Microfluidic Generation and Selective Degradation of Biopolymer-Based Janus MicrobeadsMelanie Marquis,* Denis Renard, and Bernard Cathala
INRA, UR1268 Biopolymeres Interactions Assemblages, F-44300 Nantes, France
ABSTRACT: We describe a microfluidic approach for generatingJanus microbeads from biopolymer hydrogels. A flow-focusingdevice was used to emulsify the coflow of aqueous solutions of oneor two different biopolymers in an organic phase to synthesizehomo or hetero Janus microbeads. Biopolymer gelation wasinitiated, in the chip, by diffusion-controlled ionic cross-linking ofthe biopolymers. Pectin−pectin (homo Janus) and, for the firsttime, pectin−alginate (hetero Janus) microbeads were produced.The efficiency of separation of the two hemispheres, which reflectedmixing and convection phenomena, was investigated by confocalscanning laser microscopy (CSLM) of previously labeled biopoly-mers. The interface of the hetero Janus structure was clearly defined, whereas that of the homo Janus microbeads was poorlydefined. The Janus structure was confirmed by subjecting each microbead hemisphere to specific enzymatic degradation. Thesenew and original microbeads from renewable resources will open up opportunities for studying relationships between combinedenzymatic hydrolysis and active compound release.
■ INTRODUCTIONDuring the past decade, multicompartment1 and anisotropicparticles,2 have received significant attention due to their novelmorphologies and diverse potential applications.3 Janusparticles have two distinguishable surface areas of equal size,which makes them suitable for applications in switchable dis-play devices,4 interface stabilizers,5 self-motile microparticles,6
and smart nanomaterials, such as biological sensors, nano-motors, antireflection coatings,7 and anisotropic building blocksfor complex structures.8 Biphasic particles were first reportedby Xerox society in 1970s with black and white plastichemispheres for use in twisting-ball display.9 The name Janusparticles was initially given by Lee and co-workers in 1985 withpolymerization of asymmetric poly(styrene)/poly(methylmethacrylate) emulsion10 and many methods of producingthese anisotropic particles have been developed over the lasttwo decades.11−13 Janus particles are currently produced bytemplating methods,14−17 colloidal assembly,18,19 particlelithography techniques,20 glacing-angle deposition,21 nano-sphere lithography,22 and capillary fluid flow. Capillary flow-based approach such as microfluidic4,23−25 devices offer anumber of advantages over conventional flow controltechnology because they ensure highly versatile geometry andcan be used to produce monodisperse spherical polymericmicroparticles with diameters ranging from several tens toseveral hundreds of micrometers.26,27 In most of the previousworks, Janus particles produced by microfluidics were obtainedfrom the polymerization of organic monomers by fast UVillumination.4,28 This strategy is, however, limited to light-sensitive compounds. Hydrogel-based microparticles, in con-trast, are hydrophilic polymer networks with a high affinity forwater. These microparticles have recently been used in tissue
engineering, drug delivery, and bionanotechnology.29−32 Ingeneral, biopolymer particles with the appropriate propertiesfor releasing or immobilizing the products of interest have beenproduced by solution-based batch methods33−35 but also bymicrofluidic techniques that allow precise control of the fluidvelocities and droplet volumes. Pectin and alginate areenvironmentally friendly because they are highly water-soluble,biocompatible, and biodegradable. One feature of these bio-polymers is their high content of carboxylic groups that can beionically cross-linked to achieve the formation of gels.36−38 Inthe past decade, versatile microfluidic technologies haveemerged for the fabrication of biopolymer particles of con-trolled size, shape, and composition.26,39−41 Zhao’s group23
recently reported the production of Janus microbeads withmagnetic anisotropy. They obtained Janus architecture byembedding magnetic beads on one side of symmetric ionicallycross-linked alginate beads. Due to the large size of themagnetic beads, almost no diffusion occurred and convectionphenomena were limited, resulting in a clearly defined sepa-ration between the two hemispheres. In fact, although littlemixing occurs between liquids undergoing laminar flow, in thecase of Janus particles with two miscible phases, both diffusiveintermixing and convective transport in the microchannels needto be considered.4,42 It should be noted that the formation ofJanus particles using two chemically distinct biopolymers andionically cross-linked hydrogels has not yet been achieved andremains a challenge.
