Online Concentration and Affinity Separation of Biomolecules Using

PVA-coated capillary, 37 cm × 50 μm i.d. (29 cm to the detector); detection wavelength, ... MFMPs f, 2.1, −5.7, 0.64 ... Although the injection ti...
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Anal. Chem. 2007, 79, 3041-3047

Online Concentration and Affinity Separation of Biomolecules Using Multifunctional Particles in Capillary Electrophoresis under Magnetic Field Yukihiro Okamoto, Fumihiko Kitagawa,* and Koji Otsuka

Department of Material Chemistry, Graduate School of Engineering, Kyoto University, Katsura, Nishikyo-ku, Kyoto 615-8510, Japan

To overcome several problems in affinity capillary electrophoresis (ACE), i.e., low detectability, need for sample derivatization, and difficulty in the fixation of affinity ligands (ALs), multifunctional magnetic particles (MFMPs) were prepared by immobilizing both fluorescent molecules and ALs for low-density lipoproteins onto the surface of magnetic polymer microspheres with a polyelectrolyte multilayer coating technique and applied to the ACE analysis. The prepared MFMPs showed a remarkable change in the electrophoretic mobility (µep) by the addition of low-density lipoproteins (LDL), whereas for highdensity lipoproteins (HDL), µep of the MFMPs kept constant, so that it was confirmed that the MFMPs possess an affinity with LDL. On the other hand, the MFMPs can be trapped by the magnetic field even under a higher electric field for electrophoresis. By a successive on-off control of the magnetic field, online preconcentration of the LDL bound MFMPs and the selective separation of LDL from HDL were successfully achieved. In the ACE analysis of LDL employing UV detection, an 82-fold increase in the sensitivity was obtained by the oncapillary sample preconcentration using the MFMPs. When laser induced-fluorescence detection was employed, furthermore, the limit of detection for LDL was improved to the order of subpicomolar. In the separation of biological samples, there are many analytical requirements, e.g., high resolution, high sensitivity, easy operation, short analysis time, small amount of analytes, and so on. Among several separation techniques, slab gel electrophoresis has been routinely employed for the separation and analysis of biomolecules. However, some experimental procedures such as the preparation of the gel and staining the analytes are laborintensive and time-consuming. On the other hand, capillary electrophoresis (CE) has been regarded as one of the useful analytical tools for biological samples. Especially, affinity capillary electrophoresis (ACE), which utilizes biospecific interactions, is the most powerful tool in CE and can attain selective separations of biomolecules.1-9 * To whom correspondence should be addressed. E-mail: [email protected]. (1) Heegaard, N. H. H.; Nilsson, S.; Guzman, N. A. J. Chromatogr., B 1998, 715, 29-54. (2) Kajiwara, H. Anal. Chim. Acta 1999, 383, 61-66. (3) Guijt-van Duijn, R. M.; Frank, J.; Dedem, G. W. K.; Baltussen, E. Electrophoresis 2000, 21, 3905-3918. (4) Guzmann, N. A.; Stubbs, R. J. Electrophoresis 2001, 22, 3602-3628. 10.1021/ac061693q CCC: $37.00 Published on Web 03/10/2007

