Surfactant-free, UV-curable core–shell microcapsules in a hydrophilic PDMS microfluidic device

Core–shell microcapsules play a significant role in research and commercial products. These microcapsules serve as miniature containers for the storage and delivery of substances.^1,2 The current production methods include emulsion-based,^3,4 liquid marble-based,^5,6 and microfluidic^7,8 approaches. Emulsion-based methods generate double emulsions through homogenization or ultrasonification of two liquid phases. Surfactants are accordingly needed to ensure interfacial stability by reducing interfacial tension between the different liquid layers.⁹ However, surfactants may affect chemical processes, such as enzymatic activities in the droplet.¹⁰ This makes them unsuitable for certain enzyme-related research. Liquid marbles, a liquid droplet coated with hydrophobic powder, are also constrained by their relatively large size.^5,6 On the contrary, smaller microcapsules can be obtained through electro-spraying with a strong electric field.¹¹ These methods, however, require the use of additional equipment for electric field generation.¹² 

Microfluidics has been a viable option for generating microcapsules. However, the challenges for this approach are with the breaking and misalignment of glass capillary devices^7,14 and dual hydrophobic/hydrophilic surface treatment of polydimethylsiloxane (PDMS) devices.¹³ The dual surface treatment is required for the complete wetting of the microchannels to ensure a steady and reliable liquid flow within each microchannel. The applied treatment accordingly depends on the interfacial tension of the bulk liquid flowing through each microchannel. In general, hydrophobic treatment is employed when oil-based liquids are used to wet the microchannel.¹⁸ Hydrophilic treatment is used for the water-based counterpart.¹⁶ Reducing the amount of surface treatments in a microfluidic device significantly simplifies the fabrication process of the devices. Furthermore, the shell material of the existing solutions has been often limited to alginate, a salt of alginic acid and sodium or calcium. Addressing all these bottlenecks, we present here a method for generating surfactant-free, ultraviolet (UV) light curable core–shell microcapsules in a fully hydrophilic PDMS microfluidic device. Using a UV-curable polymer instead of an oil-based liquid, we avoid the need for dual surface treatment. Figures 1(a) and 1(b) show the experimental setup and the device design. The microfluidic device consists of three cross junctions. The first junction generates the core droplet [Fig. 1(c)]. The second junction forms the core–shell droplet [Fig. 1(d)]. The final junction introduces a spacer liquid to prevent accidental coalescence [Fig. 1(e)]. Upon collection, the core–shell droplets are cured under UV light to form solid microcapsules. The microcapsules are subsequently dried and collected in air. For retrieving the core material, the microcapsules are immersed in a water container and stirred to break the shells.

As the core liquid, we used HFE7500 fluorinated oil (Novec, 3M) with a viscosity of μ_CO = 1.31 mPa s, a surface tension of σ_CO = 15.52 mN/m, and a density of ρ_CO = 1.63 g/ml.¹⁹ The shell polymer (μ_SH = 42 mPa s, σ_SH = 32.5 mN/m, and ρ_SH = 1.07 g/ml)²⁰ was prepared by mixing 0.05 g camphorquinone (Merck), 0.06 g ethyl-4(dimethylamino)benzoate (Merck), and 10 g of trimethylolpropane trimethacrylate (TMPTMA, Merck) using a magnetic stirrer at 600 rpm for 30 min. The second and third junctions were supplied with 50% glycerol solution²¹ (Chemsupply, μ_CN = 6 mPa s, σ_CN = 67 mN/m, and ρ_CN = 1.12 g/ml) into the side channels.

We first optimized the flow rates used for the liquids to achieve consistent generation of thin shell droplets. Due to the high viscosity of the polymer, adjusting the shell flow rate resulted in a slow response time of approximately 30 min. Therefore, we fixed the shell flow rate at Q_SH = 150 μl/hr, which allowed us to produce core–shell droplets of around 10 000 μm². This flow rate also enables us to accommodate the subsequent high flow rates of the spacer liquid at the third junction, which was adjusted up to 2000 μl/h. We proceeded to experimentally optimize the formation process by increasing the flow rates of the core liquid (oil) and the suspension liquid (glycerol solution).

