Microcompartmentation in Artificial Cells: pH-Induced Conformational

Nov 20, 2009 - Our model cells are giant lipid vesicles (GVs, ca. ... also relocalized within compartmentalized artificial cells in response to extern...
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Microcompartmentation in Artificial Cells: pH-Induced Conformational Changes Alter Protein Localization Lisa M. Dominak, Erica L. Gundermann, and Christine D. Keating* Department of Chemistry, Pennsylvania State University, University Park, Pennsylvania 16802 Received October 7, 2009. Revised Manuscript Received November 3, 2009 We report artificial cells in which protein localization in a primitive synthetic model for the cytoplasm is controlled by pH. Our model cells are giant lipid vesicles (GVs, ca. 5-30 μm diameter) with two coexisting aqueous compartments generated by phase separation of an encapsulated poly(ethylene glycol) (PEG) and dextran solution. Proteins are localized to a microcompartment by partitioning between the phases. We quantified the local concentration of fluorescently labeled human serum albumin (HSA) via confocal fluorescence microscopy. At pH 6.5, the labeled HSA was more concentrated in the dextran-rich phase, but at partially/fully denaturing pH (4.1 or 12) it was localized in the PEG-rich phase. This partitioning behavior is consistent with a more expanded, hydrophobic conformation at low and high pH. Labeled HSA could be relocalized from the PEG-rich into the dextran-rich phase domain by increasing the pH from 4.1 to 6.5 to renature the protein. This approach to controlling protein localization does not require extensive reorganization of the vesicle interior; coexisting PEG-rich and dextran-rich compartments are maintained throughout the experiments. It is also quite general; we demonstrated that several other proteins varying in size and isoelectric point also relocalized within compartmentalized artificial cells in response to external pH change. This work presents stimulus-responsive protein relocalization between compartments in an artificial cell; such experimental models can provide a framework for investigating the consequences of protein localization in cell biology.

Introduction Due to the complexity of living cells, many hypotheses in cell biology are difficult to evaluate. Simplified, nonbiological experimental model systems are needed in which potential mechanisms for, and consequences of, intracellular organization can be evaluated. Here, our interest is in developing a model system in which local protein concentration can be altered in a synthetic model cytoplasm without otherwise changing the intracellular organization by, for example, changing the number or type of subcellular compartments present. Biological organisms are characterized by sophisticated intracellular organization.1-5 Biomolecules, biomolecular assemblies, and organelles are often organized into distinct regions, or compartments, within the cell. These can be as complex as the mitochondria of eukaryotic cells, which is itself compartmentalized and is separated from the cytoplasm by two membranes, or as simple as the nucleoid of bacterial cells, which contains the condensed genome of the organism and is irregular in shape with no membranous boundary.6 Indeed, many types of compartments not bounded by membranes have been identified in living cells.7-10 These can occur via binding interactions to form *To whom correspondence should be addressed. E-mail: keating@ chem.psu.edu. (1) Misteli, T. Cell 2007, 128, 787–800. (2) Shapiro, L.; Losick, R. Science 1997, 276, 712–718. (3) Ovadi, J.; Saks, V. Mol. Cell. Biochem. 2004, 256/257, 5–12. (4) Misteli, T. J. Cell Biol. 2001, 155, 181–185. (5) Hess, B.; Mikhailov, A. Science 1994, 264, 223–224. (6) Alberts, B.; Johnson, A.; Lewis, J.; Raff, M.; Roberts, K.; Walter, P. Molecular Biology of the Cell, 4th ed.; Garland Science: New York, 2002. (7) Sheth, U.; Parker, R. Science 2003, 300, 805–808. (8) Boisvert, F.-M.; van Koningsbruggen, S.; Navascues, J.; Lamond, A. I. Nature Rev. Mol. Cell Biol. 2007, 8, 574–585. (9) Brangwynne, C. P.; Eckmann, C. R.; Courson, D. S.; Rybarska, A.; Hoege, C.; Gharakhani, J.; Julicher, F.; Hyman, A. A. Science 2009, 324, 1729–1732. (10) An, S.; Kumar, R.; Sheets, E. D.; Benkovic, S. J. Science 2008, 320, 103– 106.

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complexes in the cytoplasm or nucleoplasm or be bound to structural elements such as cytoskeletal fibers or membranes.3,11 In other cases, differences in local biomolecule concentrations arise due to local synthesis or degradation, restricted diffusion, or other, sometimes unknown, mechanisms.3,12,13 Recently, germline P granules in C. elegans embryos have been demonstrated to be liquid droplets, a new class of intracellular compartment.9 Intracellular compartmentalization is not static. For example, a proteolysis complex has been found to be dynamically localized in Caulobacter, degrading proteins at different cellular locations at different stages in the cell cycle.13 Chemical stimuli, such as the import or export of small ions or molecules across the cell membrane, can lead to movement and relocalization of biomolecules within the cytoplasm.10,11,14-16 For example, oxygenation is necessary for assembly of a glycolytic enzyme complex onto the red blood cell membrane.11 It was recently reported that enzymes of the purine de novo biosynthesis pathway colocalized in the cytoplasm when cells were moved to purine-depleted media; these clusters dispersed when cells were transferred back to purine-rich media.10 Compartmentalization is considered crucial for many cellular functions, but it can be difficult to study in vivo due to the complexity of the system and the challenge of affecting localization without making other, unintended changes to the living cell. An attractive potential solution is the construction of simple experimental model systems in which specific aspects of subcellular (11) Campanella, M. E.; Chu, H.; Low, P. S. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 2402–2407. (12) (a) St. Johnson, D. Nat. Rev. Mol. Cell Biol. 2005, 6, 363–375. (b) Martin, K. C.; Ephrussi, A. Cell 2009, 136, 719–730. (13) McGrath, P. T.; Iniesta, A. A.; Ryan, K. R.; Shapiro, L.; McAdams, H. H. Cell 2006, 124, 535–547. (14) Walter, H.; Brooks, D. E. FEBS Lett. 1995, 361, 135–139. (15) (a) Jones, D. P., Ed.; Microcompartmentation; CRC Press: Boca Raton, FL, 1988.(b) Harold, F. M. Microbiol. Mol. Biol. Rev. 2005, 69, 544–564. (16) Al-Habori, M. Int. J. Biochem. Cell Biol. 1995, 27, 123–132.

