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Modification and characterization of polystyrene-based magnetic microspheres and comparison with albumin-based magnetic microspheres
Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
Modi"cation and characterization of polystyrene-based magnetic microspheres and comparison with albumin-based magnetic microspheres Jhunu Chatterjee, Yousef Haik*, Ching-Jen Chen Department of Mechanical Engineering, Biomagnetic Engineering Laboratory, FAMU-FSU College of Engineering, 2525 Pottsdamer Street, Tallahassee, FL 32310, USA
Abstract Polystyrene- and albumin-based magnetic microspheres for red blood cell separation were modi"ed and characterized by scanning electron and atomic force microscopy. Albumin microspheres show higher coupling e$ciency with the protein, and protein-modi"ed albumin microspheres bind the red blood cells more e$ciently than the polystyrene-based microspheres. 2001 Elsevier Science B.V. All rights reserved. Keywords: Human serum albumin; Polystyrene microspheres; Albumin microspheres; Magnetic microspheres; Maghemite; Covalent coupling; Lectin; Erythrocyte; Scanning electron microscopy; SEM; Atomic force microscopy; AFM; Protein assay
1. Introduction E!ective cell separation is a primary and most important footstep for many clinical and immunological applications [1,2]. One of these applications is the separation of red blood cells (RBC) from the whole blood for photopheresis treatment of white cells. In the photopheresis treatment of white blood cells, light is used to activate a compound that binds to the DNA and may cause an increase in immunogeneity and thus stimulate the patient's own immune system [3]. A magnetic separation device was developed to isolate the RBCs from whole blood while the photopheresis treatment is administered on the patient's leukocytes [4,5]. This
* Corresponding author. E-mail address: [email protected] (Y. Haik).
device was designed for on-line, continuous separation of RBCs from whole blood. For the magnetic separation, the RBCs were coupled to albuminbased magnetic particles. A magnetic bed was used to retain the red cells in the separation chamber while the white cells and plasma proceeds to the photopheresis unit. The albumin-based magnetic microspheres (ALBMMS) were synthesized in house and were chosen for their biodegradability. Recently, a hybrid system for separation was being tested. In the hybrid system, a centrifuge is used to isolate the plasma and the bu!y coat (a thin band ((1%) of white cells and platelets at the plasma interface) from whole blood and a magnetic separation unit to isolate red blood cells from the bu!y coat. The amount of RBCs in the bu!y coat is small and may be removed with more economical magnetic particles than the albumin based ones. Polystyrene magnetic microspheres (PSMMS) were
0304-8853/01/$ - see front matter 2001 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 8 8 5 3 ( 0 0 ) 0 1 2 2 3 - 3
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J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
chosen to substitute for the ALBMMS in the hybrid separation system. In this paper, a morphological study of the two types of microspheres that are used for blood cell separation, namely polystyrene- and albumin-based ones, was performed. This characterization is important to ensure e$cient binding to the RBC. The characterization procedures followed here for microspheres bound with RBCs included scanning electron microscopy (SEM), atomic force microscopy (AFM) and determination of protein coupling e$ciency. ALBMMS were synthesized and modi"ed for coupling to RBCs. Nanosized magnetic particles in the order of 50 nm were coated with albumin. Modi"cation of these microspheres was done by biotinylated lectin, a carbohydrate binding protein that can e!ectively couple with RBC. These microspheres were tested for toxicity and were found safe and biodegradable. They are recommended for use when RBCs are to be given back to the patient. Other synthetic polymer-based microspheres (PSMMS) have been chosen to couple with the RBCs for times, when the RBCs are to be discarded. The synthetic polymer-based microspheres cost less in manufacturing than the albumin-based microspheres. Modi"cation of these microspheres was done by covalent coupling to lectin. In order to achieve covalent coupling with protein di!erent functional groups have been reported [5}7]. For this study, carboxyl modi"ed polystyrene microspheres were selected and were further modi"ed by lectin. Both SEM and AFM are useful to study the surface characteristics of the microspheres. Use of SEM for determining shape and size of particles in emulsion and dispersion system is extremely common and reported for various polymeric materials [8,9]. High-resolution surface structures of non-conductive specimens, as well as specimen in aqueous medium can be obtained by atomic force microscopy. The structure of the biological molecules can thus be determined under native condition [10]. Atomic force microscopy is based on attractive repulsive forces occurring between a cantilever mounted probe and a sample and is not restricted to conductive probe/sample systems. It has been
applied to the surface topology of various materials [11}13].