Evaluating the frequencies in the inset of Fig. 2(b) revealed that consistent core–shell formation occurs when f₁ ≈ f₂. When f₁ > f₂, irregular splitting of core droplets was observed. This causes the inconsistent formation of core–shell droplets [Fig. 2(c)], as the core droplets were too closely packed after the first junction [insets of Fig. 2(a)]. Further investigation also revealed that the separation distance between cores needs to be greater than the major axis length of the core–shell droplet for consistent formation [Fig. 2(b)].

Next, we observed the influence of varying Q_CN from 400 μl/h to 1200 μl/h. Fixing Q_CR at 40 μl/h led to consistent core size and separation distance as shown in Figs. 3(a) and 3(b). Increasing Q_CN decreased the size of the core–shell droplet as given in Fig. 3(a). This simultaneously increases f₂ according to (4). However, the core droplet generation frequency f₁ only slightly increased from 75 drops/s at 400 μl/h to 83 drops/s at 1200 μl/h [inset of Fig. 3(b)]. We also observed that the major axis length of the droplet is much less than the core separation distance. This proved that the generation frequency played a greater role than the major axis length in the consistent production of core–shell droplets. The microscope images of the outlets at different flow rates are given in Fig. 3(c).

At the third junction, the glycerol flow rate Q_SP has a negligible influence on the droplet size and the generation frequency. This behavior can be explained by the difference between the upstream and downstream pressures at the junctions. The expansion chamber downstream has a lower fluidic resistance than the upstream microchannel. Thus, the spacer liquid only flows downstream, increasing the separation between the core–shell droplets. This effectively prevents accidental coalescence of the core–shell droplets before collection.

We subsequently optimized the oil and glycerol flow rates to achieve thin-shell microcapsules [Figs. 4(a)–4(d)]. The optimized flow rates were Q_CR = 100 μl/h, Q_SH = 150 μl/h, Q_CN = 800 μl/h, and Q_SP = 1600 μl/h. These values lead to a core area of 4700 µm² and a total microcapsule area of 9500 µm². The optimized generation frequencies were approximately f₁ ≈ f₂ ≈ 100 drops/s. The core–shell droplets were guided into a collection chamber [Fig. 4(a)] and exposed to ultraviolet (UV) light at 36 W for 5 min [Fig. 4(b)]. Individual droplets formed solidified microcapsules due to the UV-curing process of the TMPTMA polymer. Subsequently, the collection chamber was dried overnight on a hotplate at 90 °C. The dimensions of the droplets after drying were measured using scanning electron microscopy (SEM) and were consistent with those before drying as observed under the microscope [Fig. 4(c)]. This fact indicated that there was no evaporation or shrinkage during the drying process. The solidified capsules also did not rupture upon heating, proving their robustness for use as storage and delivery in pharmaceutical and beauty products.

Another application for microcapsules would be polymerase chain reaction (PCR).²² Each microcapsule can serve a single microreactor, eliminating the need for well plates. This reduces the need for plastic consumables in laboratories, while lowering experimental costs by using less reagents. As PCR mixtures are predominantly water-based, we hypothesize that replacing the hydrophilic surface treatment with hydrophobic surface treatment using Aquapel¹² would enable the stable production of water-based core–shell microcapsules.

We demonstrate a simplified production method of core–shell microcapsules in a PDMS microfluidic device. Our method provides multiple advantages over the existing state-of-the-art approaches with no surfactant, no additional equipment, and no dual surface treatment. In addition, the biocompatible shell polymer²³ makes our microcapsules suitable for applications, such as polymerase chain reaction (PCR).²² The solid shells hermetically isolate the core liquid, reducing contamination and evaporation. This feature makes the microcapsules reported here a highly practical and extremely attractive option for the storage and delivery of pharmaceutical products. Our current proof-of-concept experiments can produce 100 microcapsules per second at low flow rates, which is predicted to increase with higher flow rates. We are also able to easily obtain the core liquid from the microcapsules by manually shaking the chamber by hand.

The data that support the findings of this study are available within the article.

N.-T.N. acknowledges financial support from the Australian Research Council, Project No. DP180100055. The devices were fabricated in the Queensland Microtechnology Facility, part of the Queensland node at Griffith University of the Australian National Fabrication Facility, a company established under the National Collaborative Research Infrastructure Strategy to provide nanofabrication and microfabrication facilities for Australia’s researchers.