Article pubs.acs.org/Biomac
Microfluidic Generation and Selective Degradation of BiopolymerBased Janus Microbeads Mélanie Marquis,* Denis Renard, and Bernard Cathala INRA, UR1268 Biopolymères Interactions Assemblages, F-44300 Nantes, France ABSTRACT: We describe a microfluidic approach for generating Janus microbeads from biopolymer hydrogels. A flow-focusing device was used to emulsify the coflow of aqueous solutions of one or two different biopolymers in an organic phase to synthesize homo or hetero Janus microbeads. Biopolymer gelation was initiated, in the chip, by diffusion-controlled ionic cross-linking of the biopolymers. Pectin−pectin (homo Janus) and, for the first time, pectin−alginate (hetero Janus) microbeads were produced. The efficiency of separation of the two hemispheres, which reflected mixing and convection phenomena, was investigated by confocal scanning laser microscopy (CSLM) of previously labeled biopolymers. The interface of the hetero Janus structure was clearly defined, whereas that of the homo Janus microbeads was poorly defined. The Janus structure was confirmed by subjecting each microbead hemisphere to specific enzymatic degradation. These new and original microbeads from renewable resources will open up opportunities for studying relationships between combined enzymatic hydrolysis and active compound release.
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engineering, drug delivery, and bionanotechnology.29−32 In general, biopolymer particles with the appropriate properties for releasing or immobilizing the products of interest have been produced by solution-based batch methods33−35 but also by microfluidic techniques that allow precise control of the fluid velocities and droplet volumes. Pectin and alginate are environmentally friendly because they are highly water-soluble, biocompatible, and biodegradable. One feature of these biopolymers is their high content of carboxylic groups that can be ionically cross-linked to achieve the formation of gels.36−38 In the past decade, versatile microfluidic technologies have emerged for the fabrication of biopolymer particles of controlled size, shape, and composition.26,39−41 Zhao’s group23 recently reported the production of Janus microbeads with magnetic anisotropy. They obtained Janus architecture by embedding magnetic beads on one side of symmetric ionically cross-linked alginate beads. Due to the large size of the magnetic beads, almost no diffusion occurred and convection phenomena were limited, resulting in a clearly defined separation between the two hemispheres. In fact, although little mixing occurs between liquids undergoing laminar flow, in the case of Janus particles with two miscible phases, both diffusive intermixing and convective transport in the microchannels need to be considered.4,42 It should be noted that the formation of Janus particles using two chemically distinct biopolymers and ionically cross-linked hydrogels has not yet been achieved and remains a challenge.
INTRODUCTION During the past decade, multicompartment1 and anisotropic particles,2 have received significant attention due to their novel morphologies and diverse potential applications.3 Janus particles have two distinguishable surface areas of equal size, which makes them suitable for applications in switchable display devices,4 interface stabilizers,5 self-motile microparticles,6 and smart nanomaterials, such as biological sensors, nanomotors, antireflection coatings,7 and anisotropic building blocks for complex structures.8 Biphasic particles were first reported by Xerox society in 1970s with black and white plastic hemispheres for use in twisting-ball display.9 The name Janus particles was initially given by Lee and co-workers in 1985 with polymerization of asymmetric poly(styrene)/poly(methyl methacrylate) emulsion10 and many methods of producing these anisotropic particles have been developed over the last two decades.11−13 Janus particles are currently produced by templating methods,14−17 colloidal assembly,18,19 particle lithography techniques,20 glacing-angle deposition,21 