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organization can be investigated. Giant lipid vesicles (GVs, ca. 5-30 μm diameter)17,18 can provide a cell-sized, membranebounded model system to encapsulate a synthetic model cytoplasm. While GVs have typically been used to model reactions in dilute solutions,18-23 several groups have incorporated hydrogels24-26 or polymer solutions,27-30 which brings these structures closer to a cytoplasmic environment. We have developed a model cytoplasm based on GV-encapsulated aqueous-two phase systems (ATPSs),31-34 and have used it to control the local concentration of proteins and the spatial organization of membrane domains relative to distinct compartments in the “cell” interior.27-30 The ATPSs are formed from aqueous solutions of poly(ethylene glycol) (PEG) and dextran that phase separate to form two coexisting phase domains under appropriate conditions of polymer concentration and temperature. When encapsulated within a cell-sized liposome, the PEG/dextran ATPS provides a crowded environment (typically between 8 and 20 wt % polymer), with chemically distinct PEG-rich and dextran-rich aqueous phases that need not be separated from each other by a membrane. Previously, we have shown that local protein concentration within the vesicle interior can be changed with slight changes in temperature or osmotic pressure that induce a phase change.27 When two phases are present, the protein is compartmentalized into one of the phases, but when a single phase is present it is distributed throughout the interior volume. Therefore, not only does our primitive model of the cytoplasm incorporate macromolecular crowding,35-38 it also provides a route to dynamic compartmentalization of solutes. However, in living cells, alterations in local protein concentration are not in general the result of extensive reorganization of the cytoplasm. Rather, proteins move between existing compartments. Mechanisms for controlling the subcellular localization of proteins include covalent modifications, (17) (a) Menger, F. M.; Angelova, M. I. Acc. Chem. Res. 1998, 31, 789–797. (b) Dimova, R.; Aranda, S.; Bezlyepkina, N.; Nikolov, V.; Riske, K. A.; Lipowsky, R. J. Phys.: Condens. Matter 2006, 18, S1151–S1176. (c) Dobereiner, H.-G. Curr. Opin. Colloid Interface Sci. 2000, 5, 256–263. (18) Giant Vesicles; Luisi, P. L., Walde, P., Eds.; Perspectives in Supramolecular Chemistry; John Wiley and Sons, Ltd.: West Sussex, England, 2000; Vol. 6. (19) (a) Sott, K.; Lobovkina, T.; Lizana, L.; Tokarz, M.; Bauer, B.; Konkoli, Z.; Orwar, O. Nano Lett 2006, 6, 209–214. (b) Karlsson, A.; Sott, K.; Markstrom, M.; Davidson, M.; Konkoli, Z.; Orwar, O. J. Phys. Chem. B. 2005, 109, 1609–1617. (20) Noireaux, V.; Libchaber, A. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 17669– 17674. (21) Chen, I. A; Salehi-Ashtiani, K.; Szostak, J. W. J. Am. Chem. Soc. 2005, 127, 13213–13219. (22) Nomura, S. M.; Tsumoto, K.; Hamada, T.; Akiyoshi, K.; Nakatani, Y.; Yoshikawa, K. ChemBioChem. 2003, 4, 1172–1175. (23) Wick, R.; Luisi, P. L. Chem. Biol. 1996, 3, 277–285. (24) Faivre, M.; Campillo, C.; Pepin-Donat, B.; Viallat, A. Prog. Colloid Polym. Sci. 2006, 133, 41–44. (25) Jesorka, A.; Markstrom, M.; Orwar, O. Langmuir 2005, 21, 1230–1237. (26) Markstrom, M.; Gunnarsson, A.; Orwar, O.; Jesorka, A. Soft Matter 2007, 3, 587–595. (27) Long, M. S.; Jones, C. D.; Helfrich, M. R.; Mangeney-Slavin, L. K.; Keating, C. D. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 5920–5925. (28) Helfrich, M. R.; Mangeney-Slavin, L. K.; Long, M. S.; Djoko, K. Y.; Keating, C. D. J. Am. Chem. Soc. 2002, 124, 13374–13375. (29) Long, M. S.; Cans, A.-S.; Keating, C. D. J. Am. Chem. Soc. 2008, 130, 756– 762. (30) Cans, A.-S.; Andes-Koback, M.; Keating, C. D. J. Am. Chem. Soc. 2008, 130, 7400–7406. (31) Zaslavsky, B. Y. Aqueous Two-Phase Partitioning: Physical Chemistry and Bioanalytical Applications; Marcel Dekker: New York, 1995. (32) Albertsson, P.-A. Partition of Cell Particles and Macromolecules, 2nd ed.; John Wiley and Sons: New York, 1971. (33) Hatti-Kaul, R., Ed. Aqueous Two-Phase Systems: Methods and Protocols; Humana Press: Totowa, NJ, 2000. (34) Walter, H.; Johansson, G. Methods Enzymol. 1994, 228. (35) Zhou, H.-X.; Rivas, G.; Minton, A. P. Annu. Rev. Biophys. 2008, 37, 375– 397. (36) Ellis, R. J. Trends Biochem. Sci. 2001, 10, 597–604. (37) Johansson, H.-O.; Brooks, D. E.; Haynes, C. A. Int. Rev. Cytol. 2000, 192, 155–170. (38) Minton, A. P. Curr. Biol. 2006, 16, R269–R271.

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Dominak et al. Scheme 1. pH-Controlled Dynamic Protein Localization in Artificial Cellsa

a

Lipid membranes are shown in red and protein in green.

for example, phosphorylation, and/or changes in protein conformation.39-41 Here, we take advantage of pH changes to alter the conformation of proteins in order to affect their localization within artificial cells that contain two coexisting aqueous phase “compartments”. This was accomplished not by undergoing a phase transition but instead by changing the conformation of the protein such that it accumulated in a different aqueous phase. As such, it is an important step forward in the development of biologically relevant models of intracellular protein localization. Bulk ATPSs are often used in protein separations, and it has been demonstrated that several proteins alter their partitioning in response to a pH change. For example, Albertsson and others have shown that several proteins exhibit differences in localization due to pH.31,32,42,43 We used pH change as an external chemical stimulus to dynamically control the local concentration of a protein encapsulated within an artificial cell (Scheme 1). Localization of fluorescent human serum albumin (HSA) within twocompartment vesicles was quantified via confocal fluorescence microscopy. We find in general that HSA typically partitions to the PEG-rich phase (i.e., PEG-rich compartment) of vesicles formed at pH 4.1, while those at pH 6.5 prefer the dextran-rich compartment. Vesicles formed at pH 4.1 can be changed to pH 6.5, with HSA relocalizing from the PEG-rich to the dextran-rich phase, while a further change from pH 6.5 to 12 results in HSA migration to the PEG-rich phase. Several other proteins varying in size and isoelectric point were also found to relocalize within artificial cells as a result of external pH change, indicating the generality of this mechanism. These results are important in that they demonstrate dynamic control over local protein concentration within an artificial cell interior due to an external chemical stimulus without requiring a reorganization of the intracellular compartments. Our two-compartment vesicle system can serve as an experimental model system for investigating the effects of local protein concentrations and local microenvironments.

