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International Journal of Pharmaceutics 452 (2013) 211– 219

Contents lists available at SciVerse ScienceDirect

International Journal of Pharmaceutics

j o ur nal ho me page: www.elsev ier .com/ locate / i jpharm

esearch of novel biocompatible radiopaque microcapsules forrterial embolization

iao-Jing Lua, Yuan Zhanga, Dai-Chao Cuia, Wen-Jing Menga, Ling-Ran Dua,ai-Tao Guanb, Zhuo-Zhao Zhengc, Nai-Qi Fuc, Tian-Shi Lvb, Li Songb,ing-Hua Zoub, Wan-Liang Lua, Tian-Yuan Fana,∗

Department of Pharmaceutics, School of Pharmaceutical Sciences, Peking University, Beijing 100191, ChinaDepartment of Interventional Radiology and Vascular Surgery, Peking University First Hospital, Beijing 100034, ChinaDepartment of Radiology, Peking University Third Hospital, Beijing 100191, China

a r t i c l e i n f o

rticle history:eceived 17 February 2013eceived in revised form 9 April 2013ccepted 2 May 2013vailable online 22 May 2013

a b s t r a c t

Embolic agents, such as microparticles, microspheres or beads used in current embolotherapy are mostlyradiolucent, which means the agents are invisible under X-ray imaging during and after the process ofembolization, and the fate of these particles cannot be precisely assessed. In this research, a radiopaqueembolic agent was developed by encapsulating lipiodol in polyvinyl alcohol. The lipiodol-containingpolyvinyl alcohol microcapsules (LPMs) were characterized and evaluated for their morphology, size dis-

eywords:icrocapsules

mbolization-ray imagingipiodololyvinyl alcohol

tribution, lipiodol content, lipiodol release, elasticity, and deliverability through catheter. The radiopacityof LPMs in vials and in living mice was both detected by an X-ray imaging system. The biocompatibility ofLPMs was investigated with L929 cells and in mice after subcutaneous injection. Embolization of LPMs toa rabbit kidney was performed under digital subtraction angiography (DSA) and the radiopacity of LPMswas verified by computed tomography (CT).

© 2013 Elsevier B.V. All rights reserved.

. Introduction

Embolization has emerged as a highly effective interventionaladiographic technique in a wide variety of diseases in current med-cal therapy. It has been used for treatment of inoperable tumorsBendszus et al., 2000; Skalicky et al., 2010; Treska et al., 2010),rteriovenous malformations (AVMs) (Fleetwood and Steinberg,002; Guzinski et al., 2010; Khan et al., 2010) and haemopty-is (excessive bleeding) (Loffroy et al., 2012; Shin et al., 2011;inaya et al., 2004; Yoon, 2004), etc. Embolic agents, such as micro-articles, microspheres or beads have been successfully employed

n the embolization therapy (Bonomo et al., 2010; Hagit et al.,010; Lookstein and Guller, 2004; Matsumaru et al., 1997; Nicolinit al., 2011; Qureshi, 2004; Shah et al., 2011; Sousa et al., 2011).owever, most of these embolic agents are radiolucent, whicheans that they cannot be visualized under X-ray imaging and can

nly be indicated indirectly by the flow of contrast agent injectedn blood vessels. Consequently, complications such as “reflux

ith non-target embolization” and “through embolization” are

∗ Corresponding author. Tel.: +86 10 82801584; fax: +86 10 82801584.E-mail addresses: tianyuan fan@bjmu.edu.cn, tianyuan fan@hotmail.com

T.-Y. Fan).

378-5173/$ – see front matter © 2013 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.ijpharm.2013.05.001

essentially undetectable (Grüll et al., 2010; Saralidze et al., 2007).In addition, some possible migration of the embolic agent overtime cannot be found to guide another injection of embolic agentif repeated treatment is necessary (Emans et al., 2005).

