Designing Food Structure Using Microfluidics | SpringerLink

Food Eng Rev (2015) 7:393–416 DOI 10.1007/s12393-014-9100-0

REVIEW ARTICLE

Designing Food Structure Using Microfluidics F. Y. Ushikubo • D. R. B. Oliveira M. Michelon • R. L. Cunha

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Received: 3 June 2014 / Accepted: 3 November 2014 / Published online: 19 December 2014 Ó Springer Science+Business Media New York 2014

Abstract Microfluidics is an emerging technology that can be employed as a powerful tool for designing structures in the food industry. Those structures can modify the texture and flavor perception in a food product or can be used as carrying vehicles for a wide range of bioactive compounds. Microfluidic processes occur in strictly laminar flow, which is quite interesting to process control. This results in particles with more homogeneous size and shape, which improve the properties of controlled release of active compounds and their stability. Innumerable complex structures can be designed, with single or multiple shells, one or more cores inside the particles. In addition, self-assembled structures can be formed in microfluidic devices. However, most of the published works have fundamental character. Fundamental studies are important to understand the physical phenomena that occur at the microscale, in order to obtain successful results in the design of microstructures. Especially in the case of food-grade ingredients, more complex variables have to be considered, such as their rheological and mass- and heattransport properties. In this review, the fundamentals of microfluidics will be presented, such as the main forces and dimensionless numbers that are involved in the microscale processes and droplet formation regimes. In addition, the most common microfluidic devices will be described, covering their geometries and materials. The review will also present results from studies on structure design using microfluidics, including emulsion-based techniques for the production of microparticles and strategies for the production of self-assembled structures.

F. Y. Ushikubo D. R. B. Oliveira M. Michelon R. L. Cunha (&) Faculty of Food Engineering, University of Campinas, Campinas, SP 13.083-862, Brazil e-mail: [email protected]

Keywords Microfluidics Food structure Encapsulation Emulsion Microparticle Self-assembled structure

Introduction Microfluidics is defined as the science of designing, manufacturing and operating processes and devices with small amounts of fluids in laminar regime. Microfluidic devices have dimensions ranging from a few millimeters to micrometers, and they are characterized by exhibiting at least one channel with dimension smaller than 1 mm [160]. The first applications of microfluidics started in the beginning of the 1990s such as in blood rheology [64] and chemical detection [83]. A decade later, the applications of microfluidics expanded to a wide range of fields: chemistry, biology [120, 140], medical [155], agriculture and food sciences [75, 98, 123] among others. The use of microfluidic techniques allows the integration of procedures into planar chips or other small devices, reducing reaction volumes and the associated cost of chemical and biological experimentations by several orders of magnitude, while simultaneously increasing performance of the process [32, 91, 160]. The greatest advantage of the application of microfluidic processes is the perspective of process intensification, which aims to develop sustainable technologies with less financial demand and increased production in terms of amplification process, reducing costs and time for the implementation of the laboratory to the industrial process [23, 153]. Several types of food structures may be obtained using microfluidics devices: self-assembled structures [93], emulsions [99, 100, 119, 170] and emulsion-based structures such as solid lipid microparticles [65, 129] and

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biopolymeric microbeads [38, 172]. The development of methods to producing food structure has great importance for the development of the food products, because they can be used to modify the texture and flavor perception, to mask undesirable taste/smell and to avoid oxidation of compounds [111]. In addition, they can be used as encapsulation systems for oral administration with controlled release of the bioactive and functional compounds, protecting against the action of enzymes and adverse conditions of gastrointestinal tract such as ionic strength and acid pH [82]. Production techniques of food structures with microfluidic approach are able to overcome the hurdles inherent of conventional methods, such as control of polydispersity and particle size, and are still reported as more reproducible processes with highly controllable operational conditions [34]. Moreover, particles or other food structures obtained by microfluidics often dispense a subsequent processing step for size reduction and homogenization such as membrane extrusion, high-pressure homogenization or microfluidization [22, 79]. Furthermore, the use of microfluidic techniques is related to much faster reactions, minimum size of devices, formation and control of a defined interface between two phases due to strictly laminar flow, low consumption and power dissipation and, finally, relative low cost of production per device [19, 84, 123]. However, most of the published works in food processing area still have a fundamental and exploratory character. Moreover, a small number of studies are related to the influence of interfacial phenomena and rheological behavior of fluids on the generation and stability of the structures [73, 102, 110, 139, 145]. One of the challenges for implementation of microfluidic processes in the food industry is the adjustment of the process conditions considering the fluid properties. Complex systems such as food suspensions exhibit complex rheological properties that may lead to the channels clogging, difficulties in the fluid injection, flow stabilization or fluid flow breakup, depending on process conditions employed. The development of low-cost microfluidic devices that can operate at high flow rates resulting in higher yields, as well as the use of food-grade ingredients, remains as the major technology challenges. In this paper, the aim was to discuss the fundamentals of microfluidics, such as the main forces and dimensionless numbers that are involved in the microscale processes. In addition, the most common microfluidic devices found in the literature were described, covering their geometries and materials. Recent literature was reviewed considering the production of several types of food structures by microfluidic routes, including single and multiple, emulsionbased techniques for the production of particles and self-

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assembled structures. At last, a brief review on the scaling up of microfluidic systems was presented.

Microfluidics: Fundamentals Dimensionless Numbers Processes involving hydrodynamics can be described by a large number of variables and properties that reveal the dominating acting forces. The relation between these acting forces is expressed by the dimensionless numbers, which locates the system in the fluidic parameter space [125]. However, in microscale, the number of relevant parameters is reduced, since the viscosity and interfacial tension are the prevailing effects [15]. In microscale, the more relevant dimensionless numbers are Reynolds, Weber, Capillary, Pećlet and the flow rate ratio. While Weber and Capillary numbers characterize multiphase systems, Pećlet number is calculated for systems in which mass transfer occurs. The Reynolds number (Re) expresses the ratio of inertial to viscous forces in fluid flow: Re ¼ qUl=g, in which q is the fluid density, U is the characteristic velocity, l is the characteristic dimension of the channel and g is the fluid viscosity. The flow regime in microfluidic devices is mostly laminar (Re \ 2,100), considering that their characteristic dimension is very small so that inertial effects are normally irrelevant [16]. The Weber number (We) indicates the relative importance of inertial effects when compared to the interfacial tension in a multiphase system: We ¼ U 2 ql=c;, where U is the characteristic velocity, q represents the fluid density, l is the characteristic dimension of the channel and c is the interfacial tension. In droplet generation, We is usually calculated regarding the dispersed phase to evaluate the detachment forces. The ratio of viscosity to interfacial tension is given by the capillary number, Ca ¼ gU=c, where g is the fluid viscosity, U is the characteristic velocity and c the interfacial tension. Ca is often calculated in multiphase systems for the phase with the higher viscosity [15, 16]. The Pećlet number (Pe) expresses the relative importance between diffusion and convection: Pe ¼ Ul=D, where U is the characteristic velocity of the flow, l is the characteristic length of the channel and D is the diffusion coefficient of the particle/molecule of interest. This dimensionless number is highly relevant for mixing of miscible fluids [13]. In systems with mixing fluids, the flow rate ratio is usually represented by the abbreviation FRR, and it is calculated by the ratio of the aqueous to the organic phase flow rates (Qa and Qo, respectively), which gives FRR = Qa/Qo [14, 57, 95]. In systems with droplet