Received: January 30, 2012Revised: March 6, 2012Published: March 8, 2012
We describe here a microfluidic device for the generation ofmonodisperse homo and hetero Janus microbeads usingpectin−pectin and pectin−alginate hydrogels. As the poly-saccharides used are completely miscible in a wide range ofconcentrations, the challenge in microfluidic design was toobtain a well-defined interface between the two biopolymerhemispheres in the homo and hetero Janus microbeads. Thiswas successfully achieved in the case of hetero Janusmicrobeads due to specific interactions between alginate andpectin chains at the interface. In the case of homo Janusmicrobeads, however, the interface was poorly defined due tomutually repulsive interactions between the pectin chains.Further experiments were carried out to achieve the gel degrad-ations in two independent steps by directing enzymatic hydro-lysis according to the biopolymer compositions of the Janusmicrobeads.
Mn = 2.03) was purchased from Cargill France SAS, and had a degreeof 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-Dimethylamino-propyl)-N′-ethyl-carbodiimide hydrochloride (EDC; Sigma-Aldrich,France) and N-hydroxysuccinimide (NHS; Sigma-Aldrich) were usedfor 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 ofthe polysaccharide carboxyl groups, as described by Ogushi et al.43
Sodium FA-alginate, citrus FA-pectin, and citrus Bodipy-pectin wereprepared at 2 wt % concentrations and dissolved in deionized waterfor 2 h. The pH value for both biopolymer solutions was thenadjusted to around 7 with NaOH 1 M to obtain low viscosities and tominimize the onset of gelation due to the presence of acidcompounds. Freeze-dried calcium carbonate (CaCO3) powder (5 μmdiameter particles) was dispersed in deionized water at 1 wt % con-centration. Calcium carbonate and biopolymer solutions were thenmixed 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 Span80 (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 Sigma-Aldrich, France. Enzyme solutions of PGII and AL were prepared indeionized 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 siliconwafers. PDMS polymer (in a mixture of 10:1 base polymer/curingagent) was cast from this mold, and access holes were punched on thePDMS 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 amixture 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 tostrengthen the cross-linking. The microchannels were rectangular inshape with a uniform height of 120 μm and respective widths of 75 μmfor the biopolymer phases, 150 μm for the oil phase, 100 μm for therestriction, and 200 μm for the central channel as determined byprofilometry (see Figure 2).Emulsion of Aqueous Solutions of Biopolymer and
Preparation of Microgels. Aqueous solutions of biopolymers withCaCO3, a sunflower seed oil with surfactant (Span 80), and a
sunflower seed oil with surfactant and acetic acid were supplied to themicrochannels using digitally controlled syringe pumps (HarvardApparatus PHD 2000, France). The biopolymer microbeads generatedby the microfluidic flow focusing device were produced by internalgelation.40 The droplets contained pectin and/or alginate and CaCO3as the cross-linking agent in an inactive form. The continuous phase(sunflower seed oil) contained acetic acid (0.5 wt %), which diffusedinto the droplets and triggered the release of Ca2+ ions, resulting incross-linking of the polysaccharide chains and thus the formation of abiopolymer network. The microbeads were collected in a bath ofCaCl2 (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 performedusing either polygalacturonase type II (PGII) or alginate lyase (AL)and Bodipy-pectin/FA-alginate microbeads as substrates. Reactionmixtures containing 28 μL of deionized water, 2 μL of microbeadsolutions, and 10 μL of PGII at 3 wt % or AL at 0.25 wt % con-centrations were incubated at 40 °C for 25 min or at roomtemperature for 10 min, respectively. Enzymatic degradation of themicrobeads, which depended on the enzyme used, was qualitativelyevaluated using visualization by fluorescence coupled with phasecontrast microscopy and by confocal microscopy .
Imaging. Phase contrast and fluorescence microscopy images werecaptured with an Olympus IX51 inverse microscope (Olympus,France) equipped with phase contrast illumination, a standard greenfilter (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 theImageJ freeware v1.35c.
Confocal microscopy images were captured using a Nikon Ti-E withC1si scanning laser confocal microscope (Nikon, France) and a NikonEclipse Ti inverse microscope (Nikon, France). FA emissionfluorescence was recorded between 500 and 530 nm after excitationat 488 nm. Bodipy emission fluorescence was recorded between 570and 620 nm after excitation at 561 nm.
