MS	-	-	[270]
Potato crisps	VVC	HS-SPME	CAR-PDMS	20	50	-	-	3	300	MS e-nose / GS e-nose	-	-	[271]
Food packaging migrants
Cork	chloroanisoles	CF-HS-SPME	PDMS	10	130 / 10	-	-	3	260	GC-TOF-MS	-	>90%	[56]
Wine	chlorophenols and chloroanisoles	MHS-SPME	DVB-CAR-PDMS	60	70	-	KHCO₃ and acetic acid anhy-dride	5	280	GC-MS-MS	0.004–0.077 ng	-	[80]
Wine	4-ethylphenol, 4-ethylguaiacol	SPME	PDMS-CAR	30	60	-	-	15	220	GC-MS	-	-	[272]
Wine	2,4,6-trichloroanisole, 2,3,4,6-tetrachloroani-sole, pentachloroanisole, 2,4,6-tribromoanisole, 4-ethylphenol (4-EP), 4-ethylguaiacol, 4-vinyl-phenol, 4-vinylguaiacol	MHS-SPME	DVB-CAR-PDMS	60	70	-	-	5	270	GC-MS-MS	4-EP: 1800 g/L; others: 1000 g/L	93.85–101.27%	[273]
Wine	haloanisoles	MHS-SPME	DVB-CAR-PDMS	35	60	99.8% NaCl	-	4	250	GC-ion-trap MS	120.70–150 pg	88.8%	[274]
Water and honey	chlorophenols	DMSPE-HS-SPME	PVC/MWCNTs nanocomposite	15	60	5 mol/L NaCl solution	-	4	215	GC-ECD	0.08–0.6 ng/mL	91–109%	[120]
Volatlile toxic compounds
Contaminants
Milk	diethylstilbestrol	DI-SPME	CNT reinforced hollow fiber	30	60	-	-	10	-	HPLC	5.1 mg/L	57.50%–120.42%	[125]
Milk	enzyme-generated volatile organic com-pounds associated with Listeria monocytogenes	HS-SPME	PA	10	37	-	-	2	230	GC-MS	-	-	[275]
Milk	PAHs	DI-SPME	diethoxydiphenyl-silane prepared by sol-gel technology	60	60	-	-	2	330	GC-MS	0.01–0.08 µg/L	-	[276]
Milk and honey	benzimidazole	DI-SPME	MEMF	70	-	-	-	20	-	HPLC-DAD	0.11–0.30 μg/L	72.3–121%; 83.1–119%	[277]
Chicken muscle and milk	tetracyclines (antibiotic)	SPME	molecularly imprinted polymer	30	-	-	-	10	-	HPLC	1.0 - 2.3 µg/L	-	[141]
Baby formula	furfural and hydroxymethylfurfural	HS-SPME	dodecylbenzene-sulfonate-doped polypyrrole	30	50	2 mol/L NaCl	-	-	200	IMS	6 ng/g; 5 ng/g	95% / 92%	[278]
Baby food and fruit juice	furan	HS-SPME	PEG and PEG/CNTs fibers prepared by sol-gel technology		25 / 30	3 g NaCl	-	0.25	230	GC-FID	0.001 ng/mL; 0.00025 ng/mL	92–98.5%	[126]
Fruit juices	carbamate and phenylurea pesticide residues	DI-SPME	PDMS-DVB and CW-TPR	90	20	0.3 g 30% NaCl	-	15	250	LC/QIT-MS	0.001–0.01 mg/kg	0–82%	[279]
Soft drinks	4-methylimidazole	HS-SPME	PDMS-DVB	50	110	saturated NaCl solution	-	-	270	GC-MS / LC-MS/MS	1.9 µg/L	-	[280]
Brazilian sugarcane juice	pesticideand benzo[a]pyrene	SBSE andMASE	-	180 / 30	280 / 45	saturated NaCl solution	-	11	250	TD-GC-MS / LVI-GC-MS	0.002–0.4 µg/L; 0.004 - 0.56 µg/L	0.2–55.3%; 13.6–103.1%	[281]
Carbonated drink, juice drink, sauce, jam, succade	benzoic and sorbic acids	in-tube SPME	diethylamine-modified poly(GMA-co-EDMA) monolithic capillary	5	-	-	-	7	-	HPLC-UV	1.2; 0.9 ng/mL	84.4–106%	[282]
Water and juice	benzoylurea insecticides	MMF-SPME	MMF/MAED	70	-	-	-	-	-	HPLC-DAD	water: 0.026–0.075 mg/L; juice: 0.053–0.29 mg/L	65.1–118%	[283]
Drinking water	organic micro-pollutants	DI-SPME	PDMS-DVB	30	60	-	-	-	280	GC-MS	0.5–10 µg/L	-	[284]
Apple, apple juice, tomato	organophosphorus pesticides	HS-SPME	PDMS-DVB including B15C5 prepared by sol-gel technology	45	70	5 g NaCl	-	5	270	GC-FPD	0.003–0.09 ng/g	apple juice: 71.5 –01.6% apple: 83.3–97.7% tomato: 55.3–105.3% (spiked 5 ng/g)	[285]
Apples	polycyclic aromatic hydrocarbons, benzene, toluene, ethylbenzene, xylene	HS-SPME	CAR-PDMS	45	60	-	-	5	250	GC-MS	0.02 mg < LOD < 0.26 mg	0.012–0.140 µg	[286]
Fruit and vegetables	pesticide residues	HS-SPME	PDMS	34	62	10% NaCl	-	7	270	GC-MS	0.35–8.33 µg/kg	73–118%	[287]
Packaged fresh vegetables	volatiles derived from Salmonella typhimurium	HS-SPME	CAR-PDMS	15	20	-	-	3	-	GC-MS	-	-	[288]
Vegetables	PAHs	HS-SPME	benzoxy-C[6]/OH-TSO prepared by sol-gel technology	40	60	NaCl	-	10	280	GC-FID	0.04–2.32 ng/g	81.07–107.5%	[289]
Vegetables	organophosphorus pesticide residues	DI-SPME	PA	30	RT	10% NaCl	-	11	260	GC-FPD	0.01–0.14 µg/L	-	[290]
Radish	organochlorine pesticides	HS-SPME	calix[4]arene/hy-droxy-terminated silicone oil prepared by sol-gel technology	30	70	1.0 g K₂SO₄	-	2	270	GC-ECD	1.27–174 ng/kg	83.05–119.3%	[291]
Roasted coffee	furan	HS-SPME	CAR-PDMS	30	35	-	-	-	-	GC-MS	3–10 μg/kg	76–101%	[292]
Wine	2,4,6-trichloroanisole, dibutyl phthalate	SR-SPME	graphene and graphene oxide prepared by sol-gel technology	20	45	20% NaCl	-	5	250	GC-MS	0.3 ng/L	96.96% / 98.20%	[293]
Still and fortified wines	fungicides captan, chlorthalonil, folpet, iprodione, procymidone and vinclozolin, acaricide dicofol	DI-SPME	PDMS	60	35	-	-	3	250	GC-MS/MS	-	70–120%	[294]
Breaded fish products	furanic compounds	HS-SPME	CAR-PDMS	40	37 ± 1	3 g NaCl	-	10	280	GC-MS	-	-	[295]
Fresh, deep frozen, canned, boiled, roasted fish	formaldehyde	HS-SPME	CAR-PDMS	30	80	-	PFBHA	3	310	GC-MS	17 µg/kg	94.8 ± 1.7%	[81]
Cooked, peeled tropical shrimps	3-methyl-1-butanal, 2,3-butanedione, 2-methyl-1-butanal, 2,3-heptanedione and trimethylamine induced by isolated bacteria	HS-SPME	CAR-PDMS	25	40	-	-	5	280	GC-MS	-	-	[296]
Packaged, aged, fresh beef	VOCs associated with Salmonella	HS-SPME	CAR-PDMS	30	23 ± 2 °C	-	-	10	270	GC-MS	-	-	[297]
Smoked meat products	PAHs	DI-SPME	PDMS	60	25	-	-	30	250	GC-MS	0.008–0.138 ng/mL	-	[298]
Meat roasted in plastic bags	plasticisers (phthalates)	DI-CF-SPME	PA	30	45	-	-	0.02	250	GC-MS	0.01–0.18 µg/kg	-	[134]
Fruit leathers	carbonyl compounds generated from ozone-based food colorants decomposition	HS-SPME	PDMS-DVB	15	60	-	PFBHA methanol	-	250	GC-MS	0.016–0.030 μg/L	-	[86]