Results and Discussion Several proteins have been shown to exhibit pH-dependent partitioning in ATPSs.31,32,42,43 HSA partitions to the dextranrich phase of a PEG/dextran ATPS between pH 5 and 8 and to the PEG-rich phase at pH 4.1.32,43 HSA has also been reported to partition into the PEG-rich phase at basic pH values (>pH 8).43 We took advantage of this to control the local concentration of HSA and other proteins in artificial cells comprising an ATPS encapsulated within giant lipid vesicles. Here, the PEG-rich and dextran-rich aqueous phases form the subcellular compartments (39) Faux, M. C.; Scott, J. D. Trends Biochem. Sci. 1996, 21, 312–315. (40) Ubersax, J. A.; Ferrell, J. E., Jr. Nature Rev. Mol. Cell. Biol. 2007, 8, 530– 541. (41) Silhavy, T. J.; Benson, S. A.; Emr, S. D. Microbiol. Rev. 1983, 47, 313–344. (42) (a) Walter, H.; Forciniti, D. Methods Enzymol. 1994, 228, 223–233. (b) Albertsson, P.-A.; Sasakawa, S.; Walter, H. Nature 1970, 228, 1329–1330. (43) Di Nucci, H.; Nerli, B.; Pico, G. Biophys. Chem. 2001, 89, 219–229.

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Figure 1. Molar ellipticity of HSA at 222 nm as a function of pH in buffer (red), PEG-rich phase (green), and dextran-rich phase (blue). The overall ATPS composition was 3.9 wt % PEG 8 kDa and 3.9 wt % dextran 500 kDa. Lines connecting the data points were added to guide the eye.

in our artificial cells, and the membrane serves to confine a “cellsized” volume and provide a semipermeable membrane to enable the external stimulus (pH) to alter the local protein concentration within the model cell. HSA Partitioning in ATPS as a Function of pH. The polymer composition of our ATPS is more dilute than typically used for partitioning in literature reports, because our composition was selected for ease of ATPS encapsulation within the lipid vesicles. Additionally, our standard vesicle preparation protocol requires incubation at 37 °C for an extended period (see Materials and Methods). Thus, we began by verifying that HSA would exhibit pH-dependent partitioning in our bulk ATPS comprising 3.9 wt % PEG 8 kDa and 3.9 wt % dextran 500 kDa at pH 4.1 and 6.5 when treated identically to samples that would be encapsulated within lipid vesicles in later experiments. We partitioned 19 nM HSA that had been labeled with AlexaFluor 488 (HSAAF488). Partitioning is quantified in terms of the partition coefficient, K, which is defined as the concentration of solute in the PEG-rich phase (Cp) as compared to that in the dextran-rich phase (Cd), K=Cp/Cd. In our system, the labeled HSA accumulated in the PEG-rich phase at pH 4.1 (K=2.2 ( 0.58) and in the dextran-rich phase at pH 6.5 (K=0.36 ( 0.01). HSA partitioning can be understood in terms of pH-induced conformational changes. In buffer solutions, HSA adopts a native conformation near neutral pH (pH 5-7).44 At pH between 5 and ∼3.5, HSA is partially acid expanded, and at pH values below 3.5 it is fully acid expanded.44,45 Between pH 7 and 9, HSA undergoes a subtle partially base expanded conformation, while over pH 10 a steep transition to the base expanded conformation has been observed.44,45 All of these transitions were reversible.45-47 Since macromolecular crowding in our ATPS is significant and can lead to more condensed protein structures,35-38,48 we used circular dichroism (CD) to check whether the pH dependence of HSA conformation in our PEG/dextran ATPS was similar to that in buffer alone.46,49 These experiments were performed on bulk ATPS of the same composition that were treated in the same way (including incubation at 37 °C) as the ATPSs that were (44) (a) Shahid, F.; Gomez, J. E.; Birnbaum, E. R.; Darnall, D. W. J. Biol. Chem. 1982, 257, 5618–5622. (b) Hvidt, A.; Wallevik, K. J. Biol. Chem. 1972, 247, 1530– 1535. (45) Wallevik, K. J. Biol. Chem. 1973, 248, 2650–2655. (46) Rezaei-Tavirani, M.; Moghaddamnia, S. H.; Ranjbar, B.; Amani, M.; Marashi, S.-A. J. Biochem. Mol. Biol. 2006, 39, 530–536. (47) Lee, J. Y.; Hirose, M. J. Biol. Chem. 1992, 267, 14753–14758. (48) (a) Tokuriki, N.; Kinjo, M.; Negi, S.; Hoshino, M.; Goto, Y.; Urabe, I.; Yomo, T. Protein Sci. 2004, 13, 125–133. (b) Ping, G.; Yuan, J.-M.; Sun, Z.; Wei, Y. J. Mol. Recognit. 2004, 17, 433–440. (c) Stagg, L.; Zhang, S.-Q.; Cheung, M. S.; WittungStafshede, P. Proc. Natl. Acad. Sci. U.S.A. 2007, 48, 18976–18981. (49) Farruggia, B.; Pico, G. A. Int. J. Biol. Macromol. 1999, 26, 317–323.

