Using Hydrogel Microparticles To Transfer Hydrophilic Nanoparticles and
Enzymes to Organic Media via Stepwise Solvent Exchange
Shuo Bai,†,^ Changzhu Wu,‡,^ Kornelia Gawlitza,§ Regine von Klitzing,§
Marion B. Ansorge-Schumacher,*,‡ and Dayang Wang*,†
†Max Planck Institute of Colloids and Interfaces, D-14424 Potsdam, Germany, ‡Department of EnzymeTechnology (TC4), Institute of Chemistry, Technical University of Berlin, Strasse des 17. Juni 124, D-10623
Berlin, Germany, and §Stranski-Laboratory for Physical and Theoretical Chemistry (TC9), Institute ofChemistry, Technical University of Berlin, Strasse des 17. Juni 124, D-10623 Berlin, Germany.
^These authors contributed equally to this paper
Received May 20, 2010. Revised Manuscript Received June 16, 2010
We present a simple and versatile approach of using hydrogel microparticles to transfer both inorganic hydrophilicnanoparticles (NPs) such asCdTe quantumdots and enzymes such as lipase B fromCandida antarctica (CalB) to organicmedia and eventually encapsulate them in the gel microparticles by consecutive exchange of the water swollen in thehydrogel microparticles with water-miscible organic solvents and water-immiscible solvents. The entrapment ofhydrophilic nanoparticles is due to their incompatibility with water-immiscible organic solvents soaked in the gelmatrices and in the surrounding environment, so the present approach obviates the need for any chemical modificationto the NP surface or to the hydrogel and furthermore does not require any size matching or chemical affinity of the NPsfor the hydrogel networks. The solvent exchange process causes little change of the intrinsic properties of hydrophilicnanoparticles; CdTe quantum dots encapsulated in hydrogel microparticles, dispersed in water-immiscible organicsolvents, remain strongly fluorescent, and CalB retains high catalytic activity. Of importance is that the hydrophilicnanoparticles encapsulated in the gel microparticles in organic media can be completely recovered in aqueous media viareversed solvent exchange. As a consequence, the present approach should hold immense promise for technicalapplications, especially in catalysis.
Introduction
Colloidal nanoparticles (NPs) ubiquitously exist in a dazzlingdiversity of chemical nature and morphology, such as slurries,inks, dusts, fogs, micelles, latex paints, enzymes, bacteria, anderythrocytes. They have been important research objects of bothfundamental science and industrial applications as in ceramics,information storage, catalysis, pigments, medicine, and manyothers in the past decades. According to the dispersion media,colloidal NPs can be simply classified into two categories: hydro-philic NPs dispersed in aqueous media and hydrophobic NPs inorganic media. Owing to easy and environmentally benignsynthesis protocols and easily accessible precursors, synthesis ofinorganic and organic NPs in aqueous media has been a scientificfascination for more than a century.1 Nevertheless, hydrophilicNPs usually are polycrystalline and polydisperse in size andshape. Their surfaces are usually not coated by additionalstabilizing ligands, thus leading to a poor passivation. Hence,there are surface defects that deteriorate the intrinsic properties ofhydrophilic NPs, which becomes particularly obvious whenquantum confinement such as photoluminescence is involved.Therefore, hydrophilic NPs are less promising in applicationssuch as nanoelectronics in which excellent quantum confinement
properties are crucial.2 On the other hand, the bare surfaces ofhydrophilicNPs being rich in surface defects can benefit a numberof applications in which high surface activity is the key issue, suchas sensing and catalysis.3 As a matter of fact, the use of hydro-philicNPs of noblemetals, transitionmetals, andmetal oxides forcatalysis of organic reactions has already been extensively studiedbefore the advent of nanoscience.
In order to adapt to different technical applications, NPs, bothhydrophilic and hydrophobic, must be able to be easily trans-ferred to a variety of environments without agglomeration anddeterioration of intrinsic and surface properties. Up to date,numerous methods have been successfully developed for phasetransfer of hydrophobicNPs to aqueousmedia in order to exploittheir biomedical applications.4 In contrast, coating with amphi-philicmolecules remains the onlyway to transfer hydrophilicNPsfrom aqueous to organic media with the intent of better surfacepassivation for the enhancement of the intrinsic properties of theNPs.5 As a result of surface passivation, the access of substratesto the surfaces of NPs is blocked, which is unsatisfactory in
*To whom correspondence should be addressed: e-mail [email protected], Fax þ49 331 5679202 (D.W.); e-mail [email protected], Fax þ49 30 31422261 (M.B.A.-S.).(1) (a) Matijivic, E. Langmuir 1994, 10, 8. (b) Horn, D.; Rieger, J. Angew. Chem.,
Int. Ed. 2001, 40, 4330. (c) Collier, C. P.; Vossmeyer, T.; Heath, J. R. Annu. Rev. Phys.Chem. 1998, 49, 371. (d) Murray, C. B.; Kagan, C. R.; Bawendi, M. G. Annu. Rev.Mater. Sci. 2000, 30, 545. (e) Burda, C.; Chen, X.; Narayanan, R.; El-Sayed, M.Chem.Rev. 2005, 105, 1025.(2) (a) Alivisatos, A. P. Science 1996, 271, 933. (b) Nirmal, M.; Brus, L. Acc.