CF: cold fiber; CNT: carbon nanotube; CSCB: Chinese traditional smoke-cured bacon; DAD: diode array detector; deriv.: derivatization; DI: direct immersion; DPrDS: dipropyl disulphide; ECD: electron capture detector; EDMA: ethylene dimethacrylate; FASST: Fast Automated Scan/SIM Type; FID: flame ionization detector; FPD: flame photometric detector; GC-O: gas chromatography-olfactometry; GMA: glycidyl methacrylate; GS: gas sensor; IMS: ion mobility spectrometry; LOD: limit of detection; LVI: large volume injection; MAED: methacrylic acid-co-ethylene dimethacrylate; MASE: membrane assisted solvent extraction; MEMF: methacrylic acid-co-ethylene dimethacrylate monolithic fiber; MHS: multiple headspace; MMF: multiple monolithic fiber; MW: multi-walled; NCI: negative chemical ionization; OH-TSO: hydroxyl-terminated silicone oil; OL: oak lactone; PAHs: polycyclic aromatic hydrocarbons; PDO: protected designation of origin; PEG: polyethylene glycol; PFBHA: pentafluorobenzyl-hydroxylamine hydrochloride; pFPD: pulsed flame photometric detector; PTR: Proton-transfer-reaction; PVC: polyvinyl chloride; QIT: quadrupole ion trap; qMS: quadrupole mass spectrometry; QTOF: quadrupole time-of-flight; ref.: reference; RT: room temperature; SAW: surface acoustic wave; SIM: selected ion monitoring; SR: space-resolved; TD: thermal desorption; temp.: temperature; TMSPMA: 3-(trimethoxysylil)propyl methacrylate; TOF: time-of-flight; TPR: template resin; VVC: various volatile compounds.

4.1. Aroma

The verification of taste and fragrance attributed to specific foods has become an important part of consumer demand. For this purpose SPME methods can serve as a valuable tool to examine the presence or absence of desired aroma compounds in food products [65]. SPME has gained increasing popularity for aroma analysis in recent years (see Figure 8). A collection of recently published studies in the field of applications of SPME in food, non-food, and fragrance analysis, etc. are listed in the Handbook of Solid Phase Microextraction [75].

Various food components like proteins, lipids, and sugars affect the chemical composition found in the headspace above food items. Page and Lacroix (1993) reported that the presence of a large quantity of lipid material in food products led to a lower extraction of volatiles from the headspace [299]. To prevent this decrease it is recommended to carefully raise the extraction temperature that will increase the number of volatile constituents present in the headspace. Thereby the number of volatile constituents present in the headspace increase. Raw meat for example had no strong aroma [300], but cooking increased meat aroma [300]. Xie et al. (2008) studied compounds that have a real impact on meat aroma. Volatiles of the roasted Mini-pig pork were determined by both SPME and SDE combined with GC-MS. Additionally GC-O was applied to identify potent contributors to the special meat aroma. A total of 86 different volatile compounds have been identified, whereas only 45 odor active regions could be recognized by olfactometry. Aldehydes from lipid oxidation followed by spice components were among the highest contributors to the volatile chemical profile [205].

Another study on meat aroma was performed with Chinese traditional smoke-cured bacon (CSCB) by Yu et al. [206]. The aim of the study was to work out the substances responsible for the characteristic aroma of CSCB. In total, 48 volatile compounds (alkanes, aldehydes, ketones, alcohols, thiolethers and thiols, furans and phenol compounds) were identified and quantified using SPME coupled to GC-MS [206]. Chemical processes like smoking, oxidation and the Maillard reaction (reaction between reducing sugars and amino acids, usually under the influence of heat) are largely responsible for the occurrence of these compounds.

Another approach in aroma analysis is to verify the origin and authenticity of foods. For example, Majcher et al. [202] developed a SPME method in combination with chemometrics for the determination of volatile compounds derived from traditional Polish cheese Oscypek and its imitations [202]. Sostaric et al. (2000) used SPME-GC-MS to differentiate between natural vanilla extracts and nature-identical and synthetic vanilla flavorings [301].

The extraction of aromatic volatile compounds from alcoholic and non-alcoholic beverages by SPME was recently extensively investigated (see table 3). Câmara et al. [212] investigated aroma compounds in whisky samples. Classical analytical methods such as LLE, simultaneous extraction or distillation show relatively low reproducibility or possible contamination with solvents. Further the lengh of time required for the analysis and the selectivity of the results are such a few drawbacks, which can be overcome selectivity can be overcome by dynamic headspace SPME and gas chromatography coupled to GC-MS. In this study the use of five different SPME fibers PDMS, PA, CAR-PDMS, CW-DVB and CAR-PDMS-DVB were compared. Optimized results were achieved by saturating the samples with salt solutions and using a CAR-PDMS fiber. More than seventy compounds including ethylesters, long-chain alcohols, isoamyl acetates and fatty acids were identified [212].

Ong et al. [231] investigated volatile compounds derived from jackfruit applying SPME and gas chromatography-time-of-flight mass spectrometry (GCTOF-MS). In five jackfruit cultivars thirty-seven compounds were identified. Compounds responsible for the sweet and fruity aroma in jackfruit were identified as ethyl isovalerate, 3-methylbutyl acetate, 1-butanol, propyl isovalerate, isobutyl isovalerate, 2-methylbutanol, and butyl isovalerate [231].

