- Email: [email protected]
Ultrasound-triggered PLGA microparticle destruction and degradation for controlled delivery of local cytotoxicity and drug release
Accepted Manuscript Title: Ultrasound-triggered PLGA Microparticle Destruction and Degradation for Controlled Delivery of Local Cytotoxicity and Drug Release Authors: Kee W. Jang, Dongrim Seol, Lei Ding, Dong Nyoung Heo, Sang Jin Lee, James A. Martin, Il Keun Kwon PII: DOI: Reference:
S0141-8130(17)32098-6 http://dx.doi.org/10.1016/j.ijbiomac.2017.08.125 BIOMAC 8118
To appear in:
International Journal of Biological Macromolecules
Received date: Revised date: Accepted date:
11-6-2017 15-8-2017 22-8-2017
Please cite this article as: Kee W.Jang, Dongrim Seol, Lei Ding, Dong Nyoung Heo, Sang Jin Lee, James A.Martin, Il Keun Kwon, Ultrasoundtriggered PLGA Microparticle Destruction and Degradation for Controlled Delivery of Local Cytotoxicity and Drug Release, International Journal of Biological Macromoleculeshttp://dx.doi.org/10.1016/j.ijbiomac.2017.08.125 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.
Ultrasound-triggered PLGA Microparticle Destruction and Degradation for Controlled Delivery of Local Cytotoxicity and Drug Release
Kee W. Jang1,*, Dongrim Seol2, Lei Ding3, Dong Nyoung Heo4, Sang Jin Lee4, James A. Martin2 and Il Keun Kwon4,*
1
Radiology and Imaging Sciences, Clinical Center, National Institutes of Health, Bethesda,
MD 2
Department of Orthopedics and Rehabilitation, The University of Iowa, Iowa City, IA
3
Jiangnan University Wuxi Medical School, Wuxi, Jiangsu, China
4
Department of Dental Materials, School of Dentistry, Kyung Hee University, Seoul,
Republic of Korea Corresponding Author: Kee W. Jang, Ph.D.; Address: Radiology and Imaging Sciences, Clinical Center, National Institutes of Health, 10 Center Dr. Bethesda, MD 20892; Tel.: 301-435-3875; Fax: 301402-3216; E-mail address: [email protected] Il Keun Kwon, Ph.D.: Address: Department of Dental Materials, School of Dentistry, Kyung Hee University, 26 Kyungheedae-ro, Dongdaemun-gu, Seoul, Republic of Korea; Tel.: +82-2-961-0771; Fax: +82-2-960-1457; E-mail address: [email protected] *These corresponding authors have made equal contributions to this work.
Graphical abstract
Input Pulses
US Power
Graphical abstract
# Pulses (DC)
MP Destruction
MP Destruction
DC Modulated US
US power (DC)
Cell Death
Cell death
Shock Waves Production
MP Destruction
Highlights ● The patterns of sustained DOX release from MPs were observed approximately 6 hr following the initial period of burst release.
● Certain portions of US-resistant MPs undergo accelerated degradation by repetitive sonication, likely in proportion to the amount of US energy affecting the MPs. ● DC-modulated sonication can be used to control cytotoxicity and the release of DOX via MP destruction and degradation.
ABSTRACT In this study, we investigated the low intensity ultrasound (US)-controlled delivery of local cytotoxicity and drug release via induced destruction and degradation of microparticles
(MPs) made of poly(lactic-co-glycolic acid) (PLGA). This study was conducted in vitro with potential application towards tumor treatment in conjunction with direct injection. MPs, either loaded with or without doxorubicin (DOX) or blank, were prepared using a double-emulsion solvent-evaporation technique. First, the MPs were exposed to US with duty cycle (DC)-modulation. The destruction and degradation of MPs were evaluated using light and scanning electron microscopy. Second, the effects of US-mediated destruction/degradation of MPs on the local cytotoxicity as well as DOX release were evaluated. US-triggered MP destruction/degradation significantly enhanced nearby cell death and DOX release. These affects occurred in proportion to the DC. Our findings indicate that controlled cytotoxicity and DOX release by US could be useful in developing the minimally invasive therapeutic applications for tumor treatment. Keywords: Ultrasound; Microparticle; PLGA; doxorubicin release
1. INTRODUCTION There have been many attempts to the use of acoustically-oscillatable lipid- or albumin- shelled microbubbles in combination with chemotherapeutic agents for tumor treatment due to the dynamic responses under the ultrasound (US) field [1-3]. When the microbubbles are exposed to low-peak US pressure (i.e., mechanical index (MI) < 0.05), the microbubbles stably oscillate resulting in stable cavitation that can lead to development of microstreaming and mild shock waves. By increasing the US peak pressures, microbubble oscillation becomes non-linear and can ultimately result in inertial cavitation leading to complete destruction of the microbubbles producing mechanical shock waves, temperature elevations, and free radicals [1-3]. The acoustic shock waves from USmicrobubbles interaction can be used therapeutically to trigger intracellular signal cascades
as well as inducing sonoporation, cell apoptosis and necrosis [3-8]. US mediated destruction of intravenously administered
microbubbles is a noninvasive
immunotherapy technique. When the microbubbles are targeted to tumors and exposed to US, they can transiently increase the tumor cell permeability for improved delivery of chemotherapeutic agents within the parenchyma [9-13]. This approach works well in tumors only if the tumors are well-formed with vasculature in which a chemotherapeutic agent-conjugated microbubble can be effectively delivered within the tumor parenchyma [14, 15]. However, if the targeted tumors are poorly vascularized, there is a potential risk of remaining chemotherapeutic agents conjugated to microbubbles in systemic circulation leading to greater toxicity in certain organs since the elimination of microbubbles is dominantly through the process of gaseous exchange in lung ventilation [16-19]. Thus, repetitive drug-microbubbles administration is unavoidable in order to achieve therapeutic outcomes [20]. The use of biodegradable polymeric micro/nano- particles has recently emerged as a means for targeted delivery of chemotherapeutic/immunotherapeutic agents into tumors. These particles have proven their therapeutic potential by exposing tumors with sustained release of encapsulated therapeutic agents over time [21-23]. Recent studies have demonstrated that combined in situ release of immuno- and chemo- therapeutic agents from the microparticles (MPs) significantly reduced tumor burden in mice models. This result suggests that constant release of the agents and immune responses can result in suppression of tumor growth [22, 24-27]. However, limitations still exist for this technique such as slow intracellular drug uptake,
limited drug-loading capacity, poor penetration into tumors, and the inability to specifically control drug release in situ by an external stimulus[28]. Thus, the use of particles in combination with physical stimuli-responsive therapy has become an option to overcome these limitations [29]. Although there have been few studies of the acoustic response of polymeric MPs, the acoustic characteristics are quite different as compared to acoustically oscillatable albumin- or lipid- shelled microbubbles with respect to destruction and degradation. At low peak US pressure, the polymeric MPs do not oscillate; however, the polymeric MPs undergo complete destruction without oscillation at high peak US pressures. Studies have suggested that the destruction of polymeric MPs starts with the creation of defects or cracks in the shell surface followed by release of encapsulated gas [9, 30]. Nonetheless, the utilization of US polymeric MP destruction as a possible delivery mechanism for drug in tissues has received little attention. Therefore, in this study we investigated the dependence of the response of MP destruction and degradation following duty cycle (DC)-modulated US in terms of cytotoxicity of nearby cells and encapsulated drug release. We hypothesized the followings: (1) the destruction and degradation of the MPs were dependent on US exposure; (2) US mediated MP destruction can cause nearby tumor cell death; and (3) the release of encapsulated doxorubicin (DOX) in MPs can be facilitated by US exposure.
