Biocompatible Solid-Phase Microextraction Coatings Based on

Aug 9, 2007 - Optimization of the Coating Procedure for a High-Throughput 96-Blade Solid Phase Microextraction System Coupled with LC–MS/MS for ...
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Anal. Chem. 2007, 79, 6903-6911

Biocompatible Solid-Phase Microextraction Coatings Based on Polyacrylonitrile and Solid-Phase Extraction Phases Mihaela L. Musteata, Florin Marcel Musteata, and Janusz Pawliszyn*

Department of Chemistry, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1

The applications of solid-phase microextraction (SPME) are continuously expanding, and one of the most interesting current aspects consists of applying SPME for fast analysis of biological fluids. The goal of this study is to develop biocompatible SPME coatings that can be utilized for in vivo and in vitro extractions, in direct contact with a biological matrix such as blood or tissue. The biocompatibility of the proposed new coatings is confirmed by X-ray photoelectron spectroscopy, and their performance is tested by developing an SPME/HPLC method for analysis of verapamil, loperamide, diazepam, nordiazepam, and warfarin in buffer solutions and in human plasma. The coatings prove to be biocompatible by not adsorbing proteins and are successfully applied for fast drug analysis and assay of drug plasma protein binding. Drug monitoring often requires biocompatible devices and is highly dependent on the development of new analytical instruments or techniques. The most widely used methods for separation of drugs from biological samples are liquid-liquid extraction, solid-phase extraction, liquid-phase microextraction (LPME),1 membrane assisted extraction, ultrafiltration, dialysis, microdialysis, supercritical fluid extraction, affinity sorbent extraction, and solid-phase microextraction (SPME).2 Since its conception, the development of SPME has seen huge growth at both the fundamental and the application oriented levels.3-5 In the few years of its practice, SPME has developed to a mature technique and a useful alternative to contemporary techniques in various scientific and research fields. An impressive number of publications describe applications ranging from air and water analysis to in vivo sampling. The approach has gained interest in the environmental, forensic, food, biological, and pharmaceutical analysis fields,6 due to its robustness, ease of use, * Corresponding author. Phone: +1-519-8851211. Fax: +1-519-7460435. E-mail: [email protected]. (1) Pedersen-Bjergaard, S.; Rasmussen, K. E. J. Chromatogr., B 2005, 817, 3-12. (2) Heringa, M. B.; Hermens, J. L. M. Trends Anal. Chem. 2003, 22 (10), 575587. (3) Pawliszyn, J. Solid Phase Microextraction: Theory and Practice; Wiley-VCH: New York, 1997. (4) Pawliszyn, J., Ed. Comprehensive Analytical Chemistry; Vol. 37, Sampling and Sample Preparation for Field and Laboratory: Fundamentals and New Directions in Sample Preparation; Elsevier: Amsterdam, 2002. (5) Mullett, W. M.; Pawliszyn, J. J. Sep. Sci. 2003, 26, 251-260. (6) Lord, H. L.; Grant, R. P.; Walles, M.; Incledon, B.; Fahie, B.; Pawliszyn, J. B. Anal. Chem. 2003, 75 (19), 5103-5115. 10.1021/ac070296s CCC: $37.00 Published on Web 08/09/2007

© 2007 American Chemical Society

and successful coupling with GC (gas chromatography), LC (liquid chromatography), and CE (capillary electrophoresis).7-9 Although SPME is widely recognized, there are no commercially available biocompatible extraction phases. In fact, only custom-made coatings based on polypyrrole (PPY)6 and poly(ethylene glycol) (PEG)10 were used so far for in vivo drug analysis. The most difficult and undesirable problem is the adsorption of proteins and other macromolecules on the surface of SPME fibers. These macromolecules constitute a diffusion barrier and decrease the extraction efficiency in subsequent experiments.4 In order to transfer all SPME advantages to the field of drug analysis, it is imperative to develop new biocompatible devices suitable for extraction of polar and medium polar compounds from biological matrices. The biocompatibility of an artificial device introduced into the body can be generally defined as the compatibility with the living tissue with which it is brought into contact.11 Bioincompatibility leads to toxic reactions or immunological rejection. A material could be considered biocompatible if the sum of adverse humoral and cellular reactions occurring during exposure is lower than for a reference material.11,12 When an artificial material is placed in blood, a race for the surface starts immediately. Within a few milliseconds after the device is put in contact with the biological material, a layer made of water, proteins, and other biomolecules from the physiological fluid is formed at the surface of the device.13 When protein adsorption does not occur, the material is considered to be biocompatible. Biocompatible extractive materials that have been developed and used so far include restricted access materials (RAM),14-17 ionic liquids (IL),18-21 polydimethylsiloxane (PDMS),22 PPY,6 and (7) Jia, C.; Luo, Y.; Pawliszyn, J. J. Microcolumn Sep. 1998, 10 (2), 167-173. (8) Pawliszyn, J. Anal. Chem. 2003, 75 (11), 2543-2558. (9) Pawliszyn, J., Ed. Applications of Solid Phase Microextraction; Royal Society of Chemistry: Cambridge, U.K., 1999. (10) Musteata, F. M.; Musteata, M. L.; Pawliszyn, J. Clin. Chem. 2006, 52 (4), 708-715. (11) Chanard, J.; Lavaud, S.; Randoux, C.; Philippe, R. Nephrol., Dial., Transplant. 2003, 18, 252-257. (12) Lipatova, T. E.; Lipatov, Y. S. Macromol. Symp. 2000, 152, 139-150. (13) iNANO, University of Aarhus. http://www.inano.dk/sw2495.asp (accessed August, 2006). (14) Hermansson, J.; Grahn, A. J. Chromatogr., A 1994, 660, 119-129. (15) Hermansson, J.; Grahn, A.; Hermansson, I. J. Chromatogr., A 1998, 797, 251-263. (16) Musteata, F. M.; Walles, M.; Pawliszyn, J. Anal. Chim. Acta 2005, 537 (12), 231-237. (17) Mullett, W. M.; Pawliszyn, J. Anal. Chem. 2002, 74, 1081-1087.

