Poly(acrylic acid) microspheres loaded with superparamagnetic iron oxide nanoparticles for transcatheter arterial embolization and MRI detectability: In vitro and in vivo evaluation

Accepted Manuscript Title: Poly(acrylic acid) microspheres loaded with superparamagnetic iron oxide nanoparticles for transcatheter arterial embolization and MRI detectability: in vitro and in vivo evaluation Authors: Zi-Yuan Li, Xiao-Ya Qin, Li-Ying Guo, Huan Wang, Xiao-Xin Liu, Zhuo-Zhao Zheng, Hai-Tao Guan, Li Song, Ying-Hua Zou, Tian-Yuan Fan PII: DOI: Reference:

S0378-5173(17)30379-4 http://dx.doi.org/doi:10.1016/j.ijpharm.2017.04.069 IJP 16635

To appear in:

International Journal of Pharmaceutics

Received date: Revised date: Accepted date:

7-2-2017 17-4-2017 28-4-2017

Please cite this article as: Li, Zi-Yuan, Qin, Xiao-Ya, Guo, Li-Ying, Wang, Huan, Liu, Xiao-Xin, Zheng, Zhuo-Zhao, Guan, Hai-Tao, Song, Li, Zou, Ying-Hua, Fan, Tian-Yuan, Poly(acrylic acid) microspheres loaded with superparamagnetic iron oxide nanoparticles for transcatheter arterial embolization and MRI detectability: in vitro and in vivo evaluation.International Journal of Pharmaceutics http://dx.doi.org/10.1016/j.ijpharm.2017.04.069 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Poly (acrylic acid) microspheres loaded with superparamagnetic iron oxide nanoparticles for transcatheter arterial embolization and MRI detectability: in vitro and in vivo evaluation

Zi-Yuan Lia,b, Xiao-Ya Qina,b, Li-Ying Guoa,b, Huan Wanga,b, Xiao-Xin Liua,b, Zhuo-Zhao Zhengc, HaiTao Guand, Li Songd, Ying-Hua Zoud, Tian-Yuan Fana.b*

a

The State Key Laboratory of Natural and Biomimetic Drugs, School of Pharmaceutical Sciences,

Peking University, Beijing 100191, China b

Beijing Key Laboratory of Molecular Pharmaceutics and New Drug Delivery Systems, School of

Pharmaceutical Sciences, Peking University, Beijing 100191, China c

Department of Nuclear Medicine, Beijing Tsinghua Changgung Hospital, Beijing 100044, China

d

Department of Interventional Radiology and Vascular Surgery, Peking University First Hospital,

Beijing 100034, China

*

Corresponding author at: Department of Pharmaceutics, Health Science Center, Peking University, 38

Xueyuan Road, Haidian District, Beijing 100191, People’s Republic of China. E-mail address: [email protected].

Graphical abstract

Abstract To develop embolic microspheres with MRI detectability, superparamagnetic iron oxide nanoparticles (SPIONs) were synthesized and mixed with monomer of acrylic acid to prepare SPIONsloaded polymerized microspheres (SPMs) by inverse suspension polymerization method. The SPMs were evaluated for the ability of embolization by investigating the morphology, particle size, elasticity and renal arterial embolization to rabbits. Meanwhile, the loading of SPIONs was verified by optical microscope, transmission electron microscope, Fourier transform infrared spectrum, vibrating sample magnetometer, X-ray diffraction and X-ray photoelectron spectroscopy, and the content of SPIONs in SPMs was measured quantitatively. Furthermore, the MRI detectability of SPMs was testified in gel phantom, mice and rabbits respectively by a clinical 3.0 Tesla MRI scanner. The results revealed the SPMs were potential MRI detectable embolic microspheres for improving the effectiveness and safety of embolotherapy in the future.

Keywords: Hydrophilic polymer microspheres; Superparamagnetic iron oxide nanoparticles; Magnetic resonance imaging; Embolization

1. Introduction

By blocking the selected blood vessel with embolic materials under the guidance of image devices (usually digital subtraction angiography, DSA) (Poursaid et al., 2016), interventional embolization has played a significant role in treating malignancy (Tandberg et al., 2012), uterine fibroids (Khan et al., 2014), hemangioma (Zielinski et al., 2016), vascular malformation (Kolber et al., 2015; Van Beijnum et al., 2011) and hemostasis (Simon et al., 2010) for several decades (Corr 2005; Osuga et al., 2002; Spies et al., 2004). A variety of solid embolic agents have been developed and applied in clinical embolotherapy, among which hydrophilic polymer microspheres have emerged as a popular device in recent years, including Embospheres, DC bead, HepaSphere and so on. However, most of the embolic agents cannot be detected by DSA (Duran et al., 2016) and their locations have to be judged indirectly by monitoring X-ray angiography with the assistance of contrast agent injected before, during and after the process of embolization (Loffroy et al., 2013; Wang et al., 2013). Meanwhile, some inadequacies of X-ray