(3) Nanoparticles and Catalysis; Astruc, D., Ed.; Wiley-VCH: Weinheim, 2007.(4) (a) Bruchez, M.; Moronne, M.; Gin, P.; Wiess, S.; Alivisatos, A. P. Science
1998, 281, 2013. (b) Chan, W. C.; Nie, S. Science 1998, 281, 2016. (c) Gerion, D.;Pinaud, F.;Williams, S. C.; Parak,W. J.; Zanchet, D.;Weiss, S.; Alivisatos, A. P. J. Phys.Chem. B 2001, 105, 8861. (d) Gittins, D. I.; Caruso, F. Angew. Chem., Int. Ed. 2001,40, 3001. (e) Dubertret, B.; Skourides, P.; Norris, D. J.; Noireaux, V.; Brivanlou, A. H.;Libchaber, A. Science 2002, 298, 1759. (f) Yu, W. W.; Chang, E.; Falkner, J. C.; Zhang,J.; Al-Somali, A. M.; Sayes, C. M.; Johns, J.; Drezek, R.; Colvin, V. L. J. Am. Chem.Soc. 2007, 129, 2871. (g) Lin, C.-A. J.; Sperling, A.; Li, J. K.; Yang, T.-Y.; Li, P.-Y.;Zanella, M.; Chang, W. H.; Parak, W. J. Small 2008, 334. (i) Lee, E. E.; Nguyen, T.-L.;Clayton, A. H. A.; Mulvaney, P. ACS Nano 2009, 3, 1121.
(5) (a) Sastry, M. Curr. Sci. 2003, 85, 1735. (b) Gaponik, N.; Talapin, D. V.;Rogach, A. L.; Eychmueller, A.; Weller, H. Nano Lett. 2002, 2, 803. (c) Duan, H.;Kuang, M.; Wang, D.; Kurth, D.; M€ohwald, H.Angew. Chem., Int. Ed. 2005, 44, 1717.
applications such as catalysis. Up to date, to our best knowledge,there is no methodology allowing transfer of hydrophilic NPs toorganic media without surface modification. Crooks et al. haverecently synthesized hydrophilic metallic NPs inside dendrimersand created dendrimer-encapsulated metallic NPs.6 They havedemonstrated that the dendrimers can efficiently stabilize themetallic NPs. At the same time, their nanoporous structuresenable the nanoparticle surfaces to interact with the surroundingsubstrates and thus catalyze organic reactions in both aqueous(original dispersion media) and organic media (after phasetransfer by modification of dendrimers with alkane carboxylicacid).6However, this dendrimer encapsulation protocol should bematerials-specific, and its applicability is limited especially by thedeliberate design of dendrimers. Furthermore, the surface modi-fication of nanoparticle guests during modification of the den-drimer hosts is inevitable. Herein hydrogels are employed asgeneric carriers and hosts for hydrophilic NPs for phase transferwithout modification of the NPs or hydrogels.
A hydrogel is a three-dimensional, physically and/or chemi-cally cross-linked network of hydrophilic natural or syntheticpolymers. As the name suggests, it usually is in a form of water-swollen jellylike solid. Hydrogels have been extensively exploitedfor various technical uses, as superabsorbents, sensors, contactlenses, medical electrodes, scaffolds for tissue engineering, dres-sings for wound healing, and implants, just to name a few.7 Thehighly porous and biomimetic aqueous interior environment andthe stimuli-responsive swelling and shrinking behavior makehydrogels ideal candidates for encapsulation and release ofhydrophilic drug NPs and proteins in a controlled manner.8 Thisencapsulation strategy has recently been extended to varioushydrophilic inorganic NPs, such as CdTe quantum dots (QDs)and gold NPs, and opened up new pathways to produce multi-functional nanocomposites.9-11 In order to guarantee encapsula-tion of hydrophilic NPs within hydrogels, attractive interactionand sizematching between the particle guests and the pores of thehydrogel hosts are needed.8-11
It can be easily understood that hydrophilic polymers can besoluble in a wide range of organic solvents of different polarity bytaking into account the fact that the molar fractions of theirhydrophobic segments are at least comparable to those of thehydrophilic segments. Wang et al. have recently demonstratedthat the dual solubility of the hydrophilic polymer coating canimpart NPs with dispersibility in both water and most organicsolvents and even the capability of reversible transfer between
aqueous and organic media across the interfaces.12 This concepthas also been extended to transfer hydrogelmicroparticles (MPs),loadedwithout andwith hydrophilic inorganicNPs, fromaqueousto organic media via stepwise exchange of the water initiallyswollen in the hydrogelswithwater-miscible organic solvents suchas ethanol and tetrahydrofuran (THF) and eventuallywith water-immiscible organic solvents such as toluene and hexane.11c,13
Inspired by these successes, we here present a new approach ofusing hydrogel MPs to directly encapsulate and transfer hydro-philic NPs into organic media via stepwise solvent exchange.Different from the previous studies,8-11 hydrophilic NPs areembedded within hydrogelMPs in the course of solvent exchangeand phase transfer. They are confined within the hydrogel hostssoaked with organic solvents after solvent exchange by theincompatibility of the NP guests with the organic surroundingmedia.As a result, chemicalmodification of the surfaces of hydro-philic NPs and/or the hydrogel networks is avoided. Further-more, no size matching or chemical affinity of the hydrophilicNPs to the hydrogel networks is required. Thus, the presentapproach is independent of the chemical nature of hydrophilicNPs and hydrogels. We have successfully employed both agaroseand poly(N-isopropylacrylamide) (PNIPAM) hydrogelMPswithdifferent sizes as carriers and hosts to encapsulate hydrophilicNPswith very different properties such asCdTeQDs and lipase Bfrom Candida antarctica (CalB) and transfer them to organicmedia with little change of their intrinsic behavior, such asphotoluminescence and catalytic activity.
