A microfluidic approach for the generation of perfluorocarbon nanodroplets as the primary emulsion with diameters as small as 300-400 nm is described. The system uses a pressure-controlled delivery of all reagents and increased viscosity in the continuous phase to drive the device into an advanced tip-streaming regime, which results in generation of droplets in the sub-micrometer range. Such nanodroplets may be appropriate for emerging biomedical applications

Bardin, David

Dayton, Paul A.

Lee, Abraham P.

Martz, Thomas D.

Sheeran, Paul S.

PubMed

Carolina Digital Repository

Microfluidic generation of acoustically active nanodroplets **Thomas D. Martz1, David Bardin2, Paul S. Sheeran3, Abraham P. Lee2, and Paul A.Dayton3,*1Curriculum of Applied Science & Engineering - Material Science, University of North Carolina-North Carolina State University, Chapel Hill, NC, USA2Department of Biomedical Engineering, University of California-Irvine, Irvine, California, USA3Joint Dept. of Biomedical Engineering, University of North Carolina-North Carolina StateUniversity, Chapel Hill, NC, USALiquid perfluorocarbon (PFC) droplets have emerged in medical acoustics as acousticallyactivatable emulsions for a variety of applications. The phase-shift of the inner core of aperfluorocarbon droplet from the liquid state to the gas state as a result of an appliedultrasound pressure is most commonly referred to as acoustic droplet vaporization (ADV).[1]While most ADV investigations have focused on droplets in the micron range forintravascular procedures, there has been much recent interest in the utility of sub-micronliquid perfluorocarbon droplets for extravascular applications such as drug delivery,enhanced thermal ablation, and diagnostic imaging and tissue characterization.[2,3]Capitalizing on the characteristically ‘leaky’ vasculature of many solid tumors, sub-microndroplets may extravasate into the tumor interstitium through inter-endothelial gaps andaccumulate[4], or may be able to traverse the lymphatic system. One main limitation ofcurrent methods to generate these nanodroplets is the use of processing techniques such asemulsification[5] and sonication[6], which result in polydisperse populations. Significantvariation is reported in the activation threshold due to the strong correlation betweenvaporization energy and perfluorocarbon droplet size.[7,8] Additionally, the biodistributionof sub-micron ADV agents may improve by narrowing the size distribution. Substantialclinical motivation thus exists for a precise method to generate perfluorocarbon nanodropletsfor use in extravascular tumor therapies. The emergence of droplet-based microfluidicsystems has enabled the generation of droplets in the micrometer diameter range with highuniformity[9,10]; yet, the generation of nanodroplets from devices with microscale geometrystill presents a fundamental challenge. Previous microfluidic approaches to producenanodroplets have relied on the inherently inefficient dynamic sorting of small satellitedroplets (as small as 100 nm) from microscale droplet populations as the basis of amonodisperse submicron emulsification system.[10,11,12] To overcome this limitation, wehypothesized that by modulating the basic mechanism of droplet generation to access a morefavorable droplet formation regime[13,14,15], we could yield an efficient approach for thegeneration of perfluorocarbon nanodroplets as the primary emulsion.Our group recently demonstrated the feasibility of microfluidic generation of microscaleADV agents to achieve precision control of droplet size and uniform activation.[7, 15] In thisstudy, we advance upon this prior work with the development of a microfluidic systemcapable of generating populations of primary sub-micron droplets with diameters as small as300–400 nm through two key refinements of the previously explored systems. In the earliest*Corresponding Author: Prof. Paul A. Dayton, Thomas D. Martz, and Paul S. Sheeran, 304 Taylor Hall, CB 7575, Chapel Hill, NC27599 (USA), padayton@bme.unc.edu.David Bardin and Prof. Abraham Lee, 3406 Engineering Hall, Irvine, CA, 92697, (USA)NIH Public AccessAuthor ManuscriptSmall. Author manuscript; available in PMC 2013 June 25.Published in final edited form as:Small. 