Superparamagnetic core–shell structured microspheres carrying carboxyl groups as adsorbents for purification of genomic DNA

Colloids and Surfaces A: Physicochem. Eng. Aspects 401 (2012) 74–80

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Superparamagnetic core–shell structured microspheres carrying carboxyl groups as adsorbents for purification of genomic DNA Haihui Jiang a , Xiaoyun Han a , Zhili Li a , Xincheng Chen a , Yunhua Hou b , Ligang Gai a,∗ , Decai Li a , Xinrui Lu a , Tiling Fu a a Key Laboratory of Fine Chemicals in Universities of Shandong, School of Chemistry and Pharmaceutical Engineering, Shandong Polytechnic University, Jinan 250353, People’s Republic of China b Shandong Provincial Key Laboratory of Microbial Engineering, School of Food and Bioengineering, Shandong Polytechnic University, Jinan 250353, People’s Republic of China

a r t i c l e

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Article history: Received 13 December 2011 Received in revised form 29 February 2012 Accepted 13 March 2012 Available online 21 March 2012 Keywords: Carboxyl-functionalized Fe3 O4 /PS-PAA microspheres Superparamagnetism Aspergillus niger Genomic DNA

a b s t r a c t Magnetic composite microspheres present a promising candidate for applications in bioseparation, drug delivery, and biocatalysis. We report here the synthesis of carboxyl-functionalized and core–shell structured Fe3 O4 /PS-PAA microspheres via a dispersion interfacial polymerization method, using styrene (St) and acrylic acid (AA) as monomers. The magnetic composite microspheres exhibit good dispersity, superparamagnetism, and high saturation magnetization. The samples were characterized with XRD, XPS, FTIR, FESEM, TEM, and VSM techniques, and further tested as adsorbents for purification of genomic DNA from Aspergillus niger NA1003 cells. Complexation mechanism of DNA with the adsorbent is discussed. The magnetic separation yields high-quality genomic DNA and satisfying productivity comparable to those isolated by the conventional phenol–chloroform extraction. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Magnetic composite nano- and/or micro-particles have been subjected to extensive research due to their potential applications in the fields of biotechnology and biomedicine, such as DNA extraction [1–3], enzyme immobilization [4,5], protein separation [6,7], drug delivery [8,9], and catalysis [10,11]. Magnetic separation techniques, which employ magnetic composite particles as both a solid adsorbent and a magnetic carrier, can reduce chemical inputs and enable ease of manipulation and automation as compared to conventional nonmagnetic separation methods which require centrifugation, precipitation, filtration, and chromatographic separations [12,13]. Among the magnetic composites, carboxyl functionalized magnetic particles are of great interest because of their biocompatibility and facile surface modification. Carboxyl groups on the surface of magnetic composites can be easily activated for covalent coupling with various ligands suitable for specific interaction with biological molecules [14,15]. It has been demonstrated that carboxyl-carrying magnetic particles have wide applications in enzyme immobilization and separation [14–17], DNA purification [18–20], and drug delivery [21,22].

∗ Corresponding author. Tel.: +86 531 89631208; fax: +86 531 89631207. E-mail address: [email protected] (L. Gai). 0927-7757/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfa.2012.03.024