nanosphere lithography,22 and capillary fluid flow. Capillary flowbased approach such as microfluidic4,23−25 devices offer a number of advantages over conventional flow control technology because they ensure highly versatile geometry and can be used to produce monodisperse spherical polymeric microparticles with diameters ranging from several tens to several hundreds of micrometers.26,27 In most of the previous works, Janus particles produced by microfluidics were obtained from the polymerization of organic monomers by fast UV illumination.4,28 This strategy is, however, limited to lightsensitive compounds. Hydrogel-based microparticles, in contrast, are hydrophilic polymer networks with a high affinity for water. These microparticles have recently been used in tissue © 2012 American Chemical Society
Received: January 30, 2012 Revised: March 6, 2012 Published: March 8, 2012 1197
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sunflower seed oil with surfactant and acetic acid were supplied to the microchannels using digitally controlled syringe pumps (Harvard Apparatus PHD 2000, France). The biopolymer microbeads generated by the microfluidic flow focusing device were produced by internal gelation.40 The droplets contained pectin and/or alginate and CaCO3 as the cross-linking agent in an inactive form. The continuous phase (sunflower seed oil) contained acetic acid (0.5 wt %), which diffused into the droplets and triggered the release of Ca2+ ions, resulting in cross-linking of the polysaccharide chains and thus the formation of a biopolymer network. The microbeads were collected in a bath of CaCl2 (1 wt %) solution, gently washed in water then centrifuged (2000 rpm, 5 min) to remove the residual oil phase and CaCl2. Enzymatic Hydrolysis. The enzymatic hydrolyses were performed using either polygalacturonase type II (PGII) or alginate lyase (AL) and Bodipy-pectin/FA-alginate microbeads as substrates. Reaction mixtures containing 28 μL of deionized water, 2 μL of microbead solutions, and 10 μL of PGII at 3 wt % or AL at 0.25 wt % concentrations were incubated at 40 °C for 25 min or at room temperature for 10 min, respectively. Enzymatic degradation of the microbeads, which depended on the enzyme used, was qualitatively evaluated using visualization by fluorescence coupled with phase contrast microscopy and by confocal microscopy . Imaging. Phase contrast and fluorescence microscopy images were captured with an Olympus IX51 inverse microscope (Olympus, France) equipped with phase contrast illumination, a standard green filter (Exciter filter (BP) 460−490 nm, Dichroic Mirror (DM) 500, Barrier Filter (BA) 520 nm), a standard red filter (BP 510−550 nm, DM 570 nm, BA 590 nm), and a digital camera (Sony, SCD-SX90). The size distributions of the microbeads were analyzed using the ImageJ freeware v1.35c. Confocal microscopy images were captured using a Nikon Ti-E with C1si scanning laser confocal microscope (Nikon, France) and a Nikon Eclipse Ti inverse microscope (Nikon, France). FA emission fluorescence was recorded between 500 and 530 nm after excitation at 488 nm. Bodipy emission fluorescence was recorded between 570 and 620 nm after excitation at 561 nm.
We describe here a microfluidic device for the generation of monodisperse homo and hetero Janus microbeads using pectin−pectin and pectin−alginate hydrogels. As the polysaccharides used are completely miscible in a wide range of concentrations, the challenge in microfluidic design was to obtain a well-defined interface between the two biopolymer hemispheres in the homo and hetero Janus microbeads. This was successfully achieved in the case of hetero Janus microbeads due to specific interactions between alginate and pectin chains at the interface. In the case of homo Janus microbeads, however, the interface was poorly defined due to mutually repulsive interactions between the pectin chains. Further experiments were carried out to achieve the gel degradations in two independent steps by directing enzymatic hydrolysis according to the biopolymer compositions of the Janus microbeads.