2 Materials & Methods 2.1 Low intensity ultrasound system One megahertz sinusoidal waveforms, either continuous or pulsed at 1kHz pulse repetition frequency (PRF), were generated using versatile sweep function generator (BK Precision, Yorba Linda, CA, US). DC was adjusted by modulating the number of cycles
per each burst: 0 and 100% of DC were equivalent to non-US and continuous US respectively. The pulses were amplified using a power amplifier (ENI Inc., Rochester, NY, US) and the pulses were then transferred to a customized unfocused water-immersible US transducer (1MHz) (Ultrasonic S-Lab, Concord, CA, US). The output US power was determined by a radiation force balance technique and the power was regarded as spatial averaged and temporal averaged (ISATA) [31-33]. 2.2 Formulation of PLGA MPs MPs composed of poly(lactic-co-glycolic acid) (PLGA) were synthesized using a double emulsion solvent evaporation method. To create the water phase, either 75µL of PBS, for blank MPs, or 75µL of 1% poly(vinyl alcohol) (Mowiol 8-88; Sigma, Allentown, PA) containing 2mg of DOX (Sigma), for DOX encapsulated MPs (DOX/MPs), were prepared. To create the oil phase, 200mg of PLGA (Resomer® RG 503; Boehringer Ingelheim KG, Germany) was dissolved in 1,500µL of dichloromethane (DCM). A primary emulsion was prepared by adding water phase into oil phase during sonication using a Sonic Dismembrator (model FB120 equipped with an ultrasonic converter probe; Fisher Scientific, Pittsburgh, PA) operated at 40% amplitude for 30s. The secondary emulsion was achieved by adding the primary emulsion into the water phase (30mL of 1% PVA) and homogenized using an Ultra-Turrax homogenizer (IKA; Wilmington, NC) at 17,500rpm. The secondary emulsion was then stirred to allow DCM to be evaporated. The size dependent PLGA MPs were harvested by sequential centrifugation at 200 and 1,000 rpm using an Eppendorf centrifuge 5804R (Eppendorf, Westbury, NY). The MPs were washed twice with nanopure water,
dispersed in 5mL of water, and lyophilized (FreeZone 4.5, Labconco Corp., Kansa s City, MO). To characterize the centrifugation speed-dependent size distribution of MPs, a scanning electron microscopy (SEM) (Hitachi S-4800, Hitachi High Technologies, Ontario, CAN) was performed and the average diameters were determined (16.7+4.7µm at 200rpm and 7.4+3.3µm at 1,000rpm) by free-hand drawing lines across the MPs using ImageJ (National Institutes of Health, MD). 2.3 Determination of US-mediated MP destruction and degradation To evaluate US-mediated MP destruction and degradation, 1% (w/v) MPs were suspended in PBS without cells and exposed to US with 0.27 W/cm2 at 100% DC for 10s. Immediately after US exposure, 10µL of suspension was analyzed. The number of remaining MPs was counted using a hemocytometer and the surface area of the MPs were measured by light microscopy and analyzed with ImageJ. The surface area was extrapolated from the measured diameters using the equation for a spherical surface (4πr2). A morphological examination following US exposure was performed using a SEM. 2.4 Cytotoxicity of US-mediated MP destruction To evaluate the cytotoxic effect of combining US exposure (0.27 W/cm2 at 100% DC for 10s) and MPs, two murine tumor cell lines were used. Melanoma cell line, B16F10 (B16), and lymphoma cell line, EL4, were purchased (ATCC, Manassas, VA). B16 cells were cultured in 90% Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco) supplemented with 10% Fetal Bovine Serum (FBS) (Atlanta Biologicals, Lawrenceville, GA), 10mM (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) (HEPES) (Gibco), 50µg/mL Gentamycin (Cellgro, Manassas, VA), 1mM GlutaMAX (Gibco), and 1mM sodium pyruvate (Gibco) at 37oC and 5% CO2. EL4 cells were cultured in 0.1%
Mercaptoethanol in Roswell Park Memorial Institute (RPMI) 1640 medium (Gibco, Life Technologies Corporations, NY). One percent (w/v) of PLGA MPs was prepared in a suspension containing either B16 or EL4 cells (0.5 x 106 cells/mL) and exposed to DC dependent US for 10s (Fig. 1). Control cells in suspension were combined with 1% (w/v) PLGA MPs and were not sonicated. After US exposure, the cells were labeled with 1µM Calcein-AM (Thermo Scientific, Waltham, MA), a live cell indicator, and 1µM ethidium homodimer-2, a dead cell indicator (Thermo Scientific). The live and dead cells were counted using a hemocytometer and a fluorescence microscope (BX60, Olympus, Center Valley, PA, US). A percentage of cell viability was calculated as [(live cells)/(live + dead cells)] x 100 [%]. 2.5 Drug release in MPs following sonication Two to three milligrams of DOX-encapsulated MPs (DOX/MPs) were suspended in 1mL of PBS in Eppendorf tubes. The DOX/MPs were exposed to US with 0.27 W/cm2 in intensity at duty cycles of 10, 50 or 100% for 10s. After US exposure, the tubes were placed in a shaker incubator set at 300rpm and 37oC. At each predetermined time point, 800µL of supernatant was collected after centrifugation at 1,000rpm for 5min for analysis. Eight hundred microliter of fresh PBS was replenished into the tubes to maintain initial volume and avoid concentrating the remaining DOX in solution. The amount of DOX in the harvested supernatants was determined by fluorescence at excitation and emission wavelength of ex = 470 and em = 585 nm, respectively using a SpectraMax M5 multidetection microplate reader (Molecular Devices Inc., Sunnyvale, CA). The percentage of DOX release was evaluated by normalizing the data to the initial time point measurement. 2.6 Statistical analysis
Data analysis was performed using Prism6 (GraphPad, La Jolla, CA). Groups were compared by one-way analysis of variance (ANOVA) with the Tukey post hoc test. Repeated-measures one-way ANOVA with Dunnett’s multiple comparison tests was performed to compare the effect of US exposure on DOX release. Data were compared to the time point immediately before US exposure. The level of significant difference was considered at p<0.05. Asterisk and pound symbols represent statistical significance (* and # p<0.05, ** and ## p<0.01, *** and ### p<0.001 and **** and #### p<0.0001).
3. Results 3.1 Determination of US intensity The output US power was determined by a radiation force balance technique. The US intensity at 100% DC was determined as 0.27 W/cm 2 and assumed as spatially averaged and temporally averaged. 3.2 Microscopic evaluation of MP destruction and degradation To evaluate the destruction and degradation of MPs (mean diameter = 7µm) following sonication, morphologic analysis of US-exposed MPs was performed. After US exposure (0.27W/cm 2 , 100% DC for 10s), the percentage of remaining MPs was significantly decrease (p<0.001) (Fig. 2A and B). There was no considerable difference among tested US intensities (0.27, 1.5 and 2.5 W/cm 2 (Fig. 2C). To examine the effect of repetitive sonication (0.27W/cm 2 , 100% DC for 10s) on MP degradation, the surface area of MPs was measured and analyzed. A greater degree of MP degradation was observed with increasing the repetition of US exposure. Compared to the control (x 0 sonication), there was a 34% reduction in surface
area of MPs with 1 sonication (p<0.01). It was further reduced by 63% following 4 sonications (p<0.0001) (Fig. 2D). 3.3 Morphology of sonicated-MPs by SEM SEM analysis showed morphologically intact and smooth surface in control MPs (7µm) (Fig. 3A and C). However, numerous cracks and holes appeared on the MP surface after US exposure (0.27W/cm 2 , 100% DC for 10s) (Fig. 3B and D). 3.4 Local cytotoxicity of US-mediated MP destruction To test the local cytotoxic effect of US-mediated MP destruction, two types of cell lines (B16 and EL4) in suspension containing MPs (1% w/v)) were sonicated. In the presence of MPs (7µm), cell viability remained above 95% when unsonicated. However, the cell viability was significantly decreased into around 50% (p<0.001) when exposed to US at 50% DC. It further dropped to approximately 5% at 100% DC (p<0.001). There were no significant differences of cell viability between B16 and EL4 following tested DCmodulated sonication (Fig. 4A). Since the cytotoxicity of US mediated MP destruction was not dependent on the types of cell lines, we arbitrarily chose B16 for performing the rest of experiments. To examine the cytotoxic effect of DC-modulated US exposure and in combination with MPs, B16 cells with or without 1% MPs (w/v) (7µm) in suspension were exposed to US at 0, 50 and 100% DC. In the absence of MPs, US exposure alone did not alter cell viability. Cell viabilities were all remained above 95% following US exposure regardless of DC. However, DC had a clear effect on cell viability when suspended with MPs (7µm). The cell viability was around 50% with 50% DC (p<0.0001) and 5% with 100% DC (p<0.0001) (Fig. 4B). To examine the mean size- or concentration- dependent cytotoxicity, either 7 or
17 µm of MPs were suspended with B16 cells at different levels of concentration followed by US exposure (0.27W/cm 2 , 100% DC for 10s). At concentration of 0.5% and 2.5% MPs (w/v) (7µm) in the cell suspension, we observed 50% (p<0.0001) and 10% (p<0.0001) of the remaining cells to be viable following US exposure. However, we observed no considerable cell death when the cells were suspended with 17µm MPs. Cell viability remained unchanged at above 95% regardless of tested concentrations (Fig. 4C). 3.5 US-triggered DOX release The pattern of sustained DOX release in MPs was observed approximately 6hr after the initial burst release. US exposure (0.27W/cm 2 , 100% DC for 10s) with DC modulation at 10, 50 and 100% was applied immediately at the beginning of experiments. During the period of burst release (<6hr), no clear difference of the release rate was observed. However, the difference became apparent when it reached the period of sustained release (>6hr) as compared to control. DOX release was significantly increased following sonication as compared to control at each subsequent time point (Fig. 5A). The DOX release was increased by 17% with 10 and 50% DC and 33% with 100% DC, as compared to control. The accelerated release kinetics remained higher until the pre-determined end time point of the experiments (Fig. 5A). To confirm that the US-accelerated of DOX release can be achieved during the period of sustained release (>6hr), sonication (0.27 W/cm2 at 100% DC for 10s) was applied at a time point of 48hr. The results showed that another burst release appeared immediately following US exposure and the accelerated release became significant at post 54hr (p<0.05) and at post 96hr (p<0.05), as compared to the time point immediately before sonication at 48hr (Fig.
5C and D). The DOX release rate remained unchanged in unsonicated control (Fig. 5B and C).
