Article pubs.acs.org/JACS
Cite This: J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Pulsatile Gating of Giant Vesicles Containing Macromolecular Crowding Agents Induced by Colligative Nonideality Wan-Chih Su,†,# Douglas L. Gettel,‡,# Morgan Chabanon,⊥,# Padmini Rangamani,*,⊥ and Atul N. Parikh*,†,‡,§,∥ †
Departments of Chemistry, ‡Chemical Engineering, §Biomedical Engineering, and ∥Materials Science & Engineering, University of California, Davis, California 95616 United States ⊥ Department of Mechanical and Aerospace Engineering, University of California San Diego, La Jolla, California 92093, United States S Supporting Information *
ABSTRACT: The ability of large macromolecules to exhibit nontrivial deviations in colligative properties of their aqueous solutions is well-appreciated in polymer physics. Here, we show that this colligative nonideality subjects giant lipid vesicles containing inert macromolecular crowding agents to osmotic pressure differentials when bathed in small-molecule osmolytes at comparable concentrations. The ensuing influx of water across the semipermeable membrane induces characteristic swell-burst cycles: here, cyclical and damped oscillations in size, tension, and membrane phase separation occur en route to equilibration. Mediated by synchronized formation of transient pores, these cycles orchestrate pulsewise ejection of macromolecules from the vesicular interior reducing the osmotic differential in a stepwise manner. These experimental findings are fully corroborated by a theoretical model derived by explicitly incorporating the contributions of the solution viscosity, solute diffusivity, and the colligative nonideality of the osmotic pressure in a previously reported continuum description. Simulations based on this model account for the differences in the details of the noncolligatively induced swell-burst cycles, including numbers and periods of the repeating cycles, as well as pore lifetimes. Taken together, our observations recapitulate behaviors of vesicles and red blood cells experiencing sudden osmotic shocks due to large (hundreds of osmolars) differences in the concentrations of small molecule osmolytes and link intravesicular macromolecular crowding with membrane remodeling. They further suggest that any tendency for spontaneous overcrowding in single giant vesicles is opposed by osmotic stresses and requires independent specific interactions, such as associative chemical interactions or those between the crowders and the membrane boundary.
■
Moreover, the cumulative weight of these findingsderived primarily by the in vitro application of model vesicular compartments and artificial crowding agents (e.g., polyethylene glycol (PEG), dextran, or Ficoll)lend support to the notion that living cells actively regulate the excluded volume and, thus, the complementary proportion of their solute-accessible free volume to optimize kinetics and equilibria of intracellular biochemical processes achieving a so-called crowding homeostasis.9 Despite the significant progress these findings portend, the question of how excluded volume effects affect the behaviors of the bounding membrane remain largely unknown.10 From the vantage point of the membrane, macromolecular crowding in the encapsulated milieu presents a variety of indirect perturbations. First, the physical crowding of macromolecules in solution gives rise to nonideal, colloidal osmotic pressure (Π),11 a phenomenon well-appreciated in polymer physics. The van’t Hoff’s law, which treats a polymer solution as an ideal gas ((Π = RTc) where c = concentration mol·m−3, R = gas constant, and T = temperature) is valid for ideal solutions
INTRODUCTION The interior of a living cell is dense, packed with large macromolecules including proteins, polysaccharides, and nucleic acids as well as cytoskeletal filaments and organelles.1 This is perhaps best exemplified by the cytoplasm of Escherichia coli in which the volume occupancy of ∼15−20% by proteins (roughly 200 mg/mL) corresponds to the dramatically low surface-to-surface separation of 2 nm for an average protein 5 nm in diameter.2 In this highly crowded environment, “excluded volume” effectsarising from the inaccessible space preoccupied by neighboring moleculesresult in steric repulsion, depletion attraction, and reduced translational degrees of freedom, all of which have important ramifications for the biochemical workings of a living cell.3 Indeed, a rapidly growing body of experimental and theoretical efforts now establishes how macromolecular crowding acts to (1) alter protein conformations, aggregation, and stability;4 (2) modulate intermolecular interactions between proteins, sugars, nucleotides, and even organelles;5 (3) render protein diffusion anomalous and subdiffusive;6 and (4) induce liquid−liquid phase separation or microcompartmentalization.7 Taken together, these findings support a picture of cytosolic fluid as a glassy, nonergodic, and viscous emulsion of coexisting fluids.8 © XXXX American Chemical Society
Received: September 28, 2017
A
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society
crowding agents is sufficient to stress the bounding membrane at biologically relevant concentrations: the membrane responds by producing transient, but long-lived, pulsewise oscillations in size, tension, poration, and the spatial organization of membrane molecules in a cyclical manner en route to osmotic equilibration. These observations recapitulate the behaviors of lipid vesicles25 and red blood cells26 experiencing sudden osmotic shocks due to large (hundreds of osmolars) differences in the concentrations of small molecule osmolytes, and importantly link intravesicular macromolecular crowding with the membrane deformations at the compartmental interface.