2012 June 25; 8(12): 1876–1879. doi:10.1002/smll.201102418.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author Manuscriptstudy, syringe pumps were used to control input rates of all reagents, but the mechanicalvariability inherent to syringe-pump driven systems[16] led to a lower limit of approximately7 μm in diameter.[7] By controlling the dispersed perfluoropentane phase with a custompressure-controlled input similar to that used in prior studies[16, 17], droplet sizes werereduced to 3–5 μm.[15] Here, we demonstrate that pressure-controlled delivery of allreagents (Figure 1a) affords the ability to drive the microfluidic device into a tip-streamingregime[18] with extremely low variability – enabling sub-micron droplet production despitethe fact that the orifice width is over an order of magnitude larger than the dropletsthemselves (Figure 1b). We also observed that increasing the viscosity of the continuousphase through the addition of glycerol greatly enhanced the ability to drive dropletproduction stably in the sub-micron regime at the high pressures used. Relative to satellitedroplet sorting, advanced tip-streaming, as we demonstrate, allows for efficiency ingenerating droplets in the sub-micron range, without filtration or dynamic in-deviceseparation.By precisely varying the pressures and, consequently, the flow rates of the continuous anddispersed phases, we were able to produce narrowly dispersed lipid-encapsulated nano- andmicrodroplets ranging from 360 nm to 11 μm with the same microfluidic device (Table 1).Although some variability was observed between devices generated from the same mask,droplets between 3 μm and 11 μm were achieved by driving the dispersed phase with 12–15psi with the continuous phase at 33–35 psi. Droplets on the order of 800–900 nm wereachieved by driving the dispersed phase with 16–17 psi with the continuous phase kept near44–45 psi. By driving the dispersed phase with a pressure of 19–20 psi while the continuousphase was kept at 50 psi, droplets on the order of 300–400 nm were produced. Channel“blowouts” were observed when pressures on the order of 60 psi were used to drive reagentflow.Droplets larger than 1 μm were quantified with high-speed light microscopy as they exitedthe orifice. Smaller droplets could not be resolved with traditional light microscopy, andhence were collected and then assessed by dynamic light scattering (DLS) with a MalvernNanosizer (Figure 2) after production (thus, production rate data was not obtained for theseflow conditions). Transmission electron microscopy (TEM) of a number of the samples wereused to verify DLS results (Figure 3). The average diameter measured by TEM over 30droplets was 312 ± 49 nm, which is in excellent agreement with the DLS measurements forthe same sample (Figure 2, Sample 1). One of the consequences of using a lipid-basedcontinuous phase in conjunction with microfluidics is a high degree of naturally-occurringlipid vesicles, typically on the order of 100 nm or less. In these experiments, the presence oflipid vesicles was confirmed by both DLS and TEM, appearing as peaks on the order of 30 –70 nm in DLS measurements (Figure 2), and resulting in a heterogeneous background inTEM images composed of a high number of clearly visible non-PFC particles below 100 nmin size (Figure 3).In order to confirm the resulting samples were acoustically vaporizable, droplets weresonicated with 100-cycle pulses of varying amplitude at 3.2 MHz. As describedpreviously[7,8,19], the pressure required to vaporize droplets increases as droplet sizedecreases. In this study, a peak rarefactional pressure of approximately 3.15 MPa wassufficient to vaporize the majority of droplets in focus when the droplet size was greater thanapproximately 7 μm in diameter. As expected, smaller samples required substantially morepressure to vaporize. Droplets near 0.9 μm in diameter required approximately 4 MPa tovaporize, while smaller droplets could not be vaporized due to the maximum output limit ofour ultrasound system (4.3 MPa). Activation thresholds established from this study are inagreement with thresholds previously identified for individual perfluoropentane droplets.