■ RESULTS AND DISCUSSION
Diffusion-Controlled Gelation of Bulk Phases. Prior tothe microfluidic experiments, we examined the time-dependentinternal gelation of biopolymers, as governed by progressivesolubilization of the cross-linking agent in the aqueous phase,due to acidification of the medium by acetic acid which diffusedfrom the organic to the aqueous phase.27 Figure 1 shows photo-graphs of an aqueous solution of alginate or pectin that wasmixed with a solution of insoluble CaCO3 then brought incontact for various time intervals with sunflower seed oil con-taining acetic acid. The decrease in pH of the biopolymersolutions led to calcium bridging and, therefore, to biopolymergelation. Diffusion of acetic acid from the sunflower seed oil tothe aqueous phase triggered the release of Ca2+ ions fromCaCO3 and binding to the residues of α-L-guluronic (G) andmannuronic (M) acids of alginate45 and to the D-galacturonicacid of pectin, thereby, causing biopolymer gelation.46 Theextent of alginate and pectin gelation depended on the timethat the macroscopic oil and aqueous phases were kept incontact. Figure 1 shows that a gelation time of 30 s was tooshort to produce an alginate gel, while a complete gelation ofpectin solution occurred. In both cases, the polysaccharidesolutions had completely gelled after a contact time of 2 min.The kinetics of network formation varied according to the twobiopolymers,46,47 and was probably dependent on the numberand accessibility of the free functional groups (carboxylicgroups) in each labeled polysaccharide.
Device Optimizations for the Generation of JanusMicrobeads. Although the microfluidics process used to obtain
Janus particles takes advantage of laminar flow,4,23,25,28,39,48−50
it is important to note that diffusive intermixing may occur withmiscible fluids in a two-phase stream before the droplets breakup. In fact, the low Reynolds number in the channel before the
junction (Re < 1) indicates that nonconvective transport canoccur across the two parallel streams. However, symmetricrecirculation is induced inside the forming droplets bycoflowing of the external streams.4 Furthermore, the contactbetween droplet and channel walls increases the impact ofconvection and the occurrence of internal recirculationloops.51 Thus, the internal recirculation loop phenomenoncan disappear when the contact with the wall and the internaldroplet velocity are reduced. Finally, the droplet diameter needsto be less than 80% of the channel width in order to inhibit theexchange of material caused by internal circulation duringcoflowing.In this context, the channel containing the two miscible
phases of the biopolymers (FA-pectin and Bodipy-pectin orFA-alginate and Bodipy-pectin), the geometry of the flowfocusing junction and the central channel were thereforeadjusted 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 anyzones of turbulence, such as a serpentine shape or the pressurevariations commonly used to optimize mixing inside thedroplets or coalescence.26,42,52,53 The outlet at the end of thePDMS microcircuit was therefore parallel to the central channeland without a swimming pool. Indeed, at the end of the centralchannel, gelation was incomplete and a swimming pool outletwould have resulted in high pressure variations. Pregelledmicrobeads would therefore be distorted and finally fuse bycoalescence. Another critical zone in the microcircuit was theparallel outlet where diffusive intermixing phenomena werelimited by using a polytetrafluoroethylene (PTFE) tube (0.3 mmi.d. × 0.76 mm o.d. and length, 20 cm) directly inserted in aPDMS short exit channel (depth, 100 μm; width, 400 μm; and
Figure 2. (a) Schematic representation of the microfluidic device for Janus droplet generation using a micro flow focusing device with inlets for thetwo biopolymers mixed with an inactive form of the cross-linking agent (CaCO3) emulsified in oil. Droplet gelation was induced by the diffusion ofacetic acid from the oil phase to the droplets, where the resulting pH decrease inside the droplets led to calcium bridging and, thus, to biopolymergelation. (b) Dimensions of the microdevice produced by soft-lithography. Channels were rectangular in shape with a depth of 120 μm and lengthsL1, L2, and L3 of 1, 2, and 20 mm, respectively. (c) Bright-field (left) and bright-field adding fluorescence with FA-filter (right) microscopy imagesof droplet formation and the two hemispheres of the Janus droplets. On the right picture, the two hemispheres were clearly visible due to the graftingof 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.
