Development of the Space-Resolved Solid-Phase Microextraction

Publication Date (Web): July 30, 2009 ... desorption protocols offer potential advantages within high throughput applications. .... Modern Extraction ...
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Anal. Chem. 2009, 81, 7349–7356

Development of the Space-Resolved Solid-Phase Microextraction Technique and Its Application to Biological Matrices Xu Zhang,† Jibao Cai,†,‡ Ken D. Oakes,§ Franc¸ois Breton,† Mark R. Servos,§ and Janusz Pawliszyn*,† Department of Chemistry and Department of Biology, University of Waterloo, Ontario, N2L 3G1, Canada, and Department of Chemistry, University of Science and Technology of China, Hefei, 230026, China To facilitate rapid in situ analyte monitoring within heterogeneous samples, a space-resolved solid phase microextraction (SR-SPME) technique was developed that utilized miniaturized segmented fibers. Initially, a multilayered agarose gel was used to determine the effects of diffusion-based mass transfer and fiber dimension on the space-resolving capability of SPME. For diazepam within agarose gel, the SR-SPME limit of detection was 2.5 ng/ mL, with a linear dynamic range up to 500 ng/mL. The efficacy of the SR-SPME technique was further evaluated within diverse biological matrices (onion bulb, fish muscle, and adipose tissues) containing stratified pharmaceutical analytes. Empirically, the results agreed well with established techniques such as microdialysis and liquid extraction, but SR-SPME was simpler to implement, displayed higher spatial resolution, and was more cost-effective than traditional approaches. Additionally, the segmented design of the SPME fibers and stepwise desorption protocols offer potential advantages within high throughput applications. As a fast, simple, and solvent-free sampling and sample preparation method, solid-phase microextraction (SPME) has been widely employed in numerous applications ranging from environmental studies to in vivo pharmacokinetics.1-8 To date, most SPME applications have been restricted to relatively homogeneous sample systems, such as water, air, or blood.4-8 Studies on some heterogeneous matrices (such as soil) can also benefit from the operational convenience and reduced matrix effects afforded by * Corresponding author. Tel: +1-519-888-4641. Fax: +1-519-746-0435. E-mail: [email protected]. † Department of Chemistry, University of Waterloo. ‡ University of Science and Technology of China. § Department of Biology, University of Waterloo. (1) Arthur, C. L.; Pawliszyn, J. Anal. Chem. 1990, 62, 2145–2148. (2) Pino, V.; Ayala, J. H.; Gonzalez, V.; Afonso, A. M. Anal. Chem. 2004, 76, 4572–4578. (3) Pawliszyn, J. (Ed.) Applications of Solid Phase Microextraction; RSC: Cornwall, UK; 1999. (4) Lord, H.; Grant, R.; Walles, M.; Incledon, B.; Fahie, B.; Pawliszyn, J. Anal. Chem. 2003, 75, 5103–5115. (5) Gorecki, T.; Martos, P.; Pawliszyn, J. Anal. Chem. 1998, 70, 19–27. (6) Liu, Y.; Shen, Y.; Lee, M. L. Anal. Chem. 1997, 69, 190–195. (7) van Eijkeren, J. C. H.; Heringa, M. B.; Hermens, J. L. M. Analyst 2004, 129, 1137–1142. (8) Setkova, L.; Risticevic, S.; Pawliszyn, J. J. Chromatogr. A 2007, 1147, 224– 240. 10.1021/ac900718q CCC: $40.75  2009 American Chemical Society Published on Web 07/30/2009

SPME through the use of the headspace-SPME technique.9 Within the SPME analysis scenarios previously utilized, spatial resolution is constrained to the single longitudinal dimension of the sampled matrix. However, within biological systems, the distribution of analytes of interest can vary markedly between adjacent tissues, and greater spatial resolution of the presence and free concentrations of contaminants, drugs, and other bioactive compounds is often desirable. Further, from both an animal welfare and cost perspective, the ability to spatially resolve bioactive compounds in living tissues by nonlethal sampling would be a significant advantage. Traditional SPME approaches offer poor spatial resolution due to the relatively large size of the fiber coating relative to the tissue being sampled. For example, commercially available SPME fibers were used to study the concentration and translocation of herbicides in a living onion,10 but the results were largely qualitative for two reasons. First, commercial SPME fibers (∼1 cm fiber length) are limited in their spatial resolution, as analyte concentrations are averaged along the 1 cm fiber axis (across three to five tissue layers) within the heterogeneous onion structure. Second, the detected tissue concentrations were timeaveraged over an interval of 1 h, during which diffusion of the analytes within the tissue might obscure the concentration difference between the sampling location and surrounding tissues. For example, with an analyte diffusion coefficient in onion of 1 × 10-9 m2/s, the diffusion distance would be approximately 8.5 cm over the 1 h sampling time. As the diffusion distance is much larger than the 1 cm fiber length, the concentrations detected in the onion tissue are effectively averaged over time and space. Uneven distribution of substances within biological matrices is more common than uniform distributions due to the spatial heterogeneity produced by differential uptake, storage, and metabolism of energy stores as well as specialized tissue function at higher levels of biological organization.11,12 Monitoring of dynamic physiochemical processes in living systems consequently dictates the use of an approach possessing both a high spatial resolution and a rapid, relatively noninvasive sampling methodol(9) Zuliani, T.; Lespes, G.; Milacic, R.; Scancar, J.; Potin-Gautier, M. J. Chromatogr. A 2006, 1132, 234–240. (10) Lord, H.; Moeder, M.; Popp, P.; Pawliszyn, J. Analyst 2004, 129, 107–108. (11) Brooks, B. W.; Chambliss, C. K.; Stanley, J. K.; Ramirez, A.; Banks, K. E.; Johnson, R. D.; Lewis, R. J. Environ. Toxicol. Chem. 2005, 24, 464–469. (12) Huggett, D. B.; Cook, J. C.; Ericson, J. F.; Williams, R. T. Hum. Ecol. Risk Assess. 2003, 9, 1789–1799.