2. Materials and methods Carboxyl modi"ed polystyrene microspheres of 1}2 m diameter were obtained from Bangs Laboratory. Human serum albumin, n-hexane, 25% glutaraldehyde, mineral oil, sorbitan sesquioleate, 1-cyclohexyl-3-(2-morpholinoethyl) carbodimide metho-p-toluenesulfonate (water soluble carbodiimide), lectin and biotinylated lectin were obtained from Sigma Chemical Company. BCA protein estimation kit, Sulfo SMCC, succinimidylS-acetylthioacetate (SATA), and immunopure avidin were obtained from Pierce Chemical Company. Maghemite was obtained from Nanophase Technologies Corporation. 2.1. Preparation of the HSA-based magnetic microsphere Chemically crosslinked albumin magnetic microspheres for blood separation were synthesized with a few di!erences from the reported methods [14,15]. A 10 ml aqueous solution of 250 mg HSA and 250 mg maghemite was added dropwise to a mixture of 40 ml n-hexane, 10 ml light mineral oil and 0.5 ml of sorbitan sesquioleate to form a water in oil inverse emulsion system. After sonication for 10 min at 50% amplitude, 10 ml of 25% glutaraldehyde saturated with toluene was added to this mixture and mechanically stirred with te#on paddle stirrer at 200 rpm for 2 h. The suspension was then centrifuged at 3000 rpm for 15 min, the supernatant decanted, the microspheres washed repeatedly with petroleum ether and acetone and "nally dispersed in distilled water. Modi"cation of HSA microspheres with biotinylated lectin was done by preparation of thiolated avidin which act as a bridge between biotinylated lectin and iron oxide bound HSA. Avidin was dissolved in PBS (pH 7.4) at a concentration of 10 mg/ml. A stock solution of SATA was prepared in dimethyl sulphoxide at a concentration of 13 mg/ml. Twenty-"ve milliliter of SATA stock solution was added to each milliliter of the avidin
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
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solution and reacted for 30 min. Maghemite-bound HSA microspheres were dispersed in PBS bu!er (pH 7.4) at a concentration of 10 mg/ml, 3.3 mg of sulfo-SMCC were added to each milliliter of HSA solution and reacted for 30 min and then added to the thiolated avidin solution and reacted for 2 h at room temperature. HSA and avidin were reacted in 4:1 molar proportion to form the conjugate properly. The mixture was then centrifuged and washed repeatedly with PBS bu!er to remove unreacted avidin. Aproximately 5.15 mg (based on the amount calculated for the monolayer formation [16]) of biotinylated lectin was added to 25 ml of the avidin-modi"ed HSA microsphere suspension and reacted for 4 h at room temperature.
were prepared following a set of steps of "xing [19], washing and dehydration [20] to maintain the physiological condition necessary for retaining the shape of the RBC and thus preventing artifacts.
2.2. Modixcation of the polystyrene microspheres
2.5. Determination of the protein coupled to microspheres
PSMMS with a carboxyl functional group were further modi"ed by coating with lectin. The functional carboxyl group was converted into an ortho acyl-urea intermediate, which can bind to a protein according to di!erent coupling procedures available in the literature [17]. Typically, 1 ml of the carboxyl modi"ed microspheres (10% solid) was repeatedly washed with phosphate bu!ered saline using a magnet for the complete removal of the surfactant and redispersed in 10 ml of PBS bu!er. One hundred milligram of the water-soluble carbodiimide was added and reacted for 20 min at room temperature. The mixture was then washed twice with PBS bu!er and "nally resuspended in 5 ml of the PBS bu!er. Five milligram of lectin (approximately "ve times of the amount required for the calculated monolayer formation) was dissolved in 5 ml of PBS bu!er. The microspheres suspension was then added to the protein solution to avoid agglomeration and reacted for 4 h at room temperature. 2.3. Scanning electron microscopy A JEOL (JSM 840) instrument was used for SEM to characterize both types of microspheres after modi"cation and after coupling to RBC. Modi"ed microsphere specimens for SEM were prepared following the procedure of Margel et al. [18]. Specimens of microspheres coupled to RBC
2.4. Atomic force microscopy A D-3000 (Digital Instruments, Santa Barbara) in `tapping modea was used for the AFM. All specimens were prepared exactly in the same way as for SEM. The specimen "lms were formed on glass slides and did not need any metal coating. The RBC specimens were prepared on a cover slip using the same procedure as in SEM and were then further mounted on a glass slide for AFM.