Fruit origin and authenticity (species idenfication) can also be successfully investigated by means of SPME analysis of volatile compounds [229]. For example, the typical sensory expectations of apricot fruits are sweetness and juiciness, which are strongly related to the ripening stage apricots at harvest [302]. Aroma compounds of apricots from different regions of the world were extensively studied [303]. Major identified aroma compounds were ethyl acetate, hexyl acetate, limonene, 6-methyl-5-hepten-2-one, menthone, E-hexen-2-al, linalool, beta-ionone and cyclo-decalactone [228].

Recently the European Union established the research program Horizon 2020 to develop a validated analytical method for determination of volatiles resulting from virgin olive oils by means of SPME-GC/MS. Romero et al. [244] published a validated method including calibration curves for 29 volatile compounds. The analytical precision of 67% of these compounds had a relative standard deviation lower than 10% and accuracies were determined for 97% of the analyzed volatile compounds. The limits of detection ranged from 0.1 to 2.54 mg/kg for determined volatile compounds [244].

4.2. Off-Flavors

As described in Section 4.1, aroma belongs to the most important sensory characteristics of food. Food products with an inappropriate aroma impression are often rejected by consumers and may lead to a loss of consumer confidence [304]. The awareness of off-flavors in food relies mainly on their concentration occurring in the food matrix and odor threshold values. Off-flavors are defined as unpleasant odors or flavors transmitted to food through internal impairing changes [304]. Many volatile compounds are associated with unpleasant odor notes such, e.g., putrid, musty, rotten, skunk, etc. However, the received off-flavor impression often depends on the concentration of the related aroma compounds. The most common off-flavor compounds are those that cause musty, earthy off-flavors (haloanisoles, halophenols, geosmin and methylisoborneol) and medicinal off-flavors (phenolic compounds, sulfur-containing compounds and carbonyl compounds) [304]. Other off-flavor compounds are generally classified according to their origin, including microbially derived off-flavors, compounds originating from packaging materials, from cleaning agents, and compounds resulting from Maillard reactions [304].

The impact of the cork composition on the sensory properties of wines was investigated by means of SPME GC-MS [272]. Cork was shown to have an active role in the sorption of volatile phenols from wine. In particular, the sorption properties of 4-ethylphenol and 4-ethylguaiacol phenols in hydro-alcoholic medium were investigated by Gallardo-Chacón and Karbowiak [272]. Through the high sorption activity of corks the concentration of 4-ethylphenol and 4-ethylguaiacol was decreased. High concentration of these compounds can lead to the formation of off-flavor. Both, in models and real wine samples the tested cork leads to a significant reduction of the mentioned off-flavor compounds in wine [272].

The oxidative stability of conventional high-oleic sunflower oil and analytical and sensory lipid oxidation parameters in conventional and high-oleic rapeseed oil were reported by Petersen et al. [260,261]. It was concluded that the combination of volatile compound analysis with HS-SPME-GC and multivariate statistical methods provides a sensitive tool in differentiating conventional sunflower oil and high-oleic sunflower oil as by means of volatile lipid oxidation marker compounds. The same applies to differentiating rapeseed oil and high-oleic rapeseed oil.

4.3. Volatile Toxic Compounds

Various sources of microbiological and chemical hazardous compounds may occur in foods. The latter may include natural toxicants, such as mycotoxins [305,306,307], marine toxins [308], and environmental contaminants, such as mercury and lead [309,310,311,312,313,314].

Among these substances for example volatile toxic aldehydes like formaldehyde has recently received increased attention. Formaldehyde is formed post-mortem by the enzymatic reduction of trimethylamine-N-oxide to formaldehyde and dimethylamine [315,316]. Due to its high volatility, formaldehyde is captured well by SPME for subsequent measurement. Firstly, in 2005 Bianchi et al. [317] developed a SPME method with in situ derivatisation with pentafluorobenzyl-hydroxylamine hydrochloride (PFBHA). In 2007, Bianchi et al. [81] used a SPME-GC-MS method based on the same derivatization to evaluate the formaldehyde content of various fish products.

Additional non-flavor contaminants are polycyclic aromatic hydrocarbons (PAHs), identified as pollutants in the environment and food items [1,2,3]. Bianchi et al. [276] developed a SPME method with a coating based on the use of diethoxydiphenylsilane produced by sol-gel technology for determination of PAHs at trace levels in milk. For vegetables, a simple, sensitive and affordable method has been developed for the quantitation of eight PAHs by Lei et al. [289]. The method was based on HS-SPME connected with GC-FID. The sol-gel designed benzoxy-C[6]/OH-TSO proved to have a good capability to capture aromatic compounds such as phthalate acid esters. Compared to the tested PDMS fiber and C[4]/OH-TSO fiber, the benzoxy-C[6]/OH-TSO showed to have the highest affinity to PAHs [289].

The VOCs 2-nitrophenol and 3-fluoroaniline were extracted, separated and detected by headspace-solid phase microextraction coupled to gas chromatography–mass spectrometry (HS-SPME-GC-MS). It was stated to be a potential rapid method for future development [275].

Microorganisms can also be responsible for food contamination and spoilage. The metabolic activity of microorganisms through breakdown of compounds in food leads to the release of volatile organic compounds [41]. Extensive research was carried out in order to identify microbial activities in foods using SPME [275,288,297,318]. Siripatrawan and Harte [288] investigated the occurrence of Salmonella typhimurium present in packaged alfalfa sprouts. For that purpose CAR/PDMS fibers were compared to PDMS/DVB fibers which extracted lesser sulfur volatiles. Sulfur-containing compounds seem to be a major marker compound for spoiled food stuffs, off-flavor in milk, and the presence of pathogens [288]. Other volatile organic compounds such as 2-nitrophenol and 3-fluoroaniline may be associated with the presence of Listeria monocytogenes contamination in food [318].

Pesticides are not only contaminants, but are also non-flavor compounds occurring in food items. The simultaneous determination of fourteen multiclass pesticide residues in fruit and vegetable samples by means of HS-SPME coupled to GC-MS were reported by Abdulra’uf and Tan [287]. The method development was based on multivariate experimental designs (Plancket-Burman and central composite design) conducted in two stages. Internal standard calibration was applied for quantitation (detection limits between 0.35 and 8.33 µg/kg). Compared to liquid-liquid extraction techniques and SPE, which are commonly used in pesticide analysis, SPME takes less time, is less labor-intensive, does not require any solvents and simplifies sample preparation to one single step and one device [41].

5. Environmental Applications

SPME has been widely applied to the sampling and analysis of environmental matrices including air, water, soil, and sediment samples, in on-site or off-site analysis [20,16,18]. Hundreds of papers reporting SPME applications in the field of environmental analysis have been published in recent years (see Table 4), the vast majority of which referred to aqueous samples (see Figure 10).