© 2007 American Chemical Society

In ACE, affinity ligands (ALs) are employed mainly by four different methods: (i) addition into the background solution (BGS), (ii) packed ACE (p-ACE), and (iii) open tubular ACE (oACE).10 In o-ACE, preparation of AL immobilized capillaries is easier than p-ACE, which generally has a problem concerning the preparation of frits, but a poor resolution is sometimes problematic according to a lower phase ratio in the AL-coated capillaries. By employing the covalent immobilization technique, a stable ALs layer can be obtained. However, the immobilization via covalent bonding often requires labor-intensive and time-consuming processes, so that the separation conditions are difficult to be optimized. On the contrary, the physical adsorption can easily provide the AL immobilized capillaries but needs frequent reimmobilizations of ALs due to their desorption from the inner surface. Thus, the introduction of a versatile method for a rigid immobilization of ALs with a higher phase ratio in ACE has generated a great deal of interest. In this study, to prepare stable AL layers, a polyelectrolyte multilayer (PEM) coating technique was applied to the surface of magnetic polymer particles (MPs). Since Decher and Hong reported that alternating depositions of polyelectrolytes form a stable multilayer in 1991,11,12 PEM has been extensively studied and it has been elucidated that PEM has several superior characteristics.13-16 In the PEM coating, the thickness and properties of the multilayer can be adjusted by the preparation conditions, e.g., ionic strength, pH, polyelectrolyte concentration, and number of depositions.17-22 In addition, the (5) Schou, C.; Heegaard, N. H. H. Electrophoresis 2006, 27, 44-59. (6) Ensing, K.; Paulus, A. J. Pharm. Biomed. Anal. 1996, 14, 305-315. (7) Wang, Q.; Luo, G.; Ou, J.; Yeung, W. S. B. J. Chromatogr., A 1999, 848, 139-148. (8) Tomlinson, A. J.; Benson, L. M.; Guzman, N. A.; Naylor, S. J. Chromatogr., A 1996, 744, 3-15. (9) Guzman, N. A.; Park, S. A.; Schaufelberger, D.; Hernandez, L.; Paez, X.; Rada, P.; Tomlinson, A. J.; Naylor, S. J. Chromaotogr. B 1997, 697, 37-66. (10) Kitagawa, F.; Inoue, K.; Hasegawa, T.; Kamiya, M.; Okamoto, Y.; Kawase, M.; Otsuka, K. J. Chromatogr., A 2006, 1130, 219-226. (11) Decher, G.; Hong, J. D. Makromol. Chem. Macromol. Symp. 1991, 46, 321. (12) Decher, G. Science 1997, 277, 1232-1237. (13) Bertrand, P.; Jonas, A.; Laschewsky, A.; Legras, R. Macromol. Rapid Commun. 2000, 21, 319-348. (14) Hammond, P. T. Curr. Opin. Colloid Interface Sci. 2000, 4, 430-442. (15) Decher, G., Schlenoff, J. B., Eds. Multilayer thin films: sequential assembly of nanocomposite materials; Wiley-VCH: Weinheim, 2003. (16) Scho ¨nhoff, M. Curr. Opin. Colloid Interface Sci. 2003, 8, 86-95. (17) Dubas, S. T.; Schlenoff, J. B. Macromolecules 1999, 32, 8153-8160. (18) Dubas, S. T.; Schlenoff, J. B. Langmuir 2001, 17, 7725-7727. (19) Tiourina, O. P.; Antipov, A. A.; Sukhorukov, G. B.; Larionova, N. I.; Lvov, Y.; Mo ¨hwald, H. Macromol. Biosci. 2001, 1, 209-214.

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activity of labile biomolecules in water is sometimes enhanced in PEM.23,24 Compared to a Langmuir-Blodgett film, the PEM coating has a wider applicability of immobilization, i.e., for not only a flat but more complicated form substrate,25 which enables us to prepare a thin film onto the surface of microparticles.26-29 In the immobilization of ALs, it is most important to keep the specific interaction activity. Additionally, easy and rapid procedures are also required to obtain a strong immobilization. From these points, the PEM coating is regarded as one of the best methods for immobilizing ALs. MPs, which respond to the magnetic field, have been utilized for cell separation,30 DNA collection,31 metal collection,32 and so on.33-35 The selective separation and collection of target analytes can be magnetically attained by using biomolecule immobilized MPs. As an application of MPs in CE, Rashkovetsky et al. reported that femtomoles of antigen could be quantified by isotachophoretic focusing.36 In the case of AL immobilized MPs, the efficiency for the specific interaction between target analytes and ALs is expected to be enhanced since the nano- or micrometer-sized particles have large surface areas.37 These properties should facilitate affinity analysis of the trace amount of analytes. In this study, we prepared multifunctional magnetic particles (MFMPs) by the PEM coating of cationic fluorescent and anionic AL polymers as the inner and outer layers, respectively, onto the surface of MPs. To achieve selective and sensitive ACE analysis, online preconcentration and selective separation of target analytes by using the MFMPs were conducted as shown in Figure 1. The prepared MFMPs are expected to possess three functions, i.e., magnetic response, fluorescence labeling, and ALs for target analytes. By adding the MFMPs into a sample solution, the target analytes bind selectively onto the MFMP surface via specific interaction with ALs. Since the MFMP has a fluorescent layer, simultaneous labeling of the target analytes can be achieved, which can omit a labor-intensive labeling procedure prior to the analysis. When the prepared sample suspensions are electroki(20) Kapnissi, C. P.; Valle, B. C.; Warner, I. M. Anal. Chem. 2003, 75, 60976104. (21) Kamande, M. W.; Zhu, X.; Kapnissi-Christodoulou, C.; Warner, I. M. Anal. Chem. 2004, 76, 6681-6692. (22) Kitagawa, F.; Kamiya, M.; Okamoto, Y.; Taji, H.; Onoue, S.; Tsuda, Y.; Otsuka, K. Anal. Bioanal. Chem. 2006, 386, 594-601. (23) Onda, M.; Ariga, K.; Kunitake, T. J. Biosci. Bioeng. 1999, 87, 69-75. (24) Caruso, F.; Schu ¨ ler, C. Langmuir 2000, 16, 9595-9603. (25) Ohyama, T.; Nishide, T.; Iwata, H.; Sato, H.; Toda, M.; Toma, N.; Taki, W. J. Neurosurg. 2005, 102, 109-115. (26) Sukhorukov, G. B.; Donath, E.; Lichtenfeld, H.; Knippel, E.; Knippel, M.; Budde, A.; Mo ¨hwald, H. Colloids Surf. A: Physicochem. Eng. Aspects 1998, 137, 253-266. (27) Caruso, F.; Susha, A. S.; Giersig, M.; Mo¨hwald, H. Adv. Mater. 1999, 11, 950-953. (28) Gittins, D. I.; Caruso, F. J. Phys. Chem. B 2001, 105, 6846-6852. (29) Dun, H.; Zhang, W.; Wei, Y.; Xiuqing, S.; Li, Y.; Chen, L. Anal. Chem. 2004, 76, 5016-5023. (30) Furdui, V. I.; Harrison, D. J. Lab Chip 2004, 4, 614-618. (31) Fan, Z. H.; Mangru, S.; Granzow, R.; Heaney, P.; Ho, W.; Dong, Q. P.; Kumar, R. Anal. Chem. 1999, 71, 4851-4859. (32) Ngomsik, A. F.; Bee, A.; Draye, M.; Cote, G.; Cabuil, V. C. R. Chim. 2005, 8, 963-970. (33) Pankhurst, Q. A.; Connolly, J.; Jones, S. K.; Dobson, J. J. Phys. D: Appl. Phys. 2003, 36, R167-R181. (34) Gijs, M. A. M. Microfluid Nanofluid 2004, 1, 22-40. (35) Pamme, N. Lab Chip 2006, 6, 24-38. (36) Rashkovetsky, L. G.; Lyubarskaya, Y. V.; Foret, F.; Hughes, D. E.; Karger, B. L. J. Chromatogr., A 1997, 781, 197-204. (37) Watanabe, J.; Ishihara, K. Biomacromolecules 2006, 7, 171-175.