A BCA protein assay kit was used to estimate the percentage of protein coupled to the microspheres. A standard calibration curve (50}500 mg) was made with bovine serum albumin (BSA) and the absorbance measured in an Ultraspec 2000 UV/VIS spectrophotometer (Pharmacia Biotech) at 562 nm. Carboxyl modi"ed microspheres (solid content 7.4%) and albumin microspheres (solid content 1%) were modi"ed with three di!erent concentrations of the protein. Three sets of microsphere suspensions were made using three di!erent protein concentrations. The absorbance of supernatant was measured and the protein coupled to the microspheres was obtained by di!erence.
3. Results and discussion Scanning electron micrograph for both ALBMMS and PSMMS are shown in Fig. 1(a) and (b). PSMMS are monodispersed particles having diameter 1}2 m and the albumin microspheres have shown a broad distribution of particle ranging with submicron to 1 m diameter and they appear as #akes also. Albumin microspheres were found to be always agglomerated even in extremely dilute dispersion. Micrographs obtained by SEM for proteincoated carboxyl modi"ed particles are shown in
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J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
Fig. 2. SEM micrograph of lectin coated polystyrene magnetic microspheres.
Fig. 1. Scanning electron micrograph (SEM) of (a) polystyrene magnetic microspheres and (b) albumin magnetic microspheres.
Fig. 2. No signi"cant change in shape observed after protein coating but they appeared as more agglomerated structure and they are connected with each other. Some of them showed increase in size also along with holes in it. Holes appeared in the protein-coupled polystyrene microspheres may be due to the immiscibility of the seed polymer with polymer formed in the later steps or due to the excess monomer compared to the polymer in the seed causing low viscosity and ease transport of the phase separated seed polymer as reported by Ugelstad and coworkers for the pure polystyrene particles [21]. The size of the holes basically determines the size of the seed particle, which is converted to the "nal microspheres by
more than one polymerization steps. No signi"cant change is observed in the morphology of the albumin particle before and after coating. The SEM micrographs of the RBC coupled with protein coated albumin and polystyrene microspheres are shown in Figs. 3(a), and (b). Albumin particles have coupled with RBC as a cluster form but the polystyrene particle can couple singly with the RBC. Atomic force micrographs for the polystyrene and albumin microspheres are shown in Fig. 4. The spherical surface of PSMMS is more clearly observed in the 3D view obtained in atomic force microscopy. AFM characterizes the albumin microspheres as distinctly spherical in shape even in agglomerated state. The section analysis in Fig. 4(c) shows that the albumin microspheres are in submicron range. A top view of the protein-coated polystyrene microspheres is shown in Fig. 5(a) clearly indicating the presence of structures on the surface of the spheres. A 3D #attened view of the protein-coated surface with an extremely nice contrast shows structures due to protein (Fig. 5(b)). The AFM micrographs for the protein-coated albumin are shown in Fig. 6. Protein molecules seem to form a layered structure on the clusters of the spheres. The AFM micrograph for the polystyrene coupled RBCs is shown in Fig. 7. Albumin microspheres coupled with RBCs are shown in
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
Fig. 3. SEM micrograph of (a) a RBC coupled with lectin coated PSMMS and (b) a RBC coupled with protein coated ALBMMS.
Fig. 8(a). This micrograph shows a distinct attachment of the microspheres to the RBC. Albumin microspheres were highly agglomerated and often were attached to the RBCs as a cluster of microspheres. In the phase mode of AFM distinct attachment of the albumin microspheres on the membrane of the RBC is shown in Fig. 8(b). Functional groups present in the polymeric microspheres surface, protein concentration and other factors such as pH of the medium could be a probable cause of formation of highly agglomerated structure for the protein-coated microspheres of both types. When protein forms 䉴 Fig. 4. Atomic force 3D micrograph (AFM) of (a) a polystyrene magnetic microsphere, (b) magnetic albumin microspheres, and (c) a sectional analysis of albumin microspheres.
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Fig. 6. AFM micrograph of protein-coated albumin microspheres.
Fig. 5. AFM 3D micrograph of (a) protein coated polystyrene microsphere (top view) and (b) of a protein coated surface of polystyrene microsphere (#attened view).
a covalent bond with the surface of the microspheres, they become neutral and may aggregate with each other in absence of any charges. Investigations of the nature of the interaction of this particular lectin with polymer and albumin microspheres particle are still in progress. Fig. 9 shows the variation of the optical density (OD) with concentration of the protein (in g/ml) added in the three di!erent sets for both albumin based and polystyrene based microspheres. Optical density is found to be very high showing lower amount of protein coupling to the carboxyl modi"ed polystyrene microspheres and the OD increase
Fig. 7. AFM micrograph of a RBC coupled with a polystyrene microsphere.
