Analytical Biochemistry 446 (2014) 87–89
Contents lists available at ScienceDirect
Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio
Notes & Tips
Preparation of stabilized magnetic nanoparticles with polymerizable lipid and anchor compounds Boram Kang, Suk-Jung Choi ⇑ Department of Chemistry, Gangneung-Wonju National University, Gangneung 210-702, Republic of Korea
a r t i c l e
i n f o
Article history: Received 27 June 2013 Received in revised form 28 September 2013 Accepted 30 October 2013 Available online 8 November 2013 Keywords: Magnetic nanoparticle Surface modification Polymerizable lipid Anchor compound
a b s t r a c t Although the lipid-based method for coating of magnetic nanoparticles (MNPs) is rapid and simple, the unstable state of the lipid layer is a major limitation for the practical application of this method. We devised a method to prepare stabilized MNPs by covalent modifications such as lipid polymerization and anchoring of the lipid layer. The stability of the modified lipid layer was demonstrated by the stable status of enzymes immobilized on the MNPs and the resistance of the MNPs to aggregation. We also determined the maximum ratio of nonpolymerizable lipophilic compounds that can be included in the layer without significantly reducing stability. Ó 2013 Elsevier Inc. All rights reserved.
Magnetic nanoparticles (MNPs) have a wide range of applications in immunoassays, separating molecules or cells, delivering therapeutic molecules, and magnetic resonance imaging (MRI) [1–4]. Prior to use in these applications, the MNP surface must be modified to ensure the stability and solubility of the particles in an aqueous environment. Another purpose of surface modification is to provide MNPs with reactive functional groups that might be used to immobilize biological molecules such as antibodies or oligonucleotides. Lipids have the potential to be used for coating of solid surfaces because of their ability to form bilayer or monolayer structures on surfaces depending on the surface’s hydrophilic or hydrophobic nature [5,6]. The lipid coating method has been applied in the preparation of MRI contrast agents or in vivo imaging agents [7– 9]. It was shown that coating with PEG–lipids (lipids containing a polyethylene glycol head group) enhanced solubility and stability of MNPs while lowering the toxicity of these MNPs to cells [7,9]. The surface was modulated to have a positive charge or fluorescence with the addition of a cationic lipid or a fluorescent lipid [8,9]. The process of lipid coating is very fast and simple in comparison to other methods. Moreover, a required functional group can be easily introduced onto the surface using lipids or lipophilic compounds containing the functional group. Nevertheless, limited stability is a major practical limitation of the lipid coating method, in which the constituent lipid molecules associate via noncovalent interactions. ⇑ Corresponding author. Fax: +82 33 640 2264. E-mail address: [email protected] (S.-J. Choi). 0003-2697/$ - see front matter Ó 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ab.2013.10.036
To prepare stabilized functional MNPs, we devised a new coating method using the polymerizable lipid 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DYPC) and the anchor compound octadecyltrimethoxysilane (ODTS). A lipid film containing 0.25 mg DYPC, 0.011 mg ODTS, and 0.051 mg 1,2-dipalmitoylsn-glycero-3-phosphoethanolamine-N-(biotinyl) (BTPE) dissolved in 0.5 ml chloroform (composition D) was prepared in a glass vial. DYPC is capable of forming lipid networks via polymerization induced by UV irradiation [10]. ODTS can be covalently linked to the MNP surface while inserted in a lipid layer via its hydrophobic tail. BTPE was added to present biotin groups that were used to immobilize streptavidin-linked enzymes. Control experiments were conducted with three lipid films of different compositions: 0.25 mg 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 