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Figure 1. Concept of online preconcentration and selective separation of lipoproteins using MFMPs in ACE.

netically injected as a long plug into the capillary by applying the magnetic field, the online trapping and preconcentration of the target analyte bound MFMPs is expected and further application of the electric field provides a foregoing migration of other unbound components, which would result in the purification of the trapped target analytes. Finally, the trapped and concentrated MFMPs are forced to migrate by removing the magnetic field, and then they can be detected with laser-induced fluorescence (LIF). By employing shorter length of the magnet for trapping, efficient concentration of large amount of sample can provide a shape peak. Therefore, a further sensitive detection of the target analytes is expected by our approach. As a model study, low-density lipoproteins (LDL) and sodium dextran sulfate (DxS) were selected as the target analytes and ALs, respectively. Separation of LDL and high-density lipoproteins (HDL) is very important since the ratio of cholesterol in LDL to that in HDL is the most common indicator of risk for coronary artery disease. As conventional methods for the separation of HDL and LDL, slab gel electrophoresis, size exclusion chromatography, and ultracentrifugation have been employed. To reduce the sample amount and operation time with high resolution, capillary and microchip electrophoretic separations have been also applied.38,39 In these electrophoretic separations, however, there are some problems to be solved such as complicated procedures and insufficient separation efficiency. By employing DxS immobilized (38) Bo ¨ttcher, A.; Schlosser, J.; Kronenberg, F.; Dieplinger, H.; Knipping, G.; Lackner, K. J.; Schmitz, G. J. Lipid Res. 2000, 41, 905-915. (39) Weiller, B. H.; Ceriotti, L.; Shibata, T.; Rein, D.; Roberts, M. A.; Lichtenberg, J.; German, J. B.; Rooij, N. F.; Verpoorte, E. Anal. Chem. 2002, 74, 17021711.

