Shell Magnetic Nanoparticles with Plasmonic

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Ag/FeCo/Ag Core/shell/shell Magnetic Nanoparticles with Plasmonic Imaging Capability Mari Takahashi, Priyank Mohan, Akiko Nakade, Koichi Higashimine, Derek Mott, Tsutomu Hamada, Kazuaki Matsumura, Tomohiko Taguchi, and Shinya Maenosono Langmuir, Just Accepted Manuscript • DOI: 10.1021/la5046805 • Publication Date (Web): 23 Jan 2015 Downloaded from http://pubs.acs.org on February 1, 2015

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Ag/FeCo/Ag Core/shell/shell Magnetic Nanoparticles with Plasmonic Imaging Capability Mari Takahashi,1 Priyank Mohan,1 Akiko Nakade,1 Koichi Higashimine,1 Derrick Mott,1 Tsutomu Hamada,1 Kazuaki Matsumura,1 Tomohiko Taguchi,2 and Shinya Maenosono1,*

1

School of Materials Science, Japan Advanced Institute of Science and Technology, 1-1

Asahidai, Nomi, Ishikawa 923-1292, Japan 2

Department of Health Chemistry, Graduate School of Pharmaceutical Sciences, The University

of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan

KEYWORDS: Magnetic separation, Plasmon imaging, Superparamagnetism, Iron cobalt, Silver

ABSTRACT: Magnetic nanoparticles (NPs) have been used to separate various species such as bacteria, cells and proteins. In this study, we synthesized Ag/FeCo/Ag core/shell/shell NPs designed for magnetic separation of subcellular components like intracellular vesicles. A benefit of these NPs is that their silver metal content allows plasmon scattering to be used as a tool to observe detection by the NPs easily and semi-permanently. Therefore, these NPs are considered a potential alternative to existing fluorescent probes like dye molecules and colloidal quantum

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dots. In addition, the Ag core inside the NPs suppresses the oxidation of FeCo because of electron transfer from the Ag core to the FeCo shell, even though FeCo is typically susceptible to oxidation. The surfaces of the Ag/FeCo/Ag NPs were functionalized with ε-poly-L-lysine-based hydrophilic polymers to make them water soluble and biocompatible. The imaging capability of the polymer-functionalized NPs induced by plasmon scattering from the Ag core was investigated. The response of the NPs to a magnetic field using liposomes as platforms and applying a magnetic field during observation by confocal laser scanning microscopy was assessed. The results of the magnetophoresis experiments of liposomes allowed us to calculate the magnetic force to which each liposome was subjected.

Introduction Cells take up extracellular solute and proteins/lipids on the plasma membranes (PM) by endocytosis and package them into endocytic vesicles.1 The internalized materials are then either delivered to lysosomes for degradation or recycled back to the PM for reuse. This process is fundamental in many biological processes, such as nutrition uptake, regulation of mitogenic signaling, cell differentiation and locomotion, and immune response.2 As expected, impaired endocytosis is known to cause many diseases, such as familial hypercholesterolemia3,4 and cancer.5 Moreover, some pathogenes such as viruses and bacteria exploit endocytosis to invade host cells. Thus, understanding the molecular machinery that regulates endocytosis and characterizing the protein/lipid components of endocytic vesicles is essential for cell biology and pathology. Recently, magnetic separation of endocytic vesicles/endosomes has been reported by several groups.6,7 The merit of magnetic separation is to obtain purified and un-damaged target


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materials compared to other techniques such as ultracentrifugation and density-gradient separation.8 Three desirable properties of a magnetic probe for endocytic research are imaging capability, small size and high magnetism. With imaging capability, one can make sure that the magnetic probe is correctly targeted to endocytic vesicles or endosomes. The second factor is the size of the probes. Clathrin-coated vesicles are one of the major endocytic vesicles.9 The vesicles are around 100–150 nm in diameter. Therefore, the size of magnetic probes should be less than 100 nm for the magnetic probes to be packaged into the clathrin-coated vesicles. In this context, smaller probes should be better. However, since the saturation magnetization (which determines the efficiency of magnetic separation) tends to decrease as the size or volume of the magnetic probe decreases, there is a significant limitation in smaller sized particles. Magnetism is also an important parameter of a magnetic probe since a magnetic probe must respond sufficiently to an external magnetic field to ensure effective magnetic separation. A magnetic probe with high saturation magnetization should therefore be used. However, this is a challenge because magnetic materials with high saturation magnetization such as Fe, Co, and FeCo are susceptible to oxidation, which decreases saturation magnetization. A magnetic probe that possesses imaging capability, small size and high saturation magnetization should allow effective magnetic separation of intracellular vesicles, which would be a breakthrough in biology-related nanotechnology. To achieve these properties simultaneously, there are several candidates in terms of structure and composition. Here we discuss the concrete design of the magnetic probe according to the required attributes. First is image capability. Metal nanostructures such as Au and Ag nanoparticles (NPs) are used for many kinds of bioimaging10,11 as well as in other fields because of their localized surface plasmon


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resonance (LSPR). Imaging using LSPR is considered to be a promising technique for bioimaging because it enables semi-permanent observation, and thus, magneto-plasmonic hybrid NPs are suitable as a magnetic probe with imaging capability. Second, it is necessary to simultaneously meet the requirements of small size and high saturation magnetization. Several types of magnetic-plasmonic hybrid NPs have been reported including Au-Fe3O4 heterostructured NPs,12 Co/Au core/shell NPs,13 and FePt-Au heterostructured NPs.14 In the present study, we combine two magnetic and plasmonic materials to meet the requirements of imaging capability, small size and high magnetism; FeCo and Ag. FeCo possesses the highest saturation magnetization of known magnetic materials, while Ag has the highest scattering cross section of existing plasmonic materials. Several Ag and FeCo hybrid NPs have been reported. For example, Sachan and coworkers synthesized Ag-FeCo and Ag-Co heterostructured NPs by pulsed laser dewetting.15 However, Ag-FeCo hybrid NPs have not been chemically synthesized. We considered Ag/FeCo core/shell NPs that were covered with an outer Ag shell to enhance ligand exchange using thiol groups; i.e., Ag/FeCo/Ag core/shell/shell NPs, because we could not prevent oxidation of the FeCo NPs or cover the FeCo NPs with Ag (partly because of the galvanic replacement reaction between FeCo and Ag ions) when we tried to synthesize FeCo/Ag core/shell NPs. This structure avoids oxidation of FeCo mainly because of electron transfer from Ag core to FeCo shell. Similar electron transfer has been investigated in our group previously,16,17 where we found that oxidation of Ag shell was suppressed by electron transfer from Au core to Ag shell in Au/Ag core/shell NPs. To use the Ag/FeCo/Ag NPs for bioapplications, we had to make them water soluble because the as-synthesized NPs could only be dispersed in a non-polar solvent. Therefore, we modified the NP surface with a hydrophilic polymer, ε-poly-L-lysine (PLL). PLL is a synthetic amino acid