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generation, the flow rate ratio can be expressed as the relation between the dispersed to the continuous phase flow rates (Qd/Qc) [10, 33, 145] or as the ratio between the continuous to the dispersed phase flow rates (Qc/Qd) [11, 102]. Since flow rate is the major operating condition at microfluidic, this dimensionless number acquires a great importance at microscale. Almost all works involving droplets, bubbles, micro-/nanoparticles and self-assembled structures relate this dimensionless number as the main factor in obtaining structures with the desired sizes and shapes [10, 46, 57, 145]. Further information about dimensionless numbers in microfluidics can be found in Addae-Mensah et al. [7], Atencia and Beebe [13], Ganãń Calvo and Gordillo [39], Garstecki et al. [40], Phapal and Sunthar [113] and Squires and Quake [125]. Microdevice Geometries Different geometries have been suggested to generate a wide range of structures. For the droplets generation, there are shear-induced geometries, such as those based on capillaries assemblies and planar geometries, and interfacial tension-induced one, which are the terrace geometry and its variations. In the case of systems that involve mass transfer, such as for the fabrication of self-assembled structures, mostly planar cross-junction geometry has been used. In shear-induced geometries, droplets can be formed at different regimes: squeezing, dripping or jetting. Squeezing and dripping regimes occur at the tip of the tube orifice or junction. The squeezing regime takes place when the interfacial tension prevails over the shear forces, so the droplet grows, until it almost blocks the cross section of the channel. Then, the continuous phase is confined to a thin film between the walls of the channel and the droplet, increasing the pressure upstream of the droplet, resulting in the detachment of the droplet neck [41]. In the dripping regime, the droplet detaches when there is still room for the continuous phase flow [121]. The jetting regime consists in the formation of a long stream of the inner fluid where the point of droplet detachment can vary, leading to a broader size distribution. In this regime, the droplets generation is explained based on the principle of Rayleigh–Plateau instability, where perturbations in the jet result in an increase in Laplace pressure within the jet. When this pressure is high, the jet becomes thin enough to break the stream into droplets [147]. In dripping and jetting regimes, droplets detachment occurs through a balance between interfacial and viscous forces. In the next sections, the details of each geometry will be presented, describing their mechanism, advantages and disadvantages.

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Capillary Devices The capillary microfluidic devices are assemblies of coaxial capillary tubes, where one is placed inside another with a larger diameter (Fig. 1a, b). These devices are widely used for droplets, vesicles and microparticles generation. For single emulsions, co-flow and flow-focusing arrangements are commonly used. In the former, fluids move in the same direction, so that the dispersed phase is driven by the interior of the small diameter capillary while the continuous phase flows between the capillaries (Fig. 1a). In the latter, fluids are introduced from the two ends of the external capillary in opposite directions (Fig. 1b). Both phases flow into the narrow inner capillary, and the continuous phases constrict the dispersed phase, which breaks into droplets in a dripping or jetting regimes, depending on the operating conditions [121]. One advantage of the coaxial microcappilaries is that the droplets are surrounded completely by the outer phase, which increases the effects of interfacial forces between the two phases. On the other hand, the process with coaxial microcappilaries is difficult to scale up because of the intensive hand labor in the fabrication and alignment of the tubes [34]. Planar Microfluidic Devices The planar microfluidic devices are channels with rectangular cross section produced with different techniques and materials, which are chosen depending on the application. These devices differ from each other according to the type of junction between the channels. T-, Y- and cross-junctions are the most common planar devices used for droplets generation [5, 27, 41, 42, 117, 126, 127, 145, 159]. In the case of the production of self-assembling structures, the most utilized geometry is the cross-junction [14, 51, 54–58, 95, 115]. In T-junction devices (Fig. 2a), the fluids are inserted in two perpendicular microchannels. These devices are lar-

Fig. 1 Schematic illustrations of a a co-flow and b a flow-focusing microcapillary devices for making droplets (reprinted with permission from Utada et al. [147]. Copyright 2007, Cambridge University Press)

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Fig. 2 Design of planar microfluidic devices. a T-junction; b Y-junction; c cross-junction

Fig. 3 Schematic mechanism of the droplet formation in a terrace-based microchannel device (reprinted with permission from Sugiura et al. [129]. Copyright 2000, Elsevier)

gely used because of the facility in the generation of monodisperse droplets and bubbles in a wide range of flow rates [133, 145]. Even at low Ca, in which the droplet detachment is less favored, droplets are detached in the squeezing regime [33, 41, 165]. In this geometry, the droplet generation occurs in two steps: The first is related to the droplet growth and the second is the droplet detachment [126]. In Y-junction (Fig. 2b), the angle between the channels is usually higher than 90°, which will influence the droplets generation, due to the effect of shear imposed by the continuous phase on the dispersed one. Differently to T-junction, the droplet detachment mechanism in Y-junction is described by one single step [127, 128]. The simpler mechanism results in a higher dependence of shear and interfacial forces, as well as of the operation conditions in the droplets size generated in Y-junction. However, since this geometry depends directly on the balance between shear forces and interfacial tension, no droplets can be formed at low Ca [145]. Cross-junction microfluidic devices (Fig. 2c) present four crossed perpendicular or oblique channels, where usually the flow of the lateral channels constricts the main channel fluid flow, in a technique known as hydrodynamic flow focusing [123]. This arrangement is the most used for self-assembled structures formation, such as liposomes,

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due to a great control of molecules diffusion between two fluids [54]. This type of geometry is also used for droplets formation. Compared with T- or Y-junctions, in crossjunction, a higher shear is imposed by the continuous phase due the flow at both sides of the dispersed phase stream. Thus, this configuration may lead the production of smaller droplets. However, depending on operating conditions, jetting regime with the generation of polydispersed droplets can occur, since the mechanism of breakup is directly controlled by the flow rate ratio of the fluids [102]. The planar geometries have the advantage of being easily designed and produced, giving the possibility to create more elaborated configurations. On the other hand, unlike the capillary devices, in planar microchannels, the fluids are limited by microdevice walls at the upper and down surfaces of the chip, which decreases the interfacial area between the two phases. Terrace Geometry The terrace-based geometry consists in droplets generation through the forced passage of the dispersed phase in a microchannel to a terrace area filled with a continuous phase, which can be stagnant (Fig. 3). The shallow depth of the terrace leads to a disk-shaped droplet with a high Laplace pressure, which is thermodynamically unstable.

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When the droplet comes to the terrace edge, it moves spontaneously to a deeper region (well) and assumes a thermodynamically more favorable spherical shape. Diversified designs based on terrace microchannels were presented to improve and increase the throughput of the droplet generation, like those with symmetric straightthrough microchannels [68] showing circular or rectangular cross sections, or asymmetric channels [69, 70] with circular and rectangular sections in the upstream and downstream, respectively. The latter configuration allows the droplets generation with good performance independently of the properties of the phases [69]. In this type of geometry, only the interfacial force drives the droplets formation, unlike capillary and planar microfluidic devices. This mechanism results in a very smooth droplet detachment and consequently in high energetic efficiency [130]. Besides, the arrangement in arrays allows increasing the droplets throughput [70]. On the other hand, the fabrication costs of the device are higher than mostly planar geometries and capillaries. Materials and Methods of Manufacturing Microfluidic Devices The first microfluidic devices were produced using expensive and time-consuming techniques such as photolithography and etching in silicon and glass derived from microelectronics systems [137]. These techniques enable the creation of devices with dimensions as low as few micrometers [61–63, 68]. Besides, with different combination of techniques, more elaborated structures are obtained, such as microgrooves, straight-through microchannels and micronozzles [156]. Silicon microfluidic devices can be obtained by isotropic or anisotropic wet etching or by dry etching, which can be physical, chemical or physical–chemical [80, 101]. The main advantages of wet etching are the high selectivity, repeatability controllable etch rate and relatively planar etching surface. Physical dry etching can be applied to almost all materials, but presents slow etch rates and low selectivity. Chemical dry etching is more similar to wet etching with higher selectivity. Reactive ion etching (RIE) [59], a physical–chemical dry etching, is one of the most popular dry etching techniques, as it can result in high aspect ratios of channels [101]. Etching can be classified as a direct technique. This category also includes laser ablation, micromachining, photolithography in deep resists and stereolithography [18, 67]. Laser ablation involves the material removal by absorption of short-duration laser pulses in the UV or IR regions that break bonds within the long-chain polymer molecules. This technique allows the fabrication of