4. Discussion Linearly oscillating lipid- or albumin- shelled microbubbles have been extensively studied as an ultrasound contrast agent as well as a targeted drug delivery vehicle due to the fact that their acoustic properties are exceptionally dynamic depending on US pressure [1, 4, 5, 34]. However, US mediated polymeric MP destruction has received little attention even though there is evidence that polymeric MPs can be destroyed by US. Studies have suggested that the destruction of the polymeric MPs starts with creating defects and cracks in the shell surface by releasing encapsulated gas or air without linear oscillation [9, 30]. Nonetheless, there have been little studies about utilizing polymeric MP destruction in response to US exposure as a therapeutic strategy. Therefore, in this study we investigated the effects of US exposure on the destruction and degradation of MPs made of PLGA for potential tumor treatment. We chose PLGA as a source of MPs because it is regarded as one of the most promising biomaterials that can be naturally hydrolyzed and metabolized in the body with minimal toxicity. Additionally, it has well established patterns of controlled and sustained release of encapsulated drugs [22, 23]. We first examined the acoustic responses of MPs (mean diameter = 7µm) in terms of destruction and degradation. Light microscopy confirmed that approximately 40% of MPs were destroyed immediately following sonication (0.27 W/cm2 at 100% DC for 10s) as compared to control. We next examined the effects of higher intensity sonication on MP destruction by applying three different levels of US intensities, 0.27, 1.5 and 2.5 W/cm2 (at
100% DC for 10s). As compared to the un-sonicated control group, approximately 40% of MPs were destroyed by US exposure at 0.27W/cm2. However, no difference in MP destruction was observed amongst the tested US intensities. Although the difference of acoustic characteristics between destroyed and un-destroyed MPs remains unknown yet, it is probable that a certain population of MPs were US-sensitive due to their structure. These could be imploded immediately and the rest of the MP population was probably more resistant to the tested US. Since the MP destruction was independent to the tested US intensities, we arbitrarily chose the US intensity at 0.27 W/cm2 for the rest of study. Following US exposure (0.27 W/cm2 at 100% DC for 10s) on MPs, we quantified the surface area of MPs that could represent the degradation of MPs. We observed that the surface area of MPs was gradually reduced by increasing the number of sonications. Approximately 40% of surface area of MPs was reduced after the first sonication and 60% reduction was observed after 4 repeated sonications. These results are consistent our assumption that a certain portion of US-resistant MPs undergo accelerated degradation by repetitive sonication. This is likely in proportion with the amount of US waves affecting the MPs. Since we confirmed that MPs can be destroyed by US exposure, we examined the cytotoxic effect of imploding MP on nearby cells. Cytotoxicity is observed due to the fact that imploding microbubbles are known to generate shock waves which can mechanically rupture nearby cells [3, 6, 8]. First, we examined the effect of MP destruction on cytotoxicity in two different cell lines, B16 and EL4. We observed no different cell viability between tested cell lines suggesting that the cytotoxicity of MP destruction was primarily from instant mechanical shock waves [6, 8, 35]. Thus, we chose B16 for the rest
of experiments. The dependency of DC (50 and 100 %) and MP concentration (0.5 and 2.5 % (w/v)) on cell death was examined. We observed that the greater cell death was observed with higher DC and MP concentration. This may be due to several factors. These include that the greater DC produced the greater averaged US power affecting MPs resulting in a greater number of MP destruction. Additionally, the presence of more US-susceptible MPs in suspension, due to higher MP concentration, increases the number that are destroyed US exposure. We were intrigued by examining the different sizes of MPs (mean diameter = 17µm). We expected that the cytotoxicity of MP destruction would be similar for 7µm MPs. However, we observed little cell death with 17µm MPs. Although the reason for the inactivity of 17µm MPs responding to US exposure remained unclear, we speculate that the innate characteristics of 7µm MPs are more susceptible to US exposure than 17µm MPs. It is likely that the surface area of MPs directly interacting with US waves could be the primary sensitizing factor in terms of destruction or degradation because the percentage of US propagation could be widely affected depending on the available surface area of MPs. Further investigation is required to understand. It is also possible that the size dependent US susceptibility might be also dependent on other US parameters such as frequency, intensity and duration. Although, further investigation would be required to understand the US-parameter dependent MP destruction, we agreed that the 7µm MPs can have complete destruction by US exposure with the mechanism as previously suggested [9, 10, 36, 37]. Since certain populations of MPs were susceptible to destruction and degradation by US exposure, it was reasonable to hypothesize that the release of encapsulated drugs in
MPs can be controlled by sonication. To test this hypothesis, DOX was loaded in MPs and the release kinetics was evaluated. DOX was chosen because the release of DOX following cytotoxicity and poration due to MP destruction would have synergistic tumor-suppressive effects in our future study. In our first experiment, US exposures (0.27 W/cm2 for 10s) at 10, 50 and 100% DC were applied at the beginning of the experiments. All groups, including un-sonicated control, showed an initial burst release (<6hr) followed by a sustained release phase (after 6hr) of DOX from the MPs. The effect of US exposure on DOX release became apparent during the sustained release phase by increasing approximately 17% at 10 and 50 % DC and 33% at 100% DC as compared to un-sonicated control. This suggests that the destroyed and degraded MPs generated by sonication facilitated a faster release of DOX as compared to intact MPs. In order to confirm that USinduced DOX release can be achieved anytime irrespective to the release states (in either burst or sustained release state), we sonicated DOX-MPs once during the period of sustained release (at 48hr). Immediately following sonication, a new burst release occurred and the enhancement of DOX release became significant within 6hr following sonication. This suggests that breaking the MPs during sustained release further releases DOX. Further investigation is needed to confirm that accelerated MP degradation, by repetitive sonications, can further increase the DOX release. This may or may not occur because the DOX could be still possibly bound to broken pieces of MPs. There have been extensive attempts to use intravenously infused gas-filled microbubble destruction by US for targeted drug delivery into tumors. The effect, so fa, have proven to be promising [11-13]. However, to be successful, it requires that the tumors possess a well-formed vasculature. This is not typically the cases [14, 38, 39]. Also, the
potential risks of infusing microbubble-conjugated anti-cancer drugs through intravenous administration, remain unknown as it may lead to unnecessarily high concentrations of toxic drugs in systemic circulation [16, 17]. In this study, we demonstrated that DOX loaded MPs, made of PLGA, can be destroyed by US application producing cytotoxicity and accelerated DOX release from MPs. Considering that DOX-PLGA is directly infused into tumors, this could potentially have long-term benefits for the tumor treatment by controlled delivery of instant cytotoxicity from MP destruction as well as sustained DOX release with minimal invasiveness in combination with US exposure. This technique also provides for reduced potential risks of toxic chemotherapeutic drugs in systemic circulation.
5. Conclusion In summary, DC-modulated sonication can be used to control cytotoxicity and the release of DOX via MP destruction and degradation. Our future study focuses on the synergistic effects of a combination of US exposure and a single intra-tumoral injection of DOX-loaded PLGA-MPs. This synergistic effect of tumor suppression can occur through an instant cytotoxicity followed by sustained release of DOX from a minimally invasive manner. Our findings suggest that DOX-loaded PLGA MPs can be used as an effective tool for use as a therapeutic agent against tumor and cancer cells.
ACKNOWLEDGEMENTS This work was supported by the Bio & Medical Technology Development Program of the National Research Foundation (NRF) & funded by the Korean government (MSIP&MOHW) (No. 2017M3A9E4048170).
REFERENCES [1] T. Kodama, Y. Tomita, K. Koshiyama, M.J. Blomley, Transfection effect of microbubbles on cells in superposed ultrasound waves and behavior of cavitation bubble, Ultrasound Med Biol 32(6) (2006) 905-14. [2] P.A. Dijkmans, L.J. Juffermans, R.J. Musters, A. van Wamel, F.J. ten Cate, W. van Gilst, C.A. Visser, N. de Jong, O. Kamp, Microbubbles and ultrasound: from diagnosis to therapy, Eur J Echocardiogr 5(4) (2004) 245-56. [3] L.B. Feril, Jr., T. Kondo, Q.L. Zhao, R. Ogawa, K. Tachibana, N. Kudo, S. Fujimoto, S. Nakamura, Enhancement of ultrasound-induced apoptosis and cell lysis by echocontrast agents, Ultrasound Med Biol 29(2) (2003) 331-7. [4] P. Qin, L. Xu, W. Zhong, A.C. Yu, Ultrasound-microbubble mediated cavitation of plant cells: effects on morphology and viability, Ultrasound Med Biol 38(6) (2012) 1085-96. [5] M.M. Forbes, R.L. Steinberg, W.D. O'Brien, Jr., Examination of inertial cavitation of Optison in producing sonoporation of chinese hamster ovary cells, Ultrasound Med Biol 34(12) (2008) 2009-18. [6] Y. Feng, Z. Tian, M. Wan, Bioeffects of low-intensity ultrasound in vitro: apoptosis, protein profile alteration, and potential molecular mechanism, J Ultrasound Med 29(6) (2010) 963-74. [7] L.B. Feril, Jr., T. Kondo, Biological effects of low intensity ultrasound: the mechanism involved, and its implications on therapy and on biosafety of ultrasound, J Radiat Res 45(4) (2004) 479-89.