only and must be extended to incorporate noncolligative effects.11 The osmotic pressure of a polymer solution of molar concentration c can be expressed as Π = RT[c + A 2 (Mc)2 ]
(1)
where A2 is the second virial coefficient (mol m3/ g2) and M is the molecular weight of the polymer.12,13 For small molecular weight solutes (e.g., KCl), the nonlinear term vanishes and the van’t Hoff’s lawwhere the osmotic pressure depends only on the concentration and not on the chemical identity of the soluteis fully recovered. For moderate molecular weight solutes, such as glucose and sucrose, colligative nonideality is weak14 and practically inconsequential at low concentrations in the millimolar range. By contrast, for solutions containing macromolecular solutes of large molecular weights, the nonlinear term involving the second virial coefficient in Eq 1 becomes significant even at moderate concentrations, and colligative nonideality becomes dominant. This solvent−solute specific phenomenon is due to solvent-mediated monomer− monomer interactions: these include interpenetration of polymers and changes in persistence lengths as well as preferential binding of water to solute and the exclusion of neighboring solute molecules from each solute molecule’s local “hydration shell”.15 These noncolligative factors become exacerbated at elevated concentrations corresponding to the crowding scenario.15 Second, nonspecific, depletion interactions resulting from the excluded volume effects16 at the membrane boundary induce additional perturbations. Specifically, the preferential exclusion of the macromolecules from the vicinity of the membrane surface17 introduces an additional contribution to the osmotic imbalance, which in turn favors membrane conformations that maximize the translational entropy of the trapped macromolecules.18 It has been long-recognized that such depletion attraction can drive membrane stacking19 and induce membrane fusion20 when the crowding agents are on the outside of the vesicles. More recent studies have shown that when macromolecules are encapsulated in the vesicle interior, depletion forces can induce large-scale membrane deformations including topological division.21,22 Third, highly hygroscopic19b,20 crowding agents, such as PEG, further complicate the picture by competing with the headgroups of membrane lipids for binding water, thereby inducing membrane dehydration.23 This decreases the effective size of the lipid headgroup, creating tighter packing, reducing lateral fluidity, and compressing the membrane.24 Taken together, the considerations above support the notion that the presence of crowding agents in the vesicular compartment must exert a complex influence over the bounding membrane. In the work reported here, we describe the behaviors of the membranes of giant vesicles encapsulating physiologically relevant concentrations of macromolecular crowding agents. Our in vitro experimental system comprises ∼20−60 μm giant unilamellar vesicles (GUVs) bounded by a membrane consisting of a phase separating mixtures of phospholipids and cholesterol to nominally identical, isomolar solute concentrations between the inside and the outside, differing only in the fact that the confined solutes are model macromolecular crowding agents7b (i.e., high-molecular weight poly(ethylene)glycol (PEG) or dextran), while the extravesicular medium bathes the small-molecule counterparts (i.e., sucrose or glucose). We find that the colligative nonideality of the osmotic pressure due to the macromolecular nature of the
■
RESULTS AND DISCUSSION We used electroformed GUVs (20−60 μm in diameter)27 consisting of a phase-separating, ternary mixture containing a common unsaturated phospholipid, namely 1-palmitoyl 2oleoyl-sn-1-glycero-3-phosphocholine (POPC), egg-sphingomyelin (SM), and cholesterol (Chol) in a 2:2:1 molar ratio (Supporting Information). Companion experiments were carried out using single-component GUVs made of POPC. In all cases, a small concentration of methoxy(polyethylene glycol) derivatized 1,2-dioleoyl-sn-glycero-3-phosphatidyl ethanol amine (mPEG-2000-DOPE, 2.2 mol %) was added to facilitate reproducible formation of large GUVs, presumably by providing steric repulsion between the membrane lamellae during hydration needed for the topological transition during electroformation. 28 Depending on the exact molecular composition and temperature, membranes of these lipid mixtures equilibrate producing a uniform single phase or exhibit microscopic phase separation,29 the latter characterized by two coexisting liquid phases: a dense phase enriched in SM and Chol designated as the Lo (liquid-ordered) phase and a second, less dense Ld (liquid-disordered) phase consisting primarily of POPC. To discriminate between the Lo and Ld phases by fluorescence microscopy, we doped the GUVs with a small concentration of two complementary phase-sensitive probes, lissamine rhodamine B 1,2-dioleoyl-sn-glycero-3phosphoethanol amine (rhodamine-B DOPE, 1 mol %), which stains the Ld phase30 and NBD-PE (2 mol %), which preferentially labels the Lo phase. To introduce crowding agents in the intravesicular milieu, we encapsulated PEG 8000 (MW: 8000 Da, 14 mM), dextran 10000 (MW: 9,000−11 000 Da, 14 mM), or their mixtures at concentrations mimicking intracellular protein levels. The measurements of osmotic pressure due to polymer solutions using a vapor-pressure deficit osmometer confirm elevated osmotic pressures corresponding to the colligative nonideality of the crowding agents (Figure 1). Specifically, we find that the 14 mM concentrations of PEG 8000 and dextran 10000 produce osmotic pressures corresponding to ∼116 and ∼60 mM sucrose, respectively. Moreover, fitting Eq 1 to these data provides quantitative estimates for the second virial coefficient for PEG 8000 (A2 = 8.18 × 10−9 mol m3/ g2) and dextran 10000 (A2 = 2.32 × 10−9 mol m3/ g2), which are in good agreement with previous reports of identical or comparable solutes.12,13,31 To subject the polymer-containing GUVs above to transvesicular osmotic gradients due to colligative nonideal contributions alone, we substitute the extra-vesicular dispersion medium by an aqueous phase containing osmotically active sucrose at the iso-molar concentration (14 mM) and record the ensuing dynamics (Figure 2a). Pulsatory Dynamics of GUVs containing Macromolecular Crowding Agents. The time-lapse video B
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society