Experimental Section
Materials.Acrylic acid (AA), agarose (typeXI), bovine serumalbumin (BSA) (g98%, lyophilized powder,∼66 kDa), cadmiumchloride hemi(pentahydrate) (CdCl2, 99þ%),N-isopropylacryla-mide (97%) (NIPAM), 3-mercaptopropionic acid (MPA) (99%),octanoic acid (g99%), 1-octanol (g99%), oleylamine (technicalgrade, g 70%), and tellurium powder (-200 mesh, 99.8%) werepurchased from Sigma-Aldrich (Munich, Germany), and N,N0-methylenebis(acrylamide) (MBA) (g99.5%) and potassium per-oxodisulfate (KPS) (g99%) were from Fluka (Munich,Germany). NIPAM was purified by recrystallization by usingn-hexane. Other chemicals were used as received. Lipase B fromCandida Antarctica (CalB, Lipozyme CALB L, protein content
Scheme 1. Schematic Illustration of Using Hydrogel MPs as
Carriers for Phase Transfer of Hydrophilic NPs from Water to
Water-Immiscible Organic Solvents and Thus Encapsulation via
Solvent Exchange
(6) Crooks, R. M.; Zhao, M.; Sun, L.; Chechik, V.; Yeung, L. K. Acc. Chem.Res. 2001, 34, 181.(7) (a) Sauders, B.; Vincent, B. Prog. Colloid Polym. Sci. 1997, 105, 11. (b)
Torres-Lugo, M.; Ward, J.; Zhang, J. Annu. Rev. Biomed. Eng. 2000, 2, 9. (c)Eichenbaum, G. M.; Kiser, P. F.; Dobrynin, A. V.; Simon, S. A.; Needham, D.Macromolecules 1999, 32, 4867.(9) (a) Peppas, N. A.; Hilt, J. Z.; Khademhosseini, A.; Langer, R. Adv. Mater.
2006, 18, 1345. (b) Nayak, S.; Lyon, L. A. Angew. Chem., Int. Ed. 2005, 44, 7686. (c)Ballauff,M.; Lu, Y.Polymer 2007, 48, 1815. (d) Karg,M.; Hellweg, T. J.Mater. Chem.2009, 19, 8714.(10) (a) Martinez-Rubio, M. I.; Ireland, T. G.; Fern, G. R.; Silver, J.; Snowden,
M. J. Langmuir 2001, 17, 7145. (b) Zhang, J.; Xu, S.; Kumacheva, E. J. Am. Chem.Soc. 2004, 126, 7908. (c) Holtz, J.; Asher, S. Nature 1997, 389, 829. (d) Sheeney-Haj-Ichia, L.; Sharabi, G.; Willner, I. Adv. Funct. Mater. 2002, 12, 27. (e) Contreras-Cacers, R.; Sanchez-Iglesias, A.; Karg,M.; Pastoriza-Santos, I.; Perez-Juste, J.; Pacifico,J.; Hellweg, T.; Fernandez-Barbero, A.; Liz-Marzan, L. Adv. Mater. 2008, 20, 1666.(11) (a) Kuang, M.; Wang, D.; Bao, H.; Gao, M.; M€ohwald, H.; Jiang, M. Adv.
Mater. 2005, 17, 267. (b) Gang, Y.; Gao, M.; Wang, D.; M€ohwald, H. Chem. Mater.2005, 17, 2648. (c) Kuang, M.; Wang, D.; M€ohwald, H. Adv. Funct. Mater. 2005, 15,1611–1616.(12) (a) Duan, H.; Kuang, M.; Wang, D.; Kurth, D.; M€ohwald, H. Angew.
Chem., Int. Ed. 2005, 44, 1717. (b) Edwards, E. W.; Chanana, M.; Wang, D.; M€ohwald,H. Angew. Chem., Int. Ed. 2008, 47, 320. (c) Edwards, E. W.; Chanana, M.; Wang, D.J. Phys. Chem. C 2008, 112, 15207.
0.24% (w/w)) was generously donated by Novozymes A/S(Bagsvaerd,Denmark). Benzaldehyde lyase (BAL) was expressedand purified according to the literature.14 The fluorescein iso-thiocyanate (FITC) labeling kit was purchased from EMDBiosciences (Nottingham, UK). The water in all experimentswas prepared in a three-stageMilliporeMilli-Q Plus 185 purifica-tion system and had a resistivity higher than 18.2 MΩ cm.