angiography have been revealed, such as the exposure of doctors and patients to X-ray, the restricted use in special patient groups like children and pregnant women, allergic effects of using iodinated contrast agents for some sensitive patients, etc. In addition, the ability of standard angiography to distinguish the pathological tissues from normal tissues is considered limited, though it is used to find the hypervascular target tissue (Fidelman et al., 2002; Kahn and Busse, 2012; Kok et al., 1996). Great efforts have been made to monitor the position of embolic agents directly so as to improve the efficacy and safety of embolization (Fidelman et al., 2002). Nowadays, promising progresses have been achieved in both imaging equipment and embolic agents. The combination of MRI equipment and MRI detectable microspheres would help radiologists to monitor the embolic procedure directly (Kim et al., 2016), estimate the occurrence of incomplete or ectopic embolization and examine the embolic effects during and after the operation (Cilliers et al., 2008). Magnetic resonance imaging (MRI) is acknowledged to have multiple advantages of multidirectional image, high tissue resolution and no ionizing radiation (Fritz et al., 2012; GonzálezVillà et al., 2016; Kahn and Busse, 2012), which has emerged in a variety of medical navigations, including neurosurgical procedures, cardiac catheterization and high intensity focused ultrasound. It has also been introduced in the guidance and surveillance of embolization by using X-ray angiography-magnetic resonance combined technology (XMR) or even MRI alone instead of DSA (Fahrig et al., 2001; Kos et al., 2008; Lillaney et al., 2016; Shin et al., 2013). MRI detectable embolic agents have also attracted attention. The earlier preparation of MRI detectable embolic microspheres found in literature was immersing the commercially available Embospheres in Omniscan, a kind of gadolinium-containing MRI contrast agent (Wilson et al., 2003). Thereafter, with superparamagnetic iron oxide nanoparticles (SPIONs) being used as MRI contrast agents in clinical practice (Kiani et al., 2016; Zhao et al., 2014), microspheres loaded with SPIONs have become the mainstream of the studies on MRI detectable embolic microspheres. From 2005, SPIONs were successively mixed with chitosan (Choi et al., 2015; Chung et al., 2012; Kang et al., 2009; Kim et al., 2007; Lee et al., 2005), alginates (Kim et al., 2014; Kim et al., 2015; Kim et al., 2016) and poly (lactide-co-glycolide) to prepare MRI detectable embolic microspheres through various synthetic technologies for microspheres. SPIONs could also be loaded on 2-methacryloyloxyethyl (2,3,5-triiodobenzoate) copolymers by in situ precipitation method (Bartling et al., 2011). Poly (acrylic acid) microspheres (PMs), belonging to the popular hydrophilic polymer

microspheres, were prepared and characterized in our previous study, presenting ideal embolic microspheres with good morphology, size distribution, elasticity, catheter deliverability, cytocompatibility and drug loading capacity for potential clinical application (Cui et al., 2012). After that, we loaded SPIONs on the PMs by in situ precipitation method and obtained MRI detectable microspheres (Wang et al., 2016 b). In this study, we developed another method to prepare MRI detectable embolic microspheres. SPIONs were synthesized and characterized first, then mixed with the monomer of acrylic acid to prepare SPIONs-loaded poly (acrylic acid) (SPMs) by inverse suspension polymerization method. The SPMs were characterized comprehensively

for

the

morphological,

physical, chemical,

embotherapeutic and MRI detectable properties in vitro. Thereafter, transcatheter arterial embolization for rabbits’ kidney was performed to investigate the embolic ability and long-term MRI detectability of SPMs. In addition, the SPMs were compared and discussed with the magnetic microspheres prepared in our previous work by in situ precipitation method (Wang et al., 2016 b) in order to find a better way for developing MRI detectable microspheres. As far as we knew, the two methods have never been compared.

2. Materials and methods

2.1. Materials

Acrylic acid (AA, purity ≥ 98%), potassium persulfate (KPS, A.R.) and Span 80 (C.P.) were supplied by Sinopharm Chemical Reagent Co., Ltd. (Beijing, China). N, N’-methylenebisacrylamide (MBA, A.R.) was purchased from Junyao Albert Biotechnology Company (Beijing, China). Liquid paraffin (C.P.) was obtained from Xilong Chemical Co., Ltd. (Shantou, China). Ferric chloride (FeCl3∙6H2O, A.R.) and ferrous sulfate (FeSO4∙7H2O, A.R.) were purchased from Tianjin Guangfu Fine Chemical Research Institute (Tianjin, China). Ammonia water (25%, (w/w), A.R.) and trisodium citrate dihydrate (A.R) were purchased from Beijing Chemical Plant (Beijing, China). 1,10phenanthroline hydrate (A.R.) was obtained from Yingdaxigui Chemical Reagent Factory (Tianjin, China). Hydroxylamine hydrochloride (A.R.) was purchased from Tianjin Fuchen Chemical Reagents Factory (Tianjin, China). Embospheres® were supplied by Biosphere Medical Inc. (USA). Agarose,

pentobarbital sodium and sodium carboxyl methyl cellulose were purchased from Sigma Aldrich (USA). All the other chemicals and solvents were analytical agents and used without further purification.

