Article pubs.acs.org/Langmuir
Sonication−Microfluidics for Fabrication of Nanoparticle-Stabilized Microbubbles Haosheng Chen,† Jiang Li,*,‡ Weizheng Zhou,§ Eddie G. Pelan,∥ Simeon D. Stoyanov,∥,⊥,# Luben N. Arnaudov,∥ and Howard A. Stone*,∇ †
State Key Laboratory of Tribology, Tsinghua University, Beijing, 100084, China School of Mechanical Engineering, University of Science and Technology Beijing, Beijing, 100083, China § Unilever R&D Shanghai, 66 Lin Xin Road, Shanghai 200335, China ∥ Unilever Research & Development, 3133AT Vlaardingen, The Netherlands ⊥ Laboratory of Physical Chemistry and Colloid Science, Wageningen University, 6703 HB Wageningen, The Netherlands # Department of Mechanical Engineering, University College London, Torrington Place, London WC1E 7JE, United Kingdom ∇ Department of Mechanical and Aerospace Engineering, Princeton University, Princeton, New Jersey 08544, United States ‡
S Supporting Information *
ABSTRACT: An approach based upon sonication−microfluidics is presented to fabricate nanoparticle-coated microbubbles. The gas-in-liquid slug flow formed in a microchannel is subjected to ultrasound, leading to cavitation at the gas−liquid interface. Therefore, microbubbles are formed and then stabilized by the nanoparticles contained in the liquid. Compared to the conventional sonication method, this sonication−microfluidics continuous flow approach has unlimited gas nuclei for cavitation that yields continuous production of foam with shorter residence time. By controlling the flow rate ratios of the gas to the liquid, this method also achieves a higher production volume, smaller bubble size, and less waste of the nanoparticles needed to stabilize the microbubbles.
1. INTRODUCTION Microbubbles have a wide range of applications, such as ultrasound contrast agents in medical diagnostics,1 foams in food, cleansers in personal care products, and as templates to synthesize novel particles in petrochemical materials.2 Because the microbubbles are thermodynamically unstable, nanoparticles,3−6 lipids,7,8 polymers,9 and proteins10,11 have been used to cover the bubble surface to provide stability and prolong lifetime. The most common approach to realize stable covering is through ultrasound cavitation of slightly denatured and concentrated protein solutions, where a shell around the bubbles can be created by in situ cross-linking proteins on the bubbles’ surface due to the very high local temperatures and pressures at the moment of bubble collapse.10,11 An alternative approach is to use nanoparticles with intermediate hydrophobicity that can irreversibly adsorb and close-pack at the bubble surface due to the oscillations of the cavitation bubbles. However, the sonication method has a limitation on the production volume, as bubbles will no longer be generated when the gas nuclei contained in solution are used up. Recently, microfluidics with flow-focusing methods have been investigated to fabricate microbubbles stabilized by colloidal particles, polymers, and lipids.12−16 The microfluidic method17−19 can produce the bubbles continuously because the © 2014 American Chemical Society
dispersed gas phase can be injected into the continuous phase and is not limited by the gas nuclei in the solution. The drawback, however, stems from the fact that it is operated under laminar flow conditions, which has relatively low energy and mass transfer to achieve high production rates. Also, the bubbles fabricated by microfluidics are much larger than the stabilized cavitation bubbles formed by sonication. Therefore, the combination of sonication with microfluidics may overcome the limitations on the production volume and the bubble size. There are a variety of applications of ultrasound to microfluidics that followed after it was revealed that sonication can enhance mixing in microfluidic systems.20 This combination of sonication and microfluidics can be used to fabricate stabilized microbubbles by driving a microfluidic gas-in-liquid slug flow through the ultrasound field;21,22 here we refer to the process as sonication−microfluidics. The advantages of the sonication−microfluidics compared to common sonication approaches lie in the following: (1) The gas−liquid slug flow provides gas nuclei in microfluidic channels continuously; thus, the production volume of the stabilized microbubbles can be Received: February 5, 2014 Revised: March 28, 2014 Published: April 2, 2014 4262
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improved. (2) Microfluidic devices provide precise spatial and temporal control of cavitation; thus, it causes a narrower size distribution of the stabilized microbubbles and a shorter residence time for the production in the sonication field, which prevents formation of products from excessive sonication and overheating. (3) The area of the gas−liquid interfaces for cavitation can be controlled by the flow rate ratio of the gas to the liquid, which helps to reduce waste of the nanoparticles that stabilize the cavitation microbubbles. In this work, we demonstrate how to combine sonication with microfluidics to realize the fabrication of microbubbles coated by nanoparticles. The mechanism of cavitation at the gas−liquid interface in the microfluidic slug flow by sonication is investigated, and the advantages of sonication−microfluidics are demonstrated for increasing the production volume of smaller stable microbubbles with shorter residence time and less waste of the nanoparticles. As the nanoparticles and its processes are in principle food-grade, and the final diameter of the particle-stabilized microbubbles is around 10 μm, the microbubbles may have potential applications in the food industry.
