Headspace sampling of the volatile fraction of vegetable matrices

Available online at www.sciencedirect.com

Journal of Chromatography A, 1184 (2008) 220–233

Review

Headspace sampling of the volatile fraction of vegetable matrices Carlo Bicchi ∗ , Chiara Cordero, Erica Liberto, Barbara Sgorbini, Patrizia Rubiolo Dipartimento di Scienza e Tecnologia del Farmaco, Via Pietro Giuria 9, I-10125 Torino, Italy Available online 16 June 2007

Abstract The evolution of vapour phase sampling of the volatile fraction of vegetable matrices, or of products directly related to them, over the period 1996–2007 is reviewed. High concentration capacity headspace (HCC-HS) and dynamic headspace (D-HS) techniques, that is headspace sampling approaches where the analytes in the vapour phase are concentrated into a sorbent, an adsorbent or a solvent, are considered. Advantages, disadvantages and applications to the vegetable field of several successful techniques based on these approaches are critically presented, including in-tube sorptive extraction (INCAT, HS-SPDE), headspace sorptive extraction (HSSE), solid-phase aroma concentrate extraction (SPACE), large surface area HCC-HS sampling (MESI, MME, HS-STE), headspace liquid-phase microextraction (HS-LPME) and dynamic headspace samplings (D-HS). The developments necessary to overcome some of the limits of the above approaches and techniques are also discussed in view of their application to new fields. © 2008 Elsevier B.V. All rights reserved. Keywords: Headspace sampling; Static headspace; Dynamic headspace; Static and dynamic high concentration capacity headspace technique; Volatile fraction of vegetable matrices

Contents 1. 2. 3. 4. 5. 6. 7. 8. 9.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Review articles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . In-tube sorptive extraction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Headspace sorptive extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Solid-phase aroma concentrate extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Large surface area high concentration capacity headspace sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Headspace liquid-phase microextraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dynamic headspace . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . General considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

221 222 223 223 224 225 226 227 230

Abbreviations: AEDA, aroma extraction dilution analysis; CLS, closed loop stripping; D-HF-HS-LPME, dynamic hollow fibre-supported HS-LPME; D-HS, dynamic headspace; DP, dual phase; DSA, descriptive sensory analysis; DSE, direct solvent extraction; D-TD-HS, direct thermal desorption headspace; EGPE, equilibrium gum phase extraction; EN, electronic nose; FTMS, Fourier transform mass spectrometry; GC-O, gas chromatography-olfactometry; GPE, gum phase extraction; HCC-HS, high concentration capacity headspace techniques; HD, hydrodistillation; HS-LPME, headspace liquid-phase microextraction; HSME, headspace solvent microextraction; HS-SPDE, headspace solid-phase dynamic extraction; HS-SPME, headspace solid-phase microextraction; HSSE, high capacity headspace sorptive extraction; INCAT, inside needle capillary adsorption trap; IS-SPDE, in solution solid-phase dynamic extraction; LLE, liquid–liquid extraction; MAE, microwaveassisted extraction; MALDI, matrix-assisted laser desorption ionization; MESI, membrane extraction sorbent interface; MHE, multiple headspace extraction; MME, membrane microextraction; MWHD, micro wave hydrodistillation; OSF, organic solvent film; OTT, open tubular trapping; PDMS, polydimethylsiloxane; PEG, polyethyleneglycol; PHWE, pressurized hot water extraction; P&T, purge and trap; SAFE, solvent assisted flavour evaporation; SBSE, stir bar sorptive extraction; SDE, simultaneous distillation solvent extraction; SDEV, simultaneous distillation and extraction under reduced pressure; SDME, single drop microextraction; SE, solvent extraction; SFE, supercritical fluid extraction; S-HS, static headspace; SPACE, solid phase aroma concentrate extraction; SPE, solid phase extraction; STE, sorptive tape extraction; TAS, total analysis system; TCMs, traditional Chinese medicines; VHS, vacuum headspace ∗ Corresponding author at: Dipartimento di Scienza e Tecnologia del Farmaco, Universit` a di Torino, Via Pietro Giuria 9, I-10125 Torino, Italy. Tel.: +39 011 670 7661/7662; fax: +39 011 670 7687. E-mail address: [email protected] (C. Bicchi). 0021-9673/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2007.06.019

10.

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

221

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

231 231 231

1. Introduction In the history of this technique, Ettre [1] reported that the first application mentioning the concept of headspace (HS) sampling was the “aerometric method” for rapid determination of alcohol in water and body fluids due to Harger et al. [2]. He also found that the terms “headspace” and “headspace analysis” were first used in 1960 by Stahl et al. [3], adapting an expression used in the food packaging industry, while the first communication in which HS sampling was combined with a GC analysis was by Bovijn et al. [4] who, in 1958, applied sampling of the “gaseous phase in equilibrium with a liquid phase” to monitoring trace concentrations of hydrogen at the 1-ppb level present in water in high-pressure boilers. In spite of its continuous evolution, it is only over the last two decades that HS sampling has enjoyed a remarkable development; this came about at the same time as the ever-increasing success of solvent-free sample preparation techniques, i.e. techniques in which the analyte(s) is isolated from a matrix without using a liquid solvent. HS sampling meets this definition perfectly, since its main aim is to sample the gaseous or vapour phase in equilibrium (or not) with a solid or liquid matrix, in order to characterise its composition [5]. Traditionally HS sampling operates either in static (S-HS) or in dynamic mode (D-HS): paradoxically, the principles of these two approaches were defined so clearly that they limited the development of HS techniques until the end of the 1980s. The renewed interest in this technique coincided with the introduction, in the early 1990s, of an additional approach, which acts as a bridge between S-HS and D-HS: High concentration capacity headspace techniques (HCC-HS). HCC-HS techniques are based on either the static or the dynamic accumulation of volatile(s) on polymers operating in sorption and/or adsorption modes, or, more seldom, on solvents. Their immediate success was mainly because they are as simple, fast, easy to automate, and reliable as SHS, and, at the same time, show analyte concentration factors comparable to those of D-HS. Several successful techniques based on the HCC-HS approach are nowadays widely used in addition to conventional S-HS and D-HS samplings: HSsolid-phase microextraction (HS-SPME) [6], in-tube sorptive extraction (INCAT, HS-SPDE) [7,8], headspace sorptive extraction (HSSE) [9,10], solid-phase aroma concentrate extraction (SPACE) [11], headspace liquid-phase microextraction (HSLPME) [12,13], and large surface area HCC-HS sampling (MESI, MME, HS-STE) [14–16]. The first HCC-HS technique to appear was HS-SPME introduced by Zhang and Pawliszyn in 1993 as an extension of SPME [6], which was developed by Arthur and Pawliszyn [17] to overcome some drawbacks of SPE in sampling organic pollutants from water. Zhang and Pawliszyn also advanced a theory for SPME applied to HS sampling [6,18] and showed that analyte recovery from headspace by a fibre depends on two closely-related but distinct equilibria:

the first is the matrix/headspace equilibrium responsible for the headspace composition (measured by its distribution coefficient K2 ), the second is the headspace/polymeric fibre coating equilibrium (measured by its distribution coefficient K1 ). Pawliszyn and his co-workers reviewed the theory, technology, evolution and applications of SPME together with some additional specific topics [19–21]. The HS-SPME approach (and the underlying theory) has been and still is the basis for the development of new HCC-HS sampling techniques, some of them introduced with the aim of overcoming some of its limitations. A further important factor that contributed greatly to the development of HCC-HS techniques is the ever-increasing knowledge of sorption (in particular with polydimethylsiloxane (PDMS)). Sorption is a partition based on the analyte dissolution in a liquid retaining polymer and, together with adsorption, is the main phenomenon involved in the recovery of an analyte from a liquid or vapour sample by a polymeric phase [22–24]. Baltussen et al. [25] first applied the sorption approach to D-HS, to preconcentrate volatiles and semivolatiles from air and gaseous samples using glass tubes packed with apolar PDMS particles. The sampled analytes were recovered by thermal desorption and analysed by GC and GC–MS. PDMS cartridge immediately gave an excellent performance, including for polar analytes, and even compared to that of adsorbents such as Tenax TA or Carbo-trap 300. The HCC-HS approach has made a dramatic contribution in extending the availability of solutions to meet most requirements for vapour phase sampling, and successfully fills the “gap” between conventional S-HS and D-HS. HS-sampling in its different approaches has played and still plays a fundamental role in studying the composition of the volatile fraction of a plant, in both research and quality-control fields. It is of interest here to mention the basic concepts involved in the volatile fraction of a plant. The volatile fraction emitted from a plant plays a fundamental role in the plant’s life, because it is an important biosensor diagnostic of the changes that take place in its metabolism [26]. It generally consists of a mixture of compounds that can be sampled, as a consequence of their capability to vaporise either spontaneously or when suitable sampling conditions or techniques are applied. Its analysis involves a range of approaches and/or techniques, which produce samples that, while they may be of different compositions, are representative of the volatiles characterising the vegetable matrix or of products deriving from it, e.g. headspace, essential oils, flavours, fragrances, aromas or extracts prepared through specific techniques. For these studies, it is therefore necessary to develop analytical methods and technologies suitable both to detect variations in the composition of the emitted volatile fraction and to monitor the dynamics of the reaction(s) of a vegetable organism when stressed. An ideal set up to analyse a plant’s volatile fraction consists of a sampling technique in which recoveryover-time of biologically-interesting metabolites is maximised,