2.2 Synthesis and characterization of superparamagnetic iron oxide nanoparticles

Citrate coated SPIONs were synthesized by the alkaline co-precipitation method as our previous report (Wang et al., 2016 a). FeSO4∙7H2O (2.78 g) and FeCl3∙6H2O (5.13 g) were dissolved in 100 ml of distilled water. The solution was transferred into a round flask under nitrogen protection and heated up to 60 °C by using a water bath. 15.5 ml of ammonia water was added with vigorous agitation to start the reaction. After 30 min, 20 ml of sodium citrate solution (0.45 mol/L) was added to the mixture. The temperature was raised to 80 ºC and maintained for 60 min. When the mixture was cooled to room temperature, the SPIONs were isolated by centrifugation at 6000 rpm and 4 °C for 15 min, washed with distilled water till the solution was neutral. The washed SPIONs were dispersed in distilled water by ultrasonic and stored at 4 ºC. The dried SPIONs were obtained by freeze drying at -70 ºC (Freeze Drier, Christ ALPHA24, Germany). The morphology, particle size, infrared characteristics, magnetic property and X-ray diffraction (XRD) pattern of SPIONs were respectively characterized as reported previously by transmission electron microscope (TEM, JEM-1200EX, JEOL, Japan), dynamic light scattering instrument (DLS, Zetasizer NanoZS, Malvern, UK), Fourier transform infrared (FTIR) spectrophotometer (NEXUS670, Nicolet, USA), vibrating sample magnetometer (VSM, PPMS-9, Quantum Design, USA) and X-ray diffractometer (Mini Flex 600 X, Rigaku, Japan) (Wang et al., 2016 a). X-ray photoelectron spectroscopy (XPS) was performed on an X-ray photoelectron spectrometer (Axis Ultra, Kratos, England) using monochromatic Al Kα radiation. Pass energy of 160 eV and step size of 1.00 eV were employed for survey spectrum of SPIONs. For Fe 2p high-resolution spectrum, pass energy of 40 eV and step size of 0.10 eV were adopted (Jiang et al, 2012; Yang et al. 2014). The iron content of SPIONs was measured by 1,10-phenanthroline method in triplicate (Oerlemans et al., 2013). Briefly, about 45 mg of SPIONs were weighted precisely and digested in 1 ml of concentrated hydrochloric acid with sonication for 5 min, and the dissolved SPIONs were diluted with water to 100 ml in a volumetric flask. Then, the diluted solution was reacted with hydroxylamine

hydrochloride solution (10%, w/v) and 1, 10-phenanthroline (0.15%, w/v) at pH 4 to form an orangered complex solution. The absorbance of the resultant solution was measured by the UV-visible spectrometer at 510 nm. The content of SPIONs was calculated, which was treated as Fe3O4 according to the result of SPIONs in Section 3.8.

2.3 Preparation of SPIONs-loaded poly (acrylic acid) microspheres

The SPMs were prepared by inverse suspension polymerization (Correia et al., 2016; Cui et al., 2012). Briefly, 40 ml of liquid paraffin (oil phase) containing Span 80 was poured into a three-necked flask equipped with a mechanical stirrer. 400 mg of the freeze-dried SPIONs were dispersed in a blended aqueous solution of AA (monomer, 3.6 g), MBA (cross linker, 0.154 g) and KPS (initiator, 0.136 g). The resultant suspension was then added dropwise into the oil phase with stirring. The polymerization reaction was carried out at 55 °C for 3 h under nitrogen protection. After cooling to room temperature, the resultant microspheres were washed successively with 0.5% Tween 80 (w/v) solution and deionized water, and then sieved to the final size range of 100-900 μm by wet sieving. For special use in the later experiments, some of the SPMs were further separated into different subgroups with the size ranges of 100-300, 300-500, 500-700 and 700-900 μm. Finally, all SPMs were freeze-dried or stored in saline for further analysis. PMs were also prepared by the aforementioned method except the addition of SPIONs.

2.4 Morphology and particle size

The morphologies of the SPMs and PMs were respectively observed by an optical microscope (XTZ-D/T, Shanghai Optical Instrument Factory No. 6, China). Furtherly, The TEM was used at an acceleration voltage of 100 kV to observe the freeze-dried SPMs (Frounchi and Shamshiri, 2015; Sirivisoot and Harrison, 2015), which were cut into thin slices to identify SPIONs particles in the SPMs (Wang et al., 2016 b). The particle size of SPMs and PMs were separately measured under the optical microscope. The number-average diameter (Dn) of microspheres was expressed as Eq. (1) by measuring the diameters of at least 1000 individual microsphere under optical micrographs (Zhou et al., 2012). The size

distribution graph of SPMs and PMs was drawn. (1) where di is the diameter of individual microsphere, and ni is the number of microspheres with di.

2.5 Fourier transform infrared spectrum

FTIR was used to illustrate the infrared characteristics of SPMs and PMs. Small quantities of freeze-dried SPMs and PMs were separately scanned over the wavelength range of 4000 to 400 cm-1 with KBr pellet in the FTIR instrument (Cui et al., 2012).

2.6 SPIONs content

SPIONs loaded on SPMs were treated as Fe3O4 according to the result of SPMs in Section 3.8. The contents of SPIONs in terms of weight per volume for each subgroup of SPMs (100-300, 300-500, 500-700, and 700-900 μm) were measured by the same method as Section 2.2., except that the quantity of SPMs was 0.5 ml (taken by a syringe), the digestion solution was a mixture of fuming nitric acid and concentrated hydrochloric acid at a volume ratio of 1:3 and the time of ultra-sonication was 30 min. The content of SPIONs in terms of weight per volume of SPMs was calculated. The content of SPIONs in terms of weight percentage of SPMs was also measured by the aforementioned method, except that 500 mg of freeze-dried SPMs were digested.

2.7 Magnetic property

In order to confirm the loading of SPIONs in SPMs and investigate the magnetic property of SPMs, the magnetization curves of freeze-dried SPMs were recorded on the VSM with a maximum magnetic field of 6 kOe at room temperature (Hagit et al., 2010; Kim et al., 2015).