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encapsulated within vesicles, which are described in the following sections. The molar ellipticity (θ) at 222 nm is a characteristic of the R-helical structure of a protein, and it can be used to show the extent of protein denaturation. Figure 1 shows molar ellipticity at 222 nm as a function of pH for HSA in buffer and in the PEG-rich and dextran-rich phases of an ATPS that had an overall composition of 3.9 wt % PEG 8 kDa and 3.9 wt % dextran 500 kDa. The greatest R-helicity (i.e., most negative ellipticity at 222 nm) was observed at pH 6.5 in all three media, and all three media were found to exhibit similar conformational changes. The protein loses R-helicity as the pH is raised or lowered from pH 6.5, consistent with the partial and full acid and base expansion (denaturation) of HSA that has been reported for this protein in polymer-free buffer solutions. The change in the structure of HSA at various pH values is consistent with its preference for the dextran-rich phase of an ATPS at pH 6.5 and for the PEG-rich phase at pH 4.1. Most biological molecules in their native conformation (e.g., HSA at pH 6.5) accumulate in the dextran-rich phase in a PEG/dextran ATPS, while more denatured proteins (e.g., HSA at pH 4.1) partition to the more hydrophobic PEG-rich phase.50,51 In an extended conformation, proteins tend to have greater surface hydrophobicity than when in a native conformation, due to the exposure of previously hidden residues.32,51 Since samples for these CD experiments were treated the same way as those for ATPS encapsulation in giant vesicles (including incubation at 37 °C), the nativelike partitioning at pH 6.5 suggests that nativelike conformation can be expected for HSA after encapsulation in the vesicles at this pH. Additionally, these data indicate that stabilization of the more compact native structure due to macromolecular crowding by the PEG and dextran polymers does not prevent pH-induced conformational changes under our experimental conditions. Protein Localization in Artificial Cells Formed at pH 6.5. Artificial cells were formed by preparing GVs by the gentle hydration method in the presence of a warm (37 °C) solution of 3.9 wt % PEG 8 kDa/3.9 wt % dextran 500 kDa containing 0.19 μM HSA-AF488, at pH 6.5. At this temperature, the polymer solution exists as a single phase. Upon cooling to room temperature after vesicle formation, the polymer solution phase separates both inside the vesicles and in the bulk.27,28 This led to some GVs that contained coexisting PEG-rich and dextran-rich aqueous phases, and some in which the encapsulated polymer concentration was not high enough for phase separation (for a discussion of encapsulation in GVs, see refs 27, 52, and 53). To improve the yields of artificial cells that contained compartments, we concentrated the internal polymer solution to induce phase separation by adding a hypertonic solution to shrink the vesicles (31 mM sucrose, a difference of 20 mOsm from solution in which vesicles were formed). This resulted in a mixture of GV morphologies, both spherical or budded, as previously described.29 In this Article, we will show images of both spherical and budded GVs. The selection of morphology for a particular experiment is unimportant; both types were present in all of the experiments, and both types have been included in the statistics. Figure 2 shows representative confocal fluorescence images of ATPS-containing GVs, that is, artificial cells, formed at pH 6.5. Protein localization is shown in green. Fluorescent PEG 5 kDa or dextran 500 kDa was added to indicate the location of the PEGrich and dextran-rich phase domains, respectively, which serve as (50) (51) (52) (53)

Jiang, J.; Prausnitz, J. M. J. Phys. Chem. B 2000, 104, 7197–7205. Tubio, G.; Nerli, B.; Pico, G. J. Chromatogr., B 2004, 799, 293–301. Dominak, L. M.; Keating, C. D. Langmuir 2007, 23, 7148–7154. Dominak, L. M.; Keating, C. D. Langmuir 2008, 24, 13565–13571.

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Figure 3. Relationship between the partition coefficients of HSA and PEG 5 kDa (left) and dextran 500 kDa (right) in twocompartment vesicles formed at pH 6.5.

Figure 2. Fluorescently labeled HSA and polymer partitioning in two-compartment vesicles formed at pH 6.5. Fluorescence images have been overlaid and false-colored. Green indicates HSA fluorescence, blue indicates labeled polymer fluorescence (500 kDa dextran or 5 kDa PEG, as noted), and red indicates lipid (DOPE-rhodamine) fluorescence. Scale bar is 5 μm.

our microcompartments in these experiments. The HSA is partitioned into the dextran-rich compartment, consistent with its behavior in the bulk experiments described above. To quantify local [HSA], line scans were taken across each GV and compared with calibration curves. The concentration of HSA in the PEGrich (Cp) and dextran-rich (Cd) phases was used to calculate partition coefficients. For example, the vesicle in the top panel had a 4-fold difference in local protein concentration in the two microcompartments (HSA K=0.25; Cp=0.05 μM, Cd=0.2 μM). The vesicle in the lower panel had a similar difference in local concentration, K = 0.27. We note that it was not possible to simultaneously image all four molecules of interest in these experiments (HSA, PEG, dextran, and lipid). Therefore, labeled PEG and dextran polymers were added in separate experiments (top and bottom panels of Figure 2). HSA and lipid channels were necessary to determine protein localization and vesicle morphology, respectively. Morphological features of interest included not only the overall shape of vesicles but also the presence and location of any lipid material inside the outer membrane. We note that lamellarity of the outer membrane was not assessed, but rather we selected for study only those vesicles that appeared unior oligolamellar based on the confocal fluorescence microscopy images. We have shown one spherical and one budded vesicle in Figure 2; however, as noted above, both vesicle morphologies were present in both experiments. The mean HSA K for all of the vesicles analyzed in this experiment was 0.25 ( 0.09 (N = 35 vesicles) for the batch in which labeled PEG was incorporated and 0.37 ( 0.14 (N=40) for the batch with labeled dextran. Although these values are statistically different at the 95% confidence level, we interpret this as arising from batch-to-batch variations in synthesis rather than any difference due to which polymer was spiked with a labeled analogue.27 Variations in protein partitioning are expected; both the encapsulation of the ATPS-forming polymers and the degree of volume loss upon exposure to hypertonic sucrose are not identical for all vesicles, and hence, 5700 DOI: 10.1021/la903800e

some encapsulated ATPS compositions will produce better protein partitioning than others. In general, higher concentrations of phase-forming polymers are conducive to better partitioning in bulk ATPS experiments.31-34,54 GVs with the highest K values will have the highest interfacial tension between the phases and in some cases even cause lipid material to aggregate at the interface.27,29,32,33,55 For artificial cells formed at pH 6.5, lipid material could often be found be at the interface between the PEG-rich and dextran-rich regions. Examples are shown in Supporting Figure 1 (Supporting Information). We attempted to relate the partitioning of HSA to the concentrations of PEG and dextran polymers in the encapsulated phases and to the partitioning of the polymers between the phases. Because labeled PEG and dextran were not present in the same experiment, we could not measure total polymer concentrations for each vesicle. Our best indicator of ATPS composition was therefore partitioning of the polymers themselves. Figure 3 shows the relationship between partitioning of HSA-AF488 and labeled PEG (left) or dextran (right). Considerable variation in partitioning of both HSA and the polymers was observed between vesicles within a batch. Protein partitioning was more strongly correlated with the distribution of dextran (correlation coefficient, r=0.83) than with that of PEG (r=0.36). The concentration of dextran in the PEG-rich phase was also correlated with HSA K value (r= 0.68); however, neither the concentration of dextran in the PEG-rich phase nor the concentration of PEG in either phase showed much correlation (i.e., all |r| < 0.5). An explanation for dextran K being a stronger indicator of HSA partitioning than PEG K could be that the protein is more similar to dextran in its surface interactions with the solvent.56-58 Protein Localization in Artificial Cells Formed at pH 4.1. Based on experiments in bulk ATPS and information from the literature, we expected that the HSA would localize in the PEG-rich compartment of artificial cells formed at pH 4.1. Figure 4 and Supporting Figure 2 (Supporting Information) show representative images of artificial cells observed at pH 4.1. There were three variations in HSA localization seen at pH 4.1: (1) all of the HSA was at the aqueous/aqueous interface between the PEG-rich and dextran-rich phases, (2) some of the HSA was at the ATPS interface, while some partitioned to either phase, or (3) all of the HSA partitioned to either the PEG-rich or dextran-rich phases. The propensity of HSA to assemble at the aqueous/aqueous interface (54) Long, M. S.; Keating, C. D. Anal. Chem. 2006, 78, 379–386. (55) Li, Y. Phase separation in giant vesicles. Ph.D. Dissertation, Max Plank Institute of Colloids and Interfaces, Science Park Golm, Potsdam, Germany, November 2008. (56) Arakawa, T.; Timasheff, S. N. Biochemistry 1982, 21, 6536–6544. (57) Back, J. F.; Oakenfull, D.; Smith, M. B. Biochemistry 1979, 18, 5191–5196. (58) Farruggia, B.; Nerli, B.; Di Nucci, H.; Rigatusso, R.; Pico, G. Int. J. Biol. Macromol. 1999, 26, 23–33.