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PEG.23-25 Biocompatible membranes have been prepared from polyurethane,26 chitosan,27 cellulose,28,29 and polyacrylonitrile (PAN).30-32 Polymers such as polysulfones, PAN, and polyamides are currently used to prepare biocompatible membranes used for separation of submicrometer particles in biomedical applications. PAN has been widely used as membrane material in the fields of dialysis and ultrafiltration. It has been found that its properties can be fine-tuned by using specific comonomers. For example, PAN can be tailored with a reactive group for enzyme immobilization. Furthermore, some comonomers lead to improved mechanical strength, solvent resistance, high permeation flux, and biocompatibility. Accordingly, PAN-based membranes have great potential for the treatment of wastewater, the production of ultrapure water, hemodialysis in artificial kidneys, and biocatalysis with separation.30 PAN is one of the most important polymers used in the biomedical area because of its exceptional qualities, such as good thermal, chemical, and mechanical stability as well as biocompatibility. Membranes made of PAN are widely used as dialyzers able to remove low to middle molecular weight proteins and for high-flux dialysis therapy.31 PAN is one of the best polymers in terms of biocompatibility32 but is not appropriate as extractive material for drugs. On the other hand, good extractive materials are generally not biocompatible; therefore, mixtures of PAN and extracting materials were studied. This article presents the development of new types of coatings which can be used for direct microextraction of drugs from biological fluids, particularly plasma. The coatings are prepared by covering flexible stainless steel wires with a mixture of PAN and different extractive particles (octadecyl silica, RP-amide, HSF5). In comparison to previously used PEG glue,10 PAN has much better elasticity and mechanical stability. Accordingly, PAN can be used both for covering existing commercial fibers with a biocompatible layer and for immobilizing extractive particles onto wires. Five different drugs (verapamil, diazepam, loperamide, nordiazepam, and warfarin, with log Pow values of 4.9, 3.0, 4.3, 3.2, and 3.5) were selected as target compounds for evaluating the performance of the newly developed coatings. These drugs were chosen because their chemistry and pharmacological action are well-known, they are widely used, and convenient to obtain. The (18) Anderson, J. L.; Armstrong, D. W.; Wei, G.-T. Anal. Chem. 2006, 78, 28932902. (19) Baker, G. A.; Baker, S. N.; Pandey, S.; Bright, F. V. Analyst 2005, 130, 800-808. (20) Peng, J. F.; Liu, J. F.; Jiang, G. B.; Tai, C.; Huang, M. J. Chromatogr., A 2005, 1072, 3-6. (21) Freemantle, M. Chem. Eng. News 2005, August 1, 33-38. (22) Makamba, H.; Hsieh, Y.-Y.; Sung, W.-C.; Chen, S.-H. Anal. Chem. 2005, 77 (13), 3971-3978. (23) Popat, K. C.; Sharma, S.; Desai, T. A. J. Phys. Chem. B 2004, 108, 51855188. (24) Sharma, S.; Desai, T. A. J. Nanosci. Nanotechnol. 2005, 5 (2), 235-243. (25) Sharma, S.; Johnson, R. W.; Desai, T. A. Langmuir 2004, 20, 348-356. (26) Mathieu, H. J. Surf. Interface Anal. 2001, 32, 3-9. (27) Peniche, C.; Arguelles-Monal, W.; Peniche, H.; Acosta, N. Macromol. Biosci. 2003, 3 (10), 511-520. (28) Elmquist, W. F.; Sawchuk, R. J. Pharm. Res. 1997, 14 (3), 267-288. (29) Shibata, T. Macromol. Symp. 2004, 208, 353-369. (30) Nie, F.-Q.; Xu, Z.-K.; Ming, Y.-Q.; Kou, R.-Q.; Liu, Z.-M.; Wang, S.-Y. Desalination 2004, 160, 43-50. (31) Lavaud, S.; Canivet, E.; Wuillai, A.; Maheut, H.; Randoux, C.; Bonnet, J.-M.; Renaux, J.-L.; Chanard, J. Nephrol., Dial., Transplant. 2003, 18, 2097-2104. (32) Yang, M. C.; Lin, W. C. J. Polym. Res. 2002, 9, 201-206.