the pulsatory behavior is fully reproduced, ruling out the possibility that the observed behavior is specific to any unique physical or chemical properties of PEG. Furthermore, recognizing that intracellular crowding is such that although the total macromolecular concentration in the cellular interior is substantial, no single crowding agent is present at very high concentrations. To minimally capture this chemical heterogeneity of intracellular crowding, we used a binary mixture of PEG 8000 (7.5 mM) and dextran 10000 (6.4 mM) mimicking intravesicular crowding. Again, qualitatively comparable pulsatory cycles characterized by oscillatory phase separation and size fluctuations (Figure 2d and supplementary video 3) are fully recapitulated, lending further support to the physical basis of the observed vesicle pulsatility. The pulsatile GUV dynamics generated through the noncolligative behavior of crowding agents with those derived by subjecting vesicles to corresponding concentration difference of small molecule osmolytes (114 mM sucrose in 14 mM sucrose bath corresponds to 14 mM PEG 8000 in 14 mM sucrose bath) as identified by osmotic pressure measurements (Figure 1) were compared. Specifically, subjecting 114 mM sucrose containing GUVs to an extra-vesicular medium of 14 mM glucose (Figure 2e and supplementary video 4), qualitatively similar pulsatile cycles follow. A closer look at the cyclical dynamics above reveals that the oscillatory process in all cases dampens with time (Figure 2f− i): The period of oscillation increases with the passage of time or the cycle number. The cycle perioddefined as the time elapsed between two consecutive instances of homogeneous fluorescenceincreases 5- to 10-fold over 90−120 min. Depending on the type of crowding agent used, time spent in each of the two, homogeneous and phase-separated, states lengthens. This slowing down of the cyclical phase separation suggests that the forces driving the oscillatory process (i.e., colloidal osmotic pressure) must weaken with each cycle. This requires a physical mechanism for the efflux of the macromolecular crowding agents across the membrane, which is otherwise impermeable to polymeric solutes at the experimental time scales. Indeed, we find that transient, single microscopic pores, in an apparent synchronization with the oscillating phase separation, periodically open up during the phase-separated (and swollen) state of each cycle suggesting pulsewise ejection of the trapped macromolecular cargo25b,33 (Figure 3 and supplementary video 5). These single pores are large, several micrometers wide, assuming as much as 10−30% vesicle surface area (as judged from a rudimentary geometric analyses of pore sizes estimated from microscopy images) and spatially localized, either in the Ld phase or at the Lo − Ld phase boundary. The pores are short-lived, closing within hundreds of milliseconds after their initial appearance, and form once during each cycle. To probe the transport properties of the transient pores, we carried out experiments doping the bath solution with a small proportion (58 μM) of fluorescently labeled glucose. These measurements reveal little or no influx (supplementary videos 6 and 7) of the labeled osmolyte. This then suggests that the material flow through the open pores is unidirectional, directed to the vesicular exterior driven by the excess Laplace pressure of the interior fluid because of the edge tension at the pore boundaries. This directed transport is perhaps best visualized in structurally complex GUVs, hierarchically embedding smaller “cargo” vesicles within their interior, where, as shown previously,34 the expulsion of the “cargo” vesicle dragged
Figure 1. Determination of second virial coefficients of PEG 8000 and Dextran 10000. Osmotic pressures of PEG 8000 Dextran 10000, measured using vapor pressure deficit osmometry, as a function of molar concentrations (open symbols). Fitting the measured value using the equation for nonideal osmotic pressure of polymer solutions (solid lines) yields second virial coefficients, which estimate for PEG 8000 at A2 = 8.18 × 10−9 mol m3/g2 (R2 = 0.975) and dextran 10000 at A2 = 2.32 × 10−9 mol m3/g2 (R2 = 0.994). For comparison, osmotic pressures estimated using the van’t Hoff equation, valid for ideal solutions, are shown as a dashed line.
(supplementary video 1) and a montage of selected frames (Figure 2b) of confocal fluorescence micrographs documents the dynamics, which follow upon the transfer of PEG 8000 encapsulating GUVs immersed in the surrounding sucroseladen bath at the identical concentration. They reveal that the initial homogeneous texture of the membrane (at optical length scales) is quickly abandoned, replaced by a striking, dynamic pattern of phase separation, rhythmically switching between the optically homogeneous state and the one characterized by surface textures decorated with microscopic domains. A view of the spatial distribution of membrane domains, offered by considering the radial pattern of fluorescence intensity of the membrane boundary, reveals that the domains are not static: they fluctuate in their position, coalesce, and grow during the phase-separated state (Supplementary Figure S2.1). Synchronized with this temporal pattern of phase separation is the periodic oscillation in vesicle size: the largest domains appear when the GUVs are fully swollen and the optically homogeneous state reappears when the GUV reaches its lowest diameters. This cyclical process of swelling and phase separation repeats multiple times (n ≥ 10) for individual GUVs lasting several tens of minutes to hours with domains forming at different spatial positions in different cycles before a quiescent steady-state is reached. This behavior is fully reproducible in multiple experiments (n = 20). In single experiments, the majority of well-formed GUVs (95%, n > 30) exhibit the cyclical dynamics. Because PEG is known to have direct interactions with membrane phospholipids,32 we carried out experiments substituting PEG by a chemically different crowding agent, namely dextran 10000, at the comparable concentrations of 10−15 mM (Figure 2c and supplementary video 2). Here too, C
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society
Figure 2. Pulsatile phase separation of single giant unilamellar vesicles subject to an osmotic imbalance due to colligative nonideality. (a) Schematic representation of crowding-induced pulsewise phase separation, n = cycle number. b-d, Selected frames from the videos of time-lapse fluorescence images (supplementary videos 1−4) of single GUVs encapsulating 14 mM macromolecular crowding agents upon immersion in isomolar sucrose solution: (b) PEG 8000; (c) dextran 10000; (d) 6.0% (w/w) PEG 8000 and 6.4% (w/w) dextran 10000. (e) Reference measurements for GUVs containing 114 mM sucrose inside and 14 mM glucose in the surrounding bath. In all cases, the GUVs imaged consist of a mixture containing POPC:egg-SM:cholesterol (2:2:1), 2.2% DOPE-mPEG, 1% Rho-DOPE, and 2% NBD-PE. All scale bars, 10 μm. (f−i) Corresponding bar plots displaying the time intervals in single and phase-separated states and increase of cycle period during oscillatory domain dynamics. Black bar: period in single phase. Light gray bar: period in the phase separated state. Dark gray bar: total cycle period (see text for details).