Preparation of CdTe QDs.MPA-stabilized CdTe QDs withdifferent sizes were synthesized in water according to the methodreported by Zhang et al.15 Typically, freshly prepared aqueousNaHTe solutions were injected into 20 mM N2-saturated CdCl2solutions in the presence of various thio-ligands as stabilizers toform the QD precursors at pH 11. The molar ratio of Cd2þ:thiol:HTe- was set as 1:2.4:0.5. The resultant precursor solutions wererefluxed at 100 �C to prepare CdTe QDs. The QD sizes werecontrolled to be 2.8, 3.3, and 4.0 nmby increase of the reflux time,and the sizes were estimated from the first electronic transition inthe absorption spectra of the QDs.
Preparation ofHydrogelMPs.AgaroseMPswith sizes in therange of 5-50 μmwere obtained according to a modified protocolreported byMao et al.16 Typically, 15 mg of agarose was dissolvedin 1mL of water at 80 �C; the agarose concentrationwas 1.5 wt%.
The resulting hot aqueous solution was dropped into 20 mLof hexane in the presence of oleylamine (5% w/v) as stabilizerunder sonication at ambient condition. Oleylamine was removedby washing with tetrahydrofuran (THF) for at least 3 times.The agarose MPs were consecutively washed with isopropanol(IPA) for 3 times and with water for 3 times and finally storedin water.
PNIPAM MPs with diameters of about 600 nm were synthe-sized via surfactant free emulsion polymerization at 70 �C.PNIPAM MPs with diameters of about 1.5 μm were synthesizedvia surfactant free emulsion polymerization under deliberatetemperature control.17 Typically, 1.8 g of NIPAM and 0.06 g ofMBAweredissolved in 125mLofwater.After deaerationwithN2
for 1 h, the aqueous mixture was slowly heated to 45 �C, followedby addition of 5 mL of KPS aqueous solution (0.078 M) undergentle stirring. Subsequently, the reaction medium was heated to65 �C at a rate of 0.5 �C/min and maintained at 65 �C overnightunder stirring to guarantee completion of the polymerization.After removal of aggregates via filtration through glass wool, theresulting PNIPAMMPs were purified by 4 times repetition of 1 hcentrifugation at 8965g, decanting the supernatants, and redis-persion in water. They were collected via lyophilization at-85 �Cunder 1 � 10-3 bar for 48 h.
Phase Transfer and Encapsulation of Hydrophilic CdTe
QDs into Hydrogel MPs. As depicted in Scheme 1, hydrogelMPs were incubated in aqueous dispersions of CdTe QDs for 2 h
Figure 1. CLSM images of agarose MPs loaded with hydrophilic CdTe QDs dispersed in water (a), isopropanol (b), and hexane (c), in thecourse of solvent exchange. The fluorescence, transmission, and their overlay images are shown in the left, middle, and right panel,respectively. Part a was shot immediately after separation of the gel MPs from the aqueous dispersion of the QD.
(14) Janzen, E.; Mueller, M.; Kolter-Jung, D.; Kneen, M. M.; Mcleich, M. J.;Pohl, M. Bioorgan. Chem. 2006, 34, 345.(15) Zhang, H.;Wang, L.; Xiong, H.; Hu, L.; Yang, B.; Li,W.Adv.Mater. 2003,
to ensure homogeneous and extensive diffusion of the QDs intothe hydrogel MPs. After decanting the aqueous QD dispersion,the MPs were collected by centrifugation and incubated inisopropanol for 1 min. After centrifugation, theMPs were readilydispersed into various water-immiscible organic solvents such aschloroform, toluene, and hexane, and thus the QDs were trans-ferred into the organic solvents and trapped within agarose orPNIPAM MPs soaked by the organic solvents. Large agaroseMPs were easily collected by simple sedimentation or by slightcentrifugation at 2000g for 5 min, and small PNIPAMMPs werecollected via centrifugation at 18000g for 12 min. In order toimprove the dispersibility of QD-loaded hydrogel MPs in water-immiscible organic solvents, they were incubated in THF for
1min after IPA incubation,whichwasusually a need in the caseofPNIPAM MPs.
Phase Transfer and Encapsulation of Proteins/Enzymes
into Hydrogel MPs. For phase transfer of BSA, CalB, andBAL, hydrogel MPs were incubated overnight in the bufferedenzyme solutions (0.1 M potassium phosphate buffer, pH 7)at 4 �C. In order to minimize the adverse effect of water-miscibleorganic solvents on the enzymes, large agaroseMPs, which couldrapidly be separated from the organic solvents, were used ascarriers and host for phase transfer and isopropanol (IPA) wasused for BSA and CalB loading and dimethyl sulfoxide (DMSO)for BAL loading. For visualization of the enzyme loading withfluorescence microscopy and spectroscopy the enzymes werelabeled with FITC according to the instructions in the FITClabeling kit (Supporting Information).