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ogy. To address these in situ sampling challenges, space-resolved SPME (SP-SPME) was theoretically conceptualized and validated by a series of experiments. A comprehensive assessment of the efficacy of this approach was conducted by evaluating the concentration distribution of pharmaceuticals in an onion bulb and fish tissues, with results validated against established microdialysis (MD) and liquid extraction (LE) approaches. THEORETICAL CONSIDERATIONS The spatial resolution of an in situ sampling technique is defined both as its capability to determine accurately local concentrations of analytes and to clearly resolve two different concentrations spatially close to each other. The spatial resolution of SPME is mainly determined by the dimension of its extraction phase, because, in principle, the sample concentration monitored by SPME is spatially averaged over the area where the extraction phase (fiber) is in direct contact with the sample matrix. Thus, spatial resolution can be improved by reducing the fiber size. Sampling time, while determining the temporal resolution of SPME during the monitoring of dynamic analytes, also affects spatial resolution. Diffusion of analyte molecules during longer sampling intervals tends to average concentration gradients in adjacent tissues, thus negating any benefits of a reduced fiber size. Consequently, neither temporal nor spatial resolution can be considered independently when improving the resolution of SPME under many sampling scenarios. A potential pitfall of fiber miniaturization and rapid sampling is that the sensitivity of the SPME technique can be adversely affected by excessive reductions in either factor. According to preequilibrium SPME theory, the extracted amount of analyte is a function of sampling time, as follows,13 n ) KfsVfCs[1 - exp(-at)]

(1)

where Cs is the sample concentration, Vf is fiber volume, a is the time constant, Kfs is the partition coefficient of the analyte between the fiber and the sample matrix, and t is the sampling time. Equation 1 describes the extraction kinetics when the sample volume is much larger than the product of the volume of the fiber coating and Kfs, which is often the case for in vivo sampling with miniaturized SPME probes. When the extracted amount of analyte, n, approaches the quantitation limit of the instrument (the minimal amount of analyte that generates a significant signal with the instrument), Q, the relationship between the spatial and temporal resolution of a SPME fiber can be described as equivalent to the relationship between lm, the minimal fiber length that can extract a quantifiable amount of analyte in a columnar fiber with a cross-sectional coating area, S, and minimal sampling time, tm, as described quantitatively in the following equation. lm )

Q SKfsCs[1 - exp(-atm)]

(2)

This equation indicates that sampling time has a negative correlation with fiber length when all the other conditions are fixed. Further, spatial and temporal resolution are related to (13) Ai, J. Anal. Chem. 1997, 69, 1230–1236.

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sample concentration, instrument sensitivity, cross-sectional area of the fiber, partitioning coefficient, and the time constant. For example, when the sample concentration or the partitioning coefficient is high, it takes less time to extract a detectable amount of analyte; under these conditions, even if the fiber is shorter, the extracted amount of analyte is sufficient for detection by the instrument. It also follows that, if using an instrument with improved sensitivity, the size of the fiber and sampling time can both be reduced. When the sampling time, t, is short (as described in the present study) and the amount of analyte to be extracted falls within the linear region of the extraction time profile, eq 2 can be approximated by eq 3 using the first-order Taylor expansion. lm )

Q SKfsCsatm

(3)

and eq 3 can be rearranged to eq 4. lmtm )

Q SKfsCsa

(4)

Equation 4 clearly demonstrates the negative correlation between the minimal sampling time and fiber dimension when the sampling conditions such as fiber material, sample concentration, and instrument performance are fixed. Accordingly, conditions under which rapid extraction SR-SPME can achieve sufficient analyte for analysis should include a fast equilibrium time and/ or high affinity of the fiber coating for the analyte (as shown by Kfs value). Another important SR-SPME parameter is the diffusion length of analyte molecules within the sample matrix, x, since this process affects the concentration distribution in heterogeneous matrices. This diffusion length could be described by the solution in one dimension to the diffusion equation (Fick’s first law of diffusion) x2 ) 4Dt

(5)

where t is the time duration of the molecule migration via diffusion and D is the diffusion coefficient, which is defined by the amount of substance diffusing across a unit area through a unit concentration gradient per unit time. To ensure representative in situ sampling in a well-defined location, the distance of molecule migration, x, in the sample matrix should be shorter than the minimal fiber length, lm; otherwise, the sphere of molecule diffusion is larger than the probe size. lm > x

(6)

For example, if the diffusion coefficient of a drug molecule in a semisolid sample is 1 × 10-9 m2/s and the fiber length is 1 mm, the maximal sampling time could be calculated as 500 s. Under these conditions, a sampling time exceeding 9 min would draw analyte molecules a distance greater than 1 mm, negating some spatial resolution benefits of the 1 mm fiber length. EXPERIMENTAL SECTION Chemicals and Materials. All chemicals purchased were of the highest possible purity and used without further purification.