5.1. Air Samples

Applications of SPME in air sampling can be performed on-site [319,320] or in the laboratory after collecting air samples in suitable containers, e.g., bags [102,321], glass bulbs [181,322], or pre-evacuated vessels [323]. The analytes are extracted either by direct exposure of the fiber to the air sample or by headspace method after absorption in a suitable liquid medium [324,325] or adsorption on a solid material, e.g., Tenax TA [326]. Some authors use dynamic flux chambers for air sampling [133,327,328,329]. Most applications published on SPME analysis of gaseous samples involve the use of commercially available SPME fibers, although novel fiber coatings have already been developed for this purpose as well. For example, a sol-gel single-walled carbon nanotube/silica composite coated SPME has been developed for the analysis of organohalogen compounds in workplace air [133].

Figure 10. Applications of SPME to different environmental matrices. Number of papers based on ScienceDirect search for years 2005–2015.

Table 4. SPME applications in environmental analysis.

**Table 4.** SPME applications in environmental analysis.
		Extraction conditions						Desorption conditions
Analyte	Environmental Sample	Technique	Fiber	Time [min]	Temp. [°C]	(Salt) Addition	Deriv.	Time [min]	Temp. [°C]	Detection	LOD	Recovery [%]	Ref.
Air
Pollutants
Volatile organohalogen compounds	workplace air	SPME	sol-gel SWCNT / silica	15	RT	-	-	3	280	GC-MS	0.09–0.2 ng/mL	-	[127]
Formaldehyde and other carbonyl compounds	indoor air	SPME	PDMS-DVB	15	RT	-	PFBHA	4	250	GC-MS	0.002–0.032 µg/m³	-	[82]
Acetaldehyde, acetone, BTX, pinene, trichloroethylene, alkanes	indoor air (classrooms)	SPME	PDMS-CAR	240	RT	-	-	2.5	320	GC-MS	0.05–5.9 µg/m³	79–120	[181]
BTEX	indoor and outdoor air	SPME	PDMS-CAR	30	14–24	-	-	2	260	GC-MS	0.05–0.1 µg/m³ (benzene)	-	[319]
PAH	ambient air particulate matter (PM10, TPS), sampling on quarz filter disks and extraction with water/methylene chloride/acetone	DI-CF-SPME	PDMS	60	70	-	-	1	270	GC-MS	0.02–1.16 ng	88–98	[330]
Alkyl- and methoxy-phenolic compounds	biomass smoke, absorption in aqueous NaOH prior to SPME	HS-SPME	CW-DVB	90	-	35% NaCl, pH 2	-	3	250	GC-MS	1.13–4.60 ng/mL	-	[324]
Organophosphate triesters	indoor air (lecture room, office)	dynamic SPME	PDMS	40–90 min / >18 h; air flow rate 10–35 cm/s	22	-	-	2	280	GC-NPD	-	-	[327]
Volatile organic compounds
VOC	indoor air	SPME	PDMS-CAR	10	RT	-	-	10	300	GC-MS / GC-FID	-	122 ± 24	[331]
VOC	indoor air	SPME	PDMS-CAR	1–45	-	-	-	2.5	320	GC-MS	-	-	[332]
VOC	air from volcanic and geothermal areas, landfill gas	SPME	PDMS-DVB-CAR	30	20	-	-	2	230	GC-MS	-	78–84	[323]
Odorous compounds
VOC and odorous compounds	air from swine barn, cattle feedlots	SPME	PDMS-CAR	60	RT	-	-	5	260	GC-MS-O	-	-	[320]
Volatile carbon, sulfur and nitrogen compounds	air from livestock operations	dynamic SPME	PDMS-DVB-CAR	5 min / air flow rate 16 cm/s	20	-	-	-	300	GC-MS	-	-	[328]
VOC and odorous compounds	swine barn particulate matter (TPS, PM10, PM2.5, PM1), adsorption on TEOM filters prior to SPME	HS-SPME	PDMS-CAR	180	25	-	-	40	260	GC-MS-O	-	-	[333]
Odorous compounds	waste gas from fat refinery	SPME	PDMS-CAR	30	RT	-	-	5	290	GC-MS / GC-FID-O	-	-	[102]
Odorous compounds	gaseous effluents from production of poultry feather and viscera meal, condensed prior to SPME	HS-SPME	PDMS-DVB-CAR	20	50	-	-	4	250	GC-MS	-	-	[325]
Volatile organic sulfur compounds	air from different areas of sewage treatment plant	SPME	PDMS-CAR	45	22 /RT	-	-	2	200	GC-MS	0.01–0.08 µg/m³	75–96	[322]
Synthetic musks	indoor air, adsorption on Tenax TA prior to SPME	HS-SPME	DVB-CAR-PDMS	20	100	100 µL acetone	-	5	270	GC-MS	0.029–0.380 ng/m³	85–103	[326]
Trimethylamine	ambient air	SPME	PDMS-DVB	10	22 / RT	-	-	3	210	GC-FID			[321]
Diacetyl	air	SPME	PDMS	2	RT	-	-	1	250	GC-MS	0.05 ppm	-	[200]
Monoterpenes	plant emissions, ambient air	SPME	PDMS-DVB	20	25 / RT	-	-	5	250	GC-MS	4–20 ppt	-	[329]
Water
Odorous compounds
Earthy-musty odorants	source, product, and tap water from different waterworks	HS-SPME	PDMS-DVB-CAR	30	60	25% NaCl	-	5	230	GC-MS	0.1–1.3 ng/L	83–112	[334]
Earthy-musty odorants	tap water, river water, lake water	HS-SPME	PDMS-DVB-CAR	30	90	-	-	-	-	GC-MS	0.25–0.61 ng/L	65–92	[335]
Earthy-musty odorants	tap water, lake water	HS-SPME	PDMS-DVB-CAR	30	50	30% NaCl	-	3	265	GC-MS	0.32–0.66 ng/L	86–113	[336]
Odorous trichlorobromo- phenols	tap water, river water	HS-SPME	PDMS-DVB-CAR	90	60	NaCl 6.5 g/30 mL	di-methyl sulfate / NaOH	1	270	GC-MS	0.22–0.95 ng/L	-	[337]
Algal taste and odor compounds	lake water	HS-SPME	PDMS-DVB-CAR	30	65	NaCl	-	3	250	GC-MS-O	sub to low ppt range	80–115	[87]
Volatile sulfur compounds	odorous freshwater lakes	HS-SPME	PDMS-CAR	30	45	-	-	3	250	GC-FPD	1.6–93.5 ng/L	87–112	[338]
Nitro musk fragrances	tap water, wastewater	HS-SPME	PDMS-CAR, PDMS-DVB	25	100	-	-	2	300 / 270	GC-μECD	0.25–3.6 ng/L	96–108	[339]
Volatile and semivolatile organic compounds
VOC	surface water, wastewater from wastewater treatment plant and from municipal solid-waste treatment plant	HS-SPME	PDMS-CAR	30	50	10% NaCl	-	5	280	GC-MS-MS	0.005–2 µg/L	70–120	[340]
Volatile and semivolatile organic compounds	landfill leachate	SBSE	PDMS	60–120	RT	-	none, acetate or BSTFA	30 °C to 280 °C at 60 °C/min, 5 min hold at 280 °C	GC-MS	-	-	[341]
Volatile and semivolatile organic compounds	snow	Di-SPME/HS-SPME	PDMS-DVB	40 / 180	RT	-	-	5	250	GC-MS	0.11–1.93 µg/L	-	[342]
Pesticides
Pyrethroid pesticides	tap water, waste water from sewage treatment plant, run-off water, water from container used for washing oranges	DI-SPME	PDMS	20	50	0.02% Na₂S₂O₃, 2% acetone	-	3	280	GC-MS	0.9–35 pg/mL	-	[343]
Pyrethroid pesticides	groundwater	DI-SPME	PDMS/DVB	30	65	-	-	5	-	LC-PIF-FD	0.003–0.009 µg/L	92–109	[344]
Benzimidazole fungicides	seawater, ground water, sewage	DI-SPME	PDMS-CAR	40	60	15% NaCl	-	10 (in metha-nol)	-	HPLC-FD	0.03–1.30 ng/mL	81–120	[345]
Organochlorine pesticides	lake water	HS-SPME	PMPVS	30	80	-	-	2	270	GC-ECD	0.84–13.0 ng/L	72–116	[346]
Organochlorine pesticides	groundwater	DI-SPME	PDMS-DVB-CAR	45	RT	-	-	2	260	GC-ECD	0.0013–0.45 ng/L	92–105	[347]
Organochlorine pesticides	seawater	HS-SPME	poly(3,4-ethylene dioxythio-phene)	15	70	-	-	2	270	GC-µECD	0.16–0.84 ng/L	63–127	[146]
Organochlorine pesticides and metabolites	seawater, sewage, groundwater	DI-SPME	PDMS-DVB, CW/TPR	40	50 /45	-	-	8 / 10	-	HPLC-UV-DAD	0.3–3.6 ng/mL	90–104	[348]
Pesticides tebuthiuron and diuron	water	DI-SPME	PA	50	-	-	-	15	-	LC-DAD	10–50 µg/L	-	[349]
Pesticides	surface and groundwater	DI-SPME	PA	30	50	-	-	5	280	GC-MS	0.02–0.3 ng/mL	78–109	[350]
Organo-phosphorus pesticides	river water	DI-SPME	sol-gel based amino fiber	40	30	20% NaCl	-	4	250	GC-MS	0.05–1.0 ng/L	80–115	[351]