222

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

and an analytical technique where analysis time is minimised. These considerations make fundamental the role that HS sampling plays as a tool for sample preparation in this field clear. Not infrequently, HS and essential oil are confused: although their compositions may sometimes be similar, the definitions clearly distinguish them, since HS involves sampling from the vapour phase in equilibrium with a (plant) matrix, while an essential oil is the “odorous product, usually of complex composition, obtained from a botanically defined plant raw material by steam distillation, dry distillation, or a suitable mechanical process without heating” [27]. This article is an overview of the most recent HCC-HS and DHS sampling techniques, in particular applied to the analysis of plant matrices used in the food, medicinal and cosmetic fields. Principle, evolution, advantages and limits of each technique, together with a survey of its applications in the plant field, are reported in a dedicated section. Since the HS-SPME applications that appeared during 2000–2005 [28] in the same fields were specifically reviewed in 2006 by the same authors, they will be left out of this article. However, in consideration of the fundamental contribution that HS-SPME and its theory have given, and are still giving, to the development of HCC-HS sampling techniques, HS-SPME will be taken as reference standard to evaluate advantages and disadvantages of new techniques and/or on how they overcome some of its limitations. After a survey on review articles on the various topics involved in HS sampling, analyte concentrating HS sampling techniques are presented with their applications to the plant field, following their chronological appearance; in-tube sorptive extraction, headspace sorptive extraction, solid-phase aroma concentrate extraction, large surface area HCC-HS-sampling, headspace liquid-phase microextraction and dynamic headspace sampling are discussed in detail. The review is chiefly based on articles quoted by Sci-Finder Chemical Abstract Data Base [29] and published over the last decade (1996–2007). A list of the full name, abbreviations and/or acronyms of the techniques reviewed is reported in abbreviations. The language of articles not available in English is mentioned in the table reference. 2. Review articles Over the last ten years, HS sampling in both S-HS and DHS modes has been the object of several reviews concerning techniques, general and specific approaches, theory, and applications dealing with topics characterising the flavour and fragrance fields and products deriving from plant matrices. Several subjects of general interest for HS-sampling have been reviewed. Vitenberg [30] discussed the equilibrium model in gas extraction for the static, dynamic, and flow versions of headspace analysis, together with their efficiency. Hinshaw [31] critically reviewed thermal desorption methods and their ability to make compatible large gas volumes with capillary columns, including on-column cryotrapping, sample stream splitting, multiple adsorbent sample tubes, and secondary adsorbent traps. Nongonierma et al. [32] dealt with the mechanisms of extraction of aroma compounds from foods, using adsorbents. In particular, they discussed the recent acquisition on the

two main stages of an adsorption HS process, i.e. the kinetic stage, involving the diffusion of analytes within the adsorbent pores, and the thermodynamic stage, described by adsorption isotherms. Several parameters were critically evaluated, including: (a) physico-chemical characteristics of the adsorbent such as porosity and hydrophobia; (b) physico-chemical characteristics of the aroma compounds, including sample/adsorbent partition coefficient, and volumetric mass influencing the diffusion in the adsorbent pores; and (c) sampling conditions such as pH, temperature, time, gas or solvent flows, and composition of the sample. Baltussen et al. [24] critically reviewed sorptive sample preparation techniques, in particular covering open tubular trapping (OTT), SPME, gum phase extraction (GPE), equilibrium gum phase extraction (EGPE) and stir bar sorptive extraction (SBSE). The performance of HS techniques has also been compared and discussed. In a survey dealing with modern methods for sample preparation for separation techniques, Smith [33] described several examples of HS analysis and vapour trapping. In a critical evaluation of some of the most recent HCC-HS techniques in the analysis of flavours and fragrances, Bicchi et al. [34] discussed advantages, limits, and fields of application of HS-SPME, HSSE and HS-SPDE. Sun et al. [35] recently reviewed HSLPME, reporting theory and applications of these techniques, in particular from the perspective of their adoption in the tobacco industry. Several reviews have addressed specific subjects of application in the field of plant matrices and products derived from them. Jakobsen [36] reviewed the approaches for isolating plant volatiles and discussed a system developed to control environmental factors during in situ HS analysis. Brunke et al. [37] discussed odour and taste detection methods and HS analysis of blossoms as a source of ideas for research and perfumery. Tholl et al. [26] critically reviewed practical approaches and methods to plant volatile analysis, in particular focussing on the latest technology underlying some new developments; this review provides guidance for the selection of appropriate instrumentation and protocols for on-line and off-line biochemical, physiological and ecological applications to study volatiles emitted by the whole organism, organs or enzymatic preparation and related physiological parameters. Rousseff and Cadwallader [38] reviewed headspace techniques to sample the volatile fraction in foods, fragrances and flavours, and presented an overview of contemporary HS techniques including SPME, purge and trap automation, electronic noses or MS, without separation prior to HS analysis, and applications together with advantages, limitations and alternatives. Song [39] discussed the last development in food flavour analytical techniques, in particular dealing with D-HS, solvent assisted flavour evaporation (SAFE), gas chromatography-olfactometry (GC-O) and aroma extraction dilution analysis (AEDA). Augusto et al. [40] recently reviewed advances and applications in sampling and sample preparation for analysis of aromas and fragrances, including analytical distillation (vapour distillation (HD) and simultaneous distillation extraction (SDE)), HS-manipulation methods (S-HS and D-HS and HS-SPME) and direct extraction methods (such as liquid–liquid, LLE, solid-

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

phase, SPE, and supercritical fluid, SFE). Wilkes et al. [41] critically evaluated sample preparation methods for the analysis of flavours and off-flavours in several sorts of food, including SHS and D-HS, purge and trap (P&T), HS-SPME, direct thermal desorption (D-TD-HS) headspace techniques. Reid [42] dealt with a similar subject, discussing instrumental methods to detect taints and off-flavours by HS, SPME, SBSE and HSSE, as well as electronic noses. Mestres et al. [43] extensively reviewed SHS, and D-HS and HS-SPME methods to study organic sulphur compounds in wine aroma, and their effect on it. 3. In-tube sorptive extraction In-tube sorptive extraction is one of the approaches based on D-HS that was developed to overcome the relatively limited concentration capability of HS-SPME, due to the small volume of sorbent coating a fibre. The first technique based on this approach is known as inside needle capillary adsorption trap (INCAT) and was introduced in 1997 by McComb et al. [7]. INCAT is a pre-concentration device consisting of a hollow needle, with either a short length of GC capillary column placed inside it, or an internal coating of carbon. Sampling may be performed on ambient air, on a solution, or on the S-HS of a liquid sample, by passing the gaseous or liquid sample through the device, either actively with a syringe, or passively via diffusion. The VOCs are sorbed and concentrated onto either the carbon layer or the liquid stationary phase of the capillary column inside the needle. The sampled analytes are recovered by direct thermal desorption into the GC injector. To the best of the authors’ knowledge, INCAT has never been applied to sampling the HS of plant matrices, having chiefly been used for fingerprint analyses of complex mixtures of petroleum-based products [44]. The principle of in-tube sorptive extraction is the foundation of solid-phase dynamic extraction (SPDE), also known as “the magic needle”, introduced by Lipinsky [45]. SPDE is a technique that can be used equally for sampling from liquid (IS-SPDE) or vapour phases (HS-SPDE), where the analytes are concentrated on a thick film (50 ␮m) of a polymer coated onto the inside wall of the stainless steel needle (5.5 or 7.5 cm long) of a gas tight syringe (2.5 mL). In HS-SPDE, analytes are accumulated in the polymer coating the inner needle wall by pulling in and pushing out a fixed volume of HS to be sampled, through the gas tight syringe for an appropriate number of times within a fixed time. The trapped analytes are then thermally desorbed, on-line transferred by a fixed volume of carrier gas into the GC injector body, and analysed by GC or GC–MS. The volume of polymer coated on 5.5 cm SPDE needle wall is about 4.5 ␮L, i.e. about 10 times greater than that coating an SPME fibre (0.4–0.6 ␮L). Several polymeric coatings are available: PDMS, PDMS/activated charcoal, PDMS/OV 225, PDMS/phenyl-methylpolysiloxane, polyethylenglycole (PEG), and polydimethylsiloxane, 7% phenyl, 7% cyanopropyl (OV 1701). HS-SPDE is a D-HS approach because the vapour phase flowing over the accumulating phase layer is continuously renewed. Few HS-SPDE applications have been reported in the literature; they include the analyses of cannabinoids and

223

amphetamines both in hair samples of drug abusers [8,46] and of furan, benzene and toluene from aqueous solutions [47]. Bicchi et al. [48] applied HS-SPDE to the analysis of aromatic plants and food matrices, in particular rosemary, banana, green and roasted coffee and red and white wine, all with a PDMS coated needle. More recently other approaches based on the same principles known as inside needle dynamic extraction (INDEX) [49,50] and in-tube extraction (ITEX) [www.ctc.ch/misc/documents/Itex.pdf] have been introduced. To the best of the authors’ knowledge, these techniques have not yet been reported to the field of vegetable matrices. Table 1 lists the applications of in-tube sorptive extraction techniques to HS analysis of plant matrices; for each article, common and Latin name of the plant investigated is given, together with reference number, the in-tube sorptive extraction approach adopted, a list of the techniques with which it has been compared, and the main component(s) identified or class of compounds investigated. The main advantages of in-tube sorptive extraction are its effectiveness with highly volatile compounds (unlike HSSPME) and the possibility of optimising its concentration capability, depending on the analyte amount in the vapour phase, by selecting a suitable number of pull/push cycles. Its main limit compared to HS-SPME is that it requires dedicated instrumentation, cryoconcentration and tuning of several parameters. 4. Headspace sorptive extraction HSSE was introduced in 2000 by Bicchi et al. [9] and Tienpoint et al. [10] as an extension of stir bar sorptive extraction (SBSE), in its turn introduced by Baltussen in 1999 to increase the concentration capability in comparison to solidphase microextraction (SPME) [51]. In HSSE, the analytes are statically accumulated onto a thick film of PDMS coating a glasscoated magnetic stir bar, whose PDMS volume ranges from 25 to 250 ␮L depending on its size. HS components are recovered by suspending the PDMS stir bar in the vapour phase, in equilibrium or not with the matrix, for a fixed time. After sampling, the stir bar is placed in a glass tube and transferred to a thermo-desorption system where the analytes are thermally recovered and analysed by GC or GC/MS. PDMS stir bars are commercialised under the name ‘Twister’ (Gerstel, M¨ulheim a/d Ruhr, Germany). In a further in-depth study concerning the influence of sampling conditions on HSSE performance, carried out on some volatile compounds characteristic of the essential oil field (iso-butyl-methyl-ketone, 3-hexanol, isoamyl acetate, 1,8cineole, linalool and carvone), Bicchi et al. [52] showed that analyte recovery depends on its physico-chemical characteristics and affinity for PDMS (octanol–water partition coefficient), sampling temperature and time, and HS/PDMS phase ratio. More recently, Bicchi et al. [53] introduced a new generation of twisters to overcome some of the limitations of HSSE, in particular those concerning recovery of polar analytes from complex or multi-ingredient matrices and that of very volatile components (C1–C4 analytes). Dual-phase twisters (DP-twisters) combine the concentration capabilities of two or more sampling materi-

224

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

Table 1 List of the applications of HS-SPDE, HS-SPACE, MESI and HS-STE to the analysis of plant matrices Ref.