Gemfibrozil, atorvastatin, ibuprofen, carbamazepine, diclofenac, naproxen, and bisphenol A (BPA) were ordered from SigmaAldrich (Oakville, ON, Canada). Diazepam, fluoxetine, lorazepam, and atrazine were obtained from Cerilliant Corp. (Round Rock, TX). Isotopically labeled standards (gemfibrozil-d6, atorvastatind5, ibuprofen-d3, carbamazepine-d10, diclofenac-d4, 13C1-naproxend3, fluoxetine-d5, diazepam-d5, atrazine-d5 and BPA-d16) were purchased from CDN Isotopes Inc. (Pointe-Claire, Quebec, Canada). For the in vivo experiment, 0.1% ethyl 3-aminobenzoate methanesulfonate (Sigma-Aldrich) was used to anaesthetize the fish. HPLC-grade acetonitrile for the HPLC mobile phase, methanol for standard preparation and desorption solutions, formaldehyde, ethanol, and acetic acid (glacial) were all purchased from Fisher Scientific (Unionville, ON, Canada). For preparation of C18-coated fibers, the C18 particles (10 µm) and polyethylene glycol (PEG) were obtained from Supelco (Bellefonte, PA). Preparation of the Miniaturized SPME Probes. A schematic representation of a miniaturized SPME probe is illustrated in Figure 1A. For analysis within agarose gel and onion tissues, C18-coated fibers were utilized due to their proven performance in these applications and were prepared as previously described.14,15 Stainless steel wire (127 µm o.d.) was purchased from Small Parts Inc. (Miami Lakes, FL) and were cut into 10 cm lengths with the terminal 2 cm etched with 400-grit silicon carbide polishing paper. Wires were then sonicated in acetone for 20 min and rinsed with deionized water to remove oxides or other contaminants on the wire surface prior to being dried at room temperature for 30 min. The bonded phase silica particles were mixed thoroughly with PEG in the ratio of 60:40 (w/w) and then packed into a 200 µL micropipet tip. The wire was carefully introduced into the tip from

the small opening and withdrawn to form a uniform coating layer. The coated metal wires were preheated at 70 °C for 10 min and then conditioned at 200 °C for 2 h. The fibers were subsequently cooled to room temperature and carefully trimmed with a blade to form the segmented coating configuration shown in Figure 1A. Before experimental use, the fibers were conditioned in 50% acetonitrile and 50% deionized water for 4 h. Home-made poly(dimethylsiloxane) (PDMS) fibers were employed for in vivo fish applications using Helix medical silicone tubing (0.31 mm i.d., 0.64 mm o.d.; Carpinteria, CA) as the extraction phase. Previous use of PDMS fibers yielded good sensitivity and reproducibility (with no biofouling) during in vivo monitoring of pharmaceuticals in fish.16,17 Both C18 and PDMS fibers were cut into 1 mm long segments, the latter supported internally by a 3.5 cm long stainless steel wire (0.483 mm o.d.; Small Parts Inc.). Individual fiber segments were separated by a 5 mm (C18 fibers) or 4 mm (PDMS fibers) space to allow for simultaneous sampling of two differing tissues or stratified matrices (Figure 1A). All fibers were preconditioned in 100% methanol for 24 h, followed by sterile nanopure water for 2 h prior to in vivo application. For rapid sampling with kinetic calibration, fibers were preloaded with deuterated standards (i.e., diazepam-d5) by immersing the probes for 30 min in a 50 µg/L loading solution prepared by spiking deuterated standard into 25 mL of sterile deionized water. The preloaded SR-SPME probes were subsequently stored in clean test tubes at 4 °C for up to 24 h prior to use. LC-MS/MS Analysis. A CTC-PAL autosampler/Shimadzu 10 AVP LC/MDS Sciex API 3000 tandem mass spectrometry (MS) system was used for the analysis of diazepam and its deuterated standards as previously described.4 Compounds other than diazepam were quantified with an Agilent 1200 HPLC/MDS Sciex 3200 Q-trap tandem MS system as described in detail elsewhere.15 The transitions monitored were as follows: diazepam, 285/154; fluoxetine, 310/44; carbamazepine, 237/195; gemfibrozil, 249/121; ibuprofen, 205/161; atrazine, 216/174; atorvastatin, 559/440; naproxen, 229/169; diclofenac, 294/250; bisphenol A, 227/212; gemfibrozil-d6, 255/121; atorvastatin-d5, 564/445; ibuprofen-d3, 207/164; carbamazepine-d10, 247/204; diclofenac-d4, 298/217; 13 C1-naproxen-d3, 233/169; fluoxetine-d5, 315/44; diazepam-d5, 290/198; atrazine-d5, 221/179; and BPA-d16, 241/142. In Vitro Simulation of Heterogeneous Samples Using Agarose Gel. Agarose gel (1%) prepared in phosphate-buffered saline (PBS, pH 7.4) was used to simulate the analyte gradients found in many biological tissues. In the first experiment, a series of nine individual 8 mL gels containing a range of diazepam concentrations (0.1-500 µg/L) was cast in 10 mL screw cap vials (Supelco, Bellefonte, PA), with each vial containing an individual concentration. The extraction behavior of the SPME probes including the determination of extraction and desorption time profiles, matrix effects of the agarose gel, the dynamic range of extraction, and verification of symmetry between extraction and desorption were all conducted in the gel matrices. In a second experiment, additional 1% agarose gels were cast (20 mm thick) within a round (65 mm diameter) Pyrex crystal-