2. EXPERIMENTAL SECTION Fabrication of the Sonication−Microfluidics Device. The sonication−microfluidics device to generate microbubbles consists of (i) a microfluidic device to generate gas-in-liquid slug flow and (ii) sonication to realize cavitation and the coating of the bubbles. As shown in Figure 1a, the microfluidic device consists of three capillary tubes: the injection tube, the collection tube, and the outer tube. The inlet of the injection tube is for the gas, and its end is tapered to an inner diameter of 50 μm using a pipet puller. The collection tube is PE/5 tubing (Scientific Commodities Co.) with an inner diameter of 860 μm. The injection and the collection tubes are fitted into the opposite ends of the outer tube, which is a square capillary with an inner dimension of 1.05 mm. In our experiments, air is injected into the injection tube with a flow rate Qg. The continuous phase is a water dispersion containing ethyl cellulose (EC) nanoparticles with diameter of 100−300 nm and is injected into the device at one end through the interstices between the outer tube and the injection tube at a flow rate Ql, while the other end of the outer tube is sealed. When the gas and the EC dispersion meet at the junction, the gas-in-liquid slug flow is formed. The gas-in-liquid slug flow is transported to the sonication field in the PE/5 tubing, which is immersed in pure water, and a 6mm-diameter vibration horn of an ultrasonic dismembrator is placed above the tubing at a distance of 1 mm. The vibration horn is driven to vibrate with a fixed frequency of 20 kHz and a maximum amplitude of 12 μm; the power can be adjusted from 50 to 600 W, which determines the intensity of sonication. The velocity and the number of bubbles passing the sonication field can be controlled by the flow rate of the two phases. Ethyl Cellulose Dispersions Used in the Sonication−Microfluidics Experiment. EC powder with an ethoxyl content of 48% (Sigma-Aldrich, 247499-100G, batch 08521KH) was dissolved in acetone at 30 °C for 30 min to obtain a clear 1.0 wt % EC solution. Then, the same volume of deionized water was quickly poured into this solution under stirring to cause the formation of EC colloidal particles. The resulting turbid dispersion was put in a rotary evaporator (Buchi R-200, Heidolph), where all of the acetone and part of the water were removed to obtain a 1.0 wt % EC dispersion in water. The residual amount of acetone was below 0.01 mg/mL, indicating that the acetone had been removed from the solution containing EC particles. The obtained EC dispersions were stored in a refrigerator at a temperature of 4 °C. Before the EC dispersions were used in an experiment, 0.035 M citric acid aqueous solution was added in the diluted dispersion to tune the pH to 3 to ensure good foamability and foam stability. The size of the EC particles was then controlled to be
Figure 1. (a) Schematics of the sonication−microfluidics device to fabricate nanoparticle-coated bubbles. (b) Bubbles in the EC dispersion pass through the ultrasound field in the PE/5 tubing, and clusters of the micro-cavitation bubbles are formed in the intervals between each pair of air bubbles. The slug flow and bubbles are observed using a high-speed camera (Phantom M110). A video file is provided as Video-A in the Supporting Information. (c) Microscope images of the EC bubbles. (d) Cryo-SEM image of the cross section of the surface of a microbubble coated by nanoparticles. around 100−300 nm. Note that the surface properties of the EC particles have been studied in detail in ref 23. Measurement of the Turbidity of the EC Dispersions. The concentrations of the EC nanoparticles in the dispersions before and after the sonication experiments were determined by a turbidimeter (Hach 2100N). Before the measurement, the relationship between the concentration of the EC nanoparticles in the dispersion and its turbidity was calibrated. Then, the dispersions with 1.0 wt % EC nanoparticles were used in sonication experiments with six different gas-to-liquid flow rate ratios, which were collected after the two phases flow out of the tubing. The collected residual EC dispersions were measured by the turbidimeter, and according to the calibration, the concentrations of the EC nanoparticles in the dispersions were determined.