[7]Martz et al. Page 2Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptOur results demonstrate that sub-micron (and micron) droplets for acoustic dropletvaporization applications can be effectively generated using droplet-based microfluidics.The pressure-controlled reagent delivery system crucially enabled the quick and precisetuning of the continuous and dispersed phases to maintain constant rates of flow andconsistent tip-streaming. By contrast, in our prior studies utilizing mechanical pumping, wewere unable to maintain the tip-streaming of sub-micron droplets for sufficient time tocollect samples. To our knowledge, this study is one of the first examples of sub-micronproduction of primary liquid droplets from a microfluidic device. Future studies are neededto determine the extent that similarly small droplets can be produced with reagents otherthan those used here. The ability to achieve sub-micron droplet production, as well as theability to produce droplet populations of uniform size, will have a significant impact inburgeoning medical applications requiring acoustically active nano- and microdroplets.Experimental SectionReagent PreparationThe continuous phase lipid solution was prepared using a 9:1 molar ratio of 1,2-distearoyl-sn-glycero-3-phosphocoline (DSPC) (Avanti Polar Lipids, Alabaster, AL) and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N[methoxy-(polyethylene glycol)-5000](DSPE-PEG5000) (Avanti Polar Lipids, Alabaster, AL) in saline (lipid concentration of 1.5mg/mL). Lipid solution was combined with glycerol (Fisher Scientific, Pittsburgh, PA) andPluronic F-68 (Sigma Aldrich Corporation, St. Louis, MO) in a 1:1:1 mL volumetric ratio.Liquid perfluoropentane (FluoroMed, Round Rock, Texas) was utilized as the dispersedphase.Microfluidic Device FabricationThe process for developing the microfluidic wafer has been previously described byHettiarachchi et al.[20] Briefly, microfluidic channels were designed using Illustrator(Adobe, San Jose, CA) and printed at 20,000 dpi by CAD/Art Services (Brandon, OR). In aclass 10,000 clean room, a silicon wafer was spin-coated with a UV-curable epoxy(SU-8-25, Micro-Chem, Newton, MA) and exposed to UV light to develop the channels as anegative mold. Bulk polydimethylsiloxane (PDMS) (Dow Corning, Midland, MI) and acuring agent were cast over the wafer at a 10:1 ratio, and the mold was cured at 60°C for atleast 8 h. Microfluidic chips were separated from the wafer and cleaned along with the glassslides (2947-75 × 25, Corning, Corning, NY) in a plasma cleaner (Harrick Plasma, Ithaca,NY). The device components were exposed to a plasma field under a controlled pressure of500 mTorr for 3 min. Upon removal, the microfluidic channels and glass slide were joined,creating a sealed microfluidic device. Then the channels were filled with deionized water tostabilize the hydrophilic environment.Optical ObservationsThe microfluidic device was secured to a moving stand on an inverted microscope (Olympus1 × 71 microscope, Center Valley, PA) with a 50X NA = 1.0 objective. A high-speedcamera (Fastcam APX-RS, Photron, Inc., San Diego, CA) was used to capture images andvideos of the device output.Pressure Control Flow SystemThe reagent pump system was established in a similar manner to Korczyk et al.