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EXPERIMENTAL SECTION
Materials. Low-methoxyl citrus pectin (Mw = 169250 g/mol, Mw/ Mn = 2.03) was purchased from Cargill France SAS, and had a degree of esterification (DE) of 30% and contained 78.5% galacturonic acid. Alginate (Mw = 151550 g/mol, Mw/Mn = 2.11), of medium viscosity, was obtained from FMC biopolymer (U.S.A.). N-(3-Dimethylaminopropyl)-N′-ethyl-carbodiimide hydrochloride (EDC; Sigma-Aldrich, France) and N-hydroxysuccinimide (NHS; Sigma-Aldrich) were used for covalent coupling of fluoresceinamine (FA; Sigma-Aldrich France; λexc 485, λem 535 nm) and Bodipy TR cadaverine (Invitrogen, France; λexc 588, λem 616 nm) to citrus pectin or alginate through activation of the polysaccharide carboxyl groups, as described by Ogushi et al.43 Sodium FA-alginate, citrus FA-pectin, and citrus Bodipy-pectin were prepared at 2 wt % concentrations and dissolved in deionized water for 2 h. The pH value for both biopolymer solutions was then adjusted to around 7 with NaOH 1 M to obtain low viscosities and to minimize the onset of gelation due to the presence of acid compounds. Freeze-dried calcium carbonate (CaCO3) powder (5 μm diameter particles) was dispersed in deionized water at 1 wt % concentration. Calcium carbonate and biopolymer solutions were then mixed at a 1:1 (v/v) ratio to give final concentrations of 0.5 and 1 wt % for the calcium carbonate and biopolymer solutions, respectively. The oil phase was sunflower seed oil (Fluka) either mixed with Span 80 (Sigma-Aldrich; 1 wt %) or with Span 80 (1 wt %) and acetic acid (0.5 wt %). Polygalacturonase type II (PGII) from Aspergillus niger was purchased from Novozymes (Bagsvaerd, Denmark). Alginate lyase (AL) from Sphingobacterium multivorum was purchased from SigmaAldrich, France. Enzyme solutions of PGII and AL were prepared in deionized water at 3 wt % (w/w) and 0.25 wt % concentrations, respectively. Microfluidic Device. A microfluidic system, comprising a Flow Focusing Device (FFD) and a second inlet for the continuous phase, was prepared using poly(dimethylsiloxane) (PDMS; RTV 615, Elecoproduit, France) and a soft lithography technique.44 SU-8 (CTS, France) positive relief structures were produced on silicon wafers. PDMS polymer (in a mixture of 10:1 base polymer/curing agent) was cast from this mold, and access holes were punched on the PDMS layer. The PDMS layer (in a mixture of 10:1 base polymer/ curing agent) was then placed in contact with a thin PDMS layer (in a mixture of 20:1 base polymer/curing agent) to generate the microchip. The cross-linker diffused as a result of the gradient from PDMS (20:1) to PDMS (10:1). The chip was then oven-treated at 70 °C for 24 h to strengthen the cross-linking. The microchannels were rectangular in shape with a uniform height of 120 μm and respective widths of 75 μm for the biopolymer phases, 150 μm for the oil phase, 100 μm for the restriction, and 200 μm for the central channel as determined by profilometry (see Figure 2). Emulsion of Aqueous Solutions of Biopolymer and Preparation of Microgels. Aqueous solutions of biopolymers with CaCO3, a sunflower seed oil with surfactant (Span 80), and a
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RESULTS AND DISCUSSION Diffusion-Controlled Gelation of Bulk Phases. Prior to the microfluidic experiments, we examined the time-dependent internal gelation of biopolymers, as governed by progressive solubilization of the cross-linking agent in the aqueous phase, due to acidification of the medium by acetic acid which diffused from the organic to the aqueous phase.27 Figure 1 shows photographs of an aqueous solution of alginate or pectin that was mixed with a solution of insoluble CaCO3 then brought in contact for various time intervals with sunflower seed oil containing acetic acid. The decrease in pH of the biopolymer solutions led to calcium bridging and, therefore, to biopolymer gelation. Diffusion of acetic acid from the sunflower seed oil to the aqueous phase triggered the release of Ca2+ ions from CaCO3 and binding to the residues of α-L-guluronic (G) and mannuronic (M) acids of alginate45 and to the D-galacturonic acid of pectin, thereby, causing biopolymer gelation.46 The extent of alginate and pectin gelation depended on the time that the macroscopic oil and aqueous phases were kept in contact. Figure 1 shows that a gelation time of 30 s was too short to produce an alginate gel, while a complete gelation of pectin solution occurred. In both cases, the polysaccharide solutions had completely gelled after a contact time of 2 min. The kinetics of network formation varied according to the two biopolymers,46,47 and was probably dependent on the number and accessibility of the free functional groups (carboxylic groups) in each labeled polysaccharide. Device Optimizations for the Generation of Janus Microbeads. Although the microfluidics process used to obtain 1198