(14) Es-haghi, A.; Zhang, X.; Musteata, F.; Bagheri, H.; Pawliszyn, J. Analyst 2007, 132, 672–678. (15) Zhang, X.; Es-haghi, A.; Musteata, F.; Ouyang, G.; Pawliszyn, J. Anal. Chem. 2007, 79, 4507–4513.

(16) Hutchinson, J. P.; Setkova, L.; Pawliszyn, J. J. Chromatogr. A 2007, 1149, 127–137. (17) Zhou, S. N.; Oakes, K. D.; Servos, M. R.; Pawliszyn, J. Environ. Sci. Technol. 2008, 42, 6073–6079.

Figure 1. The segmented SR-SPME fiber and the two-step desorption process in a 96-well plate for the extracted analyte and calibrant: (A) the segmented fiber, (B) the fiber placed into 50 µL of MeOH (100%) in a well for 2 min to desorp the analyte from the first (lower) coating segment, (C) the fiber placed into another well containing 200 µL of MeOH (100%) to desorp the analyte from the second (upper) coating segment.

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Figure 2. Study of diffusion-controlled mass transfer. Agarose gel (1%) was cast in a round crystallization plate to a depth of 20 mm. SPME fibers (3 fibers/location) were placed along the circumference of the plate center with a radius of 15 mm. A 45° angle separated adjacent sampling locations, which are labeled on the bottom of the plate numbers from 1 to 8. Diazepam (in 10 µL MeOH) was applied in the plate center at t ) 0.

Figure 3. C18 SR-SPME fibers and microdialysis probes were inserted in a multilayered gel system (as illustrated) to determine diazepam distribution.

lization plate to determine the diffusion velocity of diazepam at both room temperature and at 3.5 °C (Figure 2). C18 fibers (10 mm in length) were placed at 45° intervals around the perimeter of the gel, at a radius of 15 mm. The SPME fiber coatings were introduced to a depth of roughly 15 mm into the gel, and care was taken to ensure that the fiber coating did not touch the bottom of the plate. Diazepam (20 µg in 10 µL MeOH) was applied in the geometric center of the gel, where a 22 gauge needle created a hole with an approximate diameter and depth of 1 and 10 mm, respectively. SPME fibers were removed sequentially at 45 min intervals and then desorbed in 200 µL of HPLC grade MeOH for 30 min. Finally, the MeOH was evaporated under nitrogen gas, and the sample was reconstituted in 30 µL of 50% acetonitrile:50% nanopure water containing 7.5 ng/mL lorazepam as internal standard. In the third experiment, a multilayer gel was cast to simulate a heterogeneous sample system (Figure 3). Three 1.5% agarose gel layers were cast in three plates. In the first plate, the gel layer was 2 mm thick and contained 100 ng/mL of diazepam; the second gel layer was 5 mm thick and had no diazepam, while the third layer was only 1 mm thick and contained 200 ng/mL of diazepam. The thickness of the gel layers was controlled by their volume during casting. After 1.5 h of solidification at room temperature, a 4 cm2 section of gel was excised from each plate and stacked 7352

