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Microgel stabilized emulsions: Breaking on demand
Colloids and Surfaces A: Physicochem. Eng. Aspects 495 (2016) 193–199
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Colloids and Surfaces A: Physicochemical and Engineering Aspects journal homepage: www.elsevier.com/locate/colsurfa
Microgel stabilized emulsions: Breaking on demand Susanne Wiese a , Yoanna Tsvetkova a,1 , Nadine J.E. Daleiden a , Antje C. Spieß b,c,d , Walter Richtering a,∗ a
RWTH Aachen University, Physical Chemistry, Landoltweg 1, 52056 Aachen, Germany RWTH Aachen University, AVT —Enzyme Process Technology, Aachen, Worringerweg 1, 52074 Aachen, Germany DWI—Leibniz-Institute for Interactive Materials Research, Forckenbeckstraße 50, 52074 Aachen, Germany dTU Braunschweig, Institute for Biochemical Engineering ibvt, Gaußstr. 17, 38102 Braunschweig, Germany d TU Braunschweig, Institute for Biochemical Engineering ibvt, Gaußstr. 17, 38102 Braunschweig, Germany b c
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
• Microgels are employed to stabilize and break emulsions on demand.
• Microgel composition and structure influence temperature sensitivity of emulsions. • We control whether microgel stays in solution or aggregates while breaking the emulsion. • Microgels as reversible emulsion stabilizers can be adopted to requirements of enzyme reactions in biocatalysis.
a r t i c l e
i n f o
Article history: Received 28 July 2015 Received in revised form 2 February 2016 Accepted 3 February 2016 Available online 17 February 2016 Keywords: Switchable microgel Switchable emulsion Biocatalysis Poly-N-isopropyl-acrylamideco-N-isopropyl-methacrylamide
a b s t r a c t Here, we report on how to stabilize and break emulsions that are compatible with enzymatic reaction conditions. Many substrates of enzymatic reactions are soluble in unpolar organic solvents whereas the enzymes themselves often need an aqueous environment. We use a buffer solution (triethanolamine hydrochloride) as aqueous and MtBE (tert-butyl methyl ether) as organic phase which provide good enzyme compatibility. We are able to break emulsions in a desired temperature range by using NiPAM–NiPMAM microgels with different monomer compositions and architecture, respectively. Our microgels need to deswell to about 55% of its swollen size at room temperature to let the emulsion break. Emulsions can be broken such that the microgels are either colloidally stable in the aqueous phase or flocculated. The temperature interval in which the microgels stay colloidally stable while the emulsion is broken is broader for the core–shell microgel than for the copolymer microgel. The behavior of the microgels in aqueous solution allows predicting: (i) the temperature at which the emulsion breaks and (ii) whether microgels flocculate or not during breaking the emulsion. However, the partial miscibility of the organic phase with the aqueous phase has to be taken into account.
∗ Corresponding author. Fax: +49 214 80 92327. E-mail addresses: [email protected] (S. Wiese), [email protected] (Y. Tsvetkova), [email protected] (N.J.E. Daleiden), [email protected] (A.C. Spieß), [email protected] (W. Richtering). 1 UK Aachen, Institut für Biomedizinische Technologien—Experimentelle Molekulare Bildgebung, Pauwelsstraße 30, 52074 Aachen, Germany. http://dx.doi.org/10.1016/j.colsurfa.2016.02.003 0927-7757/© 2016 Elsevier B.V. All rights reserved.
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Thus, we are able to stabilize and break emulsions by employing microgels as responsive emulsifiers and to adapt the microgels to the requirements of biocatalytic processes. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Many substrates of enzymatic reactions are soluble in organic solvents whereas the enzymes themselves often need an aqueous environment [1]. When performing enzyme reactions in industrial scale a large reservoir of substrates is needed that leads to a two phase system, the aqueous phase containing the enzyme and an organic phase containing the substrates and eventually products. Surfaces as well as interfaces may lead to enzyme unfolding and therefore, enzyme inactivation [2]. One approach to protect enzymes from denaturation at the interface is to introduce microgels as surface-active species. Microgels allow for emulsion stabilization [3]. Emulsions are systems of two immiscible liquids, mostly an aqueous phase and a non-aqueous phase, a so called organic or oil phase. Surfactants [4], proteins [5], star-copolymers [6] or nanoparticles [7,8] stabilize droplets of one liquid (dispersed phase) dispersed in the other liquid (continuous phase). A new class of stabilizing agents are microgels [3]. Microgels are soft cross-linked, solvent swollen polymer particles [9,10]. Microgel stabilized emulsions have great potential for industrial application. e.g., in drug delivery [11] or biocatalysis [12,13]. One special property of some microgels is their sensitivity to external stimuli [14–22]. For example N-isopropylacrylamide (NiPAM)-based microgels undergo a volume change with increasing temperature [14]. The VPTT (volume phase transition temperature) can be adjusted by using different co-monomers. The switchability of the microgels leads to switchability of the emulsions [3,23]. Since the enzyme requires specific conditions with respect to reaction temperature and stability temperature the emulsion needs to be switched in a certain temperature range. Furthermore, the stability and reactivity of the enzyme depends on the pH value and the used solvents. In this study we use Lactobacillus brevis alcohol dehydrogenase (Lb-ADH) as model enzyme whose optimal pH value is maintained using the buffer TEA·HCl (triethanolamine hydrochloride). TEA·HCl buffer has the advantage that the temperature has only a moderate influence on the pH value (pK = −0.02 pH/K) [24]. We introduce tert-butyl methyl ether (MtBE) [25] as organic phase due to its good enzyme compatibility [26]. MtBE is a polar and aprotic organic solvent and thus different from heptane and octanol which have previously been used as organic phase in microgel stabilized emulsions [23,27,28]. In general enzymes denature with rising temperatures. Therefore, microgels used in emulsions for biocatalysis: (i) must be stable in the buffer, (ii) must stabilize an emulsion of an polar, aprotic organic phase with a buffer, (iii) must stabilize these emulsions at the optimal enzymes’ working temperature (in our case 30 ◦ C), (iv) break the emulsion at a temperature above 30 ◦ C, which however is yet low enough to avoid enzyme denaturation. Mostly P(NiPAM-co-MAA) microgels (poly-(N-isopropylacrylamide-co-methacrylic acid)) were used to stabilize emulsions but in recent studies also uncharged microgels were used [25,28].