[8] M.A. Hassan, L.B. Feril, Jr., K. Suzuki, N. Kudo, K. Tachibana, T. Kondo, Evaluation and comparison of three novel microbubbles: enhancement of ultrasound-induced cell death and free radicals production, Ultrason Sonochem 16(3) (2009) 372-8. [9] S. Hernot, A.L. Klibanov, Microbubbles in ultrasound-triggered drug and gene delivery, Adv Drug Deliv Rev 60(10) (2008) 1153-66. [10] W.G. Pitt, G.A. Husseini, B.J. Staples, Ultrasonic drug delivery--a general review, Expert opinion on drug delivery 1(1) (2004) 37-56. [11] J. Du, L.F. Du, F.H. Li, X.Z. Zheng, H.L. Li, Q.S. Shi, Y. Wu, Ultrasound targeted microbubble destruction-mediated gene delivery system: application to therapy for ocular disease, Asian Biomed 5(5) (2011) 577-587. [12] C. Pu, S. Chang, J. Sun, S. Zhu, H. Liu, Y. Zhu, Z. Wang, R.X. Xu, Ultrasoundmediated destruction of LHRHa-targeted and paclitaxel-loaded lipid microbubbles for the treatment of intraperitoneal ovarian cancer xenografts, Mol Pharm 11(1) (2014) 4958. [13] H. Liu, S. Chang, J. Sun, S. Zhu, C. Pu, Y. Zhu, Z. Wang, R.X. Xu, Ultrasoundmediated destruction of LHRHa-targeted and paclitaxel-loaded lipid microbubbles induces proliferation inhibition and apoptosis in ovarian cancer cells, Mol Pharm 11(1) (2014) 40-8. [14] R.M. Terek, Angiogenesis in chondrosarcoma, Current Opinion in Orthopaedics 13(6) (2002) 449-453. [15] J.A. Buckwalter, The Structure of Human Chondrosarcoma Proteoglycans, J Bone Joint Surg Am 65(7) (1983) 958-974.
[16] S. Podell, C. Burrascano, M. Gaal, B. Golec, J. Maniquis, P. Mehlhaff, Physical and biochemical stability of Optison, an injectable ultrasound contrast agent, Biotechnol Appl Biochem 30 ( Pt 3) (1999) 213-23. [17] J.C. Hutter, H.M. Luu, P.M. Mehlhaff, A.L. Killam, H.C. Dittrich, Physiologically based pharmacokinetic model for fluorocarbon elimination after the administration of an octafluoropropane-albumin microsphere sonographic contrast agent, J Ultrasound Med 18(1) (1999) 1-11. [18] H. Chen, J.H. Hwang, Ultrasound-targeted microbubble destruction for chemotherapeutic drug delivery to solid tumors, J Ther Ultrasound 1 (2013) 10. [19] F. Zhu, Y. Jiang, F. Luo, P. Li, Effectiveness of localized ultrasound-targeted microbubble destruction with doxorubicin liposomes in H22 mouse hepatocellular carcinoma model, J Drug Target 23(4) (2015) 323-34. [20] J. Kang, X. Wu, Z. Wang, H. Ran, C. Xu, J. Wu, Z. Wang, Y. Zhang, Antitumor effect of
docetaxel-loaded
lipid
microbubbles
combined
with
ultrasound-targeted
microbubble activation on VX2 rabbit liver tumors, J Ultrasound Med 29(1) (2010) 6170. [21] J. Khandare, T. Minko, Polymer-drug conjugates: Progress in polymeric prodrugs, Prog Polym Sci 31(4) (2006) 359-397. [22] H.K. Makadia, S.J. Siegel, Poly Lactic-co-Glycolic Acid (PLGA) as Biodegradable Controlled Drug Delivery Carrier, Polymers-Basel 3(3) (2011) 1377-1397. [23] X. Zhu, R.D. Braatz, A mechanistic model for drug release in PLGA biodegradable stent coatings coupled with polymer degradation and erosion, J Biomed Mater Res A 103(7) (2015) 2269-79.
[24] K. Shahani, S.K. Swaminathan, D. Freeman, A. Blum, L.A. Ma, J. Panyam, Injectable Sustained Release Microparticles of Curcumin: A New Concept for Cancer Chemoprevention, Cancer Res 70(11) (2010) 4443-4452. [25] J.M. Lu, X. Wang, C. Marin-Muller, H. Wang, P.H. Lin, Q. Yao, C. Chen, Current advances in research and clinical applications of PLGA-based nanotechnology, Expert Rev Mol Diagn 9(4) (2009) 325-41. [26] M. Figueiredo, R. Esenaliev, PLGA Nanoparticles for Ultrasound-Mediated Gene Delivery to Solid Tumors, J Drug Deliv 2012 (2012) 767839. [27] F. Danhier, E. Ansorena, J.M. Silva, R. Coco, A. Le Breton, V. Preat, PLGA-based nanoparticles: an overview of biomedical applications, J Control Release 161(2) (2012) 505-22. [28] C. Deng, Y.J. Jiang, R. Cheng, F.H. Meng, Z.Y. Zhong, Biodegradable polymeric micelles for targeted and controlled anticancer drug delivery: Promises, progress and prospects, Nano Today 7(5) (2012) 467-480. [29] N. Rapoport, Physical stimuli-responsive polymeric micelles for anti-cancer drug delivery, Prog Polym Sci 32(8-9) (2007) 962-990. [30] S.H. Bloch, M. Wan, P.A. Dayton, K.W. Ferrara, Optical observation of lipid- and polymer-shelled ultrasound microbubble contrast agents, Appl Phys Lett 84(4) (2004) 631-633. [31] B. Zeqiri, C.J. Bickley, A new anechoic material for medical ultrasonic applications, Ultrasound Med Biol 26(3) (2000) 481-5. [32] R.C. Preston, Measurement and Characterization of the Acoustic Output of Medical Ultrasonic Equipment .2., Med Biol Eng Comput 24(3) (1986) 225-234.
[33] R.C. Preston, Measurement and Characterization of the Acoustic Output of Medical Ultrasonic Equipment .1., Med Biol Eng Comput 24(2) (1986) 113-120. [34] C.R. Mayer, N.A. Geis, H.A. Katus, R. Bekeredjian, Ultrasound targeted microbubble destruction for drug and gene delivery, Expert Opin Drug Deliv 5(10) (2008) 1121-38. [35] L.B. Feril, T. Kondo, Q.L. Zhao, R. Ogawa, K. Tachibana, N. Kudo, S. Fujimoto, S. Nakamura, Enhancement of ultrasound-induced apoptosis and cell lysis by echocontrast agents, Ultrasound in Medicine and Biology 29(2) (2003) 331-337. [36] P. Hauff, S. Seemann, R. Reszka, M. Schultze-Mosgau, M. Reinhardt, T. Buzasi, T. Plath, S. Rosewicz, M. Schirner, Evaluation of gas-filled microparticles and sonoporation as gene delivery system: Feasibility study in rodent tumor models, Radiology 236(2) (2005) 572-578. [37] D.J. May, J.S. Allen, K.W. Ferrara, Dynamics and fragmentation of thick-shelled microbubbles, Ieee T Ultrason Ferr 49(10) (2002) 1400-1410. [38] R.L. McGough, C.Z. Lin, P. Meitner, B.I. Aswad, R.M. Terek, Angiogenic cytokines in cartilage tumors, Clin Orthop Relat R (397) (2002) 62-69. [39] M. Ryzewicz, B.J. Manaster, E. Naar, B. Lindeque, Low-grade cartilage tumors: Diagnosis and treatment, Orthopedics 30(1) (2007) 35-46.
Fig. 1. Schematic illustration of experimental setup. The US transducer was immersed in degassed water and a cell suspension, either with or without PLGA MPs, was exposed to US from a distance of approximately 1 cm. US mediated MP destruction killed nearby cells.
Fig. 2. MP destruction and degradation by US exposure. (A) Light microscopic image showing 1% (w/v) MP suspension before US exposure. (B) Light microscopic image showing 1% (w/v) MP suspension after US exposure. (C) The number of remaining MPs was significantly reduced after US exposure and no considerable difference among tested intensities was observed. (D) The surface area of MPs was gradually reduced with more repetitive US exposure. *Statistically significant at p < 0.05 and NS represents no significance. Error bars represent standard deviation (n=3). Scale bar = 500µm.
Fig. 3. Morphological analysis of MPs by SEM. (A) Un-sonicated group showing intact and smooth surfaced MPs. (B) Appearance of cracks and holes at the surface of MPs was observed following US exposure. (C) Higher magnification of non-US exposed MPs. (D) Higher magnification of US exposed MPs. White arrows indicate holes in MPs that were created following sonication. Scale bar = 10µm (A and B) and 5 µm (C and D).
Fig. 4. The dependency of cell-type, DC, MP diameter, and MP concentration on local cell death following sonication. (A) The local cell death was increased by increasing DC. No considerable difference between tested cell lines (B16 and EL4) was observed. (B) Following sonication, little cytotoxicity was observed in the absence of MPs whereas significant cell death was observed by increasing DC in the presence of MPs. (C) Cell death was also proportional to the concentration of MPs (7µm); however, no cell death was observed with MPs (17µm) following sonication (n=3-5). Asterisk (*) and pound (#) represent statistical significances compared to control (
*** and ###
p<0.001 and
represent standard deviation (n=3-5).
**** and ####
p<0.0001) and NS represents no significance. Error bars
Fig. 5. Accelerated DOX release by US exposure. (A) Accelerated DOX release was dependent on DCmodulated US. (C) Sonication at a state of sustained release shows a new burst release. (B) Detailed release profile of control group be tween 45 to 55 hrs. (D) Detailed release profile of US exposed group between 45 to 55 hrs. Asterisk (*) represents statistical significances as compared to *
**
control ( p<0.05, p<0.01,
***
p<0.001 and
****
p<0.0001). Error bars represent standard deviation (n=3-6).