3. EXPERIMENTAL RESULTS First, pure water in the absence of air bubbles was injected into the device and flowed through the sonication field, and we did not observe cavitation, even when 0.5 wt % sodium dodecyl sulfate (SDS) was added to the water (see Figure S1 in Supporting Information). Then, EC dispersions were tried and no bubbles were generated (see Figure S2 in Supporting Information). Finally, air bubbles were injected in water and the resulting gas-in-liquid slug flow was driven to pass through the sonication field. At this time, cavitation bubbles were generated. When the liquid phase did not contain EC particles, the cavitation bubbles collapsed and disappeared immediately after they exited the low-pressure zone of the sonication field, due to the fact that there was not enough resistance at the interface to 4263
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prevent bubble implosion and collapse. But when the liquid phase contained EC particles, we observed that cavitation bubbles were stabilized by the EC particles attached at their surfaces (EC bubbles). As shown in Figure 1b, when the gasdispersion slug flow was formed and entered a 300 W ultrasound field, cavitation occurred at the gas−liquid interface, and the micro-cavitation bubbles were generated from both ends of the injected macrobubbles. As a result of the recirculating flow field between the two macrobubbles,24 the generated micro-cavitation bubbles aggregated at intervals between the two injected macrobubbles. The cavitation bubbles collapsed in pure water, while bubbles in dispersions of EC nanoparticles are stabilized by the aggregation of the nanoparticles, which are compacted together at the bubble surface. The detailed mechanism for the particles to stabilize the bubbles has already been documented.25 Bubbles were collected and observed under a microscope, as shown in Figure 1c. Some of the bubbles were not spherical and as smooth as ordinary gas bubbles, because of the coverage of the nanoparticles on their surfaces. The Cryo-SEM image of Figure 1d shows the cross section of a collected microbubble, and the nanoparticles are seen clearly on the surface, where they are compacted tightly to form multilayers that stabilize the bubble. The EC bubbles can survive for a long time, and their size approached a stable value around 10 μm after the first hour and then changed less than 10% in the following 5 days (see Figure S3 in Supporting Information). The spatial and temporal control of cavitation in sonication− microfluidics reduces the residence time for bubble production in the sonication field. For example, with a flow rate of 25 mL/ h, the residence time for the bubbles to pass a 6 mm section of the ultrasound field is less than 1 s. The short residence time avoids the overheating that is common in ultrasound-assisted reactions in bulk solutions.26 In our experiment, the temperature of the EC dispersion was raised from 20 to 80 °C in 7 min when the conventional sonication was used, and obvious flocculation was found in the dispersion (see Figure S4 in Supporting Information). Therefore, the sonication−microfluidic method is also suitable for biomaterials that cannot sustain excessive sonication and overheating, such as proteins or living bacteria, which are now commonly contained in droplets to react with chemicals in cell assays.27
Figure 2. (a) A capillary wave formed on the bubble surface with the wavelength of λ in an ultrasound field. A video file is provided as Video-B in the Supporting Information. The scale bar is 1 mm. (b) The size distribution of the stabilized bubbles under different sonication powers. Typical microscopic images are shown in Figure S5 in the Supporting Information.