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polymer that has been used as a coating for Ag NPs18 to make them water soluble. We introduced succinic anhydride onto PLL, resulting in NPs with a tunable charge, which led to low cytotoxicity.19 In addition, 2-iminothiolane was introduced to bind PLL to the surface of the NPs through metal-thiol interactions. We also investigated magnetophoresis of liposomes by attaching the NPs onto the surface of liposomes non-specifically under observation by confocal laser scanning microscopy (CLSM). Based on these results, we calculated the magnetic force that the liposome was subjected to by the NPs and estimated the number of NPs needed to isolate intracellular vesicles from cells. Experimental Section Chemicals Cobalt acetylacetonate (Co(acac)2, purity 97%), iron acetylacetonate (Fe(acac)3, purity ≥99.9%), silver nitrate (AgNO3, purity ≥99.9999%), 1,2-hexadecanediol (purity 90%), succinic anhydride (purity ≥99%), sodium hydroxide (NaOH, purity ≥98%), oleylamine (OLA, purity 70%), oleic acid (OA, purity 90%), and tetraethylene glycol (TEG, purity 99%) were purchased from Sigma-Aldrich and used as received. ε-Poly-L-lysine (PLL) and 2-iminothiolane were purchased from JNC Co. and Toronto Research Chemicals, respectively. 1,2-Dipalmitoyl-snglycero-3-phosphocholine (DPPC, 16:0) was purchased from Avanti Polar Lipids. Hexane and ethanol were purchased from Nacalai Tesque. Acetone was purchased from Kanto Chemical, and toluene and hydrochloric acid (HCl) were purchased from Wako Pure Chemical. Synthesis of Ag/FeCo/Ag core/shell/shell NPs


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Ag/FeCo/Ag core/shell/shell NPs were synthesized by the hot injection method in combination with Chaubey et al.’s polyol method for the synthesis of FeCo NPs20 with modification. Scheme S1 (Supporting Information) shows the synthetic procedure used to prepare Ag/FeCo/Ag NPs in which 1,2-hexadecanediol served as a reducing agent, OLA and OA were reducing and capping agents, and TEG was a reducing agent and solvent during the reaction. First, AgNO3 (0.1 mmol), 1,2-hexadecanediol (1.0 mmol), OA (8 mmol), OLA (10 mmol) and TEG (10 mL) were added to a three-necked flask. The flask was connected to a trap sphere that was also connected to a condenser tube. The temperature of the reaction solution was monitored by a thermocouple. The reaction mixture was degassed with Ar for 5 min at room temperature with vigorous stirring. The reaction mixture was heated to 100 °C for 10 min to remove volatile substances from the mixture, and then the temperature was increased to 170 °C. When the reaction temperature reached 170 °C, a solution of Fe(acac)3 and Co(acac)2 (both 0.2 mmol) in OLA (1 mL) and toluene (2 mL) was injected into the reaction solution, keeping the temperature at 170 °C. The temperature was increased to 250 °C and a solution of AgNO3 (0.1 mmol) in OLA (1 mL) and toluene (1 mL) was injected into the reaction solution to make the Ag outer shell. The temperature was reduced to 230 °C for 10 min. The reaction mixture was then cooled to room temperature. The mixture was separated into two centrifuge tubes and acetone was added to make a total volume 45 mL followed by centrifugation at 4500 rpm for 5 min. The supernatant was removed and hexane (400 µL) was added to each tube to disperse the NPs. The total solution was divided into four tubes so that each tube contained 200 µL of hexane in which the NPs were dispersed. Acetone was added to make a total volume of 45 mL following by centrifugation at 4500 rpm for 5 min. The NPs were dried in a vacuum drying system after removing the supernatant.


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To compare the electronic states of each element in the Ag/FeCo/Ag NPs, FeCo NPs were synthesized in the absence of Ag precursor for comparison using a similar system to that employed to synthesize the Ag/FeCo/Ag NPs. Fe(acac)3 and Co(acac)2 (0.2 mmol of each), 1,2hexadecanediol (1.0 mmol), OA (8 mmol), OLA (10 mmol) and TEG (10 mL) were added to a three-necked flask, and then the same process was followed until after the volatile substances were removed. The temperature was increased to 290 °C, held for 20 min, and then allowed to cool naturally. The resulting NPs were washed in the same manner as the Ag/FeCo/Ag NPs. Characterization of Ag/FeCo/Ag core/shell/shell NPs The Ag/FeCo/Ag NPs were characterized by transmission electron microscopy (TEM), highresolution TEM (HR-TEM), scanning TEM equipped with a high-angle annular dark-field (STEM-HAADF) detector, energy-dispersive X-ray spectroscopy (EDS) elemental mapping, Xray photoelectron spectroscopy (XPS), X-ray diffraction (XRD), inductively coupled plasma optical emission spectroscopy (ICP-OES), superconducting quantum interference device (SQUID) magnetometry and ultraviolet-visible spectroscopy (UV-vis). TEM analysis was performed on an electron microscope (Hitachi H-7650) operated at 100 kV. HR-TEM, STEMHAADF and EDS elemental mapping were performed on a different microscope (JEOL JEMARM200F) operated at 200 kV with a spherical aberration corrector, and a nominal resolution of 0.8 Å. XPS analysis was performed on a high-performance XPS system (Shimadzu Kratos AXIS-ULTRA DLD). Photoelectrons were excited by monochromated Al Kα radiation. XRD patterns were collected on an X-ray diffractometer (Rigaku MiniFlex600) operated in reflection geometry at room temperature with Cu Kα radiation (1.5418 Å). ICP-OES analysis was performed on a sequential plasma spectrometer (Shimadzu ICPS-7000). SQUID analysis was