S0141-8130(17)32098-6 http://dx.doi.org/10.1016/j.ijbiomac.2017.08.125 BIOMAC 8118
To appear in:
International Journal of Biological Macromolecules
Received date: Revised date: Accepted date:
11-6-2017 15-8-2017 22-8-2017
Please cite this article as: Kee W.Jang, Dongrim Seol, Lei Ding, Dong Nyoung Heo, Sang Jin Lee, James A.Martin, Il Keun Kwon, Ultrasoundtriggered PLGA Microparticle Destruction and Degradation for Controlled Delivery of Local Cytotoxicity and Drug Release, International Journal of Biological Macromoleculeshttp://dx.doi.org/10.1016/j.ijbiomac.2017.08.125 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.
Ultrasound-triggered PLGA Microparticle Destruction and Degradation for Controlled Delivery of Local Cytotoxicity and Drug Release
Kee W. Jang1,*, Dongrim Seol2, Lei Ding3, Dong Nyoung Heo4, Sang Jin Lee4, James A. Martin2 and Il Keun Kwon4,*
1
Radiology and Imaging Sciences, Clinical Center, National Institutes of Health, Bethesda,
MD 2
Department of Orthopedics and Rehabilitation, The University of Iowa, Iowa City, IA
3
Jiangnan University Wuxi Medical School, Wuxi, Jiangsu, China
4
Department of Dental Materials, School of Dentistry, Kyung Hee University, Seoul,
Republic of Korea Corresponding Author: Kee W. Jang, Ph.D.; Address: Radiology and Imaging Sciences, Clinical Center, National Institutes of Health, 10 Center Dr. Bethesda, MD 20892; Tel.: 301-435-3875; Fax: 301402-3216; E-mail address: [email protected] Il Keun Kwon, Ph.D.: Address: Department of Dental Materials, School of Dentistry, Kyung Hee University, 26 Kyungheedae-ro, Dongdaemun-gu, Seoul, Republic of Korea; Tel.: +82-2-961-0771; Fax: +82-2-960-1457; E-mail address: [email protected] *These corresponding authors have made equal contributions to this work.
Graphical abstract
Input Pulses
US Power
Graphical abstract
# Pulses (DC)
MP Destruction
MP Destruction
DC Modulated US
US power (DC)
Cell Death
Cell death
Shock Waves Production
MP Destruction
Highlights ● The patterns of sustained DOX release from MPs were observed approximately 6 hr following the initial period of burst release.
● Certain portions of US-resistant MPs undergo accelerated degradation by repetitive sonication, likely in proportion to the amount of US energy affecting the MPs. ● DC-modulated sonication can be used to control cytotoxicity and the release of DOX via MP destruction and degradation.
ABSTRACT In this study, we investigated the low intensity ultrasound (US)-controlled delivery of local cytotoxicity and drug release via induced destruction and degradation of microparticles
(MPs) made of poly(lactic-co-glycolic acid) (PLGA). This study was conducted in vitro with potential application towards tumor treatment in conjunction with direct injection. MPs, either loaded with or without doxorubicin (DOX) or blank, were prepared using a double-emulsion solvent-evaporation technique. First, the MPs were exposed to US with duty cycle (DC)-modulation. The destruction and degradation of MPs were evaluated using light and scanning electron microscopy. Second, the effects of US-mediated destruction/degradation of MPs on the local cytotoxicity as well as DOX release were evaluated. US-triggered MP destruction/degradation significantly enhanced nearby cell death and DOX release. These affects occurred in proportion to the DC. Our findings indicate that controlled cytotoxicity and DOX release by US could be useful in developing the minimally invasive therapeutic applications for tumor treatment. Keywords: Ultrasound; Microparticle; PLGA; doxorubicin release
1. INTRODUCTION There have been many attempts to the use of acoustically-oscillatable lipid- or albumin- shelled microbubbles in combination with chemotherapeutic agents for tumor treatment due to the dynamic responses under the ultrasound (US) field [1-3]. When the microbubbles are exposed to low-peak US pressure (i.e., mechanical index (MI) < 0.05), the microbubbles stably oscillate resulting in stable cavitation that can lead to development of microstreaming and mild shock waves. By increasing the US peak pressures, microbubble oscillation becomes non-linear and can ultimately result in inertial cavitation leading to complete destruction of the microbubbles producing mechanical shock waves, temperature elevations, and free radicals [1-3]. The acoustic shock waves from USmicrobubbles interaction can be used therapeutically to trigger intracellular signal cascades
as well as inducing sonoporation, cell apoptosis and necrosis [3-8]. US mediated destruction of intravenously administered
microbubbles is a noninvasive
immunotherapy technique. When the microbubbles are targeted to tumors and exposed to US, they can transiently increase the tumor cell permeability for improved delivery of chemotherapeutic agents within the parenchyma [9-13]. This approach works well in tumors only if the tumors are well-formed with vasculature in which a chemotherapeutic agent-conjugated microbubble can be effectively delivered within the tumor parenchyma [14, 15]. However, if the targeted tumors are poorly vascularized, there is a potential risk of remaining chemotherapeutic agents conjugated to microbubbles in systemic circulation leading to greater toxicity in certain organs since the elimination of microbubbles is dominantly through the process of gaseous exchange in lung ventilation [16-19]. Thus, repetitive drug-microbubbles administration is unavoidable in order to achieve therapeutic outcomes [20]. The use of biodegradable polymeric micro/nano- particles has recently emerged as a means for targeted delivery of chemotherapeutic/immunotherapeutic agents into tumors. These particles have proven their therapeutic potential by exposing tumors with sustained release of encapsulated therapeutic agents over time [21-23]. Recent studies have demonstrated that combined in situ release of immuno- and chemo- therapeutic agents from the microparticles (MPs) significantly reduced tumor burden in mice models. This result suggests that constant release of the agents and immune responses can result in suppression of tumor growth [22, 24-27]. However, limitations still exist for this technique such as slow intracellular drug uptake,
limited drug-loading capacity, poor penetration into tumors, and the inability to specifically control drug release in situ by an external stimulus[28]. Thus, the use of particles in combination with physical stimuli-responsive therapy has become an option to overcome these limitations [29]. Although there have been few studies of the acoustic response of polymeric MPs, the acoustic characteristics are quite different as compared to acoustically oscillatable albumin- or lipid- shelled microbubbles with respect to destruction and degradation. At low peak US pressure, the polymeric MPs do not oscillate; however, the polymeric MPs undergo complete destruction without oscillation at high peak US pressures. Studies have suggested that the destruction of polymeric MPs starts with the creation of defects or cracks in the shell surface followed by release of encapsulated gas [9, 30]. Nonetheless, the utilization of US polymeric MP destruction as a possible delivery mechanism for drug in tissues has received little attention. Therefore, in this study we investigated the dependence of the response of MP destruction and degradation following duty cycle (DC)-modulated US in terms of cytotoxicity of nearby cells and encapsulated drug release. We hypothesized the followings: (1) the destruction and degradation of the MPs were dependent on US exposure; (2) US mediated MP destruction can cause nearby tumor cell death; and (3) the release of encapsulated doxorubicin (DOX) in MPs can be facilitated by US exposure.