Figure 1. Time-dependent gelation of pectin and alginate driven bythe diffusion of acetic acid from oil to an aqueous solution ofbiopolymer containing an inactive form of calcium (CaCO3). (a)Samples of Bodipy-pectin (1) and FA-alginate (2) at 1 wt % mixedwith CaCO3 (0.5 wt %; bottom aqueous phase) placed in contact withsunflower seed oil containing 0.5 wt % of acetic acid (top oil phase).Acetic acid freely diffused from the oil phase into the biopolymersolution and triggered the release of calcium ions for polysaccharidegelation. (b) Photographs from left to right show vials with an aqueoussolution of pectin and alginate mixed with CaCO3 in contact with oilcontaining acetic acid for 30 s, 1, and 2 min. To evaluate gelation time,vials were reversed and complete gelation was estimated when only oilphase is fallen.
length, 5 mm). Internal gelation was obtained by using calciumcarbonate and acidification.27 Gelation beyond the FFD junc-tion had to be delayed to limit cap-formation at the junction.Thus, a second inlet of the continuous phase was added 2 mmaway from the junction to delay diffusion of the acetic acid inthe sunflower seed oil (Figure 2a). The extensive work reportedin the literature has shown that the size of the resulting Janusdroplets can be controlled by adjusting the fluid flow rates andchannel geometry.54 The flow rates for FA-pectin or FA-alginate, Bodipy-pectin, and oil in each channel were 1, 1, and18 μL/min, respectively. As shown in Figure 2b, theseadaptations of the flow-focusing junction device to Janusmicrobeads allowed the production of monodisperse droplets.More precisely, the resulting homo and hetero Janus microbe-ads had an average diameter of 92 μm and coefficients ofvariance (c.v. = σ/μ × 100, where σ is the standard deviationand μ the average diameter) of 3.41 and 3.28%, respectively(for a number of beads N = 50), which thus highlights therelevance of microfluidics in the production of monodispersemicrobeads.Generation of Homo and Hetero Janus Microbeads.
Fluorescently labeled (Fluoresceinamine, FA and Bodipy Trcadaverine, Bodipy) pectins were prepared to visualize thecoflowing aqueous stream and the production of fluorescenthomo Janus particles. The fluorescence micrographs takenbefore (Figure 2c) and after gelation (Figure 3a) showedfluorescently labeled hemispheres within the droplets,composed of FA-labeled and Bodipy-labeled pectin. Thisprocess was highly reproducible. The microbeads were thencollected at the end of the microcircuit in an aqueous solutionof CaCl2 to ensure maximal cross-linking and to limit thecoalescence of homo Janus microbeads. Initial observations byfluorescence microscopy, with the appropriate filters (resultsnot shown), showed that FA-pectin and Bodipy-pectin wereconcentrated on opposite sides of the hemispheres. However,the interface was not clearly defined even though convectionphenomena were controlled by having a large central channeland rapid gelation, which suggested that some diffusiveintermixing occurred (Figure 3a). Analysis of the fluorescence
intensity profile across the microbead clearly revealed asuperimposition of fluorescence labeling near the interface.Fluorescent FA-labeled alginate and Bodipy-labeled pectin
microbeads were prepared under the same conditions as forhomo Janus microbeads. The alginate/pectin microbeads alsodisplayed two distinct hemispheres but the interface was well-defined, clearly indicating that only limited diffusive intermixingoccurred (Figure 3b). The sharper definition of the interface inthe hetero Janus particles was confirmed by the analysis offluorescence intensity. This difference in interface definitionbetween hetero and homo Janus microbeads led us to reflect onthe mechanisms occurring during droplet and gel formation.
Convective and Diffusive Intermixing Phenomena inJanus Droplets. The occurrence of convective and diffusiveintermixing processes inside the droplets in the microchannelhave already been described by Nisisako4 and Sarrazin51 andcould explain the observed cross-contamination of the twosides of Janus microbeads (Figure 3a). Cross-contaminationcould take place during droplet formation when the continuousoil phase flows past the growing drop and creates a convectiveflow inside the droplets.4 In addition, internal recirculationloops were still present within the droplets in the rectangularmicrochannels even though the droplet size was smallcompared to the width of the central channel. These convectiveflows in each hemisphere caused recirculating movementsat the biopolymer/biopolymer interface, thus, allowing cross-contamination of the fluids in each hemisphere. Moreover, inboth homo and hetero Janus formation, gelation was stillincomplete at the end of the microcircuit (residence timebetween the onset of gelation and the outlet (L3) = 1.6 s,Figure 2b), even though the bulk gelation time was shown torange from 10 to 30 s for pectin and to exceed 1 min in the caseof alginate (Figure 1). As previously mentioned, the outletconstitutes a critical zone of turbulence, due to pressurevariations caused by the larger diameter of the PTFE tubing,where convection can occur if gelation is incomplete. Thisphenomenon, coupled with diffusive intermixing55 prior to thetermination of gelation, could result in an irregular biopolymer/biopolymer interface and the occurrence of invaginations.