Matrices

Headspace solid-phase dynamic extraction (HS-SPDE) [48] Aromatic plants, foods

Main components

Other techniques

Annotations

␤-Pinene, isoamyl acetate, linalool

HS-SPME

SPDE polymeric phase: PDMS

HS-SPME, D-HS, SE

SPACE adsorbent coating: 18% graphite + 2% activated carbon in ethanol

␣-Pinene, eucalyptol, ␥-terpinene ␣-Pinene, eucalyptol, ␥-terpinene

HS-SPME

MESI polymeric phase: PDMS MESI polymeric phase: PDMS

Dihydro-carvyl acetate 1,8-cineole, 5-hydroxy-methyl furfural

HS-SPME, HSSE

Headspace solid-phase aroma concentrate extraction (HS-SPACE) [11] Coffee, standard mixture 2-Octanone, tetradecane, furfurale [67]

Plant matrices

Membrane extraction sorbent interface (MESI) [70] Eucalyptus dunnii leaves [71]

Eucalyptus dunnii leaves

Headspace sorptive tape extraction (HS-STE) [16] Mentha spicata L. (mentha spearmint), Rosmarinus officinalis L. (rosemary), apple

HS-SPME

STE polymeric phase: PDMS

Enclosed data: reference number, the common and Latin name of the plant investigated, the main component(s) identified or class of compounds investigated and a list of the techniques with which they have been compared.

als operating in different modes (e.g. sorption and adsorption) and consist of a short PDMS tube, the ends of which are closed with two magnetic stoppers, thus creating an inner cavity that is packed with different activated carbons as adsorbents. DPtwisters have been shown to be very effective in HS sampling of vegetable matrices in the food, cosmetic and pharmaceutical fields. The possible application of HSSE and SBSE to the analysis of flavour carriers was discussed by Sandra et al. [54] through the results of sorptive extraction analysis on tea, beer, yoghurt, and bananas, and off-flavours and bitter compounds in beer. Eri et al. [55] also evaluated the performance of HSSE by comparing it to those of D-HS combined with GC–O in the analysis of fruit HS and correlated analysis results and sensory data. In a survey on HCC-HS techniques, Bicchi et al. [34] critically discussed advantages, limits, and fields of application of HS-SPME, HSSE and HS-SPDE in the field of flavours and fragrances. The high HSSE concentration capability was immediately evident from the original articles [9,10]. Table 2 reports the list of the applications of HSSE to plant matrices and related products. For each article quoted, the common and Latin name of the vegetable matrix investigated is given, together with the reference number, main component(s) identified or class of compounds investigated, and a list of the techniques with which HSSE has been compared. If not specified otherwise, analysis of the fractions sampled by HSSE was by GC and GC–MS. HSSE has successfully been applied in the fields of medicinal and aromatic plants [9,56,57], fruits [58,59], to discriminate toxigenic from nontoxigenic fungi [60,61], truffle fruiting body versus mycelial aromas [62] and in shiitake mushrooms [63] and to some food samples [10,64–66]. The results reported show not only the high concentration capability of HSSE compared to other techniques, mainly

due the high volume of PDMS, but also that: (a) PDMS twisters can successfully be applied in both S-HS and DHS modes to trace analysis and passive sampling; (b) they can be used to sample headspace with unfavourable β values and/or large headspace volumes; (c) their concentration capability can be varied by using twisters with different PDMS volumes (from 25 to 250 ␮L); (d) the absolute amounts of analyte contained in the headspace sampled by the twister can easily be determined through analyte calibration curves constructed by direct injection through the TDS system of a standard solution of the analyte(s) investigated; (e) sampling and analysis can be carried out separately, thus affording infield or process sampling; (f) the range of analyte polarity can be extended by a suitable combination of trapping phases in DP-twisters. On the other hand, the main limitations on HSSE are the need for dedicated and expensive instrumentation, (i.e. a thermodesorber in combination with a cryoconcentration system) and the lack of “polar” polymer coating for twisters, to improve HSSE’s effectiveness with medium-to-high polarity compounds. 5. Solid-phase aroma concentrate extraction SPACE was introduced by Ishikawa et al. [11] as a modification of SPME, with the aim of increasing the area of the adsorbent so as to improve sensitivity. The device consists of a stainless steel rod coated with a mixture of adsorbents, mainly graphite carbon. Sampling is obtained by suspending the SPACE rod in the matrix headspace for a fixed time; the sampled analytes are then recovered by thermal desorption on-line to the GC or GC–MS system. This technique has been shown to be successful with roasted coffee beans and other plant matrices [67]. Table 1 lists the applications of SPACE to the HS analysis of plant matrices.

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

225

Table 2 List of the applications of HSSE to plant matrices and related products Ref.

[9]

[10] [53] [54] [55] [56] [57] [58] Jap [59] Jap [60] [61] [62] [63] [64] [65] [66] Jap

Headspace sorptive extraction (HSSE) Matrices

Main components

Other techniques

Rosmarinus officinalis L. (rosemary), Salvia officinalis L. (sage), Thymus vulgaris L. (thyme), Valeriana officinalis L. (valerian) Standard mixture

Limonene, ␤-bourbonene, thymol, bornyl acetate

HS-SPME, GC-RI

Linear hydrocarbons, chlorinated hydrocarbons, esters

HS-SPME, GC-O

Salvia officinalis L. (sage), coffee Tea, beer, yogurt and bananas Fruits Thymus vulgaris L. (thyme), Eucalyptus globulus L. (eucalyptus), Melaleuca alternifolia C. (tea tree) Benzoin gum Siam, benzoin gum Sumatra, (balsam) Mangifera indica L. (mango) Yatay palm fruits Penicillium roqueforti (surface cultures) Fusarium sambucinum, Fusarium sporotrichioides, Fusarium graminearum (surface cultures) Tuber borchii Vitt. (Tuscany white truffle), T. melanosporum Vitt. (Black truffle), T. indicum (China truffle) Lentinus edodes (Shiitake mushrooms) Roasted Arabica coffee French olive oil Flower, fruit, wine

Limonene, terpynen-4-ol, ␣-terpineol

SBSE SBSE SE Enantio-MDGC-MS

Coniferyl benzoate, p-coumaryl benzoate, benzoic acid Sulfur compounds Esters, alcohols, hydrocarbons (+)-Aristolochene, ␤-myrcene, ␤-elemene Unidentified sesquiterpene (base peak 161), trichodiene, (E)-␤-farnesene VOCs

SHS, HS-SPME

Furans, thiazoles, thiazolines 2,3-Pentanedione, pyrazine, 2-methylpyrazine (E)-Hex-2-enal, (Z)-hex-3-enol, (E)-hex-2-enol

SDE, SBSE SBSE, HS-SPME SHS, HS-SPME, DTD SBSE

LLE SBSE HS-SPME HS-SPME

Enclosed data: reference number, the common and Latin name of the vegetable matrix investigated, the main component(s) identified or class of compounds investigated, and a list of the techniques with which HSSE has been compared.

6. Large surface area high concentration capacity headspace sampling One of the parameters that has been shown to have a strong influence on analyte recovery in HCC-HS techniques is the area of the sorbent (adsorbent) exposed to the HS [15,16]. The first technique to consider the surface of the sampling polymer was membrane extraction sorbent interface (MESI) introduced in 1994 by Yang et al. [68] for in solution sampling; its use was extended to HS-sampling by Segal et al. [14] in 2000. MESI is based on a D-HS approach and consists of two simultaneous steps: extracting analytes from the vapour phase of a matrix flowing on a thin film PDMS membrane, and stripping them from the other side of the membrane into a flowing gas. The analytes stripped from the flowing gas are concentrated by a sorption (adsorption) trap. The sampled analytes are recovered by thermal desorption of the trap and directly injected into the GC or GC–MS for analysis. Transport through the nonporous membrane occurs by a solution/diffusion mechanism, and selectivity depends either on differences in membrane/sample material partition coefficient or on diffusivity [69]. Wang et al. [70] showed the effectiveness of MESI by monitoring the volatiles emitted by Eucalyptus dunnii into indoor air, using a PDMS membrane and two different traps, PDMS and Tenax. The same group [71] developed a system for on-site monitoring of biogenic emissions from Eucalyptus dunnii leaves, consisting of a portable MESI device (PDMS membrane and Tenax microtrap) on-line coupled with a portable GC.