basically with increase in amount of the added protein. But for the albumin microspheres optical density of the decanted clear liquid is very low indicating higher amount of protein coupling and the OD values are almost same for the higher concentrations of the added protein, which means that the amount of the unbound protein is almost equal in the decanted clear liquid. This observation also shows more protein coupling to the albumin microspheres with increasing concentration of
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
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microspheres is not certainly enough for the 100% protein coupling and hence there is always unbound protein. Lack of site on the microspheres is de"nitely caused by the agglomeration of the spheres amongst themselves during the washing and other processing steps. As all other parameters such as temperature, pH of the medium, mixing time and speed are indentical in the process of protein coupling, the available site on the microspheres cannot change for the three di!erent samples with same solid content. We have to also eliminate the possibility of protein decoupling as the binding takes place by covalent bonding. So, the probable reason for higher protein coupling when the concentration of the added protein is increased could be a protein multilayer formation [22] on the available site. Chances of cluster formation are higher in higher concentration of protein and if these clusters couple with the microspheres as observed in Fig. 6 rather than small number of molecules, there will be less amount unbound protein. But this conclusion needs to be veri"ed by studying the morphology of the protein in di!erent concentrations. Albumin microspheres are certainly showing more coupling than the carboxyl modi"ed polystyrene microspheres. To establish the coupling e$ciency more quantitatively for the above microspheres more data is required and the experiment is in progress.
Fig. 8. AFM micrograph of (a) a RBC coupled with albumin microspheres and (b) a RBC coupled with albumin microspheres using phase mode.
added protein. Amount of the protein added is usually the calculated amount required for the monolayer formation [16] and this concentration is represented as the "rst concentration and other two concentrations are double and three times to this for both cases. Fig. 10 shows the percentage of the protein coupled for a certain amount of added protein to both kinds of microspheres. In both cases, percentage of protein coupled increases with increase in amount of the added protein. Available site on the
4. Conclusion Both carboxyl modi"ed polystyrene microspheres and albumin microspheres have shown effective coupling to RBCs when they are modi"ed with lectin and biotinylated lectin. Our study shows that albumin microspheres have some advantage over the synthetic polymer-based microspheres as RBCs separated by albumin microspheres can be reinjected into a patient. Characterization by AFM is able to show the details of the protein coupling to the microspheres and the coupling of the red blood cell to these microspheres. The presence of submicron sized particles in both SEM and AFM micrographs point at the possibility of forming nanosized albumin particles using modi"ed synthesis conditions. The amount of the protein
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Fig. 9. Plot of OD for unbound protein.
References [1] [2] [3] [4] [5] [6] [7] Fig. 10. Plot of absorbed protein as a function of amount of added protein.
[8] [9] [10]
coupled could not be optimized yet as the coupling procedure showed kinetics that might be optimized with more data for other di!erent concentrations.
[11]
[12]
Acknowledgements Part of this work is funded by Therakos Incorporation, USA.
[13] [14] [15]
A. Rembaum, W.J. Dreyer, Science 208 (1980) 364. A. Rembaum, S. Margel, Br. Polym. J. 10 (1978) 275. R. Edelson, Sci. Amer. 259 (1988) 68. C.J. Chen, Y. Haik, V.M. Pai, US Patent 6,036,857, 2000. Y. Haik, V.M. Pai, C.J. Chen, J. Magn. Magn. Mater. 194 (1999) 254. M. Okubo, Y. Iwasaki, Y. Yamato, Colloid Polym. Sci. 270 (1992) 733. T. Delair, V. Margnet, C. Pichot, B. Mandrand, Colloid Polym. Sci. 272 (1994) 962. F. Sauzedde, A. Elaissari, C. Pichot, Colloid Polym. Sci. 277 (1999) 846. H. Bamnolker, B. Nitzan, S. Gura, S. Margel, J. Mater. Sci. Lett. 16 (1997) 1412. F. Sauzedde, A. Elaissari, C. Pichot, Colloid Polym. Sci. 277 (1999) 846. H.J. Butt, E.K. Wol!, S.A.C. Gould, B. Dixon Northen, C.M. Peterson, P.K. Hansma, J. Struct. Bio. 105 (1990) 54. G. Binning, C.F. Quate, C.H. Gerber, Phys. Rev. Lett. 56 (1986) 930. H. Butt, K.H. Downing, P.K. Hansma, Biophys. J. 58 (1990) 1473. T. Ishizaka, K. Endo, M. Koishi, J. Pharm. Sci. 70 (1981) 358. W.A. Knepp, Master's Thesis, University of Florida, 1991.