P(NiPAM-co-MAA) microgel-stabilized emulsions can be broken by increasing the temperature or by changing the pH value [23,27,29]. We use microgels made of two uncharged monomers, NiPAM and NiPMAM (N-isopropyl-methacrylamide), to stabilize emulsions. The general structure of both monomers is quite similar; they differ only in a methyl group at the polymer backbone. The VPTT of pure PNiPAM microgels is ca. 30–32 ◦ C [9,30] whereas the VPTT of pure PNiPMAM is ca. 40–43 ◦ C [30,31]. With these two monomers we now create microgels with different composition and structure. Our aim is to stabilize and break emulsions on demand. In this study we show that the switching behavior of microgels and emulsions correlate with each other. We found that emulsions, stabilized with NiPAM–NiPMAM-microgels stabilize emulsions at room temperature and can be broken at temperatures slightly above their microgels’ transitions temperatures. We are able to adapt microgels to the requirements of enzymatic reactions, thus emulsions can be stabilized and be broken on demand in a desired temperature range. Furthermore, we can predict emulsion behavior from microgel properties measured in aqueous solution. Emulsions stabilized with the core–shell microgel break already when the temperature is between the transition temperatures of core and shell, respectively. In other words the emulsion breaks when the stabilizer is not fully collapsed but partially swollen. This particular behavior opens a wide field of possible applications. 2. Material and methods 2.1. Chemicals N-Isopropylacrylamide (NiPAM) (>99.0%) and the cross-linker N,N -methylenebisacrylamide (BIS) (>99.5%) were purchased from Acros Chemicals; N-isopropylmethacrylamide (NiPMAM) (>99%) from Aldrich; the initiator potassium peroxodisulfate (KPS) (99.0%), tert-butyl methyl ether (MtBE) (>99%) from Merck. Triethanolamine hydrochloride (TEA·HCl) (>99%) and sodium dodecylsulfate (SDS) were obtained from Fluka. All monomers and oils were used as received. Water for all purposes was doubly distilled Milli-Q-water. The buffer solution was TEA·HCl buffer with a concentration of 50 mM and pH 7. 2.2. Microgel synthesis The homo- and copolymers as shown in Table 1 were prepared by the same procedure. A 500 mL three neck vessel was equipped with a reflux condenser, an overhead stirrer and a nitrogen inand outlet. 160 mL water were purged with nitrogen and heated to 70 ◦ C. The monomer(s), the cross-linker, and – where needed – SDS were dissolved in water and added to the vessel. Afterwards, KPS was dissolved in degassed, cold water and added to the hot reaction solution. The reaction mixture became turbid after some minutes. The reaction continued at 70 ◦ C under nitrogen and constant stirring at 330 rpm for 5 h. Afterwards, the reaction mixture was cooled to room temperature under constant stirring. The core microgel was synthesized as stated above and cleaned by means of centrifugation (3 × 50.000 rpm, 30 min). The shell was grafted onto the core by the same synthesis routine. The reaction product was centrifuged (50.000 rpm, 30 min), the supernatant was removed,and the sediment was redispersed in
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Table 1 Microgel composition. NiPAM/g PNiPMAM PNiPAM P(NiPAM–NiPMAM) 50/50 PNiPAM–PNiPMAM CS 70/30 core PNiPAM–PNiPMAM CS 70/30 shell P(NiPAM–NiPMAM) 70/30
3.931 1.965 2.751 Purified corestock solution 2.751
NiPMAM/g
BIS/g
KPS/g
4.414
0.075 0.075 0.075 0.052 0.023 0.075
0.151 0.150 0.150 0.105 0.045 0.150
2.208 1.325 1.324
SDS/g
Water/mL
0.076 0.053 0.023 0.075
160 160 160 112 48(total: 160) 160
water; this procedure was repeated three times. For storage, the sediment was freeze dried. Prior to use the microgel was redispersed in the desired aqueous phase overnight. 2.3. Microgel characterization The microgel size was analyzed by means of dynamic light scattering. Therefore, an ALV-5000 device with a laser wavelength of 633 nm was used. The highly diluted aqueous (H2 O and TEA·HCl) samples were filtered with 5 m syringe filters to avoid dust. The measurements were performed for temperatures between 20 ◦ C and 50 ◦ C at a scattering angle of 60◦ . The hydrodynamic radii were calculated via the Stokes–Einstein equation from diffusion coefficients via cumulant analysis. For dynamic light scattering measurements in saturated aqueous phase (MtBE saturated TEA·HCl) the microgel was dissolved in buffer solution overnight. In a DLS cuvette the aqueous phase was covered with MtBE and equilibrated for 2 days. Afterwards the DLS measurement was performed in aqueous solution at 20, 32 and 42 ◦ C and angles of 45, 65, 75, 85, 95 and 105◦ . The hydrodynamic radii were calculated via Stokes–Einstein equation from the diffusion coefficient derived from CONTIN analysis. 2.4. Emulsion preparation and breakage MtBE and microgel in TEA·HCl buffer solution (50 mM, pH 7) were added together. The oil/water ratio was 1/1, the microgel amount 1 wt% in the aqueous phase. The mixing was performed with an Ultra Turrax T-25 equipped with a 10 mm head with 8000 rpm for 1 min. Emulsions were either heated in a thermomixer at different temperatures with a shaking of 200 rpm or analyzed with a multisample analytical centrifuge (LumiFuge, LUM). A LumiFuge with STEP technology was used to measure the intensity of transmitted light as a function of time and position over the entire sample length [32]. Centrifugation is possible at different speeds and temperatures, up to 40 ◦ C. At different temperatures, 600 rpm and a detection wavelength of 865 nm 120 profiles were taken in 20 min to detect the transmittance through 10 mm PA cuvettes. Region 1 and region 2 were analyzed independently in terms of integral transmittance as a function of time (cp Fig. 4). 3. Results and discussion 3.1. Microgels We synthesized microgels of different composition and structure, made of the two monomers NiPAM and NiPMAM. Two homopolymer and two copolymer microgels with a monomer ratio of 70/30 and 50/50 were prepared. Additionally, a core–shell microgel with a 70/30 NiPAM/NiPMAM ratio was synthesized in order to investigate the influence of the microgel morphology. The NiPAM–NiPMAM ratio affects the VPTT (Fig. 1) [33,34] [35–37]. In water all microgels show a temperature dependent swelling (see Fig. SI 1); the higher the NiPMAM amount the higher is the VPTT. Fig. 1 shows the influence of temperature on the hydro-
Fig. 1. Temperature-dependent hydrodynamic radius of microgel with different compositions and structures in TEA·HCl. The microgels start to aggregate reversibly at temperature above Tstab . Lines are guidance for the eye.
Table 2 Colloidal stability temperature (Tstab ) and volume phase transition temperature (VPTT) of the polymers in TEA·HCl or water, respectively.
