Controllable Formation of Monodisperse Polymer Microbubbles as

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Biological and Medical Applications of Materials and Interfaces

Controllable formation of monodisperse polymer microbubbles as ultrasound contrast agents Ruyuan Song, Chuan Peng, Xiaonan Xu, Jianwei Wang, Miao Yu, Youmin Hou, Ruhai Zou, and Shuhuai Yao ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b17258 • Publication Date (Web): 11 Apr 2018 Downloaded from http://pubs.acs.org on April 11, 2018

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ACS Applied Materials & Interfaces

Controllable formation of monodisperse polymer microbubbles as ultrasound contrast agents Ruyuan Song,a‡ Chuan Peng,c‡ Xiaonan Xu,b Jianwei Wang,c Miao Yu,b Youmin Hou, b Ruhai Zou,*c and Shuhuai Yao*ab

a

Bioengineering Graduate Program, Department of Chemical and Biological Engineering, The

Hong Kong University of Science and Technology, Hong Kong, China. b

Department of Mechanical and Aerospace Engineering, The Hong Kong University of Science and Technology, Hong Kong, China.

c

State Key Laboratory of Oncology in South China, Collaborative Innovation Center of Cancer

Medicine, Department of Ultrasound, Sun Yat-sen University Cancer Center, Guangzhou, China.

ABSTRACT: Microbubbles have been widely used as ultrasound contrast agents in clinical diagnosis and hold great potential for ultrasound-mediated therapy. However, polydisepersed population and short half-life time (3 times) for ultrasound imaging in comparison with the commercial lipid microbubbles.

KEYWORDS: polymer microbubbles, microfluidics, ultrasound contrast agent, stability, monodisperse size distribution

INTRODUCTION Ultrasound imaging is widely used for medical diagnosis of internal organs, muscles, and the blood circulatory system, thanks to its inherent merits such as noninvasiveness, nonradiation, real-time monitoring, relatively low cost, and wide availability.1-3 The employment of ultrasound contrast agents has improved the capabilities of ultrasound as a powerful imaging tool for diagnosis and provided a promising strategy for image guided drug delivery.4 Currently, commercial ultrasound contrast agents such as SonoVue®, Definity® and Optison® are gaseous octafluoropropane or sulfur hexafluoride microbubbles stabilized by phospholipids or albumin with a typical diameter of 1-8 µm.5-6 However, these kinds of microbubbles suffer from the stability issues such as bubble

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coalescence and gaseous core dissolution, especially under ultrasound irradiation, resulting in limited half-life time in circulation (e.g., less than 10 min).7-11 To enhance the stability of microbubbles, materials with higher mechanical strength than phospholipids have been developed to stabilize the gaseous cores including silica12 and polymers13-15, which is called hard-shelled microbubbles. Hard-shelled microbubbles hardly present volume expansion and remained intact at a low acoustic pressure. Above a certain pressure threshold, the shell of the hard-shelled microbubbles encounters ruptures.16 Among hard-shelled microbubbles, the biodegradable polymers, e.g., poly (lactic-coglycolic acid) (PLGA)and polylactide (PLA)17are most abundantly used for producing hard-shelled ultrasound contrast agents. The microbubbles with polymer shells not only possess much better stability with greatly improved resistance to pressure and mechanical variations under ultrasound but also enrich the versatility of the microbubbles with functional agents by grafting or encapsulation of therapeutic drugs for drug delivery, sensitizers for ultrasound image-guided photothermal therapy and sonodynamic therapy18 or combining other contrast agents for dual mode imaging and therapy.19-22 The polymer microbubbles or nanobubbles are typically fabricated by using water-in-oil-in-water (W/O/W) emulsion as a template through emulsion-solvent evaporation method.23-24 After the solvent is evaporated, polymers in the oil phase of a mixture of polymer/volatile organic solvent precipitate out to form a shell around the water core which would be