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29 [16] H.J. Hager, Latex Polymer Reagents for Diagnostic Tests, US Patent 3,857,931, 1974. [17] L.B. Bangs, Uniform Latex Particles, Technical Notes Seradyn, 1999. [18] M. Shahar, H. Meshulam, S. Margel, J. Polym. Sci., Polym. Chem. Edn. 24 (1986) 203. [19] D. Platt, Blood Cells, Rheology and Aging, Springer, Berlin, 1998, p. 32.
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[20] J.J. Bozzola, L.D. Russell, Electron Microscopy, 2nd Edition, Jones and Bartlett, Sudbury, MA, 1992, p. 54. [21] W.S. Prestvik, A. Berge, P.C. Mork et al., in: U. Hafeli, W. Schutt, J. Teller, M. Zborowski (Eds.), Scienti"c and Clinical Applications of Magnetic Carriers, Plenum Press, New York, 1997, p. 16. [22] D. Kowalczyk, J.P. Marsault, S. Slomakowski, Colloid Polym. Sci. 274 (1996) 513.
Modi"cation and characterization of polystyrene-based magnetic microspheres and comparison with albumin-based magnetic microspheres Jhunu Chatterjee, Yousef Haik*, Ching-Jen Chen Department of Mechanical Engineering, Biomagnetic Engineering Laboratory, FAMU-FSU College of Engineering, 2525 Pottsdamer Street, Tallahassee, FL 32310, USA
Abstract Polystyrene- and albumin-based magnetic microspheres for red blood cell separation were modi"ed and characterized by scanning electron and atomic force microscopy. Albumin microspheres show higher coupling e$ciency with the protein, and protein-modi"ed albumin microspheres bind the red blood cells more e$ciently than the polystyrene-based microspheres. 2001 Elsevier Science B.V. All rights reserved. Keywords: Human serum albumin; Polystyrene microspheres; Albumin microspheres; Magnetic microspheres; Maghemite; Covalent coupling; Lectin; Erythrocyte; Scanning electron microscopy; SEM; Atomic force microscopy; AFM; Protein assay
1. Introduction E!ective cell separation is a primary and most important footstep for many clinical and immunological applications [1,2]. One of these applications is the separation of red blood cells (RBC) from the whole blood for photopheresis treatment of white cells. In the photopheresis treatment of white blood cells, light is used to activate a compound that binds to the DNA and may cause an increase in immunogeneity and thus stimulate the patient's own immune system [3]. A magnetic separation device was developed to isolate the RBCs from whole blood while the photopheresis treatment is administered on the patient's leukocytes [4,5]. This
* Corresponding author. E-mail address: [email protected] (Y. Haik).
device was designed for on-line, continuous separation of RBCs from whole blood. For the magnetic separation, the RBCs were coupled to albuminbased magnetic particles. A magnetic bed was used to retain the red cells in the separation chamber while the white cells and plasma proceeds to the photopheresis unit. The albumin-based magnetic microspheres (ALBMMS) were synthesized in house and were chosen for their biodegradability. Recently, a hybrid system for separation was being tested. In the hybrid system, a centrifuge is used to isolate the plasma and the bu!y coat (a thin band ((1%) of white cells and platelets at the plasma interface) from whole blood and a magnetic separation unit to isolate red blood cells from the bu!y coat. The amount of RBCs in the bu!y coat is small and may be removed with more economical magnetic particles than the albumin based ones. Polystyrene magnetic microspheres (PSMMS) were
0304-8853/01/$ - see front matter 2001 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 8 8 5 3 ( 0 0 ) 0 1 2 2 3 - 3
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J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
chosen to substitute for the ALBMMS in the hybrid separation system. In this paper, a morphological study of the two types of microspheres that are used for blood cell separation, namely polystyrene- and albumin-based ones, was performed. This characterization is important to ensure e$cient binding to the RBC. The characterization procedures followed here for microspheres bound with RBCs included scanning electron microscopy (SEM), atomic force microscopy (AFM) and determination of protein coupling e$ciency. ALBMMS were synthesized and modi"ed for coupling to RBCs. Nanosized magnetic particles in the order of 50 nm were coated with albumin. Modi"cation of these microspheres was done by biotinylated lectin, a carbohydrate binding protein that can e!ectively couple with RBC. These microspheres were tested for toxicity and were found safe and biodegradable. They are recommended for use when RBCs are to be given back to the patient. Other synthetic polymer-based microspheres (PSMMS) have been chosen to couple with the RBCs for times, when the RBCs are to be discarded. The synthetic polymer-based microspheres cost less in manufacturing than the albumin-based microspheres. Modi"cation of these microspheres was done by covalent coupling to lectin. In order to achieve covalent coupling with protein di!erent functional groups have been reported [5}7]. For this study, carboxyl modi"ed polystyrene microspheres were selected and were further modi"ed by lectin. Both SEM and AFM are useful to study the surface characteristics of the microspheres. Use of SEM for determining shape and size of particles in emulsion and dispersion system is extremely common and reported for various polymeric materials [8,9]. High-resolution surface structures of non-conductive specimens, as well as specimen in aqueous medium can be obtained by atomic force microscopy. The structure of the biological molecules can thus be determined under native condition [10]. Atomic force microscopy is based on attractive repulsive forces occurring between a cantilever mounted probe and a sample and is not restricted to conductive probe/sample systems. It has been
applied to the surface topology of various materials [11}13].