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new coatings were tested for biocompatibility, reusability, extraction efficiency, and suitability for fast drug analysis and determination of drug plasma protein biding. EXPERIMENTAL SECTION: DEVELOPMENT OF BIOCOMPATIBLE SPME FIBERS Materials and Reagents. Diazepam, nordiazepam, and lorazepam were purchased from Cerilliant (Austin, TXsall benzodiazepines were 1 mg/mL in methanol, purity >98%). Polyacrylonitrile (monomer as impurity less than 4.85 ppm), verapamil (>98%), warfarin (>98%), loperamide (>99%), and phosphate buffer saline (pH ) 7.4) were bought from Sigma (ON, Canada). Ammonium acetate was obtained from BDH Inc. (Toronto, ON, Canada); N,N-dimethylformamide (DMF) was supplied by Caledon Laboratories LTD (Georgetown, ON, Canada). HPLC grade acetonitrile, acetone, methanol, acetic acid, and hydrochloric acid were purchased from Fisher Scientific (Nepean, ON, Canada). Deionized water was obtained using a Barnstead/ Thermodyne NANO-pure ultrapure water system (Dubuque, IA). Human plasma (with EDTA as anticoagulant) was bought from Bioreclamation Inc. (Hicksville, NY). Stainless steel wires (254 µm diameter) were purchased from Small Parts Inc. (Miami Lakes, FL); LiChrospher 60 XDS 25 µm (SO3/diol) particles were supplied by Merck KGaA (Darmstadt, Germany) as research samples and were used to prepare RAMbased coatings as previously described.5 PDMS fibers were purchased from Supelco (Belefonte, PA), and PPY fibers were prepared in-house as previously mentioned.6 The C18-silica (5 µm), RP-amide-silica (5 µm), and HS-F5-silica (5 µm) particles were provided by Supelco (Bellefonte, PA) as research samples. Octadecyl-silica (C18) 5 µm particles contain octadecyl as the bonded phase. For RP-amide C16 5 µm, the bonded phase is palmitamido-propyl. Both types of particles are spherical, with 180 Å pore size and 200 m2/g surface area. HS-F5 5 µm particles contain pentafluorophenyl-propyl as bonded phase, the shape of the particles is spherical, the pore size is 120 Å, and the surface area is 300 m2/g. SPME Fibers Based on PAN. PAN as a Solid Matrix. An amount of 0.47 g of particles (C18, RP-amide, or HS-F5, commonly used as HPLC stationary phases) was brought into suspension with 2 g of a solution of 10% PAN in DMF. SPME coatings with a length of 1.5 cm were prepared by applying a uniform layer of slurry of PAN and different particles on the surface of stainless steel wires, allowing it to dry under flowing nitrogen, and finally curing it for 1.5 min at 180 °C (in order to ensure better adherence of the PAN coating to the wire). The SPME coating was applied by dipping the wires into the slurry and removing them slowly. The wires were previously processed as follows: they were cut in 7.5 cm segments, washed with acetone, etched for 1 min in concentrated hydrochloric acid (by dipping 1.5 cm of the wire), immediately washed with water, thoroughly cleaned by sonication in water, and finally dried at room temperature. Prior to use, the coated fibers were conditioned in a water/methanol 50:50 wash for 30 min. PAN as a Membrane. Existing fibers with conventional extraction phases (CW/TPRscarbowax/templated resin, from Supelco, PA) were coated with PAN by dipping them for 2 min in a solution of 10% PAN in DMF. Consequently, the fibers were removed slowly from the solution, allowed to dry under flowing nitrogen,

Figure 1. SEM images of PAN/C18 coatings at 100× and 1000× magnification.

and finally cured by a short exposure (5 s) to a flow of nitrogen at 200 °C. The conditioning procedure was the same as for the fibers with PAN and extractive particles. Chemical Analysis. Standard Solutions and Sample Preparation. Stock solutions of the investigated drugs (diazepam, verapamil, warfarin, nordiazepam, loperamide, and lorazepam as internal standard) with a concentration of 1 mM were prepared weekly in a water/methanol 1:1 mixture and kept refrigerated at 4 °C (in 2 mL silanized vials). Human plasma (with EDTA as anticlotting agent, in 2 mL polypropylene vials) was stored at -20 °C until analysis. For analysis, plasma was thawed at room temperature and aliquots of 1.5 mL plasma were transferred into clean vials. Appropriate amounts of spike were added to obtain final concentrations in the range of 1 nM to 50 µM, followed by vortex mixing for 1 min. Samples and standards in PBS (phosphate buffer saline) were prepared similarly, to a final concentration in the range of 0.1 nM to 5 µM. The time required to reach equilibrium for plasma and PBS samples at 2400 rpm vortex stirring was determined at room temperature for all target compounds (diazepam, verapamil, and nordiazepam 5 × 10-7 M; warfarin 5 × 10-6 M; loperamide 5 ×