It is known that duplex DNA carries two univalent negative charges per base pair at most of the solution pH values [23]. Although carboxyl-modified magnetic particles are also negatively charged in basic and near neutral solutions, complexation of DNA with negatively charged sorbents can take place in the presence of multivalent counterions due to a counterion-mediated Coulomb coupling mechanism [24–26]. Also, a polymer-andsalt-induced (psi) condensation mechanism is well-accepted for the like-charge sorbent/DNA complexation [18–20,27]. While the large specific surface area belonging to magnetic nanosorbents is beneficial for sorption, theoretical calculations [28,29] predicted that the increase in radius of the sorbent sphere facilitates its complexation with semiflexible polyelectrolyte, which is analogous to DNA with size larger than several tens of kb, due to the decrease in entropy penalty [28] and in bending energy cost [29]. This suggests that magnetic composite microspheres are superior to their nano-counterparts in particular for in vitro bioseparations [3]. In this paper, we report the synthesis of polystyrene (PS)–polyacrylic acid (PAA)–coated magnetite (Fe3 O4 ) microspheres via a dispersion interfacial polymerization method. The composite microspheres show good dispersity, superparamagnetism, and high saturation magnetization. The magnetic separation of genomic DNA from Aspergillus niger NA1003 cells using the composite microspheres yields satisfying results as compared to the conventional phenol–chloroform extraction.

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2. Materials and methods 2.1. Materials All chemicals for use in preparation of Fe3 O4 and composite microspheres were of analytical grade, including ferric chloride hexahydrate (FeCl3 ·6H2 O), sodium acetate trihydrate (NaAc·3H2 O), glycol (HOCH2 CH2 OH), polyethylene glycol (PEG, Mw = 6000), phenylethylene (C6 H5 CH CH2 ), polyvinyl pyrrolidone (PVP, Mw = 3000), benzoyl peroxide (BPO), acrylic acid (AA). A. niger NA1003 was derived from the A. niger S4309-2 (CICC 41236), which was obtained from the China Center of Industrial Culture Collection (Beijing, China). Agarose of molecular biology grade was purchased from Gene Tech Company (Shanghai, China). Proteinase K, ribonuclease (RNase A), sodium chloride (NaCl), tri(hydroxymethyl)aminomethane (Tris), Tris–HCl, PEG6000, ethylene diamine tetraacetic acid (EDTA), acetic acid (HAc), isopropanol, and sodium dodecyl sulfate (SDS) for use in DNA isolation and analysis were of analytical grade. 2.2. Synthesis of Fe3 O4 microspheres A glycol reduction method was employed for the synthesis [30]. Briefly, 5 mmol FeCl3 ·6H2 O was dissolved in 40 mL of glycol to form a clear solution, followed by addition of 40 mmol NaAc·3H2 O and 1.0 g of PEG-6000. The mixture was stirred vigorously for 30 min and then placed into a Teflon-lined autoclave (50 mL, capacity). The autoclave was heated at 200 ◦ C for 12 h, and then allowed to cool to room temperature. The black precipitate was collected with a permanent magnet, and washed with ethanol and distilled water several times. Finally, the Fe3 O4 microspheres were dried in a vacuum oven at 40 ◦ C for 10 h. 2.3. Synthesis of carboxyl-functionalized microspheres In a typical procedure, 0.23 g of Fe3 O4 microspheres and 40 mL of anhydrous ethanol were transferred to a four-neck flask, the mixture was treated with ultrasonication for 5 min, followed by addition of 0.25 g of PVP, 0.025 g of BPO, and 0.5 mL of redistilled AA. The mixture was treated with ultrasonication and nitrogen protection, and then mechanically stirred at 30 ◦ C for 1 h. Subsequently, 2 mL of redistilled phenylethylene was added into the above mixture. The reaction was performed under continuous stirring and intermittently ultrasonic treatment at 78 ◦ C for 4 h, and allowed to cool to room temperature. The resulting product was magnetically separated and washed with ethanol and water several times. Finally, the composite microspheres were dried in a vacuum oven at 40 ◦ C for 12 h, and referred to as Fe3 O4 /PS-PAA microspheres. 2.4. Characterizations X-ray diffraction (XRD) patterns were recorded on a Bruker D8 Advance diffractometer (Bruker AXS Co., Germany), using Cu ˚ with a step size of 0.3◦ 2 s−1 , operatK˛ radiation ( = 1.5406 A) ing at 40 kV and 40 mA. X-ray photoelectron spectroscopy (XPS) analyses were performed on an ESCALAB 250 X-ray photoelectron spectrometer (Thermo Fisher Scientific Co., America), using monochromatic Al K˛ radiation. Pass energy of 20 eV and step size of 0.05 eV were employed for Fe 2p high-resolution spectra. For O 1s and C 1s analyses, pass energy of 50 eV and step size of 0.05 eV were adopted. Transmission electron microscopy (TEM) images and selected area electron diffraction (SAED) pattern were obtained on a Hitachi H-800 transmission electron microscope (Hitachi Ltd., Japan) operating at 150 kV. Field-emission scanning electron microscopy (FESEM) images were taken on a JSM-6700 scanning electron microscope (JEOL Ltd., Japan). For the TEM and FESEM