0.064 mg BTPE (composition A), 0.25 mg DPPC, 0.013 mg ODTS, and 0.064 mg BTPE (composition B), and 0.25 mg DYPC and 0.051 mg BTPE (composition C). For convenience and consistency of the experiment, many films were prepared simultaneously and stored in the freezer until use. As the mole fraction of BTPE was 0.154–0.167 in all compositions, the surface density of the biotin group was sufficient for a saturating level of streptavidin binding, considering the size of the lipid and streptavidin [11,12]. Coating schemes for the four lipid compositions are depicted in Fig. 1a. Iron oxide (Fe3O4) MNPs (100-nm diameter) were suspended in PBS (phosphate-buffered saline) at a concentration of 0.5 mg/ml and then 2 ml of the solution was added to a vial. The vial was filled with nitrogen gas and placed in a bath sonicator (Model SD-D300H, S-D Ultrasonic Cleaner, Seoul, Korea) for 3 h. The vial was sonicated with 80% output power, while temperature was
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Notes & Tips / Anal. Biochem. 446 (2014) 87–89
Fig.1. Lipid coating schemes and enzyme immobilization efficiency. (a) Schematic representations of coating with the four lipid compositions: (1) composition A (DPPC + BTPE), (2) composition B (DPPC + ODTS + BTPE), (3) composition C (DYPC + BTPE), and (4) composition D (DYPC + ODTS + BTPE). (b) Activity of enzymes immobilized on the four kinds of lipid-coated MNPs. Alkaline phosphatase (AP) and horseradish peroxidase (HRP) activity are represented as optical density at 405 nm (light gray) and light intensity (dark gray). Each data point represents the mean of triplicate measures, and the error bars represent the standard deviation.
maintained at 50–60 °C during the process to ensure the lipids remained in a liquid state. For polymerization of DYPC, vials were irradiated for 30 min at 254 nm with a UV lamp. During the polymerization reaction, the MNP was agitated under a nitrogen gas atmosphere in the bath sonicator. The lipid-coated MNPs were biofunctionalized with streptavidin–alkaline phosphatase (SA–AP) or streptavidin–horseradish peroxidase (SA–HRP) to estimate protein immobilization efficiency and lipid layer stability. SA–AP or SA–HRP was added to the lipidcoated MNPs (20 lg protein per 1 mg MNP) and incubated at room temperature for 30 min. After being washed three times with PBS, the biofunctionalized MNPs were suspended in PBS at a concentration of 10 lg/ml and 0.1 ml of the modified MNPs was used for the estimation of enzyme activity. AP activity was estimated by the colorimetric assay method using pNPP solution (10 mM p-nitrophenyl phosphate, 0.5 mM MgCl2, 1 M diethanolamine, pH 9.8). The pNPP solution (0.1 ml) was added to a 96-well microplate containing MNPs and the microplate was incubated at room temperature for 5 min. Absorbance was read at 405 nm with a Synergy-HT plate reader (BioTek, Winooski, VT, USA). HRP activity was estimated by the chemiluminescence assay method in a white 96-well microplate. The microplate containing 0.1 ml of the biofunctionalized MNPs was placed in a LuBi plate reader (Seoulin Bioscience, Seongnam, Korea). Light intensity was recorded over time after injecting 80 ll of luminol solution (2.8 mM luminol, 3.6 mM 4-iodophenol, 0.002% Tween 20, 0.1 M Tris–HCl, pH 8.3) and 80 ll of 3 mM H2O2 solution into each well. As shown in Fig. 1b, activity of the AP enzyme had a tendency to increase as covalent modifications were added to the lipid layers. The enzyme activity was lowest in composition A, in which the DPPC lipid layer was associated using only noncovalent interactions. Positive effects of lipid-layer anchoring and lipid cross-linking were revealed in compositions B and C, respectively. Combined use of DYPC and ODTS resulted in the highest activity, demonstrating a synergistic effect of polymerization and anchoring (composition D). Without UV treatment of compositions C and D, AP enzyme activity was about 85% of the activity of the UV-treated samples (Supplementary Fig. S1). This result confirms that the activity increase in the DYPC-coated MNPs was dependent on the
