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Structural and morphological investigation of magnetic nanoparticles based on iron oxides for biomedical applications
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Materials Science and Engineering C 28 (2008) 489 – 494 www.elsevier.com/locate/msec
Structural and morphological investigation of magnetic nanoparticles based on iron oxides for biomedical applications Paula S. Haddad a,⁎, Tatiana M. Martins a,b , Lília D'Souza-Li c , Li M. Li d , Konradin Metze e , Randall L. Adam e , Marcelo Knobel b , Daniela Zanchet a a Laboratório Nacional de Luz Síncrotron (LNLS), Caixa Postal 6192, CEP 13083-970, Campinas-SP, Brazil Instituto de Física Gleb Wataghin (IFGW), Universidade Estadual de Campinas (UNICAMP), Caixa Postal 6165, CEP 13083-970, Campinas-SP, Brazil Laboratório de Endocrinologia Pediátrica da Faculdade de Ciências Médicas (FCM), UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil d Departamento de Neurologia da FCM, UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil e Grupo interdisciplinar “Patologia Analítica Celular”, Departamento de Anatomia Patológica da FCM, UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil b
c
Received 30 November 2006; accepted 4 April 2007 Available online 12 April 2007
Abstract The present work reports the synthesis, characterization and properties of magnetic iron oxide nanoparticles for biomedical applications, correlating the nanoscale tunabilities in terms of size, structure, and magnetism. Magnetic nanoparticles in different conditions were prepared through thermal decomposition of Fe(acac)3 in the presence of 1,2 hexadecanodiol (reducing agent) and oleic acid and oleylamine (ligands) in a hot organic solvent. The 2,3-dimercaptosuccinic acid (DMSA) was exchanged onto the nanocrystal surface making the particles stable in water. Nanoparticles were characterized by X-ray diffraction (XRD) measurements, small angle X-ray scattering (SAXS) and transmission electron microscopy (TEM). Preliminary tests of incorporation of these nanoparticles in cells and their magnetic resonance image (MRI) were also carried out. The magnetization characterizations were made by isothermal magnetic measurements. © 2007 Elsevier B.V. All rights reserved. Keywords: Magnetite; DMSA; Nanoparticles; MRI; SAXS; Biomedical applications
1. Introduction The increasing exploration of the nanotechnology in biological and medical applications has led to significant advances in diagnosis, prevention, and treatment of diseases. Superparamagnetic iron oxide nanoparticles (SPIO) currently have a surge of interest as their potential has been demonstrated in biomedical applications such as contrast agent in (MRI) [1–5], drug delivery [6] and therapy [7]. In this context, the minimization of nanoparticle polydispersity and heterogeneity will be essential for their fully exploitation and use in the construction of nanopharmaceuticals [8]. Organometallic precursor-based synthesis has proved successful for the preparation of
⁎ Corresponding author. E-mail address: [email protected] (P.S. Haddad). 0928-4931/$ - see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.msec.2007.04.014
uniform nanoparticles (NPs) [9–12]. This method has been widely used and consists in the decomposition of an organometallic precursor in organic solvents with high boiling points in the presence of capping ligands. Metal particles such as Fe and their oxides such as magnetite, hematite and other ferrites have been synthesized by this method [13]. The biocompatibility of these particles is a crucial step for many applications; therefore after the synthesis in organic solvent it is necessary to exchange the ligand to transfer the NPs from the organic to the aqueous phase [14]. Cell labeling with SPIO substances is an increasingly common method for in vivo cell separation as well as detection by MRI [15]. For example, Song et al. [16] evaluated the transport capabilities of particles into fetal rat neural stem cells. The transport efficiencies of these nanoparticles soluble in water were compared to that of the commercial particles such as Feridex–PLL system [17]. The results showed transport of
