Facile Synthesis of Sustainable High Internal Phase Emulsions by a

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Facile synthesis of sustainable high internal phase emulsions by a universal and controllable route Chen Tan, Michelle C. Lee, and Alireza Abbaspourrad ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.8b03923 • Publication Date (Web): 10 Oct 2018 Downloaded from http://pubs.acs.org on October 20, 2018

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ACS Sustainable Chemistry & Engineering

Facile synthesis of sustainable high internal phase emulsions by a universal and controllable route Chen Tan, Michelle C Lee, and Alireza Abbaspourrad* Department of Food Science, Cornell University, Stocking Hall, Ithaca NY 14853, USA * Corresponding author: [email protected]; +1 607 255-2923

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ABSTRACT We demonstrate a universal route for facile synthesis of high internal phase emulsions (HIPEs), requiring only the centrifugation of ultrasonically produced oil-loaded microspheres. By sonochemically cross-linking the dispersant layer at the oil/water interface, we can engineer these microspheres against coalescence under high centrifugal force while simultaneously obtaining a high internal phase volume fraction up to 89.5%. The properties of these HIPEs and resulting porous materials can be controlled by simply adjusting the acoustic intensity. We can also deposit the non-cross-linked dispersed materials back to the microsphere surface, creating an extra electrostatic layer and polyelectrolyte complexes coating, thus further reinforcing the processability of HIPEs. Our strategy could work for nearly any material as long as it can be crosslinked during acoustic cavitation. Thus, these scalable HIPEs can be fully sustainable as they are solely stabilized by natural materials, such as proteins and polysaccharides (as low as 0.5 wt%), without introducing any additional surfactants or synthetic particles, or requiring the additional synthesis of Pickering emulsions. Keywords: high internal phase emulsions; centrifugation; ultrasonication; cross-linking; sustainable; universality


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INTRODUCTION Centrifuging a binary emulsification of oil and water will typically result in the separation of the oil and aqueous phases as the emulsified droplets coalesce under the centrifugal field. Thus, centrifugation is widely used as a powerful method to destabilize emulsions.1,2 However, centrifugation can also be applied to concentrate and collect dispersed droplets when an appropriate centrifugal speed is utilized. If no coalescence occurs, the centrifugation process can force the emulsified droplets into contact with each other while simultaneously excluding the continuous phase from the inter-droplet volume. When the internal phase volume fraction reaches 0.74 or greater, a unique type of emulsion gel is achieved, known as a high internal phase emulsion (HIPE).3 Due to their large interfacial area and tunable flow behavior, this particular emulsified system has attracted considerable attention in food products,4 drug delivery systems,5 oil recovery,6 and as templates or scaffolds in material science.7–10 Surprisingly, very few centrifugation-based preparation methods for HIPEs have been reported,3,11 though it is a simple and fast procedure for HIPE production. The main challenge of HIPE formation during centrifugation is the simultaneous demand for a high internal phase volume fraction as well as stability against phase separation under high centrifugal forces. That is, concentrating emulsions up to 74% of the internal phase requires a sufficient centrifugal speed, which could in turn accelerate coalescence. To overcome this tradeoff during centrifugation, researchers have tried stabilizing the oil droplets using high amounts of phospholipids, oleosins, and proteins.11 However, destabilization due to coalescence still takes place, resulting in large oil patches. As an alternative, the most common preparation method for HIPEs is the progressive addition of the dispersed phase (i.e., the internal phase) to the continuous phase (i.e., the external phase) under constant agitation by vortex-



mixing, stirring, or homogenization. However, these methods are far from robust. For example, to avoid phase inversion, the dispersed phase must be continuously added dropwise with care into the continuous phase, which is a time-consuming process and greatly restricts the efficiency for large-scale production. Another issue is that conventional HIPEs must be stabilized by a large quantity of surfactants12 or colloidal particles (also known as Pickering HIPEs),13,14 which requires careful surfactant choice and additional

sophisticated syntheses of Pickering emulsions.

