DEVELOPMENT OF SOLVENT-MINIMIZED EXTRACTION PROCEDURES FOR ENVIRONMENTAL ANALYSIS MAUNG PAN (B.Sc., UNIVERSITY OF YANGON) A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT OF CHEMISTRY NATIONAL UNIVERSITY OF SINGAPORE 2007 Acknowledgements There are important persons to whom I am indebted for their help, guidance, advice, support and patience throughout this course. First of all, I would like to express my sincere gratitude to my supervisor, Professor Hian Kee Lee for his understanding and giving me a chance to be his student. I would also like to express my appreciation to Dr. Chanbasha Basheer for his suggestions, support and tolerance throughout this work. Ms Frances Lim is really an important person to all the students including me, by offering her invaluable technical assistance and advices. I give special thanks to her. I finally would like to thank all the students in our group for their kind assistance and friendship. Most of all, I thank my parents for their love, patience and encouragement. i Contents Acknowledgements i Contents ii Summary vi International Conference Papers viii Chapter 1. Sample preparation techniques 1.1. Introduction 1 1.2. Extraction of organics from liquids 4 1.2.1. Liquid-liquid extraction 5 1.2.2. Flow injection analysis (FIA) 6 1.2.3. Liquid-phase microextraction (LPME) 7 1.2.4. Hollow fiber membrane-based LPME (HFM-LPME) 9 1.2.5. Purge and trap (P&T) or dynamic headspace 10 1.2.6. Static headspace extraction 11 1.2.7. Solid-phase extraction (SPE) 12 1.2.8. Solid-phase microextraction (SPME) 13 1.3. Extraction of organics from solid matrices 16 1.3.1. Soxhlet and Soxtec 16 1.3.2. Pressurized fluid extraction (PFE) 17 1.3.3. Ultrasonic extraction (USE) 18 1.3.4. Microwave assisted extraction (MAE) 18 ii 1.3.5. Supercritical fluid extraction (SFE) 18 1.3.6. Direct thermal extraction (DTE) 19 1.4. Chromatography in environmental analysis 21 1.7. Scope of this study 22 References 23 Chapter 2. Room temperature ionic-liquid as solvent in hollow fiber-protected liquid-liquid-liquid microextraction technique coupled with high performance liquid chromatography 2.1. Introduction 27 2.2. Experimental 29 2.2.1. Chemicals and reagents 29 2.2.2 Materials 30 2.2.3. Wastewater samples 30 2.2.4. HPLC 30 2.2.5. Ionic-liquid based LLLME 30 2.3. Results and discussion 32 2.4. Method performance 38 2.5. Conclusion 42 References 43 iii Chapter 3. Novel micro-solid-phase extraction of carbamates in green tea leaves with determination by high performance liquid chromatography 3.1. Introduction 46 3.2. Expreimental 47 3.2.1. Chemicals and materials 47 3.2.2. Chromatographic analysis 49 3.2.3. Sample preparation 49 3.2.4. Micro-solid phase extraction (µ-SPE) procedure 50 3.2.5. Principle of µ-SPE 51 3.3. Results and discussion 52 3.3.1. Optimization of the method 52 3.3.2. Individual and mixed-mode sorbents approaches 54 3.3.3. Effect of extraction and desorption time 56 3.3.4. Dependence of pH and ionic strength 57 3.3.5. Dependence of sorption on sample volume 58 3.3.6. Method evaluation 59 3.4. Conclusion 60 References 61 Chapter 4. Novel amphiphilic poly(p-phenylene)s used as sorbent for solid-phase microextraction of environmental pollutants 4.1. Introduction 64 iv 4.2. Experimental 66 4.2.1. Materials and reagents 66 4.2.2. GC-MS analysis 67 4.2.3. Amphiphilic poly(p-phenylene)s 68 4.2.4. Synthetic scheme 69 4.2.5. Preparation of SPME fiber 70 4.2.6. SPME theory 71 4.2.7. SPME procedure 72 4.3. Results and discussion 73 4.3.1. C12PPPOH vs commercial fibers 73 4.3.2. Optimization of PAHs extraction using C12PPPOH coating 75 4.3.3. Method validation 77 4.3.4. SPME/GC-MS of real water sample 79 4.4. Conclusion 79 References 80 Chapter 5. Conclusion 84 v Summary The analysis of environmental pollutants is a very complex exercise. In many such applications, analytes must be determined in complicated matrices, such as soil, sludge, blood, foods, waters and wastewater at very low concentrations. The aims in environmental analysis are sensitivity (due to the low concentration of microcontaminants to be determined), selectivity (due to the complexity of the sample) and automation (to increase the throughput in control analysis). Notable among recent developments are simple, faster and greener (environmentally friendly) microextraction techniques. This thesis focuses on the developments of solvent-minimized extraction techniques including liquid-liquid-liquid microextraction (LLLME) and micro-solidphase extraction (µ-SPE) combined with high-performance liquid chromatography (HPLC) and solid-phase microextraction (SPME) combined with gas chromatography mass spectrometry (GC-MS). Chapter 1 introduces an overview and the background of sample preparation/ extraction methods in environmental analysis for solid and liquid samples. In Chapter 2, a green solvent, an ionic-liquid, is applied as an acceptor phase inside the hollow fiber membrane for the first time in LLLME. The advantages of this work are that (1) sensitivity is improved by injecting a larger volume of extract directly into the HPLC, (2) porous polypropylene hollow fiber membrane (HFM) serves as a protective sleeve for LLLME providing a very efficient sample cleanup for dirty wastewater samples compared to single drop liquid-phase microextraction (LPME) which has limited injection volume and is not a desirable for dirty samples such as vi wastewater. The ionic liquid, 1-butyl-3-methylimidazolium hexafluorophosphate ([BMIM][PF6]) mixed with acetonitrile proved to be an excellent solvent for extraction of phenolic compounds from wastewater sample. µ-SPE is developed for the determination of carbamates pesticides in green tea leaves, this is reported in Chapter 3. Polar and non-polar sorbents are packed polypropylene microporous membrane envelopes and these are used as extraction devices. After extraction, the devices are desorbed in a suitable organic solvent. This desorbing solvent is directly injected into the HPLC. µ-SPE offers good extraction efficiency and sample cleanup when C18 is used as packing material. They have several advantages over traditional SPE: (1) the envelopes are affordable and simple to prepare, (2) the porous membrane serves as both a pre-concentration and clean-up device (further purification is not necessary compared to traditional SPE) and carry over effects can be eliminated since µ-SPE devices are ultrasonically cleaned in acetone after each extraction, (3) the amount of organic solvent used is reduced and the final extract is compatible with HPLC. Chapter 4 introduces the application of novel amphiphilic polymer coated fused silica capillary tubing for the pre-concentration of PAHs, OCPs and OPPs from environmental water samples. Comparative studies were also made with commercial SPME fibers (PDMS-DVB, PA) for the above compounds. PAHs were studied as a reference analytes for method evaluation and extraction parameters such as pH and salting-out effects were investigated. The PPP coated capillary could be applied at up to 320 oC and was used for the pre-concentration/extraction of PAHs in sea water collected from St. John’s Island, Singapore. vii International Conference Papers [1] Chanbasha Basheer, Maung Pan and Hian Kee Lee, "Room temperature ionic-liquid as solvent in hollow fiber-protected liquid-liquid-liquid microextraction technique for wastewater extraction coupled with high performance liquid chromatography". 9th International Symposium on Hyphenated Techniques in Chromatography and Hyphenated Chromatographic Analyzers & 8th International Symposium on Advances in Extraction Techniques, 10 February 2006, York, UK. [2] Chanbasha Basheer, Maung Pan, Zhang Jie and Hian Kee Lee, "Single-step microwave-assisted headspace liquid-phase microextraction for the analysis of aromatic amines in sediment samples”. 9th International Symposium on Hyphenated Techniques in Chromatography and Hyphenated Chromatographic Analyzers & 8th International Symposium on Advances in Extraction Techniques, 10 February 2006, York, UK. viii Chapter 1. Sample Preparation Techniques 1.1. Introduction Sample preparation is often the most time-consuming step in environmental analysis. The goal of sample preparation is enrichment, cleanup, and signal enhancement. Sample preparation is often the bottleneck in a measurement process, as it tends to be slow and labor-intensive. It is important in all aspects of environmental, chemical, biological, materials, and surface analysis. Notable among recent developments are faster, greener extraction methods and microextraction techniques [1]. The common steps involved in a typical environmental analysis are shown in Figure.1.1.1. Sampling Sample Preservation Sample Preparation Analysis Even Sampling Suitable Container Homogenization Size reduction Instrument Calibration Extraction Instrument Analysis Representative Sample Storage time, Temperature Concentration Without Contamination Clean-up Data Processing Fig.1.1.1. Common steps in environmental analysis. . As shown in the above diagram, sample contamination is possible in every steps of an analysis. The most common sources of contamination may originate from: Sample handling Sample containers, equipments Cross-contamination from other samples 1 Chapter 1 Carryover in instruments, glassware Size reduction, dilution, homogenization Syringes, reagents Instrument memory effects, etc., Not only would contamination result in inaccurate data, there are many possible errors throughout the analysis. These include: Uneven sampling Loss of analytes due to evaporation, decomposition, adsorption on sample container Incomplete extraction or concentration Loss of sample due to operator’s mistake Purity of standards and stock preparation Carry over from previous run Variation of instrument response Interference species in the sample, etc., The errors cannot be eliminated completely, although their magnitude and nature can be characterized. Accuracy and precision are the two important parameters to improve the analysis. By minimizing the number of measurement steps and using appropriate techniques (for example, a volume of less than 1 mL can be measured more accurately and precisely with a syringe than with a pipette) also reduce errors in analysis. An excellent sample preparation method must involve the following ‘figures of merit’ [23]; Minimize the analysis errors by following good laboratory practice (GLP) 2 Chapter 1 Ecoefficiency in terms of solvent consumption and waste generation High sample preparation selectivity to distinguish the analyte from the matrices High samples throughput within a given time Ease of automation with common instruments Good accuracy, precision, limits of detection and linear range Reasonable cost of the entire analysis Table.1.1.1. show the common instrumental methods and the necessary sample preparation steps prior to analysis [2]. Table.1.1.1. Common sample preparation analytical methods Analytes Sample Preparation Instruments Organics Extraction, concentration, Cleanup, derivatization Transfer to vapor phase, Concentration Extraction, concentration, speciation Extraction, derivatization, Concentration, speciation GC, HPLC, CE, GC/MS, LC/MS Volatile organics Metals Metals Ions DNA/ RNA Amino acids, fats carbohydrates Microstructures Extraction, concentration, derivatization Cell lysis, extraction, polymerase chain reaction Extraction, cleanup GC, GC-MS AA, GFAA, ICP, ICP/MS UV-VIS molecular absorption Spectrophotometry, Ion chromatography IC, UV-VIS Electrophoresis, UV-VIS, florescence GC, HPLC, CE, electrophoresis Etching, polishing, reactive ion Microscopy, surface techniques, ion bombardments, spectroscopy etc. The major sources of environmental pollutants can be attributed to agriculture, electricity generation, derelict gas works, metalliferous mining and smelting, metallurgical industries, chemical and electronic industries, general urban and industrial 3 Chapter 1 sources, waste disposal, transport and other miscellaneous sources [4-6]. Some important environmental pollutants are shown in Table.1.1.2. Table.1.1.2. Important environmental pollutants 1) Pesticides 2) Aldrin 3) Polycyclic aromatic hydrocarbons 4) Dichlorvos 5) Volatile organic compounds 6) Atrazine 7) Phenols 8) Tributlytin compounds 9) Polychlorinated biphenyls 10) Triphenlytin compounds 11) Dioxins and furans 12) Trifluralin 13) Mercury and cadmium 14) Fenitrothion 15) γ-hexachlorohexane 16) Azinphos-methyl 17) Persistent organics, e.g. DDT 18) Malathion 19) Benzene 20) Endosulfan 22) Hydrocarbons 21) Hexachlorobutadiene Pollution of the environment poses a treat to the health and wealth of living things. Consequently, it is essential to monitor the levels of organic pollutants in the environment. The trace analysis of organic pollutants is complicated and involves many steps. The accuracy and precision of the results of analysis are not only dependent on the analytical instruments used but are also based on factors such as sampling strategy, sample storage, sample pretreatment, sample extraction/ pre-concentration and clean-up. The followings sections briefly describe sample preparations and extraction techniques for environmental solid and aqueous samples. 1.2. Extraction of Organics from Aqueous Liquids Aqueous samples can be subdivided into natural waters and wastewater, biological fluids, milk, alcoholic and soft drinks, etc. 4 Chapter 1 1.2.1. Liquid-liquid extraction The principle of liquid-liquid extraction is based on the fact that the sample is distributed or partitioned between two immiscible solvents in which the analyte and matrix have different solubilities. In an aqueous and an organic phase, an equilibrium can be obtained by shaking the two phases together. Suppose analyte A is in the aqueous phase. The partition can be written as; A (aq) = A (org) (1) where (aq) and (org) are the aqueous and organic phases, respectively. The distribution coefficient Kd between two phases can be represented by; Kd = {A}org / {A}aq (2) The fraction of analyte extracted (E), often expressed as an equation; E = CoVo / (CoVo + CaqVaq) (3) or E = Kd V / (1 + Kd V) (4) where Co and Caq are the concentrations of the analyte in the organic and aqueous phases; Vo and Vaq are the volumes of the organic and aqueous phases, respectively; and V is the phase ratio Vo / Vaq. Typically, two or three repeat extractions are required with fresh organic solvent to achieve quantitative recoveries. The below equation is used to determine the amount of analyte extracted after successive multiple extractions; E = 1 - [1 / (1 + KdV)]n (5) where n = number of extractions. For example, if the volumes of the two phases are the 5 Chapter 1 same (V=1) and Kd = 3 for an analyte, then four extractions (n=4) would be required to achieve >99% recovery. The problem with LLE is that it is very time-consuming, and it uses expensive glassware and toxic solvents. The volume of the extract is usually too large for direct injection for analysis and, in order to obtain sufficient sensitivity, an additional evaporation-concentration step, e.g. using an apparatus (Kuderna-Danish) is necessary. Particular care needs to be taken in both the solvent extraction and concentration procedures to avoid contamination of the sample and formation of emulsions [7-10]. Thus, the demand for miniaturization in analytical chemistry combined with the use of reduced organic solvent and better automation with modern instruments have led to recent developments of miniaturized liquid-liquid extractions procedures. 1.2.2. Flow Injection Analysis Flow injection analysis can be used to minimize the volumes of organic solvent required for LLE, as well as to automate the extraction process. Using this technique, sample and solvent volumes of less than 1 mL can be used. FIA is based on the injection of a liquid sample into a moving, non-segmented continuous carrier stream of a suitable liquid. The injected liquid forms a zone, which is then transported toward a detector. Mixing with the reagent in the flow stream occurs mainly by diffusion-controlled processes, and a chemical reaction occurs. The detector continuously records the absorbance, electrode potential, or other physical parameter as it changes as a result of the passage of the sample material through the flow cell [11-13]. The advantages of FIA are that since all conditions are reproduced, dispersion is very controlled and reproducible. That is, all samples are sequentially processed in 6 Chapter 1 exactly the same way during passage through the analytical channel, or, in other words, what happens to one sample happens in exactly he same way to any other sample. FIA is a general solution-handling technique, applicable to a variety of tasks ranging from pH or conductivity measurement to colorimetric and enzymatic assays. Still, FIA has disadvantages compared to the latest micro-extractions techniques because the volumes of organic solvents used in FIA are still in the order of several milliliters for each analysis [14]. 