2.8 X-ray diffraction

The XRD patterns of the freeze-dried SPMs were analyzed with the X-ray diffractometer with a Cu Kα source (λ = 1.54 Å) at 40 kV and 15 mA scanning from 10 ° to 80 ° at a scan rate of 4 °/ min (Frounchi and Shamshiri, 2015; Tian et al., 2015).

2.9 X-ray photoelectron spectroscopy

Due to the similarity in the XRD patterns of Fe 3O4 and γ-Fe2O3 (Maity and Agrawal, 2007), XPS was further performed as same as that in Section 2.2 to verify the compositions of SPIONs loaded in SPMs.

2.10 Elasticity

To determinate the elasticity of microspheres, the PMs and SPMs were sieved to the size range of 700-750 μm, respectively. Four tests (compression test for monolayer of microspheres, stress relaxation test for monolayer of microspheres, compression test for single microspheres, and repeated compression test for single microspheres) were performed using a texture analyzer (TA. XTPlus, Stable Micro Systems, UK). The analyzer was equipped with a 6 mm cylindrical probe and a 5 N load cell. The initial force was set at 0.001 N. Both compression force and displacement of probe were collected 100 times per second by a computer connected to the analyzer. PMs and SPMs were used as samples and Embospheres were used as a contrast. All the tests were performed at least three times (Lewis et al., 2006; Cui et al., 2012).

2.11 Magnetic resonance imaging

2.11.1 MR imagines in vitro Each subgroup of SPMs was diluted with agarose (0.5%, w/v) to a series concentration of 0%, 2.5%, 5%, 10% and 20% (v/v) in Eppendorf tubes to prepare imaging phantoms. The MRI scanner (GE Medical Systems, LLC, USA; 3.0 Tesla) with a head coil was used to detect the gel phantom. The T2-weighted MR images was acquired by a fast spin-echo (FSE) sequence with the following parameters: repetition time (TR) of 3000 ms, echo time (TE) of 99.7 ms, field of view (FOV) of

160×160 mm, slice thickness of 5.0 mm, flip angle of 150°, matrix of 320×224, and number of excitation of 3 (Choi et al., 2015; Kim et al., 2013).

2.11.2 MR imagines in mice MRI visibility of SPMs in vivo was evaluated in three mice with an approval of the Animal Care Committee of Peking University Health Science Center. Three male adult Kun Ming mice (about 25 g of initial weight) were purchased from Peking University Experimental Animal Center (Beijing, China) (Franklin-Ford, et al. 2012; Wang, et al., 2016b). All animals were anesthetized by intraperitoneal injection of 1% (w/v) pentobarbital sodium (75 mg/kg body weight). 0.1 ml of sterilized SPMs (100-300 μm of SPMs was chosen with the best syringeability; autoclaved at 121 °C for 30 min) were suspended in 0.7 ml of sodium carboxylmethyl cellulose solution (1%, w/v), and 0.2 ml of the suspension was injected subcutaneously to each mouse. The animals were scanned by the 3.0 T MR scanner with a head coil pre and post injection immediately using the T2-weighted FSE sequence: TR of 3000 ms, TE of 102 ms, FOV of 120×120 mm, slice thickness of 5.0 mm, flip angle of 150°, matrix of 320×224, and number of excitation of 4. The MR images were recorded and compared with the same animal.

2.12 Embolization and MR imaging of SPMs in rabbits

To evaluate the embolization capability and in vivo MRI detectability of the SPMs, renal arterial embolization was performed in two male New Zealand white rabbits weighting 3.5 ± 0.1 kg each (purchased from Peking University Experimental Animal Center, Beijing, China). To induce anesthesia, 2% (w/v) pentobarbital sodium solution (30 mg/kg body weight) was injected via the marginal ear vein. Afterwards, a 2.8 Fr microcatheter (Progreat®, Terumo Co., Tokyo, Japan) was inserted into the left renal artery under the guidance of digital subtraction angiography (DSA, Innova®4100, GE Healthcare Technologies, USA). 0.3 ml of sterilized SPMs (100-300 μm) suspended in iohexol 350 (0.3 ml) and saline (0.2 ml) was then delivered slowly to the renal trunk. A renal artery angiography was carried out before and after the embolization. Finally, the catheter was gently pulled out. The femoral artery was ligated and the incision was closed properly. All animals were scanned by the 3.0 T MR scanner with knee coil before and after embolization

2 h and 7 d with the two following sequences: T2*-weighted gradient recalled echo sequence (OA×2D MERGR: TR 551 ms, TE 5.2 ms, slice thickness 4.0 mm, FOV 20×20 cm, flip angle 20°, and matrix 256×224) and T2-weighted fast spin-echo sequence (OA×T2 PROPELLER: TR of 3478 ms, TE of 57 ms, slice thickness of 4.0 mm, FOV of 20×20 cm and matrix of 256×256; OCorT2 PROPELLER: TR of 3000 ms, TE of 59 ms, slice thickness of 4.0 mm, FOV of 20×20 cm and matrix of 256×256). The MR images were recorded and compared between the same animal.