Characterization of Transfer Process and MP and NP
Properties. The sizes of CdTe QDs were determined by UV-visabsorption spectroscopy (Cary 50 UV-vis spectrophotometer,Varian, Inc.). Hydrogel MPs in water were characterized bydynamic light scattering and confocal microscopy (Leica DMIRBE confocal laser scanning microscope (CLSM) with a 30 WUV lamp (λ = 350 nm) as the light source). Transfer andencapsulation of hydrophilic NPs into hydrogel MPs via solventexchange were analyzed by CLSM. The photoluminescencevariation of CdTe QDs during transfer and encapsulation ofhydrogel MPs via solvent exchange was analyzed by using aFluoromax-4 spectrophotometer. The secondary structures ofenzymes were analyzed by circular dichroism (CD) spectroscopy(JAS.C.O J-715 spectropolarimeter). The concentration of CdTeQDs loaded in hydrogel MPs was estimated with the help ofUV-vis absorption spectroscopy (Figure S1, Supporting In-formation). The number of hydrogel MPs was determined byusing single particle light scattering (SPLS).18 The concentrationof CalB immobilized in agaroseMPs was quantified via Bradfordassay.19
Assessment of the Catalytic Performance of Free and
Immobilized CalB. The catalytic performance of CalB wasdetermined via the esterification of 1-octanol and octanoic acidin heptane. Typically, 50 μL droplets of the aqueous solution ofnativeCalB (0.12mgof protein) or 500mgof agaroseMPs loadedwith CalB (0.42 mg protein) were thrown in 5 mL of substratesolution in heptane containing 100 mM 1-octanol and 100 mMoctanoic acid.Thesedispersionswere stirredat 400 rpm for 50minat 40 �C. During the reaction, 50 μL aliquots of the reaction
Figure 2. Optical and fluorescent photographs of an agarosehydrogel block loaded with hydrophilic CdTe QDs after a definedamount of isopropanol was added atop for 3 h.
Figure 3. (a) Optical photographs of glass vials in which differentwater-miscible polar organic solvents are added atop agarosehydrogel blocks loaded with hydrophilic CdTe QDs. From left toright, the organic solvents used are ethanol, IPA, acetone, THF,DMF, and DMSO. The photographs were shot after 3 h incuba-tion. (b) Chart of the thickness percentages of the colorless upperlayers in the hydrogel blocks versus different water-miscible or-ganic solvents.
Figure 4. Fluorescence spectra of original CdTe QDs in water(black curve) and CdTe QD-loaded agarose MPs in hexane viasolvent exchange by using pure IPA (red curve) and the IPAsolution of MPA (blue curve). The MPA concentration in theIPA solution is identical to that in the aqueous dispersion of CdTeQDs. The CdTe QDs are about 3.3 nm in size.
solutions were taken every 10 min and subjected to gas chroma-tography (GC) (Shimadzu 2010; BTX column from SGE: length25 m, i.d. 0.22 mm; film thickness: 0.25 μm; detector: FID at300 �C; injector: 275.0 �C, injection volume of 1 μL, split model;temperature program: start temperature 80.0 �C, hold for 0.5min,temperature rise 20 �C/min from 80 to 170 �C and 5 �C/min risefrom170 �C to end temperature 200 �C). The concentration of theproduct octyl octanoate was calculated at a typical retention timeof 10.01 min. All reactions were performed in triplicate. Oneunit permg (U/mg) of specific activity of free or immobilizedCalBwas defined as 1 μmol of product produced per min per mg offree or immobilized CalB.
For determination of enzyme reusability, agarose MPs loadedwith CalB were separated from old substrate media, washedthree times with 10 mL of heptane, and redispersed in freshsubstrate media for the next experimental run. Enzyme stabilitywas measured by analyzing the residual catalytic activity afterstorage of agarose MPs loaded with CalB in hexane at 30 �C for10 days under gentle shaking. All experiments were performedin triplicate.
Results and Discussion
Using Hydrogel MPs for Phase Transfer and Encapsula-
tion of CdTe QDs. Fluorescent CdTe QDs were used as modelsto demonstrate the present methodology because they allowedboth fluorescence and absorption based detection of the QDloading and transfer, and their fluorescence behavior is verysensitive to changes in surface chemistry. Because of the highwater content of hydrogelMPs (98-99wt% in the currentwork),CdTe QDs can easily diffuse into the gel network like they do inwater. This was demonstrated by the fact that after 2 h incubationfollowed by fast separation from the aqueous dispersions ofCdTeQDs agarose MPs became fluorescent and the fluorescence washomogeneously distributed over the MPs (Figure 1a). This easyand fast diffusion of CdTe QDs through agarose MPs suggeststhat the pores of the agarose hydrogel network (agarose concen-tration of 1.5 wt %) are much larger than the QDs size (in therange of 2.8-4.0 nm). Since the stabilizing ligands of the QDs,MPA, are completely deprotonated under the applied conditions(pH 11), which renders the QD surfaces negatively charged,the hydrogen bonding between the QDs and the network of theagarose hydrogel network is expected too weak to confine theQDs within the hydrogel MPs. As a result, the redispersion ofCdTe QD-loaded agarose MPs back into water caused a fastrelease of the QDs out of the agarose MPs via diffusion; no QDswere left after washing the agarose MPs with water (Figure S2,Supporting Information).
In order to confine CdTe QDs in the hydrogel network, afterCdTe QD-loaded agarose MPs were separated from the aqueousdispersion of the CdTe QDs, they were immediately dispersedin polar water-miscible organic solvents, such as ethanol, IPA,
acetone, tetrahydrofuran (THF), dimethylformamide (DMF),and DMSO. The incompatibility of the negatively chargedsurfaces of the CdTe QDs with these solvents forced the QDs toremain within the agarose MPs, as shown in the left panel ofFigure 1b.