cross-linking of the lipids. Furthermore, the effects of the covalent modifications on the protein immobilization efficiency were reproducibly observed in the experiments with SA–HRP. One possible explanation for this result is that the covalent modifications effectively reduced the removal of enzymes by stabilizing the lipid layer. The DPPC coating is maintained only by noncovalent interactions and is susceptible to environmental stresses, causing disintegration of the layer and removal of the enzymes. In composition D, however, the enzymes were preserved because the lipid layer was stabilized by two covalent modifications: crosslinking of lipid molecules within the layer and covalent linkage of the layer to the MNP surface. The stabilizing effect of the covalent modifications was also shown by the resistance of these MNPs to detergent treatment. Only 8% of the enzyme activity remained after detergent treatment in the DPPC-coated MNPs, while 40% of the enzyme activity was still associated with the MNPs in composition D (Supplementary Fig. S2). For functionalization of MNPs, functional lipophilic compounds such as BTPE should be included in the lipid layer. However, most of the commercially available functional lipophilic compounds are nonpolymerizable and may have a negative effect on the stability of the lipid layer by preventing the cross-linking of DYPC. Therefore, we estimated the effect of the DYPC ratio on the activity of immobilized enzymes. Lipid-coated MNPs were prepared with lipid films containing DYPC, DPPC, ODTS, and BTPE. The amounts of ODTS and BTPE were kept constant at 10 lg (2.7 lmol) and 2.56 lg (0.3 lmol) in all films. However, the amounts of DYPC and DPPC were varied while the total moles of the two lipids were kept constant at 27.3 lmol. SA–HRP or SA–AP was immobilized on the MNPs and their activity was determined as described previously. The enzyme activity decreased with a decreasing mole fraction of DYPC, but this decrease was discontinuous at DYPC mole fractions lower than 0.72 (Supplementary Fig. 2a). Although the reason for this discontinuity is not clear, the result indicates that the mole fraction of DYPC should be at least 0.72 to prepare a stable lipid coating. In other words, nonpolymerizable functional lipophilic compounds can be included up to a mole fraction of 0.19 without significantly reducing stability, based on the mole fraction of ODTS here (0.09). In the case of composition D, the
Notes & Tips / Anal. Biochem. 446 (2014) 87–89
DYPC mole fraction was 0.76, high enough to form a stable lipid layer. MNPs have hydrophobic surfaces with a large surface area to volume ratio and have a tendency to form aggregates, which is a critical obstacle to biomedical applications [1]. Lipid coating is expected to provide MNPs with resistance to aggregation, as there are several reports describing improved resistance to aggregation in zwitterion-coated nanoparticles [13–15]. Therefore, the aggregation of the biofunctionalized MNPs was estimated by the filtration method. MNPs were coated with composition A or D and biofunctionalized with SA–HRP as described previously. The MNPs were filtered through 0.4-lm Isopore filters (Millipore, Billerica, MA, USA) to remove aggregates and were stored at 4 °C. The MNPs were filtered again each day and HRP activity remaining in the filter was estimated. The degree of aggregation is proportional to the HRP activity associated with the filter because the aggregated MNPs are trapped in the filter. The enzyme activity associated with the filter increased rapidly within 4 days in the DPPC-coated MNP (composition A) but remained almost constant for 3 weeks in the MNP stabilized with composition D (Supplementary Fig. 2b). Aggregation of the DPPC-coated MNPs may be induced by the exposed MNP surfaces that resulted from the partial disintegration of the lipid layer as discussed