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microdevices with a sort of polymers such as polystyrene (PS), cellulose acetate (CA), poly(ethylene terephthalate) (PET), polycarbonate (PC) and polymethylmethacrylate (PMMA) [76, 118]. This technique is also applied to glass, ceramic, silicon and metals. Structures can have with dimensions as small as 6 lm [112]. Mechanical micromachining methods, such as sawing, cutting, milling and turning, allow the fabrication of structures of a wide range of materials with dimensions of few tens of micrometers [18]. A special application is for machining stainless steel. This material has the advantage of being more resistant to alkalis and acids and higher mechanical strength than silicon, but cannot be etched because of its multicrystal property [143]. Both laser ablation and micromachining methods are fast techniques, but their drawback is that the surface roughness obtained is greater than that obtained in replication techniques [156]. Photolithography in deep resists is the application of a negative photoresist such as SU-8 by spin coating on the substrate, the exposition of the coated substrate under UV light, followed by the baking and development steps. The region exposed to the UV light become rigid and insoluble, while the non-exposed areas are removed in the development step. The use of SU-8 enables the fabrication of structures with high aspect ratios and fine control, with feature size as low as 1 lm [67]. In contrast, among the disadvantages of SU-8, there are the difficulty of processing this material, its high internal stress and, after the exposition to UV light, it becomes hard to remove from structures [18]. The stereolithography technique consists in focusing a directed UV beam in a photocuring liquid polymer, which is cured and forms two-dimensional solid structures. By stacking the layers, a three-dimensional structure is fabricated [18, 101]. The advantage of this technique is that the device can be fabricated with all inlet and outlet ports incorporated in one structure, so that no bonding or connectors are needed [156]. However, the buildup time is very slow, in the range of several hours [18]. On the other hand, replication techniques use a single master structure, fabricated with a direct technique, to replicate polymer structures many times [18]. The replication technologies were adapted to the microscale to produce polymer microdevices at lower cost. Among the replication techniques, there are injection molding [90], hot embossing [37] and soft lithography [91]. In injection molding, the molten polymer is injected at high pressure into the cavity of the mold. In hot embossing, the mold and the flat polymer are heated up above the glass transition temperature and they are brought together at a controlled force [18, 67, 156]. While hot embossing is a simpler technique with lower setting up costs, injection molding is more feasible for mass production with shorter

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cycle time [101]. The most common polymers used in both techniques are PMMA, PC, polystyrene (PS) [7, 18] and cyclic olefin copolymer (COC) [60, 108]. Soft lithography is one of the most common techniques to fabricate planar microfluidic devices [91]. In this technique, a master structure is produced by photolithography. An elastomer precursor with curing agent is poured into the mold and cured, so the structure can be peeled off. The most popular elastomer polymer is poly(dimethylsiloxane) (PDMS), which offers optically transparency, non-toxicity and flexibility [92]. Despite these advantages, PDMS is a hydrophobic polymer that absorbs nonpolar organic solvents [136] and hinders the production of stable O/W emulsions [17]. Besides, its deformability leads to the reduced control over the hydrodynamic field [22, 57]. To circumvent the former problem, some techniques were proposed to make the PDMS surface more hydrophilic: oxygen plasma treatment [134], UV/UV–ozone treatments [35], glass coating by sol–gel [1], layer-by-layer deposition [17] and covalent surface modification via graft photopolymerization [52].

Applications Emulsions Emulsions play an important role in food industry, in the form of oil-in-water (O/W) emulsions, such as mayonnaise and dressings, or as water-in-oil (W/O) emulsions, such as butter and margarine. They can be used as a part of more complex foods, such as in yogurts and processed cheeses, or as a base to create new structures such as ice creams [30]. Moreover, emulsions can also be employed in functional applications, such as vehicles for bioactive compounds [89]. Conventional methods of emulsification include highpressure homogenizers, colloid mills, high-speed mixers and ultrasonic homogenizers [88]. Those methods do not allow a precise control in the particle size distribution, resulting in highly polydisperse emulsions. Furthermore, in order to break up the droplets, high shear stress is applied in the system, which can cause proteins denaturation, degradation or loss of activity in thermo- or shear-sensible compounds. As an alternative technology, microfluidic devices can be applied to the production of individual droplets. As a result, highly monodisperse emulsions are obtained through a smoother process [121, 130]. The size of the droplets formed in the microfluidic devices is in the range of tens of micrometers, when obtained in terrace-based geometry devices [131] and in a wider range from tens to hundreds of micrometers in shear-based geometries, such as microcapillaries [78, 116] and planar geometries [103, 148].

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Fig. 4 Basic concept for preparing double emulsions (W/O/W) using T-shaped microchannels (reprinted with permission from Okushima et al. [109]. Copyright 2004, American Chemical Society)

Several studies have focused in emulsions produced by microdevices using food-grade ingredients, such O/W emulsions stabilized by proteins [119, 170], or containing edible oils with bioactive components [99, 100]. Furthermore, due to the high manipulating control provided by the microfluidic devices at laminar regime, more elaborated structures can be created. One example is the double or multiple emulsions, such as water-in-oil-in-water (W/O/W) and oil-in-water-in-oil (O/W/O), which are interesting structures for encapsulating hydrophilic and/or hydrophobic compounds. Double emulsions can be produced with two successive T-junctions (Fig. 4). The desired number of internal droplets can be introduced in the larger drop with the adjustment of the internal and external droplets breakup rates [109]. Moreover, the production of a double emulsion with two different compositions in the internal phase can be performed by the use of a cross-junction geometry in the first step [107]. Multiple emulsions can be also formed in sequential cross-junctions along a microchannel [2]. With alternate wettability between the junctions, double, triple and up to indefinitely order emulsions can be produced (Fig. 5). By setting the jetting regime in the initial junctions and dripping instability in the final junction, the jet is broken into monosized drops at a constant rate [3]. This method is specially useful to break up viscoelastic fluids [4], which normally do not detach droplets easily. The viscoelastic fluid is injected as the inner phase, and the intermediate phase is a fluid that can be easily emulsified. With the instability, the intermediate phase breaks up and so does the inner phase. This method is quite interesting for the food systems, because many of food-grade ingredients show viscoelastic properties. A method to produce multiple emulsions in a single step consists in the use of two capillaries of circular cross section disposed within a rectangular capillary [146] (Fig. 6a). In this configuration, the internal phase is

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Fig. 5 Drop maker arrays used to produce multiple emulsions with controlled order. Photomicrographs of a single, b double, c triple, d quadruple and e quintuple emulsion drop maker arrays. The

multiple emulsions produced by the arrays are shown in the right. The scale bars denote 100 lm (reprinted with permission from Abate and Weitz [2]. Copyright 2009, Wiley–VCH)

pumped through the interior of one capillary, while the intermediate phase flows through the external capillary in the same direction. The external phase flows externally the second capillary, in the opposite direction to the other fluids. When the three fluids reach the collection tube, a double emulsion is formed. By placing sequential capillaries, indefinite ordered multiple emulsions can be produced [28] (Fig. 6b). The number of internal droplets and the middlephase thickness are controlled by the ratio between the flow rates of the phases. By adding one more microcapillary parallel to the inner phase capillary, two different inner phases can be injected into the larger droplet [6].