PNiPAM P(NiPAM-co-NiPMAM) 70/30 PNiPAM-PNiPMAM CS 70/30 P(NiPAM-co-NiPMAM) 50/50 PNiPMAM
Tstab (TEA·HCl)/◦ C
VPTT (H2 O)/◦ C
34 36 40 40 46
32 35 33 and 41 38 42
dynamic radius of all microgels in TEA·HCl buffer. The microgels have the same size as in water at T < VPTT; thus, the swelling is not affected by the presence of the buffer. However, the buffer reduces the colloidal stability of the microgel above the VPTT. The ionic strength of the buffer reduces the electrostatic stabilization of the collapsed microgels, and they aggregate when heated above their corresponding VPTT. We define the temperature of colloidal stability (Tstab ) as the lowest measured temperature before aggregation is observed (see Fig. SI 2). The accuracy of Tstab is sufficient for our further investigations; the values are given in Table 2. The aggregation is reversible when the temperature is lowered. The core–shell microgel consists of a PNiPAM core and a PNiPMAM shell and reveals a different behavior as compared to the copolymer microgels. There are two swelling transitions in water (Fig. 2) that are due to the collapse of the PNiPAM core at ca. 32 ◦ C and of the PNiPMAM shell at 42 ◦ C [38]. In TEA·HCl buffer the microgel shrinks as well at 32 ◦ C because the core collapses, but stays colloidally stable. The microgel aggregates at temperatures above the VPTT of the shell (see Fig. 2). The core–shell morphology allows inducing a significant collapse of the microgel at temperatures where the microgel is still colloidally stable. The collapse of the copolymer microgels, however, directly leads to flocculation.
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Fig. 2. Temperature-dependent hydrodynamic radius of microgels ((core–shell PNiPAM–PNiPMAM) 70/30) in TEA·HCl-buffer (50 mM), water and with MtBE saturated TEA·HCl-buffer (50 mM). As expected the core–shell microgel measured in water shows two VPTTs, one for the NiPAM–core, the second for NiPMAM–shell. In TEA·HCl the behavior is different. One can also see two transitions, the first at the same temperature like in water, whereas the microgels aggregate when the shell collapses. In saturated buffer the microgels show the same behavior as in pure buffer but the transitions are shifted to lower temperatures. Lines are guidance for the eye.
Since in an emulsion the aqueous phase is saturated with organic phase we also analyze microgels in this environment. Measurements of microgels in TEA·HCl buffer saturated with MtBE reveals the presence of aggregates which were not observed in pure TEA·HCl buffer (not saturated with MtBE). The partial miscibility of MtBE in TEA·HCl buffer reduces the colloidal stability of the microgels. This effect does not influence the general trend but only the stability temperature at which the microgels start to aggregate (cp Fig. SI 6). 3.2. Preparation of emulsions The five microgels were employed as stabilizers for emulsions of MtBE in TEA·HCl buffer in a volume ratio of 1/1. The organic phase should be fully emulsified at room temperature but the emulsions should break on demand at elevated temperatures. A 1/1 mixture cannot be fully emulsified with 0.5 wt% microgel in the aqueous phase under the conditions of emulsification used in this study (see Fig. SI 3 and Section 2.4). Since the entire oil phase can then be dispersed, we used 1 wt% microgel in the aqueous phase in all further experiments (see Fig. 3a). We found no differences for emulsification at room temperature whether we used TEA·HCl buffer or pure water as aqueous phase. Since microgels are colloidally stable in both solvents at room temperature this behavior can be expected (Figs. Fig. 11 and SI 1). 3.3. Emulsions: breaking properties We continue to analyze the breaking behavior of the emulsions. Therefore, we first heated the emulsions under slight shaking in a Thermomixer to 50 ◦ C, which is well above Tstab of the microgels. All MtBE/TEA·HCl emulsions break into three phases as shown in Fig. 3b. The top phase is pure MtBE, the bottom phase is TEA·HCl buffer. Aggregated microgel is located between both phases. Reemulsification is possible after cooling down to room temperature. Since microgels behave differently in water and TEA·HCl above their VPTT/Tstab , we also studied the influence of the presence of buffer in aqueous phase on the emulsion breaking behavior. Emulsions prepared with water as aqueous phase break into two phases,
see Fig. 3c: a turbid water-microgel phase on the bottom and a clear MtBE phase on top. As described above, emulsions prepared with TEA·HCl buffer as aqueous phase break into two clear phases with a third phase containing collapsed aggregated microgel in between. The colloidal instability of microgel particles above their Tstab causes microgel aggregation in the broken emulsion. An analytical centrifuge (LumiFuge) with spatial and temporal resolution allows analyzing the breaking process in more detail. The transmittance was detected over the entire cuvette length during the centrifugation, revealing different effects like creaming or phase separation. Fig. 4 shows results of an analytical centrifuge measurement. Different colors of the transmittance profiles indicate the time dependence: red profiles were taken first, blue ones at last. The centrifugation experiments start with fully emulsified samples with an oil water volume ratio of 1/1. Thus, a complete breakage of the emulsions will lead to two separated phases of equal volume. Region 1 near the top of the cuvette will contain the oil phase and region 2 the aqueous phase at the bottom of the cuvette. In the top of the cuvette, there is air, which results in a high transmittance in this region (∼105–110 nm) Fig. 4 left summarizes the results of the centrifugation experiment at 30 ◦ C. The first profiles (shown in red) reveal a low transmittance over the whole sample range due to turbid, stable emulsion. The blue profiles taken after 20 min reveal a minor increase of transmittance in the bottom part of the cell due to slight creaming of oil droplets. In a second experiment we treated the emulsion at 35 ◦ C (Fig. 4, middle). In the beginning the transmittance was again low (red profiles) over the whole sample range. However, with time the transmittance increased in region 1 (∼112–120 mm) whereas it stays low in region 2 (120–130 mm). The oil is spilled out, and a transparent, macroscopic oil phase is formed in region 1. The volume of region 1 agrees with the amount of oil used in the first place; thus the emulsion is fully broken. Accordingly, region 2 contains the aqueous phase, and the low transmittance in this region can be traced back to the high concentration of dissolved microgel. At 40 ◦ C (Fig. 4, right) the first profiles show again low transmittance (red profiles) over the whole sample range. Like at 35 ◦ C, the transmittance increased with time in region 1 (∼ 112–118 mm); a macroscopic oil phase was built. In contrast to 35 ◦ C, the transmittance increases also in region 2 with time. In between the two regions with high transmittance there is a small region from 118–120 mm with low transmittance. When looking at the cuvette after finishing the experiment, collapsed, aggregated microgel is located here. Since the microgel is no longer dissolved in the aqueous phase, this phase shows high transmittance (region 2). The spatial variation of transmittance detected at the end of the centrifugation measurements is thus in perfect agreement with the results from the thermomixer experiments shown in Fig. 3. In order to compare in more detail the stabilizing behavior of two microgels with the same Tstab , but different structure, we did an integral transmission analysis separately for region 1 and region 2. The integral transmission, which is the area under the profile, correlates to the clearing in this region. The higher the integral transmission value, the clearer is the sample. The used copolymer P(NiPAM-co-NiPMAM) 50/50 and the core–shell microgel 70/30 both had a Tstab of 40 ◦ C (cp. Fig. 1 and Table 2). First, we compare the emulsion behavior in region 1 (top of the cuvette) stabilized with either copolymer (Fig. 5a) or core–shell microgel (Fig. 5c). At 30 ◦ C, the integral transmission shows low values over time for both samples, indicating a stable emulsion at this temperature. In contrast, at 35 ◦ C both samples show increasing integral transmission values with time; both emulsions break at this temperature. Although the two microgels have a different
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Fig. 3. Pictures of a stable emulsion (a) and two broken emulsions with MtBE as organic phase and either TEA·HCl buffer solution as aqueous phase (b) or pure water (c). The emulsion prepared with TEA·HCl buffer solution and MtBE (b) breaks into three phases, clear MtBE is on top, clear TEA·HCl buffer solution on bottom and aggregated microgel between both phases. The emulsion prepared with pure water and MtBE breaks into two phases, clear MtBE phase on top and turbid microgel-TEA·HCl buffer dispersion on bottom. To break the emulsion they were treated for 30 min with 50 ◦ C at 200 rpm.