above. It is also possible that the DPPC layers of different MNPs fused to form larger clusters to release the tension caused by high curvature. In this work, the lipid-based method for MNP coating was improved by covalent modifications of the lipid layer. The stability of the lipid layer that was modified with cross-linking of DYPC and anchoring with ODTS was demonstrated using three methods. First, a higher enzyme immobilization efficiency was achieved by coating with DYPC, ODTS, and BTPE compared with coating with DPPC and BTPE. Second, the immobilized enzyme on the modified layer was more stable following detergent treatment. Finally, the MNPs coated with the modified lipid layer showed higher resistance to aggregation. It was also shown that the mole fraction of DYPC should be at least 0.72 to obtain the stabilizing effect of polymerization. Acknowledgments This research is part of the project titled ‘‘Development of Biosensors for Paralytic Shellfish Toxins’’ funded by the Ministry of Oceans and Fisheries, Korea, and was also supported by the
89
Research Institute of Natural Science of Gangneung-Wonju National University. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ab.2013.10.036. References [1] A.K. Gupta, M. Gupta, Synthesis and surface engineering of iron oxide nanoparticles for biomedical applications, Biomaterials 26 (2005) 3995–4021. [2] S. Laurent, D. Forge, M. Port, A. Roch, C. Robic, L. Vander Elst, R.N. Muller, Magnetic iron oxide nanoparticles: synthesis, stabilization, vectorization, physicochemical characterizations, and biological applications, Chem. Rev. 108 (2008) 2064–2110. [3] J. Gao, H. Gu, B. Xu, Multifunctional magnetic nanoparticles: design, synthesis, and biomedical applications, Acc. Chem. Res. 42 (2009) 1097–1107. [4] S.T. Selvan, T.T. Tan, D.K. Yi, N.R. Jana, Functional and multifunctional nanoparticles for bioimaging and biosensing, Langmuir 26 (2010) 11631– 11641. [5] F.M. Linseisen, M. Hetzer, T. Brumm, T.M. Bayerl, Differences in the physical properties of lipid monolayers and bilayers on a spherical solid support, Biophys. J. 72 (1997) 1659–1667. [6] E. Reimhult, M. Zäch, F. Höök, B. Kasemo, A multitechnique study of liposome adsorption on Au and lipid bilayer formation on SiO2, Langmuir 22 (2006) 3313–3319. [7] N. Nitin, L.E. LaConte, O. Zurkiya, X. Hu, G. Bao, Functionalization and peptidebased delivery of magnetic nanoparticles as an intracellular MRI contrast agent, J. Biol. Inorg. Chem. 9 (2004) 706–712. [8] G.A. van Tilborg, W.J. Mulder, N. Deckers, G. Storm, C.P. Reutelingsperger, G.J. Strijkers, K. Nicolay, Annexin A5-functionalized bimodal lipid-based contrast agents for the detection of apoptosis, Bioconjugate Chem. 17 (2006) 741–749. [9] H.C. Huang, P.Y. Chang, K. Chang, C.Y. Chen, C.W. Lin, J.H. Chen, C.Y. Mou, Z.F. Chang, F.H. Chang, Formulation of novel lipid-coated magnetic nanoparticles as the probe for in vivo imaging, J. Biomed. Sci. 16 (2009) 86. [10] A. Mueller, D.F. O’Brien, Supramolecular materials via polymerization of mesophases of hydrated amphiphiles, Chem. Rev. 102 (2002) 727–757. [11] M.J. Janiak, D.M. Small, G.G. Shipley, Temperature and compositional dependence of the structure of hydrated dimyristoyl lecithin, J. Biol. Chem. 254 (1979) 6068–6078. [12] W.A. Hendrickson, A. Pähler, J.L. Smith, Y. Satow, E.A. Merritt, R.P. Phizackerley, Crystal structure of core streptavidin determined from multiwavelength anomalous diffraction of synchrotron radiation, Proc. Natl. Acad. Sci. U.S.A. 86 (1989) 2190–2194. [13] Z.G. Estephan, J.A. Jaber, J.B. Schlenoff, Zwitterion-stabilized silica nanoparticles: toward nonstick nano, Langmuir 26 (2010) 16884–16889. [14] H. Wei, N. Insin, J. Lee, H.S. Han, J.M. Cordero, W. Liu, M.G. Bawendi, Compact zwitterion-coated iron oxide nanoparticles for biological applications, Nano Lett. 12 (2012) 22–25. [15] L.L. Rouhana, J.B. Schlenoff, Aggregation resistant zwitterated superparamagnetic nanoparticles, J. Nanopart. Res. 14 (2012) 835.