Additionally, the use of synthetic surfactants and non-degradable particles presents potential environmental risks. The limitations of these current methods and synthetic materials prompted us to re-explore the potential of centrifugation to develop an efficient and “green” HIPE synthesis method. Here, we present a simple, universal, scalable, and environmentally friendly route for the synthesis of HIPEs that feature both excellent stability and processability, via centrifuging ultrasonically produced oilloaded microspheres (Figure 1A, details are provided in the Supporting Information). By creating a cross-linked layer at the oil/water interface during the acoustic cavitation process, we can engineer these microspheres against coalescence within a high centrifugal field while simultaneously obtaining a high internal phase volume fraction. This strategy could provide a universal route to fabricate processable HIPEs, as long as the dispersed materials in the continuous phase can be cross-linked by ultrasonication. With this technique, we can synthesize “green” HIPEs and porous materials based entirely on natural products without the use of any surfactants or synthetic particles, or requiring the additional synthesis of Pickering emulsions. EXPERIMENTAL SECTION Materials. Bovine serum albumin (BSA, fraction V, 98% purity) was purchased from CalBiochem (catalog number 12659). Fluorescein-conjugated bovine serum albumin (FITC-BSA)


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was purchased from Life Technologies (catalog number A23015, Invitrogen, Life Technologies). Chitosan of medium molecular weight (Mw = 190–310 kDa, 75–85% degree of deacetylation, viscosity 200–800 cP) was purchased from Sigma-Aldrich (St. Louis, MO, US). Chondroitin sulfate type A from bovine trachea cartilage (Mw = 5–10 KDa) was purchased from Bulk Supplements (Henderson, NV, US). Pectin powder from citrus peel, dextran (from Leuconostoc spp., Mw~ 100,000), and sodium dodecyl sulfate (SDS) were purchased from SigmaAldrich. 100% pure corn oil (Mazola, ACH Food Companies, Inc., Cordova, TN) and vegetable oil were purchased from a local supermarket. The kappa-carrageenan was provided by TIC Gums Incorporated (White Marsh, MD). Xanthan gum, guar gum, and konjac were provided by Colony Gums (Monroe, NC). All other reagents used were of analytical grade. Preparation of oil-loaded microspheres by ultrasonication. The proteinaceous microspheres were prepared according to our earlier study.15 Briefly, 8 mL of BSA aqueous solution (0.5%, w/w) was added to 4 mL of cyclohexane or other types of oil (chloroform, decane, hexane, toluene, corn oil, and vegetable oil), and exposed to high-intensity ultrasound using a 750 Watt ultrasonic processor with a high power sonic tip operated at 20 kHz frequency (VC 750, Sonics vibra-cell, Sonics & Materials, Newtown, CT, USA). The bottom of the 13 mm diameter ultrasonic horn was positioned at the oil/water interface and the solution was sonicated in an ice bath for 10 min at different amplitudes (20%, 30%, 40% and 50%) corresponding to acoustic powers of 150, 225, 300, and 375 W cm-2 (5 s on, 2 s off). For the preparation of the polysaccharide-based microspheres, seven combinations of polysaccharides were chosen, including chondroitin sulfate/dextran, chondroitin sulfate/guar gum, chitosan/guar gum, chitosan/konjac, κ-carrageenan/pectin, chitosan/pectin, and chitosan/xanthan gum. The concentration of each individual polysaccharide solution was 0.5% (w/w). To prevent