radius). This apparently stable pore stays open for about 8 s, during which time the cargo vesicle is expelled. Assuming the cargo vesicle does not significantly perturb the flow field, this expulsion provides a rough estimate for the efflux rate during poration. Following a theoretical analysis based on flow through an open orifice under the action of Laplace pressure, previously developed by Karatekin and co-workers,35 the efflux rate can be used to estimate the edge tension at the pore boundary
toward and through the pore provides a quantifiable measure of the flow field. An example shown in Figure 4 (supplementary video 8) reveals a directed transport of a 5.4 μm radius cargo vesicle within the 30 μm radius mother vesicle translating 26.5 μm distance within the vesicle in ∼8 s with an average velocity of 3.3 μm·s−1 before being expelled. Strikingly, the pore first expands to a maximum radius of about 30% of the vesicle radius before stabilizing at a smaller radius (about 5% of the vesicle D
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society
Figure 4. Osmotically driven transport of cargo vesicles. Selected frames from a time-lapse movie (supplementary video 9) of epifluorescence microscopy images of hierarchical, multicompartment GUVs containing 14 mM PEG 8000 upon immersion in the external dispersion medium containing 14 mM sucrose. GUV are composed of POPC:SM:Ch (2:2:1) with 2.2% DOPE-mPEG labeled with 1% RhoDOPE. Scale bar: 15 μm.
response of the membrane to the transmembrane osmotic pressure gradient. First, the differential in osmotic pressure Δπ across the membrane boundaryin the present case due to the colligative nonideality of the entrapped crowding agents triggers an entropically driven influx of water across the semipermeable membrane boundary, which hinders the concentration equalization of the macromolecular solute but allows for the facile transport of water. As a consequence, the vesicle volume grows and the osmotic pressure difference decreases. The resulting increase in volume-to-area ratio, corresponding to the PdV work stored in the elastic membrane, tenses the membrane. The lateral tension in turn drives spatial segregation of membrane molecules isothermally producing microscopic domains likely by elevating the line tension between coexisting domains or through the effect of pressure on thermodynamic phase separation.25b,38 Beyond a critical membrane tension, the membrane lyses by transiently opening a single microscopic pore producing edge tension (γ) resulting from the hydrophobically determined reorientation of edge lipids into a hemimicelle. The transport of intravesicular fluid through the pore relaxes the tension, which in turn drives poreclosure regenerating topologically closed and optically homogeneous GUVs, but with reduced concentrations of crowders and lowered osmotic pressure difference. Subsequently, the residual trans-membrane osmotic gradient repeats the cycle, which reiterates multiple times until the intravesiclar macromolecular concentrations is sufficiently lowered and the sublytic osmotic pressure difference reached. Together, this well-coordinated sequence of elemental biophysical processes, namely swelling, stretching, domain formation, and poration, constitutes an extraordinary negative feedback loop dissipating the osmotic pressure gradient by cyclically dispersing the crowding agents and achieving osmotic equilibrium in a pulsewise manner. Notably, this cyclical process in the present case is fueled entirely by the colligative nonideal component of the osmotic pressure of the crowded, macromolecular solutions. A comparison of the details of pulsatile dynamics of macromolecule-filled GUVs with those containing sucrose reveal noteworthy differences. Specifically, polymer-filled GUVs pulsate with the pore life times, which are noticeably longer (600−1000 ms vs. 100−200 ms in low viscosity sucrose solution); oscillation periods, which are longer in duration
Figure 3. Formation of microscopic pores during each swell-burst cycle. Selected frames from a video of time-lapse fluorescence images and linearized fluorescence intensity profiles across the pore (supplementary videos 7 and 8) reveal the transient appearance of a single, microscopic pore during a pulsatory swell-burst cycle. The pores appear abruptly when the vesicles are fully swollen and the membrane is phase separated. Immediately after pore opening, vesicle diameters drop instantaneously, and the pores heal in a few frames corresponding to the lifetime of ∼ 600 to 800 ms: (a) GUVs imaged consist of a mixture containing POPC:egg-SM:cholesterol (2:2:1), 2.2% DOPE-mPEG, 1% Rho-DOPE, and 2% NBD-PE, and encapsulate 14 mM PEG 8000 immersed in 14 mM sucrose; (b) viscosity-match control experiment. The GUVs imaged consist of a single-component POPC encapsulating a mixture of 14 mM PEG 8000 and 100 mM sucrose immersed in the surrounding bath of 14 mM PEG 8000 (see the text for details). Arrows are guides to the eye. Scale bar: 10 μm.