The fluorescence and transmission overlay imaging reveals anonfluorescent periphery of all CdTe QD-loaded agarose MPs,dispersed in for instance IPA, suggesting that the organic polarsolvents dispel theQDs to the center of the agaroseMPs (the rightpanel of Figure 1b). In the current work, we used agarosehydrogel blocks to assess the effect of organic polar solvents onthe migration and intrinsic fluorescence behavior of CdTe QDsloaded in the hydrogels. Noteworthy, polar water-miscible sol-vents such as IPA are usually used to precipitate CdTe QDs fromtheir aqueous dispersions accompanied by the precipitation andfluorescence quenching of theQDs.However, Figure 2 shows thatafter IPA is brought on the top of a CdTe QD-loaded hydrogelblock, the CdTe QDs did not precipitate but moved toward thebottom of the hydrogel block. The hydrogel block was clearlydivided into two layers: the colorless and nonfluorescent upperlayer and the yellow and fluorescent lower layer. The strongfluorescence of the lower layers suggests that they remain rich inwater rather than organic polar solvents; otherwise, precipitationand fluorescence quenching of CdTe would be observed. Thecolorlessness of the upper layers suggests no presence andprecipitation of the CdTe QDs inside these upper layers; other-wise, they would be either yellow due to the QD presence or darkblue due to the QD precipitation or shape transformation.20
Intriguingly, the thickness of the newly formed colorless upperlayer increases with the dielectric constant of the organic polarsolvent (Figure 3). The migration of CdTe QDs in the presence oforganic polar solvents is reminiscent of the process of thin layerchromatography.21
After dispersion of CdTe QD-loaded agarose MPs in polarwater-miscible organic solvents such as IPA, their peripheralshells are expected to be rich in the organic solvents and thusmore compatible with water-immiscible organic polar and evenapolar solvents. Therefore, after incubation in IPA, CdTe QD-loaded agarose MPs were able to readily disperse in variouswater-immiscible organic solvents, e.g., chloroform, toluene,hexane, and many others. Figure 1c shows that the dispersionof CdTe QD-loaded hydrogel MPs in water-immiscible organicsolvents such as hexane causes a slight collapse of the agarose gelnetwork, which is evidenced by the disappearance of the non-fluorescent peripheral shells.
Figure 5. Fluorescence spectra of original CdTe QDs in water (black curve), CdTe QD-loaded agarose MPs in different water-immiscibleorganic solvents via solvent exchange (red curve), andCdTeQDs redispersed inwater after release out of the agaroseMPs via reversed solventexchange (blue curve). The water-immiscible organic solvents used are hexane (a), toluene (b), and chloroform (c). The CdTe QDs are about3.3 nm in size.
(20) (a) Zhang, H.; Wang, D.; M€ohwald, H. Angew. Chem., Int. Ed. 2006, 45,748. (b) Zhang, H.; Wang, D.; Yang, B.; M€ohwald, H. J. Am. Chem. Soc. 2006, 128,10171.
In addition to large and polydisperse agarose MPs, small andmonodisperse PNIPAMMPswith sizes of 600 nmand 1.5 μmcanbe used for phase transfer and encapsulation of hydrophilic CdTeQDs in water-immiscible organic solvents (Figure S3, SupportingInformation). The advantage of using small monodisperse PNI-PAM MPs is that one can quantitatively analyze the concentra-tion of CdTe QDs loaded in each hydrogel MPs with the aid ofabsorption spectroscopy and static light scattering.11,13 The con-centration of QDs in PNIPAMMPs was almost identical to thatin original aqueous solution; each MP contained about 650 QDs(Figure S1, Supporting Information). This further confirms thatloading of CdTe QDs in hydrogel MPs is a diffusion-drivenprocess; no size matching and attractive affinity are neededbetween the QDs and the hydrogel networks. Otherwise, theconcentrations of CdTe QDs loaded in hydrogel MPs would bedifferent from those of their aqueous solutions.
CdTe QD-loaded hydrogel MPs remain fluorescent in water-immiscible organic solvents such as hexane, but the fluorescenceintensity is much weaker than that of aqueous dispersions ofCdTe QDs; it is about 15% of that of the aqueous dispersions(Figure 4). This suggests that a considerable amount of thestabilizing ligands of MPA are removed from the surfaces ofCdTe QDs during the solvent exchange, thus leading to a poorsurface passivation and in turn a weaker fluorescence. When thesolutions ofMPA in IPA or THF with the same concentration asthat of aqueous dispersions of CdTe QDs were used for the QDs
transfer from water to water-immiscible organic solvents, thefluorescence intensity of CdTe QDs-loaded hydrogel MPs re-mains almost identical to that of the QDs in original aqueousdispersions (Figure 4). This also proves that there is no QD lossduring solvent exchange.