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SPIO covered with biocompatible ligand into the neural stem cells takes place efficiently. Considering interest in biomedical applications of SPIO particles, the present paper reports the synthesis and characterization of water soluble magnetic iron oxide NPs and first in vitro MRI experiments. The ultimate goal of this work is to optimize SPIO probes for MRI applications on stem cells therapy. It has been recently showed in the literature that SPIO particles with homogeneous size and anionic surface give better probes for MRI [17], but the applications and the full characterization including statistical size distribution and magnetic properties still have to be persuaded. Magnetic nanoparticles with three distinct sizes (5.2, 8.1 and 10.4 nm) were prepared in organic solvent. This approach renders a much better size distribution than the one typically obtained in waterbased synthesis, although it requires an additional step that is the exchange of the hydrophobic ligand by a hydrophilic one. In the present work, 2,3-dimercaptosuccinic acid (DMSA) was exchanged onto the nanocrystal surface. SPIO nanoparticles were characterized by X-ray diffraction (XRD), small angle Xray scattering (SAXS) and transmission electron microscopy (TEM). The magnetization analysis was carried out by isothermal magnetic measurements in applied field. Preliminary studies of incorporation into HeLa cells and MRI were also accomplished. 2. Experimental 2.1. Synthesis of Fe3O4 nanoparticles The synthesis was carried out using standard airless procedures and commercially available reagents. Magnetic Fe3O4 nanocrystals were synthesized through thermal decomposition of Fe(acac)3 (acac = acetylacetonate) in benzylether in the presence of oleic acid and oleylamine by following previously developed methods [18–19]. Fe3O4 nanoparticles of increasing sizes (5–10 nm range) were obtained (samples 1, 2 and 3). Sample 1 was obtained by heating the mixture containing the precursor with surfactants at 200 °C for 30 min and then, under nitrogen, heated to reflux (300 °C) for another 30 min. Sample 2 was prepared heating the same mixture at 200 °C for 2 h. Under nitrogen flux, the mixture was further heated to reflux (300 °C) and kept for 1 h. Sample 3 was obtained by heating the mixture containing the precursor with surfactants and seeds of sample 2 dispersed in 4 mL of hexane. This mixture was first heated at 100 °C for 30 min to remove hexane, then to 200 °C for 1 h and further was heated to reflux (300 °C) for 30 min. The particles can be easily purified by precipitation using ethanol and centrifuged (3000 rpm, 15 min) to remove the solvent, and redispersed in hydrophobic solvents. 2.2. Phase transfer of Fe3O4 nanocrystals from the organic to the aqueous phase Fe3O4 nanoparticles (∼ 10 mg) were dissolved in 1 ml of toluene while DMSA (∼ 10 mg) was dissolved in 1 ml of
dimethyl sulfoxide (DMSO). The solutions were mixed and vigorously stirred for 14 h producing a black powder that was isolated by centrifugation. The resulting nanoparticles were dried in vacuum and redispersed in 1 mL of water. The nanoparticles were stable in water for about 5 days. After this time the particles started to precipitate and could be redispersed by ultrasound. 2.3. Cell preparation and nanoparticle up-take HeLa cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 10% fetal bovine serum (FBS) and penicillin/streptomycin (complete medium). Cells were plated in 6 well plates and treatment was performed at 80–90% confluence in serum free medium with 50 μg/ml of iron oxide nanoparticles in water. A negative control without treatment was carried out. After 1, 3, 6 and 20 h, cells were harvested by scrapping them from the plate for optic microscopy staining and magnetic resonance. 2.4. Methods 2.4.1. XRD Powder XRD studies were realized in the XRD2 beamline at LNLS using 7100.2 eV photons (λ = 1.74619 Å) and θ–2θ geometry. The incident X-ray beam was monochromatized by a Si (111) double crystal Monochromator with sagittal focusing. Data were recorded from 2θ equal to 20° to 80° with 0.05° steps. 2.4.2. SAXS The SAXS experiments were performed at the SAXS beamline at LNLS, using X-ray wavelength of 1.7712 Å. The sample suspension (in toluene or water) was placed in sealed cell with mica windows specially designed for studies of liquid samples [20]. The sample detector distance (370 mm) was chosen to re4psenh cord the scattering intensities for q ¼ values ranging from k −1 −1 0.017 Å to 0.30 Å . The parasitic scattering was subtracted from the total SAXS intensity by measuring just the solvent. The data were analyzed by GNOM software [21]. 2.4.3. TEM Transmission electron microscopy of the as synthesized NPs (TEM, JEM-3010 operating at 300 kV, 1.7 Å point resolution) was carried out at the Laboratory of Electron Microscopy at LNLS, to observe the morphology of the NPs. The samples for observation were suspended in octane or water and stirred for 10 min. Then a drop of the supernatant dispersion was placed on an amorphous carbon film supported by a copper grid. 2.4.4. SQUID Magnetization measurements were carried out in a superconducting quantum interference device (SQUID) magnetometer; model MPMS XL7, from Quantum Design at 300 K, at IFGW, UNICAMP. The measurements were carried out on the dried powder, slightly pressed and conditioned in cylindrical holders of Lucite.