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 emissionfluorescence 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).
The extent to which diffusive intermixing occurred prior tothe completion of gelation was therefore assessed by calculatingthe average linear diffusion distance of the polysaccharidechains, 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 droplet-forming cross-junction and the onset of gelation (L2). How-ever, as the diffusion coefficients of alginate and pectin in salt-free solution are 780 and 800 μm2·s−1, respectively,56,57 theaverage linear distance covered in 0.16 s was of the same orderof magnitude, that is, 16 μm. This gave rise to considerablediffusive intermixing when the droplet size was ∼90 μm. Inthe second step, that is, gelation (L3), the viscosities of thebiopolymer solutions increased from outside to inside themicrobeads. This led to an increase in capillary number andresulted in node movement at the oil/biopolymer interface.51
Diffusive intermixing was slowed down as a consequence of thisincreased viscosity but still occurred in the centers of themicrobeads. How then, and despite these convection anddiffusion phenomena, can the more sharply defined interface inhetero Janus microbeads be satisfactorily explained?The most likely reason probably stems from the chemical
structure of each polysaccharide and the interactions takingplace during the gelation process. Pectins are polysaccharidesfrom terrestrial plant cell walls, in which the backbone iscomposed of α1→4-linked D-galacturonic (GalA) acid units,known as galacturonans.38 Alginate is a linear polysaccharidederived from brown algae and consists of D-mannuronic acid(M) and L-guluronic acid (G) linked in β1→4.58 In the case ofpectin gelation, only the carboxyl groups from the galacturonicacid repeat units are involved in binding with calcium ions,while the methanol-esterified carboxyl groups create stericrepulsions between adjacent pectin chains. This intricatebalance between long-range attractions and short-rangerepulsions would produce a less well-defined, irregularly shapedinterface in homo Janus microbeads due to repulsive forcesbetween the methoxyl groups, not involved in the ionic gelationprocess, on both sides of the hemispheres.59 These conforma-tional constraints, coupled with the convective and interdiffu-sive mixing phenomena, would generate invaginations at thehomo Janus interface, as revealed by the fluorescent confocalimages (Figure 3a).
In the case of alginate gelation, L-guluronic and D-mannuronic acid repeat units are involved in the ionic gelationprocess with divalent ions. In addition, pectin and alginate havebeen previously shown to be able to interact with each other asa result of the formation of hydrogen bonds between themethoxyl groups in pectin and the hydroxyl groups in theguluronic acids of alginate.60 This specific interaction wouldlead to a decrease in long-range molecular motions at theinterface and, therefore, reduce the linear distance of diffusiveintermixing, thus, creating a better separation at the interface ofhetero Janus microbeads (Figure 3b). This specific interactionwould thus produce a sort of “arrested” interface and overridethe convective and interdiffusive mixing phenomena, whichoccur during droplet and microbead formation.In summary, the key to successful generation of hetero Janus
microbeads from biopolymers with a brief time of gelationwould be to use two different biopolymers, which specificallyinteract together, or to use totally immiscible biopolymers.To our knowledge, these original biopolymer-based micro-
beads have never before been reported in the literature and thenext step in our strategy was to selectively degrade the twomicrobead hemispheres using specific enzymes according to thechemical structure of the two biopolymers.
Enzymatic Hydrolysis of Hetero Janus Microbeads.The feasibility of selective degradation was demonstrated andthe Janus architecture confirmed by investigating the effect ofspecific enzymes against each of the polysaccharides present inthe hetero Janus microbeads. A polygalacturonase (PGII) fromAspergillus niger was used to hydrolyze the backbone, especiallythe 1−4 linkages between adjacent α-D-GalA residues presentin pectin.61 Alginate degradation was achieved by using analginate lyase (AL) from Sphingobacterium multivorum, alsoknown as alginase or alginate depolymerase. This enzyme cata-lyzes the hydrolysis of alginate by a β-elimination mechanismtargeting the glycosidic 1→4 O-linkages between monomers.58
The enzymatic hydrolyses, using FA and Bodipy filters, clearlyrevealed the degradation of the desired hemisphere in themicrobeads (Figure 4). Indeed, FA-alginate/Bodipy-pectinmicrobeads mixed with PGII showed a single fluorescenthemisphere, implying total degradation of the pectin, after25 min of incubation (Figure 4a). On the other hand, microbeadsmixed with alginate lyase displayed a single fluorescent hemi-sphere, which suggested the total degradation of alginate, after
Figure 4. Fluorescence coupled to phase contrast microscopy images and confocal scanning laser microscopy images of the enzymatic degradation offluorescently 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 ofmicrobead solution, and 10 μL of enzyme solution. Scale bars: 100 μm for photonic microscopy and 50 μm for confocal microscopy.