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removed subsequently for gas infusion. Alternatively, Kooiman et al. fabricated polymer microbubbles using a combination of O/W emulsion-solvent evaporation and internal phase separation.17 The sacrificial cores are created by hexdecane/octane oils which are added along with polymers in volatile organic solvent prior to emulsification. However, the emulsification methods for preparing polymer microbubbles including sonication and homogenization usually lead to broad size populations from one to tens of micrometers, which highly compromise the performance of the polymer microbubbles as ultrasound contrast agents for the following reasons: First, oversized microbubbles fail to pass through the pulmonary capillary vessels freely, which may even cause serious local embolism. Second, since both the resonance frequency for imaging and trigger frequency for therapy with ultrasound are size-dependent, monodisperse microbubbles yield higher sensitivity of ultrasound imaging due to their homogeneous echogenic behaviors,25-26 while only a small portion of polydispersed microbubbles fit the detection window of the clinical ultrasound diagnostic system with a limited frequency bandwidth.9 Therefore, monodispersity is a critical measure for ultrasound contrast agent development. Recently, the utilization of microfluidic systems for droplet generation has attracted substantial attention by virtue of their capability to accurately control over the size, shape, and components of emulsions.27-28 Monodisperse core-shell microcapsules have been extensively fabricated by W/O/W double emulsion method.29


For example,

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Abbaspourrad et al. developed monodisperse pH-responsive polymer microcapsules using W/O/W emulsion as the template via a coaxial microcapillary fluidic device.30 Despite the promise of a high degree of control over the size and composition using the double emulsion method in microfluidic devices, the produced microcapsules are usually over 50 μm in diameter, much larger than the optimal size (2-5 μm in diameter)31 of the microbubbles used as ultrasound contrast agents. Alternatively, monodisperse oil-filled PLA microcapsules were fabricated using a combination of O/W emulsion-solvent evaporation and internal phase separation via microfluidic emulsification.

32-33

Since

monodisperse single emulsion synthesis can be well controlled from hundreds of microns to even submicron in diameter using flow focusing microfluidic devices,34-35 therefore, the O/W emulsion-solvent evaporation method is more feasible to be implemented in microfluidics to produce monodisperse polymer microbubbles in optimal size range of 25 μm in diameter. In this paper, we present a microfluidic flow focusing droplet generation strategy to produce monodisperse microbubbles with biodegradable polymer PLGA shells as ultrasound contrast agents. The polymer microbubbles prepared by microfluidics have merits of controllable size, narrow size distribution and good stability. The potential of polymer microbubbles as ultrasound contrast agents was examined in vitro and in vivo. The drug loading capacity of the polymer microbubbles and ultrasound-mediate release



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was also examined, showing the potential use of polymer microbubbles for ultrasound imaging-guided therapy. MATERIALS AND METHODS Materials. Perfluorooctyl bromide (PFOB), poly(D, L-lactide-co-glycolide) (RG502H, MW 7,000-17,000), poly(ethylene glycol) bis(amine) (PEG-bis(amine), MW ~3,400), polyvinyl alcohol (PVA) (MW 30,000-70,000, 89% hydrolyzed), Nile red, sodium cholate,

N-(3-dimethylaminopropyl)-N’-ethylcarbodiimihydrochloride

(EDC),

N-

hydroxysuccinimide, trimethylamine(TEA) and chloroform-d (CDCl3) were purchased from Sigma-Aldrich (St. Louis, MO). Perfluoropropane (PFP) gas was purchased from Aoborui (Tianjin, China). Paclitaxel (PTX) was purchased from Meilunebio (Dalian, China). Dichloromethane (DCM), glycerol and sulfoxide (DMSO) were obtained from Acros Organics. Aqueous solutions were all prepared in Milli-Q deionized water (18 MΩcm-1 deionized water, Millipore CO., MilliQ system). Microfluidic device fabrication. A typical flow focusing device, composed of a central channel and two side channels with 10 μm wide nozzles, was designed for droplet generation.34 All microchannels were etched in a 525-µm-thick silicon wafer using deep reactive-ion etching (DRIE), resulting in 10 μm in depth. The sample feeding and collecting holes were etched through the wafers using tetramethylammonium hydroxide (TMAH) etching with a 3500 Å silicon oxide layer as the mask. The silicon wafer was