MESI is an interesting technique, whereby the PDMS membrane acts as a selective “filter” but the analytes are dynamically accumulated on a conventional polymeric trap. The large surface of PDMS membranes has also been successfully exploited to achieve high yields in recovering analytes from both liquid and vapour phases. Bruheim et al. [15] compared the performance of a thin sheet of a PDMS membrane to that of a thick-film PDMScoated SPME fibre, in sampling PAHs-spiked water samples. They found that higher extraction efficiency in shorter time and with better sensitivity could be achieved with the PDMS membrane, for both in-solution and S-HS sampling, because of the larger surface area/extraction phase volume ratio. Recently, Sandra et al. [72] introduced sorptive tape extraction (STE), to study sebum composition before and after cosmetic treatment, through in vivo sampling at the human skin surface. Sebum components were sampled by a thin flexible PDMS tape directly in contact with the skin surface for a fixed time, to study the effect of a mattifying product on skin shininess. The sampled analytes were then recovered by either thermal or solvent desorption and analysed on-line by GC or GC–MS. The wide exchange surface offered by PDMS tape makes STE very interesting to analyse plant volatile fractions, in particular concerning the chemical messages emitted from plants (or animals) when a variation in their metabolism occurs. Bicchi et al. [16] successfully applied PDMS tapes to S-HS sampling of plant matrices and demonstrated some applications to aromatic plants and fruits, also in combination with direct-contact surface sampling. Table 1 lists the applications of MESI and STE to the HS analysis of vegetable matrices.

226

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

The main advantages of large surface area HCC-HS samplings compared to HS-SPME are increased recovery and sampling speed, due to the higher polymer volume and extended surface, while their main limits are that they require cryoconcentration and dedicated instrumentation, in particular when operating in D-HS mode. 7. Headspace liquid-phase microextraction Authors’ note: Liquid-phase microextraction (LPME) is known under several names and acronyms (e.g. single drop microextraction (SDME), headspace solvent microextraction, etc.). Here, in agreement with the nomenclature of another fundamental modern approach of sampling, i.e. solid-phase microextraction, the term used is liquid-phase microextraction, or better headspace liquid-phase microextraction (HS-LPME), unless the article(s) quoted employs different terminology. LPME was introduced about 10 years ago by Jeannot and Cantwell who successfully used a 8 ␮L drop of n-octane at the tip of a teflon rod to sample 4-methyl acetophenone from water, and analysed an aliquot of the resulting solution by GC [73]. In 1997, the same authors [74] and He and Lee [75], reduced the size of the drop to 1 ␮L of solvent, which was generated at the tip of a micro-syringe needle, thus enabling the drop to be retracted back into the needle after sampling, and then to be directly injected into the GC system. In 2001, Theis et al. [12] and Tankeviciute et al. [13] extended the use of LPME to S-HS sampling. They demonstrated the effectiveness and reliability of S-HS-LPME as a tool to preconcentrate trace levels from the headspace of aqueous solutions of model compounds (benzene, toluene, ethylbenzene, and o-xylene (BTEX)) in a 1 ␮L microdrop of 1-octanol, containing n-decane as internal standard [12], as well as those of a set of C1–C5 alcohols in a 1 ␮L microdrop of ethylene glycol as solvent and butan-2-one as internal standard [13]. This technique immediately appeared very attractive, because of its simplicity and flexibility, which is mainly related to the wide range of non-polar, polar and water miscible solvents that may be adopted. This also makes the system suitable for selective sampling, e.g. Zhang et al. [76] introduced the use of water as a solvent to extract polar and ionic analytes from the headspace of drainwater. The main limitation on the solvent is that its vapour pressure must be low enough to avoid evaporation during sampling but, at the same time, it must be compatible with GC analysis; furthermore, when aqueous samples have to be analysed, if the solvent is miscible with water the drop size may increase, causing the drop to fall from the needle [13]. In 2003, Shen and Lee [77] introduced manual DynamicHS-LPME (D-HS-LPME) to sample chlorobenzenes from soil, in the aim of increasing recovery and overcoming some of the drawbacks of conventional or S-HS-LPME mentioned above. With this approach, analyte accumulation takes place inside the micro-syringe barrel (10 ␮L), which operates first as an extractor and then as a syringe to inject the resulting solution into the GC system. Shen and Lee showed that when the syringe plunger is withdrawn (at 1 ␮L/s), a very thin organic solvent film (OSF) is generated on the inner syringe walls, thus

enabling the analytes contained in the vapour phase (5 ␮L) that is pulled in and pushed out from the syringe to be extracted by the OSF (1 ␮L). OSF and vapour phase are in mutual contact and renewed at each pull/push cycle. After a given number of cycles, the solvent plug in the barrel, containing the recovered analytes, is directly injected into the GC system. In 2005, Jiang et al. [78] introduced dynamic hollow fibre-supported HS-LPME (D-HF-HS-LPME) based on the same principle as D-HS-LPME, as described above, but where the extracting solvent is contained in a hollow fibre affixed to the syringe needle. One of the advantages of HS-LPME is that the system can easily be automated and can reliably be applied to quantitative analysis. To the best of the authors’ knowledge, the first fully-automated HS-LPME application was by Wood et al. [79]. They developed a fully-automatic S-HS-LPME method to determine residual solvents in pharmaceutical products, using n-methylpyrrolidone as accumulating liquid phase. In 2005, Ouyang et al. [80] successfully applied full automation to both S-HS-LPME and D-HS-LPME, in a study that investigated in depth the kinetics of the absorption and desorption processes involved with these techniques and their iso-tropic nature. In the same year, Saraji [81] described a semi-automatic device developed to analyse nine alcohols in water samples by D-HS-LPME using n-octanol as accumulating solvent. Very recently Ouyang et al. [82] discussed the performance of several fully-automated LPME approaches, including S-HS-LPME, exposed D-HSLPME (the solvent drop is exposed to the headspace of a sample for a given number of times, and then withdrawn into the barrel of the syringe and injected into the GC), unexposed D-HS-LPME (see ref [77]), together with the corresponding direct-immersion LPME techniques. They investigated and optimised the critical factors affecting the precision and the extraction efficiency of the methods, including temperature, choice of extraction solvent, solvent volume, plunger movement rate, and extraction time. Among the three HS-LPME techniques evaluated, exposed D-HS-LPME provided a better performance than either unexposed D-HS-LPME or S-HS-LPME. Hansson and Hakkarainen [83] demonstrated the reliability and sensitivity of S-HS-LPME quantitative analysis, by applying the multiple headspace extraction approach (MHE) to the quantitative determination of styrene in polystyrene, using butyl acetate as solvent. Sun et al. recently reviewed HS-LPME [35], discussing classification, principles of extraction, influencing factors, research status of these techniques, and their applications in particular in the tobacco industry. HS-LPME in S-HS mode has also been widely used to analyse the volatile fraction of plant matrices, or those of products derived from their transformation. Table 3 lists the applications of HS-LPME to the HS analysis of plant matrices; for each article quoted, common and Latin name of the plant investigated are given, together with reference number, solvent and internal standard (I.S.) and sampling temperature adopted, the techniques with which it has been compared and the main component(s) identified or class of compounds investigated. Unless specified otherwise, analysis of the HS-LPME extract was by GC or GC–MS.

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

227

Table 3 List of the applications of HS-LPME to the HS analysis of plant matrices Ref.

Headspace liquid-phase microextraction (HS-LPME) Matrices

Main components

[84] [86]

Pimpinella anisum L. (anise seed) Tanacetum parthenium (feverfew)

[87] [88]

SPME, SD

Paeonol

MAE

1-Octanol, menthol (I.S.) (50 ◦ C)

Panaxynol

PHWE

1-Octanol, menthol (I.S.) (80 ◦ C)

[92]

Foeniculum vulgare Mill (TCMs) Syzygium aromaticum L. (dried clove buds) Oenthera odorata Jacqui (evening primrose flowers) Cynanchum paniculatum, Paeonia suffruticosa (TCMs), dry roots Saposhnikovia divaricata Turcz., Panax ginseng L. (TCMs) Lavandula angustifolia Mill.

t-Anethole Camphor, camphene, t-chrysanthenyl-acetate t-Anethole, limonene, estragole Eugenol, ␤-caryophyllene, eugenol acetate Linalool

[93]

Tobacco powder

[94] [95]

White wine Beer and beverage

[89] [90] [91]

Other techniques

Solvent, internal standard (I.S.) extraction temperature (◦ C) Dodecane, benzophenone (I.S.) (60 ◦ C) p-Xylene, o-xylene (I.S.) (50 ◦ C) Benzyl alcohol, 1-octanol (I.S.) (70 ◦ C) 1-Octanol (25 ◦ C) Hexadecane (40 ◦ C)

Linalool, linalyl acetate, lavandulyl acetate VOC

MALDI-FTMS

Esters Dimethyl sulphide, diethyl sulphide, dimethyl disulphide

DE HS-SPME, HS-LPME, direct-LPME

n-Hexadecane, n-heptadecane (I.S.) (100 ◦ C) Dihydroxy benzoic acid, H2 O/acetone (1:1, v/v) with 1% trifluoroacetic acid (100 ◦ C) p-Cymene, n-nonane (I.S.) (35 ◦ C) N,N-Dimethylformamide (HS), thiophene (I.S.) (25 ◦ C), n-Hexane (DI), thiophene (I.S.)

Enclosed data: reference number, the common and Latin name of the plant investigated, the main component(s) identified or class of compounds investigated, the techniques with which it has been compared, solvent and internal standard (I.S.) and sampling temperature adopted

Several studies have been carried out by S-HS-LPME on plant matrices [84–93], on wine [94] and beer and beverages [95]. All articles emphasise that, compared to others, this technique is cheaper, easy to apply, flexible, and selective, because of the possibility of choosing from a large number of nonvolatile solvents; it also requires a small amount of sample. Most studies focussed on the choice of most effective solvent and internal standard (I.S.) and at the same time the selection of parameter values that maximise analyte recovery, including: solvent drop volume, sampling time and temperature, phase ratio, stirring speed and salting out for liquid samples, particle size for solid samples, and plunger movement rate for D-HS-LPME. The flexibility of the HS-LPME makes this technique easy to combine on-line and off-line with other extraction and distillation sampling techniques, including off-line microwaveassisted extraction (MAE-S-HS-LPME) [90], pressurised hot water extraction (PHWE-S-HS-LPME) [91] and on-line with continuous hydrodistillation (HD-S-HS-LPME) [92]. S-HSLPME was also directly coupled to matrix-assisted laser desorption/ionisation (MALDI) with Fourier transform mass spectrometry (FTMS) [93]. A further advantage of this technique is that the solvent drop can also be made chemically reactive to run on-line derivatisation of specific classes of compounds. Deng et al. [96] developed and validated a method to sample volatile aldehydes in water by S-HS-LPME, using n-decane as solvent, followed by in-drop derivatisation with O-2,3,4,5,6-(pentafluorobenzyl)hydroxylamine hydrochloride (PFBHA).