complex formation after mixing, the pH of each polysaccharide solution was adjusted to 2 using hydrochloric acid before mixing.16 The two polysaccharide aqueous solutions were mixed at a ratio of 1:1 and added to 4 mL cyclohexane. The ultrasonic horn was positioned at the oil/water interface followed by sonication at the same condition as described above. To generate extra electrostatic layer and complex coating on the microsperes, the pH of the sonicated polysaccharide microsphere solutions was adjusted to higher values (3–6) before further centrifugation. To prepare microspheres stabilized by polyelectrolyte complexes alone, we adjusted the pH of the individual chitosan and xanthan gum aqueous solution (0.5%) to 5 before mixing. The xanthan gum solution was then added dropwise into the chitosan solution at a ratio of 1:1. The resulting polyelectrolyte complex (8 mL) solution was then added to 4 mL cyclohexane and sonicated at the same condition as described above. Preparation of HIPEs. The freshly prepared microspheres were then transferred into a 50 mL Falcon tube and centrifuged at different speeds (100–15000 g) for 5 min. A rigid cream layer, i.e., the HIPE, formed on top of the aqueous phase after centrifugation. The aqueous phase was also collected in order to calculate the internal phase volume fraction of the HIPE. For comparison, conventional HIPEs were prepared by a standard homogenization emulsification method,14 using a T25 digital Ultra Turrax (IKA-Werke, Wilmington, NC) at 13000 rpm. The cyclohexane was continuously added dropwise to the required volume of BSA (0.5%, w/w) during homogenization until the desired internal oil volume fraction was achieved. Morphological characterization. To perform scanning electron microscopy (SEM), HIPEs were placed onto a silicon wafer, freeze-dried, and sputtered with gold.17 The porous structure of the freeze-dried HIPEs was observed on a LEO 1550 SEM equipped with a GEMINI field emission column. Confocal imaging of fresh microspheres or resultant HIPEs were carried out on a Zeiss


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confocal microscope (LSM 710, Carl Zeiss, Göttingen, Germany). The structural morphology was studied by adding Nile red and FITC-BSA. Nile red was directly added to the final obtained microspheres or HIPEs. The FITC-BSA was first added into the non-fluorescent BSA solution at a mass ratio of 1:500 before further preparation of the microspheres. Excitation/emission wavelengths for Nile red and FITC-BSA were 488/566 nm and 488/515 nm, respectively. Microsphere size. The average particle size was determined by a commercial zeta-sizer (NanoZS90, Malvern Instruments Ltd., U.K.) with a He/Ne laser (λ = 633 nm) and scattering angle of 90o. The samples treated by different acoustic intensities were diluted with the same buffer solution before measurement.18 FTIR spectra. Fourier-transform infrared (FTIR) spectra of pure polysaccharides and their corresponding freeze-dried HIPE foams were recorded on an IRAffinity-1S spectrometer equipped with a single-reflection attenuated total reflectance (ATR) accessory (Shimadzu Corp., Kyoto, Japan). The FTIR spectra were scanned between 500 to 4000 cm-1 at a resolution of 4 cm-1. Rheology measurement. Dynamic rheological measurements were performed on an AR1000N controlled stress rheometer (TA Instruments, Inc., Ghent) using a plate geometry. A range of experiments, including amplitude (stress = 0.1–1000 Pa, frequency = 1 Hz), and frequency sweeps (0.1–10 Hz, strain = 1%), were carried out at 25 °C. The elastic modulus (G′) and loss modulus (G″) were recorded using the RheoWin 3 Data Manager. Viscosity was measured at a range of shear rates of 0–10 s-1 at 25 °C. Stability assays of HIPEs. The stability of the HIPEs was evaluated against the surfactant SDS and pH 7 phosphate buffer. Briefly, 0.1 g of the HIPE was accurately weighted in a centrifuge tube. Then, 1 mL aqueous solution with 1% SDS or phosphate buffer with pH 7 was added and gently swirled by hand until the HIPE was completely dissociated. The BSA that was not adsorbed