The solvent exchange procedure described above can bereversed from water-immiscible organic solvents to water-mis-cible solvents and eventually back towater or aqueous solution ofMPA. When CdTe QD-loaded hydrogel MPs were redispersedback into water, CdTe QDs were released from the hydrogelMPsand redispersed in aqueous media, for instance MPA aqueoussolution. As shown in Figure 5, the profiles of the fluorescencespectra of CdTe QDs are little changed before and after phasetransfer from water into different water-immiscible organicsolvents via solvent exchange and transfer back toMPA aqueoussolutions via reversed phase transfer Nevertheless, an about 5 nmred shift of the fluorescence band of CdTeQDs is clearly visible inparticular by comparison between original QDs in water and theQDs redispersed in water after reserved solvent exchange. Thereason accounting for this small red shift could be threefold: (1)the rearrangement of MPA ligands on the QDs and/or theformation of CdS shells due to decomposition of the thiol groupsof theMPA lead to a better passivation;22 (2) the growth of CdTe
Figure 6. CLSM images of agaroseMPs loadedwithFITC labeledCalBdispersed inPBS (a), IPA (b), and hexane (c) in the course of solventexchange. The fluorescence, transmission, and their overlay images are shown in the left,middle, and right panel. Part awas shot immediatelyafter separation of the gel MPs from the aqueous dispersion of the CalB. The scale bar is 200 μm.
(22) Gao, M.; Kirstein, S.; M€ohwald, H.; Rogach, A. L.; Kornowski, A.;Eychm€uller, A.; Weller, H. J. Phys. Chem. B 1998, 102, 8360.
QDs in hydrogel MPs due to the removal and regeneration of thesurface passivation shells of MPA; and (3) slight aggregation ofCdTe QDs because all water-miscible organic polar solvents canbe regarded as precipitating agents for CdTe QDs. Elucidation ofthe red shift of the fluorescence of CdTeQDs obviously requires ameticulous study of the change of the structures and especiallysurface chemistry of the QDs during solvent exchange, which iscurrently underway.Hydrogel MPs for Phase Transfer and Encapsulation of
Enzymes. Enzymes, as peculiar and environmentally friendlycatalysts for organic reactions, are increasingly used in chemicalsynthesis because of their high activity and stereoselectivity.Lipases are frequently employed to catalyze the chemo-, regio-,and/or stereoselective hydrolysis of esters and amides.23However,the technical application of many hydrophilic and vulnerableenzymes requires immobilization on or in appropriate carriers toimprove the dispersibility in organic media, the long-term opera-tional stability, and the efficiency of recovery and reuse.24 Herewe extended the solvent exchange process to the encapsulation ofCalB in hydrogel MPs and transfer them to water-immiscibleorganic solvents in order to provide a simple and efficientimmobilization technique. For the benefit of easy visualizationof enzyme loading,CalBwas labeledwith the fluorescencemarkerFITC (Supporting Information).
Similar to CdTe QDs, CalB can be transferred to water-immiscible organic solvents such as hexane and in turn encapsu-lated in MPs of agarose (Figure 6) or PNIPAM (Figure S4) viasolvent exchange by using IPA as the intermediate solvent.Furthermore, the reversed solvent exchange allows release ofCalB from the hydrogel MP hosts and redispersion in aqueousmedia. The CD spectra of CalB, redispersed in PBS buffer afterrelease out of hydrogel MPs via reversed solvent exchange, arerather similar to those of nativeCalB; the bands in thewavelengthrange of 190-240 nm, characteristic of the R-helix secondarystructure,25 are clearly observed in Figure 7.
In the case of native CalB, the enzymatic catalysis occurredonlywhen substrates slowly diffused fromheptane to the aqueous
droplets of the CalB across the interface.24 On the other hand, theaqueous droplets of the native CalB were not stable in heptaneand tended to coalesce, thus leading to rather small interface areafor enzymatic catalysis. In contrast, immobilization of enzymes inhydrogel MPs drastically increased the interfacial area of thehydrophobic substrates in contact with hydrophilic CalB. Figure 8atherefore shows that the specific catalytic activity of CalB-loadedhydrogel MPs, dispersed in heptane, is significantly higher thanthat of native CalB. Intriguingly, the specific activity of CalB inhydrogel MPs drastically increases from 2.7 U/mg in the firstcatalysis trial to 13.4 U/mg in the second trial and then slightlyincreases with every reuse (7 times). In contrast, the specificactivity of native CalB just slightly increases in the secondcatalysis trial while afterward it slightly decreases with every reuse(Figure 8a). The excellent reusability of CalB-loaded hydrogelMPs could be of great interest for the practical use in chemicalsynthesis. Figure 8b shows that the residual activity of CalB inhydrogel MPs is little changed during storage in hexane for atleast 10 days. In contrast, the residual activity of native CalBgreatly decreased with time when being brought in contact withhexane. This stabilization of CalB should be due to the immobi-lization effect of the gel network of agarose MPs on the enzymes.
In order todetermine the overall applicability of themethod forprotein entrapment and transfer to organic solvents, the solvent
Figure 7. CD spectra of native CalB (black curve) in PBS andCalB redispersed in PBS after being released fromagaroseMPs viareversed solvent exchange (red curve).
Figure 8. (a) Plot of the specific activity of native CalB (red curve)and CalB-loaded agarose MPs (black curve) versus the number ofreuses. (b) Plot of the stability of native (red curve) and immobi-lized CalB (black curve) versus the time of storage in hexane; thedata of the specific activity were normalized by taking the highestvalue as 100%.