10-8 M) by measuring the amount extracted at different time points (shown in Figure 2). For extraction, the samples were placed on a digital vortex platform and the extracting phase of the SPME fiber was immersed in the sample for a precise period of time. The fiber was then briefly rinsed with water and desorbed for analysis. The lowest carryover and the sharpest chromatographic peaks for the investigated drugs were obtained for a desorption time of 15 min, vortex stirring at 2400 rpm, and with a desorption solution prepared from acetonitrile/water/acetic acid (50:49:1). Unless otherwise specified, the sample volume was 1.5 mL and the fiber was desorbed for 15 min in an insert with 60 µL desorption solution containing lorazepam as internal standard (50 ng/mL). All reproducibility, reusability, extraction efficiency, and calibration experiments were performed at equilibrium in similar conditions, following the general procedure for new SPME methods.4 Calibration curves were constructed by spiking PBS or human plasma with drug concentrations in the range of 0.5 nM to 50 µM, which generally covers the therapeutic concentrations. All extractions and desorptions were performed manually. LC/MS. LC/MS (liquid chromatography coupled with mass spectrometry) analyses were performed using an Agilent 1100 Analytical Chemistry, Vol. 79, No. 18, September 15, 2007

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Figure 2. Extraction time profiles for diazepam, verapamil, warfarin, nordiazepam, and loperamide with PAN/C18 (octadecyl-silica (A)), PAN/ RP-amide (palmitamido-propyl (B)), and PAN/HS-F5 (pentafluorophenyl-propyl (C)) coatings on 0.01 in. wires (n ) 3).

series liquid chromatograph (Agilent Technologies, Palo Alto, CA), equipped with a vacuum solvent degassing unit, a binary highpressure gradient pump, an autosampler, a column thermostat, and a variable wavelength UV-vis detector coupled on-line with an Agilent 1100 series MSD single quadrupole instrument with atmospheric pressure electrospray ionization (ESI). High-purity nitrogen used as nebulizing and drying gas was obtained from an in-house generator. Chromatographic separations were carried out on a Discovery C18 column (5 cm × 2.1 mm, 5 µm particles, from Supelco), guarded by an on-line filter (0.2 µm). Data were collected and analyzed using the CHEMSTATION software from Agilent Technologies. LC and ESI-MS conditions were as follows: column temperature 25 °C, mobile phase acetonitrile/20 mM ammonium acetate pH ) 7.0 with gradient programming (initial composition 10:90, 6906 Analytical Chemistry, Vol. 79, No. 18, September 15, 2007

ramped to 80:20 over 6 min, and maintained until the end of the run), flow rate 0.25 mL min-1, nebulizer gas N2 (35 psi), drying gas N2 (13 L min-1, 300 °C), capillary voltage 3500 V, fragmentor voltage 80 V, quadrupole temperature 100 °C, positive ionization mode. Total run time was 9 min. For optimization experiments, scan mode in the range of 1001500 amu was used; for quantification experiments, selected ion monitoring is used, with a scan time of 0.42 s/cycle and a dwell time of 65 ms. The following positive ions were monitored: diazepam, m/z 285.1; verapamil, m/z 455.3; warfarin, m/z 309.1; nordiazepam, m/z 271.1; loperamide, m/z 477.3; lorazepam, m/z 321.0. All other parameters of the mass-selective detector were automatically optimized using a calibration standard. Lorazepam was used as an internal standard for compensation of variations in the injection volume (20 µL).

Scanning Electron Microscopy (SEM). For SEM imaging, the fibers were cut into 7 mm long pieces, coated with gold (∼10 nm) and analyzed using a LEO 1530 emission scanning electron microscope at the Waterloo Watlab Facility.6 X-ray photoelectron Spectroscopy. XPS (X-ray photoelectron spectroscopy) analyses were performed by using a multitechnique ultra-high-vacuum imaging XPS microprobe system (Thermo VG Scientific ESCALab 250) equipped with a hemispherical analyzer with a mean radius of 150 mm and a monochromatic Al KR (1486.60 eV) X-ray source. The spot size for the XPS analysis used for the present work was approximately 0.5 mm by 1.0 mm. The samples were mounted on a stainless steel sample holder with double-sided carbon tapes. The sample was stored in vacuum (2 × 10-8 mbar) in the load-lock chamber of the imaging XPS microprobe system overnight to remove any remaining moisture before introduction into the analysis chamber maintained at 2 × 10-10 mbar. A combination of low-energy electrons and ions was used for charge compensation on the nonconducting coating material during the analysis conducted at room temperature. Averages of five high-resolution XPS scans were performed for each element of interest (C, N, O, S). Curve fitting was performed using CasaXPS VAMAS processing software, and the binding energies of individual elements were identified with reference to the NIST X-ray Photoelectron Spectroscopy database. All investigated fibers were exposed to undiluted human plasma at 37 °C for 1 h (this is considered a rigorous biocompatibility test).22 They were then briefly washed with phosphate buffer and deionized water and dried in nitrogen before analysis. Survey scans and high-resolution XPS scans were used to determine the atomic percentages of the surfaces before and after exposure to plasma. Sterilization. Sterilization was performed chemically or by steam. For chemical sterilization, the fibers were immersed in alcohol (methanol or ethanolssame results) for 30 min and then allowed to dry. Sterilization by steam was performed in an autoclave at 121 °C and 15 psi for 30 min. Calculation of Distribution Constant. For the direct extraction mode, the distribution constant can be calculated as3