2.13 Statistical analysis

All results were presented as mean value and standard deviation (SD) as indicated. Analyses were performed using a SPSS statistical package (version 20.0; IBM Corp., Armonk, New York, USA). The Student’s t-test was used for the comparison of means and nonparametric Wilcoxon signed-rank test was used for the comparison between two groups. Tests were considered statistically significant with a P value < 0.05 and highly significant with a P value < 0.01.

3. Results and discussion

3.1 Synthesis and characterization of SPIONs

The citrate modified SPIONs were prepared successively and proved to be in inverse spinel crystalline structure and superparamagnetic with a saturation magnetization of 63.58 emu/g. The particle size of freeze-dried SPIONs was determined around 12.37 nm by TEM and XRD (calculated by Debye-Scherrer half width formula based on the strongest peak (311)), while the hydrodynamic particle size was 72.4 nm by DLS as mentioned in our previous work (Wang et al., 2016 a). The content of SPIONs (Fe3O4) was assayed to be 80.98 ± 2.52% (w/w) in this study.

3.2 Preparation of SPMs

Several types of MRI detectable microspheres have been reported in literatures by now. MRI detectable materials could be distributed in the whole microspheres (Ménager et al., 2004; Zielhuis et

al., 2007; Lee et al., 2008; Chung et al., 2012; Kim et al., 2016; Li et al., 2016; Wang et al, 2016 b), adsorbed (Fidelman et al., 2002; Wilson et al., 2003) or modified on the surface of microspheres (Cilliers et al., 2008). In this study, SPIONs were distributed in the whole SPMs by polymerizing the monomer of acrylic acid mixed with SPIONs. The method was considered relatively simple compared with the modification method and could provide long-term MRI detectability in contrast with the adsorption method. Besides, the properties of SPMs, such as MRI detectability and elasticity, could be adjusted by changing the quantity of SPIONs added in the polymerization reaction.

3.3 Morphology and particle size

The optical micrographs of PMs and SPMs were separately shown in Fig. 1 A and Fig. 1 B. Both of the PMs and SPMs were well dispersed with spherical shape and smooth surface. The SPIONs were distributed in the whole SPMs. Compared with irregular particles, spherical microspheres were reported to be calibrated more easily and precisely (Laurent et al., 1996), prevent catheter clogging and achieve better controllable and predictable occlusion to target vessel (Andrews and Binkert, 2003; Bendszus et al., 2000; Bendszus et al., 2005; Laurent A, 2007; Pelage et al., 2002). Fig. 1 C and 1 D showed the TEM images of SPMs with the dark region referring to SPIONs. The SPIONs could also be seen distributed in the whole SPMs. No obvious change was found in appearance and diameter of SPIONs before and after the polymerization reaction (Wang et al, 2016 a). The size distributions of PMs and SPMs were displayed in Fig. 2. Being sieved within the range of 100-900 μm, the PMs and SPMs were found mainly distributed around 100-300 μm. The numerical mean diameter of PMs and SPMs was calculated as 314 ± 165 μm and 298 ± 123 μm, respectively. There was no significant difference between the size distribution of PMs and SPMs (P > 0.05), which suggested the addition of SPIONs did not impact on the size distribution of polymerized microspheres. It was worth mentioning that the particle size of SPMs successfully matched the clinical demands when compared with several clinical used microspheres. For example, the particle size of Embospheres and DC Bead were respectively offered in the range of 40-1200 μm and 70-900 μm.

3.4 Fourier transform infrared spectrum

Fig. 3 displayed the FTIR spectra of PMs (Fig. 3 A) and SPMs (Fig. 3 B), respectively. For SPMs, the absorption peak at 2920 cm-1 was attributed to the stretching vibration of ―CH2―, demonstrating a successful polymerization of double bands (―C=C―) of acrylic acid. Combined with the asymmetric and symmetric stretching vibration peaks of COO- at 1549 cm-1 and 1414 cm-1, the peaks at 3430 cm-1 and 1724 cm-1 separately assigned to the stretching vibration of O―H and C=O manifested the existence of carboxyl (Cui et al., 2012). All the above absorption peaks could be found corresponding results in the FTIR spectrum of PMs, illustrating the addition of SPIONs had no influence on the polymerization reaction. Meanwhile, the peak at 631 cm-1 due to the characteristic absorption of Fe―O in the spectrum of SPMs indicated the SPIONs were successfully loaded on the SPMs (Wang et al., 2016 b; Xue et al., 2016).

3.5 SPIONs content

The SPIONs (Fe3O4) contents per volume of wet SPMs in each subgroup were shown in Table 1. The average content of SPIONs in the four subgroups was calculated as 1.41 ± 0.16 mg/ml, which converted to be 1.02 ± 0.11 mg Fe/ml or 18.25 ± 2.03 mmol Fe/L. As the concentration of Resovist, a T2 contrast agent commonly used in clinic, was reported to be 0.2 mmol Fe/L in human liver (Kim et al., 2007), the SPMs were thus supposed to be detectable by clinical MRI scanner. Meanwhile, dosage of the T2 contrast agent was reported to be 8-15 μmol Fe/kg (Weinmann et al., 2003), corresponding to 480-900 μmol Fe for a patient with a body weight of 60 kg. Referring to the dose of a commercial used microspheres for the embolization of hepatocellular carcinoma (Lammer et al., 2010), for example, when 4 ml of SPMs are needed, the equivalent of Fe in SPMs should be 73 μmol which is much lower than that of the T2 contrast agent. Thus, combing the good cytocompatibility of PMs (Cui et al., 2012) with the low amount of SPIONs, the SPMs were speculated to be safety for use. Table 1 also listed the SPIONs (Fe3O4) contents at weight percentage of freeze-dried SPMs in each subgroup. The SPIONs contents in different subgroups did not show significant difference (P>0.05). The average content of SPIONs in the four subgroups was 3.63 ± 0.08 % (w/w), which was much lower than that of magnetic microspheres (18.25 ± 0.37%, w/w) we prepared previously by in situ method (Wang et al., 2016 b).