on the oil droplets releases from the droplet interface after dilution. The morphological change of the oil droplets after dilution was monitored from CLSM images by dropping the diluted emulsions on a glass slide. To determine the adsorbed concentration of BSA, the resulting mixture was centrifuged at 8000 g for 5 min. The supernatant (0.1 mL) was collected and diluted with 0.4 mL deionized water, followed by reaction with protein reagent as described follows. Determination of adsorbed BSA on the microspheres. The protein reagent was prepared according to the classic method,19 as follows: Coomassie Brilliant Blue G-250 (100 mg) was dissolved in 50 mL 95% ethanol, followed by the addition of 100 mL 85% (w/v) phosphoric acid. The resultant solution was diluted to a final volume of 1 L with deionized water. Final concentrations in the reagent included 0.01% (w/v) Coomassie Brilliant Blue G-250, 4.7% (w/v) ethanol, and 8.5% (w/v) phosphoric acid. To determine the concentration of excess protein from the HIPEs, 0.5 mL of supernatant obtained after centrifugation of the microspheres was mixed with 5 mL of the protein reagent in a tube and then vortexed. After 2 min, the absorbance was measured at 595 nm using a UV-2600 spectrophotometer (Shimadzu, Japan). The standard protein curve was obtained through pure protein solutions in a concentration gradient (0.01–0.1 mg/mL) (y = 5.9016x + 0.3241, R2=0.993), in which y and x correspond to the absorbance and protein concentration (mg/mL), respectively. Each experiment was carried out in triplicate. Determination of adsorbed polysaccharides on the microspheres. The amount of polysaccharides (chitosan and xanthan gum in this case) on the microspheres was determined based on the phenol-sulfuric acid method, as described previously.20 The microsphere suspension was centrifuged at 10,000 g for 20 min. A 30 µL aliquot of the supernatant was taken and placed in a tube. Then, 0.16 mL of freshly prepared 5% (v/v) phenol aqueous solution and 0.8 mL of concentrated sulfuric acid was added. After incubation for 20 min at room temperature, the


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absorbance was recorded at 490 nm with a UV-2600 spectrophotometer (Shimadzu, Japan). Distilled water was used as a blank. RESULTS AND DISCUSSION Generation of protein-based HIPEs. For proof-of-concept, we chose bovine serum albumin (BSA) aqueous solution as the continuous phase and cyclohexane as the dispersed phase. We first applied ultrasound to this two-phase system by positioning the ultrasonic horn at the aqueous/organic interface to produce the oil-loaded proteinaceous microspheres. During the acoustic cavitation process, the generation of superoxides helped cross-link the BSA molecules through the formation of covalent disulfide bonds between cysteine residues, forming a permanent and stiffened layer at the oil/water microsphere interface.16,21 When we applied a centrifugal field to this system, the microspheres were forced into contact with each other, and excess protein solution was forced out from between the microspheres to produce a separate phase. As a result, a firm cream layer was formed on top of the aqueous phase, which was able to support its own weight even when the tube was inverted (Figure 1B (i)). These HIPEs were solid-like emulsions, enabling them to be easily collected from the centrifugal tube (Figure 1A). Due to the high viscoelastic gel texture, these HIPEs could be processed into different shapes, which remained stable without any oiling off after air-drying at room temperature for 1 h. For comparison, we prepared a control by mixing BSA and cyclohexane using a conventional homogenization emulsification method.18 However, the homogenization-produced system underwent phase separation after centrifugation, even though it was stable when initially prepared (Figure 1B (ii)). This phenomenon demonstrates the high stability of the ultrasonically-produced microspheres within a high centrifugal field.



Figure 1. Formation and characterization of centrifugation-produced HIPEs assisted by ultrasonication. (A) Schematic illustration of the protein-based HIPE synthesis through successive ultrasonication and centrifugation. The images on the right show the HIPEs were easily removed from the centrifugal tube. The inset demonstrates the processability of the HIPEs. Images were taken 1 h after air drying at room temperature. (B) Visual appearance of the ultrasonicationproduced microspheres after centrifugation at different speeds for 5 min (i), and the homogenization-produced system after centrifugation at 10000 g for 5 min (ii). (C) CLSM images of ultrasonication-produced microspheres at different centrifugal speeds, showing the oil phase