2. Materials and methods Carboxyl modi"ed polystyrene microspheres of 1}2 m diameter were obtained from Bangs Laboratory. Human serum albumin, n-hexane, 25% glutaraldehyde, mineral oil, sorbitan sesquioleate, 1-cyclohexyl-3-(2-morpholinoethyl) carbodimide metho-p-toluenesulfonate (water soluble carbodiimide), lectin and biotinylated lectin were obtained from Sigma Chemical Company. BCA protein estimation kit, Sulfo SMCC, succinimidylS-acetylthioacetate (SATA), and immunopure avidin were obtained from Pierce Chemical Company. Maghemite was obtained from Nanophase Technologies Corporation. 2.1. Preparation of the HSA-based magnetic microsphere Chemically crosslinked albumin magnetic microspheres for blood separation were synthesized with a few di!erences from the reported methods [14,15]. A 10 ml aqueous solution of 250 mg HSA and 250 mg maghemite was added dropwise to a mixture of 40 ml n-hexane, 10 ml light mineral oil and 0.5 ml of sorbitan sesquioleate to form a water in oil inverse emulsion system. After sonication for 10 min at 50% amplitude, 10 ml of 25% glutaraldehyde saturated with toluene was added to this mixture and mechanically stirred with te#on paddle stirrer at 200 rpm for 2 h. The suspension was then centrifuged at 3000 rpm for 15 min, the supernatant decanted, the microspheres washed repeatedly with petroleum ether and acetone and "nally dispersed in distilled water. Modi"cation of HSA microspheres with biotinylated lectin was done by preparation of thiolated avidin which act as a bridge between biotinylated lectin and iron oxide bound HSA. Avidin was dissolved in PBS (pH 7.4) at a concentration of 10 mg/ml. A stock solution of SATA was prepared in dimethyl sulphoxide at a concentration of 13 mg/ml. Twenty-"ve milliliter of SATA stock solution was added to each milliliter of the avidin
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
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solution and reacted for 30 min. Maghemite-bound HSA microspheres were dispersed in PBS bu!er (pH 7.4) at a concentration of 10 mg/ml, 3.3 mg of sulfo-SMCC were added to each milliliter of HSA solution and reacted for 30 min and then added to the thiolated avidin solution and reacted for 2 h at room temperature. HSA and avidin were reacted in 4:1 molar proportion to form the conjugate properly. The mixture was then centrifuged and washed repeatedly with PBS bu!er to remove unreacted avidin. Aproximately 5.15 mg (based on the amount calculated for the monolayer formation [16]) of biotinylated lectin was added to 25 ml of the avidin-modi"ed HSA microsphere suspension and reacted for 4 h at room temperature.
were prepared following a set of steps of "xing [19], washing and dehydration [20] to maintain the physiological condition necessary for retaining the shape of the RBC and thus preventing artifacts.
2.2. Modixcation of the polystyrene microspheres
2.5. Determination of the protein coupled to microspheres
PSMMS with a carboxyl functional group were further modi"ed by coating with lectin. The functional carboxyl group was converted into an ortho acyl-urea intermediate, which can bind to a protein according to di!erent coupling procedures available in the literature [17]. Typically, 1 ml of the carboxyl modi"ed microspheres (10% solid) was repeatedly washed with phosphate bu!ered saline using a magnet for the complete removal of the surfactant and redispersed in 10 ml of PBS bu!er. One hundred milligram of the water-soluble carbodiimide was added and reacted for 20 min at room temperature. The mixture was then washed twice with PBS bu!er and "nally resuspended in 5 ml of the PBS bu!er. Five milligram of lectin (approximately "ve times of the amount required for the calculated monolayer formation) was dissolved in 5 ml of PBS bu!er. The microspheres suspension was then added to the protein solution to avoid agglomeration and reacted for 4 h at room temperature. 2.3. Scanning electron microscopy A JEOL (JSM 840) instrument was used for SEM to characterize both types of microspheres after modi"cation and after coupling to RBC. Modi"ed microsphere specimens for SEM were prepared following the procedure of Margel et al. [18]. Specimens of microspheres coupled to RBC
2.4. Atomic force microscopy A D-3000 (Digital Instruments, Santa Barbara) in `tapping modea was used for the AFM. All specimens were prepared exactly in the same way as for SEM. The specimen "lms were formed on glass slides and did not need any metal coating. The RBC specimens were prepared on a cover slip using the same procedure as in SEM and were then further mounted on a glass slide for AFM.