3.6 Magnetic property

The magnetization curve of SPMs measured by VSM was shown in Fig. 4. As no coercivity and remanence magnetization were found in the hysteresis loop, the SPMs were proved to be superparamagnetic as same as the SPIONs before loading (Wang et al., 2016 a). The results indicated that the polymerization reaction did not change the magnetic properties of SPIONs. Meanwhile, the saturation magnetization of SPMs (3.53 emu/g) was much higher than those of magnetic embolic microspheres reported in literatures, which varied from 0.67 to 1.66 emu/g (Hagit et al., 2010; Wang et al., 2016 b). Compared with the magnetic microspheres we obtained previously by in situ method (Wang et al., 2016 b), the SPMs prepared in this study had higher saturation magnetization (3.53 emu/g versus 1.66 emu/g) but lower SPIONs contents (3.63% versus 18.25%, w/w). Probably, two reasons might be used to explain the phenomenon. It was reported that the saturation magnetization could be increased by well-crystallized particles (Hong et al., 2006). However, the crystallinity of SPIONs in situ method was supposed to be affected by the steric hindrance of polymer and result more crystal defects compared with the present method. Besides, the particle size of SPIONs in situ method (6.92 nm) was different from that of present method (14.15 nm according to the results in Section 3.7), and the small SPIONs were reported to have lower saturation magnetization than large ones (Jun et al., 2008).

3.7 X-ray diffraction

Fig. 5 showed the XRD pattern of SPMs. By comparing the resultant XRD pattern with standard JCPDS cards, the crystalline phase of SPIONs in SPMs could be found out. The diffraction peaks of SPMs at 2 of 30.3°, 35.7°, 43.2°, 57.3° and 62.9° were proved to correspond with the indices of (220), (311), (400), (511) and (440) planes of Fe3O4 (JCPDS No. 85-1436) respectively, indicating the inverse cubic spinel structure of SPIONs in SPMs (Peng et al., 2015; Xue et al., 2016). The characteristic peaks of SPMs were all observed in the XRD pattern of SPIONs (Wang et al., 2016 a) though they were diffused and even the peaks at position of 18.1° and 53.6° could not be detected due to the existence of amorphous polymer (Xu et al., 2010; Xue et al., 2016). The results suggested the polymerization reaction did not affect the crystalline phase of SPIONs. The particle size of SPIONs in

SPMs was calculated to be 14.15 nm according to Debye-Scherrer half width formula based on the strongest peak (311).

3.8 X-ray photoelectron spectroscopy

The XPS survey spectra of SPIONs and SPMs were shown in Fig. 6. The main peaks could be ascribed to Fe, C, O and Na elements as labeled in Fig. 6A and 6C. The Fe 2p spectrum of SPIONs before and after being loaded in SPMs was shown in Fig. 6B and 6D, respectively. The two broad peaks located at 710.3 eV and 723.8 eV for SPIONs (710.6 eV and 723.7 eV for SPMs) were separately ascribed to the Fe 2p3/2 and Fe 2p1/2 of Fe3O4. Usually, the Fe3+ in γ-Fe2O3 exhibits a shakeup satellite peak around 719.0 eV. The absence of this characteristic peak in the XPS spectra confirmed the pure magnetite phase in SPMs (Gao and Chambers, 1997; Yang et al., 2014).

3.9 Elasticity

Fig. 7 A showed the compression curves of SPMs, PMs and Embospheres with the maximum deformation of 50%. Based on the data of compression, Young’s modulus of the three kinds of microspheres was separately calculated as 74.90 ± 1.86 kPa for SPMs, 146.40 ± 7.49 kPa for PMs and 158.27 ± 13.54 kPa for Embospheres (showed in Table 2). The rigidity of SPMs displayed was lower than both PMs and Embospheres (P < 0.01). It was reported that softer embolic microspheres were easier to diffuse and enter in distal vessels of organs than harder ones (Hidaka et al., 2011). Therefore, the SPMs prepared in this study were more likely to occlude the distal vessels compared with PMs and Embospheres. To our knowledge, the elasticity of MRI detectable microspheres was seldom reported, so compared with the Young’s modulus of the magnetic microspheres (477.13 ± 57.11 kPa) synthesized by in situ method in our precious work (Wang et al., 2016 b), the rigidity of SPMs was lower. The reason was speculated that the addition of SPIONs in polymerization reaction might decrease the polymerizing degree of AA. Fig. 7 B showed the stress relaxation curves of SPMs, PMs and Embospheres. The data of residual force, RHT, failure stress, failure deformation, springiness, cohesiveness and resilience were calculated and displayed in Table 2.