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Figure 5. Histograms comparing the partition coefficient (ln K) of HSA-AF488 in two-compartment artificial cells formed at different pH.

Figure 4. Fluorescently labeled HSA (left panels) and PEG (right panels) partitioning in two-compartment vesicles formed at pH 4.1. Fluorescence images have been overlaid and false-colored. Green indicates HSA fluorescence, blue indicates PEG fluorescence, and red indicates lipid fluorescence (DOPE-rhodamine). Scale bar is 5 μm.

is consistent with aggregation of the protein, which is in a partially acid-expanded (i.e., partially denatured) state at this pH.31-34,59 Fluorescence line scans were again used to quantify local HSA concentration (see Supporting Figure 2, Supporting Information). The average HSA K value for all 36 GVs imaged (including all morphologies and HSA partitioning variations) was 2.17 ( 0.85, indicating accumulation in the PEG-rich compartment. The large standard deviation in mean K is due not only to differences in polymer composition within the different vesicles52,53 but also to variability in the degree of localization to the interface. Of the 36 GVs imaged in this experiment, seven had all of the HSA aggregated at the interface with an average K of 1.13 ( 0.2. Twelve GVs had HSA aggregation at the interface along with some HSA in the PEG-rich or dextran-rich phases, with an average of 1.8 ( 0.23. The remaining GVs had no discernible HSA at the interface, with an average K value of 2.85 ( 0.67. Supporting Figure 3 (Supporting Information) shows the relationship between partitioning of HSA and PEG or dextran at pH 4.1. There is a slight positive correlation (r=0.36) between HSA K and PEG K and a stronger correlation between HSA and labeled dextran partitioning (r=0.59). This suggests that, even in its more hydrophobic, acid-expanded conformation, the HSA partitions more similarly to the dextran polymer than to PEG. (59) Goldberg, A. L. Nature 2003, 426, 895–899.

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Although protein localization into the PEG-rich or dextranrich compartment of our artificial cells is heterogeneous, samples prepared at pH 4.1 differ substantially from those prepared at pH 6.5. Figure 5 shows a histogram of ln K for HSA-AF488 in twocompartment vesicles formed at each pH; the two populations do not overlap. We conclude from these data that pH change is a viable means of controlling the local concentration of protein within our two-compartment artificial cells. Relocalization of HSA in Response to pH Change. The pH-dependent localization of HSA-AF488 in artificial cells with coexisting PEG-rich and dextran-rich compartments observed above suggests that pH change could be used to move protein from one compartment to the other. Hence, we attempted to renature partially acid-expanded HSA that was already encapsulated in a two-compartment vesicle by increasing the external pH from 4.1 to 6.5. Figure 6 shows representative sequential confocal microscopy images of a budded artificial cell formed at pH 4.1 (HSA partially denatured) and changed to pH 6.5 (where HSA in a native state) along with a plot showing the change in HSA partitioning (ln K) over time. Prior to the increase in pH, HSAAF488 (green) was localized to the PEG-rich phase (K=2.8, Cp= 0.2 μM, Cd=0.07 μM) and the interface. By 93 s after the change to pH 6.5, the HSA shows reduced partitioning to the PEG-rich phase (K=1.5) and to the interface, and by 104 s it is slightly more concentrated in the dextran-rich phase (K=0.75). Just 133 s after the pH change, HSA relocalization to the dextran-rich phase is complete (K=0.3, Cp=0.06 μM, Cd=0.21 μM), and lipid material can be seen forming at the interface.55 Accumulation of lipid material at the interface was also seen in a subset of the artificial cells formed at pH 6.5, as mentioned above. The overall morphology of this vesicle also changes during this process due to changes in the balance of interfacial tensions as the HSA that had been accumulated at the aqueous/aqueous interface is replaced by membranous material. We observed different time scales for protein relocalization in different vesicles. For example, in Figure 6, relocalization to the dextran-rich phase was not complete until over 100 s, while in Supporting Figure 4 (Supporting Information) no further change was observed after 30 s; other individual vesicles had intermediate relocalization times. The time scale of protein relocalization depends on several factors, including vesicle size and polymer content, the latter of which impacts both the degree of partitioning DOI: 10.1021/la903800e

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Figure 6. HSA relocalizes from the PEG-rich to the dexran-rich compartment of an artificial cell in response to a pH change from pH 4.1 to 6.5. (Top) Sequential confocal fluorescence images of a two-compartment vesicle formed at pH 4.1 and changed to pH 6.5. HSA-AF488 fluorescence is indicated in green, and DOPE-rhodamine is in red. (Bottom) HSA partitioning (ln K) over time. The line was added to guide the eye. Scale bar is 5 μm.

and the viscosity in both compartments of the artificial cells. Additionally, the presence of aggregated protein and/or lipid material at the interface can hinder diffusion between the PEGrich and dextran-rich compartments. We note that partitioning of HSA in Figure 4 continues to change after 104 s, when the line of membrane appears to separate the two compartments. Thus, this membranous material does not represent a diffusion barrier to HSA. We do not envision this material as a bilayer separating the two aqueous compartments; Li and co-workers have been able to image networks of lipid nanotubes at the interface of an encapsulated PEG/dextran ATPS by orienting the vesicles with the dextran-rich end below the PEG-rich end, so that the interface lies in a plane accessible to the confocal microscope.55 We also attempted to perform the experiment in reverse, that is, by changing the external pH from 6.5 to 4.1, to cause destabilization of the HSA and relocalization to the PEG-rich phase. However, changing the external solution from pH 6.5 to 4.1 resulted in destabilization of the GVs; no acceptable ATPScontaining GVs were found after the pH change. This was unexpected, since two-compartment vesicles could be formed at pH 4.1 without difficulty and could be transitioned between the two pH values when no HSA was encapsulated (Supporting Figure 5, Supporting Information). Presumably, the difficulty in changing from pH 6.5 to 4.1 in our experiments is related to acid-denaturation-induced HSA interactions with the aqueous/ aqueous interface and/or membrane. Since HSA conformation can also be altered by transitioning to more basic pH, we tested whether generation of the base-denatured form of HSA would lead to relocalization into the PEG-rich compartment of the artificial cells. HSA-AF488 relocalization 5702 DOI: 10.1021/la903800e