1.2.3. Liquid-Phase Microextraction The term “liquid phase microextraction” (LPME) was first introduced in 1997 to describe two-phase systems in microscale LLE [15-18] which involves the use of a droplet of organic solvent hanging at the end of a microsyringe needle. This organic microdrop is placed in an aqueous sample, and the analytes present in the aqueous sample are extracted into the organic microdrop. Alternatively, LPME is performed in a three-phase system in which analytes in their neutral form were extracted from aqueous samples, through a thin layer of an organic solvent on the top of the sample, and into an aqueous microdrop at a (different pH from the sample) placed at the tip of a microsyringe [19-20]. Subsequently, the aqueous microdroplet was withdrawn into the syringe which was then transferred an HPLC or CE system for direct analysis. Static and dynamic LPME modes were developed by He and H.K.Lee in 1997 [21-22]. It was these authors also actually called the term “Liquid-phase microextraction”. In static mode (similar to the microdrop approach), the extraction occurrs by mass transfer and diffusion. In dynamic LPME, the organic solvent is 7 Chapter 1 confined within the microsyringe barrel, the extraction of analytes is carried out by moving the microsyringe plunger repeatedly to and from a renewable organic film and plug within the barrel. When the plunger is withdrawn, a solvent film is generated on the inner wall of the syringe. Analytes are extracted from the aqueous sample plug to the organic film, then quickly diffuse into the bulk organic solvent upon expulsion of the aqueous aliquot from the syringe barrel. In general, the dynamic mode produces better enrichment than static LPME. Another type of LPME was developed and also termed solvent microextraction with simultaneous back extraction (SME/BE) which applied unsupported organic liquid membrane held within a Teflon ring to separate the aqueous sample and acceptor phase. After extraction, an aliquot of acceptor phase was directly injected into the HPLC or GC. The higher extraction efficiency can be obtained by increasing the volume ratio between sample solution and acceptor phase in SME/BE [23-24]. LPME has the advantages over LLE as the consumption of organic solvents is dramatically reduced. It produces higher enrichment factor. It is simple, low cost and compatible with the final analytical instrument. Moreover, no solvent evaporation is needed. However, the disadvantages are that LPME based on hanging organic microdrops is not very robust [25], and the latter may be lost from the needle tip of the syringe during extraction. This is especially the case when samples are stirred vigorously to speed up the extraction process. In addition, biological samples, such as plasma, may emulsify substantial amounts of organic solvents, and this may also affect the stability of hanging drops during extraction. Therefore, hollow fiber membrane-protected LPME was developed recently to eliminate the above problems. 8 Chapter 1 1.2.4. Hollow Fiber Membrane-Protected LPME An alternative concept for LPME based on the use of single, low-cost, disposable, and porous, hollow fiber made of polypropylene was introduced recently [26-31]. In this hollow fiber-protected (HFM) LPME device, the extractant solvent is contained within the lumen (channel) of a porous hollow fiber, such that it is not in direct contact with the sample solution. As a result, samples may be stirred or vibrated vigorously without any loss of the solvent during extraction. Thus, hollow fiber-protected LPME is a more robust and reliable alternative for LPME since the solvent is “protected”. In addition, the equipment needed is very simple and inexpensive. Polypropylene was selected for HFMLPME because it is highly compatible with a broad range of organic solvents. In addition, with a pore size of approximately 0.2 µm, polypropylene strongly immobilizes the organic solvents used in LPME. Immobilized organic solvent Acceptor solution Porous hollow fiber membrane Aqueous sample Fig.1.2.4.1. Basic extraction set up in HFM-LPME The acceptor solution may be the same organic solvent as that immobilized in the pores, resulting in extraction of the analyte (A) in a two-phase system in which the analyte is collected in an organic phase; 9 Chapter 1 A sample A acceptor organic phase Two-phase LPME may be applied to most analytes with a solubility in a water immicible organic solvent, that is substantially higher than in an aqueous medium. The acceptor solution in this mode is directly compatible with GC, whereas evaporation of solvent and reconstitution in an aqueous medium is required for HPLC or CE. Alternatively, the acceptor solution may be another aqueous phase providing a three-phase system, in which the analytes (A) are extracted from an aqueous sample, through the thin film of organic solvent impregnated in the pores of the fiber wall, and into an aqueous acceptor solution which generally is set at a different pH from that of the sample solution; A sample A organic phase A acceptor aqueous phase Therefore, the two phase system is more suitable for GC, whereas, three-phase LPME system is suitable for HPLC and CE analysis. Generally, both methods based on diffusion in which extraction is promoted by high partition coefficients. The three-phase system is known as liquid-liquid-liquid microextraction (LLLME). 1.2.5. Purge and Trap or Dynamic Headspace Purge and trap (P&T) is widely used for the extraction of volatile organic compounds from aqueous samples followed by GC. It is also used for solid and gaseous samples. The method involves the introduction of an aqueous sample (typically 5 mL) into a glass sparging vessel. The sample is then purged with high purity nitrogen at a specified flow rate and time. The extracted volatile organics are then transferred to a trap, e.g. Tenax, at ambient temperature. This is followed by the desorption step. In this step, the trap is rapidly heated to desorb the trapped volatile organic compounds in a narrow 10 Chapter 1 band. The desorbed compounds are transferred via a heated transfer line to the injector of a gas chromatograph for separation and detection [32-34]. The advantages of the P&T are its high sensitivity; normally detection of the analytes in the lower ppb range can be achieved. By purging samples at higher temperatures, higher molecular weight compounds can be detected. However, the technique has some disadvantages. It requires more time for sample preparation and cannot normally be automated. In addition, very light volatiles and gases will not be trapped on the adsorbent resins (Tenax) and therefore will be missed in the analysis. Nevertheless, this technique is used in many standard methods approved by the EPA [35]. 1.2.6. Static Headspace Extraction Static headspace extraction is most suited for the analysis of very light volatiles in samples that can be efficiently partitioned into the headspace gas volume from the liquid or solid matrix sample. This technique has been available for over 30 years [36], so the instrumentation is both mature and reliable. The method of extraction is straightforward; solid or liquid sample is placed in a headspace autosampler (HSAS) vial of about 10 mL, and the volatile analytes diffuse into the headspace of the vial. Once the concentration of the analyte in the headspace of the vial reaches equilibrium with the concentration in the sample matrix, a portion of headspace is swept into a gas chromatograph for analysis. However, higher boiling volatiles and semi-volatiles are not detectable with this technique. In addition, the sensitivity of the technique is limited, typically a factor of 1000 time lower than P&T. Multiple headspace extraction (MHE) may also be applied to determine the total amount of analyte in an exhaustive headspace extraction [37-38]. The 11 Chapter 1 advantage to MHE is that sample matrix effects are eliminated since the entire amounts of analytes are examined. 1.2.7. Solid-Phase Extraction In conventional solid-phase extraction (SPE), a liquid sample is passed into a solid or “sorbent” that is packed in a polypropylene cartridge or embedded in a disk. As a result of strong attractive forces between the analytes and the sorbent, the analytes are retained on the sorbent. Later, the sorbent is washed with small volume of a solvent that has ability to disrupt the bonds between the analytes and the sorbent. The final result is that the analytes are concentrated in a relatively small volume of clean solvent and are therefore ready to be analyzed without any additional sample work up [39-40]. In some cases, the extract still has to be concentrated but evaporation to a small volume. The most common goals of an extraction protocol are clean-up, concentratration, and solvent exchange (e.g., aqueous to organic) prior to analysis. SPE achieves these goals in four simple steps as illustrated in figure below. The advantages of SPE are that it is simple, inexpensive, can be used in the field, can be automated with HPLC or GC and uses relatively little solvents. However, it has 1 c o n d i t i o n 2 r e t e n t i o n 3 r i n s i n g 4 e l u t i o n Fig.1.2.7.1. Four basic steps in traditional SPE 12 Chapter 1 disadvantages because of low recovery- resulting from interaction between the sample matrix and analytes, some solvent is still necessary, and usually evaporation of the final eluate is needed. There is also the possible of plugging of the cartridge by solid and oily components. 1.2.8. Solid-Phase Microextraction Arthur and Pawliszyn developed this microscale technique in the late 1980’s [4142]. They introduced it as a solvent-free sample preparation technique that could serve as an alternative to traditional extraction procedures such as LLE, P&T, static headspace, and SPE procedures. SPME preserves all of the advantages of SPE while eliminating the main disadvantages of low analyte recovery, plugging, and solvent use. This technique utilizes a short thin solid rod of fused silica (typically 1 cm long and 0.1um outer diameter), coated with an adsorbent polymer. The coated fused silica (SPME fiber) is attached to a metal rod. The entire assembly (fiber holder) may be described as a modified syringe. In the stand by position, the fiber is withdrawn into a protective sheath. For sampling, a liquid or solid sample is placed in a vial, and the vial is closed with a cap with a septum. The sheath is pushed through the septum and the plunger is lowered, introducing the fiber into the vial, where it is immersed directly into the liquid sample or is held in the headspace. Analytes in the sample are adsorbed on the fiber. After a predetermined time, the fiber is withdrawn into the protective sheath which is then removed from the sampling vial. Immediately after, the sheath is inserted through the septum of a GC injector, the plunger is pushed down, and the fiber is forced into the injector where the analytes are thermally desorbed and separated on the GC column. The 13 Chapter 1 desorption step is usually 1-2 min. After the desorption, the fiber is withdrawn into its protective sheath and the sheath is removed from the GC injector. Modified Syringe Headspace Fiber Sample Heater/ Stirrer Fig.1.2.8.1. Headspace SPME VS Direct SPME There are two approaches to SPME sampling of volatile organics: direct and headspace as shown in Fig.1.2.8.1 [43-44]. In direct sampling, the fiber is placed into the sample matrix, and in headspace sampling, the fiber is placed in the headspace of the sample. In addition, membrane protected SPME sampling is also applied in some works where the fiber is separated from the sample with a selective membrane which lets analytes through while blocking interferences. SPME has been interfaced to HPLC, CE and fourier transform infrared spectroscopy (FTIR) in addition to GC [45-47] and used to extract from a wide variety of sample matrix [48]. Several adsorbent polymers are commercially available on SPME such as polydimethylsiloxane (PDMS). Which is normally used for alkyl benzenes, PAH’s, and volatile halogenated compounds; polyacrylate (PA), or mixture of polyacrylate with Carbowax (CW) and/or 14 Chapter 1 polydivinylbenzene (DVB). The latter is used for alcohols and small polar compounds. It has been established that the fiber can usually be used for 100 times or more. The advantages of SPME techniques are; It is an equilibrium technique and is therefore, selective Time required for analyte to reach an equilibrium between the coated fiber and sample, relatively short Ideal for field sampling: large volume sampling, direct sampling, portable apparatus Solvent-less extraction and injection, eliminating solvent disposal Smooth liquid coating can be used, eliminating the problem of plugging By sampling from headspace, SPME can extract analytes from very complex matrices All analytes collected on the solid phase can be injected into GC for further analysis Method is fast, inexpensive, and easily automated, simple The disadvantages of SPME are; Often only a small fraction of the sample analytes are extracted by the coated fiber Quantification in SPME requires calibration Carryover resulting from incomplete desorption Fiber easily broken Limited number of polymeric coatings for SPME- lack of fibers that are sufficiently polar 15 Chapter 1 1.3. Extraction of Organics from Solid Matrices The extraction and recovery of a solute from a solid matrix can be regarded as a five-stage process: [49] i. the desorption of the compound from the active sites of the matrix ii. diffusion into the matrix itself iii. solubilization of the analyte in the extractant iv. diffusion of the compound in the extractant and v. collection of the extracted solutes In practical environmental applications, the first step is usually the rate-limiting step, as solute–matrix interactions are very difficult to overcome and to predict. As a consequence, the optimization strategy will strongly depend on the nature of the matrix to be extracted. Solid sample includes soils, sediments, fruits, meats, tissue, leaves, etc. Currently available methods for organic environmental analysis are; a) Soxhlet extraction b) Automated Soxhlet extraction, Soxtec c) Pressurized fluid extraction d) Ultrasonic extraction e) Microwave-assisted extraction f) Supercritical fluid extraction g) Direct thermal extraction 1.3.1. Soxhlet and Soxtec Soxhlet is commonly used as the benchmark method for validating and evaluatin other extraction techniques. Soxtec not only reduces the extraction time to 2 to 3 hours as 16 Chapter 1 compares to 60 to 48 hours in Soxhlet but also decreases solvent use from 250 mL to 500 mL per extraction to 40 to 50 mL per extraction. Two to six samples can be extracted simultaneously with a single Soxhtec apparatus [50]. In general, however, solvent consumption is significant. 1.3.2. Pressurized fluid extraction A new technique, pressurized fluid extraction (PFE) appeared around 10 years ago. It is called accelerated solvent extraction (ASE™, which is a Dionex trade mark), pressurized liquid extraction (PLE), pressurized solvent extraction (PSE) or enhanced solvent extraction (ESE). It was partly derives from supercritical fluid extraction (SFE). In PFE, the extractant is maintained in its liquid state. In order to achieve elevated temperatures, pressure is applied inside the extraction cell. In this way, temperatures around 100–200 °C may be attained with classical organic solvents. In fact, at such high temperatures and pressures, the solvent may be considered as being in a subcritical state, with advantageous mass transfer properties. PFE affords the ability to perform fast, efficient extractions due to the use of elevated temperatures, as the decrease in solvent viscosity helps to disrupt the solute– matrix interactions and increases the diffusion coefficients. In addition, the high temperature favours the solubilization of the compounds due to a change in their distribution coefficients. Finally, the pressure favours the penetration of the solvent into the matrix, which again favors extraction. Consequently, this very recent technique is of growing interest, and numerous commercial systems have been sold. PFE has been recognized as an official method by the EPA, and the method has enabled the efficient 17 Chapter 1 screening of soils to be performed for selected semivolatile organic priority pollutants [51-52]. 