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to form a three-layer gel with stratified diazepam concentrations of 100, 0, and 200 ng/mL (from bottom to top). Once the layered gel system was constructed, four segmented C18 fibers (preloaded with deuterated diazepam) were carefully introduced into the gel such that the two fiber segments were positioned precisely within the two layers containing diazepam for a 5 min interval. To avoid contamination, the fiber segments were protected with a needle when introduced and withdrawn from the gel. Following fiber withdrawal, a two-step successive desorption of the extracted analyte and calibrant from each of the two coating segments was performed into two separate wells of a 96-well plate. The first fiber segment was desorbed into a well containing 50 µL of 100% MeOH for 2 min, where the level of the MeOH was sufficient to immerse only the first fiber segment located at the tip of the wire (Figure 1b). Subsequently, the fiber was placed into a second well containing 200 µL of 100% MeOH for 2 min, a volume sufficient to immerse the second fiber segment for desorption. The MeOH in both wells was evaporated, and the samples were reconstituted into 30 µL of 50% acetonitrile:50% nanopure water (containing 7.5 ng/mL lorazepam as internal standard) prior to analysis. To validate the SR-SPME quantification with an established technique, two microdialysis probes were placed into the two diazepam-containing gel layers while performing SPME sampling. The microdialysis perfusion fluid was PBS buffer, with a flow rate of 2 µL/min. Dialysates were collected in 5 min intervals and mixed with 20 µL of acetonitrile for LC-MS/MS analysis. Quantification of the analyte concentration in microdialysis samples was based on the recovery of the analyte, Cdialysate/Cperfusate.18,19 In Vitro Application: Diazepam in an Onion Bulb. To evaluate the effectiveness of SR-SPME within relevant biological matrices, an onion bulb (∼6 cm in diameter) was chosen as a representative heterogeneous vegetative tissue due to its layered structure (Figure 4). To simulate an analyte gradient, as occurs among many vegetable crops due to differential accumulation of compounds,20 0.2 mL of 1 µg/mL diazepam (in MeOH:water, 10: 90 v/v) was injected into the center of the onion bulb from the stem side 4 h prior to sampling. On a lateral surface of the onion (perpendicular to the stem), two parallel holes with a depth of 19 mm were made with a 22 gauge needle. The segmented SPME fiber was introduced into one hole (to simultaneously measure analyte in two layers), while a 4 mm microdialysis probe (CMA/ 12, CMA/Microdialysis AB, Stockholm, Sweden) was placed into the other (Figure 4). After 5 min, the SPME fiber was removed and briefly rinsed with nanopure water to remove any adhering material, followed by the successive two-step desorption procedure described earlier. Segmented and nonsegmented 4 mm SPME fibers were inserted three times sequentially into the same holes in the side of the onion to assess the reproducibility of the technique. The microdialysis probe was kept in position during the SPME sampling, so that dialysate samples were collected in parallel with each SPME sampling (5 min for each sample). The microdialysis perfusion fluid, flow rate, and the calibration procedure were identical to that described for the gel experiment. (18) de Lange, E. C. M.; de Boer, A. G.; Breimer, D. D. Adv. Drug Delivery Rev. 2000, 45, 125–148. (19) Andren, P. E.; Emmett, M. R.; DaGue, B. B.; Steulet, A.; Waldmeier, P.; Caprioli, R. M. J. Mass Spectrosc. 1998, 33, 281–287.

Figure 4. Illustration of the application of SR-SPME and microdialysis for in situ sampling of diazepam within an onion bulb.

In Vivo Study of the Distribution Coefficient of Pharmaceuticals between Adipose and Muscle Tissue in Fish. The effectiveness of SR-SPME within adjacent fish adipose and muscle tissue was investigated as an example of an animal matrix that differentially accumulates analyte. All experimental procedures involving animals were conducted in the Biology Department Wetlab Facility at the University of Waterloo in accordance with protocols approved by the institutional Animal Care Committee (AUP # 07-16). The juvenile rainbow trout (Oncorhynchus mykiss) used in this study were 19.8-24.5 cm in length and 80.0-134.2 g in weight (n ) 20). Of these 20 fish, nine were randomly divided into three groups (three fish per 34 L aquaria) and exposed for 8 d to water spiked with 3 ng/mL of each compound in the multianalyte mixture of pharmaceuticals and other bioactive compounds (atorvastatin, atrazine, BPA, carbamazepine, diclofenac, fluoxetine, gemfibrozil, ibuprofen, and naproxen), while three fish were treated in the same way but exposed to water spiked with MeOH as a solvent control. Another eight fish were held in reference diluent water (without test compound or solvent addition) to serve as a vehicle and test compound control. The in vivo sampling was conducted using pre-equilibrium SPME with kinetic calibration. The sampling procedure with kinetic calibration was as described previously with the exception of the SR-SPME deployment in the fish.15 Briefly, after the fish was anaesthetized (0.1% ethyl 3-aminobenzoate methanesulfonate) until loss of vertical equilibrium, a 20 gauge needle was used to pierced the fish at an angle approximately 30°-45° (from the vertical) through the adipose fin and into the dorsal epaxial muscle, penetrating the latter tissue approximately 8 mm in depth. Subsequent to removing the needle, the SPME fiber was introduced into the hole until the two coating segments were positioned in muscle and adipose fin tissue, respectively. The fiber coating was embedded in the tissue and the close contact between the tissue and fiber prevented water entry. After fiber placement, the fish was placed into fresh reference water for 8 min. The fish was then reanaesthetized prior to removing the fiber coating, for a total contact time between the fiber and tissue of 10 min. After a brief rinsing with deionized water and drying with a Kimwipe