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Figure 7. HSA relocalizes from the dextran-rich compartment to the PEG-rich compartment of an artificial cell in response to a pH change from pH 6.5 to 12. (Top) Sequential confocal fluorescence images of a two-compartment vesicle formed at pH 6.5 and changed to pH 12. HSA-AF488 fluorescence is indicated in green, and DOPE-rhodamine is in red. (Bottom) HSA partitioning (ln K) over time. The line was added to guide the eye. Scale bar is 5 μm.

can be seen in Figure 7, which shows sequential confocal fluorescence images of a two-compartment vesicle formed at pH 6.5 and changed to pH 12. Before the pH change, HSA was partitioned into the dextran-rich compartment with K=0.55 (Cp =0.14 μM, Cd=0.26 μM). At 80 s after pH change, the HSA K is near unity, after which HSA-AF488 is localized into the PEGrich phase more clearly over time. By 290 s, relocalization is complete (Cp=0.23 μM, Cd=0.1 μM (K=2.33)). While protein relocalization in most of the vesicles was complete in under 100 s when changing from pH 4.1 to 6.5, relocalization after a pH change from 6.5 to 12 took between 157 and 290 s. This could be because of slower diffusion caused by aggregation of the basedenatured HSA and/or because the HSA interacts with the interface before moving into the PEG-rich phase. Generality of pH-Responsive Protein Relocalization. Protein conformation is generally responsive to pH. We therefore tested whether this simple approach could be applied to control local concentrations of other proteins in two-compartment artificial cells. We selected proteins with a range of molecular weights (MW) and isoelectric points (pI), as these factors can also impact partitioning in ATPSs. Table 1 lists the molecular weight (MW), isoelectric point (pI), and partitioning coefficient for the proteins investigated here as a function of pH both during vesicle (60) Devlin, G. L.; Chow, M. K. M.; Howlett, G. J.; Bottomley, S. P. J. Mol. Biol. 2002, 324, 859–870.

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Article Table 1. Localization of Proteins Encapsulated in Two-Compartment Artificial Cells as a Function of pH

protein

MW (kDa)

pI

K, pH 4.1

K, pH 6.5

K, pH 4.1 to 6.5

1.8 ( 1.3 0.25 ( 0.09 0.54 ( 0.33 HSA 67 4.9-5.1 1.8 ( 1.4 0.38 ( 0.07 0.53 ( 0.3 AAT 52 4.9-5.1a b 1.9 ( 1.4 0.56 ( 0.12 ;e fibrinogen 340 5.5-6.1 c all at interface 0.4 ( 0.1 all at interface R-Crystallin 800 5.6-6.7 1.4 ( 0.2 1.0 ( 0.2 1.4 ( 0.3 HRP 44 8.8d a Reference 43. b Reference 61. c Reference 62. d Reference 65. e Did not attempt. a

formation and after a change in the external pH. R1-Antitrypsin (AAT) has many similarities to HSA; its MW and pI are quite similar, while all of the charges and partitioning habits are also the same.43,60 Fibrinogen and R-Crystallin were selected, since they have much larger MWs than HSA or AAT, but similar pI.61,62 These proteins have also been shown to have pH-dependent conformational changes similar to those of HSA.63,64 The pH-dependent partitioning of these larger proteins was qualitatively similar to that of HSA and AAT, demonstrating the generality of the conformation-responsive localization approach. Supporting Figure 6 (Supporting Information) shows confocal microscopy images for R-Crystallin-AF488 formed at pH 4.1 and 6.5. At pH 4.1, this protein accumulated almost exclusively at the aqueous/aqueous interface. At pH 6.5, R-Crystallin was localized in the dextran-rich compartment, with mean K=0.4 ( 0.1. Larger proteins and biomolecules aggregate at bulk ATPS interfaces more frequently than their smaller counterparts.31-34,66 However, one interesting note is that while artificial cells containing either HSA or AAT did not survive the transition from pH 6.5 to 4.1, those containing fibrinogen or R-Crystallin remained intact. We also studied the enzyme horseradish peroxidase (HRP) due to its much higher pI (8.8).65 The surface charge on a protein is dependent on its pI and the pH of the surrounding solution, which can dictate its localization in an ATPS, especially in those containing one or more charged phase-forming polymers.31,67 In our ATPS containing only neutral polymers, protein partitioning was more dependent on protein conformation, or more precisely whether it was in a native or partially/fully denatured state.

Conclusions We have demonstrated that protein localization within giant vesicles encapsulating a PEG/dextran aqueous two-phase system can be dynamically controlled via an external chemical stimulus. Here, the preferential localization of human serum albumin and other proteins to a particular aqueous phase encapsulated within a giant vesicle was caused primarily by changes to the conformation of the protein as a result of an external pH change. This control over local concentration within ATPS-encapsulated giant vesicles occurred over short time scales (seconds to minutes), and protein relocalization could be followed with confocal (61) (a) Fuss, C. M.; Palmaz, J. C.; Sprague, E. A. J. Vasc. Interventional Radiol. 2001, 12, 677–682. (b) Ortega-Vinuesa, J. L.; Tengvall, P.; Lundstrom, I. Thin Solid Films 1998, 324, 257–273. (62) (a) Bera, S.; Ghosh, S. K. Biophys. Chem. 1998, 70, 147–160. (b) Horwitz, J. Exp. Eye Res. 2009, 88, 190–194. (63) (a) Stevens, A.; Augusteyn, R. C. Biophys. J. 1993, 65, 1648–1655. (b) Goncalves, S.; Santos, N. C.; Martins-Silva, J.; Saldanha, C. J. Fluoresc. 2006, 16, 207–213. (64) Chattopadhyay, K.; Mazumdar, S. Biochemistry 2000, 39, 263–270. (65) (a) Cans, A.-S.; Dean, S. L.; Reyes, F. E.; Keating, C. D. Nanobiotechnology 2007, 3, 12–22. (b) Delince, H.; Radola, B. J. Biochim. Biophys. Acta 1970, 200, 404– 407. (66) Helfrich, M. R.; El-Kouedi, M.; Etherton, M. R.; Keating, C. D. Langmuir 2005, 21, 8478–8486. (67) (a) Johansson, G.; Hartman, A.; Albertsson, P.-A. Eur. J. Biochem. 1973, 33, 379–386. (b) Johansson, G. Aqueous Two-Phase Systems: Methods and Protocols. Methods in Biotechnology; Hatti-Kaul, R., Ed.; Humana Press: Totowa, NJ, 2000; Vol. 11, pp 303-313.