MFMPs as shown in Figure 1, selective and sensitive analysis of LDL is expected. In this study, the electrophoretic properties and affinity with LDL of the prepared MFMPs were evaluated by a conventional CE-UV analysis. The selective separation and the online preconcentration of the LDL bound MFMPs under the magnetic field were also investigated with both UV and LIF detection. EXPERIMENTAL SECTION Materials and Chemicals. MPs (IMMUTEC MAG; mean diameter, 1.1 µm; surface functional group, carboxylate), which consist of acrylate polymer and superparamagnetic material, were kindly supplied from JSR (Tokyo, Japan). Poly(vinyl alcohol) (PVA; 98.5% hydrolyzed) was a gift from Japan VAM & POVAL (Osaka, Japan). Poly(ethyleneimine) (PEI; Mw ) 750 000, Mn ) 60 000, Mw/Mn ) 12.5, branched chain) and DxS (Mw ) 500 000, sulfur content of 17%) were purchased from Sigma-Aldrich (St. Louis, MO), HDL and LDL (did not contain very low-density lipoproteins) were from EMD Biosciences (San Diego, CA), rhodamine B isothiocyanate (RITC) was from Nacalai Tesque (Kyoto, Japan), and N,N-dimethylformamide (DMF) was from Wako Pure Chemical (Osaka, Japan). All the solutions were prepared with deionized water purified by a Direct-Q System (Nihon Millipore, Tokyo, Japan). All BGSs were filtered with a 0.45-µm membrane filter prior to use. Apparatus. Conventional CE analyses were carried out with a P/ACE 5000 (Beckman Coulter, Fullerton, CA) for characterization of the prepared particles. Fused-silica capillaries (Polymicro Technologies, Phoenix, AZ) with a dimension of 50-µm i.d. and 360-µm o.d. were coated with PVA to suppress the electroosmotic flow (EOF) and the sample adsorption.40 Injections were performed hydrodynamically by applying 15 psi at the capillary inlet for 5 s. The applied voltage was 15 kV and the temperature was set at 25 °C. Sample particles were detected at 200 nm. In the concentration of the MFMPs by applying a magnetic field, the CE-UV analysis was performed with a homemade apparatus, which consists of a UV detector (CE-2070 plus, Jasco, Tokyo, Japan) and high-voltage power supply (HCZE-30PN0.25, Matsusada Precision, Shiga, Japan). Output signals were acquired by using a PC with ChromatoDAQ (Ulvac, Kanagawa, Japan). The magnetic field was applied by placing a 1-mm neodium magnet (Magtec, Wakayama, Japan; 10 mm in length, 5 mm in width, 1 mm in depth; surface magnetic flux density, 220 mT; magnetic flux density at focusing point, 146 mT) onto the capillary at a distance of 16 cm from the inlet. In the CE-LIF measurement, a 532-nm laser beam (10-mW diode-pumped solid-state laser, 58GCS411, Melles Griot, Tokyo, Japan) for excitation was employed and irradiated to the detection window of the capillary through an objective lens (×20, NA ) 0.40, LCPlanFL-20X, Olympus). Fluorescence from the MFMPs collected by the same objective lens was passed through a dichroic filter and led to a multichannel photodetector (PMA-11, Hamamatsu Photonics, Shizuoka, Japan). To reduce the background signal caused by the reflection and refraction of the laser beam on the capillary surface, rectangular capillaries (Polymicro Technologies) were employed.41,42 (40) Okamoto, Y.; Kitagawa, F.; Otsuka, K. Electrophoresis 2006, 27, 10311040. (41) Tsuda, T.; Sweedler, J. V.; Zare, R. N. Anal. Chem. 1990, 62, 2149-2152.

Preparation of MFMPs. Prior to the preparation of the MFMPs, RITC-labeled PEI (RITC-PEI) was prepared by reacting PEI (0.42 µmol) and RITC (1.3 µmol) in methanol (20 mL) for 2 h. Crude product mixture was dialyzed by using a cellulose membrane for 7 days. The removal of free RITC was confirmed by thin-layer chromatography. MFMPs were prepared by the following procedures. An MP suspension (9 × 109 particles/mL, 1 µL) was washed with water twice and redispersed in water (1 mL). An RITC-PEI solution (∼4 µM, 40 µL) and the washed MP suspension (1 mL) were mixed and incubated for 15 min in the dark at room temperature. After the incubation, the MPs were precipitated with a magnet, washed, and redispersed in water (1 mL). A DxS solution (0.2 µM, 80 µL) and RITC-PEI immobilized MPs suspensions (1 mL) were mixed and incubated for 15 min. After this procedure, the modified particles were collected by magnet force, washed, and redispersed with 10 mM phosphate buffer (pH 7.4). By employing the above procedures, the PEM coating with its thickness of a few nanometers can be obtained on the MPs surface.28 Counting of the number of particles in the obtained suspension was performed with a hemocytometer, and the concentration of a stock MFMPs suspension was evaluated to be ∼1 × 107 particles/mL. Procedures. Online concentration of target analytes by capturing the MFMPs with the magnetic field was conducted by the following procedures (Figure 1). First, a sample solution containing the target analytes (LDL) and impurities were added into the MFMP suspension. In this study, HDL was treated as impurities. LDL was bound onto the MFMP surface via specific interaction with DxS and indirectly labeled with RITC-PEI. In the next step, the prepared sample was electrokinetically injected as a long plug by setting the magnet at a desired position of the capillary. As a typical condition, a voltage of 10 kV was applied for 5-20 min to introduce a large amount of the sample. The LDL bound MFMPs (LDL-MFMPs) reaching at the magnet position were trapped and concentrated to a narrow zone, while other components not immobilized on the MFMPs would migrate to the detection point. In the third step, after exchanging the inlet vial from the sample to the BGS, the electric field was applied with the magnetic field for further concentration and separation of the LDL-MFMPs. Finally, the trapped and concentrated MFMPs binding the target analytes were released by removal of the magnetic field, so that they are selectively detected with UV or LIF detection. For evaluation of the efficiency of online preconcentration, the sensitivity enhancement factor (SEFheight) was calculated by the following equation.43,44

SEFheight )

peak height obtained with magnetic trapping peak height obtained with a conventional injection (10 kV, 10 s) (1)

RESULTS AND DISCUSSION Characterization of MFMPs. Since the PVA coating can suppress the adsorption of proteins,45,46 the PVA-coated capillaries (42) Li, L.; McGown, L. B. Electrophoresis 2000, 21, 1300-1304. (43) Quirino, J. P.; Terabe, S. Science 1998, 282, 465-468. (44) Quirino, J. P.; Terabe, S. Anal. Chem. 2000, 72, 1023-1030. (45) Ikada, Y.; Iwata, H.; Horii, F.; Matsunaga, T.; Taniguchi, M.; Suzuki, M.; Taki, W.; Yamagata, S.; Yonekawa, Y.; Handa. H. J. Biomed. Mater. Res. 1981, 15, 697-718.