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2.4.5. MRI The MRI scan of labeled cells was performed using a 2T scanner (Elscint, Prestige, Haifa, Israel) at FCM, UNICAMP. It was used as a head coil and acquired two different pulse sequences. The parameters of gradient recall-echo T2 was: TR = 500 ms, TE = 10 ms, flip angle = 18, 2 number of excitations, matrix = 256 × 190; and the fast spin-echo T2 was: TR = 4800 ms, TE = 128 ms, flip angle = 170, 1 number of excitation, matrix = 256 × 256. Both sequences had field of view adjusted of the size of plaque and the thickness of slice was 3 mm. MRI experiments were carried out adding NPs dispersed in agarose and also cells before and after NPs incorporation. 2.4.6. Microscopic evaluation and image analysis Cell suspensions fixed in Karnovsky's solution and postfixed in OsO4 were incubated with uranyl acetate, dehydrated in alcohol and acetone and embedded in araldite. Ultrathin sections counterstained with lead acetate were examined by transmission electron microscopy [22]. A part of the cell suspensions was spread on glass slides, fixed in ethanol and stained by Perl's Prussian blue technique. The analysis was done by an examiner blinded to the study protocol. Cell segmentation was done interactively. Images were gray scale transformed using the complement of the blue channel (luminance level 255—blue luminance), with the levels of luminance ranging between 0 and 255 as described previously [23]. From each preparation bitmap images were taken from 100 non-overlapping randomly sampled cells (100 × oil immersion objective; 0.067 μm/pixel spatial resolutions). 3. Results and discussion Fig. 2. SAXS curves (A) before ligand exchange; (B) after ligand exchange.
Fig. 1 shows the XRD diffractograms of the three samples. The patterns were similar and the reflections were assigned to the magnetite phase, Fe3O4 (JCPDS 20-596). By using the Scherrer equation [24], the crystalline sizes of samples 1, 2 and 3 were estimated to be 5.2 nm, 8.1 nm and 10.4 nm respectively (Table 1). These results confirm that both temperature and time affect the particle formation [19]. Particles with smaller crystalline domains are obtained at lower temperatures and shorter heating times.
As synthesized, these NPs are coated with hydrophobic capping ligands and are insoluble in water. Therefore, it was necessary to use a multifunctional ligand system that allows one to transfer the NPs from the organic to the aqueous phase and also provides high biostability and a conjugation moiety for a targeting ligand. For this purpose, the 2,3-dimercaptosuccinic acid (DMSA) ligand was exchanged onto the NPs surface. The DMSA was chosen because it forms a stable coating through its carboxylic chelate (COOH) bonding and further stabilization of the ligand shells is attained through intermolecular disulfide cross-linkages between the ligands under ambient conditions [25]. Strategically the free thiol (SH) groups of DMSA ligand can be used for the attachment of target specific [26].
Table 1 Particle sizes (diameter) of samples 1, 2 and 3 obtained by XRD and SAXS before and after ligands exchange
Fig. 1. XRD patterns of the nanoparticles.
Sample 1 Sample 2 Sample 3
XRD (before) (nm)
SAXS (before) (nm)
SAXS (after) (nm)
5.2 8.1 10.4
5.5 8.3 10.9
5.7 8.5 Agglomeration
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Fig. 5. Incorporation curves of samples 1 and 2. A) Volume as a function of incorporation time; B) area as a function of incorporation time.
Fig. 3. Volume distribution as a function particle radius of sample 2: (A) before ligand exchange; (B) after ligand exchanged.
The size distribution for these systems before and after ligand exchange was evaluated by SAXS (Fig. 2A and B). The results showed evolution of size particles from sample 1 to sample 3, in agreement with the XRD data. The ligand exchange did not change substantially the particle size distribution in sample 1 and sample 2, but led to partial aggregation in sample 3.
Fig. 4. Incorporation of sample 2 in the HeLa cells.
The SAXS curves were fitted using the GNOM software, considering a polydisperse system of hard spheres. For a polydisperse system of n spheres of different sizes, the scattered intensity is given by [21]: I ð qÞ ¼
X
U2 ðqRn ÞDðRn Þ
ð1Þ
n
where Φ2 is the scattering amplitude, D(Rn) = (4/3)πRn3N(Rn) is the volume distribution function and N(Rn) is the number of particles with a given radius Rn. The quantitative results are
Fig. 6. Magnetic measurements of samples 1, 2 and 3 before ligand exchange.
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Fig. 7. MRI tests of sample 2: (a) HeLa cells in agarose; (b) NPs in agarose; (c) HeLa cells with incorporated NPs; (d) and (e) agarose background.