Contents lists available at ScienceDirect
Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio
Notes & Tips
Preparation of stabilized magnetic nanoparticles with polymerizable lipid and anchor compounds Boram Kang, Suk-Jung Choi ⇑ Department of Chemistry, Gangneung-Wonju National University, Gangneung 210-702, Republic of Korea
a r t i c l e
i n f o
Article history: Received 27 June 2013 Received in revised form 28 September 2013 Accepted 30 October 2013 Available online 8 November 2013 Keywords: Magnetic nanoparticle Surface modification Polymerizable lipid Anchor compound
a b s t r a c t Although the lipid-based method for coating of magnetic nanoparticles (MNPs) is rapid and simple, the unstable state of the lipid layer is a major limitation for the practical application of this method. We devised a method to prepare stabilized MNPs by covalent modifications such as lipid polymerization and anchoring of the lipid layer. The stability of the modified lipid layer was demonstrated by the stable status of enzymes immobilized on the MNPs and the resistance of the MNPs to aggregation. We also determined the maximum ratio of nonpolymerizable lipophilic compounds that can be included in the layer without significantly reducing stability. Ó 2013 Elsevier Inc. All rights reserved.
Magnetic nanoparticles (MNPs) have a wide range of applications in immunoassays, separating molecules or cells, delivering therapeutic molecules, and magnetic resonance imaging (MRI) [1–4]. Prior to use in these applications, the MNP surface must be modified to ensure the stability and solubility of the particles in an aqueous environment. Another purpose of surface modification is to provide MNPs with reactive functional groups that might be used to immobilize biological molecules such as antibodies or oligonucleotides. Lipids have the potential to be used for coating of solid surfaces because of their ability to form bilayer or monolayer structures on surfaces depending on the surface’s hydrophilic or hydrophobic nature [5,6]. The lipid coating method has been applied in the preparation of MRI contrast agents or in vivo imaging agents [7– 9]. It was shown that coating with PEG–lipids (lipids containing a polyethylene glycol head group) enhanced solubility and stability of MNPs while lowering the toxicity of these MNPs to cells [7,9]. The surface was modulated to have a positive charge or fluorescence with the addition of a cationic lipid or a fluorescent lipid [8,9]. The process of lipid coating is very fast and simple in comparison to other methods. Moreover, a required functional group can be easily introduced onto the surface using lipids or lipophilic compounds containing the functional group. Nevertheless, limited stability is a major practical limitation of the lipid coating method, in which the constituent lipid molecules associate via noncovalent interactions. ⇑ Corresponding author. Fax: +82 33 640 2264. E-mail address: [email protected] (S.-J. Choi). 0003-2697/$ - see front matter Ó 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ab.2013.10.036
To prepare stabilized functional MNPs, we devised a new coating method using the polymerizable lipid 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DYPC) and the anchor compound octadecyltrimethoxysilane (ODTS). A lipid film containing 0.25 mg DYPC, 0.011 mg ODTS, and 0.051 mg 1,2-dipalmitoylsn-glycero-3-phosphoethanolamine-N-(biotinyl) (BTPE) dissolved in 0.5 ml chloroform (composition D) was prepared in a glass vial. DYPC is capable of forming lipid networks via polymerization induced by UV irradiation [10]. ODTS can be covalently linked to the MNP surface while inserted in a lipid layer via its hydrophobic tail. BTPE was added to present biotin groups that were used to immobilize streptavidin-linked enzymes. Control experiments were conducted with three lipid films of different compositions: 0.25 mg 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 0.064 mg BTPE (composition A), 