1.3.3. Ultrasonic extraction Ultrasonic extraction (USE) uses ultrasonic vibration to ensure intimate contact between the sample and the solvent. Sonication is relatively fast, but the extraction efficiency is not as high as some of the other techniques and ultrasonic irradiation may lead to the decomposition of some compound [53]. Therefore, the selected solvent system and the operating conditions must usually be demonstrated to exhibit adequate performance for the target analytes in reference samples before it is implemented for the real samples. The most common solvent system is acetone-hexane (1:1 v/v) but for nonpolar analytes such as PCBs, hexane alone can also be used. 1.3.4. Microwave-assisted extraction Microwave-assisted extraction (MAE) uses microwave radiation as the source of heating of the solvent–sample mixture. Due to the particular effects of microwaves on matter (namely dipole rotation and ionic conductance), heating with microwaves is instantaneous and occurs in the middle of the sample, leading to very fast extractions [5455]. In most application, the extraction solvent is selected as the medium to absorb microwaves. Alternatively (for thermolabile compounds), the microwaves may be absorbed only by the matrix, resulting in heating of the sample and release of the solutes into the cold solvent. Microwave energy may be applied to samples in two ways: either in closed vessels (under controlled pressure and temperature), or in open vessels (at atmospheric pressure) [56-57]. These two technologies are commonly named pressurized MAE or 18 Chapter 1 focused MAE, respectively. Whereas in open vessels the temperature is limited by the boiling point of the solvent, at atmospheric pressure, in closed vessels, the temperature may be elevated by simply applying the appropriate pressure. 1.3.5. Supercritical fluid extraction Supercritical fluid extraction (SFE) is also a very popular technique for environmental analysis. It is an appropriate technique for the analysis of the less volatile compounds, much like solvent extraction. It has limitations for the range of analytes that can be extracted simultaneously. However, for a particular semi-volatile analyte or a narrow selection of analytes, this technique is preferable over solvent extraction. This technique can be automated which also makes it advantageous in many instances [58]. 1.3.6. Direct thermal extraction Direct thermal extraction (DTE) is a new technique, which is unique to Scientific Instrument Services, Inc (SIS), [59]. In DTE, volatiles and semi-volatiles can be thermally extracted directly from solid matrix samples without the use of any solvents or any other sample preparation. The advantages of this technique are that a wide range of volatiles and semi-volatiles can be analyzed and the high sensitivity of the technique (typically ppb ranges on samples less than 1.0 gram). Its main disadvantage is the extraction of water into the GC column which will form an ice plug. Since no sample preparation is required, the sampling time is small, just weigh the sample into the desorption tube and analyze it and the DTE extraction technique is more sensitive by at least a factor 10 to 100 than P&T [60]. This table below compares advantages and disadvantages among all the techniques discussed. 19 Chapter 1 Table1.3.1. Advantages and disadvantages of various techniques Technique Soxhlet Advantages Disadvantages Not matrix dependent Slow (up to 24-48 hrs) Inexpensive equipment Large amount of solvent (500 mL) Unattended operation Mandatory evaporation of extract Rugged, benchmark method Filtration not required Soxtec Not matrix dependent Relatively slow (2 hrs) Inexpensive equipment Less solvent (50 mL) Evaporation integrated Filtration not required USE SFE Not matrix dependent Large amount of solvent (300 mL) Inexpensive equipment Mandatory evaporation of extract Fast (10-45 min) Labor intensive Large amount of sample (2-30 g) Filtration required Fast (30-75 min) Matrix dependent Minimal solvent use (5-10 mL) Small sample size (2-10 g) CO2 is environmentally friendly Expensive equipment Controlled selectivity Limited applicability Filtration not required Evaporation not needed ASE Fast (12-18 min) Expensive equipment Small amount of solvent (30 mL) Cleanup necessary Large amount of sample (100 g) Automated Easy to use Filtration not required MAE Fast (10-30 min) Polar solvent needed 20 Chapter 1 Technique Advantages Disadvantages High sample throughput Cleanup mandatory Small amount of solvent (30 mL) Filtration required Large amount of sample (20 g) Expensive equipment Degradation possible DTE Very fast Form ice plug at GC column No solvent needed Small sample size ( 1-5 g) High sensitivity Expensive instrument Automated Limited applicability 1.4. Chromatography in Environmental Analysis Due to the excellent separation characteristics and versatility of chromatographic methods, all types of substances, from the small hydrogen and helium molecules to large and complex protein molecules, can be separated by chromatography which have gained growing acceptance and application for residue analysis in air, ground and surface waters, soil matrices, foods and food products and in human and veterinary health care. There are no two compounds, however similar in structure (even optical isomers), which cannot be separated by one chromatographic technique or another. The study of chromatography is too diverse and multi-faceted to be adequately presented by a single work but hundreds of [61]. For environmental analysis, HPLC and GC are the most popular techniques because of their high resolution, excellent sensitivity, faster sample throughput and user friendliness. HPLC VS GC 21 Chapter 1 Compared with older chromatographic methods, GC provides separations that are faster and better in terms of resolution. It can be used to analyze a variety of samples. However, GC simply cannot handle many samples without derivatization, because the samples are not volatile enough and cannot move through the column because they are thermally unstable and decompose under the conditions of separations. According to estimates, GC can sufficiently separate only 20% of known organic compounds without prior chemical alteration of the sample. An important advantage of HPLC over GC is that it is not restricted by sample volatility or thermal stability. It is also ideally suitable for the separation of macromolecules and ionic species of biomedical interest, labile natural products, and less stable and/or high molecular weight compounds. 1.5. Scope of This Study This thesis encompasses three sections. The first section discusses a study of the suitability of ionic-liquid supported HFM-protected LLLME as a single-step enrichment/clean-up approach, eliminating matrix effects normally encountered by other immersion-based microextraction techniques. In the second section, the development of micro-solid phase extraction (µSPE), a novel procedure, which is simple, rapid, costeffective, highly sensitive and selective for the determination of polar carbamate pesticides in tea sample is described. In this procedure, porous polypropylene membrane is used as a protective sheath for the adsorbent material for extracting from dirty matrices. Finally, in the third section, we discuss the application of a new polymeric material for SPME. The sorbent is evaluated for the extraction and preconcentration of 22 Chapter 1 organochlorine pesticides, organophosphorous compounds and polycyclic aromatic hydrocarbon analytes in environmental water samples, combined with GC-MS. References: [1] J. R. Dean, Extraction Methods for Environmental Analysis, John Wiley, New York, 1998 [2] S. Mitra, Sample Preparation Techniques in Analytical Chemistry, Hoboken, NJ, USA, 2003 [3] A. J. Handley, Extraction Methods in Organic Analysis, Sheffield Academic Press, Sheffield, 1999 [4] T. Cserhati, E. Forgacs, Chromatography in Environmental Protection, Harwood Academic Publishers, Amsterdam, 2001 [5] R. L. Grob, Chromatographic Analysis of the Environment, M. Dekker, New York, 1975 [6] F. W. Fifield, P. J. Haines, Environmental Analytical Chemistry, Blackie Academic & Professional, London, 1995 [7] W. Kleibohmer, Handbook of Analytical Separations; 3, Environmental Analysis, Elsevier, New York, 2001 [8] E. Psillakis, N. Kalogerakis, Trends Anal. Chem. Elsevier. 22 (2003) 10 [9] N. Alizadeh, S. Salimi, A. Jabbari, Anal. Sci. 18 (2002) 307 [10] K. E. Rasmussen, S. Pedersen-Bjergaard, Trends Anal. Chem. Elsevier. 23 (2004) 1 [11] B. Karlberb, S. Thelander, Anal. Chim. Acta. 98 (1978) 1 [12] F. H. Bergamin, J. X. Medi, B. F. Reis, E. A. Zagatto, Anal. Chim. Acta. 101 (1998) 9 23 Chapter 1 [13] R. Jaromir, Flow Injection Analysis, Wiley Inter Science, USA, 1998 [14] H. Liu, P. K. Dasgupta, Anal. Chem. 68 (1996) 1817 [15] M.A. Jeannot, F. Cantwell, Anal. Chem. 68 (1996) 2236 [16] H. Liu, P.K. Dasgupta, Anal. Chem. 68 (1996) 1817 [17] M.A. Jeannot, F. Cantwell, Anal. Chem. 69 (1997) 235 [18] L. Zhao, H.K. Lee, J. Chromatogr. A. 919 (2001) 381 [19] M. Ma, F. Cantwell, Anal. Chem. 70 (1998) 3912 [20] M. Ma, F. Cantwell, Anal. Chem. 71(1999) 388 [21] Y. He, H. K. Lee, Anal. Chem. 69 (1997) 4634 [22] Y. Wang, Y. C. Kwok, Y. He, H. K. Lee, Anal. Chem. 70 (1998) 4610 [23] M. Ma, F. F. Cantwell, Anal. Chem. 70 (1998) 3912 [24] M. Ma, F. F. Cantwell, Anal. Chem. 70 (1999) 388 [25] K.E. Kramer, A.R.J. Andrews, J. Chromatogr. B 760 (2001) 27 [26] S. Pedersen-Bjergaard, K.E. Rasmussen, Anal. Chem. 71 (1999) 2650 [27] S. Pedersen-Bjergaard, K.E. Rasmussen, Electrophoresis. 21 (2000) 579 [28] T.G. Halvorsen, S. Pedersen-Bjergaard, K.E. Rasmussen, J. Chromatogr. B 760 (2001) 219 [29] L. Zhu, L. Zhu, H.K. Lee, J. Chromatogr. A 924 (2001) 407 [30] G. Shen, H.K. Lee, Anal. Chem. 74 (2002) 648 [31] C. Basheer, H.K. Lee, J.P. Obbard, J. Chromatogr. A 968 (2002) 191 [32] S.M. Abel, A.K. Vickers, D. Decker, J. Chromatogr. Sci. 32 (1994) 328 [33] I. Silgoner, E. Rosenberb, M. Grasserbauer, J. Chromatogr. A. 768 (1997) 259 [34] Z. Bogdan, J. High Resolut. Chromatogr. 20 (1997) 482 [35] EPA Methods for Determination of Organic Compounds in Drinking Water, U.S. 24 Chapter 1 Environmental Protection Agency, Cincinnati, Ohio, 1995 [36] H. Hachenberb, A. P. Schmidt, GC Headspace Analysis, Heyden, London, 1977 [37] C. McAuliffe, Chem Technol. 46 (1971) 8 [38] M. Suzuki, S. Tsuge, and T. Takeuchi, Anal. Chem. 42 (1970) 1705 [39] Thurman, E. M; Mills, M. S. Solid Phase Extraction: Principle and Practice, John Wiley and Sons, New York, 1998 [40] J. I. Fritz, Analytical Solid Phase Extraction; John Wiley and Sons, New York, 1999 [41] C. Arthur and J. Pawliszyn, Anal. Chem. 62 (1990) 2145 [42] R. Berlardi and J. Pawliszyn, Water Pollut. Res. J. Can. 24 (1989) 179 [43] Z. Zhang and J. Pawliszyn, Anal. Chem. 65 (1993) 1843 [44] B. Page and G. Lacroix, J. Chromatogr. 648 (1993) 199 [45] J. Chen and J.Pawliszyn, Anal. Chem., 67 (1995) 2350 [46] J. Pawliszyn, Solid Phase Microextraction, Theory and Practice, J. Wiley and Sons, New York, 1997 [47] J. Burck, in Ref. 48, pp. 638-653 [48] SPME Application Guide, Supelco, Bellefonte, PA, USA, 2001 [49] J. Pawliszyn, J. Chromatogr. Sci. 31 (1993) 31 [50] EPA Method 3540C, Soxhlet Extraction, Test Methods for Evaluating Solid Waste, EPA, Washington DC, 1996 [51] EPA Method 3545A, Pressurized Fluid Extraction, Test Methods for Evaluating Solid Waste, EPA, Washington DC, 1998 [52] J. A. Fisher, M. J. Scarlett and A. D. Stott, Environ. Sci. Technol. 31 (1997)1120 [53] A. Kotronarou, Environ. Sci. Technol. 26 (1992) 1460 25 Chapter 1 [54] J. R. J. Pare, J. M. R. Belanger and S. S. Stafford, Trends Anal. Chem. 13 (1994) 176 [55] C. S. Eskilsson, E. Bjorklund, J. Chromatogr. A 902 (2000) 227 [56] V. Camel, Trends Anal. Chem.19 (2000) 229 [57] M. Letellier, H. Budzinski, Analusis, 27 (1999) 259 [58] R. M. Smith, J. Chromatogr. A 856 (1999) 83 [59] J. J. Manura, S. Overton, DTE Application Note, Scientific Instrument Services, Ringoes, NJ, USA, 1999 [60] A. Hoffmann, DTE Application Note, Gerstel GmbH & Co.KG, Germany, 1996 [61] T. Cserhati, E. Forgacs, Chromatography in Environmental Protection, Hungarian Academy of Sciences , Budapest, Hungary, 200 26 Chapter 2. Room temperature ionic-liquid as solvent in hollow fiberprotected liquid-liquid-liquid microextraction technique coupled with high performance liquid chromatography 2.1. Introduction Alkylphenols are used in the production of surfactants in a wide variety of industrial, agricultural and household applications [1]. The primary concern about these compounds is that their estrogenic properties have been demonstrated in in-vitro and invivo studies [2]. They function by being able to displace estradiol from the estrogen receptor. They are present in very low concentrations in the aquatic environment; therefore efficient sample preparation techniques to preconcentrate them before analysis are need. Recently, liquid-phase microextraction (LPME) a miniaturised approach to liquid-liquid extraction (LLE) has been introduced [3, 4]. LPME through the use of a single drop of solvent [5, 6] or a short plug of solvent held within a porous hollow fiber membrane (HFM) [7], has been emerging as attractive extraction approaches in environmental and other analyses. In two-phase LPME [8-11], the analytes are extracted from an aqueous sample matrix into an organic acceptor phase; this type of extraction is similar conceptually to LLE. Three-phase LLLME [12-15] is more suitable for watersoluble polar compounds and involves extraction of such analytes from an aqueous sample, through an organic immiscible phase impregnated in the pores of the HFM, and further extracted into an aqueous phase held inside the channel of the HFM. This process is similar to LLE with back extraction. Substantial sample cleanup can occur in both HFM-protected LPME and 27 Chapter 2 LLLME techniques [8-15], since the membrane prevents extraneous materials in the sample from interfering with the extraction. Room temperature ionic-liquids are waterand air-stable salts that consist of an organic cation and either an organic or an inorganic anion [16]. As they are non-organic, and water-immiscible, relatively volatile, and are able to solvate a variety of organic and inorganic species, they are being promoted as alternative environmentally friendly solvent [16]. Recently a number of reports in the literature have appeared on the applications of ionic-liquids in separation and analysis, including their being used as running electrolytes in capillary electrophoresis [17-19] and additives in HPLC [20, 21]. Poole and co-workers [22] studied the use of ethylammonium nitrate and propylammonium nitrate in HPLC. Armstrong and coworkers [23-25] have also evaluated ionic-liquids as GC stationary phases. Recently, ionic-liquid based single drop-LPME technique has been successfully demonstrated for the extraction of polycyclic aromatic hydrocarbons [26], alkylphenols [27] and chloroanilines [28]. Semi and non-volatile compounds in complex samples have also been extracted using headspace single drop-LPME [26, 28]. Generally, headspace extraction procedures are less sensitive than the direct immersion approach [29]. Moreover, the sensitivity and precision using single drop-LPME methods could be improved. One reason is the prolonged extraction times and fast stirring rates that result in drop dissolution [30]. Direct immersion using single drop-LPME is not a desirable choice for complex or “dirty” samples such as wastewater. The use of polypropylene HFM as protective sleeves for LPME provides for very efficient sample cleanup for a wide range of complex samples [31, 32]. This present work demonstrates the suitability of ionic-liquid in HFM-protected LLLME as a single step enrichment/clean-up technique, 28 Chapter 2 which could allow the extraction of alkylphenols from wastewater samples, thereby eliminating matrix effects normally encountered by other immersion-based microextraction techniques. We have tested four different room temperature ionic-liquids (IL) in this work. Most of the ionic-liquids are not suitable for the work described because of their very high viscosity. Therefore, two ionic-liquids are mixed with acetonitrile (ACN) to reduce their viscosity. This is the first time such a microextraction approach has been reported, to the best of our knowledge. Parameters affecting the extraction efficiency (such as, the most suitable ionic-liquid, the dilution ratio of acetonitrile and ionic-liquids, extraction time, salting-out effect and sample pH) were studied. 