Microparticles

Fig. 6 a Microcapillary geometry for generating double emulsions from coaxial jets. Schematic of the coaxial microcapillary fluidic device (from Utada et al. [146]. Reprinted with permission from

AAAS). b Schematic diagram of the extended capillary microfluidic device for generating triple emulsions (reprinted with permission from Chu et al. [28]. Copyright 2007, WILEY–VCH)

Microparticles of several to hundreds of micrometers can be generated based on the solidification of droplets of single or multiple emulsions. They can be used as a vehicle to transport a compound to a specific target. The solidification of the droplets increases the system stability and protects the compound from external agents (oxygen, light, reacting environment). Moreover, the release of the compound in the target site can be achieved by the exposition of the microparticle to a specific condition (temperature, pH, presence of enzymes). By the production of

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microparticles through microfluidics, the narrow particle size distribution of droplets results in particles with more homogeneous mechanical properties [36]. In the next sections, the main solidification methods to obtaining of foodgrade microparticles and some types of microparticles and other flow-induced structures will be presented. Solidification Methods The crystallization of the oil phase is an interesting method for the particle solidification in applications in which the compound release occurs by the increase of the temperature. The lipid in the liquid state is injected into the microfluidic device to generate the droplets and then is crystallized by cooling to form solid particles [65, 129, 162] (Fig. 7). The bioactive compounds are released at the melting temperature of the lipid. This temperature can be adjusted by selecting the composition of lipids with different saturation degrees. Another method is the fabrication of microgels, which consists in generating biopolymer droplets in the microfluidic device and subsequently inducing its gelation, inside or outside the device. This is a very convenient method for the encapsulation of food compounds, since several types of food-grade ingredients, such as polysaccharides and proteins, can be employed as the carrier material. Although the polymerization of droplets into microbeads is extensively studied in synthetic polymers systems, we focused only in biopolymers due to their higher compatibility with food products. A list of publications related to microgels fabrication is presented in Table 1. One of the most applied gelling methods is the ionic crosslinking of charged polysaccharides. Examples of biopolymers used in the ionic crosslinking method include sodium alginate [26, 48, 77, 78, 116, 122, 132, 135, 163, 168, 172] and low methoxyl pectin [38], which form gels in the presence of a divalent ion such as Ca2? by binding to guluronic and galacturonic acids, respectively. Other examples reported in the literature are the ionic cross-linking of chitosan with Cu2? [167] and carboxymethylcellulose with Fe3? [171]. Fig. 7 Microfluidic production of solid fat microparticles (reprinted with permission from Kim et al. [66]. Copyright 2013, American Chemical Society)

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Although less frequently applied in food systems, biopolymers can also gel through covalent cross-linking, such as the gelation of gelatin with genipin [50] and peroxidase-catalyzed gelation in chemically modified gelatin [24]. Several methods to gel the biopolymer by ionic or covalent cross-linking in microfluidic devices were proposed: (1) coalescence of droplets, (2) in situ mixing, (3) internal and (4) external gelation (Table 1). Each gelling method is described in the following. The gelation through the droplets coalescence is based on the collision of one droplet containing the biopolymer solution with a second droplet containing the cross-linking ion [77, 122, 132] (Fig. 8a). In this method, no surfactant is used in the continuous oil phase, since the coalescence of the droplets is desirable. The flow rate of the cross-linking agent is set higher than that of the biopolymer, in order to avoid the coalescence of biopolymer droplets. One of the disadvantages of this method is the difficulty in producing well-defined and homogeneous microbeads if gelation occurs before the stabilization of the droplet after the coalescence [135], resulting in higher particle size polydispersity, with coefficient of variation (CV = standard deviation/average size*100) up to 14 % [132], while other gelation methods usually result in CV lower than 5 %. Furthermore, this method is dependent on the probability of the droplets collision, which makes it difficult to obtain 100 % successful merging droplets. In the ‘‘in situ mixing’’ method, microfluidic devices with more than two inlet channels that encounter at the junction allow the injection of the biopolymer and the cross-linking agent solutions separately into the chip. The two streams merge and form a droplet, which is rapidly detached at the channel junction by the shear stress of the oil continuous phase flow (Fig. 8b). A winding channel after the droplet generation can improve the reagents mixing [26, 38]. The drawback of this method is the gelation of the biopolymer at the junction of the device. Water or a solution with the encapsulating material can be

Ionic cross-linking (internal gelation)

Ionic cross-linking (in situ mixing)

(1) 1.5 % sodium alginate in buffered solution

Ionic cross-linking (coalescence)

2 % w/w lecithin in corn oil with acetic acid (0.67 and 2.68 lL/ml oil) 0.5 % v/v acetic acid in sunflower oil

(i) 8 % w/v PGPR in Soybean oil (outer phase)

2 % w/w sodium alginate in RPMI solution with CaCO3 nanoparticles (1.14 and 2.27 mg/mL)

2 % w/v sodium alginate with 0.5 % w/v CaCO3

2 % w/w sodium alginate, 1.5 g/L CaCO3, 1 % w/w Pluronic and 30 mmol/L PAG (middle phase) (ii) 1:1 v/v soybean oil ? benzyl benzoate (inner phase)

3 % w/w Span 80 and 5 % w/w acetic acid in soybean oil

2 % Span 80 in Mineral oil

0.2–2 % w/w Span 80 in hexadecane

Edible oil

Liu et al. [78]

Qinner = 300 lL/h, Qmiddle = 400 lL/h, Qouter: 4,000 lL/h

Injection tube tip: 60 lm

Microcapillary

Hollow capsules: 383 lm (inner diameter), 467 lm (outer)

Workman et al. [163]

80–400 lm (not informed)

Inlet channel crosssectional area: 500 lm2

Stainless steel manifold with HPLC connectors in crossjunction

PDMS, T-junction

Tan and Takeuchi [135]

(not informed)

PDMS, oblique flow focusing

94–139 lm

Zhang et al. [172]

Fang and Cathala [38]

Qoil: 200–600 lL/h, Qalginate?CaCO3: 20 lL/h

40–100 lm

Choi et al. [26]

60–90 lm

Dispersed and continuous phases channels: 50 lm and 150 lm width, channel height: 48 lm

Qoil = 1.5–2 mL/h, Qbiopolymer = QCaCl2 = 0.03–0.05 mL/ h, Qwater: 0.07–0.08 mL/h

100 lm width, 78 lm height

Shintaku et al. [122]

Liu et al. [77]

Sugiura et al. [132]

Reference

104–167 lm

Spheres: 21.7 lm diameter, Disks: 24.1 lm height, 41.3 lm diameter

77.4–216 lm

Particle average size

60–110 lm

Qoil = 1–5 lL/min (from graph), Qalginate ? QCaCl2 = 0.6–2 lL/min (droplet formation)

Qoil: 6–96 lL/min, Qalginate: 2 lL/min, QCaCl2 : 6 lL/min (spherical beads)

50 lm height, junction: 50 lm width, collecting channel: 200 lm width 90 lm width and height

Qoil-in-alginate: 100 lL/h, Qalginate: 2 lL/h, QCaCl2 : 30 lL/h, Qoil-in-CaCl2 : 50 lL/h (spherical beads)

Qoil: 200 and 1,000 mL/s, Qalginate: 5 mL/ h, QCaCl2 : 100 mL/h

30 lm inner width, 60 lm outer width and 15 lm nozzle height 50–200 lm width, 40 lm height

Operating conditions

Channel dimensions

Qoil: 10.5–16.6 mL/h, Qalginate?CaCO3: 0.7–0.8 mL/h

PDMS, 5 inlets oblique flow-focusing

PDMS, 3 inlets T-junction


PDMS, two crossjunctions (droplets formation), one T-junction (fusion channel)