Fig. 4. Transmittance profiles (transmittance versus distance from the centrifuge center to positions in the cuvette) of microgel stabilized MtBE/TEA·HCl emulsions (P(NiPAMco-NiPMAM) 50/50) by analytical centrifuge: LumiFuge, 600 rpm, t = 20 min, 10 mm PA cells, spectra taken every 10 s; red profiles were taken first, blue ones at last. From left to right: 30 ◦ C, 35 ◦ C and 40 ◦ C. Each temperature was measured separately with new cuvettes. Peaks/fluctuations in the region of pure oil (region 1: ∼110–120 mm) are results of small droplets or microgel attached to the cuvette wall. High transmittance left of region 1 (∼105–110 nm) is due to air in the cuvette. (Profiles of core–shell microgel: Fig. SI 5). At 30 ◦ C the droplets are stabilized by swollen microgels, furthermore, free, swollen microgel stays in solution. At 35 ◦ C the microgel is collapsed, but still soluble in the aqueous phase. At 40 ◦ C the microgel aggregates at the oil water interface. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
structure, both stabilize emulsions at 30 ◦ C whereas they allow a breaking at 35 ◦ C. Second, we compare the emulsion behavior in region 2 (bottom of the cuvette) stabilized with either copolymer (Fig. 5b) or core–shell microgel (Fig. 5d). At 30 ◦ C as well as at 35 ◦ C the integral transmission values for both samples stay low over time. Including the results of region 1 the emulsion stays stable at 30 ◦ C, whereas at 35 ◦ C the emulsions breaks, while both microgels are colloidally stable in the aqueous phase. At 37 ◦ C the integral transmission values for region 2 of a copolymer stabilized emulsions increase with time; the emulsion breaks and the microgel aggregates. The integral transmission values for region 2 of a core–shell microgel stabilized emulsions stays low over time. At 40 ◦ C the microgel is still colloidally stable and dispersed in the aqueous phase. Concluding the results, an emulsion stabilized with copolymer microgel (Tstab of 40 ◦ C) is stable at 30 ◦ C, breaks at 35 ◦ C into a macroscopic oil phase and an aqueous microgel phase, and at 37 ◦ C as well as at 40 ◦ C beside macroscopic phase separation the microgel aggregates and therefore, the aqueous phase clears. The
behavior of a core–shell microgel stabilized emulsion (Tstab of 40 ◦ C) is the same as for the copolymer stabilized emulsion, looking at region 1. At 30 ◦ C the emulsion is stable, at 35 ◦ C oil is spilled out. Yet in region 2 no clearing could be detected, the microgel stays dissolved in the aqueous phase. The higher colloidal stability of the core–shell microgel as compared to the copolymer microgel is in good agreement with the stability of the emulsions as probed by the analytical centrifuge (cp Fig. SI 4). Thus, determining the colloidal stability of the microgels in MtBE saturated TEA·HCl buffer allows predicting the temperature where the emulsions break. 4. Conclusion In this study we show that the switching behavior of the NiPAM–NiPMAM microgels and of the microgel stabilized MtBE/TEA·HCl emulsions correlate with each other. Furthermore, we can predict emulsions breaking behavior from microgel properties measured in aqueous solution.
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Fig. 5. Analysis of the oil-phase (region 1) (a, c) and aqueous phase (region 2) (b, d) from transmittance profiles taken by analytical centrifuge. Plotted is the area under the transmittance profiles (integral transmittance, y-axis in all graphs) versus the time. Upper row (a, b): copolymer microgel P(NiPAM-co-NiPMAM) 50/50, lower row (c, d): core–shell microgel. (䊐): 30 ◦ C (): 35 ◦ C (): 37 ◦ C () 40 ◦ C.
NiPAM–NiPMAM based microgels were synthesized and used for emulsion stabilization. The VPTT of microgels can be adjusted by varying the molar ratio of the two monomers NiPAM and NiPMAM between 32 ◦ C and 42 ◦ C—the VPTTs of the two homopolymers [33–36,39]. The core–shell microgel shows both VPTTs, for the core as well as for the shell [38]. TEA–salt influences the microgels’ behavior and thus, the emulsion breaking process with temperature [40]. The microgels aggregate reversibly in TEA·HCl buffer solution at a temperature Tstab . The core–shell morphology allows inducing a significant collapse of the microgel at temperatures between the core- and shell transition temperature while the microgel is still colloidally stable. The same trend but with a shift to lower temperatures is observed in MtBE saturated TEA·HCl buffer solution. Experiments with an analytical centrifuge show that emulsions break at temperatures slightly below the Tstab . At these temperatures emulsions break into two phases, a clear oil phase and a turbid, aqueous phase containing microgel. This result shows that the breaking of the emulsions does not depend on a colloidal instability of the microgels. This observation agrees with previous studies reporting that microgels adsorb to the oil–water interface even at temperatures above the VPTT. [27,41,42] It has been suggested that changes in the viscoelastic properties of the microgel covered interface are relevant for droplet coalescence and emulsion breakage [43–45]. This observation is in good agreement with
previous studies. Brugger et al. [27] and Destribats et al. [46] found that heptane/water emulsions break at temperatures well above the microgels VPTT. For copolymer microgels flocculation of the microgel was obtained while breaking the emulsion. The core–shell microgel stays dispersed in the aqueous phase above Tstab . The temperature interval in which the emulsions are broken but the microgel stays dispersed is broader for core–shell microgels than for copolymers. The behavior of the microgels in aqueous solution when the partial miscibility of the organic phase with the aqueous phase is taken into account allows predicting the temperature at which the emulsion breaks and whether microgels flocculate or not during breaking process. The microgels need to deswell to about 55% as compared to its swollen size at room temperature to let the emulsion break. The copolymer microgel aggregates during the process of collapsing. Meanwhile the core–shell microgel aggregates from the plateau size between the two transition temperatures to larger entities. This special behavior allows adapting microgels to the requirements of biocatalysis in emulsion. A possible industrial application could be to control an oil release or a separation of oil as well as microgel from an emulsion. Enzymes can be protected during catalytic activity and afterwards separated and reused from the reaction system.