A BCA protein assay kit was used to estimate the percentage of protein coupled to the microspheres. A standard calibration curve (50}500 mg) was made with bovine serum albumin (BSA) and the absorbance measured in an Ultraspec 2000 UV/VIS spectrophotometer (Pharmacia Biotech) at 562 nm. Carboxyl modi"ed microspheres (solid content 7.4%) and albumin microspheres (solid content 1%) were modi"ed with three di!erent concentrations of the protein. Three sets of microsphere suspensions were made using three di!erent protein concentrations. The absorbance of supernatant was measured and the protein coupled to the microspheres was obtained by di!erence.
3. Results and discussion Scanning electron micrograph for both ALBMMS and PSMMS are shown in Fig. 1(a) and (b). PSMMS are monodispersed particles having diameter 1}2 m and the albumin microspheres have shown a broad distribution of particle ranging with submicron to 1 m diameter and they appear as #akes also. Albumin microspheres were found to be always agglomerated even in extremely dilute dispersion. Micrographs obtained by SEM for proteincoated carboxyl modi"ed particles are shown in
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J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
Fig. 2. SEM micrograph of lectin coated polystyrene magnetic microspheres.
Fig. 1. Scanning electron micrograph (SEM) of (a) polystyrene magnetic microspheres and (b) albumin magnetic microspheres.
Fig. 2. No signi"cant change in shape observed after protein coating but they appeared as more agglomerated structure and they are connected with each other. Some of them showed increase in size also along with holes in it. Holes appeared in the protein-coupled polystyrene microspheres may be due to the immiscibility of the seed polymer with polymer formed in the later steps or due to the excess monomer compared to the polymer in the seed causing low viscosity and ease transport of the phase separated seed polymer as reported by Ugelstad and coworkers for the pure polystyrene particles [21]. The size of the holes basically determines the size of the seed particle, which is converted to the "nal microspheres by
more than one polymerization steps. No signi"cant change is observed in the morphology of the albumin particle before and after coating. The SEM micrographs of the RBC coupled with protein coated albumin and polystyrene microspheres are shown in Figs. 3(a), and (b). Albumin particles have coupled with RBC as a cluster form but the polystyrene particle can couple singly with the RBC. Atomic force micrographs for the polystyrene and albumin microspheres are shown in Fig. 4. The spherical surface of PSMMS is more clearly observed in the 3D view obtained in atomic force microscopy. AFM characterizes the albumin microspheres as distinctly spherical in shape even in agglomerated state. The section analysis in Fig. 4(c) shows that the albumin microspheres are in submicron range. A top view of the protein-coated polystyrene microspheres is shown in Fig. 5(a) clearly indicating the presence of structures on the surface of the spheres. A 3D #attened view of the protein-coated surface with an extremely nice contrast shows structures due to protein (Fig. 5(b)). The AFM micrographs for the protein-coated albumin are shown in Fig. 6. Protein molecules seem to form a layered structure on the clusters of the spheres. The AFM micrograph for the polystyrene coupled RBCs is shown in Fig. 7. Albumin microspheres coupled with RBCs are shown in
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
Fig. 3. SEM micrograph of (a) a RBC coupled with lectin coated PSMMS and (b) a RBC coupled with protein coated ALBMMS.
Fig. 8(a). This micrograph shows a distinct attachment of the microspheres to the RBC. Albumin microspheres were highly agglomerated and often were attached to the RBCs as a cluster of microspheres. In the phase mode of AFM distinct attachment of the albumin microspheres on the membrane of the RBC is shown in Fig. 8(b). Functional groups present in the polymeric microspheres surface, protein concentration and other factors such as pH of the medium could be a probable cause of formation of highly agglomerated structure for the protein-coated microspheres of both types. When protein forms 䉴 Fig. 4. Atomic force 3D micrograph (AFM) of (a) a polystyrene magnetic microsphere, (b) magnetic albumin microspheres, and (c) a sectional analysis of albumin microspheres.
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Fig. 6. AFM micrograph of protein-coated albumin microspheres.