presented in Table 1. The volume distribution of sample 1 before and after ligand exchange is showed in Fig. 3A and B, respectively. It shows that a fraction of the smaller particles is lost during the ligand exchange procedure (probably in the centrifugation step), leading to a slightly larger mean size and narrower size distribution. Sample 2 was used in preliminary tests of incorporation in HeLa cells to evaluate the incorporation of the NPs as a ferrofluid (Fig. 4). The Hela cells are an immortal cell line used in medical research. The cell line was derived from cervical cancer cells [24]. The arrows in Fig. 4 show the incorporation of aggregated NPs in a HeLa cells. The Fig. 5 shows the incorporation curves of samples 1 and 2. The curves were obtained by Prussian blue technique, where the Fe (III) reacts with a composition of 2% HCl acid mixed and 2% K[Fe(CN)6] producing Fe4[Fe(CN)6]3, the so called Prussian blue, which is an insoluble blue reaction product deposited inside the cell at the location of the Fe (III) ions. The blue stained area/cell (in pixels) was determined by thresholding. Since the intensity of the staining reaction depends on the concentration of Fe (III) containing nanoparticles, it was calculated the volume (in arbitrary units) of the stained regions using the luminance (minus the threshold level) of each pixel as the height of a z-axis, as an estimate of the number of nanoparticles phagocytized. For each incubation time we compared area and volume between the two particle types by the Mann-Whitney test. The significance level was defined as 5%. Area and volume for all incubation times were showed in Fig. 5A and B. It can be seen that the major nanoparticle up-take by the cells occurs in the first 6 h of incubation, for both samples. The magnetization analysis was carried out through isothermal magnetic measurements in applied field at room temperature (Fig. 6). The particles presented a superparamagnetic behavior, with a coercive field from 11 to 13 Oe, approximately. As it can be seen through the saturation magnetization values, an important increase of the effective magnetic moment is shown from sample 1 to sample 3 , in agreement with the increasing size. Using a clinical MR scanner and routine built-in pulse sequences, the superparamagnetic iron oxide NPs yielded a
strong signal decrease in T2 relaxation time (Fig. 7b). The effect of field distortion was intense going outer-limits of the well. This was best seen in gradient recall-echo, whereas the well below it showed a signal loss at its upper quadrant. The two pulse sequences (spin-echo and gradient recall-echo) images depicted scattered dots of hypointense signal in the well with labeled cells suspended in agarose (Fig. 7c). This was clearer in gradient recall-echo image set than the spin-echo image. The control wells (Fig. 7a, d and e) showed no similar pattern of signal loss or reduction. Adjustments in T2-weighted pulse sequence in particular gradient recall-echo and combination with post-processing imaging techniques (texture analysis) can further improve detection of labeled cells with these particles using clinical MR scanners. 4. Conclusions We present the results on the synthesis, characterization and properties of magnetic iron oxide nanoparticles for biomedical applications. Three samples were prepared by varying the synthesis conditions. The XRD showed that the crystalline domains increase with temperature and synthesis time, but the mean size is not deeply affected. The ligand exchange produces water soluble particles. The magnetization analyses showed that the particles are superparamagnetic at room temperature. Preliminary tests of incorporation in HeLa and MRI were successful. Acknowledgments P.S. Haddad and T.M. Martins thank the financial support from CNPq. The authors thank the Laboratory of Electron Microscopy and XRD2 beamline of LNLS, the Laboratory of Materials and Low Temperatures of IFGW, FCM-UNICAMP, and FAPESP. References [1] J.W. Bulte, D.L. Kraitchman, Curr. Pharm. Biotechnol. 5 (2004) 567. [2] E.J. Delikatny, H. Poptani, Radiol. Clin. North Am. 43 (2005) 205.
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[3] G.M. Lanza, P.M. Winter, S.D. Caruthers, A.M. Morawski, A.H. Schmieder, K.C. Crowder, J. Nucl. Cardiol. 11 (2004) 733. [4] J. Vymazal, J.W. Bulte, J.A. Frank, G. Di Chiro, R.A. Brooks, J. Magn. Reson. Imaging 3 (1993) 637. [5] D.J.L. Hogemann, R. Weissleder, J.P. Basilion, Bioconjug. Chem. 11 (2000) 941. [6] V.G. Roullin, J.R. Deverre, L. Lemarie, F. Hindré, M.C.V. Julienne, R. Vienet, J.P. Benoit, Eur. J. Pharm. Biopharm. 53 (2002) 293. [7] M. Shinkai, M. Yanase, M. Suzuki, H. Honda, T. Wakabayashi, J. Yoshida, T. Kobayashi, J. Magn.Magn. Mater. 194 (1999) 44. [8] S. Mornet, S. Vasseur, F. Grasset, E. Duguet, J. Mater. Chem. 14 (2004) 2161. [9] C.B. Murray, C.R. Kagan, M.G. Bawendi, Annu. Rev. Mater. Sci. 30 (2000) 545. [10] X. Wang, J. Zhuang, Q. Peng, Y.D. Li, Nature 437 (2005) 121. [11] T. Hyeon, Chem. Commun. 3 (2003) 927. [12] M. Brust, C.J. Kiely, Colloids Surf. 202 (2002) 175. [13] J. Park, E. Kang, S.U. Son, H.M. Park, M.K. Lee, J. Kim, K.W. Kim, H.J. Noh, J.H. Park, C.J. Bae, J.G. Park, T. Hyeon, Adv. Mater. 17 (2005) 429. [14] Y.W. Jun, Y.M. Huh, J.S. Choi, J.H. Lee, H.T. Song, S. Kim, S. Yoon, K.S. Kim, J.S. Shin, J.S. Suh, J. Cheon, J. Am. Chem. Soc. 127 (2005) 5732.