according to γ = (3π/2)ηsRVcargo where ηs represents the solution viscosity (10 × 10−3 N·s·m−2), R the vesicle radius (30 μm), and Vcargo the efflux rate (3.3 μm·s−1). Using this analysis, we estimate the edge tension to be 4.7 pN, which is slightly lower but close to the range of 6−20 pN reported previously for fluid lipid bilayers.36 Swell-Burst Cycles Induced by Colligative Nonideality of GUVs Containing Macromolecular Crowding Agents. The pulsatory behaviors of giant vesicles containing macromolecular crowding agents above is in excellent correspondence with the well-known swell-burst dynamics of vesicular compartments subject to hypotonic stresses generated by the concentration difference in small colligative solutes.25b,c,37 Mechanistically, the process involves a finely coordinated E
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society (TPEG/sucrose vs. Tglucose/sucrose, n = 1: 117s vs. 24s); and the number of cycles, fewer (n = 10 vs. n = 33) prior to apparent equilibration. That the lifetimes of transient pores formed in osmotically swollen macromolecule-filled GUVs are longer, when compared to those in sucrose-filled GUVs, can be straightforwardly understood in terms of the interplay between pore dynamics and the viscosity of the encapsulated polymer solution. The buildup of membrane tension in osmotically swollen vesicles, beyond a threshold value, provides a driving force for pore nucleation. The subsequent growth of the pore relieves the membrane tension by two independent mechanisms: (1) by reducing the effective area over which the membrane is spread and (2) by enabling the efflux of the vesicular content due to the excess Laplace pressure, which in turn reduces vesicle volume and thus the membrane area. The former promotes pore-growth while the latter fosters pore-closure. For GUVs swollen due to the activities of small molecule osmolytes such as sucrose, low solution viscosities (∼1 mPa·s) ensure rapid efflux through open pore. As a result, pore growth is prematurely aborted. By contrast, in GUVs crowded with macromolecules, the higher solution viscosity (∼10 mPa·s)39 reduces the efflux rate. As a consequence, the pores grow remaining open for longer periods of time.40 To assess whether the viscosity contrast between the effluxing solution and the vesicular bath contributes to pore lifetimes we carried out a control experiment. Here, GUVs encapsulating 14 mM PEG 8000 and 100 mM sucrose are bathed in 14 mM PEG 8000 solution. The sucrose acts as a source for an equivalent osmotic pressure (See above) and isomolar PEG 8000 concentrations allow to minimize the contrast in solution viscosity. Here too, the pore remains open for 600−800 ms (Figure 3b, supplementary video 9) further confirming that the pore lifetime is primarily determined by the viscosity of the inner polymer-laden solution of the GUVs. Why do polymer-filled GUVs, activated osmotically through colligative nonideality, pulsate less frequently compared to their small molecule counterparts? This seems counterintuitive since polymer solutions have considerably higher solution viscosities and lower solute diffusivities, both of which might be expected to limit the efflux rate during poration and delay equilibration. To understand this intriguing behavior, we first consider the relative roles of diffusion and convection in driving osmotic equilibration in GUVs. During each pore opening event, solute is leaked-out by both convection and diffusion. Their relative contribution can be understood in terms of the Péclet number, defined as Pe R0vL/D, where R0 is the characteristic size of the vesicle, vL is the leak-out velocity, and D is the diffusivity of the solute. Convective transport is characterized by Pe ≫ 1, while diffusion-dominant transport is governed by a regime where Pe ≪ 1. The Péclet number for a vesicle leak-out can be written as Pe = R0ΔpLr/(D3πηs), based on the expression for the vesicle leak-out velocity (Eq S5),41 where ΔpL is the Laplace pressure, r is the pore radius, and ηs is the solution viscosity. PEG 8000 and dextran 10000 solutions have a 10-fold higher viscosity than sucrose solution but also a 10-fold lower solute diffusivity, resulting in roughly the same Péclet number. This then suggests that the relative roles of diffusive and convective transport are comparable for both small molecule and polymeric solutions. This has an important consequence on the dynamics of osmotic pressure during the pulsatile cycles: even with the same initial osmotic pressure, an equivalent decrease of solute concentration will produce a much larger
Figure 5. Effect of nonideal osmotic pressure on pulsatory swell-burst cycles of single GUVs. (a) Osmotic pressure differential as a function of the solute concentration for different inside/outside solution configurations. The expression of the osmotic pressure differential is in Eq 2, while virial coefficients and molecular weights are reported in Supplementary Table S1. (b−g) Model results for the dynamics of vesicle radii b−d, and pore radii e−g. (b, e) 14 mM PEG 8000 inside, 14 mM sucrose outside, (c, f) 14 mM dextran 10000 inside, 14 mM sucrose outside, (d, g) 114 mM sucrose inside, 14 mM glucose outside. All initial vesicle radii are 20 μm. (See the Supporting Information. for details and parameter values used in the model.)