10 min of incubation (Figure 4b). These results were confirmedby the confocal images where, in the case of PGII degradation,the red fluorescence arising from labeled pectins had almostcompletely disappeared after 25 min of incubation at 40 °C(Figure 4a). The persistence of red fluorescence at the end ofthe experiment could be due to a progressive decrease of PGIIactivity due to the release of calcium ions from the pectinnetwork. The presence of calcium salts (except for dibasiccalcium phosphate and calcium tartrate) is known to inhibitPGII activity.62 In the case of AL degradation, the greenfluorescence arising from labeled alginates had completelydisappeared after 10 min of incubation at room temperature(Figure 4b). Moreover, the presence of either PGII or AL didnot affect the stability of the nondegraded hemispheres in thehetero Janus microbeads over time. These original resultsdemonstrate the increased flexibility of microbeads derivedfrom polysaccharides and open up possibilities in controlledrelease, particularly the time-controlled release of two activecompounds embedded in each of the microbead hemispheresin response to specific enzymatic hydrolyses. However, thedesign and control of such novel active microbeads will requirea better understanding of enzyme diffusion and, hence, thedegradation of the biopolymer networks, within the microbe-ads, to modulate the release of active substances.
■ CONCLUSION
This study demonstrated the use of microfluidics to generateJanus microbeads from polysaccharides. The design of themicrodevice, coupled with optimization of the chemical routeof biopolymer gelation, allowed the production of homo andhetero Janus microbeads from pectin−pectin and pectin−alginate with a well-defined interface, particularly in the case ofhetero Janus microbeads, thanks to specific interactionsbetween the two polysaccharides. This study opens the doorto the generation of hetero Janus microbeads from distinctbiopolymers with a short gelation time but that can specificallyinteract together. New Janus microbeads could also beproduced by using protein and polysaccharide and finelycontrolling the gelation mechanism by new microfluidicsroutes. We also demonstrated the selective degradation ofeach hemisphere of the biopolymer-based microbeads byenzymatic hydrolysis. This process will therefore provide newopportunities for the release of active substances in a controlledenvironment and could find applications in food, medicine, andcosmetics. Future investigations will focus on the release ofcompounds embedded in biopolymer networks and theircontrol by fine-tuning of the network structure and enzymatichydrolysis conditions.
NotesThe authors declare no competing financial interest.
■ ACKNOWLEDGMENTS
We would like to thank the platform BIBS and, in particular, B.Bouchet for use of the confocal scanning laser microscopyfacility and are grateful to E. Bonnin for her expertise inexperimental enzymatic hydrolysis.