Organochlorine (OCP), organo-phosphorus (OPP), triazine, pyrethroid, and miscellaneous pesticides	groundwater	HS-SPME	PDMS-DVB	60	60	-	-	-	-	GC-ECD- TSD / GC-MS / GC-MS-MS	<0.1 µg/L	-	[352]
PAH and related compounds
PAH	tap water, surface water, underground water, rainwater	HS-SPME	PDMS	30	60	10% NaCl	-	2	280	GC-FID	0.06–0.5 μg/L	71–109	[353]
PAH	rainwater, strom water	DI-SPME	PDMS	60	65	0.5 M sodium mono-chloro-acetate	-	-	-	GC-MS	0.001–0.041 µg/L	72–111	[354]
PAH	wastewater from scrubber of pilot-scale fluidized bed incinerator system	MA-HS-SPME	PDMS-DVB	30	20	-	-	5	290	GC-FID	0.3–1.0 µg/L	88–103	[355]
PAH	surface waters, leaching waters of contaminated soils	HS-SPME	PA	60	50	-	-	2	250	GC-FID	0.08–0.20 µg/L	-	[356]
Hydroxy metabolites of PAH	mini pore water, minimal salts medium, soil extract culture medium	DI-SPME	PA	40	40	8% NaCl	BSTFA	3	280	GC-MS	0.002–0.134 µg/L	-	[357]
Phenols
Volatile phenols	well water, drinking water	HS-SPME	PANI	60	80	30% NaCl, pH 2	-	5	275	GC-FID	1.3–12.8 ng/mL	88–103	[88]
Phenols	real-life water samples	HS-SPME	PANI	50	50	NaCl, pH 2	-	5	200	GC-FID	0.69–3.7 ng/mL	69–112	[147]
Phenols	river water, waste water	DI-SPME	oxidized MWCNTs	30	20	36% NaCl	-	3 (in ACN / water 70:30)	RT	HPLC-DAD	0.25–3.67 ng/L	86–119	[138]
Phenols and nitrophenols	rainwater	SPME	PA	40	RT	NaCl, pH 3.0	MDBSTFA	5	280	GC-MS	0.2–99 μg/L	-	[358]
Chlorophenols	landfill leachate	IL-HS-SPME	1-butyl-3-methylimida-zolium hexafluoro-phosphate	4	25	pH 2	-	4	240	GC-MS	0.008–0.026 µg/L	87–99	[124]
Chlorophenols	landfill leachate	purge-assisted HS-SPME	PA	30	75	-	-	3	130 °C (0.05 min) to 300 °C at 8 °C/s, 4 min hold	GC-MS	0.1–0.4 pg/mL	83–114	[359]
Miscellaneous
Organometallic compounds (mercury, lead, tin)	river water, seawater	HS-SPME	PDMS-DVB-CAR	15	40	-	sodium tetra-ethyl-borate	2	260	GC-MS-MS	4–33 ng/L	50–109	[360]
Organo-phosphorus fire retardants and plasticizers	wastewater, MAE prior to SPME	DI-SPME	PDMS	30	65	10% NaCl	-	0.5	250	GC-ICP-MS / GC-TOF-MS	29–50 ng/L	38–43	[83]
Non-halogenated solvents	textile wastewater	HS-SPME	PDMS	10	35	-	-	5	240	GC-MS	0.1–300 µg/L	-	[361]
Acetone	seawater	SPME	PDMS-DVB	30	RT	NaCl, pH 3.7	PFBHA	5	250	GC-MS	3.0 nM	-	[362]
BTEX	waste water	HS-SPME	PDMS-DVB	1	RT	35% NaCl	-	0.16	200	portable GC-µFID	0.4–1.4 µg/L	98–111	[89]
BTEX	groundwater	HS-SPME	PDMS-CAR	15	25	267 g/L NaCl	-	2	290	cryo-trap-GC-MS	0.01–0.05 ng/L	-	[363]
Acrolein	surface water, drinking water	HS-SPME	PDMS-CAR	50	60	saturated NaCl solution	2,2,2-trifluo-roethyl-hydra-zine	-	220	GC-MS	0.06 µg/L	91–104	[364]
Pharmaceutical compounds	wastewater	dSPME	CW-TPR	30	75	300 g/L NaCl, pH 3/11	-	10	-	LC-MS-MS	LOQ: 0.005–0.05 µg/L	89–110	[90]
Endocrine disrupting chemicals and steroid hormones	river water	DI-SPME	PA	90	45	10 g/L NaCl 10, pH 5	BSTFA	5	290	GC-MS	0.002–0.378 µg/L	-	[365]
Bisphenol A	landfill leachate	HS-SPME	PA	60	25	100 g/L NaCl, pH 2	BSTFA	5	280	GC-MS	0.03 µg/L	-	[91]
Estrogens	surface water, wastewater	IT-SPME	DVB	20 cycles à 40 µL (100 µL/min)	-	-	-	-	-	LC-MS-MS	2.7–11.7 pg/mL	86–107	[92]
Organic pollutants (pesticides, octyl/nonyl phenols, pentachloro-benzene, PAHs)	wastewater, landfill leachate	DI-SPME	CW-DVB	45	RT	NaCl 10%	-	5	250	GC-TOF-MS	-	-	[366]
Organochlorine pesticides and polychlorinated bisphenyls	river water	HS-SPME	PDMS	60	80	-	-	5	250	GC-MS-MS	0.4–26 pg/L	75–105	[367]
Volatile halogenated hydrocarbons, benzene	groundwater from waste-oil recycling facility	HS-SPDE	PDMS-AC	15 cycles à 1 mL (50 µL/s)	60	5% NaCl	-	1 mL (10 µL/s)	300	GC-MS	12–870 ng/L	-	[368]
BTEX and halocarbons	contaminated water	HS-SPME	PAC	20	40–45	25% NaCl	-	1	260	GC-FID-ECD	LOQ: 0.01–0.94 µg/L	88–113	[369]
Chlorobenzene	tap water, river water	HS-SPME	nanofiber coated by electrospun PU	5	30	25% NaCl	-	2	200	GC-MS	10 ng/L	94–102	[370]
1,2-cis-dichloroethylene, trichloroethylene	effluent from soil column experiments	HS-SPME	PDMS	5	25	-	-	1	250	GC-FID	2.4–4.2 µg/L	-	[371]
1,3-dichloro-2-propanol	tap water, river water, paper mill sewage	HS-SPME	PDMS-DVB-CAR	30	25	NaCl	BSTFA	5	260	GC-MS-MS	0.4 ng/mL	93–103	[318]
Epichlorohydrin	water and sewage	HS-SPME	PDMS-CAR	15	50	-	-	1	240	GC-MS, GC-ECD	water: 1.0 ng/L	-	[93]
Polybrominated diphenyl ethers	river water, waste water	DI-SPME	MWCNT	30	RT	-	-	2	295	GC-ECD	3.6–8.6 ng/L	90–119	[372]
Soil / Sediment and other Solid Matrices
Odorous compounds
VOC and odorous compounds	dairy manure	HS-SPME	PDMS-DVB-CAR	30	30	-	-	-	230	MD-GC-MS-O	-	-	[373]
Odorous volatile compounds	compost	HS-SPME	PDMS-CAR	60	20	-	-	40	250	GC-MS	0.06–2.38 ppb	-	[374]
Volatile and semivolatile organic compounds
VOC	soil, manure, compost, biochar	HS-SPME	PDMS-DVB-CAR	20	20	-	-	5	230	GC-MS	0.01–310 ng/g	-	[375]
VOC	cow slurry	HS-SPME	PDMS-DVB-CAR / PDMS-CAR	15	35	NaCl	-	3	300	GC-MS	0.02–1441 µg/L	-	[376]
Volatile and semivolatile organic compounds	urban landfill soil	HS-SPME	PDMS-DVB	30	50	NaCl	-	-	250	GC-MS	-	-	[377]
Pesticides
Pyrethroids, organochlorine pesticides	agricultural soils	HS-SPME	PA	30	100	-	-	5	290	GC-µECD	0.004–1.2 ng/g	71–147	[378]
Pyrethroids	sediment pore water	SPME	PDMS	20	-	-	-	3	260	GC-ECD	-	-	[379]
PAH and related compounds
PAH	sediment pore water, sediments	SPME	PDMS	500 h	18	1 mL 10 mM NaN₃	-	15		50 °C to 250 °C at 200 °C/min	-	-	[380]
PAH	coastal sediments, MAE with acetone prior to SPME	DI-SPME	PDMS	60	60	-	-	-	270	GC-MS	0.07–0.76 µg/kg	70–110	[381]
PAH	sand, sediment	CF-HS-SPME	PDMS	40	150 (fiber: 5)	-	-	2	300	GC-FID	0.3–3 pg/g	-	[54]
Parent and alkyl PAH	sediment pore water	HS-SPME	PDMS	30	-	-	-	5	320	GC-MS	0.002–0.6 ng/mL	-	[382]
Miscellaneous
4-t-octylphenol, nonylphenol, bisphenol A	activated sludge	nd-SPME	PA	-	20	-	-	5	2800	GC-MS	-	-	[383]
Phenols and indoles	cow slurry	HS-SPME	PDMS-DVB-CAR	15	35	NaCl	MTBSTFA	9	300	GC-MS	0.004–707 µg/L	>64	[94]
Polybrominated diphenyl ethers	soil	HS-SPME	sol-gel M-β-CD-OH-TSO	60	95	methanol	-	12	300	GC-MS	13.0–78.3 pg/g	78–99	[384]
Perfluoro-carboxylic acids	harbour sediments, PFE prior to SPME	HS-SPME	PDMS	30	30	saturated NaCl solution	boron trifluo-ride	3	300	GC-MS	0.5–0.8 ng/g	99–103	[95]
Organotin compounds	sediment	HS-SPME	PDMS	30	80	ethanol, pH 5.3	sodium tetra-ethyl-borate	1	250	GC-MS	1.0–6.3 µg/kg	98–117	[84]
Mono-, di- and tri-butyltin	sediment, extraction with hydrochloric acid/ethanol prior to SPME	HS-SPME	PDMS	30	40	pH 4	sodium tetra-ethyl-borate	2	250	GC-MS-MS	0.03–1.0 pg/g	-	[96]
Cyclopentadienyl-manganese tricarbonyl, (methylcyclo-penta-di-enyl)manganese tricarbonyl	seawater, soil	HS-SPME	PDMS-DVB	20	60	5% NaCl	-	0,25	250	GC-MIP-AED	0.62–0.65 pg Mn/g	76–113	[385]
Nitrous oxide	estuarine soils and sediments	HS-SPME	PDMS-CAR	2	50	-	-	1	200	GC-MS	18 ppb	-	386]