which produces clusters of the EC bubbles, as shown in Figure 2a. Sufficiently high driving power was found to be necessary to generate the EC bubbles. When the power was low, for example, 50 W, the surface of the injected bubble oscillates, but no EC bubbles are generated in the dispersion. As the sonication power increased to 600 W, cavitation occurred and the size of the cavitation bubbles was related to the power of the sonication. At relatively lower power (100−200 W), the cavitation bubbles were polydisperse. At higher power (400− 600 W), bubbles have a much narrower size distribution, as shown in Figure 2b. We have performed systematic experiments that demonstrate that (1) the production rate of microbubbles is linearly related to the flow rate of the slug flow and (2) the nanoparticles can be used more efficiently by adjusting the flow rate ratio of gas to the liquid phase. For example, the flow rates of the gas and the liquid phases were changed to demonstrate the advantage of the sonication−microfluidics on the production rate of the microbubbles. In these experiments, the sonication power was kept at 300 W, and the flow rates of the gas were Qg = 7.5, 15, and 30 mL/h; the corresponding flow rates of the liquid were Ql = 12.5, 25, and 50 mL/h to maintain the gas-to-liquid flow rate ratio equal to 0.6. Thus, as the flow rate increases, more macrobubbles will pass through the ultrasound field to let more gas−liquid interfaces take part in the cavitation in the same processing time; thus, the production rate of microbubbles is increased, and if collected at the end of the process, more foam can be obtained. Figure 3 shows the resultant foam volume versus time corresponding to the different flow rates. The results illustrate that the production rate of the foam of EC bubbles is linearly related to the flow rate of the slug flow, which can be understood because the area of the gas−liquid interface taking part in the cavitation has a linear relationship to the number of the bubbles passing through the ultrasound field at the same sonication power. This flow-rate-controlled production rate in sonication− microfluidics overcomes the limitation of the production rate by common sonication. For example, as shown by Figure 3a,
4. DISCUSSIONS AND ANALYSIS We next investigated the mechanism of cavitation at the gas− liquid interfaces in the sonication−microfluidics method. When the gas-in-liquid slug flow enters the sonication field, a Faraday wave is generated on a gas−liquid interface and the wavelength λ of capillary waves is given by the dispersion equation: λ = [(2πσ)/( f 2ρ)]1/3, where σ = 0.065 N/m is the interfacial tension, ρ = 1000 kg/m3 is the density of water, and the surface oscillation frequency f is half of the driving frequency.28 Thus, the expected wavelength of the capillary wave is 344 μm, which compares favorably with the value of about 300 μm observed in the experiment, as shown in Figure 2a. As the wave amplitude increases, liquid crests will be formed at the gas−liquid interface, which has been well-explained during the observation of vibration-induced drop bursting.29 Under this condition, part of the gas would be entrapped between neighboring and coalescing crests of the interface,21,22 and the entrapped gas is separated from the injected bubble. The entrapped gas nuclei grow and oscillate in the sonication field, and then microcavitation bubbles are formed and coated by the nanoparticles, 4264
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Figure 4. (a) The turbidity of the dispersions collected from the slug flow with different flow rate ratios of the gas to the solution containing EC nanoparticles after the sonication−microfluidics experiment. The red circle represents the turbidity of the EC dispersions before the sonication−microfluidics experiment. The slug flow passing the sonication field in the microchannels with different flow rate ratio of gas to EC dispersion Qg:Ql = (b) 5:25, (c) 10:25, (d) 20:25, and (e) 25:25 mL/h.