2.2. Experimental 2.2.1. Chemicals and reagents Four different room temperature ionic-liquids (>98% purity); 1-butyl-3methylimidadolium tetrafluoroborate ([BMIM][OcSO4]), phosphate ([BMIM][BF4]), and ([BMIM][PO4]), 1-butyl-3-methylimidadolium 1-butyl-3-methylimidadolium 1-butyl-3-methylimidazolium octylsulfate hexafluorophosphate ([BMIM][PF6]) were purchased from Strem Chemicals (Newburyport, MA, USA). Alkylphenols were obtained from Fluka (Buchs, Switzerland). HPLC-grade solvents were purchased from Fisher Scientific (Fair Lawn, NJ, USA). Ultrapure water was produced on a Milli-Q system (Millipore, Milford, MA, USA). Stock standard mixtures of 1 mg ml-1 of each phenol were prepared by dissolving in methanol and stored at 4oC. Dilute working solution containing a mixture of 10 µg ml-1 of each phenol was prepared in methanol from the stock solutions. 29 Chapter 2 2.2.2 Materials A 50-ml glass vial (Supelco, Bellafonte, PA, USA) was used as the sample receptacle for LLLME experiments. A Heidolph (Kelheim, Germany) magnetic stirrer and a stirring bar measuring 10 mm×3 mm were used to agitate the samples during extraction. Q3/2 Accurel polypropylene HFM (600 µm inner diameter (I.D), 200 µm wall thickness and 0.2 µm wall pore size) was purchased from Membrana (Wuppertal, Germany). For each extraction, a 5.5-cm length of HFM was used for extraction and used in conjugation with a 50-µl HPLC microsyringe (0.8 mm O.D) purchased from Hamilton (Reno, NV, USA). 2.2.3. Wastewater samples Domestic wastewater samples were collected at five different locations in a township, transported to the laboratory in pre-cleaned glass bottles, and stored at -4°C. Unfiltered samples were used for experiments. The original sample pH was 6.6 and no other physical characteristics were measured. 2.2.4. HPLC The HPLC system used consisted of a Waters (Milford, MA, USA) 600E quaternary pump and a Waters M486 UV detector. Data collection and integration were accomplished using a Compaq computer with Empower Software. The reverse phase Spherisorb Spheris column (200× 4.6 mm × 5 µm) of ODS 2 packing material was from PhaseSep (Deeside, UK). The flow rate was 1 ml min-1 and the detection wavelength was set at 280 nm. An isocratic mobile phase composition of 65:35 acetonitrile:water was used for separations. 2.2.5. Ionic-liquid based LLLME 30 Chapter 2 The schematic of the LLLME experimental setup is shown in Figure 2.1. Extractions were performed according to the following procedure: a 50-ml wastewater sample (ionic strength and sample pH were not adjusted) was transferred to the 50-ml vial and a stirring bar was placed in it. Then, 25 µl of the ionic-liquid (the acceptor phase) in acetonitrile (ACN) (1:1) was drawn into a syringe. A 5.5-cm hollow fiber was inserted into the syringe and the ionic liquid was introduced into it. The fiber was then immersed in n-nonane for 10 s in order for the solvent to impregnate the pores of the fiber wall. After impregnation, the fiber (together with the syringe) was immersed in the sample (donor) solution. Samples were stirred at 73 rad s-1 (700 rpm; 1 rpm = 0.1047 rad s-1) for 50 min. After extraction, the syringe–fiber assembly was removed from sample. 25 µl of the acceptor solution was withdrawn from the fiber and then the HFM was discarded. 20 µl of the extract was injected into a 20-µl sample loop of the HPLC injector. Figure 2.1. Schematic of ionic-liquid LLLME experimental setup 31 Chapter 2 2.3. Results and discussion The design of an experiment is very important for the method development of microextraction techniques. The following represents the advantages of our modified configuration as shown in Figure 2.2. Plunger Clamp Plunger Clamp Syringe Syringe Holder with clamp Holder with clamp Aqueous Sample Aqueous Sample Hollow Fiber Hollow Fiber Strring Bar Strring Bar Magnetic Stirrer Magnetic Stirrer B A Figure 2.2. Different HFM-LPME/ LLLME sets up The length of the hollow fiber is up to 10 cm in set up A, which contains an acceptor solution of at least 25 µL, so that acceptor phase has more surface area to contact with sample solution and sensitivity is increased. The other end of hollow fiber is sealed in set up B, where as the new design is not sealed, so that when the plunger is pressed, some air inside the syringe may be passed through the pore of the hollow fiber and it will cause air bubble formation. The set up A does not face this problem. In some works, the other end of hollow fiber is not sealed for set up B, so that the acceptor solution can be easily reached to the sample solution and if the density of the acceptor solution is higher, the problem may be even worse. 32 Chapter 2 In the set up A the acceptor solution can be pushed in to the HFM without effecting the wall of HFM since the horizontal level of both ends are the same and other end of the hollow fiber is not sealed. Only need one syringe needle and any normal sample bottle can be used. It is more suitable for HPLC and CE than GC. Our initial studies showed that HFM-protected two-phase LPME with an acceptor phase, (BMIM[PF6]: ACN, 1:1), showed poorer (2 x time lower) analyte enrichment than the three-phase LLLME with the same acceptor phase. The type of solvent immobilized within the pores of the hollow fiber is important in order to obtain satisfactory enrichment factor (the ratio between the equilibrium analyte concentration in the acceptor phase and the initial concentration in the sample solution). Several parameters for the selection of the immobilized solvent were considered: (i) it should be easily retained in the hollow fiber pores, and be non-volatile, (ii) it should be immiscible with water because it serves as an intermediary between the aqueous donor and the aqueous acceptor phases and (iii) the solubility of analytes in the solvent should be higher than that in the donor phase and lower than that in the acceptor phase. Based on the above considerations six organic solvents, namely ethylacetate, dichloromethane, toluene, 1-octanol, isooctane and n-nonane were investigated for their effect on enrichment. Isooctane and n-nonane gave better analyte enrichment than the rest of the solvents. n-nonane was considered to be the best solvent and was therefore used for subsequent experiments. Other conditions that affect the extraction efficiencies such as the most suitable ionic-liquid, sample pH, salt addition and extraction time were evaluated. 33 Chapter 2 The principle of ionic-liquid supported LLLME is similar to solvent/or aqueous accepter phase LLLME procedure. In the three-phase LLLME sampling mode, analyte i is extracted from an aqueous solution (donor phase) through the organic solvent immobilised in the pores of the HFM (organic phase) and further into extraction solvent (acceptor phase) present within the channel of the HFM. Overall, the three-phase LLLME extraction process for analyte i may be illustrated as follows: id ↔ d iorg ↔ ia refers to the donor phase, org to the organic phase and a to the acceptor phase The enrichment factor EF, defined as the ratio Ca,eq/Cd,initial, (Ca,eq = concentration of analyte in the acceptor phase at equilibrium; Cd,initial initial concentration of analyte in the donor phase) and can be calculated as [4]; EF = 1/(K2/K1)+(K2Vorg/Vd)+(Va/Vd) where as K= distribution coefficient, V= volume, C= concentration Since Vorg is very small, then the above equation can be simplified into; EF = 1/ (1/K)+(Va/Vd) Where K= K1/K2 = Ca,eq/ C d,eq (eq = equilibrium) It is obvious that decreasing the volume ratio of the acceptor and the donor phases can increase in extraction efficiency. The first step in the development of the present LLLME method is the selection of a suitable ionic-liquid. Four different ionic liquids (BMIM[BF4], BMIM[PF6] BMIM[PO4] and BMIM[OcSO4]) were initially evaluated for the extraction of alkylphenols in spiked ultrapure water samples under identical extraction conditions. 34 Chapter 2 BMIM[BF4] and BMIM[PF6] gave higher enrichment values than BMIM[PO4] and BMIM[OcSO4]. The viscosities of BMIM[PO4] and BMIM[PF6] were high, however, and drawing them into the syringe was problematical; they were therefore, necessarily, diluted with ACN. The other two ionic-liquids have lower viscosities and could be directly injected into the HPLC system. Figure 2.3 clearly shows that BMIM[PF6], one of the two viscous ionic-liquids (in combination with ACN, 1:1) gave higher analyte enrichment than the rest of the ionic-liquids, and was thus chosen for further experiments. (EF) 160 4-tert-butylphenol 4-tert-octylphenol 120 4-n-octylphenol 4-n-nonylphenol 80 40 0 BMIM[PO4]+ ACN(1:1) BMIM[BF4] BMIM[OcSO4] BMIM[PF6]+ ACN(1:1) EF= Enrichment Factor (-fold) Figure 2.3. Extraction efficiency of various ionic-liquids in HFM-LLLME. Samples spiked at 25 µg l-1 of each analyte and 50 min extraction time. As mentioned above, BMIM[PF6], selected as the extraction solvent, has a higher viscosity than typical organic solvents. To avoid interferences with the target analytes, ACN was used as diluent since it was already being used as part of the mobile phase. BMIM[PF6] was diluted with different amounts of ACN. Table 1 shows the extraction efficiency of various ionic-liquid/ACN mixtures. Dilution of BMIM[PF6] with ACN reduces the viscosity, which increases the dielectric constant of the co-solvent (ACN) 35 Chapter 2 [33]. The viscosity of ionic-liquid is essentially determined by its tendency to form hydrogen bonds and by the strength of Van der Waals interactions. This could be due to the delocalization of the charge over the anion and this seems to be favored by lower viscosity, by weakening hydrogen bonding with the cation and increasing the interaction with alkylphenols [34]. Table 1 shows that BMIM[PF6] diluted with ACN at 1:1 ratio gave higher extraction efficiency than mixtures of other ratios, and thus BMIM[PF6]:ACN (1:1) was used for further experiments. Table 2.1. Effect of dilution of BMIM[PF6] on, and suitability of n-nonane for HFMLLLME: Enrichment factor. Analyte Enrichment factor (-fold) IL:ACN 4-tert-butylphenol 4-tert-octylphenol 4-n-octylphenol 4-n-nonylphenol HFM-LLLME IL:ACN IL:ACN HFM-LPME IL:ACN 2:1 1:1 1:2 1:1 125 110 93 82 146 120 102 87 83 89 91 71 96 83 60 68 IL = ionic-liquid (BMIM[PF6]) ACN = acetonitrile A series of extraction times from 10 to 60 min was investigated extracting water with spiked at a concentration of 25 µg l-1 of individual analytes. For all target analytes, the amount extracted increased with increasing extraction time from 10 to 50 min (Figure 2.4). After 50 min, the enrichment factor decreased slightly. After reaching equilibrium, the analyte has the tendency to be extracted back from the extraction solvent (Le Chatlier’s principle), resulting in enrichment factor reduction after 50 min. 50 min, therefore, appeared to be the optimum extraction time. 36 Chapter 2 The effect of pH in the range from 2 to 12 was investigated. Changes in extraction efficiency with varying pH are shown in Figure 2.5. Samples pH at 7 gave higher analyte enrichment than either strongly acidic or basic conditions. For convenience, no Enrichment factor (-fold) adjustment of pH of wastewater is (pH 6.6) was made before extraction. 180 160 140 120 100 80 60 40 20 0 4-tert-butylphenol 4-tert-octylphenol 4-n-octylphenol 4-n-nonyl phenol 0 20 40 60 80 Extraction time (min) Figure 2.4. Ionic-liquid HFM-LLLME extraction time profile of alkylphenols. Samples spiked at 25 µg l-1 of each analyte. IL:ACN (1:1) as acceptor phase and n-nonane as immobilized solvent. 4-tert-butylphenol 200 Enrichment factor (-fold) 4-tert-octylphenol 4-n-octylphenol 160 4-n-nonylphenol 120 80 40 0 0 2 4 6 8 10 12 14 Sample pH Figure 2.5. Influence of sample pH. Extraction conditions are same as Figure 2.4. 37 Chapter 2 The salting-out effect has been used commonly in LLE and LPME. In LLE, addition of sodium chloride (NaCl) can decrease the solubility of analytes in the aqueous sample and consequently increase their hydrophobicity [35]. This is due to the salting-out effect where fewer water molecules are available for dissolving the analyte molecules, preferably forming hydration spheres around the salt ions [36]. A series of experiments were carried out in which the aqueous samples contained different amounts of NaCl [(5%, 10%, 15%, 20% and 30%) (w/v)] and extraction for them evaluated. The results show addition of 5-20% (w/v) NaCl increased the peak area of 4-tert-butylphenol but showed a decrease for the other three analytes in the study (data not shown). Moreover, addition of 30% NaCl appeared to be no significant increases in extraction efficiency for all the test phenols. This could be due to the increase in the viscosity of the sample solution, which then reduced the mass transfer of the analytes to BMIM[PF6]:ACN. 2.4. Method performance The optimized hollow fiber protected ionic-liquid supported LLLME procedure proved to be simple and effective for the extraction of the alkylphenols. Calibration was performed with five samples of water, each spiked with analyte concentrations ranging from 5 to 100 µg l-1. The correlation coefficient (r) values ranged between 0.9723 and 0.9948 (see Table 2). Inter-day precision was studied for 10 µg l-1 spiked water samples with six replicates and the relative standard deviation RSD ranged from 0.3% to 5.9%. Intra-day precision was carried out on experiments done on three consecutive days at the same concentration levels with six replicates. As can be seen from Table 2.2, the intraday precision for the analysis were in the range of 5.6 and 13.2%. Limits of detection (LODs) were calculated by progressively decreasing the analyte concentration in the 38 Chapter 2 spiked sample until HPLC signals were clearly discerned at S/N=3 at the final lowest concentration. LODs varied between 0.05 and 0.26 µg l-1 for spiked ultrapure water and spiked 0.06 and 0.35 µg l-1 for wastewater samples, respectively. By comparing peak areas in the chromatograms, it can be seen that most of the target compounds were preconcentrated with an enrichment factor of more than 100-fold in the acceptor solution. Table 2.2. Enrichment factor, linearity, and reproducibility for extraction of alkylphenols by the proposed BMIM[PF6]:ACN(1:1) HFM-LLLME method Enrichment Factor (EF) Interday %,RSDs n=6 Intraday %,RSDs n=6 Correlation Coefficient (r) LODa (ng ml-1) LODb (ng ml-1) LOD* (ng ml-1) 4-tertbutylphenol 163 0.3 5.6 0.9862 0.05 0.08 - 4-tertoctylphenol 145 5.9 8.9 0.9834 0.06 0.06 0.7 4-noctylphenol 128 4.1 9.1 0.9723 0.10 0.15 - 4-nnonylphenol 101 0.7 13.2 0.9948 0.26 0.35 0.3 Analytes a = ultrapure water = wastewater * = ref [27], ionic-liquid based single drop LPME with fluorescence detection b Five different wastewater samples (from different sites) were extracted under the optimized extraction conditions. Concentrations of alkylphenols detected in the real samples are shown in Table 2.3. The range was form ‘not detected’ to 4.2 µg l-1. Common components of wastewater sample, such as humic acids and inorganic salts, could reduce the applicability of the method in analysis by affecting the recovery. 