[15] A.K. Gupta, M. Gupta, Biomaterials 26 (2005) 3995. [16] H.T. Song, J.S. Choi, Y.M. Huh, S. Kim, Y.W. Jun, J.S. Suh, J. Cheon, J. Am. Chem. Soc. 127 (2005) 9992. [17] A.S. Arbab, L.A. Bashaw, B.R. Miller, E.K. Jordan, J.W.M. Bulte, J.A. Frank, Transplantation 76 (2003) 1123. [18] S. Sun, H. Zeng, J. Am. Chem. Soc. 124 (2002) 8204. [19] S. Sun, H. Zeng, D.B. Robinson, S. Raoux, P.M. Rice, S.X. Wang, G. Li, J. Am. Chem. Soc. 126 (2004) 273. [20] G. Kellermann, F. Vicentin, E. Tamura, M. Rocha, H. Tolentino, A. Barbosa, A. Craievich, I. Torriani, J. Appl. Cryst. 30 (1997) 880. [21] D.I. Svergun, J. Appl. Cryst. 25 (1992) 495. [22] K. Metze, L.A. Andrade, Pathol. Res Pract. 187 (1991) 1031. [23] M.R. Alborghetti, M.E. Correa, R.L. Adam, K. Metze, F.L. Coracin, C.A. Souza, M.L. Cintra, J. Oral Pathol. Med. 34 (2005) 486. [24] H.P. Klug, L.E. Alexander, X-ray Diffraction Procedures for Polycrystalline and Amorphous Materials, John Wiley & Sons, New York, 1974, p. 1. [25] N. Fauconnier, J.N. Pons, J. Roger, A.J. Bee, J. Colloid Interface Sci. 194 (1997) 427. [26] D.R. Smith, D. Woolard, M.L. Luck, N.K. Laughlin, Toxicol. Appl. Pharm. 166 (2000) 230.
Materials Science and Engineering C 28 (2008) 489 – 494 www.elsevier.com/locate/msec
Structural and morphological investigation of magnetic nanoparticles based on iron oxides for biomedical applications Paula S. Haddad a,⁎, Tatiana M. Martins a,b , Lília D'Souza-Li c , Li M. Li d , Konradin Metze e , Randall L. Adam e , Marcelo Knobel b , Daniela Zanchet a a Laboratório Nacional de Luz Síncrotron (LNLS), Caixa Postal 6192, CEP 13083-970, Campinas-SP, Brazil Instituto de Física Gleb Wataghin (IFGW), Universidade Estadual de Campinas (UNICAMP), Caixa Postal 6165, CEP 13083-970, Campinas-SP, Brazil Laboratório de Endocrinologia Pediátrica da Faculdade de Ciências Médicas (FCM), UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil d Departamento de Neurologia da FCM, UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil e Grupo interdisciplinar “Patologia Analítica Celular”, Departamento de Anatomia Patológica da FCM, UNICAMP, Caixa Postal 6111, CEP 13083-970, Campinas-SP, Brazil b
c
Received 30 November 2006; accepted 4 April 2007 Available online 12 April 2007
Abstract The present work reports the synthesis, characterization and properties of magnetic iron oxide nanoparticles for biomedical applications, correlating the nanoscale tunabilities in terms of size, structure, and magnetism. Magnetic nanoparticles in different conditions were prepared through thermal decomposition of Fe(acac)3 in the presence of 1,2 hexadecanodiol (reducing agent) and oleic acid and oleylamine (ligands) in a hot organic solvent. The 2,3-dimercaptosuccinic acid (DMSA) was exchanged onto the nanocrystal surface making the particles stable in water. Nanoparticles were characterized by X-ray diffraction (XRD) measurements, small angle X-ray scattering (SAXS) and transmission electron microscopy (TEM). Preliminary tests of incorporation of these nanoparticles in cells and their magnetic resonance image (MRI) were also carried out. The magnetization characterizations were made by isothermal magnetic measurements. © 2007 Elsevier B.V. All rights reserved. Keywords: Magnetite; DMSA; Nanoparticles; MRI; SAXS; Biomedical applications
1. Introduction The increasing exploration of the nanotechnology in biological and medical applications has led to significant advances in diagnosis, prevention, and treatment of diseases. Superparamagnetic iron oxide nanoparticles (SPIO) currently have a surge of interest as their potential has been demonstrated in biomedical applications such as contrast agent in (MRI) [1–5], drug delivery [6] and therapy [7]. In this context, the minimization of nanoparticle polydispersity and heterogeneity will be essential for their fully exploitation and use in the construction of nanopharmaceuticals [8]. Organometallic precursor-based synthesis has proved successful for the preparation of
⁎ Corresponding author. E-mail address: [email protected] (P.S. Haddad). 0928-4931/$ - see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.msec.2007.04.014
uniform nanoparticles (NPs) [9–12]. This method has been widely used and consists in the decomposition of an organometallic precursor in organic solvents with high boiling points in the presence of capping ligands. Metal particles such as Fe and their oxides such as magnetite, hematite and other ferrites have been synthesized by this method [13]. The biocompatibility of these particles is a crucial step for many applications; therefore after the synthesis in organic solvent it is necessary to exchange the ligand to transfer the NPs from the organic to the aqueous phase [14]. Cell labeling with SPIO substances is an increasingly common method for in vivo cell separation as well as detection by MRI [15]. For example, Song et al. [16] evaluated the transport capabilities of particles into fetal rat neural stem cells. The transport efficiencies of these nanoparticles soluble in water were compared to that of the commercial particles such as Feridex–PLL system [17]. The results showed transport of