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bonded with a 170-µm-thick Pyrex glass (SENSOR Prep Services, Inc.) using anodic bonding and individual devices were then diced using a wafer cutting machine. Amphiphilic polymer PLGA-PEG synthesis. For the PEGylation of polymer microbubbles, amphiphilic biodegradable polymer PLGA-PEG was synthesized by coupling PLGA and PEG-bis(amine) following the synthesis routine (Figure S1a). Briefly, DCM was dried over activated 4Å molecular sieves for three days before use. 600 mg of PLGA, 13.5 mg of NHS, and 22 mg of EDC were dissolved in 10 ml of dry DCM and stirred under nitrogen atmosphere for 24 h at room temperature for activation of the carboxyl group. Then, the PLGA-NHS solution was added dropwise into 5ml of dry DCM containing 680 mg of PEG-bis(amine) and 50 μl of TEA and stirred for another 24 h under nitrogen atmosphere. After the reaction was complete, DCM was evaporated and 5 ml of dimethyl sulfoxide (DMSO) was added. The mixture was transferred into the dialysis bag (MWCO, 7k) and dialysed against water for 72 h to remove the unreacted PEG-bis(amine) and other small molecules. Finally, the product was lyophilized to obtain white powders. The PLGA-PEG was verified by 1H NMR spectroscopy (Mercury VX 300, Varian UK) in CDCl3. The 1H NMR spectrum of PLGA-PEG in CDCl3 is shown in Figure S1b. Preparation of polymer microbubbles. Monodisperse polymer microbubbles with PEGylated surface were prepared by a two-step process. First, a mixture of perfluorooctyl



bromide (PFOB) and PLGA/PLGA-PEG polymer dissolved in DCM was emulsified into monodisperse precursor microdroplets using a flow focusing device, as illustrated in Figure 1a. Second, followed by solvent diffusion and internal phase separation, further solvent evaporation, freezing-drying, and infusion of PFP gases, monodisperse precursor microdroplets turned into uniform polymer microbubbles (Figure 1b). The flow focusing device consisted of one central inlet for the dispersed phase and two side inlets for the continuous phases. The disperse phase is a mixture of PLGA-PEG/PLGA, PFOB and DCM. To prepare the PFOB and polymer mixture in DCM, 20 μl of PFOB, along with the desired amount of mixture PLGA/PLGA-PEG (typically, 10 mg PLGA/PLGA-PEG with a mass ratio of 8/2) was dissolved in 1 ml DCM, sonicating in an ultrasonic cleaner for 20 min to ensure the solution was fully mixed. The continuous phase was 1.5 w/v% sodium cholate aqueous solution with 10 w/v% of glycerol (for increase of the viscosity of the continuous phase). For fluorescence imaging, Nile red (10 μg/ml in DCM) was added to the dispersed phase for labelling of formed microcapsules. The fluid flow in the microfluidic device was driven by a home-made pressure controller system with 0-60 psi range. During the operation, the microfluidic device was mounted on a Nikon microscopy equipped with a camera to record the droplet generation under different flow conditions. The generated droplet solution was collected into a 5 ml glass vial with magnetic stirring at 300 rpm for solvent evaporation. For 3 hours till DCM was completely evaporated,


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microcapsules were collected after being centrifuged at 1000 x g for 5 minutes, and washed three times with water. 0.2 w/v% PVA solution was added into the microcapsules as the cryoprotectant and then lyophilized for 3 days to remove the water and sacrificial PFOB cores. Then, the hollow microcapsules were transferred into a 5 ml glass vial sealed with a rubber septum. The air in the glass vial was replaced with PFP gas in a home-made gas-exchange system consisting of a three-way valve which was connected with the vacuum line, the sample bottle, and the PFP gas bottle, respectively. The PFP gas-filled microbubbles with polymer shells were stored at 4 ℃ for further use. Characterization of polymer microbubbles. A confocal scanning microscope (Zeiss LSM-710, Carl Zeiss, Germany) with a 63x objective lens was used for fluorescence imaging of the microcapsules stained with Nile Red. The concentration of the polymer microbubbles was measured using a hemocytometer. Scanning electron microscopy (SEM, JSM-6700F, JEOL, Japan) was used to examine the morphology of the microcapsules, operating at 10 kV with a filament current about 20 μA. A small drop (~5 μl) of samples was deposited on a small piece of a clean silicon wafer and then dried overnight. Then the silicon substrate was mounted on a copper holder using carbon conductive double-sided tape. Before observation, the surface of the samples was coated by a layer of palladium-platinum layer.



Drug encapsulation and release. To demonstrate the drug loading capacity, the desired amount of PTX was also dissolved into DCM with various ratios of the polymer and PFOB in forming the precursor microdroplets.