decrease in the osmotic pressure for polymer-filled vesicles than for sucrose-filled vesicles (see Figures 1 and 5a). This is because the colligative nonideality of the osmotic pressure is nonlinearly dependent on macromolecular concentration (see Figures 1 and 5a), decreasing disproportionately with the lowering of intravesicular concentrations. As a result, PEG and dextranfilled vesicles approach the equilibrium state in a manner characterized by lower pulsatility: slower swelling, slower efflux, and longer total cycle periods on the one hand and fewer cycles due to rapid diminution in the osmotic pressure difference compared to those for sucrose-filled vesicles. Theoretical Model for Osmotic Response of GUVs encapsulating Macromolecular Crowding Agents. To further characterize this effect quantitatively, we developed a mathematical model for the dynamics of a vesicle encapsulating a polymer solution, which explicitly incorporates the relative contributions of solution viscosity, solute diffusivity, and colligative nonideality. This model extends on our recent work on vesicles in hypotonic stress25c incorporating the colligative nonideality introduced by the crowded, intravesicular environment. Briefly, the model considers three conservation equations associated with the vesicle volume, the solute molarity, and the pore energy (Supporting Information, Model Development and Simulation). Using Eq 1, we determine the osmotic pressure differential between the inside and the outside of the vesicle as ΔΠ = Π in − Πout = RT[Δc + A2in (M inc in)2 − A 2out (Moutcout)2 ] F
(2)
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society where the subscripts “in” and “out” designate terms related to the inside and the outside of the vesicle, respectively. Figure 5a shows the osmotic pressure differential for the fitted values for the second virial coefficient for PEG 8000, dextran 10000, and for sucrose estimated using data in Figure 1. Simulation results of the swell-burst cycles of vesicles filled with PEG 8000, dextran 10000, and sucrose are shown in (Figure 5b−g). These calculationsrepresenting swell-burst cycles in terms of periodic variations in vesicle size (Figure 5b−d) and pore radii (Figure 5e−g)fully reproduce our experimental observations (Figure 2) above and lend quantitative support to the foregoing inference that colligative nonideality and viscosities of polymer solutions nontrivially couple determining the pathways of osmotic equilibration in polymer-filled GUVs. Note that the predicttion of a long-lived pore in the poration cascade has been theoretically predicted previously42 and ascribed to a “wash-out” effect due to compensating effects of the osmotic influx and Laplace pressure mediated efflux through the open pore.42a Although we have not observed this effect in our experiments, our observations of a rather longer lived pore revealing transport of cargo vesicles (Figure 4 and supplementary video 8) might suggest such a wash out. Salient features of these findingshighlighting the role of intravesicular crowding in inducing nonequilibrium membrane dynamicsoffer important insights in several different contexts. First, our observations extend previous observations of swell-burst dynamics in lipid vesicles produced by large concentration gradients of osmotically active solutes (>100 mM)25b to physiologically relevant concentrations of macromolecules through colligative nonideality. At 14 mM concentration, PEG 8000 (hydrodynamic radius RH ∼ 3.5 nm)43 and dextran 10000 (RH = 1.86 nm)44 have, on average, the surface-to-surface separation of 3−6 nm, which is slightly larger than the average molecular separations in the cytoplasm of E. coli. This is noteworthy since it exemplifies the role of excluded volume, depletion interactions, and colligative nonideality, which characterize intracellular crowding. Second, our observations present a caveat on perceived roles of cholesterol in altering membrane poration characteristics by elevating edge tension35 and lowering membrane rupture tension. In binary homogeneous mixtures with phospholipids, cholesterol is known to increase the edge tension36 (because the invertedconical shape of the molecule disfavors the micellar edge of the pre-boundaries) but lowers the membrane rupture tension required to nucleate a pore (likely because of structural heterogeneity). In the present ternary (and by extension, in more complex lipid mixtures) this simple picture becomes inapplicable: by dynamically phase separating into the Lo phase, cholesterol appears to have little or no effect on edge tension (or pore closure). It is also likely that the domain boundaries produced through lateral phase separation provide defect sites reducing barriers for both pore nucleation and rupture tension enabling pulsatile osmotic equilibration under lower pressure gradients than required for compositionally homogeneous vesicles. Third, our findings suggest that any propensity for spontaneous overcrowding of macromolecular cargo in situ during the self-assembly of amphiphilic vesicles is opposed by osmotic forces. Recent independent observations45 in synthetic cell and origin-of-life contexts have led to a hypothesis that a small fraction of vesicular compartments encapsulate high concentrations of solutes, at concentrations far higher than the expected Poisson statistics, giving rise to functional protocells.
Thus, for any sustained spontaneous overcrowding to occur, independent interactions that out-compete osmotic forces must be present. Indeed, recent studies suggest that cytosolic crowding due to proteins of varied sizes, charges, and shapes in living cells, unlike synthetic crowding agents studied here, is not inert: rather enthalpically driven chemical interactions of associative kind acts to reduce the effects of the entropically dominated repulsive interactions.5,46 Moreover, any specific interactions between the macromolecules and the inner face of the membrane wall such as through electrostatic interactions and tendency for liquid−liquid phase separation7a,47 in the cytoplasmic face further counteracts the osmotic effects due to crowding in living cells.