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junction (Re < 1) indicates that nonconvective transport can occur across the two parallel streams. However, symmetric recirculation is induced inside the forming droplets by coflowing of the external streams.4 Furthermore, the contact between droplet and channel walls increases the impact of convection and the occurrence of internal recirculation loops.51 Thus, the internal recirculation loop phenomenon can disappear when the contact with the wall and the internal droplet velocity are reduced. Finally, the droplet diameter needs to be less than 80% of the channel width in order to inhibit the exchange of material caused by internal circulation during coflowing. In this context, the channel containing the two miscible phases of the biopolymers (FA-pectin and Bodipy-pectin or FA-alginate and Bodipy-pectin), the geometry of the flow focusing junction and the central channel were therefore adjusted so as to minimize diffusive intermixing (Figure 2a). The central channel was therefore short and large (depth, 100 μm; width, 200 μm; and length, 22 mm) and without any zones of turbulence, such as a serpentine shape or the pressure variations commonly used to optimize mixing inside the droplets or coalescence.26,42,52,53 The outlet at the end of the PDMS microcircuit was therefore parallel to the central channel and without a swimming pool. Indeed, at the end of the central channel, gelation was incomplete and a swimming pool outlet would have resulted in high pressure variations. Pregelled microbeads would therefore be distorted and finally fuse by coalescence. Another critical zone in the microcircuit was the parallel outlet where diffusive intermixing phenomena were limited by using a polytetrafluoroethylene (PTFE) tube (0.3 mm i.d. × 0.76 mm o.d. and length, 20 cm) directly inserted in a PDMS short exit channel (depth, 100 μm; width, 400 μm; and
Figure 1. Time-dependent gelation of pectin and alginate driven by the diffusion of acetic acid from oil to an aqueous solution of biopolymer containing an inactive form of calcium (CaCO3). (a) Samples of Bodipy-pectin (1) and FA-alginate (2) at 1 wt % mixed with CaCO3 (0.5 wt %; bottom aqueous phase) placed in contact with sunflower seed oil containing 0.5 wt % of acetic acid (top oil phase). Acetic acid freely diffused from the oil phase into the biopolymer solution and triggered the release of calcium ions for polysaccharide gelation. (b) Photographs from left to right show vials with an aqueous solution of pectin and alginate mixed with CaCO3 in contact with oil containing acetic acid for 30 s, 1, and 2 min. To evaluate gelation time, vials were reversed and complete gelation was estimated when only oil phase is fallen.
Janus particles takes advantage of laminar flow,4,23,25,28,39,48−50 it is important to note that diffusive intermixing may occur with miscible fluids in a two-phase stream before the droplets break up. In fact, the low Reynolds number in the channel before the
Figure 2. (a) Schematic representation of the microfluidic device for Janus droplet generation using a micro flow focusing device with inlets for the two biopolymers mixed with an inactive form of the cross-linking agent (CaCO3) emulsified in oil. Droplet gelation was induced by the diffusion of acetic acid from the oil phase to the droplets, where the resulting pH decrease inside the droplets led to calcium bridging and, thus, to biopolymer gelation. (b) Dimensions of the microdevice produced by soft-lithography. Channels were rectangular in shape with a depth of 120 μm and lengths L1, L2, and L3 of 1, 2, and 20 mm, respectively. (c) Bright-field (left) and bright-field adding fluorescence with FA-filter (right) microscopy images of droplet formation and the two hemispheres of the Janus droplets. On the right picture, the two hemispheres were clearly visible due to the grafting of one of the polysaccharides by fluoresceinamine. The flow rates of the biopolymer solutions and oil in each channel were 1 and 18 μL/min, respectively. Scale bar: 100 μm. 1199
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Figure 3. Fluorescence confocal microscopy images and profiles of the fluorescence intensities for (a) FA-pectin/Bodipy-pectin homo Janus and (b) FA-alginate/Bodipy-pectin hetero Janus microbeads. FA and Bodipy excitations were set at 488 and 561 nm, respectively, while emission fluorescence was recorded between 500 and 530 nm (green) and between 570 and 620 nm (red). The flow rates of FA-pectin (or FA-alginate), Bodipy-pectin, and oil in each channel were 1, 1, and 18 μL/min respectively. Scale bars: 100 μm (left) and 50 μm (middle).