39 Chapter 2 Therefore, to assess the matrix effects, spiked wastewater samples were extracted using present procedure, and recoveries were calculated by the standard addition method. Table 2.3. Concentrations of alkylphenols detected in the wastewater samples collected in Singapore Concentration in µg l-1 (n=2) Analyte 4-tert-butylphenol 4-tert-octylphenol 4-n-octylphenol 4-n-nonylphenol site 1 site 2 site 3 site 4 site 5 2.0 2.1 1.9 2.6 1.9 1.6 1.6 3.4 2.8 3.6 2.3 3.2 2.3 2.7 1.9 nd 3.1 1.9 2.5 4.2 Table 2.4. Extraction recoveries obtained by BMIM[PF6]:ACN(1:1)-supported HFMLLLME of wastewater spiked samples (n=3) Analyte % Relative recoveries (n=3)* 4-tert-butylphenol 4-tert-octylphenol 4-n-octylphenol 4-n-nonylphenol spiked at 5 µg l-1 RSDs (%) spiked at 10 µg l-1 RSDs (%) 89 87 102 97 5.5 9.2 7.0 10.3 94 99 85 90 6.1 8.9 3.9 6.6 *Recoveries calculated by standard addition method Extracted chromatograms of real wastewater and spiked wastewater samples at 5 µg l-1 and 25 µg l-1 of each analyte are shown in Figure 2.6. There was a persistent interfering ionic-liquid peak (at 2.5 min) since 1 µL of pure ionic liquid, (1 µL of BMIM[PF6] can be carefully drawn by the syringe but 25 µL of it was impossible to draw) was directly injected into HPLC for identification at the beginning of the experiment. Fortunately, its retention time did not coincide with those of the alkylphenols 40 Chapter 2 in the study. The HFM afforded some selectivity, in that, the porous wall allowed a certain degree of clean-up. Humic acids typically have molecular masses up to several million daltons and thus cannot be usually extracted by the organic solvent, probably because they cannot pass through the HFM [35]. Therefore, cleaner chromatograms were obtained (see Figure 2.6). 0.0045 2 AU 1 3 4 (a) 0.0025 (b) 0.0000 (c) -0.0005 2.00 6.00 12.00 16.00 20.00 Minutes Figure 2.6. BMIM[PF6]:ACN(1:1), HFM-LLLME-HPLC-UV chromatograms of wastewater extract. (a) Extract spiked at 25 µg l-1 of each phenol; (b) extract spiked at 5 µg l-1 of each phenol; (c) extract of real unspiked wastewater sample. Peaks: (1) 4-tertbutylphenol, (2) 4-tert-octylphenol, (3) 4-n-octylphenol and (4) 4-n-nonylphenol. Furthermore, the relative recovery of the extraction procedure, determined as the ratio of the concentrations found in real wastewater and ultrapure water samples spiked at the same concentration level was also evaluated under the optimised experimental conditions. Three replicate runs of wastewater samples are two different spiked 41 Chapter 2 concentrations (5 and 10 µg l-1 of each analyte, respectively) were analysed and the percentage of extracted analytes was then calculated. The recoveries of the analytes from this wastewater were higher than 85% compared with that of spiked ultrapure water. This implies that the proposed method is more precise and the wastewater matrix did not have a significant effect on the extraction efficiency. 2.5. Conclusion The present work evaluated the feasibility of using an ionic-liquid as acceptor phase in hollow fiber protected liquid-liquid-liquid microextraction for extracting alkylphenols from wastewater samples, with analyzed by HPLC. Since some of the ionicliquids were viscous, they had to be mixed with acetonitrile to facilitate the extraction. 1butyl-3-methylimidazolium hexafluorophosphate mixed with acetonitrile (1:1) was found to be the optimum extraction solvent. The proposed method was simple and the use of disposable HFM completely eliminated the carryover effects. Very effective sample clean-up and high analyte enrichment factor could be achieved. The proposed method possessed high sensitivity, with LODs obtained in this study being lower than previously reported ionic-liquid-based single drop-liquid phase microextraction. Moreover, matrix effects were not a significant factor with insignificant influence of sample matrix effect Recoveries of 85–102% were achieved . The present method was rapid and easy to conduct as BMIM[PF6]:ACN extract was compatible with HPLC, the extract could be injected directly for analysis. Combination of ionic-liquids and hollow fiber membrane proved to be an excellent extraction technique. The most important task in future work will be to immobilize the pores of hollow fiber membrane with ionic-liquid and two-phases HFM- 42 Chapter 2 LPME could be applied. It will be a difficult task since ionic-liquids may not penetrate into HFM in normal condition. HFM may be needed chemical treatments such as soaking in de-ionized water for several hours followed by vacuum heating of HFM at higher temperature to clean and dry. The dry membranes are then swollen with a mixture of an ionic-liquid and a volatile organic solvent (e.g. acetonitrile or methanol) that is highly miscible with ionic-liquid. The organic solvent will serve as swelling agent and it can be easily removed by evaporation after immobilization process. Ionic-liquids may also be coated to the organic polymer by the above processes and could be applied in solid-phase extraction or solid-phase microextraction techniques. Moreover, ionic-liquids could remove contaminants in petroleum productions. Chiral ionic-liquids may separate the optically active compounds from natural or pharmaceutical products. Carbon dioxide, which causes global warming, is highly soluble in ionic-liquids. Therefore, ionic-liquid supported-liquid membrane for the separation of CO2 from natural gas will be another potential research area for the future. References: [1] S. Jobling, J. P. Sumpter, Aquat. Toxicol. 27 (1993) 361 [2] J. Vos, E. Dbing, H. A. Greim, O. Ladefoge, C. Lambré, J.V. Tarazona, I. Brandt, D. Vethaak, Crit. Rev. Toxicol. 30 (2000) 71 [3] S. Liu, P.K. Dasgupta. Anal. Chem. 67 (1995) 2042 [4] M. A. Jeannot, F. F. Cantwell, Anal. Chem. 69 (1997) 235 [4] S. Pedersen-Bjergaard, K.E. Rasmussen, Anal. Chem. 71 (1999) 2650 [5] Y. He, H. K. Lee. Anal. Chem. 69 (1997) 4634 [6] M.A. Jeannot, F.F. Cantwell. Anal. Chem. 68 (1996) 2236 43 Chapter 2 [7] L. Hou, X. Wen, C. Tu, H. K. Lee, J. Chromatogr. A 979 (2002) 163 [8] C. Basheer, H. K. Lee, J. P. Obbard, J. Chromatogr. A 968 (2002) 191 [9] C. Basheer, R. Balasubramanian, H. K. Lee, J. Chromatogr. A 1016 (2003) 11 [10] C. Basheer, H. K. Lee, J. P. Obbard, J. Chromatogr. A 1022 (2004) 161 [11] E. Psillakis, N. Kalogerakis, J. Chromatogr. A 999 (2003) 145 [12] .G. Halvorsen, S. Pedersen-Bjergaard, K.E. Rasmussen, J. Chromatogr. A 909 (2001) 87 [13] H.G. Ugland, M. Krogh, K.E. Rasmussen, J. Chromatogr. B 749 (2000) 85. [14] T. S. Ho, S. Pedersen-Bjergaard, K.E. Rasmussen, Analyst 127 (2002) 608 [15] L. Zhu, K. H. Ee, L. Zhao, H. K. Lee, J. Chromatogr. A 963 (2002) 335 [16] T. Welton, Chem. Rev. 99 (1999) 2071 [17]. E.G. Yanes, S. R. Gratz, M. J. Baldwin, S.E. Robison, A. M. Stalcup. Anal. Chem. 73 (2001) 3838 [18]. M. Vaher, M. Koel, M. Kaljurand. J. Chromatogr. A 979 (2002) 27 [19]. T. F. Jiang, Y. L. Gu, B. Liang, J. B. Li, Y. P. Shi, Q.Y. Ou. Anal. Chim. Acta. 479 (2003) 249 [20]. L. J. He, W. Z. Zhang, L. Zhao, X. Liu, S. X. Jiang. J. Chromatogr. A 1007 (2003) 39 [21]. W. Z. Zhang, L. J. He, Y. L. Gu, X. Liu, S. X. Jiang. Anal. Lett. 36 (2003) 827 [22]. P.H. Shetty, P. J. Youngberg, B. R. Kersten, C. F. Poole. J. Chromatogr. 411 (1987) 61 [23]. D.A. Armstrong, L. F. He, Y. S. Liu. Anal. Chem. 71 (1999) 3873 44 Chapter 2 [24] J. L. Anderson, T. Welton, J. Ding, D.A. Armstrong. J. Am. Chem. Soc. 124 (2002) 14247 [25]. J. L. Anderson, D. A. Armstrong. Anal. Chem. 75 (2003) 4581 [26] J.-F. Liu, G.-B. Jiang, Y.-G. Chi, Y.-Q. Cai, Q.-X. Zhou, J.-T. Hu. Anal. Chem. 23 (2003) 5870 [27] J.-F. Liu, Y.-G. Chi, G.-B. Jiang, C. Tai, J.-F. Peng, J.-T. Hu, J. Chromatogr. A 1026 (2004) 143 [28] J.-F. Peng, J. F. Liu, G.-B. Jiang, C. Tai, M-J. Huang. J. Chromatogr. A 1072 (2005) 3 [29] C. Basheer, H. K. Lee, J. Chromatogr. A 1047 (2004) 189 [30] E. Psillakis, N. Kalogerakis. Trends Anal. Chem. 21 (2002) 53 [31] C. Basheer, J. P. Obbard, H. K. Lee, J. Chromatogr. A. 1068 (2005) 221 [32] M. Charalabaki, E. Psillakis, D. Mantzavinos, N. Kalogerakis, Chemosphere 60 (2005) 690 [33] C. F. Poole, J. Chromatogr. A 1037 (2004) 49 [34] P. Bonhote, A.-P. Dias, M. Papageorgiou, K. Kalyanasundaram, M. Gratzel, Inorg. Chem. 35 (1996) 1168 [35] G. Shen, H.K. Lee, Anal. Chem. 74 (2002) 648 [36] A.A. Boyd-Boland, J. Pawliszyn, J. Chromatogr. A 704 (1995) 163 45 Chapter 3. Novel micro-solid-phase extraction of carbamates in green tea leaves with determination by high performance liquid chromatography 3.1. Introduction Carbamates pesticides have been linked with fetal death, hormonal changes, DNA damage, birth defects and several adverse effects have been reported [1]. Which were originally extracted from the calabar bean. The use of carbamates as insecticides in agriculture began in the 1950s and approximately 25 carbamate compounds are in use currently as pesticides and biocides for industrial and other applications [2]. Their residues in food and agriculture products are of great interest because pesticides enter the human system through direct consumption of contaminated food, drinks, meat, and other products obtained from vegetables, fruits and animals origin. Green tea is consumed as a popular beverage worldwide because of its characteristic aroma, flavor and antioxidants health benefits. Harmful residual limits for carbamates have been set by several organizations such as the Food and Agricultural Organization [3], the European Union [4] and the US Food and Drug Administration [5]. Their acute toxicities are of great concern for food control because toxic values of carbamates are under 5 mg/kg (5 ppm) in the diet. Carbamates are thermally unstable or non-volatile and hence are not satisfactorily separately by GC. They are polar pesticides and which mean they are more suited to HPLC [6-11]. Even so, the trace analysis of environmental pollutants is challenging and simple chromatographic methods are not adequate to perform the task; 46 Chapter 3 effective sample preparation steps are necessary [12]. Recent developments in the analysis of carbamates pesticides in environmental samples include the followings: (i) methanolic ultrasonication [13], (ii) LPME with determination by GC [14], (iii) microwave-assisted extraction and supercritical fluid extraction of these pesticides in soil sample[15] [16], (iv) solid-phase extraction [17], (v) hot water extraction of carbamates[18] and (vi) in-tube SPME-HPLC for water sample [19]. In this present work, we have developed a novel micro-solid-phase extraction (µSPE) procedure, which can provide simpler, more cost-effective, faster, higher selectivity and better sensitivity than the recent extraction methods for the determination of polar carbamates pesticides in tea sample. 3.2. Experimental 3.2.1. Chemicals and materials HPLC-grade methanol was bought from Mallinckrodt (Paris, KY, USA) and HPLC-grade acetonitrile and tetrahydrofuran (THF) were obtained from J.T. Baker (Philipsburg, PA, USA). Ultrapure water was produced on a Nanopure system (Barnsted, Dubuque, IA, USA). Analytical-grade of glacial acetic acid (HAc), hydrochloric acid, sodium hydroxide and sodium chloride were purchased from Merck (Darmstadt, Germany). The carbametes; carbaryl (purity 98%), promecarb (purity 99%), methiocarb (purity 99%), propham (purity 99.5%), chlorpropham (purity 99.5%) and barban (99%) were supplied by ChemService (West Chester, PA, USA). Stock standard solutions (1 mg/mL) of each carbamate were prepared in methanol. A standard solution containing 10 mg/L of each carbamate was dissolved in 50:50 methanol-deionized water. All solutions were stored at 4oC. OSK Japan green tea samples were purchased from a supermarket in 47 Chapter 3 Singapore. Q3/2 Accurel polypropylene membrane sheet, (157 µm, thickness, 0.2 µm pore size), (Membrana, Wuppertal, Germany) was selected for the experiments. Different sorbent materials including C18, C8, C2, activated charcoal, HayeSep A (divinylbenzene ethyleneglycoldimethacrylate, HayeSep B (divinylbenzene polyethyleneimine), Porapak were purchased from Alltech (Deerfield, IL, USA ). Multiwalled carbon nanotubes (MWCNTs) were obtained from Honeywell Private Limited (Singapore). Plastic 200-µL graduated microcentrifuge tubes from Bioplastics (Landgraaf, Netherlands) were used for both ultrasonication and centrifugation. The ultrasonicator was supply by Midmark (Versailles, OH, USA) and the magnetic stirrer/hot plate was obtained from Heidolph (Cinnaminson, NJ, USA). Carbaryl Promecarb Propham Methiocarb Chlorpropham 48 Chapter 3 Barban Figure 3.1. Structures of carbamates considered in this work. 3.2.2. Chromatographic analysis Waters HPLC system, that is same as used in Chapter 2, equipped with 1525 µ binary pump, 200 µL injection loop, in-line degasser and waters 2487 UV dual λ absorbance detector is utilized throughout the whole experiment and data processing is carried out by Empower software. The analytical column selected for analysis was PhaseSep ODS2 5 µm, 250mm ×4.00 mm ID and column temperature kept at 25oC. The detector wavelength of 225 nm was chosen and mobile phase was water-acetonitrile (56:44) at a flow rate of 0.8 mL/min. The injection volume was 100 µL in all experiments. 3.2.3. Sample preparation Prior to µ-SPE, carbamates were first extracted from green tea leaves into pure water by hot water infusion (HWI) and by ultrasonic extraction (USE) since the sample needed to be in liquid form for µ-SPE. For the hot water extraction, 0.2 gm of tea leaves were infused into 20 mL of boiling water (1 % solution). This solution was placed for 30 min without further heating. This represents the normal brewing condition for a cup of tea. Ultrasonic extraction using a water-bath was performed with 0.2 gm grounded green tea and 20 mL of ultra pure water in a 50-mL screw-capped bottle. Sonication was carried out continuously for 1 h. The solutions were filtered after extractions before the µ-SPE. 49 Chapter 3 Tea is normally classified into green tea (unfermented), Oolong tea (semi-fermented), and black tea (fully fermented). Green tea was chosen in this work because its properties are not yet amended by fermentation. Pure water was used as extracting solvent for ultrasonication because this represents the conventional brewing condition. Also, the solubility of polar carbamates in water is very high. 3.2.4. Micro-solid phase extraction (µ-SPE) procedure Polypropylene membrane sheet C18 Figure 3.2. µ-SPE procedure for analysis of carbamates in aqueous sample (Pictures are not to scales) As shown in Figure 3.2, µ-SPE procedure consisted of: (i) preparation of µ-SPE device, (ii) extraction in sample solution, (iii) desorption, (iv) centrifugation and (v) injection of final desorbing solvent into the HPLC. µ-SPE devices were prepared in the following manner: a porous polypropylene membrane sheet (1.5 cm×1.5 cm) was folded in half and heat-sealed to become a small envelope with an opening. Exactly, 20 mg of 50 Chapter 3 solid phase materials (C18, C8, C2, activated charcoal, HayeSep A, HayeSep B, Porapak and MWCNTs) were individually packed into these envelopes. Later, another openings of which were heat-sealed. µ-SPE devices were also prepared by packing a mixture of two different sorbents into a single envelope. The devices were conditioned/ cleaned up by ultrasonication in acetone before the extraction. µ-SPE device was placed into a 50-mL bottle that contained 20 mL of sample solution. The solution was stirred at 1000 rpm for an extraction time of 40 min. After extraction, µ-SPE device was cleaned carefully with tissue paper and placed in a 200-µL graduated microcentrifuge tube filled with 150 µL of methanol. Then, µ-SPE device was desorbed in 150 µL of methanol by ultrasonication for 20 min. After that, the device was removed. The desorbing solvent (methanol) was centrifuged for 5 min to precipitate the suspended particles. Finally, 100µL of desorbing solvent was injected into HPLC. According to the experiments µ-SPE devices were reusable for more than 20 times after cleaning and conditioning in acetone. 3.2.5. Principle of µ-SPE µ-SPE is a modification of conventional solid-phase extraction (SPE). A liquid sample is in contact a sorbent during extraction. In µ-SPE, the latter is held within a membrane envelope. As a result of strong attractive forces between the analytes and the sorbent, the analytes are retained. After extraction, the sorbent is desorbed with a small amount of a solvent. The final result is that the analytes are concentrated in a relatively small volume of solvent and are therefore ready to be determined without any additional sample work up. When applying a µ-SPE method a number of factors must be considered. 