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performed on a magnetometer (Quantum Design MPMS). UV-vis analysis was performed on an absorption spectrometer (PerkinElmer Lambda35). Synthesis of PLL-based polymer PLL is composed of amino acids so it has high biocompatibility and numerous amino groups that can be used to introduce functional groups. Succinic anhydride was introduced onto some of the amino groups of PLL to produce polyampholyte (PLL-COOH) as reported previously.19,21 Furthermore, 2-iminothiolane was introduced onto PLL-COOH to enhance binding to the surfaces of the Ag/FeCo/Ag core/shell/shell NPs through thiol-metal interactions. An aqueous solution of PLL (10 mmol, 25 wt%), which has 32 lysine residues,21 was prepared, and then succinic anhydride (7 mmol) was added. The solution was stirred at 50 °C for 1 h to introduce carboxyl groups with a substitution ratio of 70% (PLL-COOH). 2-Iminothiolane (2 mmol) was added and the reaction solution was stirred at room temperature for 2 h to introduce thiol groups with a substitution ratio of 20% (PLL-COOH-SH). The products were obtained following dialysis for 2 days and freeze-drying for 2 days. The substitution ratio of each functional group was calculated from the nuclear magnetic resonance (NMR) spectra of the polymer samples. 1H NMR spectra of PLL-COOH and PLL-COOH-SH dissolved in D2O were recorded at 25 °C using a 400-MHz NMR spectrometer (Bruker Biospin AVANCE III). To estimate the substitution ratio of succinic anhydride, an NMR spectrum of PLL-COOH was measured before introducing 2-iminothiolane. The structure and 1H NMR spectrum of the PLL-COOH-SH polymer are presented in Figure S1a and b (Supporting Information), respectively. The degree of substitution of PLL with succinic anhydride and 2-iminothiolane was estimated to be 70% and 12%, respectively.


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The charge of the polymers was determined by plotting zeta potential against pH (Figure S1c). The zeta potential of PLL-COOH-SH dissolved in pure water (1 mg/mL) was measured by a Malvern Zetasizer Nano ZS ZEN3600. To tune the pH of each polymer solution (1 mL), a minute amount of HCl or NaOH was added. These measurements were repeated several times and then average and standard deviation were estimated for each pH. PLL-COOH-SH has a pH of 5 when it is dissolved in water, and shows a zeta potential of −0.6 ± 0.2 mV in water at pH 5. This means that the isoelectric point of the polymer is pH 5. Ligand exchange of the NPs Because the as-synthesized NPs were capped with OLA, the NPs could only be dispersed in non-polar solvent. To make them water soluble and functionalize the NPs, ligand exchange of the surface molecules on the NPs was carried out. A hexane dispersion of NPs (17 mg/mL) was prepared. PLL-COOH-SH (15 mg) was dispersed in pure water (1 mL). Ethanol (100 µL) was added to the polymer solution and the pH of the polymer solution was adjusted to 12 with NaOH. The hexane dispersion of NPs (200 µL) was added to the polymer solution, followed by sonication to enhance the phase transfer reaction at the interface between the water and hexane phases. During sonication, more pure water (2 mL) was added. The mixture was transferred into six 1.5-mL tubes. The total volume of each tube was increased to 1 mL by adding pure water, and then the tubes were centrifuged at 40000 rpm for 3 min. The supernatant was removed and a small amount of pure water was added to redisperse the NPs. Finally, all NPs were collected in one tube. A photograph of aqueous dispersion of PLL-COOH-SH-modified Ag/FeCo/Ag NPs is shown in Figure S2 in the Supporting Information. Real-time plasmonic imaging of the magnetophoresis of NP-loaded liposome


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To assess the feasibility of PLL-COOH-SH-modified Ag/FeCo/Ag NPs as magnetic probes for subcellular magnetic separation, the NPs were adsorbed onto the surface of DPPC (16:0) liposome. The motion of the NP-loaded DPPC liposome was observed by CLSM (Olympus FV1000D) under an external magnetic field. DPPC liposome containing glucose (0.1 M) was prepared by an electroformation method.22 DPPC liposome solution with a total lipid concentration of 0.1 mM (10 µL) was mixed with an aqueous dispersion of Ag/FeCo/Ag NPs (10 µL, 500 µg/mL). Aqueous glucose solution (10 µL, 0.2 M) was immediately added to the liposome/NP mixture, which was aged for 5 min to adsorb the NPs onto the surfaces of the DPPC liposome. Aqueous sucrose solution (30 µL, 0.1 M) was added to increase the buoyancy of the NP-loaded liposome. NP-loaded liposome solution (2.5 µL) was added dropwise onto a glass coverslip on which a silicone rubber spacer with a thickness of 0.1 mm was placed. Then, another glass coverslip whose surface was coated with bovine serum albumin (BSA) was placed on top to enclose the NP-loaded liposome solution. As a result, most of the NP-loaded liposome stayed in close proximity to the upper BSA-coated glass coverslip. This facilitated CLSM observation of magnetophoretic motion of liposomes because the liposome moved horizontally along the surface of the upper glass coverslip. The BSA layer suppressed the adsorption of liposome onto the glass surface. Figure S3 (Supporting Information) shows the setup of the CLSM. To obtain plasmonic scattering images, a laser wavelength of 473 nm was used. By adjusting the focal plane, reflection from the glass substrate was avoided. Transmission images of the sample were also taken. A magnetic field was applied using neodymium magnets and an instrument made of stainless steel, as shown in Figure S4 (Supporting Information). The magnetic field is 30-40 mT at a 1 mm distance from the tip of the instrument which corresponds to the distance between liposomes and the tip.


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Results Structure and composition of Ag/FeCo/Ag core/shell/shell NPs Figure S5a and b (Supporting Information) show TEM and HR-TEM images of the Ag/FeCo/Ag NPs, respectively. Figure S5c shows the size distribution of the Ag/FeCo/Ag NPs. The mean diameter of the NPs calculated from TEM images was around 13.5 ± 2.5 nm (n = 531). The histogram shows that the NPs are of uniform size. STEM-HAADF (Z-contrast) images of the NPs are presented in Figure S5d–f. Because the intensity (brightness) is approximately proportional to the square of the atomic number (Z2), the Ag core is brighter than the FeCo shell. These images indicate that most of the NPs have a Ag core and FeCo shell. The EDS images of Ag/FeCo/Ag NPs in Figure S5g–r clearly confirm the Ag/FeCo/Ag core/shell/shell heterostructure of the NPs. The mean thickness of the FeCo shell was estimated to be about 3.1 nm by subtracting the average size of the Ag cores (7.4 nm) from the average size of the Ag/FeCo/Ag NPs (13.5 nm assuming the thickness of the outer Ag shell is negligible). To examine the composition distribution in a single NP more closely, cross-sectional line profile analysis was performed for a single NP (Figure 1). This image confirms that the Fe/Co ratio is smaller at the interface between the Ag core and FeCo shell than at the periphery of the NPs. The presence of the outer Ag shell is also confirmed. We do not rule out the possibility that the outer shell is FeCoAg ternary alloy. Similar results were obtained for other single NPs, indicating that these structural features are typical and uniform. The composition of Ag/FeCo/Ag NPs was determined by EDS (Fe:Co:Ag = 31:40:29) and ICP-OES (Fe:Co:Ag = 36:45:19) analyses. The composition of the NPs is presented in Table S1 (Supporting Information). Assuming that the Ag core is 7.4 nm in diameter, the shell of FeCo is 3.1 nm thick, the ratio of Fe to Co is 1:1 and the thickness of the outer Ag shell is negligible, the composition of the Ag/FeCo/Ag NPs was