In order to improve embolotherapy, some methods havebeen employed to develop radiopaque particles. For example,radiopaque substances have been used to disperse in microspheres,including heavy-metal powders, inorganic salts or organic com-pounds which contain atoms of heavy elements (Hahn et al.,2011; Jayakrishnan et al., 1990; Thanoo and Jayakrishnan, 1990).Radiopacity of particles has been acquired in this way, but thephysical or mechanical properties of particles have been deteri-orated in the meantime. Moreover, the leaking of inorganic saltsinto body fluids may cause a serious threat due to the toxicityof metal ions (Jayakrishnan and Thanoo, 1992). The synthesis ofintrinsic radiopaque materials is another try, such as copolymer-izing methyl methacrylate (MMA) with vinyl monomer of metalsalt (barium or zinc acrylates) (Moszner et al., 1995) or graftingiodine-containing molecular onto preformed polymers (Hagit et al.,2010; Horák et al., 1987; James et al., 2006; Mottu et al., 2002). Still,

appropriate chemical modification of non-radiopaque polymersadversely affects their certain properties, such as hydrophilicity,swelling ability, elasticity and biocompatibility (Jayakrishnan andThanoo, 1992).

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In present work, radiopaque embolic microcapsules wereesigned to encapsulate lipiodol in polyvinyl alcohol (PVA). Lipi-dol is a worldwide used X-ray contrast and liquid embolic agentith advantages of high X-ray contrast, low toxicity and cost, whichas employed to provide radiopaque character for the PVA micro-

apsules (Ahn et al., 2011; Galperin et al., 2007; Gandhi et al., 2006;allouard et al., 2010). PVA in the form of microparticles has beensed for embolization for several decades (Bilhim et al., 2011; Lewist al., 2006). PVA has also been employed in the field of pharma-eutics as film-forming material (DeMerlis and Schoneker, 2003),hich was used as a material to encapsulate lipiodol. Therefore,

he lipiodol-containing PVA microcapsules (LPMs) were supposedo be radiopaque and biocompatible for embolization.

In this study, LPMs were developed by encapsulating lipiodoln PVA with a simple coacervation method (Lazko et al., 2004;eimann et al., 2009). The properties of LPMs in vitro were eval-ated by a series of methods, and the in vivo embolization of LPMsas investigated in a rabbit kidney.

. Materials and methods

.1. Materials

PVA was of pharmaceutical grade purchased from Shanxianwei Corporation (China). Formaldehyde (A.R. 37.0–40.0%),ichloromethane (DCM, A.R.), potassium iodide (A.R.), chloralydrate (A.R.) and soluble starch (A.R.) were purchased frominopharm Chemical Reagent Co., Ltd. (China). Lipiodol wasbtained from Shanghai Xudonghaipu Pharmaceutical Indus-ry Co., Ltd. (China). Sulfuric acid (A.R.), dimethyl sulfoxideDMSO, A.R.) and sodium sulphate were purchased from Bei-ing Chemical Plant (China). m-Chloroperbenzoic acid (m-CPBA,.R.) was purchased from Shanghai Jiachen Chemical Industryo., Ltd. (China). Dimethyl sulfide (A.R., ≥99.0%) was suppliedy Sigma–Aldrich. EmbosphereTM was obtained from Biosphereedical Inc. (USA). Iohexol was supplied by GE Healthcare

Shanghai, China). L929 cell line was supplied by Tumor Markeresearch Center (Beijing, China); 3-(4,5-dimethylthiazol-2-yl)-,5-diphenyl-2H-tetrazolium bromide (MTT) and Pento-barbitalodium were purchased from Sigma Chemical Company (USA);PMI 1640 medium was purchased from Macgene Science andechnology Co., Ltd. (Beijing, China) and fetal bovine serum (FBS)as purchased from Thermo Fisher Scientific Biological Chemical

roducts Co., Ltd. (Beijing, China). Iopamiro 370 was obtained fromracco Sine Pharmaceutical Co. Ltd. (Shanghai, China).