51 Chapter 3 Sorbent selectivity: the character of sorbent-analyte interactions can be divided into three groups: non-polar, polar and ionic. In the majority of cases non-polar or slightly polar analytes are dissolved in water, a highly polar solvent. For these applications, non-polar sorbents can be employed. On the other hand, analytes containing polar functional groups will be retained on sorbents of opposite polarity. For retention to occur with ionic interactions, an anionic sorbent should be selected to retain cations and a cationic sorbent to retain anions. Sorbent capacity: When selecting the optimum packing size for a particular applications, factors to be considered are the ability of the sorbent to retain all of the analytes present in the sample and volume of the original sample [22-26]. 3.3. Results and discussion 3.3.1. Optimization of the method Six carbametes pesticides, carbaryl, promecarb, methiocarb, propham, chlorpropham and barban were selected as analytes for the present work. All the target compounds can be separated well by HPLC within 28 min with the isocratic mobile phase concentration of water-acetonitrile (56:44) without any modifier. The optimum UV detection wavelength was 225 nm. Similarly, all the tea components and caffeine extracted were separated well from the target analytes with the above HPLC conditions. Pre-sample preparation carried out by hot water infusion as mentioned in section 3.2.3 produced broader peaks with more matrix interfering effects compared to ultrasonic extraction. This is probably because carbamates are thermally unstable and more tea components are extracted when infused with boiling water. Therefore ultrasonication was 52 Chapter 3 selected for further work. The solution was filtered through 0.45 µm filter before µ-SPE analysis. Initially, polypropylene membrane was selected to fabricate the µ-SPE device as it is compatible with most organic solvents. Before use, the membrane was ultrasonically cleaned in acetone for 2 min in order to remove any possible contaminants. For the desorbing solvent, three polar organic solvents, methanol, acetonitrile and THF were evaluated. The best results were obtained from using methanol. After analyte desorption, the solutin was centrifuged to ensure the settlement of particles in the extract. 1 3 A U 2 5 (c) 6 4 (b) 0.0 (a) 6 8 10 12 14 16 18 20 22 24 26 28 Minutes Figure 3.3. Chromatography of [a] blank green tea sample, [b] 100 µg/L spiked green tea before µ-SPE extraction and [c] 20 µg/L spiked tea solution after extraction under optimized condition. Peaks identities; (1) carbaryl, (2) propham,(3) methiocarb, (4) promecarb,(5) chlorpropham and (6) barban. Figure 3.3. shows a comparison between a liquid chromatogram of 100 µg/L spike sample solution (containing 100 µg/L each carbamates) [b] and that of a sample (spiked at 20 µg/L of each carbamates), after µ-SPE [c]. Most of the analytes in the 53 Chapter 3 second standard solution (spiked at 20 µg/L of each carbamates ) were not detected when 100 µL of this solution was directly injected into HPLC. 3.3.2. Individual and mixed-mode sorbents approaches 250000 200000 carbaryl propham methiocarb promecarb chlorpropham barban Area 150000 100000 50000 0 C18 C8 C2 Activated- HayeSep A HayeSep B charcoal Porapak MN-CNT Figure 3.4. Effect of individual sorbents packing on the final results. µ-SPE conditions: 20 µg/L ppb spiked sample, 20 mL sample volume, extraction time-30 min, desorbing time-30 min, centrifuge-5 min, injection volume-100 µL. Appropriate sorbent selection is the bottleneck in the µ-SPE process. In this experiment, eight individual sorbents (C18, C8, C2, activated charcoal, HayeSep A, HayeSep B, Porapak and multiwalled carbon nanotubes) and nine mixed-mode sorbents of different polarities (HayeSepA+C18, HayeSepA+C8, HayeSepA+C2, HayeSepB+C18, HayeSepB+C8, HayeSepB+C2, Porapak+C18, Porapak+C8 and Porapak+C2 ) we have tested. Figure 3.4. shows that reversed-phase (RP) non-polar sorbents having alkyl groups such as octadecyl (C18), octyl (C8), ethyl (C2) gave better results than other sorbents because carbamates analytes are polar (hydrophilic) compounds and they are more likely to be retained by the non-polar sorbents. RP sorbents interact with polar analytes via van 54 Chapter 3 1000000 900000 carbaryl propham 800000 methiocarb promecarb 700000 chlorpropham barban Area 600000 500000 400000 300000 200000 100000 0 H sepA+C 18 H sepA+C8 H sepA+C2 H sepB+C 18 H sepB+C8 H sepB+C2 porapak+C 18 porapak+C8 porapak+C2 Figure 3.5. Effect of mixed-mode sorbents packing on extraction efficiency. µ-SPE conditions as Figure 3.4. (where H sepA = Haye SepA, H sepB = Haye SepB) der Waals forces with the energy of interaction at about 41.8 kJ/mol [28-29]. Activated charcoal does not extract well even when compared to more polar sorbents such as HayeSepA&B and Porapak. Multiwalled carbon nanotubes (MWCNT) produced moderate efficiency. Among all sorbents, C18 give the highest extraction efficiency. Thus, may be because C18 has the longest alkyl chain that is more compatible with the analytes. The C18 sorbent may exhibit secondary or dual-retention mechanism due to unreacted surface silanol groups [30]. Therefore, electrostatic or dipole-dipole interaction mechanisms are also possible for this extraction. Of mixed-mode sorbents, as Figure 3.5 shows, the mixture of HayeSep A and C2 provides the highest peak areas. The mixture gave much higher extraction efficiency than C18 for some target analytes except for the more retained analytes, chlorpropham and barban. It is assumed that mixed-mode sorbents have dual or multiple retention mechanisms and exploit the interactions with different functional groups on a particular single analyte. However, in general, C18 has better results for all the target analytes taken 55 Chapter 3 together and was chosen for further work. The mixed-mode mixture of HayeSep A and C2 gave higher peak areas for four analytes in this work, and would be the sorbent of choice for µ-SPE if only these four compounds were considered. 3.3.3. Effect of extraction and desorption time 250000 carbaryl propham methiocarb 200000 promecarb chlorpropham 150000 Area barban 100000 50000 0 10 min 20 min 30 min 40 min 50 min 60 min Figure 3.6. Effect of exposure time on peaks areas. µ-SPE conditions: C18 sorbent, 20 µg/L spiked sample, 20 mL sample volume, desorbing time-30 min, centrifuge-5 min, injection volume-100 µL. Both extraction and desorption time are critical to efficient µ-SPE. Therefore, extraction times of 10, 20, 30, 40, 50, 60 min and desorption times of 5, 10, 15, 20, 25 and 30 min were investigated in this work. Figure 3.6 showed that extraction efficiency increases from 10 to 40 min and remained more or less constant after that, indicating that equilibrium was obtained at that time. Twenty min of desorption time was deemed to be optimum for the removal of analytes from the sorbent, as Figure 3.7 shows. 56 Chapter 3 250000 carbaryl propham methiocarb 200000 promecarb chlorpropham Area 150000 barban 100000 50000 0 5min 10min 15min 20min 25min 30min Figure 3.7. Effect of desorption time on the results. µ-SPE conditions: C18 sorbent, 20 µg/L spiked sample, 20 mL sample volume, extraction time-40 min, centrifuge-5 min, injection volume-100 µL. 3.3.4. Dependence of pH and ionic strength carbaryl 250000 propham methiocarb 200000 promecarb chlorpropham Area 150000 barban 100000 50000 0 pH 2 pH 4 pH 6 pH 8 pH 10 pH 12 Figure 3.8. Dependence of pH on analytes peak areas. µ-SPE conditions: C18 sorbent, 20 µg/L spiked sample, 20 mL sample volume, extraction time-40 min, centrifuge-5 min, injection volume-100µL. According to Figure 3.8, highest analytes peak areas were obtained at the pH 6. This pH value was almost same as the pH of pure water (pH=5.8) used for this work, 57 Chapter 3 carbaryl 250000 propham methiocarb 200000 promecarb chlorpropham Area 150000 barban 100000 50000 0 5%NaCl 10%NaCl 15%NaCl 20%NaCl 25%NaCl 30%NaCl Figure 3.9. Dependence of NaCl addition on peak areas. µ-SPE conditions: same as Figure 3.8. whereas sample with pH 2 produced lowest peak areas. In fact, these results were expected based on the behavior of C18 sorbent which is only stable over a pH range of between 2.5 and 10.5. Therefore, there was no pH adjustment of the samples for further experiments. Addition of different concentrations NaCl to the sample solutions was evaluated. As Figure 3.9 shows, salt did not significantly improve the extraction efficiency. This is probably because carbamates are polar (hydrophilic) compounds and theoretically, addition of salt to the sample solution can decrease their solubility and consequently increase their hydrophobicity. This is what happened in liquid-liquid extraction (LLE), in which the extraction solvent is organic and hydrophobic. No salt was, therefore, added to the sample solution in subsequent experiments. 3.3.5. Dependence of sorption on sample volume Six different sample volumes of 10, 20, 30, 40, 50 and 60 mL with constant 58 Chapter 3 250000 carbaryl propham 200000 methiocarb promecarb 150000 Area chlorpropham barban 100000 50000 0 10mL 20mL 30mL 40mL 50mL 60mL Figure 3.10. Dependence of sorption on sample volume. µ-SPE condition: same as Figure 3.8. concentrations were evaluated for µ-SPE. Figure 3.10 shows that sorption efficiency decreased slowly with increasing sample volumes. It could be possible that magnetic stirring, used in µ-SPE, was only suitable for smaller sample volume. Therefore, sample volume of 20 mL was selected as optimum volume for this work. 3.3.6. Method evaluation In analytical chemistry, the evaluation of a method is determined by the parameters such as repeatability, linearity and limit of detection (LODs). In this work, it was assumed that the performance of the HPLC for the carbamates considered was already validated. The µ-SPE procedure was evaluated after optimizations of the final conditions. The enrichment factor (EF) was also determined. This is defined as the ratio of the peak areas of the analytes before and after µ-SPE for the same spiked sample using the optimized conditions. The reproducibility of the method was determined by performing the extraction of six tea samples spiked at the same concentration of 20 µg/L and the method produced relative standard deviations (R.S.D) of 5.1 to 8.5%. 1, 5, 10, 15, 20 and 25 µg/L samples were extracted to evaluate the linearity. All analytes 59 Chapter 3 exhibited good linearity with correlation coefficients (r) of 0.9841–0.9979, as shown in Table 3.1. LODs calculated based on the signal to noise ratio (S/N of 3) in HPLC measurements, were in the range of 0.005 µg/L (carbaryl) to 0.1 µg/L (promecarb). All the results obtained are shown in Table 3.1, These result are comparable to typical analytical extraction methods for carbamates [32]. Therefore, the present µ-SPE method is feasible for the routine analysis of carbamates in green tea leaves samples. Table 3.1. µ-SPE; Repeatability, Linearity, Limit of detection and Enrichment factor Analytes RSD (%) (n=6) Linearity range (µg/L, 6 points) Correlation Coefficient (r) LOD (µg/L) Enrichment Factor (EF) carbaryl 5.1 1-25 0.9910 0.005 101.0 propham 8.5 1-25 0.9849 0.032 32.0 methiocarb 2.6 1-25 0.9947 0.015 45.9 promecarb 5.9 1-25 0.9979 0.100 43.0 chlorpropham 6.9 1-25 0.9841 0.028 45.4 barban 7.9 1-25 0.9850 0.018 53.8 3.4. Conclusion Ultra trace analysis of six common carbamates in green tea leaves was performed by using µ-SPE-HPLC. When µ-SPE was applied to the analysis of fresh OSK green tea sample (without spiking), we have found that there were no detectable amount of carbamates in these samples. The present µ-SPE method is simple, cost-effective, sensitive, selective, reproducible and involves minimized organic solvents use. LODs of down to 0.005 µg/L levels and reproducibility (R.S.D) of average 6.2% show the merits of the procedure. It is conceivable that the procedure is suitable for the determination of other pollutants in the environments as well. 60 Chapter 3 References; [1] G. P. Casale, J. L. Vennerstrom, S. Bavari, T. L. Wang, Immunopharmacol. Immunotoxicol. 15 (1993) 199 [2] K. A. Hassal, The Chemistry of Pesticides: Their Metabolism, Mode of Action and Uses in Crop Protection, Macmillan, New York, 1983 [3] Joint FAO/WHO Food Standards Programme, Codex Alimentarius Commission, Pesticides Residues in Food, vol. II, Food and Agriculture Organization and World Health Organization, Rome, 1993 [4] European Union Council Directive no. 95/39/EC of 7 July 1995, Off. J. Eur. Communities no. L197 of 22 August 1995; European Union Council Directive no. 96/33/EC of 21 May 1996, Off. J. Eur. Communities no. L144 of 18 June 1996; European Union Council Directive no. 94/29/EC of 23 June 1994, Off. J. Eur. Communities no. L189 of 23 July 1994 [5] Food Quality Protection Act, 3 August 1996, Pub. L. no. 104-170 (1996) [6] M. Fernandez, Y. Pico, J. Manes, J. Chromatogr. A 871 (2000) 43 [7] J. Zrostlıkova, J. Hajslova, T. Kovalczuk, R. Stepan, J. Poustka, J. AOAC Int. 86 (2003) 612 [8] A.C. Hogenboom, M.P. Hofman, S.J. Kok, W.M.A. Niessen, U.A.Th. Brinkman, J. Chromatogr. A 892 (2000) 379 [9] K. A. Barnes, R. J. Fussell, J. R. Startin, M. K. Pegg, S.A. Thorpe, S.L. Reynolds, Rapid Comm. Mass Spectrom. 11 (1997) 117 [10] K. Bester, G. Bordin, A. Rodriguez, H. Schimmel, J. Pauwels, G. V. Vyncht, Fresenius J. Anal. Chem. 371 (2001) 550 61 Chapter 3 [11] J. Klein, L. Alder, J. AOAC Int. 86 (2003) 1015 [12] L. Mondello, A. C. Lewis, K. D. Bartle, Multidimensional Chromatography, 2002 [13] K. Granby, J. H. Andersen, H. B. Christensen, Anal. Chim. Acta. 520 (2004) 165 [14] D. A. Lambropoulou, T. A. Albanis, J. Chromatogr. A 1072 (2005) 55 [15] L. Sun, H. K. Lee, J. Chromatogr. A 1014 (2003) 165 [16] T. Okumura, K. Imamura, Y. Nishikawa, Analyst 120 (1995) 2675 [17] J.M.F. Nogueira, T. Sandra, P. Sandra, J. Chromatogr. A 996 (2003) 133 [18] S. Bogialli, R. Curini, A. Di Corcia, A. Lagan, M. Nazzari, M Tonci, J. Chromatogr. A 1054 (2004) 351 [19] Y. Gou, R. Eisert, J. Pawliszyn, J. Chromatogr. A 873 (2000) 137 [20] Gy. Matolcsy, M. Nadasy, V. Andriska, Pesticide Chemistry, Elsevier Science, Amsterdam, 1988 [21] C. Basheer, H. K .Lee, Chromatogr. A 1047 (2004) 189 [22] T. Cserhati, E. Forgacs, Chromatography in Environmental Protection, Budapest, Hungary, 2001 [23] S. Mitra, Sample Preparation Techniques in Analytical Chemistry, Hoboken, NJ, 2003 [24] J. R. Dean, Extraction Methods for Enviromental Analysis, University of Northumbria, John Wiley, New York, 1998 [25] A. J. Handley, Extraction Methods in Organic Analysis, Sheffield Academic Press, Sheffield, 1999 [26] M. Moors, D. L. Massart, R. D. McDowall, Pure Appl. Chem. 66 (1994) 277 62 Chapter 3 [27] R. L.Grob, Chromatographic Analysis of the Environment, M. Dekker, New York, 1975 [28] A. J. Handley and R.D. McDowall, Solid-Phase Extraction in Organic Analysis, Boca Raton, Sheffield Academic Press; Sheffield, 1999 [29] C. F. Poole, S. K. Poole, Solid-Phase Extraction; Theory Meets Practice, Marcel Dekker, New York, 2000 [30] M. Cooke, C. F. Poole, Vol.10, Encyclopedia of Separation Science, Academic Press, San Diego, 2000 [31] H. Horie, K. Kohata, J. Chromatogr. A 881 (2000) 425 [32] J. M. Soriano, B. Jimenez, G. Font and J. C. Molto, Crit. Rev. Anal. Chem. 31 (2001) 19 63 Chapter 4. Novel Amphiphilic Poly(P-Phenylene)s Used as Sorbent for Solid-Phase Microextraction of Environmental Pollutants 4.1. Introduction Polycyclic aromatic hydrocarbons (PAHs), organophosphorous pesticides (OPPs) and organochlorine pesticides (OCPs) are important classes of persistent organic pollutants (POPs) that are commonly found in the environment [1]. POPs are extremely hazardous because of their toxicity, in combination with high chemical and biological stability, and a high lipophilicity [2]. POPs are polluted into the environment and become incorporated into food webs [3-4]. Therefore, the accurate measurement and monitoring of these compounds are become important in today’s society. For example, the presence of OCPs in the environmental waters had been strictly regulated by legislation to concentrations below 0.01 µg/L in many countries [5-7]. Thus, these very low trace levels call for the extraction/ pre-concentration techniques that can provide an easy, rapid and sensitive determination of POPs in the environment. In not only the environmental waters, but POPs are also detected routinely in fish and wildlife, as well as human adipose tissue, blood and breast milk [8-9]. Traditionally, amounts of POPs in solid environmental samples are determined by liquid-solid extraction (Soxhlet extraction) [10]. In recent years, new extraction techniques such as supercritical fluid extraction (SFE), pressurized fluid extraction (PFE) [11–12] and microwave-assisted extraction (MAE) [13-14] had been developed for the determination of POPs from solid matrices. The disadvantages of these techniques are that they require large sample size and solvent volume. Recently, C. Basheer, J. P. 64 Chapter 4 Obbard and H. K. Lee have developed a novel microwave-assisted solvent extraction (MASE) in combination with simple liquid-phase microextraction (LPME) cleanup and enrichment procedure supported by hollow fiber membrane (HFM), (MASE-HFMLPME), for the determination of POPs in marine sediments [15]. Sample preparations for the analysis of POPs environmental waters involves techniques such as liquid-liquid extraction (LLE), solid-phase extraction (SPE), head space (HS), purge and trap (P&T), solid-phase microextraction (SPME) and direct GC analysis using large injection volume with modified injectors [16-17]. Arthur and Pawliszyn developed SPME in the late 1980’s [18]. They introduced it as a solvent-free sample preparation technique that could serve as an alternative to traditional extraction procedures such as LLE, SPE, HS and P&T procedures. SPME preserves all of the advantages of SPE while eliminating the main disadvantages of low analyte recovery, plugging, and solvent use [19-20]. This technique utilizes a short thin solid rod of fused silica (typically 1 cm long and 0.1um outer diameter), coated with a sorbent polymer. The coated fused silica (SPME fiber) is attached to a metal rod; the entire assembly (fiber holder) may be described as a modified syringe. There are two approaches of sampling of volatile organics in SPME: direct and headspace [21-22]. In addition, membrane-protected SPME sampling has been also applied where the fiber is separated from the sample with a selective membrane which lets analytes through while blocking interferences [23]. The main advantages of SPME include its simplicity, easy automation and on-site application due to its portability. SPME has been interfaced to HPLC, CE and FT-IR in addition to GC [24] and applied to extract from a wide variety of the sample matrices [25]. 