8. Dynamic headspace D-HS is probably still the most widely-used vapour phase sampling approach (in particular in the plant field), because of its flexibility in terms both of sampled volume, which enables the desired concentration factors to be applied even for trace components, and of the number of possible trapping approaches and materials, allowing sampling systems to be especially optimised for the problem under study. On the other hand, this flexibility requires more complex instrumentation and sampling procedures, high standardisation of several parameters to obtain good sampling reproducibility, and complex procedures for quantitative analysis. D-HS sampling is generally based on two main approaches: (a) the so called purge and trap approach (P&T) where the volatile fraction is accumulated from the gaseous flow stream stripped through the matrix onto a suitable trapping medium, such as a cold trap, a sorbent, an adsorbent or a specific reagent or solvent for a given class or classes of compounds, and (b) the dynamic approach, where analytes are sampled from the gaseous flow stream passed over the matrix. The sampled volatiles are generally recovered either by solvent elution or (more often) by thermal desorption on-line or off-line to the GC or GC–MS system. One of the first applications of D-HS was due to Herout [97]; in 1967 he collected the volatile fractions of Viola odorata, Lycaste macrobulbum and Hyacinthus orientalis through an Apiezon trap. Since then, the D-HS performance has greatly improved, thanks to the numerous new approaches and collecting materials developed to maximise analyte recovery, and to the automation, which has helped to facilitate standardisation of the parameters conditioning analyte recovery; however,

228

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

Table 4 List of the application of D-HS to the analysis of the plant matrices Ref.

Dynamic headspace (D-HS) Matrices

[25]

Cacao and hop

[55] [63]

Fruits Lentinus edodes (Shiitake mushrooms) Floral scent Ophrys sphegodes Miller (orchid) Bulbophyllum cheiri Lindl. (fruit fly orchid) Primula elatior L., Primula farinosa L. (Primula) Primula obconica Hance Yucca elata Engelmann, Yucca filamentosa Engelmann

[98] [99] [100] [101] [102] [103]

[104]

[105]

Sapromyiophilous stapeliads (Asclepiadoideae-CeropegieaeStapeliinae) Petunia axillaris Lam. (petunia)

[106]

Lysimachia punctata L. (whorled loosestrife)

[107]

Pinus Massoiana Lamb, Pinus tabulaeformis Carr., Pinus elliottii Engelm Melia azedarach L. syringa tree, commercial formulation from the Azadirachta indica Neemix 4.5® Chrysanthemum coronarium L.

[108]

[109] [110]

Fritillaria imperialis L. Crown imperial

[111]

Kiwifruit

[112]

Psidium guajava L.

[113] [114]

Tree nuts Mangifera indica L. (mango)

[115]

Apple tree

[116]

Red coffee berries

[117] [118] [119] [120]

Black truffle Tomato juices Tomato juices Spanish tomatoes

[121] [122] [123]

Orange juices Olive oil Olive oil

[124] [125] [126]

Brassica napus ssp. oleifera (colza) Starch pastes Hedera helix L. (ivy), Lycopersicon esculentum Mill. (tomato), Jasminum polyanthum (jasmine)

Main components

Furans, thiazoles, thiazolines Farnesol, geraniol, indole Nonanal, decanal, heptanal Methyl eugenol

Other techniques

Technique, trap and recovery approach

SBSE, GC–O SDE, SBSE

D-HS, PDMS, Tenax, Carbotrap300, Lichrolut EN, Chromosorb 101, thermal desorption D-HS D-HS, Tenax

SE

D-HS, Porapack Q, hexane D-HS, activated charcoal, CS2 D-HS, Porapack Q, CH2 Cl2

Limonene, benzaldehyde

D-HS, Porapack Q, CH2 Cl2

Primin, Micodin (E)-4,8-Dimethyl-1,3,7nonatriene, C11-alcohol, 1-heptanol p-Cresol, heptanal, hexanoic acid

D-HS, CH2 Cl2 , Porapack Q D-HS, Super Q, hexane

Methyl benzoate, benzaldehyde, i-eugenol (E,E)-␣-Farnesene, (Z)-3-hexenyl acetate, d-limonene ␣-Pinene, camphene, ␤-pinene

D-HS, Tenax, pentane + diethyl ether

2-Pentenal, 3-methyl-1-butanol, 6-methyl-5-hepten-2-one

D-HS, Tenax, thermal desorption

Camphor, c-chrisanthenyl acetate, Hydrocarbons 3-Methyl-2-butene-1-thiol, 3,4-dimethyl-1,5-heptadiene, 3,4-dimethyl-1,5-heptadiene, acetic acid Valencene, ␣-cadinol, flourensiadiol (E)-2-Hexanal, (Z)-3-hexenal, (Z)-3-hexenyl acetate

D-HS, Porapack Q, Pentane-CH2 Cl2

D-HS, Tenax + Carbotrap, thermal desorption

D-HS, Tenax + Carbotrap (1:1), pentane

D-HS

D-HS, Tenax, thermal desorption

D-HS, carbon, CS2 D-HS, Porapack Q, CH2 Cl2 DSE-SAFE

1-Pentanol, ethyl boronate, thujol ␣-Copaene, germacrene B, (E)-caryophyllene Limonene, isobuthyl acetate, caryophyllene Aldehydes, sulphur compounds

Hexanal, t-2-hexenal, c-3-hexenol Limonene, ethanol, linalool Hexanal, ethyl-2-methylbutyrate, decenal ␤-Myrcene, sabinene, limonene Isoamyl acetate ␣-Pinene, ␤-phellandrene, benzyl acetate

D-HS D-HS, Carbopack-X, thermal desorption D-HS, Tenax, thermal desorption D-HS, XAD-2 resin, CH2 CL2 D-HS, Tenax, thermal desorption D-HS, thermal desorption D-HS, thermal desorption D-HS, activated charcoal, CS2

S-HS S-HS, HS-SPME HS-SPME

D-HS, P&T, Tenax, thermal desorption D-HS, Tenax and Carbotrap 300, thermal desorption D-HS, Tenax, thermal desorption D-HS, Tenax, thermal desorption D-HS, PDMS, thermal desorption

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

229

Table 4 ( Continued ) Ref.

Dynamic headspace (D-HS) Matrices

Main components

Other techniques

Technique, trap and recovery approach

[127]

Jasminum polyanthum (jasmine)

HS-SPME

[128]

Citrus Sudachi (orange)

4-Methyl phenol, linalool, 3-hexenyl butyrate Limonene, linalool, p-cymene

[129] [130] [131]

Soya oil Olive oil Vitis vinifera varieties (grapes)

D-HS, PDMS and Tenax, thermal desorption P&T, Tenax and Porapack Q, thermal desorption P&T, Tenax, thermal desorption P&T P&T, Tenax, thermal desorption

[132]

Ociminum basilicum L. (basil)

[133]

Sambucus nigra L. (black sambuca) eldelflower Turkish hazelnut

[134]

Pentane, hexanal, 2-propanone Hexyl acetate, benzyl alcohol, phenethyl alcohol Linalool, methylchavicol, 1,8-cineole Hotrienol, linalool, c-rose oxide

[135]

Genipa americana L. (genipap tree)

2-Pentanone, 2-pentanal, 2-ethyl-5-methylfuran Vanillin, methylbutanoic esther, hexanoic acid

[136]

Colombian Xylopia aromatica Lamarck (fruits)

␤-Phellandrene, ␤-myrcene, p-mentha-1(7),8-diene

[137]

Flowers scent

[138]

Anthurium species

[139]

Aesculus hippocastanum L. (oak chestnut of India) Passiflora edulis flavicarpa (Yellow passion fruit)

Limonene, 1,8-cineole, ␣-pinene 1,8-Cineole, ␣-pinene, ␤-pinene Benzaldehyde, 1,8-cineole, benzyl alcohol Hexyl butanoate, hexyl hexanoate, ethyl 3-hydroxybutanoate Thiophene, methylacetate, isobutanal Ethyl acetate, ethyl isobutanoate, isobuthyl acetate 3-Methylbutanal, R-(−)-massoilactone, acetic acid 3-Carene, ␣-pinene, ␥-terpinolene

[140]

[141]

Green and roasted coffee

[142]

Artabotrys hexapetalus (traditional Chinese medicines (TCMs)) Eurycoma longifolia (Jack Tongkat Ali)

[144]

[146]

Mangifera indica L. (mango), standard mixture

SDE

P&T, Tenax, CH2 Cl2 P&T, Porapack Q, CH2 Cl2 EN, DSA

P&T, Tenax, thermal desorption

LLE, GC-FID, GC-MSA, GC-O/AEDA HD, MWHD, SFE, SDE, S-HS, HS-SPME

D-HS, Porapack Q, acetone

Liquid trapping CH2 Cl2

CLS, charcoal, CS2 SE

CLS, XAD-4 resin, CH2 Cl2 CLS, charcoal, CH2 CL2 /MeOH (2:1)

SDE, SDEV

VHS, cold trapping ether/pentane (1:1), D-HS, Tenax, pentane/diethyl ether (1:1) D-HS, Crio trap, thermal desorption

HS-SPME, HD, SE

D-HS-LPME, CH2 Cl2

HS-SPME, EN (crystal quartz sensor) HS-SPME, D-HS-SPME

D-HS

D-HS, silicone, thermal desorption

Enclosed data: reference number, the common and Latin name of the plant or derived product investigated, the main component(s) identified or class of compounds investigated, a list of the techniques with which D-HS has been compared, the D-HS approach, trapping material and recovery approach adopted.