.2. Preparation of LPMs

A 100 ml open-mouth flask equipped with a paddle was placedn a thermostatic water bath. PVA solution (4%, w/v) and sodiumulphate solution (20%, w/v) were mixed in the flask at 10 ◦C.ipiodol (3 ml) was added into the above mixture with vigorousgitation to form an o/w emulsion. Coacervation was achieved byradually increasing temperature (1 ◦C/min) to the cloud point ofVA solution, and the temperature was kept till the end of prepa-ation. The cross-linking reaction was initiated by adding 10 ml oformaldehyde and 2 ml of sulphuric acid (50%, v/v) to the flask, andhe reaction was lasted for 24 h. Then insoluble LPMs were filtered,ashed with distilled water and stored in a phosphate buffer solu-

ion (PBS, pH 7.4) (Bachtsi et al., 1996; Bachtsi and Kiparissides,

996).

For special experiments, LPMs were separated into subgroups of00–500, 500–700 and 700–900 �m by wet-sieving as our previousork (Zhou et al., 2012).

armaceutics 452 (2013) 211– 219

2.3. Morphology

Morphology of LPMs was observed with an optical microscope(XTZ-D/T, Shanghai Optical Instrument Factory No. 6) and an envi-ronmental scanning electron microscope (ESEM) (FEI Quanta 200F,EDAX/AME-TEK, US), respectively. The cross section of LPMs wasalso observed under ESEM, where the microcapsules were cut openand the lipiodol left inside was washed out.

2.4. Size distribution

Particle size of LPMs was measured under optical microscope.The number-average diameter (Dn) of microspheres was expressedas Eq. (1) by measuring at least 500 individual particles undermicroscope. The size distribution graph of LPMs was drawn.

Dn =∑

nidi∑ni

(1)

In Eq. (1), di is the diameter of individual microcapsule, and niis the number of microcapsules with di.

2.5. Determination of elasticity

To evaluate the elasticity of LPMs, three tests were performedwith texture analyzer (TA. XTPlus, Stable Micro Systems, UK),including compression test for monolayer of microparticles, stressrelaxation test for monolayer of microparticles and compressiontest for single microparticle, which were described in details in ourprevious work (Cui et al., 2012). The analyzer was equipped with a6 mm cylindrical probe and a 5 N load cell. The initial force was setat 0.001 N. Both stress and displacement of probe were collected100 times per second by a computer connected to the analyzer.LPMs and Embospheres were used as sample and control, respec-tively. All particles were chosen within the size of 700–750 �m andmeasured at least three times. Data of LPMs and Embospheres werestatistically analyzed with T-test. The significance level was definedas P < 0.01.

2.6. Catheter deliverability

A device was used to measure the pressure on line as the LPMspassing through a catheter (Cui et al., 2012). A stable suspensionwas obtained by mixing 1 ml of LPMs with 6 ml of contrast agent(Iohexol) and saline (5:1, v/v) in a syringe. The suspension wasthen delivered through a catheter by pushing the syringe withan injection pump (Guangxi Veryark Technology Co. Ltd., China)at a speed of 2.5 ml/min. 3 Fr of catheter (lumen size 0.027 in.;Renegade; Boston Scientific) was employed to deliver LPMs in sub-group of 300–500 and 500–700 �m, respectively. And 4 Fr (lumensize 0.89 mm; Cordis; Johnson & Johnson) was employed to deliverthose in subgroup of 700–900 �m. The pressure in the catheter wasconverted into electric signals by a sensor (Powerlab, AD Instru-ments Co. Ltd., Australia) and recorded by a computer. The pressurewas measured three times for each sample.

2.7. Lipiodol content in LPMs

2.7.1. Preparation of a standard curveLipiodol of 40, 80, 120, 160 and 200 �l was dissolved in 10.0 ml

of DCM, respectively. Each lipiodol solution (0.20 ml) was removedin a test tube, diluted with 5.00 ml of DCM, and reacted with

1.00 ml of 0.20 M m-CPBA DCM solution for 1 h. Then 1.00 ml of0.20 M dimethyl sulfide DCM solution was added to quench thereaction. This organic mixture (0.30 ml) was taken out and addedto an aqueous solution consisting of 5.00 ml of 0.10 M potassium

X.-J. Lu et al. / International Journal of Pharmaceutics 452 (2013) 211– 219 213

Table 1Young’s modulus, relaxation half time (RHT), failure deformation and failure stressof LPMs and Embospheres.