65 Chapter 4 The following polymers are commercially available for SPME. Polydimethylsiloxane (PDMS) has been used to extract non-polar analytes, such as, alkyl benzenes, PAHs, and volatile halogenated compounds [26-27]. Polyacrylate (PA), a mixture of PA & Carbowax (CW), and polydivinylbenzene (DVB) polymers are used for alcohols and small polar compounds [28-30]. Recently, sol–gel technology has been used to provide an efficient incorporation of organic components into the inorganic polymeric structures in solution under extraordinarily mild thermal conditions. Reports on the application of sol–gel technology to prepare SPME coatings have been increasing in recent years [31-35]. The significant drawbacks of commercial SPME are; (a) their recommended operating temperatures are relatively low, because the extraction phases of commercial SPME are prepared by physical deposition of the polymer coating rather than bonding and cross-linking and (b) a reduction of the life time of the fibers due to desoption of higher salt content samples or complex matrices [36-37]. The lack of proper chemical bonding between the stationary phase and fused silica fiber surface and cross-linking among the stationary phase itself may be responsible for the low thermal and chemical stability of commercial SPME. Here, we describe the development of an amphiphilic polymer as a novel stationary phase for SPME. The polymer prepared is applied for the extraction/ pre-concentration of PAHs, OCPs and OPPs from environmental water samples. 4.2. Experimental 4.2.1. Materials and reagents 66 Chapter 4 Fused silica capillary tubes (77µm I.D. and 194µm O.D.) were purchased from Polymicro Technologies (Phoenix, AZ, USA). The SPME holder for manual sampling was obtained from Supelco (Bellefonte, PA, USA). The SPME fiber holder and fibers (PDMS–DVB, PA) were used without modification for comparison with the sorbent used for this work. Before extraction, the fibers were conditioned in the GC injection port based on the manufacturer’s recommended procedure. All solvents used in this study were of analytical-reagent grade. A stock solution of eleven OCPs [hexachlorobenzene, lindane, heptachlor, aldrin-R, trans-chlordane, cis-chlordane, p, p′-DDE (p,p′dichlorodiphenyldichloroethylene) , dieldrin, endrin, p, p′-DDD (p, p′- dichlorodiphenyldichloroethane), p, p′-DDT (p, p′-dichlorodiphenyltrichloroethane)] and a stock solution of six OPPs [triethylphosphorothioate, thionazin, sulfotep, phorate, disulfoton, methyl parathion] were purchased from PolyScience (Niles, IL, USA). A stock solution seven PAHs [naphthalene, acenaphthene, fluorene, phenenthrene, anthracene, fluoranthene, pyrene] was obtained from Sigma–Aldrich (St. Louis, MO, USA). Ultrapure water was prepared on a Milli-Q (Millipore, Milford, MA, USA) system. A standard stock solution of 10 mg/L each of OCPs, OPPs and PAHs was prepared in methonol and diluted to 100 µg/L for working standard solutions. 4.2.2. GC-MS analysis Analysis was performed on a Shimadzu (Tokyo, Japan) QP2010 gas chromatography–mass spectrometry (GC–MS) system equipped with a Shimadzu AOC20i auto sampler and a DB-5 fused silica capillary column 30m×0.32mm I.D., film thickness 0.25µm (J&W Scientific, Folsom, CA, USA). Helium (purity 99.9999%) was used as the carrier gas at a flow rate of 1.5 ml min−1 and splitless injection mode was 67 Chapter 4 used. For the analysis of OCPs and OPPs, the injection temperature was set at 250 oC and the interface temperature at 280 oC. The oven temperature program used was as follows: initial temperature of 70 oC was held for 2 min, then increased to 250 oC at a heating rate of 30 oCmin−1, followed by another ramp of 30 to 280 oC min−1. The later temperature was held for 2 min. For PAHs analysis, the injection temperature was set at 320 oC with the interface temperature of 280 oC and the oven temperature program used was as follows: initial temperature of 70 oC was held for 2 min, then increased to 120 oC at a rate of 20 oCmin−1, followed by increased to 245oC at 5 oC min−1, finally increased to 320oC at 10 oCmin-1and held for 2 min. The total program time was 40 min. All standards and samples were analysed in selective ion monitoring (SIM) mode with a detector voltage of 1.5 kV using a mass scan range of m/z 50–500. 4.2.3. Amphiphilic poly(p-phenylene)s Hydroxylated amphiphilic poly(p-phenylene)s (C12PPPOH) are an interesting class of conjugated polymers, extensively studied in our lab [38-43]. The chemical structure of the functionalized C12PPPOH used for coating of the capillary for SPME is shown in Figure 4.1. The amphiphilicity of the PPP backbone originates from the incorporation of a long alkoxy chain and hydroxyl groups on either side of the polymer backbone. The rigid-rod structure of the polymer backbone with polar and non-polar groups showed interesting self-assembly in the solid state and in solution [44]. C12H25O n OH Figure 4.1. The chemical structure of C12PPPOH 68 Chapter 4 4.2.4. Synthetic scheme C12PPPOH was synthesized with the use of the Suzuki polycondensation reaction as summarized in following scheme. Polyphenols are used as starting materials and the reactions consist of six stages which are; (i) reaction with bromine/acetic acid [Br2/AcOH], 80 %, (ii) sodium hydroxide [NaOH], 1-bromododacane [CH3(CH2)11Br], 50° C, 10 h, 65 %, (iii) [K2CO3], benzyl bromide [C6H5CH2Br], 50° C, 10 h, 90 %, (iv) nbutyllithium [n-BuLi], tetrahydrofuran [THF], –78 °C, triisopropyl borate, 30o C, 10 h, 70 %, (v) 2M potassium carbonate [2M K2CO3], 3.0 mol % tetrakis(triphenylphosphine) palladium(0) [3.0 mol % Pd(PPh3)4], toluene, reflux, 3 day and (vi) hydrogen, 10% palladium on carbon [10% Pd/C], chloroform/ethanol/THF. OH OH OR (i) OR (iii) (ii) Br HO Br Br HO 1 Br HO 2 BnO 3 4 (iv) OR RO Br Br RO BnO n OBn (v) OBn OR (HO)2B BnO 6 Br Br B(OH)2 5 (vi) RO RO n OH OH R=C12H25 Figure 4.2. Synthetic scheme of C12PPPOH polymer 69 Chapter 4 4.2.5. Preparation of SPME fiber Thin films of the polymers on bare capillaries (i.e. both end were opened) were prepared by drop casting from 0.5 mg/ml polymer in chloroform solution under ambient conditions, without any airflow or temperature control aids. Scanning electron micrograph (SEM) images were taken with a JEOL JSM 6700 scanning electron microscope and the thickness of the fiber was scanned at approximately 7 µm. The capillaries were carefully mounted on copper stubs with a double-sided conducting carbon tape and sputter coated with 2 nm platinum before examination. The casted film morphology on the capillary is shown in Figure 4.3 (ii) and (iii). When compared to bare capillary Figure 4.3 (i), the polymer-casted capillary is shown to have been coated with a layer of intricately patterned film of polymer. The ordered patterns are the result of condensed water on the surface of the polymer solution as the solvent evaporates, cooling the surface below dew point. Droplets of the condensate organize into most stable positions in the small time frame of the evaporation process. The growth of patterns proceeded in multiple stages [45-49]. The first stage involved the formation of small isolated droplets of condensed water, the second stage involved a marked increase in droplet sizes and the last stage where droplets interact and coalesce, driven by convection. The difference in the eventual patterns was therefore a compound result of the droplet-polymer-substrate interaction as well as droplet-droplet interaction. This process induced the phase separation of the polymer films leading to precipitation of polymer at the organic-water interface leaving behind these patterns. The strong interplay between three competing interaction forces, moisture, polymer and substrate generated a highly ordered film for some of the samples. 70 Chapter 4 There were no appreciable changes in pattern sizes as the concentrations were increased to 5 mg/ml where multi-layered films were obtained as a result of the more concentrated solution. (i) (ii) (iii) (iv) Figure 4.3. Scanning electron micrograph images of (i) bare capillary (500×); (ii) coated capillary (500×); (iii) coated capillary (5000×); (iv) coated capillary using a concentrated polymer solution (5000×). 4.2.6. SPME Theory The principle behind SPME is the partitioning of analytes between the sample matrix and extraction medium. If a liquid polymer coating is used, we can use the following equation to relate the amount of analyte adsorbed by the coating at equilibrium to its concentration in the sample: n = Kfs Vf Co Vs Eq. 4.1 Kfs Vf + Vs n: the mass of the analyte absorbed by the coating Vf: volume of the coating Vs: volume of the sample 71 Chapter 4 Kfs: the distribution constant of the analyte between the coating and the sample matrix Co: the initial concentration of the analyte in the sample As can be seen from this equation, there exists a linear relationship between the amount of analytes absorbed and their initial concentration in the sample. Coatings used in SPME typically have strong affinities for organic compounds and therefore, have large Kfs values for targeted analytes. This means that SPME is selective and has a very high concentrating effect. However, on many occasion, the Kfs values are not large enough to exhaustively extract most analytes in the matrix and only through proper calibration can SPME be used to accurately determine concentrations of target analytes. Calibration can be by the external standard method in a relatively clean sample and by standard addition or internal standards in a more complex matrix. If Vs is very large (Vs >> Kfs Vf): n = Kfs Vf Co Eq. 4.2 This means that when the volume of the sample is very large, the amount of analyte extracted by the fiber coating is not related to the sample volume. This feature, combined with its simple geometry makes SPME ideally suited for field sampling and analysis because the fiber can be exposed to air or dipped directly into a lake or river, without collecting a defined sample volume prior to analysis [19]. 4.2.7. SPME Procedure The coated fused silica (SPME fiber) is attached to a metal rod, and is protected by a metal sheath. The fiber is attired to the plunger syringe. For sampling, 10 mL of water sample is placed in a vial, magnetically agitated at 105 rad s-1 and the vial is seal with a cap with a septum. The protective sheath is pushed through the septum and the 72 Chapter 4 plunger is lowered, forcing the fiber into the vial, where it is immersed directly in the liquid sample. Analytes in the sample are adsorbed on the fiber. After a predetermined time, the fiber is withdrawn into the protective sheath which is pulled out of the sampling vial. Immediately after, the sheath is inserted into the septum of a GC injector, the plunger is pushed down, and the fiber is exposed in the injector where the analytes are thermally desorbed and swept into the GC column where they are separated. The desorption step lasts 5 min, afterwhich; the fiber is withdrawn into the protective sheath which is removed from the injector. 4.3. Results and discussion 4.3.1. C12PPPOH VS Commercial fibers Comparison studies were made between our C12PPPOH polymer and commercial PDMS-DVB and PA for the extraction of PAHs, OPPs and OCPs pure water spiking. Figure 4.4 to 4.6 show the total ion chromatograms of studied compounds using three different polymer coatings. Each extraction was performed three times. SPME conditions are described in the captions each chromatogram. As we can see in the figures, our C12PPPOH coated fiber obviously exhibits more than 20 times analytes peak signals than commercial PSMS-DVB and PA for all target compounds. Generally, the partition coefficients of the compounds considered in this work are very large. Thus, a thin-film of C12PPPOH (7 µm) not only provide the required sensitivity but also reduced analytes carry over between samples. The C12PPPOH coated fiber can be stable up to 320oC, which is much higher than the temperature limits of commercial fibers. In practical terms, non-polar OCPs and PAHs are better extracted by PDMS-DVB coating, whereas the more polar OPPs are better suited to PA coated fiber. 73 Chapter 4 The new C12PPPOH coating can be applied to both non-polar and more polar compounds, demonstrating its versatility. inten(x10,000,000) Figure 4.4. Total ion chromatogram of PAHs; (1) SPME using C12PPPOH coated fiber, (2) SPME using commercial PA coated fiber and (3) SPME using commercial PDMSDVB coated fiber. Peak identities; (a) Naphthalene, (b) Acenaphthene, (c) Fluorene, (d) Phenenthrene, (e) Anthracene, (f) Fluoranthene, (g) Pyrene. SPME conditions; 10 mL of 20 µg/L spiked into pure water, extraction time 30 min, stirring speed 105 rad s-1, 10 % NaCl, pH was not adjusted. inten(x100,000) Figure 4.5. Total ion chromatogram of OPPs; (1) SPME using C12PPPOH coated fiber, (2) SPME using commercial PDMS-DVB coated fiber and (3) SPME using commercial PA coated fiber. Peak identities; (a) Triethylphosphorothioate, (b) Thionazin, (c) Sulfotep, (d) Phorate, (e) Disulfoton, (f) Methyl parathion. SPME conditions; 10 mL of 20 µg/L spiked into pure water, extraction time 60 min, stirring speed 105 rad s-1, 10 % NaCl, pH was not adjusted. 74 Chapter 4 inten(x100,000) Figure 4.6. Total ion chromatogram of OCPs; (1) SPME using C12PPPOH coated fiber, (2) SPME using commercial PA coated fiber and (3) SPME using commercial PDMSDVB coated fiber. Peak identities; (a) Hexachlorobenzene, (b) Lindane, (c) Heptachlor, (d) Aldrin-R, (e) trans-Chlordane, (f) cis-Chlordane, (g) p, p′-DDE, (h) Dieldrin, (i) Endrin, (j) p, p′-DDD, (k) p, p′-DDT. SPME conditions; 10 mL of 20 µg/L spiked into pure water, extraction time 30 min, stirring speed 105 rad s-1, 10 % NaCl, pH was not adjusted. 