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calculated theoretically (Fe:Co:Ag = 44:44:12); these values are also included in Table S1. The composition determined by EDS analysis contains a relatively high Ag content. This is because EDS analysis is surface sensitive, so the signal from the Ag outer shell becomes dominant.

Figure 1. (a) STEM-HAADF image of a single Ag/FeCo/Ag NP. (b–e) EDS elemental mapping images of the single Ag/FeCo/Ag NP: (b) Ag L edge, (c) Fe K edge, (d) Co K edge, and (e) overlaid image. (f) The EDS line profile at the center of the NP indicated by a yellow line in (a– e). Blue, green and red lines correspond to Ag L, Co K and Fe K edge intensities, respectively. Dashed and solid lines represent raw and low-pass-filtered profiles, respectively.

Figure S6 (Supporting Information) shows an XRD pattern of the NPs. No oxide phase was detected in Ag/FeCo/Ag NPs. Figure S7 (Supporting Information) shows a TEM image and


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XRD pattern for FeCo NPs synthesized in the absence of Ag precursor for comparison. These FeCo NPs exhibit a popcorn-like shape presumably caused by aggregation and sintering of small NPs during the reaction (Figure S7a). The XRD pattern of the NPs (Figure S7b) clearly indicates that the resulting NPs are formed of CoxFe1-xO monoxide phase.23 FeCo phase is not observed. In the synthesis of CoxFe1-xO NPs, the reaction temperature was kept at 290 °C because very few NPs were obtained when the reaction temperature was fixed at 250 °C as was used for the Ag/FeCo/Ag NPs (see Scheme S1). It is possible that the Ag cores act as a catalyst for the formation of the FeCo shell in the Ag/FeCo/Ag NPs. XPS analysis of Ag/FeCo/Ag NPs To examine this phenomenon more closely, core-level XPS analysis of Ag/FeCo/Ag and CoxFe1-xO NPs was performed. Figure 2a and b show Fe 2p and Co 2p XPS spectra for Ag/FeCo/Ag NPs, respectively. Ag 3d XPS spectrum for Ag/FeCo/Ag NPs is shown in Figure S8 (Supporting Information). Figure 2c and d depict Fe 2p and Co 2p XPS spectra for CoxFe1-xO NPs. The Fe 2p spectra for both types of NPs could be deconvoluted into five different Fe species, including Fe0, Fe2+, Fe3+, high-binding energy (BE) surface structures, and satellite peaks24 using a Gaussian–Lorentzian mixed function by XPSPEAK41 software. Similarly, the Co 2p spectra for both types of NPs could be deconvoluted into four different Co species, including Co0, Co2+, Co3+, and satellite peaks.25-28 The BE of each peak for Ag/FeCo/Ag and CoxFe1-xO NPs are summarized in Tables S2 and S3 (Supporting Information), respectively. The atomic ratio between Fe and Co and the relative proportions of each species determined by XPS are listed in Table S4 (Supporting Information). The Fe content is larger than Co content according to the XPS data of Ag/FeCo/Ag NPs (Fe:Co = 63:37), unlike the composition determined by ICP-OES (see Table S1). This is because XPS is a surface-sensitive technique, so


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it gives the composition at the surface of the Ag/FeCo/Ag NPs. This result is consistent with the line profile determined by EDS (Figure 1f). In addition, the fraction of both Fe0 and Co0 species is markedly larger in the Ag/FeCo/Ag NPs (16 and 33 rel. %, respectively) than in the CoxFe1-xO ones (4 and 0 rel. %, respectively) (Table S4). This result strongly suggests that the oxidation of the FeCo shell was well suppressed by electron transfer from the Ag core to the FeCo shell.

Figure 2. (a,c) Fe 2p and (b,d) Co 2p core-level XPS for Ag/FeCo/Ag NPs (upper panels) and CoxFe1-xO NPs (lower panels). Black, yellow, blue and red curves represent raw data, Shirley background, deconvoluted peaks and the sum of all components, respectively.

Magnetic and plasmonic properties


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The magnetic properties of the Ag/FeCo/Ag NPs were measured by SQUID magnetometry. Figure 3a shows the M-H curves measured at 5 and 300 K for the as-synthesized Ag/FeCo/Ag NPs. The M-H curve at 5 K exhibits hysteresis while that measured at 300 K exhibits a Langevin function, indicating the Ag/FeCo/Ag NPs are superparamagnetic at 300 K. The saturation magnetization of the NPs at 300 K is around 36.4 (emu/g), which was calculated using the total mass of the NPs including diamagnetic Ag. However, if the saturation magnetization is recalculated using only the mass of the FeCo shell, the saturation magnetization of the NPs is 45.7 (emu/g), which corresponds to 372 (emu/cm3). Even so, this value was much lower than the saturation magnetization of bulk FeCo (1700 emu/cm3).29 This is possibly because the crystallinity of FeCo was not very high, as indicated in the XRD pattern. In fact, the saturation magnetization of Ag/FeCo/Ag NPs increased when the reaction temperature was increased (data not shown). However, the size distribution of those NPs became broader than the NPs synthesized under standard conditions. Figure 3b shows field-cooled (FC) and zero-field-cooled (ZFC) curves obtained for the Ag/FeCo/Ag NPs. Using a blocking temperature (TB) of 200 K, and assuming the size of the Ag core and thickness of the FeCo shell mentioned above, the magnetocrystalline anisotropy energy (K) of these NPs was calculated to be 64.1 kJ/m3 according to the formula: 25 = ,

(1)

where kB is the Boltzmann constant, and V is the volume of the FeCo shell in a single Ag/FeCo/Ag NP. This value is quite high because K of bulk FeCo is 1.5 kJ/m3.29 Vichery and coworkers reported that Co-doped iron oxide (Fe2O3:Co) NPs have K = 114 kJ/m3.30 Combining the results of XPS and SQUID analyses, we can conclude that the oxidation of the FeCo shell is suppressed near the interface between the Ag core and FeCo shell, but a thin oxide layer is