■ REFERENCES(1) Stahler, K.; Selb, J.; Candau, F. Mater. Sci. Eng., C 1999, 10, 171−178.(2) Jiang, S.; Chen, Q.; Tripathy, M.; Luijten, E.; Schweizer, K. S.;Granick, S. Adv. Mater. 2010, 22, 1060−1071.(3) Du, J.; O’Reilly, R. K. Chem. Soc. Rev. 2011, 40, 2402−2416.(4) Nisisako, T.; Torii, T.; Takahashi, T.; Takizawa, Y. Adv. Mater.2006, 18, 1152−1156.(5) Walther, A.; Hoffmann, M.; Mueller, A. H. E. Angew. Chem., Int.Ed. 2008, 47, 711−714.(6) Howse, J. R.; Jones, R. A. L.; Ryan, A. J.; Gough, T.; Vafabakhsh,R.; Golestanian, R. Phys. Rev. Lett. 2007, 99, 048102.(7) Yoshida, M.; Lahann, J. Acs Nano 2008, 2, 1101−1107.(8) Glotzer, S. C.; Solomon, M. J. Nat. Mater. 2007, 6, 557−562.(9) Sheridon, N. K.; Berkovitz, M. A. Proc. Soc. Inf. Disp. 1977, 18,289−293.(10) Cho, I.; Lee, K. W. J. Appl. Polym. Sci. 1985, 30, 1903−1926.(11) Zhang, H.; Nunes, J. K.; Gratton, S. E. A.; Herlihy, K. P.;Pohlhaus, P. D.; DeSimone, J. M. New J. Phys. 2009, 11, 075018.(12) Rahman, M. M.; Montagne, F.; Fessi, H.; Elaissari, A. Soft Matter2010, 7, 1483−1490.(13) Lahann, J. Small 2011, 7, 1149−1156.(14) Charnay, C.; Lee, A.; Man, S. Q.; Moran, C. E.; Radloff, C.;Bradley, R. K.; Halas, N. J. J. Phys. Chem. B 2003, 107, 7327−7333.(15) Petit, L.; Manaud, J. P.; Mingotaud, C.; Ravaine, S.; Duguet, E.Mater. Lett. 2001, 51, 478−484.(16) Boeker, A.; He, J.; Emrick, T.; Russell, T. P. Soft Matter 2007, 3,1231−1248.(17) Cui, J.-Q.; Kretzschmar, I. Langmuir 2006, 22, 8281−8284.(18) Manoharan, V. N.; Elsesser, M. T.; Pine, D. J. Science 2003, 301,483−487.(19) Cho, Y.-S.; Yi, G.-R.; Kim, S.-H.; Jeon, S.-J.; Elsesser, M. T.; Yu,H. K.; Yang, S.-M.; Pine, D. J. Chem. Mater. 2007, 19, 3183−3193.(20) Snyder, C. E.; Yake, A. M.; Feick, J. D.; Velegol, D. Langmuir2005, 21, 4813−4815.(21) Perro, A.; Reculusa, S.; Ravaine, S.; Bourgeat-Lami, E. B.;Duguet, E. J. Mater. Chem. 2005, 15, 3745−3760.(22) Haynes, C. L.; Van Duyne, R. P. J. Phys. Chem. B 2001, 105,5599−5611.(23) Zhao, L. B.; Pan, L.; Zhang, K.; Guo, S. S.; Liu, W.; Wang, Y.;Chen, Y.; Zhao, X. Z.; Chan, H. L. W. Lab Chip 2009, 9, 2981−2986.(24) Nisisako, T.; Hatsuzawa, T. Microfluid. Nanofluid. 2009, 9, 427−437.(25) Lewis, C. L.; Lin, Y.; Yang, C. X.; Manocchi, A. K.; Yuet, K. P.;Doyle, P. S.; Yi, H. Langmuir 2010, 26, 13436−13441.(26) Liu, K.; Ding, H. J.; Liu, J.; Chen, Y.; Zhao, X. Z. Langmuir2006, 22, 9453−9457.(27) Zhang, H. W.; Betz, A.; Qadeer, A.; Attinger, D.; Chen, W. L.J. Appl. Polym. Sci. 2010, 121, 3093−3100.(28) Nie, Z. H.; Li, W.; Seo, M.; Xu, S. Q.; Kumacheva, E. J. Am.Chem. Soc. 2006, 128, 9408−9412.(29) Peppas, N. A.; Hilt, J. Z.; Khademhosseini, A.; Langer, R. Adv.Mater. 2006, 18, 1345−1360.(30) Chawla, K.; Yu, T. B.; Liao, S. W.; Guan, Z. B. Biomacromolecules2011, 12, 560−567.(31) Siracusa, V.; Rocculi, P.; Romani, S.; Dalla Rosa, M. Trends FoodSci. Technol. 2008, 19, 634−643.(32) Ulery, B. D.; Nair, L. S.; Laurencin, C. T. J. Polym. Sci., Part B:Polym. Phys. 2011, 49, 832−864.(33) Bellich, B.; Borgogna, M.; Cok, M.; Cesaro, A. Food Biophys.2011, 6, 259−266.(34) Poortinga, A. T. Langmuir 2008, 24, 1644−1647.(35) Liu, X. D.; Bao, D. C.; Xue, W. M.; Xiong, Y.; Yu, W. T.; Yu,X. J.; Ma, X. J.; Yuan, Q. J. Appl. Polym. Sci. 2003, 87, 848−852.(36) Ouwerx, C.; Velings, N.; Mestdagh, M. M.; Axelos, M. A. V.Polym. Gels Networks 1998, 6, 393−408.(37) Zykwinska, A.; Gaillard, C.; Boiffard, M.-H.; Thibault, J.-F.;Bonnin, E. Food Hydrocolloids 2009, 23, 2468−2477.