Fig. 5. AFM 3D micrograph of (a) protein coated polystyrene microsphere (top view) and (b) of a protein coated surface of polystyrene microsphere (#attened view).
a covalent bond with the surface of the microspheres, they become neutral and may aggregate with each other in absence of any charges. Investigations of the nature of the interaction of this particular lectin with polymer and albumin microspheres particle are still in progress. Fig. 9 shows the variation of the optical density (OD) with concentration of the protein (in g/ml) added in the three di!erent sets for both albumin based and polystyrene based microspheres. Optical density is found to be very high showing lower amount of protein coupling to the carboxyl modi"ed polystyrene microspheres and the OD increase
Fig. 7. AFM micrograph of a RBC coupled with a polystyrene microsphere.
basically with increase in amount of the added protein. But for the albumin microspheres optical density of the decanted clear liquid is very low indicating higher amount of protein coupling and the OD values are almost same for the higher concentrations of the added protein, which means that the amount of the unbound protein is almost equal in the decanted clear liquid. This observation also shows more protein coupling to the albumin microspheres with increasing concentration of
J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29
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microspheres is not certainly enough for the 100% protein coupling and hence there is always unbound protein. Lack of site on the microspheres is de"nitely caused by the agglomeration of the spheres amongst themselves during the washing and other processing steps. As all other parameters such as temperature, pH of the medium, mixing time and speed are indentical in the process of protein coupling, the available site on the microspheres cannot change for the three di!erent samples with same solid content. We have to also eliminate the possibility of protein decoupling as the binding takes place by covalent bonding. So, the probable reason for higher protein coupling when the concentration of the added protein is increased could be a protein multilayer formation [22] on the available site. Chances of cluster formation are higher in higher concentration of protein and if these clusters couple with the microspheres as observed in Fig. 6 rather than small number of molecules, there will be less amount unbound protein. But this conclusion needs to be veri"ed by studying the morphology of the protein in di!erent concentrations. Albumin microspheres are certainly showing more coupling than the carboxyl modi"ed polystyrene microspheres. To establish the coupling e$ciency more quantitatively for the above microspheres more data is required and the experiment is in progress.
Fig. 8. AFM micrograph of (a) a RBC coupled with albumin microspheres and (b) a RBC coupled with albumin microspheres using phase mode.
added protein. Amount of the protein added is usually the calculated amount required for the monolayer formation [16] and this concentration is represented as the "rst concentration and other two concentrations are double and three times to this for both cases. Fig. 10 shows the percentage of the protein coupled for a certain amount of added protein to both kinds of microspheres. In both cases, percentage of protein coupled increases with increase in amount of the added protein. Available site on the
4. Conclusion Both carboxyl modi"ed polystyrene microspheres and albumin microspheres have shown effective coupling to RBCs when they are modi"ed with lectin and biotinylated lectin. Our study shows that albumin microspheres have some advantage over the synthetic polymer-based microspheres as RBCs separated by albumin microspheres can be reinjected into a patient. Characterization by AFM is able to show the details of the protein coupling to the microspheres and the coupling of the red blood cell to these microspheres. The presence of submicron sized particles in both SEM and AFM micrographs point at the possibility of forming nanosized albumin particles using modi"ed synthesis conditions. The amount of the protein
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Fig. 9. Plot of OD for unbound protein.
References [1] [2] [3] [4] [5] [6] [7] Fig. 10. Plot of absorbed protein as a function of amount of added protein.
[8] [9] [10]
coupled could not be optimized yet as the coupling procedure showed kinetics that might be optimized with more data for other di!erent concentrations.
[11]
[12]
Acknowledgements Part of this work is funded by Therakos Incorporation, USA.
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J. Chatterjee et al. / Journal of Magnetism and Magnetic Materials 225 (2001) 21}29 [16] H.J. Hager, Latex Polymer Reagents for Diagnostic Tests, US Patent 3,857,931, 1974. [17] L.B. Bangs, Uniform Latex Particles, Technical Notes Seradyn, 1999. [18] M. Shahar, H. Meshulam, S. Margel, J. Polym. Sci., Polym. Chem. Edn. 24 (1986) 203. [19] D. Platt, Blood Cells, Rheology and Aging, Springer, Berlin, 1998, p. 32.
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[20] J.J. Bozzola, L.D. Russell, Electron Microscopy, 2nd Edition, Jones and Bartlett, Sudbury, MA, 1992, p. 54. [21] W.S. Prestvik, A. Berge, P.C. Mork et al., in: U. Hafeli, W. Schutt, J. Teller, M. Zborowski (Eds.), Scienti"c and Clinical Applications of Magnetic Carriers, Plenum Press, New York, 1997, p. 16. [22] D. Kowalczyk, J.P. Marsault, S. Slomakowski, Colloid Polym. Sci. 274 (1996) 513.