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formed at the interface between the FeCo shell and Ag outer shell, which results in the high value of K because charge transfer can take place only near the interface between the Ag core and FeCo shell.16,17 Figure 3c shows the M-H curves measured at 5 and 300 K for the PLLCOOH-SH-modified Ag/FeCo/Ag NPs. The saturation magnetization of the PLL-COOH-SHmodified Ag/FeCo/Ag NPs was measured to be 19.2 emu/g which was almost half value of that of the as-synthesized Ag/FeCo/Ag NPs. This is because it was difficult to purify the PLLCOOH-SH-modified Ag/FeCo/Ag NPs after the ligand exchange and, thus, excessive amounts of free PLL-COOH-SH polymer were remaining in the sample. However, TB (i.e. the magnetocrystalline anisotropy energy) of the PLL-COOH-SH-modified Ag/FeCo/Ag NPs is little changed from that of the as-synthesized Ag/FeCo/Ag NPs, as shown in Figure 3d. This result indicates that the ligand exchange doesn’t affect the intrinsic magnetic properties of FeCo shell. A UV-vis spectrum of the Ag/FeCo/Ag NPs dispersed in hexane (Figure 3e) clearly shows a LSPR peak of Ag around 409 nm. In many bioapplications it is necessary to know the concentration of probes, so to estimate concentration of the Ag/FeCo/Ag NPs we made a calibration line for them by plotting the intensity of absorbance as a function of concentration, as shown in Figure 3f. The absorbance of fifteen samples of known concentration was measured and plotted. The line drawn between these points shows a strong linear relationship with R2 = 0.99 and an equation of y = 0.024x, where y is the maximum absorption of the LSPR peak and x is concentration (µg/mL). Using this calibration curve, the concentration of Ag/FeCo/Ag NPs in the magnetophoresis experiment of liposome was estimated. The extinction coefficient of the Ag/FeCo/Ag core/shell/shell NPs was estimated to be 1.6×108 L·mol−1·cm−1. This value is slightly lower than that of pure Ag NPs with a diameter of 10 nm (5.56×108 L·mol−1·cm−1),31


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presumably because the Ag core size is smaller than 10 nm and/or the FeCo shell screens the LSPR scattering of the Ag core. The contribution of the Ag outer shell to the LSPR band is negligible because the Ag outer shell is very thin (see Figure 1). If the thickness of the FeCo shell is increased, the LSPR scattering will be dampened more. This means that the LSPR has a trade-off relationship with the magnetic properties of the Ag/FeCo/Ag NPs.


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Figure 3. (a) Magnetization curves of as-synthesized Ag/FeCo/Ag NPs measured at 5 (black) and 300 K (red). (b) FC (black) and ZFC (red) curves of as-synthesized Ag/FeCo/Ag NPs. (c) Magnetization curves of PLL-COOH-SH-modified Ag/FeCo/Ag NPs measured at 5 (black) and 300 K (red). (d) FC (black) and ZFC (red) curves of PLL-COOH-SH-modified Ag/FeCo/Ag NPs. (e) UV-vis spectrum of a hexane dispersion of as-synthesized Ag/FeCo/Ag NPs. (f) LSPR peak absorbance plotted against NP concentration.

Magnetophoretic motion of DPPC liposome Magnetophoretic motion of NP-loaded DPPC liposomes were recorded under CLSM (a movie is available in the Supporting Information). A magnetic field was applied 13 s after starting to record the movie. Here, t is defined as the elapsed time from the start of recording the movie. Note that the zeta potential of DPPC liposomes in pure water was −0.15 ± 0.05 mV. Figure 4 shows some snapshots of the movie (transmission, plasmonic and their merged images) during application of an external magnetic field. Red spots in the image correspond to the plasmon scattering from the Ag/FeCo/Ag NPs.


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Figure 4. Snapshots of CLSM images: transmission (left), plasmon scattering (middle) and merged (right) images. The images were obtained at (a) t = 53 s, (b) t = 62 s, (c) t = 72 s, (d) t = 82 s, and (e) t = 92 s. Yellow arrows indicate the direction of magnetic force.

The red spots clearly accumulated at the periphery of DPPC liposomes, indicating that PLLCOOH-SH-modified Ag/FeCo/Ag NPs are effectively adsorbed onto the surface of the liposome. In addition, excess NPs formed chain-like aggregates, probably through the magnetic dipole interaction between the NPs. Both NP-loaded liposomes and chain-like NP aggregates moved rapidly in the aqueous sucrose solution to align along the direction of the external magnetic field. Ag/FeCo/Ag NPs modified with PLL-SH polymer lacking COOH groups, and thus is positively charged in water, were not effectively adsorbed onto the surface of DPPC liposomes (data not shown). The reason for this remains to be investigated.


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The trajectories of eight randomly selected liposomes were carefully analyzed and the magnetophoresis velocities of each liposome were estimated, as shown in Figure 5. There are four main types of motion observed for the liposomes: 1) constant speed, 2) sudden acceleration, 3) sudden arrest, and 4) stick slip. Figure 5a shows the time dependence of the magnetophoretic velocity of four liposomes that can be regarded as moving with nearly constant speed. Some of the liposomes were not in contact with the upper glass surface at the start of recording the movie, so the magnetophoretic velocity decreased because of contact of these liposomes with the glass surface after applying an external magnetic field, which causes the liposomes to float. Once the liposomes were in contact with the glass surface, they started to move with constant speed. Figure 5b shows the time dependence of the magnetophoretic velocity of a liposome whose motion suddenly accelerated, while Figure 5c depicts the time dependence of velocity of a liposome whose motion suddenly stopped at t = 136 s. Note that this liposome clearly showed a stick-slip motion just before stopping. In the case of Figure 5b, a chain-like NP aggregate collided with the liposome and the liposome was pulled by the NP aggregate increasing its velocity (Figure S9 in the Supporting Information). In the case of Figure 5c, on the other hand, the liposome suddenly stopped possibly because it was adsorbed onto the glass surface (Figure S10 in the Supporting Information). Figure 5d illustrates the time dependence of velocity of the two remaining liposomes. In this case, the motion of one liposome stopped at t = 112 s. The other liposome collided with the first liposome around t = 128 s. After the collision, these two liposomes stuck together and started to move with stick-slip oscillation (Figure S11 in the Supporting Information).