(38) Ralet, M. C.; Bonnin, E.; Thibault, J. F. J. Chromatogr., B 2001,753, 157−166.(39) Serra, C. A.; Chang, Z. Q. Chem. Eng. Technol. 2008, 31, 1099−1115.(40) Zhang, H.; Tumarkin, E.; Sullan, R. M. A.; Walker, G. C.;Kumacheva, E. Macromol. Rapid Commun. 2007, 28, 527−538.(41) Rondeau, E.; Cooper-White, J. J. Langmuir 2008, 24, 6937−6945.(42) Sarrazin, F.; Salmon, J. B.; Talaga, D.; Servant, L. Anal. Chem.2008, 80, 1689−1695.(43) Ogushi, Y.; Sakai, S.; Kawakami, K. J. Biosci. Bioeng. 2007, 104,30−33.(44) Xia, Y. N.; Whitesides, G. M. Annu. Rev. Mater. Sci. 1998, 28,153−184.(45) Annison, G.; Couperwhite, I. Appl. Microbiol. Biotechnol. 1984,19, 321−325.(46) Ngouemazong, D. E.; Tengweh, F. F.; Fraeye, I.; Duvetter, T.;Cardinaels, R.; Loey, A. v.; Moldenaers, P.; Hendrickx, M. FoodHydrocolloids 2011, 26, 89−98.(47) Funami, T.; Fang, Y.; Noda, S.; Ishihara, S.; Nakauma, M.;Draget, K. I.; Nishinari, K.; Phillips, G. O. Food Hydrocolloids 2009, 23,1746−1755.(48) Ahn, S.; Kim, D. W.; Kim, Y. W.; Yoo, J. Y. Int. J. Precis. Eng.Man. 2010, 11, 799−802.(49) Yuet, K. P.; Hwang, D. K.; Haghgooie, R.; Doyle, P. S. Langmuir2009, 26, 4281−4287.(50) Shepherd, R. F.; Conrad, J. C.; Rhodes, S. K.; Link, D. R.;Marquez, M.; Weitz, D. A.; Lewis, J. A. Langmuir 2006, 22, 8618−8622.(51) Sarrazin, F.; Bonometti, T.; Prat, L.; Gourdon, C.; Magnaudet, J.Microfluid. Nanofluid. 2008, 5, 131−137.(52) Tung, K. Y.; Li, C. C.; Yang, J. T. Microfluid. Nanofluid. 2009, 7,545−557.(53) Um, E.; Lee, D. S.; Pyo, H. B.; Park, J. K. Microfluid. Nanofluid.2008, 5, 541−549.(54) Seiffert, S.; Romanowsky, M. B.; Weitz, D. A. Langmuir 2010,26, 14842−14847.(55) Dambrine, J.; Geraud, B.; Salmon, J. B. New J. Phys. 2009, 11,075015.(56) Ribeiro, A. C. F.; Sobral, A. J. F. N.; Simoes, S. M. N.; Barros,M. C. F.; Lobo, V. M. M.; Cabral, A. M. T. D. P. V; Veiga, F. J. B.;Santos, C. I. A. V.; Esteso, M. A. Food Chem. 2011, 125, 1213−1218.(57) Masuelli, M. A. Int. J. Biol. Macromol. 2011, 48, 286−291.(58) Thiang Yian, W.; Preston, L. A.; Schiller, N. L. Annu. Rev.Microbiol. 2000, 54 (54), 289−340.(59) Braccini, I.; Rodriguez-Carvajal, M. A.; Perez, S. Biomacromo-lecules 2005, 6, 1322−1328.(60) Gohil, R. M. J. Appl. Polym. Sci. 2010, 120, 2324−2336.(61) Bonnin, E.; Le Goff, A.; Korner, R.; Vigouroux, J.; Roepstorff,P.; Thibault, J. F. Biochim. Biophys. Acta 2002, 1596, 83−94.(62) Biggs, A. R.; ElKholi, M. M.; ElNeshawy, S.; Nickerson, R. PlantDis. 1997, 81, 399−403.