ACN: acetonitrile; CF: cold fiber; BSTFA: N,O-bis(trimethylsily)trifluoroacetamide; BTEX: benzene, toluene, ethyl benzene, ortho-xylene and meta- and para-xylene; BTX: benzene, toluene and xylene; DAD: diode array detector; deriv.: derivatization; DI: direct immersion; dSPME: dual SPME; ECD: electron capture detector; FID: flame ionization detector; FPD: flame photometric detector; IL: ionic liquid; IT: in-tube; FD: fluorescence detection; LOD: limit of detection; LOQ: limit of quantification; MA: microwave assisted; MAE: microwave assisted extraction; M-β-CD/OH-TSO: permethylated-β-cyclodextrin/hydroxyl-termination silicone oil; MD: multidimensional; MDBSTFA: N-(t-butyldimethylsilyl)-N-methyltrifluoroacetamide; MIP-AED: microwave-induced plasma atomic emission detection; MS-O: mass spectrometry-olfactometry; MTBSTFA: N-methyl-N-t-butyldimethylsilyltrifluoroacetamide; MW: multi-walled; nd-SPME: negligible depletion-SPME; NPD: nitrogen–phosphorus detector; PAC: powered activated carbon; PAHs: polycyclic aromatic hydrocarbons; PANI: polyaniline; PDMS-AC: PDMS mixed with activated C; PFBHA: pentafluorobenzyl-hydroxylamine hydrochloride; PFE: pressurized fluid extraction; PIF: postcolumn photochemically induced fluorimetry derivatization; PMPVS: polymethylphenylvinylsiloxane; PU: polyurethane; ref.: reference; RT: room temperature; SW: single-walled; temp.: temperature; TOF: time-of-flight; TPR: template resin; TPS: total suspended particulate; TSD: thermoionic specific detection.