meaningful comparisons between results obtained with different D-HS approaches, or under different conditions, are still difficult. Raguso and Pellmyr [98] critically evaluated the influence of sampling conditions on analyte recovery, comparing the performances of different trapping sorbents, elution solvents, vacuum pump flow rates and experimental parameters, in collecting volatiles in the field of floral scent analysis, in particular (a) synthetic blends on filter papers and (b) living flowers of Clarkia breweri (Onagraceae). They found qualitatively-similar results for most approaches, although the highest recovery rates were obtained using Porapak Q as sorbent and hexane as solvent. The articles that appeared between 1996 and 2007, concerning D-HS of plant matrices and their derivatives in the pharmaceutical, cosmetic and food fields, are listed in Table 4 and grouped by the D-HS approach adopted. For each article quoted, Table 4 gives the common and Latin name of the plant or derived product investigated, reference number, D-HS approach,

trapping material adopted, main component(s) identified or class of compounds investigated, and a list of the techniques with which D-HS has been compared. If not specified otherwise, D-HS samples were analysed by GC or GC–MS. Most studies concerning matrices of plant origin were by D-HS, with adsorbent trapping and thermal or solvent elution, in particular with flowers [99–106], medicinal and aromatic plants [107–110], fruits [55,111–116], mushrooms, [63,117], fruits and vegetable juices [118–121], vegetable oils [122–125], or by sorption on PDMS particles [25,126,127]. A second very common approach is the purge and trap technique, whereby capture of the volatile fraction from the DHS flow stream is by solid adsorbent and thermal desorption [128–134], or by sorbent trapping and solvent elution [135], or alternatively by solvent trapping [136]. Other applications are based on closed loop stripping combined with different adsorption material [137–139], or sampling under vacuum-cold trapping and liquid–liquid extraction of

230

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

the aqueous phase [140] or again by cold or solvent trapping [141,142]. D-HS with different approaches is still the reference technique to study plant–insect interactions, probably because of the possibility it offers to achieve high concentration factors by sampling a suitable HS volume [99–101,103,106,108,115,137,139]. This approach has also been directly combined with electronic nose or sensor systems, to monitor volatiles emitted from plants either by (D-TD) [143] or by dynamic HS transfer [144]. In 2003, Augusto et al. patented an apparatus for solid-phase microextraction in D-HS mode [145,146] where the analytes in the nitrogen stream purging a liquid or solid sample in a glass container (125 mm × 9 mm i.d.) are on-line trapped on the SPME fibre exposed from a conventional holder in a small chamber directly connected to the sample container. Markelov and Bershevits [147] described a HS technique affording total or partial sampling of the vapour phase from a pre-equilibrated headspace vial, to analyse low ppb levels of highly volatile compounds (butadiene, butylenes, isoprene and others). The flow, generated via a dual needle system through the headspace vial, is directed either to the front of a GC column through a sample loop, or through a trapping system that can afterward be thermally desorbed directly into the GC system. 9. General considerations The constant evolution of HS-sampling over the last 10–15 years has greatly extended the perspectives of this technique, also in view of the ever-increasing demand of analytical controls that are required in a routine analysis laboratory. The most recent HCC-HS techniques can operate reliably in non-equilibrium conditions, and offer sufficiently-high recovery and concentration factors over time to achieve sampling speeds rapid enough to accumulate compounds emitted even in trace amounts (ppb level). At the same time, these techniques are also so simple, fast and easy to automate as to be suitable to characterise a plant matrix through the rapid analysis of both direct and indirect markers. Direct markers are components whose presence or amount directly characterises a plant matrix. Whenever their analysis requires time-consuming and complex methods, indirect markers can also be used. In general, they are volatile(s) (or fractions) directly or indirectly related to the direct marker(s) but easier and/or quicker to analyse. This approach makes it also possible to use the HS-chromatographic profile as a parameter to define quality or to classify a plant matrix, provided that it is appropriately “evaluated”, for instance by resorting to a statistical approach such as multivariate statistical analysis (e.g. PCA) or neural networks. On the other hand, HS sampling can also be directly combined with MS [148], thus excluding the separation step, giving an MS profile significant and diagnostic of the matrix investigated. HS-GC or HS-MS profiles can therefore be adopted as analytical decision-makers (ADM) [149] useful to limit the number of conventional and time-consuming analyses to samples that cannot be classified unequivocally with fast preliminary analysis. Moreover, the new HS approaches are also easy to automate [e.g [8,10,82]], making them suitable for use in total analysis systems (TAS), i.e. systems where sample prepa-

ration and analysis are on-line automatically combined. Last but not least, the most recent techniques are highly flexible and can be used equally in liquid or vapour phase samplings. This characteristic is particularly useful in the food and cosmetic fields, because it enables direct correlation of (1) the HS composition of a solid or liquid matrix to that of the corresponding solvent or water extracts, (2) the composition of the extract to that of its HS in equilibrium with it, and (3) it allows sensory analysts to correlate HS and matrix chemical compositions reliably to taste and/or to odour detected via both orto- and retro-nasal pathways. In spite of these advantages, a lot of work has yet to be done to overcome some of the limits of HS-sampling. In depth investigations are necessary, in particular for HCC-HS techniques, to increase our knowledge on non-equilibrium HS sampling and to develop new polymeric phases to make these techniques even more effective and flexible. This is particularly true for fast HSsampling, in order to make logical its combination with high speed GC, keeping in mind that ultra-fast GC method (also known as “sensor-like” GC) [150] that can analyse a simple mixture in about 1 s are under development. These developments would also be very useful for fast in vivo sampling, and to analyse a plant’s volatile fraction so as to monitor the dynamics of a rapid biological phenomenon, such as the plant’s reaction to a stress. On the other hand, trapping materials covering a wide range of polarities are necessary for HS sampling in the sorption mode, because only apolar PDMS is currently used on a routine basis, thus limiting recovery of polar and/or very volatile (C1–C4) components from complex solid matrices. Another important topic requiring investigation, and which is increasingly necessary, is the absolute quantitative determination of an analyte contained in a matrix. This is still a serious problem although HS sampling in both S-HS and D-HS modes is highly repeatable and reproducible, provided that rigorous standardisation of all parameters is adopted. With a given HCC-HS S-HS based technique, few easily monitored parameters must be controlled, i.e. sampling temperature and time and phase ratio, besides the analyte accumulation tools (e.g. fibres for SPME, twisters for HSSE, solvents for LPME, etc.). The situation is different for HCC-HS D-HS based techniques because, as mentioned above [98], the approach and sampling conditions adopted greatly influence HS composition. Comparable results can therefore only be obtained by adopting similar approaches (see Section 8) and strictly controlling all parameters conditioning HS composition (temperature and phase ratio), analyte stripping (sampling time, carrier gas and flow rate or vacuum conditions), and trapping (sorbent or adsorbent material, solvents or cryogenic fluid). For component absolute quantitation in liquid samples, the internal standard approach is the most widely used for both S-HS and D-HS modes, although sometimes timeconsuming operations are necessary to determine the response factor to the entire HS-sampling procedure of each analyte to be quantified, in order to overcome discrimination due to the matrix effect. With solid samples, analyte quantitation is more difficult, in particular when a reference sample is not available, or when one with a known composition cannot be “built”, as is often the case with plant matrices. The most reliable approach for HCC-

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

HS techniques operating in S-HS mode is multiple headspace extraction: although it is rather complex and time-consuming, in particular when several analytes must be simultaneously quantified [151], this approach enables reliable calibration curves to be constructed. With HCC-HS D-HS, analyte quantitation in the matrix is more complex, and requires preliminary quantitative determination of the analyte(s) under investigation contained in the matrix by a different auxiliary method (e.g. exhaustive extraction). This enables its (their) recovery to be determined correctly through the ratio of its (their) normalised areas obtained by the D-HS method applied and by the auxiliary method, taken as a reference. 10. Conclusions The renewed interest in research and development of HS-sampling is one of the consequences of the dramaticallyincreasing demand for analyses (in particular for control purposes) over the last decade. It is also due to the everincreasing importance of the volatile fraction, not only in direct and indirect matrix characterisation, but also to clarify biological phenomena, in particular in the plant field. The pressing demand for innovative analytical approaches to deal with the above problems has produced improvements in the performance of HS techniques that were unimaginable 10 years ago, resulting in ever-more effective HCC-HS techniques, robust technologies and automatic HS systems. Nevertheless, much research effort is still required to develop reliable new approaches that will facilitate quantitative analysis and reduce sampling time, as well as to find materials that can extend the fields of application of HCC-HS techniques. Acknowledgements This study was carried out within the project entitled: “Sviluppo di metodologie innovative per l’analisi di prodotti agroalimentari” of the Ministero dell’Istruzione, dell’Universit`a e della Ricerca (MIUR) (Italy). References [1] L.S. Ettre, LCGC N. Am. 20 (2002) 1120. [2] R.N. Harger, E.G. Bridwell, B.B. Raney, J. Biol. Chem. 128 (1939) XXXVIII. [3] W.H. Stahl, W.A. Voelker, J.H. Sullivan, Food Technol. 14 (1960) 14. [4] L. Bovijn, J. Pirotte, A. Berger, in: D.H. Desty (Ed.), Gas Chromatography, London, 1958, p. 310. [5] B. Kolb, L.S. Ettre, Headspace Gas Chromatography, Wiley-VCH, New York, 1997, p. 4. [6] Z. Zhang, J. Pawliszyn, Anal. Chem. 65 (1993) 1843. [7] M.E. McComb, R.D. Oleschuk, E. Giller, H.D. Gesser, Talanta 44 (1997) 2137. [8] F. Musshoff, D.W. Lachenmeier, L. Kroener, B. Madea, J Chromatogr. A 958 (2002) 231. [9] C. Bicchi, C. Cordero, C. Iori, P. Rubiolo, P. Sandra, J. High Resolut. Chromatogr. 23 (2000) 539. [10] B. Tienpont, F. David, C. Bicchi, P. Sandra, J. Microcol. Sep. 12 (2000) 577. [11] M. Ishikawa, O. Ito, S. Ishizaki, Y. Kurobayashi, A. Fujita, Flav. Fragr. J. 19 (2004) 183.