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Table 1. Electrophoretic Data for Modified MPs sample bare MPs d RITC-PEI immobilized MPs e MFMPs f

a

detection time (min)

µepb (10-4 cm2/V‚s)

% RSD of detection time c

2.5 8.8

-4.8 1.3

0.60 0.72

2.1

-5.7

0.64

a Conditions are as in the Experimental Section. b Minus sign shows an anodic migration. c n ) 5. d Surface functional groups, COOH. e Frequent spikelike peaks were observed. Detection time was defined as that obtained the highest peak. f Both RITC-PEI and DxS immobilized MPs.

Figure 3. Dependence of µep for MFMPs on the concentration of LDL and HDL. Injection, 10 kV for 10 s; MFMPs concentration, 1 × 107 particles/mL.

Figure 2. Electropherograms of (a) bare, (b) RITC-PEI immobilized, and (c) both RITC-PEI and DxS immobilized MPs. PVA-coated capillary, 37 cm × 50 µm i.d. (29 cm to the detector); detection wavelength, 200 nm; applied voltage, (a, c) -15 and (b) +15 kV; sample injection, 15 psi, 5 s; BGS, 10 mM phosphate buffer (pH 7.4).

are expected to provide the high-performance separation of biological samples using microparticles. Actually, LDL, HDL, and polymer particles were successfully analyzed in the PVA-coated capillaries with good reproducibility. It should be noted that the EOF on the PVA-coated capillary was completely suppressed since the EOF marker DMF was not detected. By employing the PVAcoated capillary, we performed the CE analysis of the MFMPs. Figure 2 shows the electropherograms of bare, RITC-PEI, and both RITC-PEI and DxS modified MPs (MFMPs). Bare MPs were detected at the anodic end at pH 7.4 due to the deprotonation of carboxylate groups on the surface of the MPs. On the other hand, the opposite direction of the electrophoretic migration was observed for the MPs immersed in a RITC-PEI solution, which indicated that positively charged fluorescent RITC-PEI was successfully immobilized onto the MP surface. The spikelike peaks obtained only for the RITC-PEI immobilized MPs indicates a slight aggregation of the particles under the high electric field or a broader distribution of the RITC labeling efficiency for PEI, which (46) Gilges, M.; Kleemiss, M. H.; Schomburg, G. Anal. Chem. 1994, 66, 20382046.

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would cause multiple peaks.47,48 PEI has a polydispersity in molecular weight according to random branched structure, so that the fluorescent labeling efficiency might be different in each polymer chain. By adding DxS into the RITC-PEI immobilized MP suspension, the migration direction changed again from cathode to anode, which clearly showed the immobilization of anionic DxS onto the outer surface of the MPs. Furthermore, the spikelike peaks disappeared with the immobilization of DxS, which also indicates the change in the surface properties of the particles. The electrophoretic mobility (µep) and the relative standard deviation (RSD) of the detection time for these particles are summarized in Table 1. The result shows that the prepared particles can be detected with good reproducibility in the PVAcoated capillaries. In addition, the obtained good reproducibility indicates that the ionic polymers with fluorescent molecules and ALs for lipoproteins were stably immobilized onto the MP surface. For evaluation of the affinity ability of the prepared MFMPs, µep of the mixed suspension of the MFMPs and lipoproteins, i.e., LDL and HDL, was estimated by CE. As shown in Figure 3, µep was almost independent of the concentration of HDL, while by adding LDL, a considerable decrease in µep was observed. This change in µep clearly indicates that the selective binding of LDL onto the surface of the MFMPs alters their surface charge density. On the other hand, the constant µep values of the MFMPs immersed into a HDL solution show that the prepared MFMPs possess no affinity with HDL, i.e., the good selectivity for LDL in the lipoprotein analysis. The result agrees well with the fact that the adsorption of HDL onto the DxS layer was negligible as reported by Tani.49 Although LDL is negatively charged at pH (47) Jorgenson, J. W.; Lukacs, K. D. Science 1983, 222, 266-272. (48) Craig, D. B.; Dovichi, N. J. Anal. Chem. 1998, 70, 2493-2494. (49) Tani, N. Artif. Organs 1996, 20, 922-929.