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analyses, the samples were ultrasonically dispersed in anhydrous ethanol solution to form uniform suspensions, and the suspensions were dripped onto carbon film coated copper grids and glass slides, respectively. Fourier transform infrared (FTIR) spectra were collected on a Nicolet Avatar 370 infrared spectrometer (Nicolet Co., America) using pressed KBr discs. The FTIR spectra were collected in the range of 4000–400 cm−1 with a resolution of 1 cm−1 . Magnetization measurements were performed on a Lake Shore 7410 vibrating sample magnetometer (VSM, Lake Shore Co., America) at room temperature. 2.5. Extraction of genomic DNA from A. niger cells with Fe3 O4 /PS-PAA microspheres Magnetically driven genomic DNA extraction from A. niger NA1003 cells was performed by a modification to the nonmagnetic separation process previously described [31]. Before DNA extraction, the spores of A. niger with a concentration of 108 spores/mL were inoculated in a minimal media (MM) at 28 ◦ C for 3–4 d. The mycelia were collected by vacuum filtration, washed with sterile water, and freeze-dried. Subsequently, the mycelia were ground in liquid nitrogen and transferred to a centrifuge tube (50 mL, capacity). 25 mL of fungi lysis buffer (pH 5.0), containing 5 mM Tris–HCl (pH 8.5), 250 mM NaCl, 25 mM EDTA, 2% SDS, and 1 mg of proteinase K, were added into the tube. The tube was vortexed for 10 s, incubated in a 65 ◦ C heating block for 1 h, and ice-cooled for 5 min, followed by centrifugation at 10,000 rpm for 20 min. After that, 24 mL of the supernatant was transferred to a new tube, followed by the addition of 14.4 mL of isopropanol. The tube was placed at −20 ◦ C for 20 min, and then centrifuged at 10,000 rpm for 10 min. 0.01 g of Fe3 O4 /PS-PAA microspheres were placed into a microcentrifuge tube (1.5 mL, capacity), and ultrasonically washed with 70% (v/v) ethanol and then sterile water twice, followed by dispersion in a mixture containing 40 ␮L of nuclear acid extract, 4 ␮L pretreated ribonuclease (RNase A, 10 mg/mL), and 1.0 mL of binding buffer (20% PEG-6000, 2 M NaCl). The suspension was gently agitated at room temperature for 10 min. The magnetic particles were immobilized by using a permanent magnet with a surface magnetization of 4000 G, and the supernatant was removed. The magnetic particles were washed twice with 70% ethanol. After removal of the supernatant, the adsorbed DNA was eluted from the magnetic particles by adding 0.2 mL of TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 7.8), followed by incubation with gentle agitation at room temperature for 10 min. The magnetic particles were immobilized again, and the eluate was collected and analyzed by ultraviolet (UV) spectroscopy (UV-7504c, Shanghai, China). A 10-␮L aliquot of the eluted DNA was analyzed by agarose gel electrophoresis on a 1.0% agarose gel in TAE buffer (40 mM Tris, 20 mM HAc, 1 mM EDTA, pH 8.0). The agarose gel was stained with 0.5 ␮g/mL of ethidium bromide (EB). Electrophoresis was performed on a horizontal gel electrophoresis unit (Liuyi Instrument Factory, Beijing, China), operating at 100 V for 50 min. 3. Results and discussion 3.1. Structure, composition, and morphology Fig. 1 shows the XRD patterns of the samples. In comparison with those of bare magnetic particles (Fig. 1(a)), the diffraction peaks of magnetic composites (Fig. 1(b)) decrease in intensity probably due to polymer coating; however, there is no change in peak position between the two samples, indicating that crystal phase of the magnetic core is well-maintained after the coating process. All the peaks marked in Fig. 1 can be indexed to (2 2 0), (3 1 1), (4 0 0), (4 2 2), (5 1 1), (4 4 0), and (5 3 3) planes of cubic Fe3 O4 (JCPDS