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Figure 5. Time dependence of magnetophoretic velocity of (a) four different liposomes that moved with nearly constant speed, (b) a liposome whose motion suddenly accelerated at t = 40 s, (c) a liposome whose motion suddenly stopped at t = 136 s, and (d) two different liposomes that stuck together at t = 128 s and started to move as one. Note that the magnetophoretic velocity of a liposome could be obtained only when it was present within the fixed observation area.

Discussion Formation mechanism of Ag/FeCo/Ag NPs The reduction potential of Ag+/Ag (0.8 V versus SHE) is highest among the three elements in the NPs, and thus, the Ag cores can easily form at relatively low temperature. After forming Ag cores, Fe and Co precursors were injected at 170 °C (see Scheme S1 in the Supporting


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Information). Because the reduction potential of Co2+/Co (−0.28 V versus SHE) is slightly higher than that of Fe2+/Fe (−0.45 V versus SHE), Co could be reduced first, resulting in a compositional gradient in the FeCo shell. Very few NPs were obtained when the synthesis was carried out in the absence of Ag cores. In contrast, when we tried to synthesize Ag/Co and Ag/Fe core/shell NPs, the former was produced while the latter was not. These results suggest that Ag cores act as a catalyst for the reduction of Co2+, and Co is required to reduce Fe cations to form the FeCo shell. This is consistent with the results of EDS elemental mapping (Figure 1). As shown in Table S4 (Supporting Information), the proportion of Co0 among Co species (33%) is much larger than that of Fe0 among Fe species (16%) in the Ag/FeCo/Ag NPs. This suggests that electron transfer from Ag to Co occurs more readily than from Ag to Fe presumably because the surface of the Ag core is rich in Co. Adsorption of NPs onto DPPC liposome The zeta potentials of PLL-COOH-SH polymer and DPPC liposome in water are −0.6 and −0.15 mV, respectively. Therefore, the adsorption of PLL-COOH-SH-modified NPs onto DPPC liposome mainly stems from van der Waals interactions. Interestingly, when we used 1,2dioleoyl-sn-glycero-3-phosphocholine (DOPC) liposome instead of DPPC liposome, the NPs were not effectively adsorbed onto the surfaces of DOCP liposome. In addition, the adsorption of NPs onto DPPC liposome was markedly suppressed when cholesterol was added to the DPPC liposome. It has been reported that two forces compete with each other when colloidal particles interact with liposome through van der Waals interactions: one is the adsorption force and the other is the elastic force of the lipid membrane. Hamada and coworkers investigated the interactions between the surfaces of lipid membranes and colloidal polystyrene (PS) beads.22 They found that PS beads bigger than a critical size tend to adsorb on disordered (soft)


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membranes rich in unsaturated DOPC lipid, and PS beads smaller than the critical size tend to adsorb on ordered (hard) membranes rich in saturated DPPC lipid. In the case of particles bigger than the critical size, the adsorption force is larger than the elastic force of the membrane, resulting in membrane deformation. Conversely, in the case of particles smaller than the critical size, the membrane deformation is minor because the adsorption force is weak. The critical radius, r*, for membrane deformation is given as32 ∗ = 2 ⁄ ,

(2)

where κ is bending stiffness and w is adhesion energy (J·m−2), which can be expressed as

= ⁄12 ,

(3)

where A is the Hamaker constant and D is the distance between the surface of a colloidal particle and that of a membrane.33 A can be calculated as = − !"# − !,

(4)

where Asilver = 186.3×10−20 J,34 Alipid = 8.0×10−20 J and Awater = 3.7×10−20 J are the Hamaker constants for Ag, lipid membrane, and water, respectively. Assuming D = 3 nm,22 one can calculate w. By substituting κ = 1.2×10−19 J35 into Eq. (2), r* was estimated to be 61 nm, so the critical diameter is 122 nm. Because the mean diameter of Ag/FeCo/Ag NPs is 13.5 nm, which is much smaller than the critical diameter, the NPs tend to adsorb on hard DPPC liposome rather than soft DOPC liposome or cholesterol-containing DPPC liposome, which is softer than pure DPPC liposome. Estimation of the magnetic force acting on a single Ag/FeCo/Ag NP


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To estimate the magnetic force acting on a single NP, we need to know the magnetic force and number of adsorbed NPs for a single liposome. First, the total surface area of a DPPC liposome was calculated assuming that the whole liposome consists of lipid bilayer and the areas of inner and outer surfaces are equal. Because the total number of lipid molecules in a liposome is 6.02×1014, the number of lipid molecules that compose the outer surface of the liposome is estimated to be 3.01×1014. The projected area of a single DPPC molecule is 0.48 nm2,36 so the total surface area of liposome was calculated to be 1.6×1014 nm2 assuming a two-dimensional close-packed fraction of 0.907. Second, the maximum area occupied by NPs was calculated to be 7.2×1013 nm2 from the total number of Ag/FeCo/Ag NPs (4.5×1011) and projected area of a single NP (143.14 nm2). Because the surface area of liposome is more than twice the maximum area occupied by NPs, it would appear that there is submonolayer coverage of NPs on the surface of the liposome. Even if it is assumed that all of the NPs are adsorbed onto the DPPC liposome, the surface coverage of NPs on the liposome is only 41% and the actual value should be much less than that. Next, the magnetic force acting on the four liposomes shown in Figure 5a was calculated assuming that these liposomes moved with constant velocity. If a liposome moves with constant speed, the magnetic force countervails the viscous drag force, fv, which can be estimated using the Stokes equation: $ = 6& ',

(5)

where r is the radius of a liposome, η = 1.114 mPa·s is the viscosity of the medium (0.1 M aqueous sucrose solution),37 and v is the constant velocity of the liposome. For example, if a liposome with a diameter of 10 µm moves with a constant velocity of 2.7 µm/s (see Figure 4), fv