7.4, it is well-known that LDL contains a positively charged part of clustering amino acids, which specifically binds negatively charged sulfate or sulfonate groups.50 The dissociation constant of LDL to the DxS with Mw of 1000 is reported to be in the order of 10-8 M, which is varied by the charge density (sulfur content) of DxS. Additionally, it was also reported that the effect of molecular weight on the dissociation constant is not remarkable in DxS with many sulfate groups.49 Since DxS with a higher molecular weight of 500 000 and a higher sulfur content of 17% was employed in this study, the dissociation constant is expected to be on the order of 10-8 M. Thus, multiple ionic interactions enable DxS to bind tightly to both the MFMP surface and LDL during the analysis. Actually, a reproducible analysis of the LDLMFMPs was attained with the RSD for the detection time of 1.2%, which is nearly as good as for the bare MFMPs, which demonstrates that LDL tightly binds onto the MFMP surface and the dissociation from their surface is almost negligible during the analysis. Since our strategy shown in Figure 1 requires a kinetically stable MFMP-target analyte complex, this suppressed dissociation between LDL and DxS is appropriate for the selective separation. Furthermore, such a smaller dissociation constant enables us to roughly estimate the adsorbed amount of LDL onto the MFMP surface from the saturated concentration for the LDL immobilization shown in Figure 3, and as a result, the saturated amount of the immobilized LDL was evaluated to be in the order of subattomoles per particle. Online Preconcentration and Selective Separation of LDL Using MFMPs and Magnetic Field. In this study, we expected that magnetic trapping of the MFMPs was applicable for the online preconcentration and the sensitivity enhancement. For the trapping of the prepared MFMPs in the capillary, a magnet was placed adjacently onto the capillary surface. To obtain a sharper peak with good reproducibility, a magnet with a shorter length of 1 mm was employed. On the basis of the bright-field observation with the optical microscope, it was confirmed that all the injected MFMPs were successfully captured by the magnetic field under a separation voltage of below 10 kV (190 V/cm). Above 15 kV, untrapped MFMPs were frequently observed and migrated toward the anodic end. To attain efficient concentration of the MFMPs, the application of a magnet length of less than 1 mm would be favorable. However, the magnetic trapping force may decrease with decreasing length of magnet, so that a weaker capturing force would result in a lower electric field less than 10 kV, which brings a longer analysis time. Thus, the applied voltage and the length of the magnet were set at 10 kV and 1 mm, respectively, in the remaining experiments. Figure 4 shows the online concentration of the MFMPs using the magnetic field. Compared to a conventional injection, a sharper and higher peak was obtained by the magnetic capturing. Although the injection time was 60-fold longer compared to the normal injection condition, the 53-fold increase in the peak height was attained for the MFMPs, which indicated that a sufficient concentration could be achieved by magnet trapping. It should be noted that, furthermore, no peak appeared during the application of the magnetic field as shown in Figure 4b, so that the efficient magnetic trapping of the MFMPs was accomplished under this experimental condition. (50) Weisgraber, K. H.; Rall, S. C., Jr. J. Biol. Chem. 1987, 262, 11097-11103.

Figure 4. CE analysis of MFMPs under (a) no and (b) applied magnetic field conditions. Sample injection, 10 kV for (a) 10 and (b) 600 s; MFMP concentration, 1 × 107 particles/mL; PVA-coated capillary, 53 cm × 50 µm i.d. (28 cm to the detector). Other conditions are as in Figure 3.

Figure 5. Effect of injection time on SEFheight of bare and LDLbound MFMPs. LDL concentration, 4.6 × 10-9 M. Other conditions are as in Figure 4.

To evaluate the concentration efficiency, SEFheight was calculated with changing injection volume of the sample dispersion. Figure 5 clearly shows that SEFheight increased with increasing injection time. As can be seen, ∼100-fold sensitivity enhancement was easily attained under the optimized condition, which also indicates that capturing and releasing of the MFMPs was successfully achieved in the capillary. In the case of LDL-MFMPs, acceptable reproducibilities can be obtained at injection times of 300-600 s. Taking into account total analysis time and analytical reproducibility, the injection time of 300 s may be a better selection in this method. Since SEFheight at 300 s might be too low to evaluate the concentration performance, however, we selected the injection time of 600 s in the remaining experiments. On the other hand, the values of SEFheight for the LDL-MFMPs were lower than those for the bare MFMPs. This would be due to a heterogeneity of particle size or charge of the LDL-MFMPs, which caused the electrophoretic mobility difference in the each MFMP as reported by Petersen and Ballou.51 Since the distribution of the molecular weight of LDL is extremely broad,52 the LDL-MFMPs would be also heterogeneous in size, shape, or surface charge, which would (51) Petersen, S. L.; Ballou, N. E. Anal. Chem. 1992, 64, 1676-1681.