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Contents lists available at ScienceDirect
Colloids and Surfaces A: Physicochemical and Engineering Aspects journal homepage: www.elsevier.com/locate/colsurfa
Microgel stabilized emulsions: Breaking on demand Susanne Wiese a , Yoanna Tsvetkova a,1 , Nadine J.E. Daleiden a , Antje C. Spieß b,c,d , Walter Richtering a,∗ a
RWTH Aachen University, Physical Chemistry, Landoltweg 1, 52056 Aachen, Germany RWTH Aachen University, AVT —Enzyme Process Technology, Aachen, Worringerweg 1, 52074 Aachen, Germany DWI—Leibniz-Institute for Interactive Materials Research, Forckenbeckstraße 50, 52074 Aachen, Germany dTU Braunschweig, Institute for Biochemical Engineering ibvt, Gaußstr. 17, 38102 Braunschweig, Germany d TU Braunschweig, Institute for Biochemical Engineering ibvt, Gaußstr. 17, 38102 Braunschweig, Germany b c
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
• Microgels are employed to stabilize and break emulsions on demand.
• Microgel composition and structure influence temperature sensitivity of emulsions. • We control whether microgel stays in solution or aggregates while breaking the emulsion. • Microgels as reversible emulsion stabilizers can be adopted to requirements of enzyme reactions in biocatalysis.
a r t i c l e
i n f o
Article history: Received 28 July 2015 Received in revised form 2 February 2016 Accepted 3 February 2016 Available online 17 February 2016 Keywords: Switchable microgel Switchable emulsion Biocatalysis Poly-N-isopropyl-acrylamideco-N-isopropyl-methacrylamide
a b s t r a c t Here, we report on how to stabilize and break emulsions that are compatible with enzymatic reaction conditions. Many substrates of enzymatic reactions are soluble in unpolar organic solvents whereas the enzymes themselves often need an aqueous environment. We use a buffer solution (triethanolamine hydrochloride) as aqueous and MtBE (tert-butyl methyl ether) as organic phase which provide good enzyme compatibility. We are able to break emulsions in a desired temperature range by using NiPAM–NiPMAM microgels with different monomer compositions and architecture, respectively. Our microgels need to deswell to about 55% of its swollen size at room temperature to let the emulsion break. Emulsions can be broken such that the microgels are either colloidally stable in the aqueous phase or flocculated. The temperature interval in which the microgels stay colloidally stable while the emulsion is broken is broader for the core–shell microgel than for the copolymer microgel. The behavior of the microgels in aqueous solution allows predicting: (i) the temperature at which the emulsion breaks and (ii) whether microgels flocculate or not during breaking the emulsion. However, the partial miscibility of the organic phase with the aqueous phase has to be taken into account.
∗ Corresponding author. Fax: +49 214 80 92327. E-mail addresses: [email protected] (S. Wiese), [email protected] (Y. Tsvetkova), [email protected] (N.J.E. Daleiden), [email protected] (A.C. Spieß), [email protected] (W. Richtering). 1 UK Aachen, Institut für Biomedizinische Technologien—Experimentelle Molekulare Bildgebung, Pauwelsstraße 30, 52074 Aachen, Germany. http://dx.doi.org/10.1016/j.colsurfa.2016.02.003 0927-7757/© 2016 Elsevier B.V. All rights reserved.
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Thus, we are able to stabilize and break emulsions by employing microgels as responsive emulsifiers and to adapt the microgels to the requirements of biocatalytic processes. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Many substrates of enzymatic reactions are soluble in organic solvents whereas the enzymes themselves often need an aqueous environment [1]. When performing enzyme reactions in industrial scale a large reservoir of substrates is needed that leads to a two phase system, the aqueous phase containing the enzyme and an organic phase containing the substrates and eventually products. Surfaces as well as interfaces may lead to enzyme unfolding and therefore, enzyme inactivation [2]. One approach to protect enzymes from denaturation at the interface is to introduce microgels as surface-active species. Microgels allow for emulsion stabilization [3]. Emulsions are systems of two immiscible liquids, mostly an aqueous phase and a non-aqueous phase, a so called organic or oil phase. Surfactants [4], proteins [5], star-copolymers [6] or nanoparticles [7,8] stabilize droplets of one liquid (dispersed phase) dispersed in the other liquid (continuous phase). A new class of stabilizing agents are microgels [3]. Microgels are soft cross-linked, solvent swollen polymer particles [9,10]. Microgel stabilized emulsions have great potential for industrial application. e.g., in drug delivery [11] or biocatalysis [12,13]. One special property of some microgels is their sensitivity to external stimuli [14–22]. For example N-isopropylacrylamide (NiPAM)-based microgels undergo a volume change with increasing temperature [14]. The VPTT (volume phase transition temperature) can be adjusted by using different co-monomers. The switchability of the microgels leads to switchability of the emulsions [3,23]. Since the enzyme requires specific conditions with respect to reaction temperature and stability temperature the emulsion needs to be switched in a certain temperature range. Furthermore, the stability and reactivity of the enzyme depends on the pH value and the used solvents. In this study we use Lactobacillus brevis alcohol dehydrogenase (Lb-ADH) as model enzyme whose optimal pH value is maintained using the buffer TEA·HCl (triethanolamine hydrochloride). TEA·HCl buffer has the advantage that the temperature has only a moderate influence on the pH value (pK = −0.02 pH/K) [24]. We introduce tert-butyl methyl ether (MtBE) [25] as organic phase due to its good enzyme compatibility [26]. MtBE is a polar and aprotic organic solvent and thus different from heptane and octanol which have previously been used as organic phase in microgel stabilized emulsions [23,27,28]. In general enzymes denature with rising temperatures. Therefore, microgels used in emulsions for biocatalysis: (i) must be stable in the buffer, (ii) must stabilize an emulsion of an polar, aprotic organic phase with a buffer, (iii) must stabilize these emulsions at the optimal enzymes’ working temperature (in our case 30 ◦ C), (iv) break the emulsion at a temperature above 30 ◦ C, which however is yet low enough to avoid enzyme denaturation. Mostly P(NiPAM-co-MAA) microgels (poly-(N-isopropylacrylamide-co-methacrylic acid)) were used to stabilize emulsions but in recent studies also uncharged microgels were used [25,28].