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SPIO covered with biocompatible ligand into the neural stem cells takes place efficiently. Considering interest in biomedical applications of SPIO particles, the present paper reports the synthesis and characterization of water soluble magnetic iron oxide NPs and first in vitro MRI experiments. The ultimate goal of this work is to optimize SPIO probes for MRI applications on stem cells therapy. It has been recently showed in the literature that SPIO particles with homogeneous size and anionic surface give better probes for MRI [17], but the applications and the full characterization including statistical size distribution and magnetic properties still have to be persuaded. Magnetic nanoparticles with three distinct sizes (5.2, 8.1 and 10.4 nm) were prepared in organic solvent. This approach renders a much better size distribution than the one typically obtained in waterbased synthesis, although it requires an additional step that is the exchange of the hydrophobic ligand by a hydrophilic one. In the present work, 2,3-dimercaptosuccinic acid (DMSA) was exchanged onto the nanocrystal surface. SPIO nanoparticles were characterized by X-ray diffraction (XRD), small angle Xray scattering (SAXS) and transmission electron microscopy (TEM). The magnetization analysis was carried out by isothermal magnetic measurements in applied field. Preliminary studies of incorporation into HeLa cells and MRI were also accomplished. 2. Experimental 2.1. Synthesis of Fe3O4 nanoparticles The synthesis was carried out using standard airless procedures and commercially available reagents. Magnetic Fe3O4 nanocrystals were synthesized through thermal decomposition of Fe(acac)3 (acac = acetylacetonate) in benzylether in the presence of oleic acid and oleylamine by following previously developed methods [18–19]. Fe3O4 nanoparticles of increasing sizes (5–10 nm range) were obtained (samples 1, 2 and 3). Sample 1 was obtained by heating the mixture containing the precursor with surfactants at 200 °C for 30 min and then, under nitrogen, heated to reflux (300 °C) for another 30 min. Sample 2 was prepared heating the same mixture at 200 °C for 2 h. Under nitrogen flux, the mixture was further heated to reflux (300 °C) and kept for 1 h. Sample 3 was obtained by heating the mixture containing the precursor with surfactants and seeds of sample 2 dispersed in 4 mL of hexane. This mixture was first heated at 100 °C for 30 min to remove hexane, then to 200 °C for 1 h and further was heated to reflux (300 °C) for 30 min. The particles can be easily purified by precipitation using ethanol and centrifuged (3000 rpm, 15 min) to remove the solvent, and redispersed in hydrophobic solvents. 2.2. Phase transfer of Fe3O4 nanocrystals from the organic to the aqueous phase Fe3O4 nanoparticles (∼ 10 mg) were dissolved in 1 ml of toluene while DMSA (∼ 10 mg) was dissolved in 1 ml of
dimethyl sulfoxide (DMSO). The solutions were mixed and vigorously stirred for 14 h producing a black powder that was isolated by centrifugation. The resulting nanoparticles were dried in vacuum and redispersed in 1 mL of water. The nanoparticles were stable in water for about 5 days. After this time the particles started to precipitate and could be redispersed by ultrasound. 2.3. Cell preparation and nanoparticle up-take HeLa cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 10% fetal bovine serum (FBS) and penicillin/streptomycin (complete medium). Cells were plated in 6 well plates and treatment was performed at 80–90% confluence in serum free medium with 50 μg/ml of iron oxide nanoparticles in water. A negative control without treatment was carried out. After 1, 3, 6 and 20 h, cells were harvested by scrapping them from the plate for optic microscopy staining and magnetic resonance. 2.4. Methods 2.4.1. XRD Powder XRD studies were realized in the XRD2 beamline at LNLS using 7100.2 eV photons (λ = 1.74619 Å) and θ–2θ geometry. The incident X-ray beam was monochromatized by a Si (111) double crystal Monochromator with sagittal focusing. Data were recorded from 2θ equal to 20° to 80° with 0.05° steps. 2.4.2. SAXS The SAXS experiments were performed at the SAXS beamline at LNLS, using X-ray wavelength of 1.7712 Å. The sample suspension (in toluene or water) was placed in sealed cell with mica windows specially designed for studies of liquid samples [20]. The sample detector distance (370 mm) was chosen to re4psenh cord the scattering intensities for q ¼ values ranging from k −1 −1 0.017 Å to 0.30 Å . The parasitic scattering was subtracted from the total SAXS intensity by measuring just the solvent. The data were analyzed by GNOM software [21]. 2.4.3. TEM Transmission electron microscopy of the as synthesized NPs (TEM, JEM-3010 operating at 300 kV, 1.7 Å point resolution) was carried out at the Laboratory of Electron Microscopy at LNLS, to observe the morphology of the NPs. The samples for observation were suspended in octane or water and stirred for 10 min. Then a drop of the supernatant dispersion was placed on an amorphous carbon film supported by a copper grid. 2.4.4. SQUID Magnetization measurements were carried out in a superconducting quantum interference device (SQUID) magnetometer; model MPMS XL7, from Quantum Design at 300 K, at IFGW, UNICAMP. The measurements were carried out on the dried powder, slightly pressed and conditioned in cylindrical holders of Lucite.