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Fig. 1. XRD patterns of the samples: (a) Fe3 O4 ; (b) Fe3 O4 /PS-PAA. Fig. 3. FTIR spectra of bare Fe3 O4 (a) and Fe3 O4 /PS-PAA (b).

88-0866) with the lattice parameter of a = 8.383 A˚ in accordance ˚ with the reference value of a = 8.384 A. Due to similarity in the XRD patterns of Fe3 O4 and ␥-Fe2 O3 [32], XPS spectra were therefore collected to verify compositions of the samples (Fig. 2). The survey spectrum of magnetite sample shows C, O, and Fe elements (Fig. 2(A), spectrum (a)), whereas the survey spectrum of composite sample exhibits only C and O elements (Fig. 2(A), spectrum (b)). The non-existence of Fe element in spectrum (b) is due to the shielding of PS-PAA shells which are so thick (∼20 nm, discussion below) as to overpass the detecting limit (<5 nm) of employed photoelectron spectrometer. The Fe 2p core level spectrum of bare magnetic particles (Fig. 2(B)) reveals

two broad peaks centered at 723.3 and 710.3 eV which can be respectively assigned to Fe 2p1/2 and Fe 2p3/2 of Fe3 O4 [32–35]. The broadness of Fe 2p peaks is attributed to the existence of dual iron oxidation states (Fe2+ and Fe3+ ) which have different but irresolvable binding energy [35]. It is worthy of note that there is no charge transfer satellite with a binding energy of ∼719 eV characteristic of Fe3+ in ␥-Fe2 O3 [32], which confirms a Fe3 O4 sample. The O 1s region (Fig. 2(C)) of Fe3 O4 sample shows a broad peak centered at 529.5 eV consistent with the reported value for Fe3 O4 [36]. The broad peak centered at 531.9 eV arises from the O elements in the form of carboxyl groups on PS-PAA shells [37,38]. The C 1s core level spectrum of composite sample can be fitted with

Fig. 2. XPS spectra of the samples: (A) survey spectra ((a) Fe3 O4 and (b) Fe3 O4 /PS-PAA); (B) Fe 2p region; (C) O 1s regions; (D) C 1s region.

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Fig. 4. FESEM images of Fe3 O4 (a) and Fe3 O4 /PS-PAA (b); TEM images of Fe3 O4 (c) and Fe3 O4 /PS-PAA (d); SAED pattern of Fe3 O4 /PS-PAA (e).

three components centered at 284.6, 286.4, and 288.6 eV. The former one is composed of the aromatic carbons [37,38], the carbons in methylene chains ( CH2 CH2 ) [39], and the carbon normally used as energy reference in XPS measurement. The latter two components can be assigned to the carboxylic carbons [37–39]. In the case of composite sample, the non-existence of Fe element in its survey spectrum and the changes in binding energy with respect to the O 1s and C 1s core levels verify that Fe3 O4 particles are completely coated by the PS-PAA polymer, leading the composite to a core–shell structure rather than a blending form. FTIR spectra were also recorded to further examine surface chemistry of the samples (Fig. 3). The strong peak at 580 cm−1 corresponds to the characteristic band of Fe O vibration [3,14]. For the sample of Fe3 O4 /PS-PAA microspheres (Fig. 3(b)), apart from the peaks (2952, 2893, 1640, 1068, 896, and 1460–1400 cm−1 ) corresponding to the benzene ring and methylene which result from the polymer shell, the broad band around 3420 cm−1 can be assigned