Typical fields of applications are the analysis of VOC, odorous compounds, or micro-pollutants in indoor air, atmospheric air, or gaseous effluents from industrial or agricultural operations. For example, Bourdin and Desauziers [82] developed a simple and fast method based on on-fiber derivatization using BSTFA for the analysis of formaldehyde and other carbonyl compounds in indoor air. For a 5 min fiber exposure time, the limits of detection were below 0.5 μg/m³, the average reproducibility was 15%, and the linearity of the calibration curves was satisfactory. Of the micro-pollutants, formaldehyde is of particular concern as it was recognized as “carcinogenic to humans” (class 1) by the International Agency for Research on Cancer. It is found in nearly every indoor atmosphere because of its numerous sources, such as furniture, human activity (tobacco), glues, and varnishes [82]. Several authors used SPME as a screening method for the characterization of odorous compounds in gaseous effluents, e.g., livestock operations [320] or production of poultry feather and viscera meal [325] in order to develop suitable abatement strategies. SPME can also be used for analysis of airborne particulate matter after collection on filters [330,333].

5.2. Aqueous Samples

Aqueous sampling by SPME can be performed by direct immersion, the HS method, or the IT SPME method. Most of the papers reported HS-SPME applications to aqueous samples and subsequent GC separation combined with different detectors (see Table 4). In recent years DI-SPME followed by LC has gained popularity [21]. Derivatization techniques have been used for example for analysis of PAH and related compounds [357], phenols [337,358], organometallic compounds [360], endocrine disruptors and steroid hormones [91,365], acetone [362], and acrolein [364] in aqueous samples. In addition to commercially available SPME fibers, fibers with new coating materials have been used. They include poly(3,4-ethylene dioxythiophene) [146] and a sol-gel based amino fiber [351] for analysis of pesticides. Polyaniline [88,147], oxidized multiwalled carbon nanotubes [138], and 1-butyl-3-methylimidazolium hexafluorophosphate [124] have been applied to the analysis of phenols. Furthermore, a nanofiber coated by electrospun PU [370] was utilized for analysis of chlorobenzene.

SPME has been used for the analysis of a wide variety of water pollutants including pesticides, PAHs, and phenols. Several publications on SPME analysis of many classes of pesticides in groundwater, lake, river and seawater, and waste water are available [146,343,344,345,346,347]. Results showed good performance of the methods with high recovery rates >70% and very low LOD, often in the range of ng/L. For example, Raposo et al. [347] applied a GC/ECD method using DI-SPME for the analysis of organochlorine pesticides in groundwater with recovery rates between 92 and 105% and LOD between 0.0013 and 0.45 ng/L [347]. Furthermore, SPME has been used to determine PAH, a large group of molecules widely distributed in the environment, and their metabolites in tap, rain, and surface waters as well as scrubber water with high recoveries (>71%) [353,354,355,356,357]. SPME has also been successfully applied to other classes of pollutants in environmental water samples, e.g., phenols [88,124,138,147,358,359], pharmaceuticals [90], endocrine disrupting chemicals and hormones [97,98,365] and organometallic compounds of mercury, lead and tin [360].

Other fields of SPME application are the analysis of volatile and semivolatile organic compounds in waste water and landfill leachate [340] and the analysis of odorous compounds like earthy-musty odorants [334,335,336] and volatile sulfur compounds [338] in tap, river, or lake water.

5.3. Solid Samples

Sampling of soil and sediments as well as other solid materials like manure, activated sludge, and biochar by SPME is usually performed by HS methods. In some applications extraction of analytes has been assisted by heating of the sample [84,377,378,384,385,386] or cooling the fiber [54], use of microwaves [381]; use of pressurized fluid extraction [95], or addition of small amounts of organic solvents prior to SPME [84,384]. Most applications published on SPME analysis of solid environmental samples involve the use of commercially available SPME fibers. A novel sol-gel permethylated-β-cyclodextrin/hydroxyl-termination silicone oil coated fiber has been used to extract polybrominated diphenyl ethers from soil [384]. With this coating, improved fiber stability has been achieved at analytical characteristics similar to commercially available fibers (RSD < 10%, LOD 13.0–78.3 pg/g, recovery > 78%, r > 0.999).

The main fields of SPME applications to solid environmental samples are analysis of pesticides [378,379], PAH and related compounds [380,381,382], phenols [94,383], perfluorocarboxylic acids [95], and organotin compounds [84] in contaminated soils and sediments (see Table 4). Since SPME is generally organic solvent free, rapid, and sensitive, it allows the detection of trace amounts of substances that are short-lived during biotransformation, thus, providing insight into degradation pathways. For example, Shen et al. [383] developed a negligible depletion-SPME (nd-SPME) method to measure free concentrations of 4-t-octylphenol, nonylphenol and bisphenol A in order to determine aerobic biodegradation kinetics in activated sludge [383]. In nd-SPME, the extracted amount of analytes is so small that the depletion (i.e., change of concentrations) of the analytes does not lead to a change in reaction kinetics or other relevant conditions in the sample. Further applications of SPME to solid environmental samples are characterization of VOC and odorous compounds in soil, manure, compost, and biochar samples [373,374,375,376,377].