231

[12] A.L. Theis, A.J. Waldack, S.M. Hansen, M.A. Jeannot, Anal. Chem. 73 (2001) 5651. [13] A. Tankeviciute, R. Kazlauskas, V. Vickackaite, Analyst 126 (2001) 1674. [14] A. Segal, T. Gorecki, P. Mussche, J. Lips, J. Pawliszyn, J. Chromatogr. A 873 (2000) 13. [15] I. Bruheim, X. Liu, J. Pawliszyn, Anal. Chem. 75 (2003) 1002. [16] C. Bicchi, C. Cordero, E. Liberto, P. Rubiolo, B. Sgorbini, P. Sandra, J. Chromatogr. A 1148 (2007) 137. [17] C.L. Arthur, J. Pawliszyn, Anal. Chem. 62 (1990) 2145. [18] T. G´orecki, J. Pawliszyn, Anal. Chem. 67 (1995) 3265. [19] H. Lord, J. Pawliszyn, J. Chromatogr A. 885 (2000) 153. [20] J. Pawliszyn, Solid Phase Microextraction—Theory and Practice, 1st ed., Wiley-VCH, New York, 1997. [21] J. Pawliszyn, Sampling and Sample Preparation for Field and Laboratory, Elsevier, Amsterdam, 2002, p. 389. [22] E. Baltussen, P. Sandra, F. David, H.G. Janssen, C. Cramers, Anal. Chem. 71 (1999) 5213. [23] E. Baltussen, PhD Thesis, Technische Universireit Eindhoven, Eindhoven, 2000, p. 10. [24] E. Baltussen, C. Cramers, P.J.F. Sandra, Anal. Bioanal. Chem. 373 (2002) 3. [25] E. Baltussen, F. David, P. Sandra, H.G. Janssen, C. Cramers, J. High Resolut. Chromatogr. 21 (1998) 332. [26] D. Tholl, W. Boland, A. Hansel, F. Loreto, U.S.R. Rose, J.P. Schnitzler, Plant J. 45 (2006) 540. [27] European Pharmacopoeia, European Directorate for the Quality of Medicines, EDQM-Council of Europe, Strasbourg, 5th ed., 2004, Suppl. 5.8. [28] C. Bicchi, F. Belliardo, C. Cordero, E. Liberto, P. Rubiolo, B. Sgorbini, J. Chromatogr. Sci. 44 (2006) 416. [29] Sci-Finder Chemical Abstract Data Base, America Chemical Society, Washington, DC, http://www.cas.org. [30] A.G. Vitenberg, J. Anal. Chem. 58 (2003) 2. [31] J.V. Hinshaw, LC–GC 18 (2000) 940. [32] A. Nongonierma, P. Cayot, J.L. Le Quere, M. Springett, A. Voilley, Food Rev. Int. 22 (2006) 51. [33] R.M. Smith, J. Chromatogr. A 1000 (2003) 3. [34] C. Bicchi, C. Cordero, P. Rubiolo, J. Chromatogr. Sci. 42 (2004) 402. [35] S.H. Sun, Y.L. Zong, J.P. Xie, Yancao Keji 5 (2006) 41. [36] H.B. Jakobsen, Plant volatile analysis Modern Methods of Plant Analysis, vol. 19, Springer, Dordrecht, 1997, p. 1. [37] E.J. Brunke, F.J. Hammerschmidt, F. Rittler, G. Schmaus, SOFW 122 (1996) 593. [38] R. Rouseff, K. Cadwallader, Adv. Exp. Med. Biol. 488 (2001) 1. [39] H. Song, Ziran Kexueban 24 (2006) 1. [40] F. Augusto, A. Leite e Lopes, C. Alcaraz Zini, Trac-Trends Anal. Chem. 22 (2003) 160. [41] J.G. Wilkes, E.D. Conte, Y. Kim, M. Holcomb, J.B. Sutherland, D.W. Miller, J. Chromatogr. A 880 (2000) 3. [42] W.J. Reid, in: B. Baigrie (Ed.), Taints and Off-Flavours in Food, Woodhead Publ., Cambridge, UK, 2003, p. 31. [43] M. Mestres, O. Busto, J. Guasch, J. Chromatogr. A 881 (2000) 569. [44] S. Shojania, M.E. McComb, R.D. Oleschuk, H. Perreault, H.D. Gesser, A. Chow, Can. J. Chem. 77 (1999) 1716. [45] J. Lipinski, Fres. J. Anal. Chem. 369 (2001) 57. [46] F. Musshoff, D.W. Lachenmeier, L. Kroener, B. Madea. Forensic Sci. Int. 133 (2003) 32. [47] K. Ridgway, S.P.D. Lalljie, R.M. Smith, J. Chromatogr. A 1124 (2006) 181. [48] C. Bicchi, C. Cordero, E. Liberto, P. Rubiolo, B. Sgorbini, J. Chromatogr. A 1024 (2004) 217. [49] S. Ampuero, S. Bogdanov, J.O. Bosset, Eur. Food Res. Technol. 218 (2004) 198. [50] J.O. Bosset, L. Pillonel, D. Altieri, R. Tabacchi, Mitt. Lebensmitt. unters. Hyg. 95 (2004) 85. [51] E. Baltussen, P. Sandra, F. David, C. Cramers, J. Microcol. Sep. 11 (1999) 737.

232

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233

[52] C. Bicchi, C. Cordero, E. Liberto, P. Rubiolo, B. Sgorbini, P. Sandra, J. Chromatogr. A 1071 (2005) 111. [53] C. Bicchi, C. Cordero, E. Liberto, P. Rubiolo, B. Sgorbini, F. David, P. Sandra, J. Chromatogr. A 1094 (2005) 9. [54] P. Sandra, F. David, J. Vercammen, Advances in Flavours and Fragrances, Special Publication – Royal Society of Chemistry 277 (2002) 27. [55] S. Eri, E. Adedeji, L. Trinnaman, presented at the 230th ACS National Meeting, Washington, DC, 2005, Abstract of papers. [56] M. Kreck, A. Scharrer, S. Bilke, A. Mosandl, Flav. Fragr. J. 17 (1) (2002) 32. [57] C. Castel, X. Fernandez, L. Lizzani-Cuvelier, A.M. Loiseau, C. Perichet, C. Delbecque, J.F. Arnaudo, Flav. Fragr. J. 21 (2006) 59. [58] Y. Akiyama, Koryo 228 (2005) 153. [59] Z. Lu, H. Tsuda, N. Matsuura, Aroma Res. 7 (2006) 368. [60] J.C.R. Demyttenaere, R.M. Morina, P. Sandra, J. Chromatogr. A 985 (2003) 127. [61] J.C.R. Demyttenaere, R.M. Morina, N. De Kimpe, P. Sandra, J. Chromatogr. A 1027 (2004) 147. [62] R. Splivallo, S. Bossi, M. Maffei, P. Bonfante, Phytochemistry D-0600636R2. [63] N. Da Costa, S. Eri, Publisher: American Chemical Society, ACS Symposium Series 926 (Modern Extraction Techniques) (2006) 163. [64] C. Bicchi, C. Iori, P. Rubiolo, P. Sandra, J. Agric. Food Chem. 50 (2002) 449. [65] J.F. Cavalli, X. Fernandez, L. Lizzani-Cuvelier, A.M. Loiseau, J. Agric. Food Chem. 51 (2003) 7709. [66] N. Matsuura, Koryo 230 (2006) 145. [67] S. Ishizaki, O. Ito, A. Fujita, Y. Kurobayashi, M. Ishikawa, in: J.-L. Le Quere, P. Etievant (Eds.), Proceedings of the 10th Weurman Flavor Research Symposium, Beaune, France, 2002, Tec & Doc, Paris, 2003, p. 634. [68] M.J. Yang, S. Harms, Y.Z. Luo, J. Pawliszyn, Anal. Chem. 66 (1994) 1339. [69] M. Mulder, Basic Principles of Membrane Technology, Kluwer, Dordrecht, 1991. [70] L. Wang, H. Lord, R. Morehead, F. Dorman, J. Pawliszyn, J. Agric. Food Chem. 50 (2002) 6281. [71] X. Liu, R. Pawliszyn, L. Wang, J. Pawliszyn, Analyst 129 (2004) 55. [72] P. Sandra, S. Sisalli, A. Adao, M. Lebel, I. Le Fur, LC–GC Eur. 19 (2006) 3311. [73] M.A. Jeannot, F.F. Cantwell, Anal. Chem. 68 (1996) 2236. [74] M.A. Jeannot, F.F. Cantwell, Anal. Chem. 69 (1997) 235. [75] Y. He, H.K. Lee, Anal. Chem. 69 (1997) 4634. [76] J. Zhang, T. Su, H.K. Lee, Anal. Chem. 77 (2005) 1988. [77] G. Shen, H.K. Lee, Anal. Chem. 75 (2003) 98. [78] X. Jiang, C. Basheer, J. Zhang, H.K. Lee, J. Chromatogr. A 1087 (2005) 289. [79] D.C. Wood, J.M. Miller, I. Christ, LC–GC Eur. 17 (2004) 573 and LC–GC North Am. 22 (2004) 516. [80] G. Ouyang, W. Zhao, J. Pawliszyn, Anal. Chem. 77 (2005) 8122. [81] M. Saraji, J. Chromatogr. A 1062 (2005) 15. [82] G. Ouyang, W. Zhao, J. Pawliszyn, J. Chromatogr. A 1138 (2007) 47. [83] E. Hansson, M. Hakkarainen, J. Chromatogr. A 1102 (2006) 91. [84] A. Besharati-Seidani, A. Jabbari, Y. Yamini, Anal. Chim. Acta 530 (2005) 155. [85] J. Cao, M. Qi, Y. Zhang, S. Shan, Q. Shao, R. Fu, Anal. Chim. Acta 561 (2006) 88. [86] A. Besharati-Seidani, A. Jabbari, Y. Yamini, M.J. Saharkhiz, Flav. Fragr. J. 21 (2006) 502. [87] L. Fang, M. Qi, T. Li, Q. Shao, R. Fu, J. Pharm. Biomed. Anal. 41 (2006) 791. [88] M.J. Jung, Y.J. Shin, S.Y. Oh, N.S. Kim, K. Kim, D.S. Lee, Bull. Korean Chem. Soc. 27 (2006) 231. [89] N.S. Kim, M.J. Jung, Z.W. Yoo, S.N. Lee, D.S. Lee, Bull. Korean Chem. Soc. 26 (2005) 1996. [90] C. Deng, N. Yao, B. Wang, X. Zhang, J. Chromatogr. A 1103 (2006) 15. [91] C. Deng, X. Yang, X. Zhang, Talanta 68 (2005) 6.