Silicon micronozzle array

Soybean oil

Soybean oil

Device material and geometry

Oil phase

2 % w/w sodium alginate with 0.1 %w/w CaCO3

(3) 1 % w/w CaCl2

(2) water

(1) 5 % w/w pectin or 4 % w/w alginate

(2) 20 mM CaCl2

(1) 0.5–2 % w/w sodium alginate

(2) 0.1 M CaCl2

(1) 1.5 % w/w sodium alginate

(2) 2 % w/w CaCl2

(1) 2 % w/w sodium alginate

(2) 0.1 M CaCl2 in 0.14 M NaCl solution

Aqueous phase

Gelling method

Table 1 Studies related to biopolymer microgels produced with microfluidic devices

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123

Chemical cross-linking (external gelation)

(1) 3 % w/v sodium alginate

Ionic cross-linking (external gelation)

PMMA, cross-junction

Sunflower seed oil

(2) 5 % w/w Span and 1.5 % w/w CaCl2 in n-decanol (collecting reservoir)

(2) 15 % w/w barium or calcium acetate in glycerol/water mixture (collecting reservoir)

(2) 10 % w/v genipin (collecting reservoir)

(1) 1 % w/v gelatin and 2 % w/v genipin

Sunflower seed oil

(1) 5 % w/w Span 80 in n-decanol

(1) 1.5 % w/w sodium alginate

(2) 10 % w/v CaCl2 (injected in collecting tube)

8 % w/v PGPR in soybean oil (inner and outer phase)

PMMA, cross-junction,

Glass Microcapillary


Qoil: 0.4–1.4 mL/min, Qalginate: 0.01–0.07 mL/min

Qoil: 1,800 lL/h, Qalginate: 30 lL/h

Injection tube tip: 50 lm

Collecting channel: 600 lm

Qinner: 60–800 lL/h, Qmiddle: 400–4,000 lL/h, Qouter and QCaCl2 : 2,000–14,000 lL/h

Injection tube tip: 20–40 lm

Qoil: 0.5–1.2 mL/min, Qalginate: 0.001–0.05 mL/min

200 lm width, 1.5 mm height

Span 80 in sunflower seed oil

PMMA, T-junction

Qoil: 0.1 = 1.0 mL/min, Qchitosan: 0.001–0.1 mL/min

200 lm

PMMA, cross-junction

Sunflower seed oil

(1) 2 % w/v sodium alginate (middle phase)

(2) 20 % w/v CaCl2 (collecting reservoir)

(1) 1.5 % w/v sodium alginate

(2) 20 % CuSO4 solution (injected after droplet formation)

(1) 3 % w/w chitosan

Qoil: 15.6–17.2 mL/h, Qalginate: 0.6–0.7 mL/

Not informed


2 % w/w calcium acetate and 3 % w/w Span 80 in soybean oil

2 % w/w sodium alginate

Qoil: 0.2–15 mL/h, Qalginate: 0.3–0.7 mL/h

At junction: 130 lm width, 110 lm height


0.5 % w/w CaI2 in undecanol with Span 80

2 % w/w sodium alginate

Qoil: 0.25 mL/h, QCMC: 0.03 mL/h

0.25 % w/w Fe(NO3)3 in undecanol with Span 80

1 % w/w Carboxymethylcellulose


0.25 % w/w CaI2 in undecanol with Span 80 PDMS, oblique flow focusing

Qoil: 0.8 mL/min, Qalginate: 0.08 mL/min


Qoil: 0.35 mL/h, Qcarrageenan: 0.03 mL/h

Collecting channel: 600 lm

Channel dimensions




Oil phase

0.8 % w/w j-carrageenan

(2) 20 % CaCl2 (collecting reservoir)

Aqueous phase

Gelling method

Table 1 continued

130–580 lm

Varied shapes (size not informed)

Core and shell: 117–173 lm (inner diameter); 250–255 lm (outer)

70–220 lm

Huang et al. [50]

Hu et al. [48]

Ren et al. [116]

Yeh et al. [168]

Yang et al. [167]

Zhang et al. [172]

50–70 lm

100–800 lm

Zhang et al. [171]

Zhang et al. [171]

30–230 lm

Not informed

Zhang et al. [171]

Huang et al. [49]

50–2,000 lm

Not informed

Reference


402 Food Eng Rev (2015) 7:393–416

1.5 % w/w Span 80 in mineral oil 0.5–1 % w/w triblock copolymer surfactant in fluorinated oil

2–5 % w/w agarose

4 % w/w chitosan in 2 % w/w acetic acid aqueous solution

5 mM hydrogel peroxide Cylindrical Teflon capillary and microneedle inserted in a cross-shaped PMMA plate

Teflon capillary: 500 lm, microneedle: 160 lm (injection)

130 lm height, 80 lm width

PDMS, oblique flowfocusing device

Qoil: 5–2,000 lL/min, Qchitosan: 3–100 lL/min

Temulsification = 37 °C, Tcool = 4 °C

Qagarose ? Qgelatin ? Qperoxide = 0.6 mL/ h, Qperoxide = 0.2 mL/h, Qoil = 0.6–24 mL/h

Temulsification = 37 °C, Tcool = 2 °C

Qoil: 0.05–04 mL/h, Qagarose: 0.025–0.2 mL/h (first junction)

60 lm height, 60–100 lm width

PDMS, sequential T-junctions

100–700 lm

105–175 lm

95–100 lm (supplementary material)

Disks: 50–250 lm diameter

Temulsification = 37 °C

30 and 60 lm heigh

PDMS or polyurethane, flow-focusing device

Elongated particle: 300–440 lm.

31.6 lm

Temulsification = 40 °C, Tcollect = 25 °C, Tcool = 5 °C

Channel: 16 lm width, 11 lm height Qbiopolymer: 6.25 and 250 lL/min, Qoil: 1–100 mL/min, Temulsification = 100 °C, Toil = 54 °C, Tcool = room



Channel dimensions

inner capillary: 0.1 mm inner diameter, outer capillary: 1 mm inner diameter.


Silicon microchannel plate (terrace geometry)


Xu et al. [166]

Chau et al. [24]

Wang et al. [158]

Xu et al. [164]

Walther et al. [157]

Iwamoto et al. [53]

Reference

TGCR tetraglycerine-condensed ricinoleic acid ester, PAG diphenyliodonium nitrate, PGPR polyglycerol polyricinoleate, HPLC high-performance liquid chromatography, DMSO dimethyl sulfoxide

Solvent extraction

1 % w/v chemically modified gelatin 2 % w/w Span 80 in 30 % w/w triotylamine in octyl alcohol

3 % w/w Span 80 in hexadecane

Agarose solution

2 % w/v agarose in DMSO

Sunflower seed oil

1.75 % w/w j-carrageenan

Temperature ? chemical cross-linking

5 % w/w TGCR in iso-octane

5 % w/w gelatin pH 7.4

Temperature

Oil phase

Aqueous phase

Gelling method

Table 1 continued

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Fig. 8 Schematic of the microfluidic production of alginate microgels by a coalescence of biopolymer and cross-linking agent droplets (reprinted with permission from Shintaku et al. [122]. Copyright 2007, Springer), b in situ mixture (reprinted with permission from Choi et al. [26]. Copyright 2007, Springer), c internal gelation (reprinted with permission from Zhang et al. [172]. Copyright 2007, WILEY–VCH), d external gelation adding the crosslinking agent

downstream (reprinted with permission from Ren et al. [116]. Copyright 2010, Elsevier), e external gelation adding the crosslinking agent off-chip (reprinted with permission from Huang et al. [50]. Copyright 2009, from Elsevier) and f external gelation adding the crosslinking agent in the continuous phase (reprinted with permission from Zhang et al. [172]. Copyright 2007, WILEY–VCH)

injected between the biopolymer and the cross-linking agent solutions to avoid their contact at the junction [38]. The internal gelation of the microgels consists in the generation of biopolymer droplets containing the crosslinking agent that initially is in an inactive form. A reaction initiator present in the continuous phase diffuses through the droplets and activates the cross-linking agent, initiating the gelling reaction (Fig. 8c). An extensively studied system is composed of alginate and CaCO3 in the dispersed phase and oil with acetic acid in the continuous phase [135, 163, 172]. With the droplet formation, the acid diffuses into the droplet, decreases its pH and releases the Ca2? ion, according to the reaction: CaCO3 ? 2H? ? CaHCO3? ? H? ? Ca2? ? H2O ? CO2 :. Thus, the ion Ca2? can bind to the guluronic acids of alginate and the microgels are formed.