0.25 mg DPPC, 0.013 mg ODTS, and 0.064 mg BTPE (composition B), and 0.25 mg DYPC and 0.051 mg BTPE (composition C). For convenience and consistency of the experiment, many films were prepared simultaneously and stored in the freezer until use. As the mole fraction of BTPE was 0.154–0.167 in all compositions, the surface density of the biotin group was sufficient for a saturating level of streptavidin binding, considering the size of the lipid and streptavidin [11,12]. Coating schemes for the four lipid compositions are depicted in Fig. 1a. Iron oxide (Fe3O4) MNPs (100-nm diameter) were suspended in PBS (phosphate-buffered saline) at a concentration of 0.5 mg/ml and then 2 ml of the solution was added to a vial. The vial was filled with nitrogen gas and placed in a bath sonicator (Model SD-D300H, S-D Ultrasonic Cleaner, Seoul, Korea) for 3 h. The vial was sonicated with 80% output power, while temperature was
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Notes & Tips / Anal. Biochem. 446 (2014) 87–89
Fig.1. Lipid coating schemes and enzyme immobilization efficiency. (a) Schematic representations of coating with the four lipid compositions: (1) composition A (DPPC + BTPE), (2) composition B (DPPC + ODTS + BTPE), (3) composition C (DYPC + BTPE), and (4) composition D (DYPC + ODTS + BTPE). (b) Activity of enzymes immobilized on the four kinds of lipid-coated MNPs. Alkaline phosphatase (AP) and horseradish peroxidase (HRP) activity are represented as optical density at 405 nm (light gray) and light intensity (dark gray). Each data point represents the mean of triplicate measures, and the error bars represent the standard deviation.
maintained at 50–60 °C during the process to ensure the lipids remained in a liquid state. For polymerization of DYPC, vials were irradiated for 30 min at 254 nm with a UV lamp. During the polymerization reaction, the MNP was agitated under a nitrogen gas atmosphere in the bath sonicator. The lipid-coated MNPs were biofunctionalized with streptavidin–alkaline phosphatase (SA–AP) or streptavidin–horseradish peroxidase (SA–HRP) to estimate protein immobilization efficiency and lipid layer stability. SA–AP or SA–HRP was added to the lipidcoated MNPs (20 lg protein per 1 mg MNP) and incubated at room temperature for 30 min. After being washed three times with PBS, the biofunctionalized MNPs were suspended in PBS at a concentration of 10 lg/ml and 0.1 ml of the modified MNPs was used for the estimation of enzyme activity. AP activity was estimated by the colorimetric assay method using pNPP solution (10 mM p-nitrophenyl phosphate, 0.5 mM MgCl2, 1 M diethanolamine, pH 9.8). The pNPP solution (0.1 ml) was added to a 96-well microplate containing MNPs and the microplate was incubated at room temperature for 5 min. Absorbance was read at 405 nm with a Synergy-HT plate reader (BioTek, Winooski, VT, USA). HRP activity was estimated by the chemiluminescence assay method in a white 96-well microplate. The microplate containing 0.1 ml of the biofunctionalized MNPs was placed in a LuBi plate reader (Seoulin Bioscience, Seongnam, Korea). Light intensity was recorded over time after injecting 80 ll of luminol solution (2.8 mM luminol, 3.6 mM 4-iodophenol, 0.002% Tween 20, 0.1 M Tris–HCl, pH 8.3) and 80 ll of 3 mM H2O2 solution into each well. As shown in Fig. 1b, activity of the AP enzyme had a tendency to increase as covalent modifications were added to the lipid layers. The enzyme activity was lowest in composition A, in which the DPPC lipid layer was associated using only noncovalent interactions. Positive effects of lipid-layer anchoring and lipid cross-linking were revealed in compositions B and C, respectively. Combined use of DYPC and ODTS resulted in the highest activity, demonstrating a synergistic effect of polymerization and anchoring (composition D). Without UV treatment of compositions C and D, AP enzyme activity was about 85% of the activity of the UV-treated samples (Supplementary Fig. S1). This result confirms that the activity increase in the DYPC-coated MNPs was dependent on the