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(and thus the magnetic force) is estimated to be 0.28 pN. By assuming the surface coverage of NPs on the liposome is 41% for all liposomes, one can calculate the number of NPs adsorbed on a single liposome. Then, the magnetic force acting on a single NP is found to be within the range of 0.2–0.7 aN by dividing the magnetic force acting on a single liposome by the number of NPs adsorbed on the liposome calculated for four different liposomes in Figure 5a. If one needs to separate intracellular vesicles with a diameter of 100 nm with an average velocity of 0.05 µm/s using a neodymium magnet, 60–210 Ag/FeCo/Ag NPs should be incorporated into (or adsorped onto) a single vesicle, which seems feasible. Conclusion Ag/FeCo/Ag core/shell/shell NPs were chemically synthesized that exhibit magnetic and plasmonic properties, giving magnetic separation and imaging capability. The surface of the NPs were modified with PLL-based polymer (PLL-COOH-SH) to make them water dispersible and biocompatible. The PLL-COOH-SH-modified Ag/FeCo/Ag NPs induced magnetic migration of a liposome by non-specific adsorption of the NPs onto the lipid membrane and application of a magnetic field. In addition, the NPs enable real-time monitoring of the motion of liposomes by plasmonic imaging. These NPs show potential for use as a magnetic probe to separate intracellular vesicles. Supporting Information. XRD pattern, STEM-HAADF image, EDS mapping image, and Ag 3d XPS spectrum of Ag/FeCo/Ag NPs, NMR spectrum and zeta potential of PLL-COOH-SH polymer, a brief discussion on cytotoxicity of NPs, and movies and snapshots of magnetophoretic motion of liposomes. This material is available free of charge via the Internet at http://pubs.acs.org.


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Corresponding Author: Shinya Maenosono, E-mail: [email protected] ACKNOWLEDGMENT: This work was supported by a Grant-in-Aid for Scientific Research, Grant Numbers 26600053 (SM), 26103516 (TH) and 25104510 (TH). ABBREVIATIONS NPs, nanoparticles; PM, plasma membranes; PLL, ε-Poly-L-lysine; DPPC, 1,2-Dipalmitoyl-snglycero-3-phosphocholine; LSPR, localized surface plasmon resonance. REFERENCES 1. Mellman, I. Endocytosis and Molecular Sorting. Annu. Rev. Cell Dev. Biol. 1996, 12, 575625 2. Maxfield, F. R.; McGraw, T. E. Endocytic Recycling. Nat. Rev. Mol. Cell Biol. 2004, 5, 121132 3. Anderson, R. G. W.; Goldstein, J. L.; Brown, M. S. A Mutation that Impairs the Ability of Lipoprotein Receptors to Localise in Coated Pits on the Cell Surface of Human Fibroblasts. Nature 1977, 270, 695-699 4. Garcia, C. K.; Wilund, K.; Arca, M.; Zuliani, G.; Fellin, R.; Maioli, M.; Calandra, S.; Bertolini, S.; Cossu, F.; Grishin, N.; Barnes, R.; Cohen, J. C.; Hobbs, H. H. Autosomal Recessive Hypercholesterolemia Caused by Mutations in a Putative LDL Receptor Adaptor Protein. Science 2001, 292, 1394-1398 5. Mellman, I.; Yarden, Y. Endocytosis and Cancer. Cold Spring Harb. Perspect. Biol. 2013, doi: 10.1101/cshperspect.a0169499


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6. Wittrup, A.; Zhang, S. H.; Svensson, K. J.; Kucharzewska, P.; Johansson, M. C.; Mörgelin, M.; Belting, M. Magnetic Nanoparticle-Based Isolation of Endocytic Vesicles Reveals a Role of the Heat Shock Protein GRP75 in Macromolecular Delivery. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 13342-13347 7. Nakamura, N.; Lill, J. R.; Phung, Q.; Jiang, Z.; Bakalarski, C.; de Mazière, A.; Klumperman, J.; Schlatter, M.; Delamarre, L.; Mellman, I. Endosomes are Specialized Platforms for Bacterial Sensing and NOD2 Signalling. Nature 2014, 509, 240-244 8. Tauro, B. J.; Greening, D. W.; Mathias, R. A.; Ji, H.; Mathivanan, S.; Scott, A. M.; Simpson, R. J. Comparison of Ultracentrifugation, Density Gradient Separation, and Immunoaffinity Capture Methods for Isolating Human Colon Cell Line LIM1863-Derived Exosomes. Methods 2012, 56, 293-304 9. Conner, S. D.; Schmid, S. L. Regulated Portals of Entry into the Cell. Nature 2003, 422, 3744 10. Wang, J.; Yu, X.; Boriskina, S. V.; Reinhard, B. M. Quantification of Differential ErbB1 and ErbB2 Cell surface Expression and Spatial Nanoclustering through Plasmon Coupling. Nano Lett. 2012, 12, 3231-3237 11. Wang, H.; Shen, J.; Li, Y.; Wei, Z.; Cao, G.; Gai, Z.; Hong, K.; Banerjee, P.; Zhou, S. Porous Carbon Protected Magnetite and Silver Hybrid Nanoparticles: Morphological Control, Recyclable Catalysts, and Multicolor Cell Imaging. ACS Appl. Mater. Interfaces 2013, 5, 9446-9453


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12. Lee, Y.; Garcia, M. A.; Huls N. A. F.; Sun, S. Synthetic Tuning of the Catalytic Properties of Au-Fe3O4 Nanoparticles. Angew. Chem. Int. Ed. 2010, 49, 1271-1274 13. Xu, Y. H.; Bai, J.; Wang, J.-P. High-Magnetic-Moment Multifunctional Nanoparticles for Nanomedicine Applications. J. Magn. Magn. Mater. 2007, 311, 131-134 14. Choi, J. S.; Jun, Y.-W.; Yeon, S.-I.; Kim, H. C.; Shin, J.-S.; Cheon, J. Biocompatible Heterostructured Nanoparticles for Multimodal Biological Detection. J. Am. Chem. Soc. 2006, 128, 15982-15983 15. Sachan, R.; Malasi, A.; Ge, J.; Yadavali, S.; Krishna, H.; Gangopadhyay, A.; Garcia, H.; Duscher, G.; Kalyanaraman, R. Ferroplasmons: Intense Localized Surface Plasmons in Metal-Ferromagnetic Nanoparticles. ACS Nano 2014, 8, 9790-9798 16. Dao, A. T. N.; Singh, P.; Shankar, C.; Mott, D.; Maenosono, S. Charge-Transfer-Induced Suppression of Galvanic Replacement and Synthesis of (Au@Ag)@Au Double Shell Nanoparticles for Highly Uniform, Robust and Sensitive Bioprobes. Appl. Phys. Lett. 2011, 99, 073107 17. Mott, D. M.; Dao, A. T. N.; Singh, P.; Shankar, C.; Maenosono, S. Electronic Transfer as a Route to Increase the Chemical Stability in Gold and Silver Core-Shell Nanoparticles. Adv. Colloid Interface Sci. 2012, 185-186, 14-33 18. Marsich, L.; Bonifacio, A.; Mandal, S.; Krol, S.; Beleites, C.; Sergo, V. Poly-L-lysine-Coated Silver Nanoparticles as Positively Charged Substrates for Surface-Enhanced Raman Scattering. Langmuir 2012, 28, 13166-13171