[16] Thepressure to control the flow system was supplied by a compressed nitrogen tank connectedin parallel to two precision pressure regulators that were used to control the pressure to eachreagent independently. A 60 max psi regulator was used to control the continuous phase, anda 30 max psi regulator was used for the dispersed phase (R-800-60 and R-800-30, Air-TrolMartz et al. Page 3Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptComponents, Inc., New Berlin, WI). The output of each regulator was connected via TygonS-54-HL tubing (Saint-Gobain Performance Plastics, Akron, OH) to provide the pressurehead to the headspace of a sealed and capped 3 mL glass vial (Figure 1a). Another length oftubing submerged in the reagent (typically 2 mL total) supplied the pressurized fluid to themicrofluidic device. By placing a positive pressure head in the vial, the reagent wastransferred to the channels of the microfluidic device, and the output rate controlled bymanually adjusting the pressure regulators. The system was allowed to stabilize for 1 minuteafter pressure changes were made before sample collection began.Optical Imaging and SizingDimensions of the final microfluidic device were determined using Image J (NIH, Bethesda,MD) and a calibrated slide reticule (Edmund Optics, Barrington NJ). The orifice, PFPchannel, and lipid solution channel, and filter spacing widths measured 12 μm, 46 μm, 45μm, and 23 μm, respectively. For micron-sized droplets, diameter was likewise determinedoptically. Sub-micron droplets could not be resolved optically, and were instead collected ina pipette tip for light scattering or electron microscopy analysis.Dynamic Light Scattering SizingA Malvern Nano ZS (Malvern Instruments Ltd., Malvern, Worcestershire, U.K.) was used todetermine the size of droplets in the sub-micron diameter range. Sub-micron samples werediluted with 0.5 mL of phosphate buffered saline and placed into a cuvette for testing.Debris less than 100 nm in diameter were assumed to be lipid vesicles.Transmission Electron Microscopy (TEM) and SizingTo verify the presence of perfluorocarbon nanodroplets and corroborate the Malvern sizingdata, a negative staining procedure was used alongside TEM (Philips Tecnai 120kV,Amsterdam, The Netherlands). The negative staining first consisted of plasma treating theindividual TEM grids for 90 seconds. Ten μl of the droplet sample was transferred viapipette onto the TEM grid and allowed to sit for 120 seconds. The TEM grid was dipped in abath of 1% uranyl acetate for 60 seconds, and allowed to dry prior to TEM imaging.Acoustic Droplet VaporizationTo verify that the generated droplet samples could be vaporized, a procedure similar to thatdescribed in Sheeran et al.[19] was utilized. In brief, a 3.2 MHz transducer with a focallength of 3.45 cm was used to activate the droplets in a cellulose tube submerged in a waterbath at 37°C. Droplets within the optical focus were subjected to manually-triggered 100-cycle pulses with 1–2 seconds of rest between each trigger. Pressure was increased in stepsof approximately 0.15 MPa until the pressure was sufficient to vaporize the majority(approximately 90% or greater) of the droplets in focus.AcknowledgmentsThe authors would like to thank Jason Streeter for producing the initial lipid composition and Roger Shih formanufacture of the master wafer. We appreciate the intellectual contributions of Dr. Terry Matsunaga on medicaluses and formulation methods for perfluorocarbon droplets. This grant was supported by R01EB008733 andR21EB011704.References1. Kripfgans OD, Fowlkes JB, Miller DL, Eldevik OP, Carson PL. Ultrasound Med Biol. 2000;26:1177–1189. [PubMed: 11053753]2. O’Neill BE, Rapoport N. Ther Deliv. 2011; 2:1165–1187. [PubMed: 22059114]Martz et al. Page 4Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author Manuscript3. Sheeran PS, Dayton PA. Curr Pharm Des. 2012 In Press.4. Campbell RB. Anticancer Agents Med Chem. 2006; 6:503–512. [PubMed: 17100555]5. Kawabata K, Sugita N, Yoshikawa H, Azuma T, Umemura S. Jpn J Appl Phys. 2005; 44:4548–4552.6. Mohan P, Rapoport N. Mol Pharm. 