Kfs )

nVs Vf(C0Vs - n)

(1)

where C0 is the initial concentration of a given analyte in the sample, Vs is the sample volume, Vf is the fiber coating volume, and n is the number of moles of analyte extracted by the coating. For new coatings, the volume of the coating can be determined from the coating length (b), coating thickness (dsobtained from SEM images), and radius of the support wire (r):

Vf ) πb[(r + d)2 - r2]

(2)

Equations 1 and 2 were used to calculate the distribution constant for five target compounds in the case of PAN-based coatings (Table 1). All determinations were performed in PBS at extraction equilibrium. RESULTS AND DISCUSSION Characterization of the New Coatings. In order to verify the topography of the extractive particles within PAN at the surface

Table 1. Distribution Constant Values for Coatings Based on PAN (Support Wire Diameter, 0.01 in.; Coating Thickness, 60 µm) (n ) 5) Kfs

PAN/C18

PAN/RP-amide

PAN/HS-F5

diazepam verapamil warfarin nordiazepam loperamide

1.14 × 3.25 × 103 1.19 × 102 5.60 × 102 1.92 × 104

4.57 × 1.43 × 103 4.02 × 101 5.66 × 102 2.34 × 103

9.96 × 102 2.13 × 103 9.37 × 101 9.70 × 102 1.14 × 104

103

102

of the fiber, optical microscope and SEM images were recorded. The SEM images of PAN/C18 coatings (Figure 1) demonstrate that the particles are completely covered with PAN and are homogeneously distributed within the coating. SEM was also used to estimate the average thickness of each coating, which was found to be 60-62 µm. No swelling of the coating was observed during analysis time (extraction up to 2 h and desorption for 15 min). To characterize the newly developed coatings, their extraction performance toward the target analytes was investigated. The amount of analyte extracted with the new coatings was compared to the amount extracted with other types of SPME fibers, under similar conditions. Coatings based on PPY and RAM were chosen as reference materials for extraction efficiency and biocompatibility tests, because their biocompatibility is already accepted.33,34 For the extraction efficiency test, performed at equilibrium as described in the Standard Solutions and Sample Preparation section, the coatings based on C18, RP-amide, and HS-F5 showed on average much higher extraction efficiency toward the test drugs: ∼90 times more than PPY, ∼50 times more than RAM or PDMS coatings, and ∼20 times more than commercially available CW/TPR. These results were indeed expected, as the particles that were used for the new coatings are widely known for their excellent properties as extraction phases in LC. Desorption Procedure. Successful coupling of SPME with HPLC is dependent on the efficiency of the desorption step. Currently, there are two choices for desorption: on-line (manual introduction of the fiber into a desorption chamber)3 or off-line (in a vial or 96-well plate).10 The carryover was found to be well below 3% (with 3 exceptions out of 20 determinations). For highly sensitive analyses, desorption is usually followed by solvent evaporation and reconstitution in a lower volume of solvent suitable for direct HPLC analysis. Nevertheless, desorption in 60 µL of solvent was found to be entirely suitable for the present study. If required, the carryover can be further decreased by using larger volumes of desorption solution or longer desorption time. Equilibration Time. Whereas the concentration of the sample analyzed by SPME has no impact on the extraction time profile and equilibration time, the agitation conditions, coating thickness (especially for liquid coatings), distribution constant, and diffusion coefficient of the analyte play very important roles in determining the equilibration time.35 While the real equilibration time is infinite, (33) Musteata, F. M.; Pawliszyn, J. J. Pharm. Biomed. Anal. 2005, 37 (5), 10151024. (34) Musteata, F. M.; Walles, M.; Pawliszyn, J. Anal. Chim. Acta 2005, 537 (12), 231-237. (35) Ai, J. Anal. Chem. 1997, 69, 1239-1236.

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Figure 3. Comparative extraction time profile for verapamil, obtained with the commercial CW/TPR (9) fiber and PAN/CW/TPR ([) fiber. Extractions were performed in phosphate buffer (n ) 3).