to the O H stretching vibrations of carboxyl groups and adsorbed water. More importantly, a relatively weak band around 1722 cm−1 and the small bands around 1270 cm−1 , which can be separately attributed to the C O and C O stretching vibration of carboxyl groups [14,40], consolidate the carboxyl-functionalized polymer shells. The weakness in intensity of the bands corresponding to carboxyl groups might be due to the disturbance of benzene rings to carboxyl groups. Fig. 4 shows the FESEM and the TEM images of Fe3 O4 and Fe3 O4 /PS-PAA samples. Nearly monodisperse microspheres with average diameters of about 220 and 260 nm are observed for the bare Fe3 O4 (Fig. 4(a) and (c)) and the composite sample (Fig. 4(b) and (d)), respectively, indicating that the average thickness of polymer shells is about 20 nm. As previously reported [31,41], every Fe3 O4 microsphere is an assembly of single domain magnetite nanoparticles with a mean size less than 30 nm. Therefore, solid and small amount of hollow microspheres coexist in the bare Fe3 O4

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Fig. 5. Room-temperature magnetization curves of Fe3 O4 (a) and Fe3 O4 /PS-PAA (b).

sample. After the interfacial polymerization process, hollow Fe3 O4 microspheres can be fully filled, leading the bare magnetite particles to form core–shell structured spheres with smooth surface (Fig. 4(d)). The SAED pattern of composite sample (Fig. 4(e)) reveals that the magnetic cores are polycrystalline magnetite crystals in accordance with the XRD result. 3.2. Magnetic properties Fig. 5 shows the room-temperature magnetization curves of the samples. The magnetic saturation (Ms ) value of Fe3 O4 /PSPAA sample is 85.4 emu/g, which is lower by 3.9 emu/g than that of pure Fe3 O4 sample. Meanwhile, the remanent magnetization (Mr ) decreases from 4.74 to 1.93 emu/g after polymer coating. The observed decrease in both Ms and Mr reflects the standard practice of normalizing the magnetization by sample mass, where the more nonmagnetic component per gram of a sample the more weakening in its magnetization readings [42]. In spite of the spheres in micrometer scale, the coercivity (Hc ) values of both bare Fe3 O4 and composite samples approximate to the theoretical value for superparamagnetism (Hc ≤ 50 Oe) [43]. The observed superparamagnetism is due to that the Fe3 O4 microspheres are composed of magnetite nanospheres with a mean size smaller than 30 nm [31,41]. The magnetic properties of the samples may be determined by dipole field energy, thermal and magnetic anisotropy energies, and interparticle interactions [44]. It is known that superparamagnetism prevents magnetic particles from self-aggregation and enables them to be redispersed rapidly after removal of the magnetic field [45,46]. The magnetic composite microspheres having good dispersity, core–shell structure, smooth surface-carrying carboxyl groups, and superparamagnetism with a high Ms value will find greatly potential applications in biomedical and bioengineering fields. 3.3. Purification of genomic DNA from A. niger cells with Fe3 O4 /PS-PAA microspheres As mentioned above, the complexation of negatively charged semiflexible genomic DNA with carboxyl-modified microspheres is apt to take place in a polymer-and-salt environment [18–20,27]. In our experiments, the binding buffer containing PEG-6000 and NaCl enables the psi-condensation process [27] for DNA adsorption. In the PEG-and-NaCl environment, the genomic DNA existing in a supercoiled structure can be folded into a more stable compact state due to the crowding effect [27]. For compact DNA in PEG solution, the negative charge on the folded parts is expected to