6. Prospects and Trends

Recent trends in sample preparation techniques comprise miniaturization, automation, high-throughput performance, on-line coupling with analytical instruments as well as low-cost operation through extremely low or no solvent consumption. Among other techniques mentioned in this review, SPME has the potential to meet these challenges [50]. Through miniaturization of sample preparation steps time and operation costs can be saved. The main advantage however is the possibility to determine trace and ultratrace analytes in complex matrices.

In addition to SPME techniques, further selective extraction procedures have been developed. They all show clear trends toward simplification of sampling and sample preparation methods, an increase in reliability and precision, and the elimination of the cleanup step. The development of more sensitive and selective phases may be a step towards further miniaturization of these techniques. Increasing interest also lies in the field of automating sample preparation. Automation can result in faster procedures as well as improved precision and cost-effectiveness. Over the last decade, sample preparation devices have been automated and coupled to separation and detection systems specifically designed for automation.

Due to the lack of automation, SPME tends to be coupled more often to GC than to LC. Moreover, more recent techniques such as SBSE are not yet as widely accepted as SPME due to the difficulty of achieving full automation. Another challenge related to new SPME techniques that has to be overcome is the carry-over effect occurring for example in SPDE, in-tip SPME and MEPS techniques due to the remaining of analytes on the inner surface of the needle after heat desorption [50,63,64,387]. Another aspect that should be considered for optimization is the moderate enrichment capacity of these techniques [50].

Research regarding SPME is not only based on the development of new devices but also on the study of derivatization strategies, new coatings with higher extraction efficiencies, selectivity and stability, and the study of novel calibration processes [2]. During the SPME process there are many different parameters that can be applied to achieve specific methods. Derivatization for example is a valuable tool in many cases to achieve efficient extraction. With regard to chemical consumption and waste generation, a recent trend however is to avoid derivatization techniques where possible [98].

The variety of commercially available fiber coatings is constantly growing which open up the range for further applications. However, in addition to these commercially available sorbents, further promising coating procedures have been developed. This approach opens up new fields of application. Furthermore, efforts are made to overcome limitations in biocompatibility, on-site compatibility, selectivity and sensitivity of currently available SPME coatings [103,104]. Besides physical coating processes which are compatible with nearly all kinds of sorbent materials there is a clear trend towards procedures developed for specific components. Most attention in past years has been given to the application of nanomaterials [115,370]. Nanostructured MIP-based coatings will continue to be of particular interest. Great attention is currently given to such specific SPME coatings as a basis for development of more designable structures. Even when issues like selectivity for target analytes and applicability for samples with complex matrix compositions have to be solved [24].

Quantitation is currently the major challenge for SPME procedures to gain wider application. This review showed that calibration techniques are still under development [22]. The model calculation methods laid down in chapter 2.5 provide a meaningful approach to overcome these challenges. A better understanding of SPME processes will be the basis of further improvements. Nonetheless, a number of norms have already been implemented and indicate that the application of calibration methods in SPME is possible. In the field of aroma analysis of food, the main quantification barrier is the high complexity of volatile compounds [388,389].

Food safety and quality control is mainly responsible for the development of further rapid and automated procedures aimed to raise the sample throughput and minimize sources of error [3]. Since 2006 there has been a strong and still valid trend towards SPME application for the analysis of wine, fruit/vegetables, dairy and meat products and other foods. In this field, SPME seems to have a leading role followed by SBSE and liquid microextraction methods such as SDME and LLME [13].

In the field of environmental analysis, further improvement of rapid techniques suitable for on-site, on-line and in situ measurements of pollutants and their transformation products will be an important future challenge. These methods will provide insight into natural environmental processes, such as natural attenuation and uptake by plants as well as toxic effects and toxicity mechanisms of emerging chemicals.

Microextraction techniques represent a step towards the miniaturization of the analytical laboratory which is one of the major challenges and directions of future method development acitivites. Applications of SPME in different fields apart from environmental and food analysis such as pharmaceutical, process monitoring, clinical, forensic or drugs are continuously increasing [2].

7. Conclusions

Solid-phase microextraction (SPME) techniques take a leading position among microextraction methods due to their simplicity and the possibility of automation. SPME is a very useful technique for analyzing volatiles occurring in very low concentrations in various foods and environmental applications occurring in very low concentrations. A reliable tool for the identification and quantitation widely applied in the analysis of chemical compounds and biological substances needs a high-quality sample preparation. In this context, SPME may provide very good solutions. The development of new rapid SPME techniques might further increase its credibility. However, the challenges of this method in handling complex sample matrices must be taken into consideration. Additionally, limitations regarding quantitation of target analytes need to be reduced. The selection of an appropriate SPME fiber requires knowledge of the sample matrix properties. Nowadays, there is a large variety of different fiber coatings, extraction and desorption techqniues as well as derivatization procedures available, enabling the development of selective, sensitive and repeatable SPME methods for food and environmental analysis.

Author Contributions

Sybille Merkle reviewed the food applications and recent SPME developments together with Dr. Kim Karen Kleeberg who wrote to the environmental applications under the supervision of Prof. Jan Fritsche.

Conflicts of Interest

The authors declare no conflict of interest.

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Merkle, S.; Kleeberg, K.K.; Fritsche, J. Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review. Chromatography 2015, 2, 293-381. https://doi.org/10.3390/chromatography2030293

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Merkle S, Kleeberg KK, Fritsche J. Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review. Chromatography. 2015; 2(3):293-381. https://doi.org/10.3390/chromatography2030293

Chicago/Turabian Style

Merkle, Sybille, Kim Karen Kleeberg, and Jan Fritsche. 2015. "Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review" Chromatography 2, no. 3: 293-381. https://doi.org/10.3390/chromatography2030293

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Recent Developments and Applications of Solid Phase Microextraction (SPME) in Food and Environmental Analysis—A Review

Abstract

1. Introduction

2. SPME Techniques

2.1. Fiber Solid-Phase Microextraction

2.2. In-tube Solid-Phase Microextraction

2.3. Cooled Coated Fiber Device

2.4. Non-fiber SPME Techniques

3. SPME Process

3.1. Extraction and Desorption Techniques

3.2. Derivatization

3.3. Salt-addition

3.4. Fiber Coatings

3.5. Quantitation

3.5.1. Traditional Calibration Methods

3.5.2. Equilibrium Extraction

3.5.3. Exhaustive Extraction

3.5.4. Diffusion-based Calibration

4. Applications in Food Analysis

4.1. Aroma

4.2. Off-Flavors

4.3. Volatile Toxic Compounds

5. Environmental Applications

5.1. Air Samples

5.2. Aqueous Samples

5.3. Solid Samples

6. Prospects and Trends

7. Conclusions

Author Contributions

Conflicts of Interest

References

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