the experimental value can be conveniently expressed as the time required to extract at least 95% of the maximum amount. To minimize the errors caused by different sampling times, the extraction time should be equal to or longer than the equilibration time. The time required to reach equilibrium was found to be between 4 and 55 min in most cases (Figure 2). No significant difference was observed when the equilibration profile in PBS was compared to the equilibration profile in plasma. When the target drugs were analyzed in mixtures, an extraction time corresponding to the maximum equilibration time was used. If a shorter extraction time is chosen, great care must be taken to ensure all the samples are stirred similarly. When existing commercial coatings were covered with a layer of PAN, the equilibration time remained essentially the same. Furthermore, the mechanical stability of the fibers coated with PAN was significantly improved: while original fibers can be used for 20 extractions before they break down, those coated with PAN last for more than 50 extractions. In addition to improved biocompatibility and durability, the resulting fibers offer almost the same extraction capacity as the initial ones (Figure 3; the extraction capacity of the PAN-coated fiber is lower because some of the extraction sites on the surface of the initial CW/TPR fiber are occupied by PAN). Reproducibility of the Analytical Method. By using five different fibers of each kind (PAN/C18, PAN/RP-amide, PAN/HS-F5, and PAN/CW/TPR), the fiber-to-fiber coating reproducibility was determined to be below 10%. Similarly, same fiber reproducibility was determined by using the same fiber five times and was found to be below 10% as well. The experimental conditions were similar to those described in the Standard Solutions and Sample Preparation section for the equilibration time profile. It should be noted that in order to obtain good reproducibility for C18-based coatings, a conditioning step of at least 30 min in water/methanol 50:50 is required before the first use or after the coatings dry up. Conditioning with water or higher proportion of methanol was found to lead to worse reproducibility. On the other hand, all the other coatings described here require only a very brief conditioning step (less than 5 min) or even none at all. 6908 Analytical Chemistry, Vol. 79, No. 18, September 15, 2007

In order to investigate the long-term reusability of the newly developed coatings, five new SPME fibers were used for repeated extractions from drug solutions in PBS and human plasma and proved to be reusable for at least 10 extractions and desorptions (more experiments were not performed, as sample preparation devices for biological samples are generally of single use). Determination of the Distribution Constant. The target analyte’s distribution constant between fiber and sample (Kfs) defines the sensitivity of an analytical method based on SPME, gives more information about the experiment, and aids optimization. Kfs can be used to calculate the sample volume and coating thickness required to reach the desired sensitivity. Generally, PAN/C18 fibers have the highest distribution constants, but they need careful conditioning before extraction from samples. Even if the distribution constants for PAN/RPamide and PAN/HS-F5 coatings are a little lower, they could be more convenient as they do not require extraction phase activation prior to extraction. As expected, the less polar extraction phases showed higher distribution constants for less polar analytes (Table 1). Sterilization. Sterilization is necessary only if the microextraction devices are to be used for in vivo experiments. In this case, the coatings should be tested for endurance during sterilization. Current sterilization methods include heat, steam, chemical (ethylene oxide, alcohols, aldehydes), and radiation. The most preferable methods would be sterilization with steam or radiation. Radiation sterilization, although highly effective, poses a risk to human health and is not readily accessible. As a result, steam sterilization is most widely used. The new coatings were tested for extraction efficiency before and after chemical and steam sterilization. No change in extraction efficiency was observed upon sterilization with alcohols, as this step is similar to the conditioning step (before extraction). In the case of sterilization in an autoclave, the proposed coatings showed no sign of deterioration (as determined from optical microscope images); this was expected, since PAN coatings are known to withstand GC injector temperatures (>250 °C). Although no signs of breakdown were observed, the extraction capacity decreased

Table 2. Atomic Composition Obtained by XPS for Selected Proteins and Coatings (before and after Exposure to Human Plasma)a

protein/coating

C% (RSD < 5%)

N% (RSD < 5%)

O% (RSD < 10%)

S% (RSD < 15%)

human serum albumin fibrinogen PAN coating (bpb) PAN coating (apc) PAN/C18 (bpb) PAN/C18 (apc) PAN/RP-amide (bpb) PAN/RP-amide (apc) PAN/HS-F5 (bpb) PAN/HS-F5 (apc) PAN/RAM (bpb) PAN/RAM (apc) PPY (bpb) PPY (apc)

63.3 62.8 78.2 73.5 77.0 73.6 78.3 68.6 79.3 70.9 78.9 72.6 61.0 69.7

16.9 18.0 17.6 15.1 17.2 13.9 17.0 15.5 20.1 15.2 20.4 16.3 3.8 12.4

19.0 18.8 4.0 11.3 5.7 12.5 4.5 15.8 0.4 13.7 0.5 10.4 35.2 17.6

0.9 0.5 0.0 0.0 0.0 0.0d 0.0 0.0 0.0 0.0 0.0 0.5 0.0 0.3

a Hydrogen is not reported, as it is not detectable by XPS. b bp ) before exposure to human plasma. c ap ) after exposure to human plasma. d The experimental value was 0.04, below the limit of quantitation of 0.1%.