A drawback of this method is that it results in weak gels with low stability [172]. In addition, channel clogging can occur due to the rapid acidification and gelation of the alginate at the channel junction. This problem can be overcome by the injection of an additional flow of pure oil near the junction to avoid the gelation before the droplet formation [163]. A different strategy that avoids channel clogging is to add a photoacid generator with the biopolymer and the CaCO3 nanoparticles dispersion. After the droplets generation, UV radiation is applied to release the Ca2? ions from CaCO3 and react with the alginate [78]. The ionic cross-linking can also be carried out through external gelation. In this method, the cross-linking agent is added after the droplet generation. There are different strategies to make the cross-linking agent reach the biopolymer. A crosslinking agent aqueous solution can be added downstream after

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the droplet formation, so the biopolymers react with the crosslinking ion at the interface of the oil and aqueous stream (Fig. 8d) [116, 167]. Another strategy is the off-chip ionic cross-linking, that is, the biopolymer droplets are formed in the oil phase and are collected in a cross-linking agent solution in the collecting reservoir (Fig. 8e) [49, 50, 168]. The droplets tend to sink to the aqueous phase because of their higher density. In a different approach, the cross-linking agent is solubilized in the continuous phase, so after the droplets are formed, the agent diffuses into the droplet, resulting in the gelation reaction (Fig. 8f) [172]. In this method, the cross-linking agent has to be partially soluble in both dispersed and continuous phases. A suitable crosslinking agent for this method is calcium acetate, which diffuses efficiently in alginate solution, resulting in stable microgels with narrow size distribution [172]. Moreover, biopolymer gelation induced by temperature can be used to generate microbeads without a cross-linking agent. The biopolymer solution is injected in the microdevice at a temperature higher than its gelation point. After the droplets formation, they are cooled below that point. This method is applicable, for example, for gelatin [53], since it presents random coil configuration at 35–40 °C, and below this temperature range, collagen-like triple helices and hydrogen bond cross-links are formed [71, 87], resulting in thermoreversible microgels. Some polysaccharides, such as j-carrageenan [157] and agarose [24, 158, 164], can also form gel by cooling, helix formation and stabilization through hydrogen bonds [12, 87]. One drawback of this technique is the temperature control along microfluidic device, which is critical due to its small dimensions and the large temperature gradient [144]. Since the temperature affects the viscosity of the solutions, this is a very important variable to achieve the repeatability of the process. In the case of pH-sensitive biopolymers, such as chitosan, their gelation could be achieved by the change in pH [166]. In this method, an acidic solution containing the

biopolymer is dispersed in a continuous organic phase containing an extractant. With the acid diffusion to the organic phase, the biopolymer gels are formed.

Fig. 9 a Core–shell particles with the shell consisting of a resolidified lipid and the core formed by an aqueous solution. Electron microscopy image of a cracked lipid shell. The scale bars denote 100 lm (reprinted with permission from Windbergs et al. [162]. Copyright 2013, American Chemical Society). b CLSM images of monodisperse core–shell Ca-alginate microcapsules loaded with

BSA-FITC molecules (reprinted from Liu et al. [78]. Copyright 2013, Elsevier). c Fluorescence confocal microscopy images for FAalginate/Bodipy-pectin hetero Janus microbeads. Scale bar 100 lm (reprinted with permission from Marquis et al. [85]. Copyright 2013 American Chemical Society)

Types of Microparticles and Other Flow-Induced Structures In this section, some configurations of microparticles that are reported in the literature are described. Although some of them use non-food-grade ingredients, they show the possibilities to be explored in the future of the food industry. The simplest structures, microbeads are formed by the production of a single emulsion, in which the dispersed phase is solidified. As discussed in the solidification methods, solid fat microparticles can be obtained through the oil crystallization in O/W emulsions [65, 129], while biopolymer microbeads are formed based on W/O emulsion [26, 38, 132, 135, 172]. Moreover, microcapsules can be obtained by the formation of a double emulsion and subsequently solidification of the middle phase, resulting in the formation of a shell [146]. Microcapsules may be a hollow shell, which is formed based on an air-in-oil-in-water (A/O/W) bubble– emulsion system [72] or a shell–core carrier, which is a capsule (shell) filled in with a solid or liquid (core), based on the solidification of the intermediate phase of a W/O/W or O/W/O double emulsion (Fig. 9a, b) [78, 116, 162]. Shell–core microcapsules are interesting structures to load both hydrophobic and hydrophilic drugs. Janus particles, which are composed by two hemispheres with different compositions, could be an interesting application in the controlled release of more than one bioactive compound at different targets by specific enzymatic hydrolysis (Fig. 9c) [85]. The formation of such structures is based on the injection of two inner phases in separately channels into the continuous phase. The solutions that compose the inner phase should be miscible in

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each other, so they form a single drop. As soon as they merge, the droplet is detached and solidified. Since the droplet is formed at laminar regime inside the microdevice, the mixing rate of the solutions is not high. Therefore, a heterogeneous structure is obtained. In addition, there are structures other than particles that can be produced by microfluidic methods. Microfibers, which can be exploited for modifying food texture and, at the same time, for compounds encapsulation, can be obtained through the confinement of a biopolymer solution flow by an outer phase in a planar cross-junction or microcapillary geometry. The outer phase contains the gelling agent, resulting in the gelling of the biopolymer in the form of a continuous microfiber [29, 169]. The diameter of the fibers is controlled by the ratio of the inner to outer phases flow rates. Furthermore, foams are formed in microfluidic devices by the injection of air bubbles in a solution [8, 124]. Foams are interesting structures that modify the texture and rheology of the products, resulting in a change in their appearance and mouthfeel [21]. Different sensorial characteristics can be obtained by controlling the microbubble size through the solution flow rate, air pressure and solution viscosity.

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of self-assembled structures, either as adaptation of traditional methods or totally innovative techniques [154]. The small dimensions of microchannel enable rapid and uniform mass transfer that can dramatically improve self-assembly yield and size distribution. In addition, the possibilities of solvent recycling and integration of separation techniques are ideal to reduce cost effective of the production of self-assembled structures [22, 23]. Particles formed using microfluidics by self-assembly include among others conventional liposomes, niosomes and polymersomes. Liposomes

Self-assembled structures consist in the arrangement in different configurations of surfactant molecules under certain conditions of concentration and system polarity. Microfluidic technologies have recently been developed for the production

Liposomes can be used as nanocarrier systems for the protection and delivery of bioactive compounds [96, 97] or modifying the texture of food components, developing new tastes and sensations [25]. Liposomes are formed by polar lipids, usually phospholipids. Phospholipids have the capacity of self-organization in bilayers or lamellae in aqueous media, exposing their hydrophilic heads, hiding their hydrophobic tails and forming flat structures. These structures close on themselves forming vesicles of spherical shell type to achieve thermodynamic stability [81, 115]. The production of liposomes by the microfluidic approach is based on the hydrodynamic flow-focusing method in microfluidic devices with cross-shaped geometry. According to Fig. 10, the formation of liposomes by this method is achieved by introducing an organic dispersion containing phospholipids through a central channel