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K, pH 6.5 to 4.1

K, pH 6.5 to 12

vesicles did not survive vesicles did not survive 0.9 ( 0.2 all at interface 1.3 ( 0.5

3.1 ( 1.3 2.4 ( 1.8 3.8 ( 1.9 2.7 ( 1.3 4.8 ( 1.8

fluorescence microscopy. Relocalization via partitioning in response to pH-induced conformational changes was possible for several different proteins, suggesting that this could be a very general approach. It should be noted that while we did not monitor the bioactivity of the proteins, the acid- and baseexpanded forms which partition into the PEG-rich phase domains are partially or fully denatured, and as such can be expected to have much lower activity as compared to their native counterparts near neutral pH, which localize to the dextran-rich compartments. Thus, it should be possible to control not only local concentration but also protein function in this manner. Here, pH was used to induce the conformational changes that triggered relocalization to a different microcompartment. Other stimuli more selective to specific proteins or conformations could be used, and would in many cases be more biologically relevant. In biological systems, almost all processes are dependent on pH, which is tightly regulated and critical for a wide range of cellular functions, including metabolism, membrane transport, and the innate immune response to microbial infection.68,69 It is therefore unlikely that large pH changes such as those used here would be a viable means of controlling localization in a living cell. Nonetheless, these results demonstrate that protein conformational changes can lead to relocalization in our simple two-compartment artificial cells; more elaborate means of inducing conformational changes could be substituted in future experiments. Several recent studies have reported transient compartmentalization of biomacromolecules within living cells.1,4,9,10 The primitive two-compartment artificial cells presented here provide an experimental model system in which fundamental consequences of protein microcompartmentation, for example, local enzymatic activities, can be explored. With this model, it is possible to change the local protein concentration in a cell-sized, macromolecularly crowded environment in real time and observe the results. Unlike in our prior work on ATPS-containing vesicles, here protein localization was based on conformation rather than affinity with one of the polymers, the relocalization was in response to a chemical stimulus, and extensive reorganization of the model cytoplasm (i.e., phase transition) was not required. Finally, we note that phase separation in living cells could lead to the microcompartmentation of proteins, nucleic acids, and other biomacromolecules in response to an external chemical stimulus such as a change in pH, ion, or other molecule concentration. Certainly, other mechanisms for microcompartmentation are at work in biological cells, for example, encapsulation within organelles or adhesion to form multienzyme complexes. However, both the biophysics of the intracellular environment14,70and recent in vivo work9,71 suggest that aqueous phase separation (68) Bizzarri, R.; Serresi, M.; Luin, S.; Beltram, F. Anal. Bioanal. Chem. 2009, 393, 1107–1122. (69) Yeung, T.; Touret, N.; Grinstein, S. Curr. Opin. Microbiol. 2005, 8, 350– 358. (70) Walter, H., Brooks, D. E., Srere, P. A., Eds. Microcompartmentation and phase separation in cytoplasm. International Review of Cytology; Academic Press: San Diego, CA, 2000; Vol. 192. (71) Ge, X.; Conley, A. J.; Brandle, J. E.; Truant, R.; Filipe, C. D. M. J. Am. Chem. Soc. 2009, 131, 9094–9099.

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could occur in living cells. If coexisting aqueous phase domains did form intracellularly, they would provide an additional route to microcompartmentation in the cell.

Materials and Methods Chemicals and Materials. L-R-Phosphatidylcholine (egg PC), 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (sodium salt) (DOPG), and 1,2-dioleoyl-sn-glycero-3(phosphoethanolamineN-lissamine rhodamine b sulfonyl) (ammonium salt) (DOPErhodamine) were purchased as CHCl3 solutions from Avanti Polar Lipids, Inc. (Alabaster, AL). Poly(ethylene glycol) (PEG) 8 kDa, dextran 500 kDa, 2-(N-morpholino)ethanesulfonic acid (MES) buffer salts, R-Crystallin from bovine eye lens, and horseradish peroxidase, type VI (HRP) were from Sigma Chemical Co. (St. Louis, MO). An Alexa Fluor (AF) 488 labeling kit, AF488, AF633, and AF647 carboxylic acid, succinimidyl esters, fibrinogen-AF488 from human plasma, amino dextran 500 kDa, and 8-hydroxypyrene-1,3,6-trisulfonic acid, trisodium salt (HPTS or pyranine) were purchased from Molecular Probes, Inc. (Eugene, OR). Human serum albumin (HSA) and R1-antitrypsin from human plasma (AAT) were from Calbiochem (San Diego, CA). Amino PEG 20 kDa and amino PEG 5 kDa were purchased from Nektar Therapeutics (Huntsville, AL). Water used in these experiments was deionized to a resistance of 18.2 MΩ with a NANOpure Diamond water system from Barnstead International (Dubuque, IA). Bulk Polymer Solutions and Analysis. ATPS phase diagrams were determined by cloud point titration for PEG 8 kDa and dextran 500 kDa in 1 mM MES buffer and water as described by Albertsson.32 The low concentration of buffer was used to facilitate formation of GVs in later experiments; GV formation is inhibited by increased ionic strength. A small amount of strong acid or base was added to adjust the pH to 4.1 and 12, respectively. From these phase diagrams, we selected 3.9 wt % PEG 8 kDa and 3.9 wt % dextran 500 kDa for use throughout these experiments, because the phase transition temperature lies between 37 and 25 °C, allowing for encapsulation in artificial cells (described below). Bulk partitioning of biomolecules and polymers between the PEG-rich and dextran-rich phases was determined by fluorescence analysis with a Jobin Yvon Horiba FL3-21 fluorimeter. Preparation of Two-Compartment Vesicles. We used the gentle hydration method to form lipid vesicles, as previously described.52,53,72 In a typical procedure, a chloroform solution of lipids containing a 3:7 molar ratio of egg PC/DOPG was prepared at a concentration of 0.26 mg/mL with 0.05 mol % DOPE-rhodamine. In a 10  75 mm test tube (Durex borosilicate glass, VWR Int., West Chester, PA), the above lipid solution was added and dried under Ar (g) to form a thin, uniform lipid film, and then the vials were vacuum desiccated for ∼2 h to remove any residual organic solvent. Then the 3.9 wt % PEG 8 kDa/3.9 wt % dextran 500 kDa solution (as a homogeneous solution/not phaseseparated) containing fluorescent protein, PEG, dextran, or some combination thereof heated to 37 °C was added along the wall of the tube, and the lipids were hydrated for 48 h at this temperature. Following incubation at 37 °C, the solutions were incubated at 4 °C for 12 h and then finally left to equilibrate at room temperature. Two-compartment vesicles (10 μL) were collected at the bulk interface and suspended in the top PEG-rich phase (150 μL) from the original bulk polymer solution/or from a bulk polymer solution of a different pH (lipid- and fluorophore-free). To this was added an aliquot of sucrose (10 μL) to a final concentration of 31 mM, and then the two-compartment vesicles were transferred to a microscope slide for analysis. We verified that PEG 20K-AF488 and PEG 5K-AF647 did not show any significant change in fluorescence or phase preference when the pH was changed; thus, any changes in phase preference when (72) Akashi, K.; Miyata, H.; Itoh, H.; Kinosita, K., Jr. Biophys. J. 1996, 71, 3242–3250.