1 ml of precursor microdroplets

solutions containing 1 mg polymer was centrifuged at 5000 x g for 30 min and then 0.5 ml of the supernatant was collected for lyophilization. Then, 200 μl of DMSO was added to dissolve the free PTX in the supernatant after lyophilization. The DMSO solution was then transferred into a 96-well plate. The amount of the free PTX in the supernatant was determined via UV absorbance at 254 nm using a multimode microplate reader (Varioskan LUX, Thermo Fisher Scientific, USA). The PTX encapsulation efficiency (EE) is defined as: Encapsulation efficiency =

weight of the drug in the polymer microbubbles weight of the feeding drug

And the PTX loading efficiency is: Loading efficiency =

weight of the drug in the polymer microbubbles weight of the polymer microbubbles.

The polymer microbubbles prepared by using10 mg/ml polymer and 0.5 mg/ml PTX were used for drug release profile measurement. 2 ml of the PTX-loaded polymer microbubbles (2.5 mg/ml) in a sealed dialysis tube (MWCO: 8000 Da) was exposed to ultrasound irradiation (1 MHz, 1000 KPa) for 20 min by an ultrasonic cavitation machine (Welld, Shenzhen, China). Then, the dialysis tube was immersed into 50 ml release medium (30 v/v% ethanol, 0.01w/v % Tween-80). At each measurement time, 1 ml of


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dialysis buffer was collected for lyophilization and 1 ml of new buffer was added. The PTX amount of the samples collected at each measurement time was determined using the same methodfor drug encapsulation. In vitro ultrasound imaging. In vitro ultrasound imaging of microbubbles was performed in a gel phantom using an ultrasound system (Acuson S2000, Siemens, USA) with a probe 9L4 (4-9 MHz, Siemens, US). The gel phantom was prepared with 2 w/v % agarose and the sample loading wells in the gel phantom were prepared by using a mold of 1.5 ml centrifuge tubes. 1 ml of the polymer microbubbles in a series of concentrations (~1.3 to ~5.0×108 bubbles/ml) was loaded into the sample loading wells of the gel phantom, immersed in a water tank at 37 °C, 2 cm far from the ultrasound transducer. The ultrasound images were typically acquired in tissue harmonic imaging (THI) mode at a frequency of 8 MHz and a mechanical index (MI) of 0.2 at specific time intervals. In vivo ultrasound imaging. To assess the microbubbles as ultrasound contrast agents in vivo, ultrasound imaging was performed on the kidney of 6-week nude mice. The mice were anesthetized using 2% isoflurane, and then injected with 200 μl of PBS solution, polymer microbubble solution (~1.0×109 bubbles/ml), SonoVue microbubble solution (~1.1×108 bubbles/ml) through the tail vein.

And mice were maintained under 2%

isoflurane anesthesia during the ultrasound scanning period. After injection, the kidney of



the mouse was monitored and imaged by Acuson S2000 ultrasound imaging system in harmonic mode at a frequency of 8 MHz and MI value of 0.2. RESULTS AND DISCUSSION Fabrication and characterization of polymer microbubbles. Monodisperse polymer microbubbles with PEGylated surface were prepared by a two-step process of microfluidic emulsification and PFP gases infusion. Using the flow focusing device, the disperse phase was emulsified to form monodisperse precursor microdroplets at generation rates from ~6 kHz to 20 kHz (Figure 2a). It typically took less than three hours to collect a clinically-relevant dose (e.g., 2.0 ×108 bubbles) for one animal test using a single device. Higher throughput can be achieved for droplet generation at generation rates up to hundreds of kHz by parallelization of such devices,36 or direct production of gas-filled (e.g., perfluorobutane) bubbles at even higher production rates up to 1 MHz in a single flow focusing device.37 As the DCM in the precursor microdroplets diffused into the surrounding aqueous phase, the PFOB in the DCM solvent became supersaturated and nucleated into tiny droplets inside the precursor microdroplets (Figure 2b). Those tiny droplets then immigrated into the center and fused into a single larger one with the assistance of surfactants. After DCM was completed evaporated, PLGA/PLGA-PEG polymers precipitated out and formed a solid shell around the PFOB core. The core-shell structure of the final microcapsules was confirmed by the confocal images of the resultant