Fig. 10 Schematic diagram of the hypothesized mechanism for liposome formation in a planar microfluidic hydrodynamic focusing device. The central stream is composed of the phospholipid dispersion

in ethanol, which is hydrodynamically compressed by the two adjacent aqueous streams (adapted with permission from Balbino et al. [14]. Copyright 2013, Elsevier)

Self-assembled Structures

123

Liposomes

Structure

Phosphatebuffered saline Phosphatebuffered saline

Isopropanol

Isopropanol

Isopropanol

Isopropanol

PFP

Ethanol

Isopropanol

Ethanol Ethanol

Ethanol

DMPC and cholesterol (5 mM)

DPPC (6.25 mM)

DLPC, DMPC, DPPC, DSPC and cholesterol (5 mM)

DMPE and cholesterol (5 mM)

DSPC and DSPE-PEG5000 (1.92 mM)

Asolectin from soybean, oleic acid and cholesterol (*6.41 mM)

POPC and DMPC (2–20 mM)

EPC, DOPE and DOTAP (8–92 mM)

DMPC, DMPE-PEG5000, DMPEPEG2000, DSPE-PEG2000 and cholesterol (20 mM)

DMPC, DPPC, DOPC and HSPC (*15.6 mM)

Ultrapure water

Aqueous buffer

Deionized water

Phosphatebuffered saline

Glycerol and deionized water





Ethanol

EPC and cholesterol (3.84 mM)


Isopropanol

Solvent

Aqueous phase

DMPC and cholesterol (5 mM)

Compounds

Organic phase

Glass

Cyclic olefin

PDMS and glass

Glass

PDMS

PDMS and glass

PDMS and glass

Silicon and borosilicate glass




Glass


Device material Oblique (*angle 45°)

42–64 lm

Perpendicular

10 lm

Circular section (500 lm)

Axial flow focusing

Oblique (*angle 45°)

190 lm 270 lm

Perpendicular

140 lm 100 lm

50 lm

220 lm

Perpendicular


30–50 lm 40–60 lm

200 lm


Perpendicular

Perpendicular

12–45 lm

50 lm

200 lm

*260 lm

65 lm

39 lm

21 lm

36 lm Perpendicular

Oblique (angle 45°)

42–65 lm 120 lm


Circular section (*1 mm)

100 lm

Intersection angle

Width/depth

Characteristics of device

Table 2 Experimental data compiled of the self-assembly structures production in a microfluidic hydrodynamic focusing devices

10–100

40–100

8–18

4–40

Not informed

Not informed

2–7

9–49

10–25

12–48

6–36

8–12

10–60

FRR range

Hood et al. [47] Phapal and Sunthar [113]

*50–*250 nm

Balbino et al. [14]

Mijajlovic et al. [95]

Davies et al. [31]

Martz et al. [86]

Wi et al. [161]

Zook and Vreeland [173]

Hong et al. [45]

Jahn et al. [57]

Huang et al. [51]

Jahn et al. [55]

Reference

*80–110 nm

85.8–508 nm

*50–*160 nm

*500 nm

360 nm–11 lm

*100– *500 nm

*62 nm

82–130 nm

60–142 nm

54–156 nm

66.27–189.9 nm

50–150 nm

Average diameter

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123

123 Phosphatebuffered saline THF

PMPC-b-PDPA (5 g/L)

PB-b-PEO (4 g/L)

Water


Water


Aqueous phase

Stainless steel

Glassy carbon

PDMS and glass

PDMS

PDMS and glass


Device material

45 lm

400 lm

Not informed

200 lm Not informed

Not informed


100 lm Not informed

Perpendicular

30–70 lm 70 lm

56 lm

400 lm Perpendicular


65 lm 120 lm

Intersection angle

Width/depth

Characteristics of device

1

1

1–6

0.25–4

15–50

15–50

FRR range

25–49 nm

29–46 nm

75–275 nm

40 nm–2 lm

272–*560 nm

54–84 nm

Average diameter

Thiermann et al. [142]

Brown et al. [20]

Thiele et al. [141]

Lo et al. [79]

Reference

HSPC hydrogenated soy phosphatidylcholine, EPC natural egg phosphatidylcholine, PS L-a-phosphatidylserine, DOPC 1,2-dioleoyl-sn-glycero-3-phosphocholine, P2VP-b-PO poly[2vinylpyridine]-b-poly[ethylene oxide], PMPC-b-PDPA poly[2-(methacryloyloxyethyl phosphorylcholine)]-b-poly[2-(diisopropylaminoethyl methacrylate], PB-b-PEO poly[butadiene]-bpoly[ethylene oxide], POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, DMPC 1,2-dimyristoyl-sn-glycero-3-phosphocholine, DMPE 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine, DOPE 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine, DLPC 1,2-dilauroyl-sn-glycero-3-phosphocholine, DSPC 1,2-distearoyl-sn-glycero-3-phosphocholine, DPPC 1,2-dipalmitoylsn-glycero-3-phosphocholine, DPPE 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine, DOTAP 1,2-dioleoyl-3-trimethylammonium-propane, DSPE-PEG5000 1,2-distearoyl-sn-glycero-3phosphoethanolamine-N [methoxy-(polyethylene glycol)-5000], DMPE-PEG5000 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N [methoxy-(polyethylene glycol)-5000], DMPEPEG2000 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N [methoxy-(polyethylene glycol)-2000], DSPE-PEG2000 1,2-distearoyl-sn-glycero-3-phosphoethanolamine- N [methoxy(polyethylene glycol)-2000], Span 20 sorbitan monolaurate, Span 60 sorbitan monostearate, Span 80 sorbitan monooleate, PFP perfluoropentane, THF tetrahydrofuran

Ethanol

P2VP-b-PO (0.5–1 g/L)

Polymersomes

Isopropanol

Solvent

Span 20, Span 60 and Span 80 and cholesterol (5 mM)

Compounds

Organic phase

Niosomes

Structure

Table 2 continued

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and subsequent compression of this dispersion by two adjacent aqueous streams. The hydrodynamic flow focusing of the lipid dispersion causes the controlled diffusion of organic solvent into the aqueous phase. As a result, water molecules will replace the organic molecules around the phospholipids causing a change in the solubility of the system. This fact triggers the self-organization of phospholipids in bilayer fragments and subsequently in spherical unilamellar vesicles [14, 51, 54–58, 95, 115]. Experimental data compiled from the literature of selfassembled structures production in a microfluidic hydrodynamic flow-focusing devices are given in Table 2. Many of the studies mention only the composition of the system and the FRR range used in the experiments, but do not present further details evaluating the effect of the geometry, the dimensions and configurations of the channels (width and depth of the rectangular cross section and angle of intersection of the side channels relative to the central one) and the manufacturing materials on the average hydrodynamic diameter and polydispersity index of the self-assembled structures. Several studies show that the size of the liposomes obtained by hydrodynamic flow-focusing is practically constant when the flow in the microchannel is between 30 and 200 lL/min, if the FRR is kept constant. A reduction in FRR will cause an increase in the width of organic stream, providing a central zone with a high concentration of

organic solvent. A higher concentration of the solvent can stabilize the phospholipids bilayer fragments and allows greater aggregation, resulting in large liposomes with a wide particle size distribution. On the other hand, increasing the value of FRR, the organic solvent will be more diluted because the organic stream is hydrodynamically focused into a thin jet. The reduced concentration of organic solvent limits the formation of phospholipids bilayer fragments, resulting in smaller liposomes with more homogeneous size distribution [55–57]. Although there is little discussion about the influence of dimensions and configurations of the planar microfluidic devices on liposomes production, some studies express the results as a function of these parameters. Jahn et al. [57] studied the influence of the dimensions of the microfluidic hydrodynamic focusing device using microchannel with different widths, depths and intersection angles between the lateral and central channels. Balbino et al. [14] studied the continuous production of liposomes employing microfluidic devices with one or two parallel hydrodynamic focusing. The results of both studies showed that in general the dimensions and geometry of the devices do not significantly affect the size and polydispersity of liposomes. However, since the devices with large dimensions are easier to fabricate and operate, achieving higher flow rates, they are generally associated with higher-throughput performance compared with smaller devices [22].