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stained with Nile red. Here, the HIPEs were prepared using cyclohexane as the dispersed phase and BSA aqueous solution (0.5 wt%) as the continuous phase. (D) CLSM images of the initial and dilute HIPEs stained with FITC-BSA. The HIPEs were prepared by standard homogenization emulsification (a-c) and centrifugation (d-f). Initial HIPEs (a, d) and those diluted with pH 7 phosphate buffer (b, e) and 1% SDS (c, f). (E) Photographs showing that HIPEs could be formed using a variety of oil phases after centrifugation at 10000 g for 5 min, in addition to a CLSM image of the HIPE made with vegetable oil as an example. The acoustic intensity was 225 W cm-2. All scale bars are 1 µm. By simply adjusting the centrifugation speed, we can modulate the volume fraction of oil (ϕ) in the cream. At 5000 g, 10000 g, and 15000 g, we calculated the volume fractions as 83.3%, 85.7%, and 89.5%, respectively, which is close to the highest theoretical value of 91%.3 To monitor the evolution of the cream morphology during centrifugation, we stained the oil phase with Nile red and visualized the materials with confocal laser scanning microscopy (CLSM). As the centrifugal force was increased to 10000 g, the cream layer represented a typical structure of HIPEs, wherein the microspheres were closely packed together and deformed, exhibiting polygonal shapes (Figure 1C). When higher centrifugal force was applied (15000 g), some microspheres merged, but there was no appearance of phase separation. These results show that we can produce stable HIPEs using this centrifugation technique by applying an appropriate centrifugal force (10000 g) and cross-linking the proteins at the microsphere interface to prevent coalescence. To visually confirm this cross-linking, we doped fluorescence-labeled BSA (FITC-BSA) into the bulk BSA solution to stain the aqueous phase. For comparison, conventional HIPEs containing FITC-BSA were also prepared with the same phases by a standard homogenization emulsification method.14 For the homogenization-produced HIPEs,



the fluorescence intensity from the protein was high in the continuous phase, but considerably less at the microsphere interface (Figure 1D). After diluting the HIPEs with pH 7 phosphate buffer, the fluorescence intensity was further decreased due to the release of protein from the aqueous phase and interface. In contrast, a large proportion of the ultrasonically treated proteins preferentially migrated onto the interface, as observed from the high fluorescence intensity around the microspheres in the ultrasonication-produced HIPEs. Dilution also did not induce detachment of these protein molecules from the interface. Additionally, the high resistance to interface competition after exposing the fluorescently-labeled microspheres to the surfactant sodium dodecyl sulfate (SDS) directly confirmed the proteins were cross-linked around the microspheres (Figure 1D), in contrast to the considerable replacement (i.e., high fluorescence loss) of native proteins from the interface of the homogenization-produced HIPEs. We also demonstrated the cross-linking reaction of proteins can occur at the interface of water and any oils studied (Figure 1E), enabling the formation of HIPEs during centrifugation. Next, we further modulated the cross-linking density by adjusting the ultrasonic energy input. Increasing the acoustic intensity from 150 to 300 W cm-2 led to decrease in particle size from 4.2 ± 0.7 µm to 2.8 ± 0.3 µm (Figure S1). In addition, the smaller microspheres facilitated subsequent packing of the HIPEs during centrifugation (Figure S2). We also evaluated the influence of acoustic intensity on the structure of porous materials prepared by evaporating the water and oil phases of the HIPEs during freeze-drying. Scanning electron microscopy (SEM) imaging demonstrates that after freeze-drying, the porosity of the resulting protein foams was determined by the original microsphere size (Figure S2). Note that the increase in the acoustic intensity allowed us to switch between open- and closed-cell structures of the porous materials. We believe that the high ultrasound power significantly enhanced cross-linking at both the interface and


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continuous aqueous phase, creating a stronger gel network that did not rupture to form pore throats during or after freezing, thus ultimately leading to a closed-cell structure. This conclusion was supported by the increased amounts of protein adsorbed at the microsphere interfaces as the acoustic intensity was increased (Figure S3). Additionally, such enhanced cross-linking endowed the HIPEs with higher viscoelasticity, as evidenced from the increased storage modulus (G’) during stress and frequency sweeps (Figure S4). Generation of polysaccharide-based HIPEs. Again, we should emphasize that the criterion for providing stability to the HIPEs during centrifugation is the formation of the cross-linked layer at the oil/water interface. Therefore, we hypothesized it was possible for other materials to stabilize HIPEs using this same strategy, as long as they could be cross-linked by ultrasonication. In this case, we chose seven combinations of natural polysaccharides to prepare microspheres under ultrasonic treatment, followed by centrifugation (Figure 2A). The pH of each polysaccharide solution was adjusted to 2 in order to make the molecules positively charged, which prevented the formation of polyelectrolyte complexes via electrostatic interaction after mixing. Although all the initially prepared formulations were stable after ultrasonication, only the combinations of κcarrageenan/pectin, chitosan/pectin, and chitosan/xanthan gum led to a firm creaming layer postcentrifugation (Figure 2B), suggesting the cross-linking between them. In the case of the chitosan/xanthan gum, Fourier transform infrared (FTIR) spectra (Figure 2C) revealed that the amide I band of chitosan located at 1633 cm-1 shifted to 1624 cm-1 after sonication, and the peaks at 1558 cm-1 (NH2 deformation) and 1031 cm-1 (NH2 angular deformation) were nearly absent. Meanwhile, the C=O stretching band of the xanthan gum shifted to a lower wavenumber (1712 cm-1) along with the disappearance of the peak at 1409 cm-1 (OH bending in carboxylic acids) in the freeze-dried chitosan/xanthan gum HIPEs. These peak changes demonstrate the ultrasonically-