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Figure 6. Online concentration and selective separation of lipoproteins using the magnetic field. Sample concentration, (a) no proteins, (b) 1.3 × 10-7 M HDL and 4.6 × 10-9 M LDL; MFMP concentration, 1 × 107 particles/mL; sample injection, 10 kV for 600 s.

reduce SEFheight by adding LDL into the MFMP suspension. However, the trapped MFMPs possess a remnant magnetization property even after the removal of the magnetic field, which leads to clustering of MPs. This hysteric magnetization is sometimes problematic; however, in this case, it works well to suppress a zone broadening of the MFMPs. Thus, sufficient concentration efficiency was obtained with the magnetic trapping of the MFMPs. Magnetic capturing of the MFMP binding target analytes via a strong affinity is expected to be applied to the simultaneous selective separation and concentration. Online concentration of the target analytes (LDL) and the separation of other components (HDL) are demonstrated as shown in Figure 6. In the analysis of the bare MFMPs with the magnetic field, only one peak was observed after the removal of the magnet as shown in Figure 6a. For the MFMPs suspension containing both LDL and HDL, on the other hand, HDL was detected as a broader peak during the application of the magnetic field, so that a successful separation was obtained with trapping and concentrating the LDL-MFMPs in Figure 6b. As can be seen in Figure 6, the detection time of the LDL-MFMPs is apparently different from that of the bare MFMPs, which enables to distinguish the LDL-MFMPs from the bare MFMPs. As well as HDL, bovine serum albumin could be also separated from the LDL-MFMPs. The reproducibility of the detection time and the peak height for concentrating the LDL-MFMPs were acceptable with RSDs of 0.49 and 11% (n ) 3), respectively. These results indicate that the selective separation and online preconcentration are easily attained by using the MFMPs with the simple experimental procedures, i.e., place and remove the magnet. Generally, the MPs with ALs for biogenic compounds are employed for the separation of target molecules from other components in a sample vial prior to the analysis, offline procedure.53 Consequently, the offline process is difficult to automate and a considerable loss of sample is sometimes (52) Assman, G. Lipid Metabolism and Atherosclerosis; Schattauer, F. K., Ed.; Verlag Gumb H.: Stuttgart, 1982; p 14.

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Figure 7. Electropherograms of (a) bare and (b, c) LDL immobilized MFMPs obtained with LIF detection. Particle concentration, (a) 1 × 107, (b) 1 × 107, and (c) 1 × 103 particles/mL; LDL concentration, (a) 0, (b) 4.6 × 10-9, and (c) 4.6 × 10-13 M; PVA-coated rectangular capillary; 53 cm × 50 µm i.d. (28 cm to the detector); sample injection, 10 kV for 600 s

problematic during handling of the MPs. On the contrary, our approach with the magnetic trapping of the MFMPs in CE would not require labor-intensive washing and labeling processes, which is suitable for the automated analysis. Application of LIF Detection. As shown in Figure 3, LDL binding onto the MFMPs surface can be identified by the difference in µep compared to the bare MFMPs. Since LDL is detected as a complex of MFMPs, it can be analyzed without a direct fluorescence labeling of the analytes even in the CE-LIF analysis. Figure 7 shows the CE-LIF analysis of the bare and LDL-MFMPs. Since the fluorescent spectrum obtained at the peak time coincided with that of PEI-RITC, it was confirmed that these MFMPs were successfully detected with LIF. The bare MFMPs were also detected after the removal of the magnetic field as well as Figure 6a. However, frequent spikelike peaks were observed. Although the total peak area versus MFMP-LDL concentration plot showed an acceptable linearity (R2 ) 0.984) ranging from 2 × 102 to 1 × 104 particles/mL, the RSD for the peak area was estimated to be 26-35%. As reported by Duffy et al.,54 this poor deviation is attributed to an inhomogeneous excitation of microspheres, which is caused by different trajecto(53) Paquette, D. M.; Banks, P. R. Electrophoresis 2001, 22, 2391-2397.