tissue, the fiber was put into a polypropylene microcentrifuge tube prior to same-day desorption and instrumental quantification. For kinetic calibration, the amount of standard loaded on the fiber, q0, was determined from parallel preloaded fibers not introduced into a fish. The amount of analyte remaining on the fiber coating after exposure to the fish, Q, and amount of analyte extracted (pre-equilibrium) by the fiber, n, were determined simultaneously from the preloaded fibers used for fish sampling. Thus, the amount of tissue analyte available to be extracted at equilibrium, ne, could be calculated by the symmetric equation 1 - Q/q0 ) n/ne.21 To validate the in vivo fast SR-SPME results, the analyte concentrations in fish tissues were measured by both ex vivo equilibrium SPME and liquid extraction (LE) with methanol. For ex vivo SPME analysis, fibers were deployed under static conditions in tissues for 15 h at room temperature, conditions deemed sufficient to achieve equilibrium based on extraction time profiles from earlier in vitro experiments in PBS buffer (carbamazepine, 14 h; fluoxetine, 15 h; gemfibrozil, 4 h; ibuprofen, 3 h; atrazine, 12 h; atorvastatin, 4 h; naproxen, 3 h; diclofenac, 3 h; and bisphenol A, 3 h). At the end of the ex vivo exposure, the fibers were collected and rinsed briefly with deionized water prior to desorption in methanol. For LE analysis, whole fish tissues were cut into approximately 2 mm2 pieces with a scalpel on tared aluminum foil with tissue mass determined in an analytical balance prior to transfer to a preweighed microcentrifuge tube. Within the microcentrifuge tube, 500 µL of MeOH containing 20 ng/mL isotopically labeled standards (for the nine compounds described in the Chemicals and Materials section) was added to each sample prior to homogenates being generated (4 × 20 s/round with a Teflon homogenizer). Following a brief vortex, low-temperature centrifugation (4 °C, 15 000 rpm, 30 min) separated the tissue pellet from the supernatant, the latter of which (100 µL) was transferred into a 200 µL polypropylene insert within a 2 mL amber vial. Finally, 60 µL of water containing 20 ng/mL lorazepam (the internal standard) was briefly vortexed into the mixture to produce the final sample for instrumental analysis. Extraction efficiency was

(20) Zayed, A.; Lytle, C. M.; Qian, J.-H.; Terry, N. Planta 1998, 206, 293–299.

(21) Chen, Y.; Pawliszyn, J. Anal. Chem. 2004, 76, 5807–5815.

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evaluated from the relative recoveries of the deuterated standards spiked into control tissue samples. RESULTS AND DISCUSSION Drug Diffusion and SPME Extraction in the Gel Medium. In the present study, pre-equilibrium SPME techniques were adopted to obtain near real-time concentrations. While rapid SPME sampling can avoid some influences of diffusion on measured analyte concentrations, each analyte will have a unique (and temperature-dependent) diffusion coefficient within each sample matrix. Consequently, at the onset of the SR-SPME in vitro evaluations, we determined the linear velocity and diffusion coefficient of diazepam in gel medium at room temperature and 3.5 °C (Figure 2). It was found that diazepam migrated slowly through the 1% agarose gel matrix and was not extracted by the SPME fibers at the gel periphery until 5 h had elapsed under room temperature (8.4 h in 3.5 °C). On the basis of diazepam traveling 15 mm in 5 h within the agarose gel, the diffusion velocity at room temperature was calculated to be ∼0.84 µm/s. Over a 5 min sampling duration, such diffusion-based changes in analyte concentration would be insignificant. Accordingly, 5 min was selected as the sampling time for our assessments within the multilayered gel system. Additionally, the diffusion coefficient, D, can be determined using the SPME technique according to eq 5. The calculated diffusion coefficient was in the range of (2.0 ± 0.3) × 10-9 m2/s (n ) 3). It should be noted that the only source of concentration variation within our gel model system is physical diffusion. Diffusion was slower at lower temperatures; for example, at 3.5 °C the D value was ∼60% that measured at room temperature. However, for in vivo applications (such as detecting a metabolite in a living animal), the situation would be more complex, as circulatory systems can actively transport analyte and metabolism can play as central a role as physical diffusion in varying the drug concentration. As for in vivo diffusion (if analyte movement due to circulation is disregarded), the higher tortuosity of semisolid tissue relative to the gel would tend to impede diffusion and make the diffusion coefficient smaller than that calculated from the gel simulation.22 Another factor affecting mass transfer is the presence of a binding matrix, such as proteins, within in vivo systems; recent studies suggest that mass transfer may be facilitated by the presence of the binding matrix as a diffusion layer effect.23-25 Thus, temporal resolution in situ is affected by a number of variables, the complexity of which is magnified under in vivo, relative to in vitro or ex vivo, scenarios. Nevertheless, reducing sampling time (real-time sampling) is necessary to accurately monitor analyte concentrations within dynamic systems. The multilayered gel system, as a simulation of heterogeneity within more complex matrices, clearly demonstrates the spaceresolving capability of the new SPME technique. Diazepam concentrations detected in the spiked gel layers (200 and 100 ng/ mL of diazepam) were very comparable using SPME (188 ± 13 (22) Benveniste, H. J. Neurochem. 1989, 52, 1667–1679. (23) Heringa, M. B.; Hermens, J. L. M. Trends Anal. Chem. 2003, 22, 575– 587. (24) Oomen, A. G.; Mayer, P.; Tolls, J. Anal. Chem. 2000, 72, 2802–2808. (25) Kopinke, F.-D.; Georgi, A.; Mackenzie, K. Acta Hydrochim. Hydrobiol. 2000, 28, 385–399.