Fig. 6. 1.0% agarose gel electrophoresis of genomic DNA isolated from A. niger NA1003 cells: (M) DNA marker (␭-DNA with a length of 32 kb without enzyme digestion); (lane 1) DNA isolated by phenol–chloroform extraction; (lane 2) DNA isolated with Fe3 O4 /PS-PAA microspheres.

be almost completely neutralized, leaving ∼10% of the negative charge retained on the unfolded random-coiled parts [47]. This leads to a weak electrostatic repulsion between DNA molecules and PS-PAA polymer under the high ionic strength condition (2 M NaCl) [23]. Thus, the adsorption of DNA on the carboxyl-modified surface is mainly controlled by dehydration and hydrogen bonds at the polymer–DNA interface [23,48]. The complexation of DNA with carboxyl-modified microspheres occurs when the adsorption free energy Fad exceeds the energy barrier Ea [25,28], where Fad derives from dehydration effects, osmotic pressure due to PEG, and hydrogen bonds, and Ea associates with elastic bending energy, thermal pressure due to monovalent counterions, electrostatic repulsion, and entropy cost in DNA confinement. In the eluting buffer, the adsorbed DNA molecules tend to release their elastic bending energy. Meanwhile, the electrostatic repulsion at the interface between DNA and polymer is resumed due to the low salt concentration [48,49]. As a result, the DNA molecules are desorbed and unraveled to their supercoiled states. The addition of EDTA is to chelate the counterions binding to DNA and further to promote the DNA desorption [48]. Considering that the pristine magnetite has a strong ability to adsorb but a poor ability to release DNA molecules [48], the Fe3 O4 /PS-PAA microspheres were tested for isolation of genomic DNA from A. niger NA1003 cells. The purified DNA samples were analyzed by agarose gel electrophoresis and quantified by UV spectrometry. Fig. 6 shows the EB-stained gel image of isolated DNA. The band in lane 1 is the DNA isolated by the conventional phenol–chloroform method, and the band in lane 2 is the DNA isolated with Fe3 O4 /PS-PAA microspheres. The length of the DNA isolated by magnetic separation is a little shorter than that of ␭-DNA with a length of 32 kb without enzyme digestion. About 10.7 ␮g of genomic DNA can be isolated from 40 ␮L of the crude extracts by using 10 mg of Fe3 O4 /PS-PAA. The A260 /A280 ratios of isolated DNA corresponding to lane 1 and lane 2 are 1.72 and 1.79, respectively,