Microparticles LPMs Embospheres

Young’s modulus (kPa) 80.19 ± 7.58** 158.27 ± 13.54RHT (s) 9.57 ± 1.35** 62.13 ± 1.06Residual force (%) 70.92 ± 4.03** 39.65 ± 1.63Failure deformation (%) 99.19 ± 0.02** 82.13 ± 2.30Failure stress (N) 2.28 ± 0.03** 0.45 ± 0.12

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ata was expressed as the mean ± SD.** P < 0.01 (statistical significance).

odide and 1.00 ml of 0.5% (w/v) starch. The two phase mixtureas shaken vigorously to extract iodine into the aqueous phase,

nd the absorption of aqueous phase was measured by a UV spec-rophotometer (UV-1100, Mapada Instruments Co. Ltd., Shanghai)t 553 nm (Kozutsumi et al., 2000).

.7.2. Measurement of lipiodol in LPMsIn order to measure the content of lipiodol, about 20 mg of LPMs

as weighted precisely and all lipiodol inside was extracted by0.0 ml of DCM. Then 5.00 ml of the extraction was taken into aest tube and the following measurement was same as that in theection of “preparation of a standard curve.”

.7.3. Calculation of lipiodol content in LPMsThe lipiodol content in LPMs was calculated by Eq. (2):

ipiodol loading content (mg/ml) = V1 × �

V2(2)

here V1 represents the volume of lipiodol measured in LPMs, � ispecific gravity of lipiodol, and V2 is the volume of LPMs. The specificravity of LPMs measured in this experiment was 702 mg/ml.

.8. Lipiodol release test

The release of lipiodol from LPMs was performed in a thermo-tat shaker with shaking speed of 100 rpm at 37 ± 0.5 ◦C (n = 3)..50 ml of LPMs and 1.50 ml of PBS were added in a dialysis bagmolecular mass cutoff 3500) and the bag was placed in 50.0 mlf PBS. At predetermined time intervals (12, 24, 48 and 72 h) theag was taken out and transferred into a fresh 50.0 ml of PBS. The

ipiodol released in PBS at each time interval was extracted with5.0 ml of DCM, which was then evaporated completely. The resid-al lipiodol was dissolved in 5.00 ml of fresh DCM and the contentf lipiodol was measured as same as that in the section of “prepa-ation of a standard curve.” The profile of lipiodol release with timeas plotted (Sharma et al., 2010).

.9. Radiopacity of LPMs in vitro

Vials of 2.5 ml were fully filled with LPMs, saline (negativeontrol) and lipodiol (positive control), respectively. An in vivomaging system (Carestream, Fx Pro, USA) was employed to evalu-te radiopacity of each sample.

.10. Radiopacity evaluation of LPMs in mice

Six female adult Kun Ming mice (about 30 g of initial weight,upplied by Department of Laboratory Animal Science, Pekingniversity Health Center) were anesthetized by intraperitoneal

njection of chloral hydrate (4%, w/v, 0.1 ml/10 g body weight).

PMs (0.5 ml) were suspended in 3.5 ml of 1% (w/v) sodium car-oxylmethyl cellulose, and 150 �l of the suspension was injectedubcutaneously to each of the 6 mice. As a control, the same amountf PBS was injected on the contralateral side. All animals were

wall of LPMs, mag ×3000.

observed under X-ray of the in vivo imaging system immediatelyafter injection. Then animals were housed under standard condi-tions with free access to food and water, according to the principles

of care and use of laboratory animals in China. At the 1st week, 4thweek and 3rd month post-injection, the mice were also subjected

2 l of Pharmaceutics 452 (2013) 211– 219

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14 X.-J. Lu et al. / International Journa

o X-ray imaging. All these X-ray images with time were comparedmong the same animal (Emans et al., 2005).