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these fluorescent tags were attached to a protein were due to the presence of the protein and not the fluorescent tag.

Following Protein Relocalization in Artificial Cells in Real Time. In cases where protein relocalization due to external pH change was monitored in real time, two-compartment vesicles (5 μL) were added to the top PEG-rich phase from the original bulk polymer solution (10 μL) containing 31 mM sucrose (e.g., vesicles formed at pH 4.1 were diluted in pH 4.1 PEG-rich phase) and then transferred to a microscope slide and covered with an Anodisc 25 membrane (0.2 μm diameter pores) from Whatman International Ltd., Maidstone, England. Then 150 μL of PEG-rich phase of another pH (i.e., pH 6.5 added to vesicles formed in pH 4.1 buffer) containing 31 mM sucrose was added through the Anodisc membrane, and the protein relocalization was followed in real time.

Monitoring pH Change Across Giant Vesicle Membranes. Vesicles containing 0.5 mM of the pH-sensitive dye HPTS/ pyranine and 24 μM of pH-insensitive 5 kDa PEG-AF647 in 1 mM MES buffer at pH 4.1 and 6.5 were prepared as the twocompartment vesicles above, although after incubation for 48 h at 37 °C the solutions were left to cool to room temperature (without incubation at 4 °C). It had previously been reported that pH can be changed across vesicle membranes in the absence of any ion channels or proton pumps; therefore, we did not include these in our system.73 Supporting Figure 7 (Supporting Information) shows representative confocal fluorescence images of these GVs formed at pH 4.1 and 6.5 before and after the external pH was changed. The average fluorescence intensities for pyranine and AF647 are listed in Supporting Table 1 (Supporting Information). The changes in pyranine fluorescence confirm that changing the external pH results in a pH change inside the vesicles. Microscopy. Imaging was performed using an LSM-5 Pascal Laser Scanning confocal microscope from Carl Zeiss, Inc. (Oberkochen, Germany) with a Plan-Apochromat 63 oil immersion objective (1.4 NA) and Pascal software as previously described.50,51 Excitation wavelengths for the dyes were as follows: HPTS/pyranine, 458 nm; AF488, 488 nm; AF633 and AF647, 633 nm; rhodamine, 543 nm. At least 35 vesicles were analyzed per batch. Only vesicles that were greater than 5 μm in diameter with both the PEG-rich and dextran-rich phases clearly visible were analyzed in the case of two-compartment vesicles, and only vesicles greater than 5 μm in diameter that appeared to be uni- or oligo-lamellar were analyzed. Solute concentrations were determined from their fluorescence intensities by taking a fluorescence line scan across each twocompartment vesicle (spanning both the PEG-rich and dextranrich phases) as previously reported.27-30 The concentration of labeled macromolecules in each phase (or buffer) was determined directly from the fluorescence intensities via a calibration curve of the labeled macromolecule or dye free in solution. The extent of protein localization to a particular phase was quantified by the partition coefficient, K, which is defined as the concentration of labeled macromolecule in the PEG-rich phase (Cp) as compared to that in the dextran-rich phase (Cd) (K = Cp/Cd). All images within this paper have been false-colored for clarity. Circular Dichroism. Samples for circular dichroism were treated the same as ATPS/GV samples in terms of incubation times and temperatures (see above). Circular dichroism spectra for 0.04 mg/mL HSA in buffer and in the PEG-rich and dextranrich phases of an ATPS that had an overall composition of 3.9 wt % PEG 8 kDa and 3.9 wt % dextran 500 kDa at various pH values were collected on a JASCO 810 spectrapolarimeter from 260 to 200 nm with a step of 1.0 nm and a 2 s averaging time. All CD spectra in this paper are the average of three scans. (73) (a) Paula, S.; Volkov, A. G.; Van Hoek, A. N.; Haines, T. N.; Deamer, D. W. Biophys. J. 1995, 70, 339–348. (b) Rossingol, M.; Thomas, P.; Grignon, C. Biochim. Biophys. Acta 1982, 684, 195–199. (c) Nichols, J. W.; Deamer, D. W. Proc. Natl. Acad. Sci. U.S.A. 1980, 77, 2038–2042.

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Acknowledgment. This work was supported by the National Science Foundation (CHE-0750196) and Pennsylvania State University. C.D.K. also acknowledges support from a Beckman Foundation Young Investigator Award, a Sloan Fellowship, and a Dreyfus Teacher-Scholar Award. E.L.G. was partially supported by funds from the Women in Science and Engineering Research (WISER) program. We thank TJ Mullen for his help with creating a MatLab program for data analysis, Jackie Keighron for assistance with CD measurements, and Clint Jones for suggesting HSA and AAT as proteins to use in these experiments. Supporting Information Available: Confocal fluorescence images of two-compartment vesicles formed at pH 6.5 with lipid material at the aqueous/aqueous interface, confocal

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Article fluorescence images and corresponding fluorescence linescans of two-compartment vesicles formed at pH 4.1, plots showing the relationship between partition coefficients of HSA and labeled PEG or dextran in two-compartment vesicles formed at pH 4.1, sequential confocal fluorescence images of HSA localization in a two-compartment vesicle formed at pH 4.1 and changed to pH 6.5, confocal fluorescence images of two-compartment vesicles containing 20 kDa PEG-AF488 surviving the transition from pH 6.5 to 4.1, confocal fluorescence images of R-Crystallin encapsulated in two-compartment vesicles formed at pH 4.1 and 6.5, and confocal fluorescence images and corresponding fluorescence values of pyranine and AF647 encapsulated in vesicles formed at and changed to various pH values. This material is available free of charge via the Internet at http://pubs.acs.org.

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