intensity profile across the microbead clearly revealed a superimposition of fluorescence labeling near the interface. Fluorescent FA-labeled alginate and Bodipy-labeled pectin microbeads were prepared under the same conditions as for homo Janus microbeads. The alginate/pectin microbeads also displayed two distinct hemispheres but the interface was welldefined, clearly indicating that only limited diffusive intermixing occurred (Figure 3b). The sharper definition of the interface in the hetero Janus particles was confirmed by the analysis of fluorescence intensity. This difference in interface definition between hetero and homo Janus microbeads led us to reflect on the mechanisms occurring during droplet and gel formation. Convective and Diffusive Intermixing Phenomena in Janus Droplets. The occurrence of convective and diffusive intermixing processes inside the droplets in the microchannel have already been described by Nisisako4 and Sarrazin51 and could explain the observed cross-contamination of the two sides of Janus microbeads (Figure 3a). Cross-contamination could take place during droplet formation when the continuous oil phase flows past the growing drop and creates a convective flow inside the droplets.4 In addition, internal recirculation loops were still present within the droplets in the rectangular microchannels even though the droplet size was small compared to the width of the central channel. These convective flows in each hemisphere caused recirculating movements at the biopolymer/biopolymer interface, thus, allowing crosscontamination of the fluids in each hemisphere. Moreover, in both homo and hetero Janus formation, gelation was still incomplete at the end of the microcircuit (residence time between the onset of gelation and the outlet (L3) = 1.6 s, Figure 2b), even though the bulk gelation time was shown to range from 10 to 30 s for pectin and to exceed 1 min in the case of alginate (Figure 1). As previously mentioned, the outlet constitutes a critical zone of turbulence, due to pressure variations caused by the larger diameter of the PTFE tubing, where convection can occur if gelation is incomplete. This phenomenon, coupled with diffusive intermixing55 prior to the termination of gelation, could result in an irregular biopolymer/ biopolymer interface and the occurrence of invaginations.
length, 5 mm). Internal gelation was obtained by using calcium carbonate and acidification.27 Gelation beyond the FFD junction had to be delayed to limit cap-formation at the junction. Thus, a second inlet of the continuous phase was added 2 mm away from the junction to delay diffusion of the acetic acid in the sunflower seed oil (Figure 2a). The extensive work reported in the literature has shown that the size of the resulting Janus droplets can be controlled by adjusting the fluid flow rates and channel geometry.54 The flow rates for FA-pectin or FAalginate, Bodipy-pectin, and oil in each channel were 1, 1, and 18 μL/min, respectively. As shown in Figure 2b, these adaptations of the flow-focusing junction device to Janus microbeads allowed the production of monodisperse droplets. More precisely, the resulting homo and hetero Janus microbeads had an average diameter of 92 μm and coefficients of variance (c.v. = σ/μ × 100, where σ is the standard deviation and μ the average diameter) of 3.41 and 3.28%, respectively (for a number of beads N = 50), which thus highlights the relevance of microfluidics in the production of monodisperse microbeads. Generation of Homo and Hetero Janus Microbeads. Fluorescently labeled (Fluoresceinamine, FA and Bodipy Tr cadaverine, Bodipy) pectins were prepared to visualize the coflowing aqueous stream and the production of fluorescent homo Janus particles. The fluorescence micrographs taken before (Figure 2c) and after gelation (Figure 3a) showed fluorescently labeled hemispheres within the droplets, composed of FA-labeled and Bodipy-labeled pectin. This process was highly reproducible. The microbeads were then collected at the end of the microcircuit in an aqueous solution of CaCl2 to ensure maximal cross-linking and to limit the coalescence of homo Janus microbeads. Initial observations by fluorescence microscopy, with the appropriate filters (results not shown), showed that FA-pectin and Bodipy-pectin were concentrated on opposite sides of the hemispheres. However, the interface was not clearly defined even though convection phenomena were controlled by having a large central channel and rapid gelation, which suggested that some diffusive intermixing occurred (Figure 3a). Analysis of the fluorescence 1200
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Figure 4. Fluorescence coupled to phase contrast microscopy images and confocal scanning laser microscopy images of the enzymatic degradation of fluorescently labeled pectin-alginate hetero Janus microbeads. (a) Enzymatic degradation of Bodipy-pectin hemisphere by polygalacturonase II (PGII); (b) Enzymatic degradation of FA-alginate hemisphere by Alginate Lyase (AL). Mixing conditions were 28 μL deionized water, 2 μL of microbead solution, and 10 μL of enzyme solution. Scale bars: 100 μm for photonic microscopy and 50 μm for confocal microscopy.