.11. Cytocompatibility

Cytocompatibility of the LPMs was determined by a MTT celliability assay. LPMs were sterilized with Co60 radiation for 8 h15 kGy) and subsequently incubated in 5 ml of RPMI 1640 mediumsupplementing 10% fetal bovine serum, 100 U/ml of penicillin and00 �g/ml of streptomycin) at 37 ◦C for 24 h to get 100% extracts.he extracts were diluted two times by culture medium to obtain0% extracts. Polyvinyl chloride (PVC) and cell culture mediumere used as positive and negative control, respectively. Mousebroblast cells L929 were cultured in medium at 37 ◦C, 5% CO2 in

thermostatic culture box (SANYO Electric Co., Ltd.) to obtain aonfluent cells monolayer prior to use. Cells were harvested and10,000 cells were inoculated in each well of a 96-well plate. After

he cells were cultured for 24 h, 100 �l of extracts or controls weredded to each well. At 1st, 3rd and 5th day post incubation, theedium was removed and replaced with culture medium con-

aining 0.5 mg/ml MTT to measure cell viability. The cells werencubated with MTT for 6 h at 37 ◦C. After removing the medium,he formed precipitated formazan was dissolved in DMSO. Thebsorbance of the samples was determined using a microtiter plateeader (Seac, Italy) at 500 nm. And the relative growth rate (RGR)as calculated by Eq. (3):

GR (%) = As − Ab

An − Ab× 100% (3)

here As, Ab, and An are absorption of sample, blank and negativeontrol, respectively.

The statistical differences between RGRs (%) were analyzed by-test.

.12. Histocompatibility

The mice, as described in Section 2.10, were sacrificed afteretection under X-ray at 1st day, 1st week, 4th week and 3rdonth. LPMs with surrounding tissue were excised for histolog-

cal evaluation. After a series of treatments, including being fixedn 10% neutral buffered formalin and embedded in paraffin, histo-ogical sections were obtained. Stained by hematoxylin and eosin,he above tissues were observed under microscopy (BH-2, Olym-us, Japan). The macroscopic and histopathological results werevaluated.

.13. Renal embolization of a rabbit

LPMs were employed for renal embolization of a femaleapanese rabbit (3.5 kg). The rabbit was fed standard food and

ater ad libitum, and fasted for more than 12 h before emboliza-ion. Pento-barbital sodium solution (2%, 30 mg/kg body weight)as injected via the marginal ear vein. After anesthesia, the right

emoral artery was surgically exposed and punctured. Under theuidance of digital subtraction angiography (DSA, Innova®4100,E Healthcare Technologies, USA), a 2.8 Fr catheter (Progreat®,erumo Co., Tokyo, Japan) was inserted into the left renal arteryy a guidewire. Iopamiro 370 was infused through the catheter tohow the renal vessels. Then, 0.25 ml of LPMs suspended in Iopami-

ol (1.5 ml) and saline (1.75 ml) was delivered slowly to the renalrunk. An arteriogram was followed to confirm the absence of bloodow to the kidney. Finally, the catheter was gently pulled out. Thearotid artery was ligated and the incision was closed properly.

Fig. 2. Size distribution of LPMs.

2.14. CT imaging

After embolization procedure, the rabbit was scanned by aclinical CT scanner (Somatom De finition Flash, Siemens Medi-cal Solutions, Forchheim, Germany). The imaging parameters wereas follows: field of view = 294 mm × 294 mm; matrix = 512 × 512;pixel spacing = 0.6 mm × 0.6 mm; collimation = 128 mm × 0.6 mm;pitch = 0.8; slice thickness = 1 mm; X-ray tube voltage and currentwas 100 kVp and 230 mA, respectively. A soft tissue reconstruc-tion filter was used and oblique coronal reconstruction was created(3-mm-thick reconstructed sections at 3-mm intervals).

3. Results and discussion

3.1. Morphology

Fig. 1 shows the morphology of LPMs under optical microscope(Fig. 1a) and ESEM (Fig. 1b and c). The LPMs were spherical and welldispersed. No lipiodol was observed on the surface of LPMs even invacuum (Fig. 1b). The shell thickness of LPMs was estimated about5 �m from Fig. 1c.

Spherical microparticles are considered to be easy for calibra-tion compared with irregular particles (Laurent, 2007), and theyare proved to match the cross-sectional configuration of an arterybetter. Even a single spherical particle might be able to occlude anartery of matching size appropriately (Andrews and Binkert, 2003).The compact surface of LPMs was supposed to prevent lipiodolleaking out and to keep long time radiopacity in vivo.