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microcapsules with Nile red labelling (Figure 2c). The precursor microdroplets and resultant microcapsules had average diameters of ~10.7 μm and ~3.2 µm respectively with a very narrow size distribution, as shown in Figure 2d. The coefficient of variation (CoV), defined as the standard deviation of the droplet size population divided by the average droplet size, was as low as 4 % for the precursors, and less than 5% for the resultant microcapsules. Amphiphilic polymer PLGA-PEG/PLGA mixture was chosen for the PEGylation of surfaces to enhance the stability of the polymer microbubbles38 and suppress the clearance from the mononuclear phagocyte system after administration.39 The mass ratio of PLGA-PEG in the polymer mixture was kept down to the 0.2 since too high portion of PLGA-PEG may induce protrusions (small bumps) on the microbubble surfaces for all PEG chains in the hydrophilic environment.40 The continuous phase was the aqueous solution with sodium cholate as the surfactant, since sodium cholate was an efficient surfactant for controlling the microcapsule morphology to obtain the core-shell structure.41 The size of droplets generated by the microfluidic device was tuned by changing the pressures applied on the disperse phase of polymer/PFOB mixture, P ,and the continuous phase of 1.5 w/v% sodium cholate in water, P . Glycerol was added into the aqueous phase (10 w/v%) to increase its viscosity (~ 30% higher than 1.5 w/v sodium cholate aqueous solution in viscosity), which enabled to obtain ultra-small droplets as small as ~2



μm . As the viscosity of continuous phase increased, stronger shear stress by the continuous phase was applied to deform the interface to form a narrower thread, resulting in smaller droplets.42 We fixed the pressure applied on the disperse phase (P ) at 8 psi and varied the pressure applied on the continuous phase (P ) from 15 to 40 psi to decrease the droplet size (Figure 3a). The droplet size was tuned from ~21 μm to ~2 μm in diameter in the range of pressure adjustment (Figure 3b). After the solvent evaporation, the droplets shrunk into the polymer-shell microcapsules about 3 times smaller than the precursor droplets, as shown in Figure 3b, which was very close to the theoretical calculation using Equation S1. The final resultant microcapsules were ~ 0.6 to 7µm in diameter. We also obtained polymer microbubbles with shells of different thickness by changing the concentration of polymer PLGA/PLGA-PEG from 10 mg/ml to 40 mg/ml while keeping the concentration of PFOB fixed at 20 μl/ml. The confocal microscope images (Figures 4a and 4b) show the shell thickness increased from ~ 300 nm to ~ 1 μm as the T/R, defined as the ratio of the thickness of the polymer shell to the radius of the whole microcapsule), varied from 0.10 to 0.26, suggesting that the T/R increased with the increase of the polymer concentration. In addition, the SEM images show that the morphology of the microcapsules differed slightly for different polymer concentrations. With a higher polymer concentration (40 mg/ml), small holes were found in the surface of


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the polymer shells while a lower polymer concentration (10 mg/ml) yielded a much smoother surface, as shown respectively in Figure 4c and Figure 4d. The small holes in the surface of the polymer matrix may be formed by the faster polymer precipitation, resulting from a higher polymer concentration of the precursor droplets during solvent diffusion, which trapped tiny PFOB droplets before they immigrated into the center. After the entrapped PFOB droplets were sublimed by lyophilization, small cavities were left in the surface of the shell. This phenomenon has been observed in use of ethyl acetate with fast diffusion rate, which also leads to fast precipitation of polymer.33 These small holes may deteriorate the stability of the polymer microbubbles since the entrapped gas may easily dissolve in solutions through the small holes. To prove the drug loading capacity of the polymer microcapsules, PTX, one of most commonly used anti-cancer chemo-therapeutic agents, was added into the organic solvent along with polymers and PFOB for the preparation of the drug-loaded polymer microbubbles. The PTX encapsulation efficiency remained very high (>86.5 %), albeit it decreased a bit with the decrease of the polymer concentration for the polymer microbubbles with thinner shells (Table 1). The PTX release of PTX-loaded polymer microbubbles was accelerated by ultrasound irradiation as shown in Figure 5. In the first 12 hours, the accumulative drug release with/without ultrasound irradiation was 32.5 % and 15.2 %, respectively. After 12 hours, the drug release rate became slower.