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19. Matsumura, K.; Hyon, S. H. Polyampholytes as Low Toxic Efficient Cryoprotective Agents with Antifreeze Protein Properties. Biomaterials 2009, 30, 4842-4849 20. Chaubey, G. S.; Barcena, C.; Poudyal, N.; Rong, C.; Gao, J.; Sun, S.; Liu, J. P. Synthesis and Stabilization of FeCo Nanoparticles. J. Am. Chem. Soc. 2007, 129, 7214-7215 21. Matsumura, K.; Hayashi, F.; Nagashima, T.; Hyon, S. H. Long-Term Cryopreservation of Juman Mesenchymal Stem Cells Using Carboxylated Poly-L-Lysine without the Addition of Proteins or Dimethyl Sulfoxide. J. Biomater. Sci.-Polym. Ed. 2013, 24, 1484-1497 22. Hamada, T.; Morita, M.; Miyakawa, M.; Sugimoto, R.; Hatanaka, A.; Vestergaard, M. C., Takagi, M. Size-Dependent Partitioning of Nano/Microparticles Mediated by Membrane Lateral Heterogeneity. J. Am. Chem. Soc. 2012, 134, 13990-13996 23. Baaziz, W.; Pichon, B. P.; Liu, Y.; Grenèche, J. M.; Ulhaq-Bouillet, C.; Terrier, E.; Bergeard, N.; Halté, V.; Boeglin, C.; Choueikani, F.; Toumi, M.; Mhiri, T.; Begin-Colin, S. Turning of Synthesis Conditions by Thermal Decomposition toward Core-Shell CoxFe1-xO@CoxFe3-yO4 and CoFe2O4 Nanoparticles with Spherical and Cubic Shapes. Chem. Mater. 2014, 26, 50635073 24. Singh, P.; Mott, D. M.; Maenosono, S. Gold/Wüstite Core-Shell Nanoparticles: Suppression of Iron Oxidation through the Electron-Transfer Phenomenon. ChemPhysChem 2013, 14, 3278-3283 25. Tufts, B. J.; Abrahams, I. L.; Caley, C. E.; Lunt, S. R.; Miskelly, G. M.; Sailor, M. J.; Santangelo, P. G.; Lewis, N. S.; Roe, A. L.; Hodgson, K. O. XPS and EXAFS Studies of the


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32. Lipowsky, R.; Döbereiner, H. G. Vesicles in Contact with Nanoparticles and Colloids. Europhys. Lett. 1998, 43, 219-225 33. Israelachvili, J. N. Intermolecular and Surface Forces; Academic Press: London, 2011 34. Pinchuk, A. O. Size-Dependent Hamaker Constant for Silver Nanoparticles. J. Phys. Chem. C 2012, 116, 20099-20102 35. Seifert, U.; Lipowsky, R. In Structure and Dynamics of Membranes; Lipowsky, R. Sackmann, E., Eds.; Elsevier: Amsterdam, 1995; p403. 36. Ramkaran, M.; Badia, A. Gel-to-Fluid Phase Transformation in Solid-Supported Phospholipid Bilayers Assembled by the Langmuir-Blodgett Technique: Effect of the Langmuir Monolayer Phase State and Molecular Density. J. Phys. Chem B 2014, 118, 97089721 37. CRC Handbook of Chemistry and Physics, 92nd ed.; Boca Raton, FL, 2011.

TOC Graphic and Synopsis Magnetic-plasmonic dual-functional Ag/FeCo/Ag core/shell/shell nanoparticles have great potential for various bioapplications including magnetic separation and plasmonic bioimaging.


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Magnetic-plasmonic dual-functional Ag/FeCo/Ag core/shell/shell nanoparticles have great potential for various bioapplications including magnetic separation and plasmonic bioimaging. 44x39mm (300 x 300 DPI)


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Figure 1. (a) STEM-HAADF image of a single Ag/FeCo/Ag NP. (b–e) EDS elemental mapping images of the single Ag/FeCo/Ag NP: (b) Ag L edge, (c) Fe K edge, (d) Co K edge, and (e) overlaid image. (f) The EDS line profile at the center of the NP indicated by a yellow line in (a–e). Blue, green and red lines correspond to Ag L, Co K and Fe K edge intensities, respectively. Dashed and solid lines represent raw and low-pass-filtered profiles, respectively. 127x83mm (300 x 300 DPI)


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Figure 2. (a,c) Fe 2p and (b,d) Co 2p core-level XPS for Ag/FeCo/Ag NPs (upper panels) and CoxFe1-xO NPs (lower panels). Black, yellow, blue and red curves represent raw data, Shirley background, deconvoluted peaks and the sum of all components, respectively. 113x85mm (300 x 300 DPI)


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Figure 3. (a) Magnetization curves of as-synthesized Ag/FeCo/Ag NPs measured at 5 (black) and 300 K (red). (b) FC (black) and ZFC (red) curves of as-synthesized Ag/FeCo/Ag NPs. (c) Magnetization curves of PLL-COOH-SH-modified Ag/FeCo/Ag NPs measured at 5 (black) and 300 K (red). (d) FC (black) and ZFC (red) curves of PLL-COOH-SH-modified Ag/FeCo/Ag NPs. (e) UV-vis spectrum of a hexane dispersion of assynthesized Ag/FeCo/Ag NPs. (f) LSPR peak absorbance plotted against NP concentration. 113x127mm (300 x 300 DPI)


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Langmuir

Figure 4. Snapshots of CLSM images: transmission (left), plasmon scattering (middle) and merged (right) images. The images were obtained at (a) t = 53 s, (b) t = 62 s, (c) t = 72 s, (d) t = 82 s, and (e) t = 92 s. Yellow arrows indicate the direction of magnetic force. 105x66mm (300 x 300 DPI)


Langmuir

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Figure 5. Time dependence of magnetophoretic velocity of (a) four different liposomes that moved with nearly constant speed, (b) a liposome whose motion suddenly accelerated at t = 40 s, (c) a liposome whose motion suddenly stopped at t = 136 s, and (d) two different liposomes that stuck together at t = 128 s and started to move as one. Note that the magnetophoretic velocity of a liposome could be obtained only when it was present within the fixed observation area. 127x92mm (300 x 300 DPI)


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Shell Magnetic Nanoparticles with Plasmonic

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