ries of the MFMPs through the laser beam. Additionally, both heterogeneous RITC labeling of PEI and the size distribution of the MFMPs might also contribute to the poor reproducibility. Thus, the fluorescence intensity would not provide the precise amount of the MFMPs detected by LIF. However, improvement of the detection configuration, e.g., the use of a narrower capillary, homogeneously fluorescent-labeled polyelectrolyte and monodispersive particles, would result in a lower RSD of the peak height and area. Since the detection time of the MFMPs obtained in this experiment was strongly dependent on the LDL concentration as shown in Figure 3, the immobilized amount of LDL onto the MFMPs can be estimated without any information about the peak intensity, which would reduce partly the significance of the reproducibility for the fluorescence intensity. As shown in Figure 7b, a successful detection of the LDLMFMPs was also achieved in CE-LIF. It should be marked that the detection time was apparently different from the bare MFMPs, which demonstrated that the indirect labeling of LDL was attained by immobilizing onto the MFMPs, modified solid-phase labeling. It is known that solid-phase labeling can avoid several drawbacks in pre- or postcolumn labeling in liquid phase, e.g., dilution of analytes caused by a labeling reagent and higher background signal due to excess derivatizing reagents.55-57 In addition to these advantages, our modified solid-phase labeling has several merits. As discussed above, selective labeling can be conducted using biologically specific interaction between the outer AL layer on the MFMPs and the target analytes, which modified the affinity probe method reported by Shimura and Karger.58 Moreover, the reduction of the affinity interaction due to a direct fluorescent derivatization of analytes would be avoided in the modified solid-phase labeling since the native target analytes should directly attach to the AL layer. In the case of a conventional solid-phase labeling, the elution process for extracted and labeled analytes is required, whereas in our approach, the target analyte-bound MFMPs are directly introduced into the separation capillary without any elution. Thus, a time-consuming elution process can be omitted and the dilution of the target analytes is avoided by this labeling technique. Among these advantages, the suitability for labeling of the target analytes with a lower concentration is expected to be applied to the analysis for a trace amount of LDL, so that the limit of detection (LOD) was estimated by the CE-LIF measurements. For the preparation of a lower concentration sample, a smaller amount of LDL was added into a 10 000 times dilution of the stock MFMPs dispersion. Figure 7c shows the electropherogram of the diluted MFMPs sample containing LDL. A peak according to the MFMPs was detected at 8.5 min, which agreed well with that of the LDL-MFMPs shown in Figure 7b. Since the LDL/MFMPs concentration ratio was equivalent in spite of the higher dilution (54) Duffy, C. F.; McEathron, A. A.; Arriaga, E. A. Electrophoresis 2002, 23, 2040-2047. (55) Nondek, L.; Frei, R. W.; Brinkman, U. A. Th. J. Chromatogr. 1983, 282, 141-150. (56) Bourque, A. J.; Krull, I. S. J. Chromatogr. 1991, 537, 123-152. (57) Pinto, D. M.; Arriaga, E. A.; Sia, S.; Li, Z.; Dovichi, N. J. Electrophoresis 1995, 16, 534-540. (58) Shimura, K.; Karger, B. L. Anal. Chem. 1994, 66, 9-15. (59) Duffy, C. F.; Gafoor, S.; Richards, D. P.; Admadzadeh, H.; O’Kennedy, R.; Arriaga, E. A. Anal. Chem. 2001, 73, 1855-1861. (60) Ahmadzageh, H.; Dua, R.; Presley, A. D.; Arriaga, E. A. J. Chromatogr., A 2005, 1064, 107-114.

of the sample dispersion, this identical detection time was obtained. Thus, LDL can be labeled even under the extremely lower concentration condition and detected as the complex with the MFMPs by LIF. In this experimental condition, the LOD for LDL was calculated to be 4.3 × 10-13 M. Therefore, the trace LDL analysis was successfully achieved by the magnetic trapping of the MFMPs. In this study, we selected lipoproteins as a model sample. Since lipoproteins are heterogeneous proteins with regard to molecular weight and size as mentioned above, the observed peak of the lipoprotein-bound MFMPs may become broader. If other homogeneous proteins are employed, peak broadening is expected to be suppressed, and as a result, a higher sensitivity can be obtained. In addition, further optimization of fluorescent molecules, optical configuration, and data analysis59,60 would provide a higher performance LIF detection. CONCLUSION MFMPs were prepared on the basis of the PEM coating technique to improve the performance ACE analysis. The prepared MFMPs showed three functions of magnetic response, solid-phase indirect fluorescent labeling, and ALs for LDL. By using the MFMPs, LDL was selectively bound to the DxS layer immobilized onto the MFMPs surface via the specific biological interaction and indirectly labeled by the inner layer of RITC-PEI. The magnetic trapping of the LDL-MFMPs provided the selective concentration and separation from HDL. Furthermore, LIF detection of the concentrated and purified MFMPs can detect subpicomolar LDL without any complicated operations. Our developed ACE method can be applied to the automatic analysis of a lowconcentration sample and rapid diagnosis by using various ALs immobilized onto the MFMP surface. In this study, the CE apparatus was mainly employed for the qualitative analysis of the MFMPs, so that the powerful separation ability of CE might not be fully utilized due to the introduction of only one specific interaction between DxS and LDL to the prepared MFMPs. Thus, the developed method is expected to be applied for the CE separation of multiple target analytes by using the MFMPs with several ALs, and work along this line is now in progress in this laboratory. ACKNOWLEDGMENT The authors thank Prof. Hiroo Iwata (Institute for Frontier Medical Sciences, Kyoto University) for particle-counting measurements. They also thank JSR and Japan VAM & POVAL for supplying the magnetic polymer particles and poly(vinyl alcohol), respectively. This work was partly supported by the Grant-in-Aid for 21st Century Program, COE for a United Approach for New Materials of Science, from the Ministry of Education, Culture, Sports, Science and Technology, Japan. K.O. is grateful to the Grant-in-Aid for Scientific Research (15350041) from the Japan Society for the Promotion of Science.

Received for review February 8, 2007.

September

7,

2006.

Accepted

AC061693Q Analytical Chemistry, Vol. 79, No. 8, April 15, 2007

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