3.2. Size distribution

Fig. 2 shows the size distribution of LPMs. The size range ofLPMs was about 300–900 �m. The number-average diameter wascalculated as 549.5 �m.

According to the different diameters of blood vessels forembolization, the size of embolic microspheres varies from 100 �mto 2000 �m in current clinical use. The commercially availableembolic microspheres are usually separated into different sub-groups, such as 100–300, 300–500, and 500–700 �m, etc. The sizedistribution of LPMs in this study matches the clinical needs well.Furthermore, the particle size of LPMs could be adjusted by chang-ing the condition of preparation to satisfy special requirements,

such as changing the ratio of lipiodol and water, the concentrationof PVA, the string speed and so on (Bachtsi et al., 1996; Bachtsi andKiparissides, 1996; Park et al., 2001).

X.-J. Lu et al. / International Journal of Ph

Fig. 3. Compression curves of LPMs and Embospheres. Symbols: (open circle) LPMsa

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.3. Determination of elasticity

.3.1. Compression test for monolayer of microparticlesCompression curves are shown in Fig. 3. Comparing the resis-

ance of microparticles at deformation of 10%, 20%, 30%, 40% and0%, LPMs exhibited a lower compression resistance than Embo-pheres (P < 0.01). Young’s modulus of LPMs and Embospheresas calculated according to previous report (Hidaka et al., 2011)

nd shown in Table 1. The modulus was (80.19 ± 7.58) kPa and158.27 ± 13.54) kPa for LPMs and Embospheres, respectively. Ituggests that LPMs are softer and might occlude blood vessels moreistally than Embosphere.

.3.2. Stress relaxation test for monolayer of microparticlesStress relaxation curves are shown in Fig. 4. The relaxation of

PMs and Embospheres was both divided into two stages (Hidakat al., 2011): (1) in the first stage, the resistance of microparticlesecreased rapidly, which is associated with water loss of micro-articles during compression and (2) subsequently, the decayecreased to an equilibrium state and the relaxation became verylow, which mainly reflects the intrinsic viscoelastic characteristicsue to deformation and movement of polymer chains. In the case

f LPMs, the second stage is also supposed to correlate with theovement of lipiodol. The first stage lasted about 20 s and 200 s for

PMs and Embospheres separately, suggesting less water loss fromPMs because water is only absorbed in the wall of microcapsules

ig. 4. Stress relaxation curves of LPMs and Embospheres. Symbols: (open circle)PMs and (filled circle) Embospheres.

armaceutics 452 (2013) 211– 219 215

but in the entity of microspheres. The relaxation half time (RHT)and residual force for LPMs and Embospheres were calculated andshown in Table 1. The RHT of LPMs (9.57 s ± 1.35 s) was muchshorter than that of Embospheres (62.13 s ± 1.06 s). The LPMs had agreater residual force than Embospheres (P < 0.01). This may resultfrom the different structure of the two microparticles (Hidaka et al.,2011).

3.3.3. Compression test for single microparticleThe failure deformation and failure stress are shown in Table 1.

The failure deformation was (99.19 ± 0.02)% and (82.13 ± 2.30)%,and the failure stresses was (2.28 ± 0.03) N and (0.45 ± 0.12) N forLPMs and Embospheres, respectively. LPMs stood greater deforma-tion and pressure than Embospheres (P < 0.01), suggesting muchlower risk for LPMs to be broken during embolization.

3.4. Catheter deliverability

The results proved that LPMs passed through catheters withoutany difficulty. LPMs of 300–500 and 500–700 �m were easily deliv-ered through a 3 Fr catheter and LPMs of 700–900 �m were easilythrough a 4 Fr catheter. Pressure curve for LPMs passing throughcatheter is shown in Fig. 5. The pressure fluctuated within a nar-row range, suggesting no aggregation or blockage happened in thecatheter. The integral pressure was drawn and shown in Fig. 6. Theaverage pressure was greater for subgroup of 500–700 �m than300–500 �m when passing through a 3 Fr catheter. And a lowerpressure was observed in subgroup of 700–900 �m through a 4 Frcatheter.