(23) (a) Gotor-Fern�andez, V.; Busto, E.; Gotor, V.Adv. Synth. Catal. 2006, 348,797. (b) Reetz, M. T. Curr. Opin. Chem. Biol. 2002, 6, 145.(24) Ansorge-Schumacher, M. B. Mini-Rev. Org. Chem. 2007, 4, 243. (b)
Klibanov, A. Nature 2001, 409, 241.(25) De Diego, T.; Lozano, P.; Gmouh, S.; Vaultier, M.; Iborra, J. L. Bioma-
exchange strategy was applied to fairly inert and stable BSA witha size of 4 nm � 4 nm � 14 nm and fairly active and vulnerableBAL with a size of 10.4 nm� 12.1 nm� 16.2 nm.26 It was shownthat bothBSA andBAL can be successfully transferred fromPBSbuffer to water-immiscible organic solvents and encapsulated inagarose or PNIPAM MPs (Figures S5 and S6, SupportingInformation). Combined with the results obtained with CdTeQDs, this demonstrates that the presented method of solventexchange is suitable for handling of different sized hydrophilicNPs. However, no residual catalytic activity was detected forBALevenwhenDMSO, themost commonly usedorganic solventfor BAL catalysis,27 was used as the intermediate water-miscibleorganic solvent. This could be explained by the high sensitivity ofBAL toward organic solvents at high concentrations and itsoverall low stability even in PBS buffer28 and suggests that afairly basic stability of enzyme catalysts is required beforeentrapment in hydrogel MPs by solvent exchange is beneficial.
Conclusions
We have demonstrated a simple and versatile approach to usehydrogel MPs as generic carriers and hosts for phase transfer ofhydrophilic NPs into organic media via solvent exchange. Hydro-philic NPs have been encapsulated within hydrogel MPs in thecourse of phase transfer via solvent exchange fromwater towater-miscible organic solvents and eventually to water-immiscibleorganic solvents. The entrapment of the NPs results from theirincompatibility with water-immiscible organic solvents soaked inthe gel matrices and in the surrounding environment. As aconsequence, no chemical modification of the surfaces of hydro-philic NPs and/or of the networks of hydrogels is needed, andfurthermore, any size matching or chemical affinity of thehydrophilic NPs for the hydrogel networks is unnecessary. Thepresent approach is therefore independent of the chemical natureof hydrophilic NPs and hydrogels MPs. The concentration of
hydrophilic NPs loaded in hydrogels after phase transfer toorganic solvents such as hexane has been demonstrated to becomparable to that in their original aqueous dispersion. It has alsobeendemonstrated that the surface stabilizing shells ofCdTeQDscan be removed and regenerated during solvent exchange in acontrolled fashion, which is exemplified by their photolumines-cence quenching and recovery. This suggests the capability andflexibility of tuning the surface chemistry of hydrophilic inorganicNPs loaded in hydrogel MPs, which should be of importance forboth fundamental research, for instance study of the surfacechemistry effect, and technical applications such as catalysis. Ofimportance is that the phase transfer and encapsulation ofhydrophilic NPs are reversible; the reversed solvent exchange ofhydrogel MPs loaded with hydrophilic NPs from organic medialeads to release and recovery of the hydrophilic NPs in aqueousmedia. The presented approach has been successfully extended tothe encapsulation of the enzyme catalyst CalB and its phasetransfer to organic media with little change of catalytic activity,good reusability, and stability, thus showing a promising poten-tial for use in biocatalysis.
Acknowledgment. This work is a part of the Cluster ofExcellence “Unifying Concepts in Catalysis” coordinated by theTechnischeUniversit€at Berlin. Financial support by theDeutscheForschungsgemeinschaft (DFG) within the framework of theGerman Initiative for Excellence is gratefully acknowledged(EXC 314). S.B. and D.W. thank the Max Planck Society forthe financial support and H. M€ohwald for valuable discussionsand research support. D.W. is in part supported by a DFG grant(WA 1704/4-1). S.B. and D.W. thank H. Lichtenfeld for assis-tance with SPLS measurement and H. Zhang (Jilin University,P. R. China) for valuable help of preparation of CdTe QDs andanalysis of their photoluminescence behavior.
Supporting Information Available: Calculation of thenumber of CdTe QDs per PNIPAM MP; fluorescenceimages of agarose MPs loaded with CdTe QDs in waterand CLSM images of agarose MPs loaded with BSA andBAL in hexane andPNIPAMMPs loadedwithCdTeQDs inhexane. This material is available free of charge via theInternet at http://pubs.acs.org.
(26) Maraite, A.; Schmidt, T.; Ansorge-Schumacher,M. B.; Brzozowski, A.M.;Grogan, G. Acta Crystallogr. 2007, F63, 546.(27) Hischer, T.; Gocke, D.; Fern�andez, M.; Hoyos, P.; Alc�antara, A. R.;
Sinisterra, J. V.; Hartmeier,W.; Ansorge-Schumacher,M. B.Tetrahedron 2005, 61,7338.(28) van den Wittenboer, A.; Nimeijer, B.; Karmee, S.; Ansorge-Schumacher,