In the case of alginate gelation, L-guluronic and Dmannuronic acid repeat units are involved in the ionic gelation process with divalent ions. In addition, pectin and alginate have been previously shown to be able to interact with each other as a result of the formation of hydrogen bonds between the methoxyl groups in pectin and the hydroxyl groups in the guluronic acids of alginate.60 This specific interaction would lead to a decrease in long-range molecular motions at the interface and, therefore, reduce the linear distance of diffusive intermixing, thus, creating a better separation at the interface of hetero Janus microbeads (Figure 3b). This specific interaction would thus produce a sort of “arrested” interface and override the convective and interdiffusive mixing phenomena, which occur during droplet and microbead formation. In summary, the key to successful generation of hetero Janus microbeads from biopolymers with a brief time of gelation would be to use two different biopolymers, which specifically interact together, or to use totally immiscible biopolymers. To our knowledge, these original biopolymer-based microbeads have never before been reported in the literature and the next step in our strategy was to selectively degrade the two microbead hemispheres using specific enzymes according to the chemical structure of the two biopolymers. Enzymatic Hydrolysis of Hetero Janus Microbeads. The feasibility of selective degradation was demonstrated and the Janus architecture confirmed by investigating the effect of specific enzymes against each of the polysaccharides present in the hetero Janus microbeads. A polygalacturonase (PGII) from Aspergillus niger was used to hydrolyze the backbone, especially the 1−4 linkages between adjacent α-D-GalA residues present in pectin.61 Alginate degradation was achieved by using an alginate lyase (AL) from Sphingobacterium multivorum, also known as alginase or alginate depolymerase. This enzyme catalyzes the hydrolysis of alginate by a β-elimination mechanism targeting the glycosidic 1→4 O-linkages between monomers.58 The enzymatic hydrolyses, using FA and Bodipy filters, clearly revealed the degradation of the desired hemisphere in the microbeads (Figure 4). Indeed, FA-alginate/Bodipy-pectin microbeads mixed with PGII showed a single fluorescent hemisphere, implying total degradation of the pectin, after 25 min of incubation (Figure 4a). On the other hand, microbeads mixed with alginate lyase displayed a single fluorescent hemisphere, which suggested the total degradation of alginate, after
The extent to which diffusive intermixing occurred prior to the completion of gelation was therefore assessed by calculating the average linear diffusion distance of the polysaccharide chains, particularly at the interface. In the first step, the droplets spent about 0.16 s in an unsolidified state (see Figure 2b) between the first dropletforming cross-junction and the onset of gelation (L2). However, as the diffusion coefficients of alginate and pectin in saltfree solution are 780 and 800 μm2·s−1, respectively,56,57 the average linear distance covered in 0.16 s was of the same order of magnitude, that is, 16 μm. This gave rise to considerable diffusive intermixing when the droplet size was ∼90 μm. In the second step, that is, gelation (L3), the viscosities of the biopolymer solutions increased from outside to inside the microbeads. This led to an increase in capillary number and resulted in node movement at the oil/biopolymer interface.51 Diffusive intermixing was slowed down as a consequence of this increased viscosity but still occurred in the centers of the microbeads. How then, and despite these convection and diffusion phenomena, can the more sharply defined interface in hetero Janus microbeads be satisfactorily explained? The most likely reason probably stems from the chemical structure of each polysaccharide and the interactions taking place during the gelation process. Pectins are polysaccharides from terrestrial plant cell walls, in which the backbone is composed of α1→4-linked D-galacturonic (GalA) acid units, known as galacturonans.38 Alginate is a linear polysaccharide derived from brown algae and consists