The catheter occlusion during embolization procedure is frus-trating and dangerous (Derdeyn et al., 1995). In order to predict thepossibility of catheter occlusion, the deliverability of microparticlesor microspheres through catheter was studied qualitatively (Barret al., 1998; Derdeyn et al., 1997). For example, the degree ofresistance on injection was graded on a scale of 0–5 (Barr et al.,1998) or assessed with the acceptable criteria of pass/failure basedon whether the lumen was occluded during delivery (Laurentet al., 1996; Taylor et al., 2007). In this study, the deliverability ofLPMs was estimated on line quantitatively. The delivery test wasdesigned to simulate the clinical condition with LPMs suspendingin a mixture of contrast agent (Iohexol) and saline. All the tubesbetween instruments were stiff enough to avoid the rebound ofpressure. The limitation of this study lies in that the vascular bloodpressure was not taken into consideration as the end of catheterwas open in the air, that is, under atmosphere instead of bloodpressure. The delivery pressure measured was near the maximallimitation of the sensor (300 mmHg) and adding extra pressure wasconsidered not to be suitable to the system.

3.5. Lipiodol content in LPMs

A linear standard curve between absorption and volume of lipi-odol was calibrated: V = 355.10A − 3.356 (r2 = 0.9996). The contentof lipiodol in LPMs was (646.8 ± 17.4) mg/ml (n = 5). Compared withthe specific gravity of lipiodol (1308 mg/ml), the content of lipiodolin LPMs is relatively high, suggesting LPMs is easy to be detectableunder X-ray.

3.6. Lipiodol release test

The cumulative release of lipiodol is shown in Fig. 7. The cumu-

lative release of lipiodol at 72 h was only (1.73 ± 0.5)%, suggestinginsignificant release of lipiodol from LMPs. The long-time retentionof lipiodol in LPMs is supposed to be advantageous for providingradiopaque guide when a second embolization is necessary.

216 X.-J. Lu et al. / International Journal of Pharmaceutics 452 (2013) 211– 219

Fig. 5. Pressure curves of catheter delivery test for LPMs. (a) 300–5

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dta

Fig. 6. Integral areas of pressure curves for catheter delivery of LPMs.

.7. Radiopacity of LPMs in vitro

Fig. 8 illustrates the radiopacity of LPMs in vitro. The radiopacityf LPMs (Fig. 8c) was stronger than saline (Fig. 8a) and a little weakerhan lipiodol contrast (Fig. 8b). It reveals detectable of LPMs under-ray and a higher content of lipiodol in LPMs.

.8. Radiopacity of LPMs in mice

None of the mice showed unusual behavior, and no premature

eaths occurred during 3 months. Fig. 9 shows the X-ray images ofhe same mouse at 0 day (Fig. 9a), 1 week (Fig. 9b), 4 weeks (Fig. 9c)nd 3 months (Fig. 9d) after subcutaneous injection of LPMs. The

Fig. 7. Cumulative release of lipiodol from LPMs.

00 �m, 3 Fr; (b) 500–700 �m, 3 Fr; and (c) 700–900 �m, 4 Fr.

LPMs are proved to be radiopaque and the visibility has no reduc-tion for 3 months, which also demonstrates that lipiodol in LPMs ishardly released in vivo, suggesting LPMs can be used as permanentradiopaque embolic agent.

3.9. Cytocompatibility

The microparticles used for embolization must be safe and non-toxic (Koole et al., 2004). Therefore, cytocompatibility of LPMs wasdetermined in vitro by a MTT cell viability assay. Extracts of LPMswere cultured with mouse fibroblast cells L929 for 5 days. The mor-phology of cells was not affected by LPMs. The RGR of cells at 5thday in contact with 100% and 50% extracts of LPMs was 101.0% and94.7%, respectively (Fig. 10). According to the standard of toxicityrating, the cell toxicity of the LPMs is in grade I, which is consid-ered to be low toxic (Cao et al., 2010). The results prove that LPMsprepared in this study are cytocompatible.