Fig. 11 Scale-up of production with a 16-channel module. a Schematic of the microchannels on a chip. Labels ‘‘b’’ and ‘‘w’’ specify the inlet positions for black and white isobornyl acrylate, respectively. The aqueous phase is infused from the inner 16 inlets, arranged circularly, b schematic of internal structure of the holder

with a glass chip (side view), c top view of the formation of biphasic droplets in the module, d magnified view of the co-flow geometries and e magnified view of the outlet port in the center of the chip (reprinted with permission from Nisisako et al. [106]. Copyright 2006, WILEY–VCH)

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Niosomes

Polymersomes

Niosomes are synthetic membrane vesicles produced by self-assembly of nonionic surfactants that do not dissociate at neutral pH. Their structure, properties and production procedure are similar to those of conventional liposomes. Niosomes can encapsulate hydrophilic molecules in their inner aqueous core and partition hydrophobic ones into their bilayer membrane [22, 79]. Microfluidic methods can be used to produce niosomes with precise control of the particles size and polydispersity. Lo et al. [79] used planar microfluidic hydrodynamic flow-focusing devices manufactured with different materials for niosomes production, following the concept proposed by Jahn et al. [54]. The results demonstrated the potential of microfluidics for the production of synthetic membrane vesicles of 54 nm size with low polydispersity through self-assembly of nonionic surfactants (Table 2).

Polymersomes, also referred to as polymeric vesicles, are spherical vesicles formed by the self-assembled amphiphilic block copolymers in aqueous environment. The preparation methods of polymersomes include similar techniques as those used to obtain liposomes and niosomes [74, 141, 142]. The production of polymersomes occurs through two different microfluidic approaches. The first is based on the mixing technique using a planar microfluidic hydrodynamic flow-focusing devices, consisting of perpendicularly crossed microchannels to focus an ethanolic block copolymer solution into a stream of water [20, 141]. Some results obtained by this technique are given in Table 2. The second technique is based on the formation of W/O/W double emulsion using a capillary device. This method relies on the evaporation of the solvent of the double emulsion stabilized by the copolymer. As the

Fig. 12 Common layouts of microfluidic channels for distribution of fluids from a single manifold into multiple parallel drop generation units. In each the device consists of 8 drop generation units (crossjunctions). The abbreviations CP, DP and E denote the inlets of continuous and dispersed phase fluid and the outlet of emulsion product, respectively (adopted from Tetradis-Meris et al. [138]. Reprinted with permission from [156]. Copyright 2013, Elsevier)

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solvent evaporates, the polymer self-assembles into a vesicular structure [9, 22, 43, 66].

Perspective for Application in Food Industries: LargeScale Production The overall market for microfluidics-based products was valued at nearly $5.1 billion in 2011 and $5.6 billion in 2012, and the total in market revenues is expected to reach nearly $10.3 billion in 2017 considering an annual growth rate of 13 % [94]. On the other hand, the current microfluidic technologies have to overcome some limitations to make its applications potentially viable in the food industry. The feasibility challenges include the precise control of heat and mass transfer functions inside the microfluidic channels and consequently their effects on the properties of food structures. Another limiting factor is the requirement of specialized facilities, such as a clean room for microfluidic device fabrication. Consequently, high manufacturing costs of devices remain a hurdle for the microfluidic technology and its application in the food industries [98]. Recently, more versatile and easy microfabrication methods that do not require a clean room facility have been suggested in order to reduce the production costs [114]. Besides, the microfluidic technologies are still limited by its low throughput. For practical use in food industry where an annual throughput of many tons is generally required, the scale-up can be done by the massive parallelization of devices, where several identical microfluidic devices are subjected to the same process [44]. This type of scale-up occurs without the need for larger equipment design, reducing costs and time of implementation study to industrial scale [104]. Parallel microfluidic devices in one-, two-, and threedimensional arrays have so far been coupled with a single set of pumps so as to increase the throughput of emulsions and might be used for others food structures [105]. An approach consists in a planar microfluidic chip droplet generator with microchannels circularly integrated, and a support holder with concentric multiple annular channels that supply each fluid evenly into the channels (Fig. 11). With this strategy, simple O/W emulsion composed by 90.7-lm-mean-diameter droplets was produced with 2.2 % coefficient of variation at a throughput of 180 mL h-1 [104–106]. Two layouts that allow to distribute fluids from a single manifold to parallelized planar microfluidic microchannels are the tree-type and the ladder-type multiple parallel drop generation units (Fig. 12) [138]. The tree-type network has one inlet for each phase at the zeroth branching level and 2m inlet channels for each phase at the mth branching level.

411

Therefore, the number of inlet channels increases by a factor of 2 between two consecutive branching levels. The ladder-type structure has two main feed channels, one for the dispersed phase and one for the continuous phase, and smaller channels branching off them [138, 156]. One hundred and eighty integrated drop generation units with cross-junctions (20 9 20 lm) arranged in 9 parallel lines, each line with 20 cross-junctions, and connected using an the ladder-type architecture, were used to produce W/O emulsions with a droplet diameter of 21 lm with droplet diameter variations of less than 5 % [138]. Based on the terrace geometry, the edge-based droplet generation (EDGE) device (Fig. 13) was designed to form multiple droplets at the edge of a shallow but rather wide rectangular plateau [149–151]. A 1.5 9 1.5 cm chip

Fig. 13 a Design of the EDGE microchip, where the dark channel guides the to-be-dispersed phase, the gray ‘‘twisting road’’ is the continuous phase channel and the rectangular areas in between are the droplet formation units and b drawing of one droplet formation parallelized unit (reprinted with permission from van Dijke et al. [150]. Copyright 2010, Elsevier)

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composed by 196 plateaus (200 9 500 9 1.2 lm) is able to produce up to 300,000 droplets per second per chip [152].

Summary and Outlook In the last decade, several strategies were presented to produce different food structures, taking advantage of the highly controlled hydrodynamics conditions of the microfluidic devices. It has been proposed many devices geometries, materials and fabrication methods. For successful results, the knowledge of the fluid dynamics in the microscale is essential. Thus, many studies have been published aiming to explore the fundamentals of the microfluidics. The operating conditions, mainly the ratio between the flow rates, and the physical properties of the fluids such as the viscosity and interfacial tension are determinant for manipulating the size and shape of the structures. The most promising application of food structure generated by microfluidic methods is the encapsulation of bioactive compounds for controlled release. Single and multiple emulsions, emulsion-template microparticles and self-assembled nanoparticles with low size polydispersity can be generated to load hydrophilic and/or hydrophobic compounds. However, more studies are needed regarding the stability of these structures in the final product as well as in the gastrointestinal environment. For the future of the food industry, some of the challenges regarding the use of microfluidics are the creation of stable and responsive structures using only food-grade ingredients, the production of devices that operate at high throughput resulting in high yield for the production at industrial scale. Acknowledgments The authors would like to thank CNPq (479459/2012-6, 140283/2013-7, 140270/2014-0 and 305477/2012-9) and FAPESP (2011/06.083-0 and 2010/16.708-4) for their financial support.

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Designing Food Structure Using Microfluidics | SpringerLink

Designing Food Structure Using Microfluidics | SpringerLink

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