2010; 7:1959–1973. [PubMed: 20957997]7. Martz TD, Sheeran PS, Bardin D, Lee AP, Dayton PA. Ultrasound Med Biol. 2011; 37:1952–1957.[PubMed: 21963036]8. Kripfgans OD, Fabiilli ML, Carson PL, Fowlkes JB. J Acoust Soc Am. 2004; 116:272–281.[PubMed: 15295987]9. Nisisako T, Torii T, Higuchi T. Lab Chip. 2002; 2:24–26. [PubMed: 15100856]10. Tan YC, Fisher JS, Lee AI, Cristini V, Lee AP. Lab Chip. 2004; 4:292–298. [PubMed: 15269794]11. Tan YC, Lee AP. Lab Chip. 2005; 5:1178–1183. [PubMed: 16175277]12. Tan YC, Cristini V, Lee AP. Sens Actuators, B. 2006; 114:350–356.13. Anna SL, Mayer HC. Phys Fluids. 2006; 18:121512.14. Cubaud TC, Mason TG. Phys Fluids. 2008; 20:053302.15. Bardin D, Martz TD, Sheeran PS, Shih R, Dayton PA, Lee AP. Lab Chip. 2011; 11:3990–3998.[PubMed: 22011845]16. Korczyk PM, Cybulski O, Makulska S, Garstecki P. Lab Chip. 2010; 11:173–175. [PubMed:20949204]17. Thorsen T, Roberts RW, Arnold FH, Quake SR. Phys Rev Lett. 2001; 86:4163–4166. [PubMed:11328121]18. Ward T, Faivre M, Stone HA. Langmuir. 2010; 26:9233–9239. [PubMed: 20334409]19. Sheeran PS, Wong VP, Luois S, McFarland RJ, Ross WD, Feingold S, Matsunaga TO, Dayton PA.Ultrasound Med Biol. 2011; 37:1518–1530. [PubMed: 21775049]20. Hettiarachchi K, Zhang S, Feingold S, Lee AP, Dayton PA. Biotechnol Prog. 2009; 25:938–945.[PubMed: 19455647]Martz et al. Page 5Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptFigure 1.A) Diagram of pressure-controlled reagent flow, achieved via compressed N2 driving thereagent through tubing to microfluidic inputs. B) Microscopy of flow-focusing orificeillustrating tip-streaming, resulting in the generation of sub-micrometer droplets.Perfluoropentane (center channel) is surrounded by lipid solution (outer channels). Arrowsindicate direction of flow. C) Microscopy of droplets on the order of 800 nm in diameter(larger than shown in the example in (B)) taken downstream of the orifice as the dropletstream widens, illustrating very small, uniform-size production.Martz et al. Page 6Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptFigure 2.Particle size distribution produced by DLS for different sub-micron samples produced by thesame microfluidic device at different driving pressures. Sample 1 had a mean diameter of850 ± 150 nm, while Sample 2 had a mean diameter of 360 ± 50 nm in the main peak. Alsopresent in the measurement for Sample 2 are naturally-forming lipid vesicles less than 100nm in diameter (confirmed by TEM images). For this sample, the lipid vesicles had a meandiameter of 49 ± 7 nm. Each plot is an average of 3 repeat data sets.Martz et al. Page 7Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptFigure 3.Transmission electron microscopy example of negative-stained sub-micron perfluorocarbondroplets. The highly-stained droplet circumference appears as a black ring around thedroplet core. Lipid vesicles present in the continuous phase can be seen in the TEM imagebackground as semi-circular particles with diameters less than 100 nm, present in highquantities – in agreement with DLS measurements.Martz et al. Page 8Small. Author manuscript; available in PMC 2013 June 25.NIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptNIH-PA Author ManuscriptMartz et al. Page 9Table 1Droplet Production Characteristics for Various Samples.Mean Diam. (μm) Standard Deviation (μm) Production Rate (droplets/sec)Input Pressure (psi)†:Continuous PhaseInput Pressure (psi)†:Dispersed Phase0.36 0.05 * 50 19–200.85 0.15 * 44–45 16–173.6 0.2 8.7×10433–35 12–156.8 0.2 4.0×10411.7 0.3 3.2×104*indicates rate is below optical resolution† range represents observed variation between devicesSmall. Author manuscript; available in PMC 2013 June 25.

Microfluidic Generation of Acoustically Active Nanodroplets

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