3.10. Histocompatibility

According to the histopathological sections, no acute inflam-matory response was observed at 2 h (Fig. 11a) and no chronicinflammatory was found at 1 week (Fig. 11b), 4 weeks (Fig. 11c)and 3 months (Fig. 11d), suggesting biocompatibility of LPMs aftersubcutaneous injection.

3.11. Embolization study in a rabbit kidney

No microcatheter occlusion occurred during the embolizationprocedure. It is supposed due to the good elasticity, spherical shapeand smooth surface of LPMs.

Fig. 8. X-ray images of saline (a), lipiodol (b) and LPMs (c) in vials.

X.-J. Lu et al. / International Journal of Pharmaceutics 452 (2013) 211– 219 217

Fig. 9. X-ray images of the same mouse at 1st day (a), 1st week (b), 4th week (c)and 3rd month (d) after subcutaneous injection of LPMs.

Fig. 11. Histological features of mice after subcutaneous injection of LPMs at 1st da

Fig. 10. Relative growth rate (%) of L929 cells in different mediums.

Fig. 12 shows the angiograms of the left kidney before and afterembolization. LPMs were considered successfully embolized intothe renal artery and mainly located in proximal part of it. Relativelarge size of LPMs (300–500 �m) was used to a rabbit kidney inthis experiment, so proximal artery was occluded as described in aprevious report (Wilson et al., 2003).

In general, the position of radiopaque microspheres can behardly observed under DSA. It was reported that clusters ofradiopaque microspheres with higher density of iodine were moreeasily to be detected (Namur et al., 2007; Sharma et al., 2010). Inthis study, the iodine content of LPMs was high up to (35.9 ± 1.0)%(w/w), but the volume of LPMs was low down to 0.25 ml for a rabbitkidney. Thus, CT was employed to detect the radiopacity of LPMsafter embolization.

3.12. CT imaging

Fig. 13 presents the CT imaging of the rabbit. As shown in Fig. 13,the LPMs provided a clear contrast to the surrounding tissue and

y (a), 1st week (b), 4th week (c) and 3rd month (d). The arrows point at LPMs.

218 X.-J. Lu et al. / International Journal of Pharmaceutics 452 (2013) 211– 219

Fig. 12. Arterial angiogram of a rabbit’s left kidney, (a) before embolization and (b)immediately after embolization with LPMs. The renal artery trunk was occludedwith LPMs after embolization.

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radiol. 21, 666–669.Bilhim, T., Pisco, J.M., Duarte, M., Oliveira, A.G., 2011. Polyvinyl alcohol particle size

for uterine artery embolization: a prospective randomized study of initial use of350–500 �m particles versus initial use of 500–700 �m particles. J. Vasc. Interv.Radiol. 22, 21–27.

ere mainly located in the renal artery trunk of the kidney. Theesult coincided with that of DSA. Thus, the LPMs are proved toe easily detected in vivo by CT imaging after embolization. Theetection of embolic agents is considered to be further improvedy using a more sensitive X-ray imaging system, such as micro-CTystem (Sharma et al., 2010).

DSA combined with CT is current trends of diagnosis and treat-ent in embolization technology (Kakeda et al., 2007; Tognolini

t al., 2010). In this situation, advanced radiopaque embolic agents certainly required to follow up the development of medical sci-nce. The above primary embolization study in a rabbit has showed

good result and prospect to LPMs. Still, further study shoulde performed in the future, with more animals and for longerime.

Fig. 13. CT images of the left kidney. The density of left renal artery trunk wasincreased by LPMs.

4. Conclusion

A clinical available X-ray diagnostic agent, lipiodol, wasphysically encapsulated into PVA microcapsules using a simplecoacervation method. The LPMs have achieved multiple require-ments for embolization, such as in spherical shape and reasonablesize range, with higher content of lipiodol, good compressibilityand biocompatibility. The radiopacity and embolization feasibilityof LPMs have been proved by initial animal experiments in miceand rabbits, respectively. In conclusion, the LPMs showed to bepotential radiopaque agent for embolization therapy.

Acknowledgment

This research was supported by the State Key Projects (No.2009ZX09310-001).

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