[92] A.R. Fakhari, P. Salehi, R. Heydari, S.N. Ebrahimi, P.R. Haddad, J. Chromatogr. A 1098 (2005) 14. [93] S. Sun, Z. Cheng, J. Xie, J. Zhang, Y. Liao, H. Wang, Y. Guo, Rapid Commun. Mass Spectrom. 19 (2005) 1025. [94] A. Tankeviciute, R. Bobnis, R. Kazlauskas, V. Vickackaite, Chem. Anal. 50 (2005) 539. [95] Q. Xiao, C. Yu, J. Xing, B. Hu, J. Chromatogr. A 1125 (2006) 133. [96] C. Deng, N. Yao, N. Li, X. Zhang, J. Sep. Sci. 28 (2005) 2301. [97] V. Herout, Planta Medica Suppl. (1967) 68. [98] R.A. Raguso, O. Pellmyr, Oikos 81 (1998) 238. [99] F.P. Schiestl, M. Ayasse, H.F. Paulus, D. Erdmann, W. Francke, J. Chem. Ecol. 23 (1997) 2881. [100] K.H. Tan, R. Nishida, T.C. Toong, J. Chem. Ecol. 28 (2002) 1161. [101] A.C. Gaskett, E. Conti, F.P. Schiestl, J. Chem. Ecol. 31 (2005) 1223. [102] L.P. Christensen, E. Larsen, Contact Dermatitis 42 (2000) 149. [103] G.P. Svensson, O. Pellmyr, R.A. Raguso, J. Chem. Ecol. 32 (2006) 2657. [104] A. Jurgens, S. Dotterl, U. Meve, New Phytol. 172 (2006) 452. [105] M. Kondo, N. Oyama-okubo, T. Ando, E. Marchesi, M. Nakayama, Ann. Bot. 98 (2006) 1253. [106] S. D¨otterl, I. Sch¨affler, J. Chem. Ecol. 33 (2007) 441. [107] X. Qin, J. Su, F.Z.Z. Ge, Fenxi Ceshi Xuebao 25 (2006) 6 (in Chinese). [108] D.S. Charleston, R. Gols, K.S. Hordijk, R. Kfir, L.E.M. Vet, M. Dicke, J. Chem. Ecol. 32 (2006) 325. [109] B. Sebastian, A.M. Urzua, M. Vines, Flav. Fragr J. 21 (2006) 783. [110] J.P.F.G. Helsper, M. Buecking, S. Muresan, J. Blaas, W.A. Wietsma, J. Agric. Food Chem. 54 (2006) 5087. [111] A. Samadi-Maybodi, M.R. Shariat, M. Zarei, M.B. Rezai, J. Essential Oil Res. 14 (2002) 414. [112] F.D. Soares, T. Pereira, M.O. Maio Marques, A.R. Monteiro, Food Chem. 100 (2007) 15. [113] K.R. Cadwallader, Q. Zhou, presented at the 231st ACS National Meeting, Atlanta, GA, United States, 2006, Abstract of papers. [114] M. Moalemiyan, A. Vikram, A.C. Kushalappa, V. Yaylayan, Plant Pathol. 55 (2006) 792. [115] A. Stewart-Jones, G.M. Poppy, J. Chem. Ecol. 32 (2006) 845. [116] F. Mathieu, C. Malosse, A.H. Cain, B. Frerot, J. High Resolut. Chromatogr. 19 (1996) 298. [117] O. Jansen, T. Talou, C. Raynaud, A. Gaset, Food Flavor and Chemistry, (Special Publication 300), Royal Society of Chemistry, Cambridge, 2005, p. 260. [118] M.K. Sucan, Mathias K, presented at the 216th ACS National Meeting, Boston, 1998, Book of abstract. [119] M.K. Sucan, G.F. Russell, Adv. Exp. Med. Biol. 488 (2001) 165. [120] A.A. Carbonell-Barrachina, A. Agusti, J.J. Ruiz, Eur. Food Res. Technol. 222 (2006) 536. [121] E. Bylaite, A.S. Meyer, Eur. Food Res. Technol. 222 (2006) 176. [122] M. Servili, R. Selvaggini, F. Jalali, G. Montedoro, EPPOS (Spec. Num.) (1997) 311. [123] A. Kanavouras, A. Kiritsakis, R.J. Hernandez, Food Chem. 90 (2005) 69. [124] M. McEwan, S.W.H. Macfarlane, Clin. Exp. Allergy 28 (1998) 332. [125] N. Cayot, F. Pretot, J.L. Doublier, J.M. Meunier, E. Guichard Elisabeth, J. Agric. Food Chem. 52 (2004) 5436. [126] J. Vercammen, H. Pham-Tuan, P. Sandra, J. Chromatogr. A 930 (2001) 39. [127] J. Vercammen, P. Sandra, E. Baltussen, T. Sandra, F. David, J. High Resolut. Chromatogr. 23 (2000) 547. [128] A. Padrayuttawat, H. Tamura, M. Yamao, J. High Resolut. Chromatogr. 19 (1996) 365. [129] V. Pinnel, J. Vandegans, J. High Resolut. Chromatogr. 19 (1996) 263. [130] T. Hofmann, M. Rothe, P. Schieberle, in: M. Dachtler, C. Engelen, C. Boucon, G. Dol (Eds.), Proceedings of the 7th Wartburg Symposium on Flavour Chemistry and Biology, Eisenach, Germany, 2004 Deutsche Forschungsanstalt fuer Lebensmittelchemie, Garching, Germany, p. 332. [131] L. Rosillo, M.R. Salinas, J. Garijo, G.L. Alonso, J. Chromatogr. A 847 (1999) 155. [132] A.N. Yousif, C.H. Scaman, T.D. Durance, B. Girard, J. Agric. Food Chem. 47 (11) (1999) 4777.

C. Bicchi et al. / J. Chromatogr. A 1184 (2008) 220–233 [133] K. Kaack, L.P. Christensen, M. Hughes, R. Eder, Eur. Food Res. Technol. 223 (2006) 57. [134] C. Alasalvar, A.Z. Odabasi, N. Demir, M.O. Balaban, F. Shahidi, K.R. Cadwallader, J. Food Sci. 69 (2004) 99. [135] A.B. Pinto, C.M. Guedes, R.F.A. Moreira, C.A.B. De Maria, Flav. Fragr. J. 21 (2006) 488. [136] E.E. Stashenko, B.E. Jaramillo, J.R. Martinez, J. Chromatogr. A. 1025 (2004) 105. [137] H.J. Bestmann, L. Winkler, O. von Helversen, Phytochemistry 46 (1997) 1169. [138] N. Kuanprasert, A.R. Kuehnle, C.S. Tang, Phytochemistry 49 (1998) 521. [139] A.B. Johne, B. Weissbecker, S. Sch¨utz, J. Chem. Ecol. 32 (2006) 2303. [140] P. Werkhoff, M. Guentert, G. Krammer, H. Sommer, J. Kaulen, J. Agric. Food Chem. 46 (1998) 1076. [141] G. Procida, B. Campisi, D. De Palo, A. Calabretti, L. Gabrielli Favretto, 19th Colloque Scientifique International sur le Cafe, Association Scientifique Internationale du Cafe, 2001 p. 219.

233

[142] C. Mahidol, N. Chimnoi, D. Chokchaichamnankit, S. Techasakul, Acta Horticul. 677 (43–50) (2005) 43–50. [143] P. Mielle, M. Souchaud, P. Landy, E. Guichard, Sens. Actuators B: Chem. B 116 (2006) 161. [144] A.K.M. Shafiqul Islam, Z. Ismail, B. Saad, A.R. Othman, M.N. Ahmad, A.Y. Shakaff, Sens. Actuators B: Chem. 120 (1) (2006) 245. [145] F. Augusto, A.L.P. Valente, P.M. Saldanha de Aguiar, R. Cesar da Silva, Braz. Pat. BR 2002 001 343, kind A (16 Dec. 2003); Braz. Appl. BR 2002-1343 (4 April 2002); Priority Appl. 2002-1343 (4 April 2002) (in Portuguese). [146] R.C. Silva, P.M.S. Aguiar, F. Augusto, Chromatographia 60 (2004) 687. [147] M. Markelov, O.A. Bershevits, Anal. Chim. Acta 432 (2001) 213. [148] I.M. Lorenzo, J.L. Perez Pavon, M.E. Fernandez Laespada, C. Garcia Pinto, B. Moreno Cordero, J. Chromatogr. A 945 (2002) 221. [149] P. Sandra, F. David, B. Tienpont, Chromatographia 60 (2004) S299. [150] G.M. Gross, B.J. Prazen, J.W. Grate, R.E. Synovec, Anal. Chem. 76 (2004) 3517. [151] B. Kolb, L.S. Ettre, Headspace Gas Chromatography, Wiley-VCH, New York, 1997, p. 40.