cross-linking of the lipids. Furthermore, the effects of the covalent modifications on the protein immobilization efficiency were reproducibly observed in the experiments with SA–HRP. One possible explanation for this result is that the covalent modifications effectively reduced the removal of enzymes by stabilizing the lipid layer. The DPPC coating is maintained only by noncovalent interactions and is susceptible to environmental stresses, causing disintegration of the layer and removal of the enzymes. In composition D, however, the enzymes were preserved because the lipid layer was stabilized by two covalent modifications: crosslinking of lipid molecules within the layer and covalent linkage of the layer to the MNP surface. The stabilizing effect of the covalent modifications was also shown by the resistance of these MNPs to detergent treatment. Only 8% of the enzyme activity remained after detergent treatment in the DPPC-coated MNPs, while 40% of the enzyme activity was still associated with the MNPs in composition D (Supplementary Fig. S2). For functionalization of MNPs, functional lipophilic compounds such as BTPE should be included in the lipid layer. However, most of the commercially available functional lipophilic compounds are nonpolymerizable and may have a negative effect on the stability of the lipid layer by preventing the cross-linking of DYPC. Therefore, we estimated the effect of the DYPC ratio on the activity of immobilized enzymes. Lipid-coated MNPs were prepared with lipid films containing DYPC, DPPC, ODTS, and BTPE. The amounts of ODTS and BTPE were kept constant at 10 lg (2.7 lmol) and 2.56 lg (0.3 lmol) in all films. However, the amounts of DYPC and DPPC were varied while the total moles of the two lipids were kept constant at 27.3 lmol. SA–HRP or SA–AP was immobilized on the MNPs and their activity was determined as described previously. The enzyme activity decreased with a decreasing mole fraction of DYPC, but this decrease was discontinuous at DYPC mole fractions lower than 0.72 (Supplementary Fig. 2a). Although the reason for this discontinuity is not clear, the result indicates that the mole fraction of DYPC should be at least 0.72 to prepare a stable lipid coating. In other words, nonpolymerizable functional lipophilic compounds can be included up to a mole fraction of 0.19 without significantly reducing stability, based on the mole fraction of ODTS here (0.09). In the case of composition D, the
Notes & Tips / Anal. Biochem. 446 (2014) 87–89
DYPC mole fraction was 0.76, high enough to form a stable lipid layer. MNPs have hydrophobic surfaces with a large surface area to volume ratio and have a tendency to form aggregates, which is a critical obstacle to biomedical applications [1]. Lipid coating is expected to provide MNPs with resistance to aggregation, as there are several reports describing improved resistance to aggregation in zwitterion-coated nanoparticles [13–15]. Therefore, the aggregation of the biofunctionalized MNPs was estimated by the filtration method. MNPs were coated with composition A or D and biofunctionalized with SA–HRP as described previously. The MNPs were filtered through 0.4-lm Isopore filters (Millipore, Billerica, MA, USA) to remove aggregates and were stored at 4 °C. The MNPs were filtered again each day and HRP activity remaining in the filter was estimated. The degree of aggregation is proportional to the HRP activity associated with the filter because the aggregated MNPs are trapped in the filter. The enzyme activity associated with the filter increased rapidly within 4 days in the DPPC-coated MNP (composition A) but remained almost constant for 3 weeks in the MNP stabilized with composition D (Supplementary Fig. 2b). Aggregation of the DPPC-coated MNPs may be induced by the exposed MNP surfaces that resulted from the partial disintegration of the lipid layer as discussed above. It