Figure 3. (a) The production rates of the EC microbubbles using the sonication−microfluidics method with different flow rates, with the result compared to that using only sonication. The power of the sonication was kept at 300 W for all the four different working conditions. Flow rate 1 was Qg:Ql = 7.5:12.5 mL/h, flow rate 2 was Qg:Ql = 15:25 mL/h, flow rate 3 was Qg:Ql = 30:50 mL/h, and the corresponding production rates of the sonication−microfluidics were, respectively, 18.5, 34.2, and 66.5 mL/h. The inset shows the bubbles collected in a glass tube at a time of 180 s using sonication− microfluidics. (b) The linear relationship between the foam volume and the injected gas volume.
consumption of the EC nanoparticles. Moreover, when the ratio of the gas to liquid continued to increase, for example, Qg:Ql = 25:25 mL/h, the gas bubbles became longer and fewer bubbles were passing through the ultrasound field at the same flow rate, as shown in Figure 4e. Thus, fewer cavitation bubbles were generated in the same processing time and fewer EC nanoparticles consumed for coating the bubbles. On the other hand, a relatively low gas-to-liquid flow rate ratio does not give the best coating efficiency, which is not surprising because the gas bubbles were fewer and smaller under the low flow rate ratio, and fewer cavitation bubbles were generated, as shown in Figure 4b. Therefore, there is an optimum gas-to-liquid flow rate ratio to obtain a high coating efficiency of the nanoparticles, as shown in Figure 4a,c.
the foam volume made by sonication in a 10 mL EC dispersion quickly (within 200 s) approached a limit of 2 mL in the beginning of sonication, as the dissolved gas in the dispersion was used up. On the other hand, when we use sonication− microfluidics and even a smaller volume of liquid, the foam volume increased linearly with time, as presented in Figure 3a, and generated even more foam than the traditional method. Thus, the production rate can be controlled by the flow rates of the fluids in the microchannels. Moreover, the foam volume has a linear relationship to the injected gas volume, as presented in Figure 3b. Next, we studied the influence of the flow rate ratio of the gas to the liquid phase to demonstrate the efficiency of the nanoparticles used on coating bubbles in sonication−microfluidics, as shown in Figure 4. In these experiments, the flow rate of the continuous liquid phase was kept at 25 mL/h while the gas flow rate was changed from 5 to 30 mL/h. The attachment of the EC nanoparticles on the microbubble surface can be determined by measuring the turbidity of the dispersion before and after the experiment. In particular, the decrease of the turbidity represents the lower concentrations of the EC nanoparticles remaining in the dispersion after the experiment, which indicates that more nanoparticles were consumed in forming EC microbubbles. We measured the turbidity of the dispersions after they pass the sonication section under different flow rate ratios of the gas to the liquid, as shown in Figure 4a. There is a maximum in the consumption of the nanoparticles in dispersion. When the flow rate ratio of the gas to the liquid was relatively high, for example, Qg:Ql = 20:25 mL/h, the bubbles were large and occupied more space in the microchannel in the sonication field, but the cavitation occurred only at the ends of the bubbles, as shown in Figure 4d. Thus, the area of the gas−liquid interfaces that took part in the cavitation was small, and fewer cavitation microbubbles were generated, which results in a low
5. CONCLUSION In summary, we developed the sonication−microfluidics method for particle-stabilized bubbles that is based on combining conventional microfluidics, where gas is injected into an aqueous suspension to generate macrobubbles, with ultrasound, which leads to formation of microbubbles. Also, the mechanism of formation of the cavitation bubbles at the microfluidic gas−liquid interfaces has been investigated. By increasing the flow rate of the gas and liquid phases, the production rate of stable EC bubbles is increased. Also, by controlling the gas-to-liquid flow rate ratios, the efficiency of the usage of the nanoparticles can be improved. Moreover, this sonication−microfluidics method reduces residence time for the products in the sonication field, which reduces exposure to high temperatures and high pressures.
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ASSOCIATED CONTENT
S Supporting Information *
The experimental results of water and EC dispersion in sonication−microfluidics, the stability of the EC-stabilized bubbles, the temperature increases in EC dispersion with conventional sonication, and videos of the gas-in-liquid slug flow passing through the ultrasound field, the capillary wave 4265
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forming on the bubble surface in ultrasound field, and water and EC dispersion flows in sonication−microfluidics. This material is available free of charge via the Internet at http:// pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Authors
*J.L. e-mail:
[email protected]. *H.A.S. e-mail:
[email protected]. Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This research is supported by NSFC Project (No. 51275266, No. 51322501, and No. 51275036) and Unilever Research Funds.
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