P(NiPAM-co-MAA) microgel-stabilized emulsions can be broken by increasing the temperature or by changing the pH value [23,27,29]. We use microgels made of two uncharged monomers, NiPAM and NiPMAM (N-isopropyl-methacrylamide), to stabilize emulsions. The general structure of both monomers is quite similar; they differ only in a methyl group at the polymer backbone. The VPTT of pure PNiPAM microgels is ca. 30–32 ◦ C [9,30] whereas the VPTT of pure PNiPMAM is ca. 40–43 ◦ C [30,31]. With these two monomers we now create microgels with different composition and structure. Our aim is to stabilize and break emulsions on demand. In this study we show that the switching behavior of microgels and emulsions correlate with each other. We found that emulsions, stabilized with NiPAM–NiPMAM-microgels stabilize emulsions at room temperature and can be broken at temperatures slightly above their microgels’ transitions temperatures. We are able to adapt microgels to the requirements of enzymatic reactions, thus emulsions can be stabilized and be broken on demand in a desired temperature range. Furthermore, we can predict emulsion behavior from microgel properties measured in aqueous solution. Emulsions stabilized with the core–shell microgel break already when the temperature is between the transition temperatures of core and shell, respectively. In other words the emulsion breaks when the stabilizer is not fully collapsed but partially swollen. This particular behavior opens a wide field of possible applications. 2. Material and methods 2.1. Chemicals N-Isopropylacrylamide (NiPAM) (>99.0%) and the cross-linker N,N -methylenebisacrylamide (BIS) (>99.5%) were purchased from Acros Chemicals; N-isopropylmethacrylamide (NiPMAM) (>99%) from Aldrich; the initiator potassium peroxodisulfate (KPS) (99.0%), tert-butyl methyl ether (MtBE) (>99%) from Merck. Triethanolamine hydrochloride (TEA·HCl) (>99%) and sodium dodecylsulfate (SDS) were obtained from Fluka. All monomers and oils were used as received. Water for all purposes was doubly distilled Milli-Q-water. The buffer solution was TEA·HCl buffer with a concentration of 50 mM and pH 7. 2.2. Microgel synthesis The homo- and copolymers as shown in Table 1 were prepared by the same procedure. A 500 mL three neck vessel was equipped with a reflux condenser, an overhead stirrer and a nitrogen inand outlet. 160 mL water were purged with nitrogen and heated to 70 ◦ C. The monomer(s), the cross-linker, and – where needed – SDS were dissolved in water and added to the vessel. Afterwards, KPS was dissolved in degassed, cold water and added to the hot reaction solution. The reaction mixture became turbid after some minutes. The reaction continued at 70 ◦ C under nitrogen and constant stirring at 330 rpm for 5 h. Afterwards, the reaction mixture was cooled to room temperature under constant stirring. The core microgel was synthesized as stated above and cleaned by means of centrifugation (3 × 50.000 rpm, 30 min). The shell was grafted onto the core by the same synthesis routine. The reaction product was centrifuged (50.000 rpm, 30 min), the supernatant was removed,and the sediment was redispersed in
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Table 1 Microgel composition. NiPAM/g PNiPMAM PNiPAM P(NiPAM–NiPMAM) 50/50 PNiPAM–PNiPMAM CS 70/30 core PNiPAM–PNiPMAM CS 70/30 shell P(NiPAM–NiPMAM) 70/30
3.931 1.965 2.751 Purified corestock solution 2.751
NiPMAM/g
BIS/g
KPS/g
4.414
0.075 0.075 0.075 0.052 0.023 0.075
0.151 0.150 0.150 0.105 0.045 0.150
2.208 1.325 1.324
SDS/g
Water/mL
0.076 0.053 0.023 0.075
160 160 160 112 48(total: 160) 160
water; this procedure was repeated three times. For storage, the sediment was freeze dried. Prior to use the microgel was redispersed in the desired aqueous phase overnight. 2.3. Microgel characterization The microgel size was analyzed by means of dynamic light scattering. Therefore, an ALV-5000 device with a laser wavelength of 633 nm was used. The highly diluted aqueous (H2 O and TEA·HCl) samples were filtered with 5 m syringe filters to avoid dust. The measurements were performed for temperatures between 20 ◦ C and 50 ◦ C at a scattering angle of 60◦ . The hydrodynamic radii were calculated via the Stokes–Einstein equation from diffusion coefficients via cumulant analysis. For dynamic light scattering measurements in saturated aqueous phase (MtBE saturated TEA·HCl) the microgel was dissolved in buffer solution overnight. In a DLS cuvette the aqueous phase was covered with MtBE and equilibrated for 2 days. Afterwards the DLS measurement was performed in aqueous solution at 20, 32 and 42 ◦ C and angles of 45, 65, 75, 85, 95 and 105◦ . The hydrodynamic radii were calculated via Stokes–Einstein equation from the diffusion coefficient derived from CONTIN analysis. 2.4. Emulsion preparation and breakage MtBE and microgel in TEA·HCl buffer solution (50 mM, pH 7) were added together. The oil/water ratio was 1/1, the microgel amount 1 wt% in the aqueous phase. The mixing was performed with an Ultra Turrax T-25 equipped with a 10 mm head with 8000 rpm for 1 min. Emulsions were either heated in a thermomixer at different temperatures with a shaking of 200 rpm or analyzed with a multisample analytical centrifuge (LumiFuge, LUM). A LumiFuge with STEP technology was used to measure the intensity of transmitted light as a function of time and position over the entire sample length [32]. Centrifugation is possible at different speeds and temperatures, up to 40 ◦ C. At different temperatures, 600 rpm and a detection wavelength of 865 nm 120 profiles were taken in 20 min to detect the transmittance through 10 mm PA cuvettes. Region 1 and region 2 were analyzed independently in terms of integral transmittance as a function of time (cp Fig. 4). 3. Results and discussion 3.1. Microgels We synthesized microgels of different composition and structure, made of the two monomers NiPAM and NiPMAM. Two homopolymer and two copolymer microgels with a monomer ratio of 70/30 and 50/50 were prepared. Additionally, a core–shell microgel with a 70/30 NiPAM/NiPMAM ratio was synthesized in order to investigate the influence of the microgel morphology. The NiPAM–NiPMAM ratio affects the VPTT (Fig. 1) [33,34] [35–37]. In water all microgels show a temperature dependent swelling (see Fig. SI 1); the higher the NiPMAM amount the higher is the VPTT. Fig. 1 shows the influence of temperature on the hydro-
Fig. 1. Temperature-dependent hydrodynamic radius of microgel with different compositions and structures in TEA·HCl. The microgels start to aggregate reversibly at temperature above Tstab . Lines are guidance for the eye.
Table 2 Colloidal stability temperature (Tstab ) and volume phase transition temperature (VPTT) of the polymers in TEA·HCl or water, respectively.