by approximately 15% after sterilization, probably because of the combined effect of heat and water vapors on the fused-silica particles. Accordingly, when maximum extraction capacity is needed, these coatings should be sterilized with alcohol. Biocompatibility Test. Many methods have been applied for the study of biocompatibility, ranging from the simple visual inspection to the most sensitive atomic force microscopes. Nevertheless, only a few methods are widely used and recognized: XPS,23 atomic force microscopy,25,36 surface plasmon resonance,37 and competitive ELISA (enzyme-linked immunosorbent assay). XPS or electron spectroscopy for chemical analysis (ESCA) is one of the most common types of spectroscopic methods for analysis of surfaces. The sampling depth for this method is approximately 1-30 nm (up to 100 nm mean-free-pass), which encompasses a surface region highly relevant for biointeractions.38 The biocompatibility of the proposed coatings was tested by XPS. A material is considered biocompatible if the amount of nitrogen and sulfur on the surface does not increase significantly after contact with a biological system.22 After exposure of PANbased coatings to plasma, the amount of nitrogen and carbon on the surface generally decreases, accompanied by an increase in the amount of oxygen (Table 2; “PAN coating” represents wires and CW/TPR fibers coated with a layer of PAN). These observations suggest that most of the molecules adsorbed from human plasma contain a high percent of oxygen (usually because of nonspecific adsorption), whereas their nitrogen content is lower than that of plasma proteins. Even more conclusive from a biocompatibility point of view is the amount of sulfur on the surface, since it is naturally present in proteins and absent from the investigated SPME coatings. When compared to RAM and (36) Sharma, S.; Johnson, R. W.; Desai, T. A. Biosens. Bioelectron. 2004, 20, 227-239. (37) Uchida, K.; Otsuka, H.; Kaneko, M.; Kataoka, K.; Nagasaki, Y. Anal. Chem. 2005, 77 (4); 1075-1080. (38) Karlgard, C. C. S.; Sarkar, D. K.; Jones, L. W.; Moresoli, C.; Leung, K. T. Appl. Surf. Sci. 2004, 230, 106-114.

PPY, materials regarded as highly biocompatible,10,34 the new coatings based on PAN showed a much lower increase in sulfur. The biocompatibility test based on XPS suggests that the most biocompatible PAN-based coatings are PAN/RP-amide and PAN/ HS-F5, followed closely by PAN/C18. Furthermore, the newly developed PAN-based coatings were inspected under the microscope after 5 min exposure to human plasma and whole mouse blood (without anticlotting agents), and no clot adhesion to the coating was observed. Applications of Biocompatible SPME. Fast Drug Analysis in Vivo and in Vitro. The advantages of the proposed SPME coatings were investigated by studying the extraction and separation of drugs (described in the LC/MS section) from human plasma. As shown in Figure 4, a very good linear relationship was obtained for a seven point calibration (n ) 3). Figure 4 also indicates that drug binding to plasma proteins changes the amount of drug available for extraction and results in different calibration slopes for plasma and PBS. The linear range covered more than 3 orders of magnitude for most drugs, with the exception of warfarin, when the linear range spanned over 2 orders of magnitude. The full details are included in Table 3. In recent years there has been considerable interest in developing techniques to monitor levels of biologically active compounds in living systems in natural environments. In vivo sampling can eliminate errors and reduce the time associated with sample transport and storage and can, therefore, result in collecting more accurate and precise analytical data.39 An ideal in vivo sampling technique should be portable, solvent-free, and offer integration of the sampling, sample preparation and analysis step. Reliable and accurate analytical methods are indispensable for in vivo research. In vivo analysis is a special application area where SPME is gaining ground because of its unique characteristics: on-site sampling, easy extraction, and analysis of the whole extracted amount. Early in vivo investigations with SPME focused on fragrances emitted by insects, fungi, and bacteria. These investigations were extended to biogenic volatile organic compounds emitted by animals and plants. In a more recent application, we showed that the SPME technology can be used for in vivo analysis of intravenous drug concentrations in a living animal. A novel SPME probe was developed, and its effectiveness was demonstrated by acquiring the pharmacokinetic profile of diazepam, nordiazepam, and oxazepam. The method was validated by comparison to conventional sampling methods.10 For this initial study, most of the time points of a pharmacokinetic profile were acquired using PPY coatings. However, the PPY coatings have a rather small linear range and are very fragile. The application of the proposed PAN/C18 and PAN/RP-amide fibers would result in significant improvements: the linear range is considerably larger (for both low and high concentrations), fiber-to-fiber RSD is below 10%, and the coating is much more robust. Such fibers are currently used for in vivo studies in rodents (manuscript in preparation). Determination of Drug Plasma Protein Binding. The determination of the amount of drug binding to plasma proteins is an essential step in both drug discovery and in clinical phases of drug development.40-45 Due to the important clinical implications of (39) Pawliszyn, J. Aust. J. Chem. 2003, 56 (2-3), 155-158.