induced chemical interaction between the amino groups of chitosan and the carboxylic groups of xanthan gum via amide linkage formation.16 Similar interactions were also found for κcarrageenan/pectin and chitosan/pectin (Figure S5) through amino, carboxyl, hydroxyl, and sulfhydryl groups, corresponding to the formation of stable HIPEs. However, HIPEs could not be produced without the help of cross-linking (no band changes were detected in other formulations). Transmission electron microscopy (TEM) and CLSM images demonstrate that the chitosan/xanthan gum microspheres did not coalesce after centrifugation and maintained their original size in the rigid HIPE layer (Figure 2D and E). The chitosan/xanthan gum HIPE was also able to form a stable monolith foam with closed-cell structure after freeze-drying (Figure 2E). Interestingly, this polysaccharide foam displayed oleophilic behavior, in which the water contact angle was 111°, while hexane was immediately adsorbed (Figure 2F), likely due to both the strong cross-linking of the continuous phase and increase in surface hydrophobicity induced by ultrasonication.22 Thus, this polysaccharide material could potentially be used as an absorbent for oil-water separation. For instance, we demonstrated that hexane was quickly adsorbed by the corresponding polysaccharide foam, which remained floating on water (Figure S6).


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centrifugation

A probe

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ultrasonication

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oil

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initially prepared

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after centrifugation

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cross-linking at interface free polysaccharides

polycationic polysaccharide (i)

polycationic polysaccharide (ii)

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chitosan Amide I band NH2 deformation NH2 angular deformation C=O stretching OH deformation

xanthan gum

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chitosan/xanthan gum 800

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-1

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Figure 2. Fabrication and characterization of various polysaccharide-based HIPEs. (A) Schematic illustration of the synthesis of polysaccharide-stabilized HIPEs. The red and blue icons in the tube represent two different types of polysaccharide, both of which are positively charged at pH 2. (B) Visual appearance of ultrasonically produced polysaccharide microspheres, before and after centrifugation. (C) FTIR spectra of pure chitosan, pure xanthan gum, and freeze-dried chitosan/xanthan gum HIPEs. The schematic shows the possible interactions between the



polysaccharides. (D) TEM image of chitosan/xanthan gum microspheres before centrifugation. Scale bar is 1 m. (E) CLSM image of chitosan/xanthan gum HIPEs and corresponding SEM image after freeze-drying. Scale bars are 1 m. (F) Photograph of a droplet of water (a) and hexane (b) on the surface of a chitosan/xanthan gum foam taken during contact measurements. The inset shows the monolith foam. Here, all the HIPEs were prepared by ultrasonication (225 W cm-2) and subsequent centrifugation at 10000 g for 5 min with cyclohexane as the dispersed phase and polysaccharide aqueous solution (0.5 wt% polysaccharides, pH 2) as the continuous phase. Strengthening HIPEs by extra electrostatic deposition. Based on these results, we believe that the cross-linked layer around the microspheres contribute to their stability against coalescence under high centrifugal force. However, we note that the degree of ultrasonication-induced crosslinking at the interfaces is typically very low, even at high ultrasound power.23,24 In our case, more than 40% of the initially added BSA was not cross-linked at the microsphere interface even though the acoustic intensity was increased to 375 W cm-2 (Figure S3). For chitosan/xanthan gum, only 48% of the polysaccharides took part in the cross-linking reaction and subsequent formation of the HIPEs. To create more rigid/thicker layer stabilized microspheres, we tried to deposit these excess non-cross-linked polysaccharides back to the microsphere surfaces prior to centrifugation. As noted in Figure 2A, the cross-linked microspheres prepared at pH 2 should be positively charged. Since the pKa values of chitosan and xanthan gum are approximately 6.225 and 2.6,26 respectively, we adjusted the pH (2) of the sonicated solution to higher values (3-6) before centrifugation. We hypothesized that the free xanthan gum would become negatively charged at pH 3-6 because of the ionization of the carboxylic acid groups, thus enabling the deposition of an extra layer of xanthan gum on the existing positively charged microspheres, as well as forming polyelectrolyte complexes with the free positively charged chitosan in solution (Figure 3 and Figure S7).