2 Materials & Methods 2.1 Low intensity ultrasound system One megahertz sinusoidal waveforms, either continuous or pulsed at 1kHz pulse repetition frequency (PRF), were generated using versatile sweep function generator (BK Precision, Yorba Linda, CA, US). DC was adjusted by modulating the number of cycles
per each burst: 0 and 100% of DC were equivalent to non-US and continuous US respectively. The pulses were amplified using a power amplifier (ENI Inc., Rochester, NY, US) and the pulses were then transferred to a customized unfocused water-immersible US transducer (1MHz) (Ultrasonic S-Lab, Concord, CA, US). The output US power was determined by a radiation force balance technique and the power was regarded as spatial averaged and temporal averaged (ISATA) [31-33]. 2.2 Formulation of PLGA MPs MPs composed of poly(lactic-co-glycolic acid) (PLGA) were synthesized using a double emulsion solvent evaporation method. To create the water phase, either 75µL of PBS, for blank MPs, or 75µL of 1% poly(vinyl alcohol) (Mowiol 8-88; Sigma, Allentown, PA) containing 2mg of DOX (Sigma), for DOX encapsulated MPs (DOX/MPs), were prepared. To create the oil phase, 200mg of PLGA (Resomer® RG 503; Boehringer Ingelheim KG, Germany) was dissolved in 1,500µL of dichloromethane (DCM). A primary emulsion was prepared by adding water phase into oil phase during sonication using a Sonic Dismembrator (model FB120 equipped with an ultrasonic converter probe; Fisher Scientific, Pittsburgh, PA) operated at 40% amplitude for 30s. The secondary emulsion was achieved by adding the primary emulsion into the water phase (30mL of 1% PVA) and homogenized using an Ultra-Turrax homogenizer (IKA; Wilmington, NC) at 17,500rpm. The secondary emulsion was then stirred to allow DCM to be evaporated. The size dependent PLGA MPs were harvested by sequential centrifugation at 200 and 1,000 rpm using an Eppendorf centrifuge 5804R (Eppendorf, Westbury, NY). The MPs were washed twice with nanopure water,
dispersed in 5mL of water, and lyophilized (FreeZone 4.5, Labconco Corp., Kansa s City, MO). To characterize the centrifugation speed-dependent size distribution of MPs, a scanning electron microscopy (SEM) (Hitachi S-4800, Hitachi High Technologies, Ontario, CAN) was performed and the average diameters were determined (16.7+4.7µm at 200rpm and 7.4+3.3µm at 1,000rpm) by free-hand drawing lines across the MPs using ImageJ (National Institutes of Health, MD). 2.3 Determination of US-mediated MP destruction and degradation To evaluate US-mediated MP destruction and degradation, 1% (w/v) MPs were suspended in PBS without cells and exposed to US with 0.27 W/cm2 at 100% DC for 10s. Immediately after US exposure, 10µL of suspension was analyzed. The number of remaining MPs was counted using a hemocytometer and the surface area of the MPs were measured by light microscopy and analyzed with ImageJ. The surface area was extrapolated from the measured diameters using the equation for a spherical surface (4πr2). A morphological examination following US exposure was performed using a SEM. 2.4 Cytotoxicity of US-mediated MP destruction To evaluate the cytotoxic effect of combining US exposure (0.27 W/cm2 at 100% DC for 10s) and MPs, two murine tumor cell lines were used. Melanoma cell line, B16F10 (B16), and lymphoma cell line, EL4, were purchased (ATCC, Manassas, VA). B16 cells were cultured in 90% Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco) supplemented with 10% Fetal Bovine Serum (FBS) (Atlanta Biologicals, Lawrenceville, GA), 10mM (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) (HEPES) (Gibco), 50µg/mL Gentamycin (Cellgro, Manassas, VA), 1mM GlutaMAX (Gibco), and 1mM sodium pyruvate (Gibco) at 37oC and 5% CO2. EL4 cells were cultured in 0.1%
Mercaptoethanol in Roswell Park Memorial Institute (RPMI) 1640 medium (Gibco, Life Technologies Corporations, NY). One percent (w/v) of PLGA MPs was prepared in a suspension containing either B16 or EL4 cells (0.5 x 106 cells/mL) and exposed to DC dependent US for 10s (Fig. 1). Control cells in suspension were combined with 1% (w/v) PLGA MPs and were not sonicated. After US exposure, the cells were labeled with 1µM Calcein-AM (Thermo Scientific, Waltham, MA), a live cell indicator, and 1µM ethidium homodimer-2, a dead cell indicator (Thermo Scientific). The live and dead cells were counted using a hemocytometer and a fluorescence microscope (BX60, Olympus, Center Valley, PA, US). A percentage of cell viability was calculated as [(live cells)/(live + dead cells)] x 100 [%]. 2.5 Drug release in MPs following sonication Two to three milligrams of DOX-encapsulated MPs (DOX/MPs) were suspended in 1mL of PBS in Eppendorf tubes. The DOX/MPs were exposed to US with 0.27 W/cm2 in intensity at duty cycles of 10, 50 or 100% for 10s. After US exposure, the tubes were placed in a shaker incubator set at 300rpm and 37oC. At each predetermined time point, 800µL of supernatant was collected after centrifugation at 1,000rpm for 5min for analysis. Eight hundred microliter of fresh PBS was replenished into the tubes to maintain initial volume and avoid concentrating the remaining DOX in solution. The amount of DOX in the harvested supernatants was determined by fluorescence at excitation and emission wavelength of ex = 470 and em = 585 nm, respectively using a SpectraMax M5 multidetection microplate reader (Molecular Devices Inc., Sunnyvale, CA). The percentage of DOX release was evaluated by normalizing the data to the initial time point measurement. 2.6 Statistical analysis
Data analysis was performed using Prism6 (GraphPad, La Jolla, CA). Groups were compared by one-way analysis of variance (ANOVA) with the Tukey post hoc test. Repeated-measures one-way ANOVA with Dunnett’s multiple comparison tests was performed to compare the effect of US exposure on DOX release. Data were compared to the time point immediately before US exposure. The level of significant difference was considered at p<0.05. Asterisk and pound symbols represent statistical significance (* and # p<0.05, ** and ## p<0.01, *** and ### p<0.001 and **** and #### p<0.0001).
3. Results 3.1 Determination of US intensity The output US power was determined by a radiation force balance technique. The US intensity at 100% DC was determined as 0.27 W/cm 2 and assumed as spatially averaged and temporally averaged. 3.2 Microscopic evaluation of MP destruction and degradation To evaluate the destruction and degradation of MPs (mean diameter = 7µm) following sonication, morphologic analysis of US-exposed MPs was performed. After US exposure (0.27W/cm 2 , 100% DC for 10s), the percentage of remaining MPs was significantly decrease (p<0.001) (Fig. 2A and B). There was no considerable difference among tested US intensities (0.27, 1.5 and 2.5 W/cm 2 (Fig. 2C). To examine the effect of repetitive sonication (0.27W/cm 2 , 100% DC for 10s) on MP degradation, the surface area of MPs was measured and analyzed. A greater degree of MP degradation was observed with increasing the repetition of US exposure. Compared to the control (x 0 sonication), there was a 34% reduction in surface
area of MPs with 1 sonication (p<0.01). It was further reduced by 63% following 4 sonications (p<0.0001) (Fig. 2D). 3.3 Morphology of sonicated-MPs by SEM SEM analysis showed morphologically intact and smooth surface in control MPs (7µm) (Fig. 3A and C). However, numerous cracks and holes appeared on the MP surface after US exposure (0.27W/cm 2 , 100% DC for 10s) (Fig. 3B and D). 3.4 Local cytotoxicity of US-mediated MP destruction To test the local cytotoxic effect of US-mediated MP destruction, two types of cell lines (B16 and EL4) in suspension containing MPs (1% w/v)) were sonicated. In the presence of MPs (7µm), cell viability remained above 95% when unsonicated. However, the cell viability was significantly decreased into around 50% (p<0.001) when exposed to US at 50% DC. It further dropped to approximately 5% at 100% DC (p<0.001). There were no significant differences of cell viability between B16 and EL4 following tested DCmodulated sonication (Fig. 4A). Since the cytotoxicity of US mediated MP destruction was not dependent on the types of cell lines, we arbitrarily chose B16 for performing the rest of experiments. To examine the cytotoxic effect of DC-modulated US exposure and in combination with MPs, B16 cells with or without 1% MPs (w/v) (7µm) in suspension were exposed to US at 0, 50 and 100% DC. In the absence of MPs, US exposure alone did not alter cell viability. Cell viabilities were all remained above 95% following US exposure regardless of DC. However, DC had a clear effect on cell viability when suspended with MPs (7µm). The cell viability was around 50% with 50% DC (p<0.0001) and 5% with 100% DC (p<0.0001) (Fig. 4B). To examine the mean size- or concentration- dependent cytotoxicity, either 7 or
17 µm of MPs were suspended with B16 cells at different levels of concentration followed by US exposure (0.27W/cm 2 , 100% DC for 10s). At concentration of 0.5% and 2.5% MPs (w/v) (7µm) in the cell suspension, we observed 50% (p<0.0001) and 10% (p<0.0001) of the remaining cells to be viable following US exposure. However, we observed no considerable cell death when the cells were suspended with 17µm MPs. Cell viability remained unchanged at above 95% regardless of tested concentrations (Fig. 4C). 3.5 US-triggered DOX release The pattern of sustained DOX release in MPs was observed approximately 6hr after the initial burst release. US exposure (0.27W/cm 2 , 100% DC for 10s) with DC modulation at 10, 50 and 100% was applied immediately at the beginning of experiments. During the period of burst release (<6hr), no clear difference of the release rate was observed. However, the difference became apparent when it reached the period of sustained release (>6hr) as compared to control. DOX release was significantly increased following sonication as compared to control at each subsequent time point (Fig. 5A). The DOX release was increased by 17% with 10 and 50% DC and 33% with 100% DC, as compared to control. The accelerated release kinetics remained higher until the pre-determined end time point of the experiments (Fig. 5A). To confirm that the US-accelerated of DOX release can be achieved during the period of sustained release (>6hr), sonication (0.27 W/cm2 at 100% DC for 10s) was applied at a time point of 48hr. The results showed that another burst release appeared immediately following US exposure and the accelerated release became significant at post 54hr (p<0.05) and at post 96hr (p<0.05), as compared to the time point immediately before sonication at 48hr (Fig.
5C and D). The DOX release rate remained unchanged in unsonicated control (Fig. 5B and C).