PNiPAM P(NiPAM-co-NiPMAM) 70/30 PNiPAM-PNiPMAM CS 70/30 P(NiPAM-co-NiPMAM) 50/50 PNiPMAM
Tstab (TEA·HCl)/◦ C
VPTT (H2 O)/◦ C
34 36 40 40 46
32 35 33 and 41 38 42
dynamic radius of all microgels in TEA·HCl buffer. The microgels have the same size as in water at T < VPTT; thus, the swelling is not affected by the presence of the buffer. However, the buffer reduces the colloidal stability of the microgel above the VPTT. The ionic strength of the buffer reduces the electrostatic stabilization of the collapsed microgels, and they aggregate when heated above their corresponding VPTT. We define the temperature of colloidal stability (Tstab ) as the lowest measured temperature before aggregation is observed (see Fig. SI 2). The accuracy of Tstab is sufficient for our further investigations; the values are given in Table 2. The aggregation is reversible when the temperature is lowered. The core–shell microgel consists of a PNiPAM core and a PNiPMAM shell and reveals a different behavior as compared to the copolymer microgels. There are two swelling transitions in water (Fig. 2) that are due to the collapse of the PNiPAM core at ca. 32 ◦ C and of the PNiPMAM shell at 42 ◦ C [38]. In TEA·HCl buffer the microgel shrinks as well at 32 ◦ C because the core collapses, but stays colloidally stable. The microgel aggregates at temperatures above the VPTT of the shell (see Fig. 2). The core–shell morphology allows inducing a significant collapse of the microgel at temperatures where the microgel is still colloidally stable. The collapse of the copolymer microgels, however, directly leads to flocculation.
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Fig. 2. Temperature-dependent hydrodynamic radius of microgels ((core–shell PNiPAM–PNiPMAM) 70/30) in TEA·HCl-buffer (50 mM), water and with MtBE saturated TEA·HCl-buffer (50 mM). As expected the core–shell microgel measured in water shows two VPTTs, one for the NiPAM–core, the second for NiPMAM–shell. In TEA·HCl the behavior is different. One can also see two transitions, the first at the same temperature like in water, whereas the microgels aggregate when the shell collapses. In saturated buffer the microgels show the same behavior as in pure buffer but the transitions are shifted to lower temperatures. Lines are guidance for the eye.
Since in an emulsion the aqueous phase is saturated with organic phase we also analyze microgels in this environment. Measurements of microgels in TEA·HCl buffer saturated with MtBE reveals the presence of aggregates which were not observed in pure TEA·HCl buffer (not saturated with MtBE). The partial miscibility of MtBE in TEA·HCl buffer reduces the colloidal stability of the microgels. This effect does not influence the general trend but only the stability temperature at which the microgels start to aggregate (cp Fig. SI 6). 3.2. Preparation of emulsions The five microgels were employed as stabilizers for emulsions of MtBE in TEA·HCl buffer in a volume ratio of 1/1. The organic phase should be fully emulsified at room temperature but the emulsions should break on demand at elevated temperatures. A 1/1 mixture cannot be fully emulsified with 0.5 wt% microgel in the aqueous phase under the conditions of emulsification used in this study (see Fig. SI 3 and Section 2.4). Since the entire oil phase can then be dispersed, we used 1 wt% microgel in the aqueous phase in all further experiments (see Fig. 3a). We found no differences for emulsification at room temperature whether we used TEA·HCl buffer or pure water as aqueous phase. Since microgels are colloidally stable in both solvents at room temperature this behavior can be expected (Figs. Fig. 11 and SI 1). 3.3. Emulsions: breaking properties We continue to analyze the breaking behavior of the emulsions. Therefore, we first heated the emulsions under slight shaking in a Thermomixer to 50 ◦ C, which is well above Tstab of the microgels. All MtBE/TEA·HCl emulsions break into three phases as shown in Fig. 3b. The top phase is pure MtBE, the bottom phase is TEA·HCl buffer. Aggregated microgel is located between both phases. Reemulsification is possible after cooling down to room temperature. Since microgels behave differently in water and TEA·HCl above their VPTT/Tstab , we also studied the influence of the presence of buffer in aqueous phase on the emulsion breaking behavior. Emulsions prepared with water as aqueous phase break into two phases,
see Fig. 3c: a turbid water-microgel phase on the bottom and a clear MtBE phase on top. As described above, emulsions prepared with TEA·HCl buffer as aqueous phase break into two clear phases with a third phase containing collapsed aggregated microgel in between. The colloidal instability of microgel particles above their Tstab causes microgel aggregation in the broken emulsion. An analytical centrifuge (LumiFuge) with spatial and temporal resolution allows analyzing the breaking process in more detail. The transmittance was detected over the entire cuvette length during the centrifugation, revealing different effects like creaming or phase separation. Fig. 4 shows results of an analytical centrifuge measurement. Different colors of the transmittance profiles indicate the time dependence: red profiles were taken first, blue ones at last. The centrifugation experiments start with fully emulsified samples with an oil water volume ratio of 1/1. Thus, a complete breakage of the emulsions will lead to two separated phases of equal volume. Region 1 near the top of the cuvette will contain the oil phase and region 2 the aqueous phase at the bottom of the cuvette. In the top of the cuvette, there is air, which results in a high transmittance in this region (∼105–110 nm) Fig. 4 left summarizes the results of the centrifugation experiment at 30 ◦ C. The first profiles (shown in red) reveal a low transmittance over the whole sample range due to turbid, stable emulsion. The blue profiles taken after 20 min reveal a minor increase of transmittance in the bottom part of the cell due to slight creaming of oil droplets. In a second experiment we treated the emulsion at 35 ◦ C (Fig. 4, middle). In the beginning the transmittance was again low (red profiles) over the whole sample range. However, with time the transmittance increased in region 1 (∼112–120 mm) whereas it stays low in region 2 (120–130 mm). The oil is spilled out, and a transparent, macroscopic oil phase is formed in region 1. The volume of region 1 agrees with the amount of oil used in the first place; thus the emulsion is fully broken. Accordingly, region 2 contains the aqueous phase, and the low transmittance in this region can be traced back to the high concentration of dissolved microgel. At 40 ◦ C (Fig. 4, right) the first profiles show again low transmittance (red profiles) over the whole sample range. Like at 35 ◦ C, the transmittance increased with time in region 1 (∼ 112–118 mm); a macroscopic oil phase was built. In contrast to 35 ◦ C, the transmittance increases also in region 2 with time. In between the two regions with high transmittance there is a small region from 118–120 mm with low transmittance. When looking at the cuvette after finishing the experiment, collapsed, aggregated microgel is located here. Since the microgel is no longer dissolved in the aqueous phase, this phase shows high transmittance (region 2). The spatial variation of transmittance detected at the end of the centrifugation measurements is thus in perfect agreement with the results from the thermomixer experiments shown in Fig. 3. In order to compare in more detail the stabilizing behavior of two microgels with the same Tstab , but different structure, we did an integral transmission analysis separately for region 1 and region 2. The integral transmission, which is the area under the profile, correlates to the clearing in this region. The higher the integral transmission value, the clearer is the sample. The used copolymer P(NiPAM-co-NiPMAM) 50/50 and the core–shell microgel 70/30 both had a Tstab of 40 ◦ C (cp. Fig. 1 and Table 2). First, we compare the emulsion behavior in region 1 (top of the cuvette) stabilized with either copolymer (Fig. 5a) or core–shell microgel (Fig. 5c). At 30 ◦ C, the integral transmission shows low values over time for both samples, indicating a stable emulsion at this temperature. In contrast, at 35 ◦ C both samples show increasing integral transmission values with time; both emulsions break at this temperature. Although the two microgels have a different
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Fig. 3. Pictures of a stable emulsion (a) and two broken emulsions with MtBE as organic phase and either TEA·HCl buffer solution as aqueous phase (b) or pure water (c). The emulsion prepared with TEA·HCl buffer solution and MtBE (b) breaks into three phases, clear MtBE is on top, clear TEA·HCl buffer solution on bottom and aggregated microgel between both phases. The emulsion prepared with pure water and MtBE breaks into two phases, clear MtBE phase on top and turbid microgel-TEA·HCl buffer dispersion on bottom. To break the emulsion they were treated for 30 min with 50 ◦ C at 200 rpm.