The residual force and RHT of SPMs (34.48 ± 0.51 %, 55.33 ± 2.92 s) were a little smaller than that of PMs (42.43 ± 0.79 %, 56.02 ± 2.28 s) or Embospheres (39.70 ± 1.60 %, 62.13 ± 1.06 s), suggesting that the SPMs had a relatively weak ability to keep rigidity during relaxation but could achieve equilibrium state faster than PMs or Embospheres (Cui et al., 2012). As for the magnetic microspheres synthesized by in situ method, the residual force and RHT were 59.11 ± 0.37 % and 10.53 ± 1.75 s (Wang et al., 2016 b). The failure stress and failure deformation of SPMs (0.37 ± 0.04 N, 84.17 ± 0.43 %) had no significant difference with those of Embospheres (0.45 ± 0.12 N, 82.10 ± 2.30 %) (P > 0.05), but they were much smaller than PMs (1.30 ± 0.18 N, 98.81 ± 0.86 %) (P < 0.01). The results revealed the addition of SPIONs lowered the failure deformation of SPMs compared with PMs. As the magnetic microspheres synthesized by in situ method failed to be broken (Wang et al., 2016 b), no data of the failure stress and failure deformation could be compared with the SPMs. Microspheres may undergo several times of compressions during the delivery to the final position of embolization. The repeated compression test was used to investigate the recovery ability of the microspheres. The ideal value of springiness, resilience and cohesiveness should tend to be one. There was no significant difference in springiness and resilience between SPMs, PMs and Embospheres (P > 0.05), but the cohesiveness of SPMs was smaller than PMs or Embospheres, indicating the recovery size and elasticity of PMs, SPMs and Embospheres were similar, but the recovery resistance of SPMs was not as good as PMs and Embospheres (Cui et al., 2012). The springiness of SPMs had no significant difference with that of the magnetic microspheres prepared by in situ method (P>0.05), but the resilience of SPMs was higher and the cohesiveness of SPMs was lower (P<0.01). Referring to the literature (Hidaka, et al., 2011), the Young’s modulus of three clinical used embolic microspheres (Embospheres, Beadblock and Embozene), measured at different condition with this study, was reported to be 39.6 ± 5.05, 18.8 ± 4.00 and 13.6 ± 1.98 kPa, respectively. The Young’s modulus of Beadblock and Embozene was therefore separately 47.5% and 34.3% of Embospheres. In this study, the Young’s modulus of SPMs (74.9 ± 1.86 kPa) was 47.3% of Embospheres (158.27 ± 13.54 kPa), approximating to that of Beadblock and higher than that of Embozene. Thus, the in vivo embolization of SPMs was supposed to be comparable with the clinical used Beadblock. While the Young’s modulus of magnetic microspheres prepared in situ method was 301% of Embospheres (477.13 ± 57.11 kPa vs 158.27 ± 13.54 kPa), much higher than any of the three clinical used

microspheres.

3.10 Magnetic resonance imaging

3.10.1 MR imagines in vitro Fig. 8 was T2-weighted MR images of SPMs in agarose gel in vitro. The SPMs at concentration of 0% (i.e. pure gel) were used as controls. The SPMs at concentration of 2.5%, 5%, 10% and 20% in all subgroups could be detected by the low signal intensity areas. The decreased signal intensity in T2weighted MR images of SPMs could be explained to the loading of SPIONs which shortened the transverse relaxation time and generated dark signal areas (Ferrucci and Stark, 1990). Moreover, in the same subgroup of SPMs, the signal intensity decreased with the concentration of SPMs increasing, which coincided with our previous report (Wang et al., 2016 b).

3.10.2 MR imagines in mice For showing low signal intensity in vitro, SPMs were further selected to MRI study in mice. Fig. 9 showed the T2-weighted MR images before and after an injection of SPMs for the same mice. It could be seen obviously the position turned out to be dark (marked by circles) after injection of SPMs, indicating that the SPMs under the skin of mice could be detected by MRI.

3.11 Embolization and MR imaging of SPMs in rabbits

Renal embolic model was chosen (Hagit et al., 2010; Kim et al., 2007) considering the blood vessels of kidney were relative simple compared with other embolic models (for example liver model), which would help to observe the flow of contrast agent and evaluate the efficiency of embolization. Besides, relative long-time survival of animal could be expected as kidneys were paired organs and when one kidney was embolized the other one kept on working. An angiogram could offer the information of blood flow in arteries and arterioles by monitoring the X-ray contrast agent injected. Fig. 10 showed the arterial angiogram of a rabbit’s left kidney before and after embolization. The renal arterioles were all unobstructed and thus visible before embolization, whereas the anterior segment of kidney could not be visualized after embolization. The results certified

a successful embolization by SPMs to the arterioles of renal parenchyma. Fig. 11 was the MR images of a rabbit’s kidney before, 2 h and 7 d after the embolization. Two kinds of common sequences used in clinic were employed in this study, of which Merge sequence was more sensitive to detect T2 contrast agents but could not reduce the effect of respiratory motion on images while Propeller sequence was on the contrary. Compared with the MR images before embolization (Figure 11 A, 11 D and 11 G), low signal intensity areas (denoted by arrows) in the arterioles of the rabbit’s left renal parenchyma could be easily found in images 2 h (Figure 11 B, 11 E and 11 H) and 7 d (Figure 11 C, 11 F and 11 I) after embolization. Combined with Fig. 9, the low signal intensity areas could be inferred as the positions of SPMs. Besides, the location of SPMs could be found in both axial plane images (Figure 11 B, 11 E at 2 h or Figure 11 F at 7 d) and coronal plane images (Figure 11 H at 2 h or Figure 11 I at 7 d) at the same time, which could be used for stereotactic localization of the SPMs in vivo. No changes of the SPMs signals were found between the images at 2 h and 7 d after embolization (Figure 11 C, 11 F, and 11 I), demonstrating the SPMs could provide contrast and long-term signal intensity for the nonbiodegradable property of SPMs. Moreover, local tissue necrosis (denoted by a circle in Figure 11 I) could be observed in the embolized position, indicating that the aim of embolization was achieved.