In vitro ultrasound image. To evaluate capacity of the polymer microbubbles as ultrasound contrast agents, the ultrasound imaging tests were conducted in vitro at first. We injected the degassed PBS solution into the lyophilized polymer microbubble power, stored in a septum sealed vial with PFP gas filled at the headspace of the vial and gently shook the vail to obtain the opaque solution. The diameter of microcapsules used for the ultrasound imaging test was around 3.2 μm which lied in the range of the optimal size of 2 – 5 μm for ultrasound contrast agents.31

We first investigated the effect of the

ultrasound frequency on the ultrasound signal intensity of the polymer microbubbles. Bright ultrasound images were acquired at frequencies ranging from 4 to 8 MHz and the brightness increased gradually with the increase of frequency (Figures 6a and 6b), which may be resulted from the reduced mismatch between the resonance frequency of the microbubbles and the frequency of the driving ultrasound. The MI value essentially determines he ultrasound signal intensity as ultrasound with higher energy drives the polymer microbubbles to oscillate more strongly. As shown in Figures 6c and 6d, the ultrasound signal intensity of the polymer microbubbles increased as the increase of MI from 0.05 to 0.4. Figures 7a and 7b show that the brightness of the ultrasound images was highly dependent on the concentration of the microbubbles. Compared to the PBS control, the average grayscale of the ultrasound images with polymer microbubbles increased significantly from 100% to 800% with the increase of the concentration of the polymer


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microbubbles (from ~1.3 to ~5.0 × 108 bubbles/ml). We also examined the echogenic properties of the polymer microbubbles with different T/R ratios, varying from 0.10 to 0.26. The ultrasound signal decreased to 1/3 of its initial ultrasound signal intensity with the increase of the shell thickness from ~300 nm to ~ 1000 nm (Figures 7c and 7d). The polymer microbubbles with thinner shells were more compressible, and therefore produced stronger ultrasound signals.

41

And the degree of mismatch between the

frequency of the driving ultrasound and the resonance frequency of polymers microbubbles, resulting from the different shell thicknesses may also contribute to the ultrasound signal variations. Furthermore, the stability of the polymer microbubbles after the ultrasound irradiation was also investigated and compared with the SonoVue lipid microbubbles. The ultrasound images of the polymer microbubbles and SonoVue lipid microbubbles at different time intervals are presented in Figure 8a.The gray value of the SonoVue microbubbles was initially larger (~1.7 times) than that of the polymer microbubbles, because lipid microbubbles with more compressive shells could oscillate in greater amplitude than polymer microbubbles under ultrasound and therefore have higher echogenicity.16 However, the ultrasound signal produced by the SonoVue mcirobubbles declined sharply with a decrease of ~ 80% of their initial intensity after ~3 min as shown in Figure 8b, while the ultrasound signal intensity of the polymer microbubbles remained for ~20 min with only 30% decrease from its initial ultrasound signal intensity (Figure



8b). The enhanced stability of the polymer microbubbles is attributed to the firm polymer shells (~ 300 nm in thickness) which provides a robust barrier against the dissolution of PFP gas entrapped inside under ultrasound irradiation, while the thickness of the lipid monolayer of SonoVue microbubbles is ~4 nm,43 through which much faster dissolution of gases can be achieved under ultrasound irradiation. Furthermore, the filled gas PFP in the polymer microbubbles has lower water solubility than that of the SonoVue microbubbles, which may also contribute to the enhanced stability of the polymer microbubbles. We further confirmed that more than 90 % of the polymer microbubbles remained intact after the ultrasound exposure, as manifested by the optical microscope images of the polymer microbubbles in Figures 8c and 8d, whilst only ~5 % of SonoVue microbubbles were left after ultrasound exposure (Figures 8e and 8f). Such the elongated lifetime of the polymer microbubbles renders a longer imaging window for clinical diagnosis. The good stability of the polymer microbubbles may have more advantages in delivering therapeutic gases with higher solubility in water, like oxygen, which suffer from more serve dissolution than perfluorocarbon gases in water. In vivo ultrasound imaging. In vivo ultrasound imaging were performed using nude mice and ultrasound images of kidney were acquired. In the control experiment using PBS solution without the polymer microbubbles, the kidney region remained dark in the ultrasound image, as shown in Figure 9a. Compared with the PBS control (Figure 9a),