4. Discussion Linearly oscillating lipid- or albumin- shelled microbubbles have been extensively studied as an ultrasound contrast agent as well as a targeted drug delivery vehicle due to the fact that their acoustic properties are exceptionally dynamic depending on US pressure [1, 4, 5, 34]. However, US mediated polymeric MP destruction has received little attention even though there is evidence that polymeric MPs can be destroyed by US. Studies have suggested that the destruction of the polymeric MPs starts with creating defects and cracks in the shell surface by releasing encapsulated gas or air without linear oscillation [9, 30]. Nonetheless, there have been little studies about utilizing polymeric MP destruction in response to US exposure as a therapeutic strategy. Therefore, in this study we investigated the effects of US exposure on the destruction and degradation of MPs made of PLGA for potential tumor treatment. We chose PLGA as a source of MPs because it is regarded as one of the most promising biomaterials that can be naturally hydrolyzed and metabolized in the body with minimal toxicity. Additionally, it has well established patterns of controlled and sustained release of encapsulated drugs [22, 23]. We first examined the acoustic responses of MPs (mean diameter = 7µm) in terms of destruction and degradation. Light microscopy confirmed that approximately 40% of MPs were destroyed immediately following sonication (0.27 W/cm2 at 100% DC for 10s) as compared to control. We next examined the effects of higher intensity sonication on MP destruction by applying three different levels of US intensities, 0.27, 1.5 and 2.5 W/cm2 (at
100% DC for 10s). As compared to the un-sonicated control group, approximately 40% of MPs were destroyed by US exposure at 0.27W/cm2. However, no difference in MP destruction was observed amongst the tested US intensities. Although the difference of acoustic characteristics between destroyed and un-destroyed MPs remains unknown yet, it is probable that a certain population of MPs were US-sensitive due to their structure. These could be imploded immediately and the rest of the MP population was probably more resistant to the tested US. Since the MP destruction was independent to the tested US intensities, we arbitrarily chose the US intensity at 0.27 W/cm2 for the rest of study. Following US exposure (0.27 W/cm2 at 100% DC for 10s) on MPs, we quantified the surface area of MPs that could represent the degradation of MPs. We observed that the surface area of MPs was gradually reduced by increasing the number of sonications. Approximately 40% of surface area of MPs was reduced after the first sonication and 60% reduction was observed after 4 repeated sonications. These results are consistent our assumption that a certain portion of US-resistant MPs undergo accelerated degradation by repetitive sonication. This is likely in proportion with the amount of US waves affecting the MPs. Since we confirmed that MPs can be destroyed by US exposure, we examined the cytotoxic effect of imploding MP on nearby cells. Cytotoxicity is observed due to the fact that imploding microbubbles are known to generate shock waves which can mechanically rupture nearby cells [3, 6, 8]. First, we examined the effect of MP destruction on cytotoxicity in two different cell lines, B16 and EL4. We observed no different cell viability between tested cell lines suggesting that the cytotoxicity of MP destruction was primarily from instant mechanical shock waves [6, 8, 35]. Thus, we chose B16 for the rest
of experiments. The dependency of DC (50 and 100 %) and MP concentration (0.5 and 2.5 % (w/v)) on cell death was examined. We observed that the greater cell death was observed with higher DC and MP concentration. This may be due to several factors. These include that the greater DC produced the greater averaged US power affecting MPs resulting in a greater number of MP destruction. Additionally, the presence of more US-susceptible MPs in suspension, due to higher MP concentration, increases the number that are destroyed US exposure. We were intrigued by examining the different sizes of MPs (mean diameter = 17µm). We expected that the cytotoxicity of MP destruction would be similar for 7µm MPs. However, we observed little cell death with 17µm MPs. Although the reason for the inactivity of 17µm MPs responding to US exposure remained unclear, we speculate that the innate characteristics of 7µm MPs are more susceptible to US exposure than 17µm MPs. It is likely that the surface area of MPs directly interacting with US waves could be the primary sensitizing factor in terms of destruction or degradation because the percentage of US propagation could be widely affected depending on the available surface area of MPs. Further investigation is required to understand. It is also possible that the size dependent US susceptibility might be also dependent on other US parameters such as frequency, intensity and duration. Although, further investigation would be required to understand the US-parameter dependent MP destruction, we agreed that the 7µm MPs can have complete destruction by US exposure with the mechanism as previously suggested [9, 10, 36, 37]. Since certain populations of MPs were susceptible to destruction and degradation by US exposure, it was reasonable to hypothesize that the release of encapsulated drugs in
MPs can be controlled by sonication. To test this hypothesis, DOX was loaded in MPs and the release kinetics was evaluated. DOX was chosen because the release of DOX following cytotoxicity and poration due to MP destruction would have synergistic tumor-suppressive effects in our future study. In our first experiment, US exposures (0.27 W/cm2 for 10s) at 10, 50 and 100% DC were applied at the beginning of the experiments. All groups, including un-sonicated control, showed an initial burst release (<6hr) followed by a sustained release phase (after 6hr) of DOX from the MPs. The effect of US exposure on DOX release became apparent during the sustained release phase by increasing approximately 17% at 10 and 50 % DC and 33% at 100% DC as compared to un-sonicated control. This suggests that the destroyed and degraded MPs generated by sonication facilitated a faster release of DOX as compared to intact MPs. In order to confirm that USinduced DOX release can be achieved anytime irrespective to the release states (in either burst or sustained release state), we sonicated DOX-MPs once during the period of sustained release (at 48hr). Immediately following sonication, a new burst release occurred and the enhancement of DOX release became significant within 6hr following sonication. This suggests that breaking the MPs during sustained release further releases DOX. Further investigation is needed to confirm that accelerated MP degradation, by repetitive sonications, can further increase the DOX release. This may or may not occur because the DOX could be still possibly bound to broken pieces of MPs. There have been extensive attempts to use intravenously infused gas-filled microbubble destruction by US for targeted drug delivery into tumors. The effect, so fa, have proven to be promising [11-13]. However, to be successful, it requires that the tumors possess a well-formed vasculature. This is not typically the cases [14, 38, 39]. Also, the
potential risks of infusing microbubble-conjugated anti-cancer drugs through intravenous administration, remain unknown as it may lead to unnecessarily high concentrations of toxic drugs in systemic circulation [16, 17]. In this study, we demonstrated that DOX loaded MPs, made of PLGA, can be destroyed by US application producing cytotoxicity and accelerated DOX release from MPs. Considering that DOX-PLGA is directly infused into tumors, this could potentially have long-term benefits for the tumor treatment by controlled delivery of instant cytotoxicity from MP destruction as well as sustained DOX release with minimal invasiveness in combination with US exposure. This technique also provides for reduced potential risks of toxic chemotherapeutic drugs in systemic circulation.
5. Conclusion In summary, DC-modulated sonication can be used to control cytotoxicity and the release of DOX via MP destruction and degradation. Our future study focuses on the synergistic effects of a combination of US exposure and a single intra-tumoral injection of DOX-loaded PLGA-MPs. This synergistic effect of tumor suppression can occur through an instant cytotoxicity followed by sustained release of DOX from a minimally invasive manner. Our findings suggest that DOX-loaded PLGA MPs can be used as an effective tool for use as a therapeutic agent against tumor and cancer cells.
ACKNOWLEDGEMENTS This work was supported by the Bio & Medical Technology Development Program of the National Research Foundation (NRF) & funded by the Korean government (MSIP&MOHW) (No. 2017M3A9E4048170).
REFERENCES [1] T. Kodama, Y. Tomita, K. Koshiyama, M.J. Blomley, Transfection effect of microbubbles on cells in superposed ultrasound waves and behavior of cavitation bubble, Ultrasound Med Biol 32(6) (2006) 905-14. [2] P.A. Dijkmans, L.J. Juffermans, R.J. Musters, A. van Wamel, F.J. ten Cate, W. van Gilst, C.A. Visser, N. de Jong, O. Kamp, Microbubbles and ultrasound: from diagnosis to therapy, Eur J Echocardiogr 5(4) (2004) 245-56. [3] L.B. Feril, Jr., T. Kondo, Q.L. Zhao, R. Ogawa, K. Tachibana, N. Kudo, S. Fujimoto, S. Nakamura, Enhancement of ultrasound-induced apoptosis and cell lysis by echocontrast agents, Ultrasound Med Biol 29(2) (2003) 331-7. [4] P. Qin, L. Xu, W. Zhong, A.C. Yu, Ultrasound-microbubble mediated cavitation of plant cells: effects on morphology and viability, Ultrasound Med Biol 38(6) (2012) 1085-96. [5] M.M. Forbes, R.L. Steinberg, W.D. O'Brien, Jr., Examination of inertial cavitation of Optison in producing sonoporation of chinese hamster ovary cells, Ultrasound Med Biol 34(12) (2008) 2009-18. [6] Y. Feng, Z. Tian, M. Wan, Bioeffects of low-intensity ultrasound in vitro: apoptosis, protein profile alteration, and potential molecular mechanism, J Ultrasound Med 29(6) (2010) 963-74. [7] L.B. Feril, Jr., T. Kondo, Biological effects of low intensity ultrasound: the mechanism involved, and its implications on therapy and on biosafety of ultrasound, J Radiat Res 45(4) (2004) 479-89.