Conclusion

In this study, hydrophilic poly (acrylic acid) microspheres were given MRI detectable property for monitoring the spatial distribution of embolic microspheres with the combination of MRI equipment during and after embolization. The MRI detectable poly (acrylic acid) microspheres (SPMs) were

prepared

successfully

by

inverse

suspension

polymerization

method,

in

which

superparamagnetic iron oxide nanoparticles (SPIONs) were first synthesized and then mixed with monomer of acrylic acid. Through a series of evaluations in vitro and in vivo, SPIONs in SPMs were confirmed unchanged after the polymerization reaction, which kept superparamagnetic with inverse spinel structure of magnetite. The more important results revealed that the SPMs processed good property for both embolization and MRI detectability, including spherical shape, smooth surface, proper particle size and elasticity, and successful embolization to the kidney of a rabbit with long-term T2-weighted MRI detectability, etc. Thus, the SPMs present to be a potential MRI detectable embolic

agent that may play an important role in improving the efficiency and safety of embolization in the future. Besides, compared with the magnetic MRI detectable microspheres we prepared previously by in situ precipitation method, SPMs in this study showed the advantage of loading much less of SPIONs but offering higher saturation magnetization. Still, further work should be done to enhance the feasibility of application, such as histocompatibility, selective embolization to larger animal model and quantitative detection by MRI, etc.

Acknowledgements

This study was supported by the National Natural Science Foundation of China (Grant No 81571779). The authors thank Zhen-Zhen Li and Ze-Yang Fan of Department of Interventional Radiology and Vascular Surgery, Peking University First Hospital for their assistances in the experiment of embolization. The authors thank Xiaonan Yao of College of Chemistry and Molecular Engineering of Peking University for his help in calculating the particle size of SPIONs.

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Figure captions:

Fig. 1. Morphology of PMs and SPMs. Optical micrographs of PMs (A) and SPMs (B); transmission electron micrographs of SPMs (C, D). Fig. 2. Size distribution of PMs and SPMs. Fig. 3. Fourier transform infrared spectra of PMs (A) and SPMs (B). Fig. 4. The magnetization curve of SPMs. Fig. 5. The X-ray diffraction pattern of SPMs. Fig. 6. The XPS spectra of SPIONs and SPMs. (A) XPS survey spectrum of SPIONs, (B) Fe 2p XPS spectrum of SPIONs, (C) XPS survey spectrum of SPMs, and (D) Fe 2p XPS spectrum of SPMs. Fig. 7. Compression curves (A) and stress relaxation curves (B) of PMs, SPMs and Embospheres. Fig. 8. T2-weighted MR images of gel phantom from left to right: the concentration of SPMs at 0%, 2.5%, 5%, 10% and 20% (v/v), respectively. Fig. 9. The T2-weighted MR images of the same mouse before and immediately after subcutaneous injection of SPMs into the back. The circle denotes the dark signal area induced by SPMs. Fig. 10. Arterial angiogram of a rabbit’s left kidney before and immediately after embolization. Fig. 11. The T2-weighted (PROPELLER sequence) and T2*-weighted (MERGE sequence) MR images of the same rabbit before embolization, at 2 h and 7 d after embolization. The white arrows denote the dark signal area induced by SPMs.

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Table 1. The SPIONs content in different subgroup of SPMs. Subgroups (μm)

100-300

300-500

500-700

700-900

Content (mg/ml) Content (wt %)

1.64 ± 0.12

1.38 ± 0.10

1.29 ± 0.04

1.32 ±0.03

3.53 ± 0.16

3.64 ± 0.17

3.64 ± 0.12

3.74 ± 0.12

Table 2. Young’s modulus, residual force and relaxation half time (RHT), failure deformation, failure stress, springiness, cohesiveness and resilience of different microspheres (n = 3). Microspheres

a

SPMsa

PMsb

Embosphere

Young’s modulus (kPa) Residual force (%) RHT (s) Failure deformation (%) Failure stress (N)

74.90 ± 1.86 34.48 ± 0.51** 55.33 ± 2.92* 84.17 ± 0.43 0.37 ± 0.04

146.40 ± 7.49 42.43 ± 0.79 56.02 ± 2.28* 98.81 ± 0.86** 1.30 ± 0.18**

158.27 ± 13.54 39.70 ± 1.60 62.13 ± 1.06 82.10 ± 2.30 0.45 ± 0.12

Springiness

0.74 ± 0.08

0.66 ± 0.07

0.71 ± 0.15

Cohesiveness Resilience

0.24 ± 0.03** 0.72 ± 0.01

0.38 ± 0.06 0.73 ± 0.03

0.57 ± 0.12 0.60 ± 0.15

**

SPIONs-loaded poly (acrylic acid) microspheres, b Poly (acrylic acid) microspheres. Data was expressed as the mean ± SD. Statistical significance: ** P<0.01, * P<0.05 (compared to Embospheres).