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eminently contrast enhancement of the kidney region began to appear in the ultrasound images after intravenous administration of both the SonoVue microbubble and polymer microbubble solution (Figure 9b and Figure9d). The bright ultrasound images of kidney indicate that the polymer microbubbles successfully pass through the pulmonary capillary vessels during the circulation, which is essential for safe performance of venous contrast agents, thanks to tight size-control by the microfluidic method. The ultrasound signal intensity of SonoVue microbubbles group was stronger than that of the polymer microbubbles group at the beginning. Since the SonoVue microbubbles with flexible lipid shells are easily compliant to area expansion and compression with minimal damping, resulting in higher echogenicity. However, ultrasound imaging time of the polymer microbubbles (Figures 9e and 9f) lasted longer than that of the SonoVue microbubbles (Figure 9b). And the polymer microbubbles achieved good contrast performance at a moderate MI of 0.2, much less than the maximum MI of 1.9 in clinical examinations that Food Drug Administration (FDA) ultrasound regulation allows. The excellent performance and stability of the polymer microbubbles prove their great potential as ultrasound contrast agents for ultrasonography of superficial organs. 9 CONCLUSION We have developed a facile microfluidic method to fabricate monodisperse polymer microbubbles as ultrasound contrast agents to tackle the limitations of current commercial



lipid microbubbles including poor stability and polydispersed size population. The size, size distribution, and shell thickness of the polymer microbubbles can be tightly tuned via microfluidic operation. The polymer microbubbles demonstrated excellent ultrasound contrast performance in vitro and in vivo ultrasound imaging experiments. Furthermore, the polymer microbubbles presented much better stability as compared to the SonoVue lipid microbubbles at the high frequency and MI, rendering a longer imaging window for clinical diagnosis. In addition, polymer microbubbles have also shown high capacity of the therapeutic agent encapsulation and ultrasound control release. Therefore, such monodisperse polymer microbubbles may hold great potential in ultrasound imaging and ultrasound imaging-guided therapy.

ASSOCIATED CONETENT Supporting Information. Synthesis routine and 1H NMR spectrum of amphiphilic copolymer PLGA-PEG, SEM images of the submicron polymer bubbles, and the theoretical calculation of the ratio of diameter of precursor droplets to the diameter of resultant microcapsules (Ddroplet/Dcapsule) and the ratio of the shell thickness to the radius of the microcapsules (T/R).

AUTHOR INFORMATION


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Corresponding Authors: E-mail: [email protected] E-mail: [email protected]

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ‡These authors contributed equally.

Notes There are no conflicts to declare. ACKNOWLEDGEMENTS This work was financially supported by Research Grants Council of Hong Kong under General Research Fund (Grant No. 16206915) and Guangdong - Hong Kong Technology Cooperation Funding Scheme (Grant No. 2017A050506020).

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19. Ao, M.; Wang, Z.; Ran, H.; Guo, D.; Yu, J.; Li, A.; Chen, W.; Wu, W.; Zheng, Y. Gd-Dtpa-Loaded PLGA Microbubbles as Both Ultrasound Contrast Agent and Mri Contrast Agent—a Feasibility Research. J. Biomed. Mater. Res. B Appl. Biomater. 2010, 93, 551-556. 20. Sun, Y.; Zheng, Y.; Ran, H.; Zhou, Y.; Shen, H.; Chen, Y.; Chen, H.; Krupka, T. M.; Li, A.; Li, P. Superparamagnetic PLGA-Iron Oxide Microcapsules for Dual-Modality Us/Mr Imaging and High Intensity Focused Us Breast Cancer Ablation. Biomaterials 2012, 33, 5854-5864. 21.

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Figure 1. Schematic of generation of monodisperse precursor microdroplets and formation of polymer microbubbles. (a) Precursor microcroplets are generated in a microfluidic device with PFOB and PLGA/PLGA-PEG in DCM as the disperse phase and sodium cholate aqueous solution as the continuous phase, inset is an optical microscope image of droplet generation (scale bar: 100 µm); (b) Illustration of the process of polymer microbubble formation from precursor microdroplets.



Figure 2. Optical microscope images of (a) monodisperse precursor microdroplets, (b) immigration of tiny PFOB droplets in precursor microdroplets, and (c) core-shell microcapsules with polymer shells labelled with Nile red. (d) Size distribution of the droplet precursors and resultant microcapsules. The microcapsules were prepared in a microfluidic device operated at P = 8 psi and P = 25 psi. The disperse phase consisted of 20 µl/ml PFOB and 10 mg/ml PLGA/PLGA-PEG ( mass ratio: 8:2) in DCM and the continuous phase consisted of 1.5 w/v% sodium cholate and 10 w/v % glycerol in water.