of D-mannuronic acid (M) and L-guluronic acid (G) linked in β1→4.58 In the case of pectin gelation, only the carboxyl groups from the galacturonic acid repeat units are involved in binding with calcium ions, while the methanol-esterified carboxyl groups create steric repulsions between adjacent pectin chains. This intricate balance between long-range attractions and short-range repulsions would produce a less well-defined, irregularly shaped interface in homo Janus microbeads due to repulsive forces between the methoxyl groups, not involved in the ionic gelation process, on both sides of the hemispheres.59 These conformational constraints, coupled with the convective and interdiffusive mixing phenomena, would generate invaginations at the homo Janus interface, as revealed by the fluorescent confocal images (Figure 3a). 1201
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10 min of incubation (Figure 4b). These results were confirmed by the confocal images where, in the case of PGII degradation, the red fluorescence arising from labeled pectins had almost completely disappeared after 25 min of incubation at 40 °C (Figure 4a). The persistence of red fluorescence at the end of the experiment could be due to a progressive decrease of PGII activity due to the release of calcium ions from the pectin network. The presence of calcium salts (except for dibasic calcium phosphate and calcium tartrate) is known to inhibit PGII activity.62 In the case of AL degradation, the green fluorescence arising from labeled alginates had completely disappeared after 10 min of incubation at room temperature (Figure 4b). Moreover, the presence of either PGII or AL did not affect the stability of the nondegraded hemispheres in the hetero Janus microbeads over time. These original results demonstrate the increased flexibility of microbeads derived from polysaccharides and open up possibilities in controlled release, particularly the time-controlled release of two active compounds embedded in each of the microbead hemispheres in response to specific enzymatic hydrolyses. However, the design and control of such novel active microbeads will require a better understanding of enzyme diffusion and, hence, the degradation of the biopolymer networks, within the microbeads, to modulate the release of active substances.
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CONCLUSION
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AUTHOR INFORMATION
REFERENCES
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This study demonstrated the use of microfluidics to generate Janus microbeads from polysaccharides. The design of the microdevice, coupled with optimization of the chemical route of biopolymer gelation, allowed the production of homo and hetero Janus microbeads from pectin−pectin and pectin− alginate with a well-defined interface, particularly in the case of hetero Janus microbeads, thanks to specific interactions between the two polysaccharides. This study opens the door to the generation of hetero Janus microbeads from distinct biopolymers with a short gelation time but that can specifically interact together. New Janus microbeads could also be produced by using protein and polysaccharide and finely controlling the gelation mechanism by new microfluidics routes. We also demonstrated the selective degradation of each hemisphere of the biopolymer-based microbeads by enzymatic hydrolysis. This process will therefore provide new opportunities for the release of active substances in a controlled environment and could find applications in food, medicine, and cosmetics. Future investigations will focus on the release of compounds embedded in biopolymer networks and their control by fine-tuning of the network structure and enzymatic hydrolysis conditions.
Corresponding Author
*E-mail:
[email protected]. Tel.: 33 2 40 67 51 07. Fax: 33 2 40 67 50 43. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS
We would like to thank the platform BIBS and, in particular, B. Bouchet for use of the confocal scanning laser microscopy facility and are grateful to E. Bonnin for her expertise in experimental enzymatic hydrolysis. 1202
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