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2.4.5. MRI The MRI scan of labeled cells was performed using a 2T scanner (Elscint, Prestige, Haifa, Israel) at FCM, UNICAMP. It was used as a head coil and acquired two different pulse sequences. The parameters of gradient recall-echo T2 was: TR = 500 ms, TE = 10 ms, flip angle = 18, 2 number of excitations, matrix = 256 × 190; and the fast spin-echo T2 was: TR = 4800 ms, TE = 128 ms, flip angle = 170, 1 number of excitation, matrix = 256 × 256. Both sequences had field of view adjusted of the size of plaque and the thickness of slice was 3 mm. MRI experiments were carried out adding NPs dispersed in agarose and also cells before and after NPs incorporation. 2.4.6. Microscopic evaluation and image analysis Cell suspensions fixed in Karnovsky's solution and postfixed in OsO4 were incubated with uranyl acetate, dehydrated in alcohol and acetone and embedded in araldite. Ultrathin sections counterstained with lead acetate were examined by transmission electron microscopy [22]. A part of the cell suspensions was spread on glass slides, fixed in ethanol and stained by Perl's Prussian blue technique. The analysis was done by an examiner blinded to the study protocol. Cell segmentation was done interactively. Images were gray scale transformed using the complement of the blue channel (luminance level 255—blue luminance), with the levels of luminance ranging between 0 and 255 as described previously [23]. From each preparation bitmap images were taken from 100 non-overlapping randomly sampled cells (100 × oil immersion objective; 0.067 μm/pixel spatial resolutions). 3. Results and discussion Fig. 2. SAXS curves (A) before ligand exchange; (B) after ligand exchange.
Fig. 1 shows the XRD diffractograms of the three samples. The patterns were similar and the reflections were assigned to the magnetite phase, Fe3O4 (JCPDS 20-596). By using the Scherrer equation [24], the crystalline sizes of samples 1, 2 and 3 were estimated to be 5.2 nm, 8.1 nm and 10.4 nm respectively (Table 1). These results confirm that both temperature and time affect the particle formation [19]. Particles with smaller crystalline domains are obtained at lower temperatures and shorter heating times.
As synthesized, these NPs are coated with hydrophobic capping ligands and are insoluble in water. Therefore, it was necessary to use a multifunctional ligand system that allows one to transfer the NPs from the organic to the aqueous phase and also provides high biostability and a conjugation moiety for a targeting ligand. For this purpose, the 2,3-dimercaptosuccinic acid (DMSA) ligand was exchanged onto the NPs surface. The DMSA was chosen because it forms a stable coating through its carboxylic chelate (COOH) bonding and further stabilization of the ligand shells is attained through intermolecular disulfide cross-linkages between the ligands under ambient conditions [25]. Strategically the free thiol (SH) groups of DMSA ligand can be used for the attachment of target specific [26].
Table 1 Particle sizes (diameter) of samples 1, 2 and 3 obtained by XRD and SAXS before and after ligands exchange
Fig. 1. XRD patterns of the nanoparticles.
Sample 1 Sample 2 Sample 3
XRD (before) (nm)
SAXS (before) (nm)
SAXS (after) (nm)
5.2 8.1 10.4
5.5 8.3 10.9
5.7 8.5 Agglomeration
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Fig. 5. Incorporation curves of samples 1 and 2. A) Volume as a function of incorporation time; B) area as a function of incorporation time.
Fig. 3. Volume distribution as a function particle radius of sample 2: (A) before ligand exchange; (B) after ligand exchanged.