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Figure 4. Calibration curve for loperamide in PBS and human plasma (with SPME, RP-amide coating). The signal was defined as the ratio between the analyte and internal standard; r2 ) 0.9997 for PBS; r2 ) 0.9986 for plasma. Table 3. Linear Ranges for SPME-Based Analytical Method (r2 > 0.99 for All) PAN/C18

PAN/RP-amide

linear range (mol/L)

PBS

plasma

PBS

plasma

diazepam verapamil warfarin nordiazepam loperamide

1 × 10-9 f 2 × 10-6 1 × 10-9 f 1 × 10-6 2 × 10-8 f 5 × 10-6 1 × 10-8 f 5 × 10-6 1 × 10-9 f 2 × 10-7

1 × 10-8 f 1 × 10-5 5 × 10-9 f 5 × 10-6 2 × 10-7 f 5 × 10-5 1 × 10-7 f 2 × 10-5 5 × 10-9 f 2 × 10-6

3 × 10-9 f 1 × 10-6 2 × 10-9 f 4 × 10-7 2 × 10-8 f 4 × 10-6 7 × 10-9 f 2 × 10-6 2 × 10-9 f 2 × 10-7

5 × 10-8 f 1 × 10-5 2 × 10-8 f 4 × 10-6 1 × 10-6 f 4 × 10-5 2 × 10-7 f 2 × 10-5 2 × 10-8 f 2 × 10-6

Table 4. Experimental and Literature Drug Plasma Protein Binding Values plasma protein binding %

PAN/C18 0.01 in., 60 µm

PAN/RP-amide 0.01 in., 60 µm

literature values (range)a

98 96 99 98 98

99 96 99 98 98

96-98 88-98 98-100 97-98 97-98

diazepam verapamil warfarin nordiazepam nordiazepam a

Refs 48 and 49.

plasma protein binding data and its role in characterizing a drug’s behavior and proper dosing, there is an increasing need to make this044 measurement as early as possible in the discovery process in order to understand drug disposition and to optimize individual drug therapy.46,47 The determination of plasma protein binding by SPME is based on determining the free concentration of drug in the presence of plasma proteins and has been comprehensively described before.48 Briefly, the percentage of drug binding to (40) Banker, M. J.; Clark, T. H.; Williams, J. A. J. Pharm. Sci. 2003, 92 (5), 967-974. (41) Kariv, I.; Cao, H.; Oldenburg, K. R. J. Pharm. Sci. 2001, 90 (5), 580587. (42) Olson, R. E.; Christ, D. D. Annu. Rep. Med. Chem. 1996, 31, 327-336. (43) Sarre, S.; Van Belle, K.; Smolder, S. I.; Krieken, G.; Michotte, Y. J. Pharm. Biomed. Anal. 1992, 10 (10-12), 735-9. (44) Tanaka, H.; Mizojiri, K. J. Pharmacol. Exp. Ther. 1999, 288 (3), 912918. (45) Fung, E. N.; Chen, Y.-H.; Lau, Y. Y. J. Chromatogr., B 2003, 795 (2), 187194. (46) Cheng, Y.; Ho, E.; Subramanyam, B.; Tseng, J. L. J. Chromatogr., B 2004, 809 (1), 67-73. (47) Schuhmacher, J.; Buhner, K.; Witt-Laido, A. J. Pharm. Sci. 2000, 89 (8), 1008-1021.

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plasma proteins (PPB) is calculated from the total and free concentration of drug:

PPB% )

Ctotal plasma - Cfree plasma × 100 ) Ctotal plasma

(

1-

)

Cfree plasma × 100 (3) Ctotal plasma

where Ctotal plasma is the total concentration of drug in plasma and Cfree plasma is the free concentration of drug in plasma. Considering that the total drug concentration is directly proportional to the slope of the drug calibration curve in PBS and the free concentration is directly proportional to the slope of plasma calibration,48 eq 3 becomes:

(

PPB% ) 100 × 1 -

slope calibration plasma slope calibration PBS

)

(4)

Equation 4 was applied for the determination of drug plasma protein binding for the five test drugs, and the results are presented in Table 4. Only the most reproducible coatings were used, and the results correlate very well with previously published values. CONCLUSIONS The current research presents the development of new biocompatible SPME coatings that are not fouled by protein adsorption during extraction from biological samples. Preparation of (48) Musteata, F. M.; Pawliszyn, J.; Qian, M. G.; Wu, J. T.; Miwa, G. T. J. Pharm. Sci. 2006, 95 (8), 1712-1722. (49) Hardman, J. G., Limbird, L. E., Gilman, A. G., Goodman, L. S., Eds. Goodman and Gilman's the Pharmacological Basis of Therapeutics, 10th ed.; McGrawHill: New York, 2001.

biocompatible and hemocompatible SPME coatings represents an important step toward developing powerful biomedical, pharmaceutical, and forensic applications, as the advantages of SPME would be directly useful for analysis of biological samples. One of the main deterrents of in vivo application of SPME has been the lack of suitable extractive phases. It is expected that the number of such applications will significantly increase with the introduction of new biocompatible coatings. It should be noted that this coating preparation approach can be used with any extractive particles currently used in solid-phase extraction or HPLC. Furthermore, other biocompatible polymers could be used as glue or support. Future developments in the area of biocompatible SPME could include analytical methods with lower limits of detection, applica-

tions for other drugs with a wider range of polarities, and automation of fiber preparation, extraction, and desorption. ACKNOWLEDGMENT The authors gratefully acknowledge the financial support received from Canada Research Chair, Natural Sciences and Engineering Research Council of Canada, NoAb BioDiscoveries, and Supelco.

Received for review February 12, 2007. Accepted May 10, 2007. AC070296S

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