[8] M.A. Hassan, L.B. Feril, Jr., K. Suzuki, N. Kudo, K. Tachibana, T. Kondo, Evaluation and comparison of three novel microbubbles: enhancement of ultrasound-induced cell death and free radicals production, Ultrason Sonochem 16(3) (2009) 372-8. [9] S. Hernot, A.L. Klibanov, Microbubbles in ultrasound-triggered drug and gene delivery, Adv Drug Deliv Rev 60(10) (2008) 1153-66. [10] W.G. Pitt, G.A. Husseini, B.J. Staples, Ultrasonic drug delivery--a general review, Expert opinion on drug delivery 1(1) (2004) 37-56. [11] J. Du, L.F. Du, F.H. Li, X.Z. Zheng, H.L. Li, Q.S. Shi, Y. Wu, Ultrasound targeted microbubble destruction-mediated gene delivery system: application to therapy for ocular disease, Asian Biomed 5(5) (2011) 577-587. [12] C. Pu, S. Chang, J. Sun, S. Zhu, H. Liu, Y. Zhu, Z. Wang, R.X. Xu, Ultrasoundmediated destruction of LHRHa-targeted and paclitaxel-loaded lipid microbubbles for the treatment of intraperitoneal ovarian cancer xenografts, Mol Pharm 11(1) (2014) 4958. [13] H. Liu, S. Chang, J. Sun, S. Zhu, C. Pu, Y. Zhu, Z. Wang, R.X. Xu, Ultrasoundmediated destruction of LHRHa-targeted and paclitaxel-loaded lipid microbubbles induces proliferation inhibition and apoptosis in ovarian cancer cells, Mol Pharm 11(1) (2014) 40-8. [14] R.M. Terek, Angiogenesis in chondrosarcoma, Current Opinion in Orthopaedics 13(6) (2002) 449-453. [15] J.A. Buckwalter, The Structure of Human Chondrosarcoma Proteoglycans, J Bone Joint Surg Am 65(7) (1983) 958-974.
[16] S. Podell, C. Burrascano, M. Gaal, B. Golec, J. Maniquis, P. Mehlhaff, Physical and biochemical stability of Optison, an injectable ultrasound contrast agent, Biotechnol Appl Biochem 30 ( Pt 3) (1999) 213-23. [17] J.C. Hutter, H.M. Luu, P.M. Mehlhaff, A.L. Killam, H.C. Dittrich, Physiologically based pharmacokinetic model for fluorocarbon elimination after the administration of an octafluoropropane-albumin microsphere sonographic contrast agent, J Ultrasound Med 18(1) (1999) 1-11. [18] H. Chen, J.H. Hwang, Ultrasound-targeted microbubble destruction for chemotherapeutic drug delivery to solid tumors, J Ther Ultrasound 1 (2013) 10. [19] F. Zhu, Y. Jiang, F. Luo, P. Li, Effectiveness of localized ultrasound-targeted microbubble destruction with doxorubicin liposomes in H22 mouse hepatocellular carcinoma model, J Drug Target 23(4) (2015) 323-34. [20] J. Kang, X. Wu, Z. Wang, H. Ran, C. Xu, J. Wu, Z. Wang, Y. Zhang, Antitumor effect of
docetaxel-loaded
lipid
microbubbles
combined
with
ultrasound-targeted
microbubble activation on VX2 rabbit liver tumors, J Ultrasound Med 29(1) (2010) 6170. [21] J. Khandare, T. Minko, Polymer-drug conjugates: Progress in polymeric prodrugs, Prog Polym Sci 31(4) (2006) 359-397. [22] H.K. Makadia, S.J. Siegel, Poly Lactic-co-Glycolic Acid (PLGA) as Biodegradable Controlled Drug Delivery Carrier, Polymers-Basel 3(3) (2011) 1377-1397. [23] X. Zhu, R.D. Braatz, A mechanistic model for drug release in PLGA biodegradable stent coatings coupled with polymer degradation and erosion, J Biomed Mater Res A 103(7) (2015) 2269-79.
[24] K. Shahani, S.K. Swaminathan, D. Freeman, A. Blum, L.A. Ma, J. Panyam, Injectable Sustained Release Microparticles of Curcumin: A New Concept for Cancer Chemoprevention, Cancer Res 70(11) (2010) 4443-4452. [25] J.M. Lu, X. Wang, C. Marin-Muller, H. Wang, P.H. Lin, Q. Yao, C. Chen, Current advances in research and clinical applications of PLGA-based nanotechnology, Expert Rev Mol Diagn 9(4) (2009) 325-41. [26] M. Figueiredo, R. Esenaliev, PLGA Nanoparticles for Ultrasound-Mediated Gene Delivery to Solid Tumors, J Drug Deliv 2012 (2012) 767839. [27] F. Danhier, E. Ansorena, J.M. Silva, R. Coco, A. Le Breton, V. Preat, PLGA-based nanoparticles: an overview of biomedical applications, J Control Release 161(2) (2012) 505-22. [28] C. Deng, Y.J. Jiang, R. Cheng, F.H. Meng, Z.Y. Zhong, Biodegradable polymeric micelles for targeted and controlled anticancer drug delivery: Promises, progress and prospects, Nano Today 7(5) (2012) 467-480. [29] N. Rapoport, Physical stimuli-responsive polymeric micelles for anti-cancer drug delivery, Prog Polym Sci 32(8-9) (2007) 962-990. [30] S.H. Bloch, M. Wan, P.A. Dayton, K.W. Ferrara, Optical observation of lipid- and polymer-shelled ultrasound microbubble contrast agents, Appl Phys Lett 84(4) (2004) 631-633. [31] B. Zeqiri, C.J. Bickley, A new anechoic material for medical ultrasonic applications, Ultrasound Med Biol 26(3) (2000) 481-5. [32] R.C. Preston, Measurement and Characterization of the Acoustic Output of Medical Ultrasonic Equipment .2., Med Biol Eng Comput 24(3) (1986) 225-234.
[33] R.C. Preston, Measurement and Characterization of the Acoustic Output of Medical Ultrasonic Equipment .1., Med Biol Eng Comput 24(2) (1986) 113-120. [34] C.R. Mayer, N.A. Geis, H.A. Katus, R. Bekeredjian, Ultrasound targeted microbubble destruction for drug and gene delivery, Expert Opin Drug Deliv 5(10) (2008) 1121-38. [35] L.B. Feril, T. Kondo, Q.L. Zhao, R. Ogawa, K. Tachibana, N. Kudo, S. Fujimoto, S. Nakamura, Enhancement of ultrasound-induced apoptosis and cell lysis by echocontrast agents, Ultrasound in Medicine and Biology 29(2) (2003) 331-337. [36] P. Hauff, S. Seemann, R. Reszka, M. Schultze-Mosgau, M. Reinhardt, T. Buzasi, T. Plath, S. Rosewicz, M. Schirner, Evaluation of gas-filled microparticles and sonoporation as gene delivery system: Feasibility study in rodent tumor models, Radiology 236(2) (2005) 572-578. [37] D.J. May, J.S. Allen, K.W. Ferrara, Dynamics and fragmentation of thick-shelled microbubbles, Ieee T Ultrason Ferr 49(10) (2002) 1400-1410. [38] R.L. McGough, C.Z. Lin, P. Meitner, B.I. Aswad, R.M. Terek, Angiogenic cytokines in cartilage tumors, Clin Orthop Relat R (397) (2002) 62-69. [39] M. Ryzewicz, B.J. Manaster, E. Naar, B. Lindeque, Low-grade cartilage tumors: Diagnosis and treatment, Orthopedics 30(1) (2007) 35-46.
Fig. 1. Schematic illustration of experimental setup. The US transducer was immersed in degassed water and a cell suspension, either with or without PLGA MPs, was exposed to US from a distance of approximately 1 cm. US mediated MP destruction killed nearby cells.
Fig. 2. MP destruction and degradation by US exposure. (A) Light microscopic image showing 1% (w/v) MP suspension before US exposure. (B) Light microscopic image showing 1% (w/v) MP suspension after US exposure. (C) The number of remaining MPs was significantly reduced after US exposure and no considerable difference among tested intensities was observed. (D) The surface area of MPs was gradually reduced with more repetitive US exposure. *Statistically significant at p < 0.05 and NS represents no significance. Error bars represent standard deviation (n=3). Scale bar = 500µm.
Fig. 3. Morphological analysis of MPs by SEM. (A) Un-sonicated group showing intact and smooth surfaced MPs. (B) Appearance of cracks and holes at the surface of MPs was observed following US exposure. (C) Higher magnification of non-US exposed MPs. (D) Higher magnification of US exposed MPs. White arrows indicate holes in MPs that were created following sonication. Scale bar = 10µm (A and B) and 5 µm (C and D).
Fig. 4. The dependency of cell-type, DC, MP diameter, and MP concentration on local cell death following sonication. (A) The local cell death was increased by increasing DC. No considerable difference between tested cell lines (B16 and EL4) was observed. (B) Following sonication, little cytotoxicity was observed in the absence of MPs whereas significant cell death was observed by increasing DC in the presence of MPs. (C) Cell death was also proportional to the concentration of MPs (7µm); however, no cell death was observed with MPs (17µm) following sonication (n=3-5). Asterisk (*) and pound (#) represent statistical significances compared to control (
*** and ###
p<0.001 and
represent standard deviation (n=3-5).
**** and ####
p<0.0001) and NS represents no significance. Error bars
Fig. 5. Accelerated DOX release by US exposure. (A) Accelerated DOX release was dependent on DCmodulated US. (C) Sonication at a state of sustained release shows a new burst release. (B) Detailed release profile of control group be tween 45 to 55 hrs. (D) Detailed release profile of US exposed group between 45 to 55 hrs. Asterisk (*) represents statistical significances as compared to *
**
control ( p<0.05, p<0.01,
***
p<0.001 and
****
p<0.0001). Error bars represent standard deviation (n=3-6).