The size distribution for these systems before and after ligand exchange was evaluated by SAXS (Fig. 2A and B). The results showed evolution of size particles from sample 1 to sample 3, in agreement with the XRD data. The ligand exchange did not change substantially the particle size distribution in sample 1 and sample 2, but led to partial aggregation in sample 3.
Fig. 4. Incorporation of sample 2 in the HeLa cells.
The SAXS curves were fitted using the GNOM software, considering a polydisperse system of hard spheres. For a polydisperse system of n spheres of different sizes, the scattered intensity is given by [21]: I ð qÞ ¼
X
U2 ðqRn ÞDðRn Þ
ð1Þ
n
where Φ2 is the scattering amplitude, D(Rn) = (4/3)πRn3N(Rn) is the volume distribution function and N(Rn) is the number of particles with a given radius Rn. The quantitative results are
Fig. 6. Magnetic measurements of samples 1, 2 and 3 before ligand exchange.
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Fig. 7. MRI tests of sample 2: (a) HeLa cells in agarose; (b) NPs in agarose; (c) HeLa cells with incorporated NPs; (d) and (e) agarose background.
presented in Table 1. The volume distribution of sample 1 before and after ligand exchange is showed in Fig. 3A and B, respectively. It shows that a fraction of the smaller particles is lost during the ligand exchange procedure (probably in the centrifugation step), leading to a slightly larger mean size and narrower size distribution. Sample 2 was used in preliminary tests of incorporation in HeLa cells to evaluate the incorporation of the NPs as a ferrofluid (Fig. 4). The Hela cells are an immortal cell line used in medical research. The cell line was derived from cervical cancer cells [24]. The arrows in Fig. 4 show the incorporation of aggregated NPs in a HeLa cells. The Fig. 5 shows the incorporation curves of samples 1 and 2. The curves were obtained by Prussian blue technique, where the Fe (III) reacts with a composition of 2% HCl acid mixed and 2% K[Fe(CN)6] producing Fe4[Fe(CN)6]3, the so called Prussian blue, which is an insoluble blue reaction product deposited inside the cell at the location of the Fe (III) ions. The blue stained area/cell (in pixels) was determined by thresholding. Since the intensity of the staining reaction depends on the concentration of Fe (III) containing nanoparticles, it was calculated the volume (in arbitrary units) of the stained regions using the luminance (minus the threshold level) of each pixel as the height of a z-axis, as an estimate of the number of nanoparticles phagocytized. For each incubation time we compared area and volume between the two particle types by the Mann-Whitney test. The significance level was defined as 5%. Area and volume for all incubation times were showed in Fig. 5A and B. It can be seen that the major nanoparticle up-take by the cells occurs in the first 6 h of incubation, for both samples. The magnetization analysis was carried out through isothermal magnetic measurements in applied field at room temperature (Fig. 6). The particles presented a superparamagnetic behavior, with a coercive field from 11 to 13 Oe, approximately. As it can be seen through the saturation magnetization values, an important increase of the effective magnetic moment is shown from sample 1 to sample 3 , in agreement with the increasing size. Using a clinical MR scanner and routine built-in pulse sequences, the superparamagnetic iron oxide NPs yielded a
strong signal decrease in T2 relaxation time (Fig. 7b). The effect of field distortion was intense going outer-limits of the well. This was best seen in gradient recall-echo, whereas the well below it showed a signal loss at its upper quadrant. The two pulse sequences (spin-echo and gradient recall-echo) images depicted scattered dots of hypointense signal in the well with labeled cells suspended in agarose (Fig. 7c). This was clearer in gradient recall-echo image set than the spin-echo image. The control wells (Fig. 7a, d and e) showed no similar pattern of signal loss or reduction. Adjustments in T2-weighted pulse sequence in particular gradient recall-echo and combination with post-processing imaging techniques (texture analysis) can further improve detection of labeled cells with these particles using clinical MR scanners. 4. Conclusions We present the results on the synthesis, characterization and properties of magnetic iron oxide nanoparticles for biomedical applications. Three samples were prepared by varying the synthesis conditions. The XRD showed that the crystalline domains increase with temperature and synthesis time, but the mean size is not deeply affected. The ligand exchange produces water soluble particles. The magnetization analyses showed that the particles are superparamagnetic at room temperature. Preliminary tests of incorporation in HeLa and MRI were successful. Acknowledgments P.S. Haddad and T.M. Martins thank the financial support from CNPq. The authors thank the Laboratory of Electron Microscopy and XRD2 beamline of LNLS, the Laboratory of Materials and Low Temperatures of IFGW, FCM-UNICAMP, and FAPESP. References [1] J.W. Bulte, D.L. Kraitchman, Curr. Pharm. Biotechnol. 5 (2004) 567. [2] E.J. Delikatny, H. Poptani, Radiol. Clin. North Am. 43 (2005) 205.
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