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Figure 3. Morphological evolution of chitosan/xanthan gum microspheres after additional electrostatic deposition of excess non-cross-linked polysaccharides. Schematic illustrating the structure of an individual microsphere after coating with an extra xanthan gum layer and chitosan/xanthan gum complexes. TEM images of the microspheres at pH 5 (A, B), and SEM images of microspheres at pH (C) 4, (D, E) 5, and (F, G) 6. (H) SEM image showing the microspheres stabilized solely by chitosan/xanthan gum complex without cross-linking. Scale bars are 1 m. The microspheres were prepared by ultrasonication (300 W cm-2) of the polysaccharide solutions (pH 2) and cyclohexane, followed by adjusting the pH (3-6). This hypothesis was verified by the pH-dependent decrease of positive charges of the chitosan/xanthan gum microspheres (Figure S8). The increased adsorbed amount of



polysaccharides further confirmed the progressive deposition of non-cross-linked polysaccharide on the microspheres with increasing pH (Figure S8). TEM visualization displayed a significantly different structure of the microspheres at pH 5 (Figure 3A, B) from those at pH 2 (Figure 2D). At pH 5, we can clearly see an extra thick xanthan gum layer deposited on the surface of the microspheres. More interestingly, many chitosan/xanthan gum complexes (i.e., small “beads” labeled in the images) also attached to the microspheres. SEM images (Figure 3C-G) show that as the pH increased from 4 to 6, more complexes were observed. For comparison, we also prepared the microspheres stabilized by chitosan/xanthan gum complexes at pH 5 (Figure 3H) to help us better understand the influence of complexes alone on the stability of the resulting HIPEs without cross-linking. After centrifugation, all these microspheres prepared at different pH values formulations were able to produce closely packed HIPEs wherein numerous complexes (black dots in the CLSM images) coated around the microsphere surfaces (Figure 4A-D). However, we observed partial coalescence in the HIPEs prepared at pH 6 (Figure 4C). The probable reason was that the highly thick interfacial layer around the microspheres (Figure 3G) would in turn reduce the surface deformability, resulting in rupture.27 Analysis of the rheological properties shows that the additional electrostatic deposition of polysaccharides at pH 3-6 contributed to the significantly stronger viscoelasticity of the HIPEs (Figure 4E). The storage modulus G’ at pH 5 was increased by approximately 7.2- and 11.9-fold as compared to that stabilized by the cross-linked layer alone (pH 2) and that stabilized by complex alone, respectively. The strength of the HIPEs determined their processability (Figure 4F). The encapsulated organic solvent (cyclohexane) rapidly evaporated within 2 h air-drying for HIPEs stabilized solely by the cross-linked layer (pH 2) and complexes, accompanied by the complete collapse of the structure. However, the interfacial


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network formed at pH 5 and 6 can strongly delay the evaporation of cyclohexane, with a dense and intact structure after 2 d drying, as demonstrated by SEM (Figure S9).