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close to 1.8 indicative of high purity, which is consistent with the electrophoresis analysis. The high quality of genomic DNA isolated with the Fe3 O4 /PS-PAA microspheres ensures further applications of the DNA in molecular biology. 4. Conclusions In summary, carboxyl-functionalized and core–shell structured Fe3 O4 /PS-PAA microspheres with a high Ms value have been prepared via a dispersion interfacial polymerization method. The magnetic composite microspheres are superparamagnetic and nearly monodisperse. The purification of genomic DNA from A. niger NA1003 cells by using the composite microspheres shows comparable to that isolated by the conventional phenol–chloroform extraction. The magnetic composite microspheres present a suitable candidate for applications in biomedical and bioengineering fields. Acknowledgments This research was supported by the Foundation for Outstanding Young Scientists (BS2009CL049), the Postdoctoral Innovation Program Foundation (200802016), and the S&T Plan Projects of Shandong Provincial Education Department (J10LC08) of Shandong Province of China. References [1] J.W. Liu, Y. Zhang, D. Chen, T. Yang, Z.P. Chen, S.Y. Pan, N. Gu, Facile synthesis of high-magnetization ␥-Fe2 O3 /alginate/silica microspheres for isolation of plasma DNA, Colloids Surf. A 341 (2009) 33–39. [2] Z. Shan, C. Li, X. Zhang, K.D. Oakes, M.R. Servos, Q. Wu, H. Chen, X. Wang, Q. Huang, Y. Zhou, W. Yang, Temperature-dependent selective purification of plasmid DNA using magnetic nanoparticles in an RNase-free process, Anal. Biochem. 412 (2011) 117–119. [3] L. Gai, Z. Li, Y. Hou, H. Jiang, X. Han, W. Ma, Preparation of core–shell Fe3 O4 /SiO2 microspheres as adsorbents for purification of DNA, J. Phys. D: Appl. Phys. 43 (2010) 445001-1–445001-8. [4] P.A. Johnson, H.J. Park, A.J. Driscoll, Enzyme nanoparticles fabrication: magnetic nanoparticle synthesis and enzyme immobilization, Methods Mol. Biol. 679 (2011) 183–191. [5] J. Huang, R. Zhao, H. Wang, W. Zhao, L. Ding, Immobilization of glucose oxidase on Fe3 O4 /SiO2 magnetic nanoparticles, Biotechnol. Lett. 32 (2011) 817–821. [6] H. Chen, C. Deng, X. Zhang, Synthesis of Fe3 O4 @SiO2 @PMMA core–shell–shell magnetic microspheres for highly efficient enrichment of peptides and proteins for MALDI-ToF MS analysis, Angew. Chem. 122 (2010) 617–621. [7] S.H. Sophia Lee, T.A. Hatton, S.A. Khan, Microfluidic continuous magnetophoretic protein separation using nanoparticle aggregates, Microfluid Nanofluid 11 (2011) 429–438. [8] Z. Xu, Y. Feng, X. Liu, M. Guan, C. Zhao, H. Zhang, Synthesis and characterization of Fe3 O4 /SiO2 @poly-l-alanine peptide brush-magnetic microspheres through NCA chemistry for drug delivery and enrichment of BSA, Colloids Surf. B 81 (2010) 503–507. [9] B. Chertok, A.E. David, V.C. Yang, Polyethyleneimine-modified iron oxide nanoparticles for brain tumor drug delivery using magnetic targeting and intracarotid administration, Biomaterials 31 (2010) 6317–6324. [10] L. Kong, X. Lu, E. Jin, S. Jiang, X. Bian, W. Zhang, C. Wang, Constructing magnetic polyaniline/metal hybrid nanostructures using polyaniline/Fe3 O4 composite hollow spheres as supports, J. Solid State Chem. 182 (2009) 2081–2087. [11] J. Garcia, Y. Zhang, H. Taylor, O. Cespedes, M.E. Webb, D. Zhou, Multilayer enzyme-coupled magnetic nanoparticles as efficient reusable biocatalysts and biosensors, Nanoscale 3 (2011) 3721–3730. [12] S. Berensmeier, Magnetic particles for the separation and purification of nucleic acids, Appl. Microbiol. Biotechnol. 73 (2006) 495–504. [13] T.L. Hawkins, K.J. McKernan, L.B. Jacotot, J.B. MacKenzie, P.M. Richardson, E.S. Lander, A magnetic attraction to high-throughput genomics, Science 276 (1997) 1887–1889. [14] J. Hong, D. Xu, P. Gong, H. Ma, L. Dong, S. Yao, Conjugation of enzyme on superparamagnetic nanogels covered with carboxyl groups, J. Chromatogr. B 850 (2007) 499–506. [15] J. Hong, J. Huang, S. Liu, J. Yu, S. Luo, Stability and activity of chymotrypsin immobilized on magnetic nanogels covered with carboxyl groups, J. Appl. Polym. Sci. 111 (2009) 2844–2850. [16] D. Shao, K. Xu, X. Song, J. Hu, W. Yang, C. Wang, Effective adsorption and separation of lysozyme with PAA-modified Fe3 O4 @silica core/shell microspheres, J. Colloid Interface Sci. 336 (2009) 526–532.

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