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Figure 5. The normalized absorption and desorption time profiles for diazepam (2) and its deuterated analogue, diazepam-d5 ([).

and 93 ± 11 ng/mL) and MD (191 ± 15 and 91 ± 12 ng/mL, respectively). The relative recovery (RR) of SPME was 91-114%, while the RR for MD was 94-97.5%. Kinetic Calibration for in Situ Sampling. Under preequilibrium extraction conditions, kinetic calibration outperforms the calibration curve approach in that the former method provides more accurate and precise calibration. The kinetic calibration approach is especially suitable for real time in situ sampling,21 although verifying a symmetric relationship between extraction and desorption of the analyte is a prerequisite to its application. Consequently, absorption and desorption experiments were conducted simultaneously (using fibers preloaded with deuterated diazepam) within a gel medium containing 200 µg/L of nonlabeled diazepam (pH 7.4). By immersing the preloaded fibers for different intervals within the gel, the fraction of the labeled diazepam remaining in the extraction phase (fiber coating) after sampling times (t) could be measured, along with the extracted (nonlabeled) diazepam. The relative desorption fraction (1 - Q/q0) and extraction fraction (n/ne) was roughly equal at each time interval (Figure 5), demonstrating symmetry between adsorption and desorption. In addition, the requirement for negligible depletion of analyte (0.2 mL, only 2-4% of diazepam was extracted in 5 min. When C18-bonded silica particle (5 µm) coated fibers were selected as the extraction phase (under static extraction conditions), the detection limit using 1 mm C18 fibers was 2.5 ng/mL for diazepam (S/N = 3, n ) 6), and the dynamic range was up to 500 ng/mL. In Situ Analysis of a Layered Plant Tissue by SPME with Microdialysis Validation. To evaluate the in situ efficacy of SRSPME within a plant tissue containing analyte gradients, local concentrations of diazepam were quantified using SR-SPME within two tissue layers (labeled as 1 and 2 in Figure 4), differing in depth within a single hole in an onion bulb. The calculated SR(26) Vaes, W. H. J.; Ramos, E. U.; Verhaar, H. J. M.; Seinen, W.; Hermens, J. L. M. Anal. Chem. 1996, 68, 4463–4467.

Table 1. Distribution of the Test Compounds after 8 d of Exposurea

Kfm Kfa real Kam apparent Kam log(Kow)b

atrazine

gemfibrozil

carbamazepine

ibuprofen

fluoxetine

703.2 ± 105.9 404.3 ± 59.4 0.05 ± 0.01 0.08 ± 0.01 2.34

15.4 ± 3.7 1.1 ± 0.2 5.98 ± 0.79 81.33 ± 21.68 4.77

11.2 ± 2.1 5.0 ± 1.2 3.78 ± 0.41 8.32 ± 1.64 2.40

10.3 ± 2.1 0.9 ± 0.2 1.62 ± 0.23 18.45 ± 4.28 3.97

25.2 ± 3.3 0.8 ± 0.1 0.08 ± 0.03 2.37 ± 0.35 4.64

The K value uncertainties were calculated using propagation of uncertainties from the monitored concentration by LE and SPME (n ) 9). Literature values. a

b

SPME concentrations within the two layers were highest at the inner layer nearer the analyte injection site (628 ± 4 ng/mL) and lower within the distal layer (126 ± 6 ng/mL), as would be anticipated. The amount of analyte extracted at each layer did not differ between replicate extractions, but the 4-5 times difference in extracted analyte between layers 1 and 2 did suggest both the appropriateness of onion bulbs as a heterogeneous model and the capability of SR-SPME to resolve differences in adjacent vegetative tissues. Conversely, the analyte concentrations determined by the microdialysis probe (in a parallel, but otherwise identical hole) was 2 times lower than that of the 1 mm SPME fibers in the inner layer (site 1), but MD values were approximately twice as high as 1 mm SPME fibers deployed in the distal layer (site 2). At first glance, concentrations determined by SPME did not match those determined by MD, when in fact analyte concentrations determined by the longer MD probe (4 mm) could be regarded as the spatially weighted average of the concentrations determined by the two 1 mm SPME segments. This rationale was supported by deploying 4 mm long C18-coated SPME fibers into a parallel hole to the MD probe for 5 min extractions. The analyte concentrations obtained with the 4 mm SPME probes (268 ± 7 ng/mL) agreed with those obtained from the 4 mm MD probe (288 ± 11 ng/mL). Therefore, these results demonstrate that the in situ quantitative capability of SPME is comparable to that of MD, but with the added advantage of greater spatial resolution. Monitoring of the Distribution Coefficient of Pharmaceuticals in Two Types of Fish Tissue. The accumulation, fate, and tissue distribution of environmental pharmaceuticals in fish is an area of emerging research interest that may benefit from in situ SPME techniques.11,12 Application of SR-SPME within adjacent fish tissues that vary in lipophilicity (and hence analyte bioconcentration potential) could dramatically advance our ability to nonlethally monitor these compounds. Consequently, we determined the true distribution coefficients of a suite of pharmaceuticals and bioactive compounds in two fish tissues, relatively lipophilic adipose and relatively hydrophilic dorsal epaxial muscle tissue.27,28 The distribution coefficients between adipose fin and true muscle (K am ), which represent the relative bioavailability of the pharmaceuticals, were defined by the ratio of the free free concentrations (C free and C m ) as follows: a true ) Kam