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Figure 3. (a) Optical microscope images of droplet generations under different flow conditions. (b) The precursor droplet size, the resultant microcapsule size, and the ratio of the precursor droplet size to resultant microcapsule size (Ddroplet/Dcapsules) under different flow conditions. The pressure applied at the disperse phase P = 8 psi and the pressure applied at the continuous phase was changed from 15 to 40 psi. The disperse phase consists of 20 μl/ml PFOB and 10 mg/ml PLGA/PLGA-PEG (mass ratio: 8:2) in DCM and the continuous phase consists of 1.5 w/v% sodium cholate and 10 w/v % glycerol in water.



Figure 4. Confocal microscope and SEM images of polymer microbubbles of different shell thickness-to-radius (T/R) ratios. The polymer microbubbles were prepared at P = 8 psi and P = 25 psi. The disperse phase consisted of 20 μl/ml PFOB and 10 to 40 mg/ml PLGA/PLGAPEG (mass ratio: 8:2) in DCM and the continuous phase consists of 1.5 w/v% sodium cholate and 10 w/v % glycerol in water. Confocal microscope images of polymer microbubbles labelled with Nile red of T/R = 0.1 (a) and 0.26 (b). SEM images of polymer microbubbles of T/R = 0.1 (c) and 0.26 (d).


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Figure 5. In vitro drug release profiles of the PTX-loaded microbubbles without ultrasound irradiation and with ultrasound irradiation (1 MHz, 1000 KPa, 20 min) for 3 days. The polymer microbubbles were prepared using the dispersed phase made of 10 mg/ml polymer and 0.5 mg/ml PTX.



Figure 6. Ultrasound images of the polymer microbubbles at different frequency and different MI values. (a) Ultrasound images of the polymer microbubbles at frequency varying from 4 to 8 MHz (MI = 0.2, concentration = 5.0 # 10% bubbles/ml). (b) The average gray values of the images obtained from panel a. (c) Ultrasound images of the polymer microbubbles at different MI varying from 0.05 to 0.4 (Frequency = 8 MHz, concentration = 5.0 # 10% bubbles/ml) (d) The average gray values of the images obtained from panel b.


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Figure 7. Ultrasound images of the polymer microbubbles at different concentrations and with different shell thickness-to-radius(T/R) ratios (frequency = 8 MHz and MI = 0.2). (a) Ultrasound images of the polymer microbubbles at different concentrations from 1.3 to 5.0 # 10% bubbles/ml (T/R ratio = 0.10, diameter = 3.2 μm). (b) The average gray values of the images obtained from panel a. (c) Ultrasound images of the polymer microbubbles with different shell thickness-to-radius(T/R) ratios (concentration = 5.0 # 10% bubbles/ml). (d) The average gray values of the images obtained from panel c.



Figure 8. (a) Ultrasound images of the polymer microbubbles (PMs) and SonoVue microbubbles at different time intervals (Frequency = 8 MHz, MI = 0.2). (b) Normalized average gray values of ultrasound images of the polymer microbubbles and SonoVue microbubbles as a function of the time. The average gray values were normalized by the initial value. Optical microscope images of the polymer microbubbles before (c) and after (d) 20 min ultrasound irradiations. Optical microscope images of the SonoVue microbubbles before (e) and after (f) 20 min ultrasound irradiation.


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Figure 9. In vivo ultrasound images of the mouse kidney after intravenous injection of the PBS, SonoVue microbubble, and polymer microbubble solutions (Frequency = 8 MHz, and MI = 0.2). Ultrasound images of the mouse kidney after intravenous injection of PBS (a); Ultrasound imaged of the mouse kidney after intravenous injection of the SonoVue microbubbles at different time intervals of 10 sec (b) and 2 min (c); Ultrasound images of the mouse kidney after intravenous injection of the polymer microbubbles at different time intervals of 1 min (d), 4 min (e) and 8 min(f). The red circles indicate the kidney regions of the mouse.



Table 1. PTX encapsulation efficiency (EE) and loading efficiency at different polymer concentrations and PTX concentrations. Polymer

PTX

EE

loading efficiency

(mg/ml)

(mg/ml)

/%

/%

40

1

98.5±1.2

2.46±0.04

20

0.5

93.5±1.2

2.67±0.06

10

0.5

86.5±1.5

4.31±0.15


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Controllable Formation of Monodisperse Polymer Microbubbles as

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