Fig. 4. Transmittance profiles (transmittance versus distance from the centrifuge center to positions in the cuvette) of microgel stabilized MtBE/TEA·HCl emulsions (P(NiPAMco-NiPMAM) 50/50) by analytical centrifuge: LumiFuge, 600 rpm, t = 20 min, 10 mm PA cells, spectra taken every 10 s; red profiles were taken first, blue ones at last. From left to right: 30 ◦ C, 35 ◦ C and 40 ◦ C. Each temperature was measured separately with new cuvettes. Peaks/fluctuations in the region of pure oil (region 1: ∼110–120 mm) are results of small droplets or microgel attached to the cuvette wall. High transmittance left of region 1 (∼105–110 nm) is due to air in the cuvette. (Profiles of core–shell microgel: Fig. SI 5). At 30 ◦ C the droplets are stabilized by swollen microgels, furthermore, free, swollen microgel stays in solution. At 35 ◦ C the microgel is collapsed, but still soluble in the aqueous phase. At 40 ◦ C the microgel aggregates at the oil water interface. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
structure, both stabilize emulsions at 30 ◦ C whereas they allow a breaking at 35 ◦ C. Second, we compare the emulsion behavior in region 2 (bottom of the cuvette) stabilized with either copolymer (Fig. 5b) or core–shell microgel (Fig. 5d). At 30 ◦ C as well as at 35 ◦ C the integral transmission values for both samples stay low over time. Including the results of region 1 the emulsion stays stable at 30 ◦ C, whereas at 35 ◦ C the emulsions breaks, while both microgels are colloidally stable in the aqueous phase. At 37 ◦ C the integral transmission values for region 2 of a copolymer stabilized emulsions increase with time; the emulsion breaks and the microgel aggregates. The integral transmission values for region 2 of a core–shell microgel stabilized emulsions stays low over time. At 40 ◦ C the microgel is still colloidally stable and dispersed in the aqueous phase. Concluding the results, an emulsion stabilized with copolymer microgel (Tstab of 40 ◦ C) is stable at 30 ◦ C, breaks at 35 ◦ C into a macroscopic oil phase and an aqueous microgel phase, and at 37 ◦ C as well as at 40 ◦ C beside macroscopic phase separation the microgel aggregates and therefore, the aqueous phase clears. The
behavior of a core–shell microgel stabilized emulsion (Tstab of 40 ◦ C) is the same as for the copolymer stabilized emulsion, looking at region 1. At 30 ◦ C the emulsion is stable, at 35 ◦ C oil is spilled out. Yet in region 2 no clearing could be detected, the microgel stays dissolved in the aqueous phase. The higher colloidal stability of the core–shell microgel as compared to the copolymer microgel is in good agreement with the stability of the emulsions as probed by the analytical centrifuge (cp Fig. SI 4). Thus, determining the colloidal stability of the microgels in MtBE saturated TEA·HCl buffer allows predicting the temperature where the emulsions break. 4. Conclusion In this study we show that the switching behavior of the NiPAM–NiPMAM microgels and of the microgel stabilized MtBE/TEA·HCl emulsions correlate with each other. Furthermore, we can predict emulsions breaking behavior from microgel properties measured in aqueous solution.
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Fig. 5. Analysis of the oil-phase (region 1) (a, c) and aqueous phase (region 2) (b, d) from transmittance profiles taken by analytical centrifuge. Plotted is the area under the transmittance profiles (integral transmittance, y-axis in all graphs) versus the time. Upper row (a, b): copolymer microgel P(NiPAM-co-NiPMAM) 50/50, lower row (c, d): core–shell microgel. (䊐): 30 ◦ C (): 35 ◦ C (): 37 ◦ C () 40 ◦ C.
NiPAM–NiPMAM based microgels were synthesized and used for emulsion stabilization. The VPTT of microgels can be adjusted by varying the molar ratio of the two monomers NiPAM and NiPMAM between 32 ◦ C and 42 ◦ C—the VPTTs of the two homopolymers [33–36,39]. The core–shell microgel shows both VPTTs, for the core as well as for the shell [38]. TEA–salt influences the microgels’ behavior and thus, the emulsion breaking process with temperature [40]. The microgels aggregate reversibly in TEA·HCl buffer solution at a temperature Tstab . The core–shell morphology allows inducing a significant collapse of the microgel at temperatures between the core- and shell transition temperature while the microgel is still colloidally stable. The same trend but with a shift to lower temperatures is observed in MtBE saturated TEA·HCl buffer solution. Experiments with an analytical centrifuge show that emulsions break at temperatures slightly below the Tstab . At these temperatures emulsions break into two phases, a clear oil phase and a turbid, aqueous phase containing microgel. This result shows that the breaking of the emulsions does not depend on a colloidal instability of the microgels. This observation agrees with previous studies reporting that microgels adsorb to the oil–water interface even at temperatures above the VPTT. [27,41,42] It has been suggested that changes in the viscoelastic properties of the microgel covered interface are relevant for droplet coalescence and emulsion breakage [43–45]. This observation is in good agreement with
previous studies. Brugger et al. [27] and Destribats et al. [46] found that heptane/water emulsions break at temperatures well above the microgels VPTT. For copolymer microgels flocculation of the microgel was obtained while breaking the emulsion. The core–shell microgel stays dispersed in the aqueous phase above Tstab . The temperature interval in which the emulsions are broken but the microgel stays dispersed is broader for core–shell microgels than for copolymers. The behavior of the microgels in aqueous solution when the partial miscibility of the organic phase with the aqueous phase is taken into account allows predicting the temperature at which the emulsion breaks and whether microgels flocculate or not during breaking process. The microgels need to deswell to about 55% as compared to its swollen size at room temperature to let the emulsion break. The copolymer microgel aggregates during the process of collapsing. Meanwhile the core–shell microgel aggregates from the plateau size between the two transition temperatures to larger entities. This special behavior allows adapting microgels to the requirements of biocatalysis in emulsion. A possible industrial application could be to control an oil release or a separation of oil as well as microgel from an emulsion. Enzymes can be protected during catalytic activity and afterwards separated and reused from the reaction system.
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