Cafree free Cm

)

na /KfbVf na ) nm /KfbVf nm

(7)

(27) Kiessling, A.; Pickova, J.; Johansson, L.; Asgard, T.; Storebakken, T.; Kiessling, K. H. Food Chem. 2001, 73, 271–284. (28) Heltsley, R. M.; Cope, W. G.; Shea, D.; Bringolf, R. B.; Kwak, T. J. Environ. Sci. Technol. 2005, 39, 7601–7608.

Table 2. Monitored Tissue Concentrations in Adipose (Ca) and Muscle (Cm) by LE (n ) 9) and LE Recoveries (%, n ) 11) for Adipose (Ra) and Muscle (Rm)a atrazine

gemfibrozil

CBZ

ibuprofen

fluoxetine

Ca 1.32 ± 0.18 61.00 ± 9.76 10.82 ± 1.22 72.34 ± 9.76 345.44 ± 33.59 Ra 75 ± 8 67 ± 12 57 ± 11 64 ± 9 47 ± 10 Cm 15.55 ± 1.74 0.75 ± 0.16 1.30 ± 0.21 3.92 ± 0.74 145.70 ± 16.33 Rm 87 ± 7 73 ± 5 89 ± 19 71 ± 16 56 ± 12 a

Concentration unit, ng/g.

where Kfb was the distribution coefficient between the SPME fiber coating and PBS buffer; Vf is the fiber volume; nm and na were the extracted amounts of analyte from muscle and adipose fin (respectively) at equilibrium. apparent In addition, the apparent distribution coefficients (Kam ) could be obtained by the ratio of the total analyte concentrations total in these two tissues (C total and C m ), obtained by LE, as a described by the following equation, apparent total Kam ) Catotal /Cm

(8)

In the in vivo experiment, C total and C total a m were obtained using pre-equilibrium extraction coupled with kinetic calibration. These in vivo concentrations agreed with those detected by the ex vivo equilibrium SPME method, indicating the validity of the in vivo SR-SPME approach. No detectable amounts of test compounds were detected in either the solvent control (n ) 3) or in reference fish (n ) 8). The positive correlation between the Kam (Kapparent and K true am am ) and log(Kow) values is likely attributable to partitioning of higher Kow analytes into the relatively lipid-rich adipose to a greater extent than the lowerlipid muscle tissue, as shown in Table 1.29 Of the nine analytes comprising the exposure test mixture, five compounds (atrazine, gemfibrozil, CBZ, ibuprofen, and fluoxetine) were detected in fish tissues by SPME and LE, while the remaining four analytes (atorvastatin, naproxen, BPA, and diclofenac) were not detected, presumably reflecting a lack of bioconcentration potential as all analytes were detected in the fish exposure water. Ionization of several of these chemicals (atorvastatin, naproxen, and diclofenac) at physiological pH values in fish tissue may be the reason they did not bioconcentrate strongly.29 Nevertheless, the five detected analytes that were accumulated in fish tissues represent a diverse but routinely detected group of compounds, demonstrating the efficacy of SR-SPME as a promising detection technique in fish. CONCLUSION A high spatial resolution SPME approach, SR-SPME, was developed and evaluated, with results indicative of a facile Analytical Chemistry, Vol. 81, No. 17, September 1, 2009

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technique yielding high temporal resolution. Systematic testing over increasing levels of in vitro and in vivo complexity demonstrated the feasibility, accuracy, and efficiency of this approach. The segmented design of the SPME fibers and stepwise successive desorption procedure offer not only high spatial and temporal resolution but also increased capability for high-throughput parallel sampling with a single probe, which suggests the potential for depth-profiling studies in complicated biological systems. ACKNOWLEDGMENT This work has been financially supported by the Natural Sciences and Engineering Research Council of Canada, the

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Canadian Water Network, and the Canada Research Chairs Program. We thank Dr. Li Xu and Shirin Taheri-Nia for their kind help for developing the figures, Fatemeh Mirnaghi and Leslie Bragg for their help in the instrumental analysis, and the anonymous reviewers for their constructive comments, which improved the quality of the manuscript. Received for review April 4, 2009. Accepted July 7, 2009. AC900718Q (29) Meylan, W. M.; Howard, P. H.; Boethline, R. S.; Aronson, D.; Printup, H.; Gouchie, S. Environ. Toxicol. Chem. 1999, 18, 664–672.