is also possible that the DPPC layers of different MNPs fused to form larger clusters to release the tension caused by high curvature. In this work, the lipid-based method for MNP coating was improved by covalent modifications of the lipid layer. The stability of the lipid layer that was modified with cross-linking of DYPC and anchoring with ODTS was demonstrated using three methods. First, a higher enzyme immobilization efficiency was achieved by coating with DYPC, ODTS, and BTPE compared with coating with DPPC and BTPE. Second, the immobilized enzyme on the modified layer was more stable following detergent treatment. Finally, the MNPs coated with the modified lipid layer showed higher resistance to aggregation. It was also shown that the mole fraction of DYPC should be at least 0.72 to obtain the stabilizing effect of polymerization. Acknowledgments This research is part of the project titled ‘‘Development of Biosensors for Paralytic Shellfish Toxins’’ funded by the Ministry of Oceans and Fisheries, Korea, and was also supported by the
89
Research Institute of Natural Science of Gangneung-Wonju National University. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ab.2013.10.036. References [1] A.K. Gupta, M. Gupta, Synthesis and surface engineering of iron oxide nanoparticles for biomedical applications, Biomaterials 26 (2005) 3995–4021. [2] S. Laurent, D. Forge, M. Port, A. Roch, C. Robic, L. Vander Elst, R.N. Muller, Magnetic iron oxide nanoparticles: synthesis, stabilization, vectorization, physicochemical characterizations, and biological applications, Chem. Rev. 108 (2008) 2064–2110. [3] J. Gao, H. Gu, B. Xu, Multifunctional magnetic nanoparticles: design, synthesis, and biomedical applications, Acc. Chem. Res. 42 (2009) 1097–1107. [4] S.T. Selvan, T.T. Tan, D.K. Yi, N.R. Jana, Functional and multifunctional nanoparticles for bioimaging and biosensing, Langmuir 26 (2010) 11631– 11641. [5] F.M. Linseisen, M. Hetzer, T. Brumm, T.M. Bayerl, Differences in the physical properties of lipid monolayers and bilayers on a spherical solid support, Biophys. J. 72 (1997) 1659–1667. [6] E. Reimhult, M. Zäch, F. Höök, B. Kasemo, A multitechnique study of liposome adsorption on Au and lipid bilayer formation on SiO2, Langmuir 22 (2006) 3313–3319. [7] N. Nitin, L.E. LaConte, O. Zurkiya, X. Hu, G. Bao, Functionalization and peptidebased delivery of magnetic nanoparticles as an intracellular MRI contrast agent, J. Biol. Inorg. Chem. 9 (2004) 706–712. [8] G.A. van Tilborg, W.J. Mulder, N. Deckers, G. Storm, C.P. Reutelingsperger, G.J. Strijkers, K. Nicolay, Annexin A5-functionalized bimodal lipid-based contrast agents for the detection of apoptosis, Bioconjugate Chem. 17 (2006) 741–749. [9] H.C. Huang, P.Y. Chang, K. Chang, C.Y. Chen, C.W. Lin, J.H. Chen, C.Y. Mou, Z.F. Chang, F.H. Chang, Formulation of novel lipid-coated magnetic nanoparticles as the probe for in vivo imaging, J. Biomed. Sci. 16 (2009) 86. [10] A. Mueller, D.F. O’Brien, Supramolecular materials via polymerization of mesophases of hydrated amphiphiles, Chem. Rev. 102 (2002) 727–757. [11] M.J. Janiak, D.M. Small, G.G. Shipley, Temperature and compositional dependence of the structure of hydrated dimyristoyl lecithin, J. Biol. Chem. 254 (1979) 6068–6078. [12] W.A. Hendrickson, A. Pähler, J.L. Smith, Y. Satow, E.A. Merritt, R.P. Phizackerley, Crystal structure of core streptavidin determined from multiwavelength anomalous diffraction of synchrotron radiation, Proc. Natl. Acad. Sci. U.S.A. 86 (1989) 2190–2194. [13] Z.G. Estephan, J.A. Jaber, J.B. Schlenoff, Zwitterion-stabilized silica nanoparticles: toward nonstick nano, Langmuir 26 (2010) 16884–16889. [14] H. Wei, N. Insin, J. Lee, H.S. Han, J.M. Cordero, W. Liu, M.G. Bawendi, Compact zwitterion-coated iron oxide nanoparticles for biological applications, Nano Lett. 12 (2012) 22–25. [15] L.L. Rouhana, J.B. Schlenoff, Aggregation resistant zwitterated superparamagnetic nanoparticles, J. Nanopart. Res. 14 (2012) 835.