Figure 4. CLSM images of HIPEs stabilized by additional electrostatic deposition at (A) pH 4, (B) 5, and (C) 6, and (D) HIPEs stabilized solely by chitosan/xanthan gum complex without crosslinking at pH 5. Scale bars are 1 m. (E) Frequency sweeps of the storage (G’) and loss (G’’)



moduli of the chitosan-xanthan gum HIPEs prepared at different pHs. “Complex” refers to the HIPEs stabilized by chitosan/xanthan gum complexes prepared at pH 5. (F) Processability of the HIPEs. Images of the heart-shaped HIPEs were taken after 2 h and 2 d air-drying at room temperature. The HIPEs were prepared by ultrasonication (300 W cm-2) and subsequent centrifugation at 10000 g for 5 min with cyclohexane as the oil-dispersed phase and chitosan/xanthan gum aqueous solution (0.5%) as the continuous phase. We attributed such higher processability to the strengthened thickness/rigidity of the interfacial layer. However, we also noticed that the repulsions of the microspheres were considerably reduced with the additional electrostatic deposition (as demonstrated by zeta potential in Figure S8). This could favor microsphere packing in the HIPEs and thus improve the processability as well. To confirm this, we added high concentration NaCl (3 M) to screen the charge of the microspheres at pH 2 and 5 (zeta potential was around 0 mV), followed by centrifugation. Indeed, we found that the decreased repulsion by the addition of salt did not significantly affect the packing of microspheres in the HIPEs, but greatly increased the viscoelasticity of the HIPEs at both pH 2 and 5 (Figure S10) compared to those in the absence of salt, which was understandable because densification occurred at the interface.28 However, the HIPEs at pH 5 still exhibited much higher viscoelasticity than those at pH 2 (Figure S10), demonstrating the predominating role of the strengthened interfacial layer in stabilizing the HIPEs. CONCLUSIONS In summary, we have explored centrifugation assisted by ultrasonication as a simple universal route for the synthesis of ultra-stable and processable HIPEs in a manner that requires neither specialized equipment nor the use of toxic substances. This route could work for various materials as long as they can be cross-linked during acoustic cavitation. Therefore, careful optimization of


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surfactant‐use or the sophisticated synthesis of Pickering emulsions was not required in our procedure. Additionally, the natural ingredients of our HIPEs made up of pure protein and polysaccharides could have enormous application in food industry, such as for the formulations of mayonnaise and salad dressings. The cross-linked layer at the microsphere interfaces contributed to the excellent stability of these emulsions under centrifugal force and hence the resultant stable HIPEs. These HIPEs produced gel and porous materials whose texture and microstructures could be tuned by varying the ultrasonic energy input. We were also able to create an extra electrostatic layer and polyelectrolyte complex coating on the microsphere surfaces to further strengthen the HIPEs. This technique combining electrostatic deposition and ultrasonically-induced cross-linking could also be utilized to enhance the delivery performance of ultrasonically produced containers for bioactive compounds. Finally, our strategy can lead to fully “green” monolith foams with tunable porosity and structure, with new opportunities for the broader development of various renewable porous materials for biological and pharmaceutical applications. CONFILICTS OF INTEREST There are no conflicts to declare. ACKOWLEDGEMENTS This publication was made possible by the research funding provided by New York State Milk Promotion Board. We thank the Cornell Center for Materials Research (CCMR) for use of their facilities. CCMR facilities are supported by the National Science Foundation under Award Number DMR-1719875. This material is based upon work that is supported by the National Institute of Food and Agriculture, U.S. Department of Agriculture, Hatch under 1010696. SUPPORTING INFORMATION



The Supporting Information is available free of charge on the ACA Publications website. CLSM of microspheres; SEM of HIPE foams; Storage and loss moduli of HIPEs; FTIR spectra of polysaccharide-based HIPEs; water-oil separation by HIPE foam; Zeta potential of polysaccharide microspheres; CLSM images of HIPEs with salt. REFERENCES (1)

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Table of Contents (TOC)

Synopsis We present a universal route for facile synthesis of sustainable HIPEs, requiring only the centrifugation of ultrasonically produced oil-loaded microspheres, without using any surfactants or synthetic particles.


Facile Synthesis of Sustainable High Internal Phase Emulsions by a

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