4.3.2. Optimization of PAHs extraction using C12PPPOH coating PAHs were selected as the reference analytes for the method optimization of SPME using our new C12PPPOH coating. Experimental variables in SPME analysis included extraction time, pH effect, salt addition to the sample, heating of sample and agitation methods as mentioned in the previous reports [50-52]. In this experiments, heating was not applied in order to study the extraction efficiency of our coated fiber at ambient temperature. Although heating of the sample solution increases analyte diffusion rate so that equilibrium is reached much faster, microextraction is an exothermic process and, eventually leads to decreased extraction. SPME is not an exhaustive extraction process. Optimum extraction time is when equilibrium is reached after 30 min (see figure 75 Chapter 4 4.4 for SPME conditions). Magnetic stirring at 105 rad s-1 is the optimum rate of agitation. In general, for extraction from water sample, addition of inorganic salt to aqueous sample improves the extraction efficiency. The ionic strength shifts the partition equilibrium; in favor of mass transfer to the organic (or fiber) phase; therefore, the analytes are more retained on the fiber coating. It is noted that higher salt concentration also reduces the lifetime of the coating material [53-54]. Figure 4.7 shows that highest peak areas for all analytes were obtained at 10% NaCl concentration. This SPME condition, therefore, was selected for further experiments. 6.E+07 5% NaCl 10% NaCl 5.E+07 15% NaCl 20% NaCl Peak area 4.E+07 25% NaCl 30% NaCl 3.E+07 2.E+07 1.E+07 0.E+00 Naphthalene Acenaphthene Fluorene Phenenthrene Anthracene Fluoranthene Pyrene Figure 4.7. Effect of salt addition on C12PPPOH coated SPME. Extraction time 30 min, stirring speed 105 rad s-1, pH was not adjusted. Generally in SPME, basic compounds are extracted at aqueous NaOH and acidic compounds like phenols are analyzed better at lower pH. As can be seen in Figure 4.8, effects of pH does not significantly improve the extraction efficiency compared to salt addition. This is due to non-ionic nature of PAHs. Therefore, pH was not adjusted for further experiments. 76 Chapter 4 5.E+07 4.E+07 4.E+07 pH 2 pH 4 pH 6 pH 8 pH 10 pH 12 Peak area 3.E+07 3.E+07 2.E+07 2.E+07 1.E+07 5.E+06 0.E+00 Naphthalene Acenaphthene Fluorene Phenenthrene Anthracene Fluoranthene Pyrene Figure 4.8. Effect of pH on C12PPPOH coated SPME. Extraction time 30 min, stirring speed 105 rad s-1, no salt was added. 4.3.3. Method validation The precision, linearity, sensitivity and limits of detection (LODs) were evaluated using spiked water samples and the range of quantitation was performed on real water samples collected from St. John’s Island, Singapore. Reproducibility of six replicate measurements are evaluated at PAHs (spiked level at 20 µg/L) calculated relative standard deviations RSD were in the range of 4.5 and 9.0. The linearity was very good over the concentration range of 0.5 and 20 µg/L. The calculated correlation coefficient (r) values were more than 0.995. LODs were measured by progressively reducing the analyte concentrations in the sample so that GC-MS peaks signals were discerned at the signal to noise ratio (S/N) of 3. These were determined to be between 0.001 to 0.005 µg/L (Table 4.1). Relative recoveries and RSD were also performed on the seawater samples at PAHs (spiked level at 5 µg/L). Recovery study using C12PPPOH-coated SPME was found to be comparable to commercial PDMS-DVB-coated SPME and better than PA-coated SPME (Table 4.2). 77 Chapter 4 Table 4.1. Precision, linearity, and limits of detection of PAHs using C12PPPOH-coated fiber. SPME conditions; 10 mL of spiked water samples, extraction time 30 min, stirring speed 105 rad s-1, 10 % NaCl, pH was not adjusted. RSD (%) Linearity LOD Analytes (n=6) ( 0.5-20 µg/L) (µg/L) Naphthalene 5.6 0.9951 0.005 Acenaphthene 6.2 0.9991 0.004 Fluorene 4.5 0.9957 0.003 Phenenthrene 9.0 0.9968 0.001 Anthracene 8.8 0.9962 0.002 Fluoranthene 7.9 0.9977 0.003 Pyrene 5.8 0.9967 0.003 Table 4.2. Recoveries and RSDs of PAHs using C12PPPOH-coated fiber & commercial fibers. SPME conditions; 10 mL of 5 µg/L spiked seawater, extraction time 30 min, stirring speed 105 rad s-1, 10 % NaCl, pH was not adjusted. Analytes C12PPPOH-coated SPME Relative RSD recovery (%) (%) PDMS-DVB-coated SPME Relative RSD recovery (%) (%) PA-coated SPME Relative RSD recovery (%) (%) Naphthalene 81.9 8.2 85.2 7.5 79.0 8.1 Acenaphthene 88.5 7.9 86.6 6.8 85.8 7.7 Fluorene 91.5 5.5 95.2 8.0 87.6 8.0 Phenenthrene 95.3 7.1 96.0 5.6 92.4 5.1 Anthracene 90.2 7.0 92.2 6.6 88.1 7.7 Fluoranthene 82.7 6.8 79.1 5.1 72.3 6.5 Pyrene 91.8 4.9 96.3 5.7 90.3 6.7 78 Chapter 4 4.3.4. SPME/GC-MS of real water sample The new C12PPPOH-coated SPME was evaluated for the preconcentration/ extraction of PAHs in seawater samples collected from St. John’s Island. There are no detectable levels of target analytes in the sample. Therefore, recovery and matrix effects (selectivity) were studied on real water samples. Table 4.2 shows comparable results between the C12PPPOH-coated fiber and commercially available fibers. These results clearly indicate that seawater matrix has no pronounced effect on the SPME/GC-MS analysis of PAHs using C12PPPOH-coated fiber. 4.4. Conclusion A new solid-phase microextraction (SPME) sorbent material has been developed and optimized for the polycyclic aromatic hydrocarbons, organophosphorous pesticides and organochlorine pesticides. C12PPPOH-coated fiber was found to provide satisfactory results in comparison with commercially available fibers. More importantly, the new coating exhibited longer application life time and thermal stability up to 320oC. The excellent extraction efficiency of C12PPPOH-coating is most probably due to porous surface structure of the film and the possession of polar and non-polar functional groups on the either side of the polymer backbone. It provides an easy, simple, rapid and inexpensive SPME method for the target analytes with sufficient sensitivity and reproducibility. It can be concluded that amphiphilic C12PPPOH-coated fiber is a substitution for existing commercial coatings with high operational temperatures along with better analytical performance and longer lifetime. Additional work is underway to investigate the suitability of the coating for other applications such as headspace SPME, 79 Chapter 4 combinations with HPLC and CE, and other types of analytes, particularly, environmental pollutants. References: [1] F. Wania, D. Mackay, Environ. Sci. Technol. 30 (1996) 390 [2] G. M. Rand, Fundamentals of Aquatic Toxicology, 2nd ed., Taylor and Francis, Washington, DC, 1995 [3] M. A. Kamrin and R. K. Ringer, Toxicol. Environ. Chem., 41 (1994) 63 [4] M. Kawano, Y. Tanaka, J. Falandysz, R. Tatsukawa, Brom. Chem. Toksykol. 30 (1997) 87 [5] Commission of the European Communities, EEC Drinking Water Guidelines, 80/799/EEC, EEC No. L229/11-29, EEC, Brusels, 30 August 1980 [6] Program Manager Rocky Mountain Arsenal, Record of Decision for the On-Post Operable unit, Sections 1–11, Version 2.0, vol. 1, Rocky Mountain Arsenal, Commerce City, CO, March 1996 [7] Book of Lists for Regulated Hazardous Substances, Government Institutes, Rockville, MD, 1990, p. 51. Taken from Part 264, Title 40, Appendix IX, US GPO, Washington, DC [8] M. Cleemann, F. Riget, G. B. Paulsen, J. Klungsoyr, R. Dietz, Sci Tot. Environ. 245 (2000) 87 [9] L. Asplund, B. G. Svensson, A. Nilsson, U. Eriksson, B. Jansson, S. Jensen, U. Wideqvist, S. Skerfving, Arch. Environ. Health 49 (1994) 477 [10] J. R. 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This thesis compiled the developments of three important microextraction approaches including liquid-liquid-liquid microextraction (LLLME), micro solid-phase extraction (µ-SPE) combined with high performance liquid chromatography (HPLC-UV) and amphiphilic poly(p-phenylene)s (C12PPPOH) coated solid-phase microextraction (SPME) combined with gas chromatography mass spectrometry (GC-MS). Each of the three different approaches was applied to the real samples and the results obtained from this work clearly demonstrated the applicability of our approaches. In the first section, we have discussed a study of the suitability of ionic-liquid supported hollow fiber membrane (HFM) protected LLLME as a single-step enrichment/clean-up approach. An advantage of this work was that it eliminates matrix effects normally encountered by other immersion-based microextraction techniques. An ionic liquid, 1-butyl-3-methylimidazolium hexafluorophosphate BMIM[PF6] was used as an acceptor phase for the first time in the HFM-protected LLLME. Since viscosity of BMIM[PF6] is too high, it was mixed with acetonitrile (ACN) to facilitate the extraction. BMIM[PF6]:ACN (1:1) was found to be the optimum extraction solvent. When this method was applied to the real wastewater samples, it was found out that wastewater matrix did not have a significant effect on the extraction efficiency and the recoveries of analytes obtained from the wastewater were higher than spiked pure water samples. Moreover, the final extract could be directly injected into the reversed phase HPLC. 84 Chapter 5 Therefore, this approach is simple, rapid, easy to use and the use of disposable HFM completely eliminate the carryover effect. In the second section, we have developed a novel micro-solid-phase extraction (µSPE) procedure and applied this approach to the determination of carbamates in tea samples. µ-SPE devices can be easily prepared by a porous polypropylene (PP) membrane sheet and the different types of sorbents were packed inside the devices. As mentioned in the first section, the use of porous membrane served as a cleanup device and prevented the matrix effects especially from the compounds with higher molecules. This approach could be used as an alternative to the traditional solid-phase extraction (SPE) techniques because the presented µ-SPE method is a simple, cost-effective and solvent minimized approach that is sensitive, selective and reproducible. In the third section, novel amphiphilic poly(p-phenylene)s (C12PPPOH) was used as sorbent for the first time for the solid-phase microextraction (SPME) of environmental pollutants. This C12PPPOH-coated fiber provided the higher extraction efficiencies for the determinations of polycyclic aromatic hydrocarbons (PAHs), organochlorine pesticides (OCPs) and organophosphorous pesticides (OCPs) from seawater samples compared to the results obtained from commercial coatings. An important advantage of this work was that the new coating exhibited longer application lifetime and higher thermal stability. Therefore, C12PPPOH-coated fiber could be used as a substitution for the commercial coatings. Future Work Ionic-liquid supported HFM protected LLLME technique could be applied to many research works in the future. In the petroleum industry, this technique can be used 85 Chapter 5 for the pilot scale separation of contaminants from petroleum products. Optically active compounds could be successfully separated by chiral ionic-liquids supported HFMLLLME from pharmaceutical products. Ionic-liquid can be coated to the HFM or PP membrane sheet and applied for LPME or µ-SPE approaches. Moreover, ionic-liquid can be coated inside the capillary column of the gas chromatograph for future applications. In the µ-SPE approach, the efficiency of µ-SPE device can be improved by coating different types of polymer to the PP membrane. µ-SPE can be combined with microwave-assisted (headspace) extraction for the extractions for volatile organic compounds from solid matrices. µ-SPE could be applied to the determinations of different types of analytes from different applications. In the final section, our new amphiphilic C12PPPOH-coated fiber proved that it is a better substitution for existing commercial coatings with high operational temperatures along with better analytical performance and longer lifetime. Additional work is underway to investigate the suitability of the coating for other applications such as headspace SPME, combinations with HPLC and CE, and other types of analytes, particularly, environmental pollutants. 86 [...]... with the use of reduced organic solvent and better automation with modern instruments have led to recent developments of miniaturized liquid-liquid extractions procedures 1.2.2 Flow Injection Analysis Flow injection analysis can be used to minimize the volumes of organic solvent required for LLE, as well as to automate the extraction process Using this technique, sample and solvent volumes of less than... point of the solvent, at atmospheric pressure, in closed vessels, the temperature may be elevated by simply applying the appropriate pressure 1.3.5 Supercritical fluid extraction Supercritical fluid extraction (SFE) is also a very popular technique for environmental analysis It is an appropriate technique for the analysis of the less volatile compounds, much like solvent extraction It has limitations for. .. on the nature of the matrix to be extracted Solid sample includes soils, sediments, fruits, meats, tissue, leaves, etc Currently available methods for organic environmental analysis are; a) Soxhlet extraction b) Automated Soxhlet extraction, Soxtec c) Pressurized fluid extraction d) Ultrasonic extraction e) Microwave-assisted extraction f) Supercritical fluid extraction g) Direct thermal extraction 1.3.1... volume of the extract is usually too large for direct injection for analysis and, in order to obtain sufficient sensitivity, an additional evaporation-concentration step, e.g using an apparatus (Kuderna-Danish) is necessary Particular care needs to be taken in both the solvent extraction and concentration procedures to avoid contamination of the sample and formation of emulsions [7-10] Thus, the demand for. .. techniques to preconcentrate them before analysis are need Recently, liquid-phase microextraction (LPME) a miniaturised approach to liquid-liquid extraction (LLE) has been introduced [3, 4] LPME through the use of a single drop of solvent [5, 6] or a short plug of solvent held within a porous hollow fiber membrane (HFM) [7], has been emerging as attractive extraction approaches in environmental and other analyses... Therefore, the selected solvent system and the operating conditions must usually be demonstrated to exhibit adequate performance for the target analytes in reference samples before it is implemented for the real samples The most common solvent system is acetone-hexane (1:1 v/v) but for nonpolar analytes such as PCBs, hexane alone can also be used 1.3.4 Microwave-assisted extraction Microwave-assisted extraction. .. The method of extraction is straightforward; solid or liquid sample is placed in a headspace autosampler (HSAS) vial of about 10 mL, and the volatile analytes diffuse into the headspace of the vial Once the concentration of the analyte in the headspace of the vial reaches equilibrium with the concentration in the sample matrix, a portion of headspace is swept into a gas chromatograph for analysis However,... significant 1.3.2 Pressurized fluid extraction A new technique, pressurized fluid extraction (PFE) appeared around 10 years ago It is called accelerated solvent extraction (ASE™, which is a Dionex trade mark), pressurized liquid extraction (PLE), pressurized solvent extraction (PSE) or enhanced solvent extraction (ESE) It was partly derives from supercritical fluid extraction (SFE) In PFE, the extractant... applicable to a variety of tasks ranging from pH or conductivity measurement to colorimetric and enzymatic assays Still, FIA has disadvantages compared to the latest micro-extractions techniques because the volumes of organic solvents used in FIA are still in the order of several milliliters for each analysis [14] 1.2.3 Liquid-Phase Microextraction The term “liquid phase microextraction” (LPME) was... radiation as the source of heating of the solvent sample mixture Due to the particular effects of microwaves on matter (namely dipole rotation and ionic conductance), heating with microwaves is instantaneous and occurs in the middle of the sample, leading to very fast extractions [5455] In most application, the extraction solvent is selected as the medium to absorb microwaves Alternatively (for thermolabile ... analysis) Notable among recent developments are simple, faster and greener (environmentally friendly) microextraction techniques This thesis focuses on the developments of solvent-minimized extraction. .. Supercritical fluid extraction (SFE) is also a very popular technique for environmental analysis It is an appropriate technique for the analysis of the less volatile compounds, much like solvent extraction. .. Limited number of polymeric coatings for SPME- lack of fibers that are sufficiently polar 15 Chapter 1.3 Extraction of Organics from Solid Matrices The extraction and recovery of a solute from