■
MATERIALS AND METHODS
Materials. 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), egg sphingomyelin (egg SM), cholesterol, 1,2-dioleoyl-snglycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)2000] (ammonium salt) (DOPE-mPEG 2000), lissamine rhodamine B 1,2-dioleyl-sn-glycero-3-phosphoethanolamine (Rhodamine-B DOPE), and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine -N-(7nitro-2-1,3-benzoxadiazol-4-yl) (ammonium salt) (NBD-PE) were acquired from Avanti Polar Lipids (Alabaster, AL). Glucose, poly(ethylene glycol) (PEG 8000, MW 8000), and dextran from Leuconostoc mesenteroides (dextran 10000, MW 9000−11000) were obtained from Sigma-Aldrich (St. Louis, MO). Alexa Fluor 488Dextran (MW 10000) and 2-NBDG (2-(N-(7-nitrobenz-2-oxa-1,3diazol-4-yl)amino)-2-deoxyglucose) (NBD-glucose) were purchased from ThermoFisher Scientific (Eugene, OR). Sucrose was obtained from EMD Chemicals (Philadelphia, PA). Chloroform was purchased from Fisher Scientific (Fair Lawn, NJ). Ninety-six-well glass bottom plates were obtained from MatTek Corp. (Ashland, MA). Indium tin oxide (ITO) coated glass slides (resistance 4−30 Ω) were obtained from Delta Technologies (Loveland, CO). All chemicals were used without further purification. GUV Preparation. Giant unilamellar vesicles (GUV) were prepared following a well-established electroformation protocol from Angelova and Dimitrov.48 Two separate stock solutions, one consisting of 2:2:1 ratio of POPC, sphingomyelin, and cholesterol with 2.2% DOPE-mPEG 2000, 1% rhodamine-B-DOPE, and 2% NBD-PE and the second consisting of 96.8% POPC with 2.2% DOPE-mPEG 2000 and 1% rhodamine-B-DOPE were prepared in chloroform at 2 mg/ mL. Small droplets (15 μL) of stock solution were spread on the conductive side of each of two ITO-coated slides and allowed to dry under vacuum overnight. The dried film on one of the two slides was then directly hydrated either with 14 mM sucrose (control experiments) or with polymer solutions. The latter included (1) 14 mM PEG 8000, (2) 14 mM dextran 10000, (3) a mixture of 6.0 wt % PEG 8000 and 6.4 wt % dextran 10000, and (4) 114 mM sucrose. The solution droplet was contained using a 1 mm thick rubber ‘O’ ring (∼20 mm diameter) from Ace Hardware (Davis, CA) sealed with high vacuum grease from Dow Corning (Midland, MI). A water-tight chamber was then created by sealing the second ITO slide over the ring, ensuring that no visible air bubbles were trapped inside. The formation of this air-free chamber allows the films on the ITO slides to become hydrated by the incubating solution. Using a function generator, a 4 Vpp AC sine-wave was then applied across the two slides at 10 Hz for ∼3 h followed by a 4 Vpp AC square wave at 2 Hz for ∼2 h, both on a hot plate set at 45 °C. During the formation the ITO sandwhich was covered with an aluminum foil to protect from light. Following the formation, the ITO sandwich was disassembled and the solution containing GUVs was collected by a pipet. The GUVs were either used immediately or stored in small centrifuge tubes at 4 °C. All vesicles were used within a week of preparation. Spinning Disk Confocal Fluorescence Microscopy. Spinning disk confocal fluorescence microscopy measurements were performed using an Intelligent Imaging Innovations Marianas Digital Microscopy Workstation (3i Denver, CO) fitted with a CSU-X1 spinning disk head G
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society (Yokogawa Musashino-sh, Tokyo, Japan) and a Quantem512SC EMCCD camera (Photometrics Tuscon, AZ). Fluorescence micrographs were obtained using oil immersion objectives (Zeiss Fluor 40x (NA 1.3), Zeiss Plan-Fluor 63x (NA 1.4), and Zeiss Fluor 100X (NA 1.46); Carl Zeiss Oberkochen, Germany). Samples of osmotically balanced GUVs were prepared as described in the previous section. In a typical experiment, the glass bottom 96-well plate containing osmolyte-laden solution was mounted onto the microscope, and once oiled, the objectives were raised to form a meniscus between the cover glass and the objective. To impose osmotic gradients in real-time, GUV suspension is added to the solution in the sample chambers. Rhodamine-B DOPE (Ex/Em; 560/583) was exposed with a 50 mW 561 laser line. Alexa Fluor 488-Dextran 10,000 (Ex/Em; 495/519), NBD-PE (Ex/Em; 460/535) and 2-NBDG (Ex/Em; 465/540) were exposed with a 50 mW 488 laser line. The images are subsequently analyzed using ImageJ (http://rsbweb.nih.gov/ij/), a public-domain software, and Slidebook digital microscopy imaging software (3i Denver, CO). Fluorescence Microscopy. Wide-field epifluorescence measurements were carried out using a Nikon Eclipse TE2000S inverted fluorescence microscope (Technical Instruments, Burlingame, CA) equipped with a Roper Cool Snap camera (Technical Instruments), Hg lamp as the light source, and filter cubes to filter absorption and emission to the source and the CCD camera, respectively. Videos were taken using a Plan Fluor 20X (NA, 0.25) objective (Nikon, Japan). Osmotic Pressure Measurement. The osmotic pressures of PEG 8000 and dextran 10000 solutions were measured using a Wescor 51OOC vapor pressure deficit osmometer (Wescor, Inc., Logan, UT). The osmometer was calibrated using NaCl solution over a concentration range of 100−1000 mmol·kg−1 (Wescor, Inc.). Ten micrliters of each sample was dropped onto a 6 mm diameter filter paper, which was then placed inside the osmometer. Osmotic pressures, II (MPa), were estimated from the osmolality measurement using Van’t Hoff equation, II = RTc, where c = osmolality mol·kg−1 and RT = 2.446 kg·MPa·mo1−1 at 21 °C. Determination of the Second Virial Coefficient. Although there exist several studies for the determination of the second virial coefficient for PEG 8000,12,14 to our knowledge, no experimental values are available for dextran 10000. Therefore, we used osmotic pressure measurement to evaluate the second virial coefficients in dextran 10000 solution and PEG 8000 for consistency. The measured osmotic pressure as a function of solute concentration was fitted to the equation Π = RT[c + A2(Mc)2] with A2 being the fitting parameter (see Figure 1). We used M = 8000 g/ mol for PEG 8000 and M = 10 000 g/mol for dextran 10000. The fitted values for the virial coefficient were A2 = 8.18 × 10−9 mol m3/ g2 for PEG 8000 (R2 = 0.975) and A2 = 2.32 × 10−9 mol m3/ g2 for dextran 10000 (R2 = 0.994). The value for PEG 8000 is in good agreement with previously measured values,12,14 and the value of A2 for dextran 10000 is in the expected range based on reported measurements for dextran solutions of comparable molecular weights.31
■
■
S4.4. Supplementary Video S4.4. Swell-Burst Dynamics of GUVs Containing High Concentrations of Small Molecule (sucrose) Osmolytes. (AVI) S4.5. Supplementary Video S4.5. Transient Poration. (AVI) S4.6. Supplementary Video S4.6. Fluorescence Leakage Assay. (AVI) S4.7. Supplementary Video S4.7. Fluorescence Leakage Assay. (AVI) S4.8. Supplementary Video S4.8. Osmotically Propelled Transport of Cargo Vesicles. (AVI) S4.9. Supplementary Video S4.9. Effect of Solution Viscosity Contrast on Pore Life. (AVI)
AUTHOR INFORMATION
Corresponding Authors
*
[email protected] *
[email protected] ORCID
Padmini Rangamani: 0000-0001-5953-4347 Atul N. Parikh: 0000-0002-5927-4968 Author Contributions #
W.-C.S., D.L.G., and M.C. contributed equally.
Notes
The authors declare no competing financial interest.
■ ■
ACKNOWLEDGMENTS This work is supported by NSF PHY-1505017 to P.R. and A.N.P. REFERENCES
(1) Ellis, R. J. Trends Biochem. Sci. 2001, 26, 597. (2) Luby-Phelps, K. Mol. Biol. Cell 2013, 24, 2593. (3) Zimmerman, S. B.; Minton, A. P. Annu. Rev. Biophys. Biomol. Struct. 1993, 22, 27. (4) Minton, A. P. Biophys. J. 2005, 88, 971. (5) Sarkar, M.; Smith, A. E.; Pielak, G. J. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 19342. (6) Dix, J. A.; Verkman, A. S. Annu. Rev. Biophys. 2008, 37, 247. (7) (a) Hyman, A. A.; Simons, K. Science 2012, 337, 1047. (b) Keating, C. D. Acc. Chem. Res. 2012, 45, 2114. (8) Parry, B. R.; Surovtsev, I. V.; Cabeen, M. T.; O’Hern, C. S.; Dufresne, E. R.; Jacobs-Wagner, C. Cell 2014, 156, 183. (9) van den Berg, J.; Boersma, A. J.; Poolman, B. Nat. Rev. Microbiol. 2017, 15, 309. (10) Wood, J. M. Microbiol. Mol. Biol. Rev. 1999, 63, 230. (11) Daoud, M.; Cotton, J. P.; Farnoux, B.; Jannink, G.; Sarma, G.; Benoit, H.; Duplessix, R.; Picot, C.; de Gennes, P. G. Macromolecules 1975, 8, 804. (12) Wang, S.-C.; Wang, C.-K.; Chang, F.-M.; Tsao, H.-K. Macromolecules 2002, 35, 9551. (13) Kushare, S. K.; Shaikh, V. R.; Terdale, S. S.; Dagade, D. H.; Kolhapurkar, R. R.; Patil, K. J. J. Mol. Liq. 2013, 187, 129. (14) Money, N. P. Plant Physiol. 1989, 91, 766. (15) Parsegian, V. A.; Rand, R. P.; Rau, D. C. Proc. Natl. Acad. Sci. U. S. A. 2000, 97, 3987. (16) Asakura, S.; Oosawa, F. J. Chem. Phys. 1954, 22, 1255. (17) Arnold, K.; Zschoernig, O.; Barthel, D.; Herold, W. Biochim. Biophys. Acta, Biomembr. 1990, 1022, 303. (18) Yamazaki, M.; Ohnishi, S.; Ito, T. Biochemistry 1989, 28, 3710. (19) (a) Evans, E.; Needham, D. Macromolecules 1988, 21, 1822. (b) Kuhl, T.; Guo, Y. Q.; Alderfer, J. L.; Berman, A. D.; Leckband, D.; Israelachvili, J.; Hui, S. W. Langmuir 1996, 12, 3003. (20) Lentz, B. R. Chem. Phys. Lipids 1994, 73, 91.
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.7b10192. Supporting figures and details of the theoretical model (PDF) S4.1. Supplementary Video S4.1. Swell-Burst Dynamics of PEG-laden GUVs. (AVI) S4.2. Supplementary Video S4.2. Swell-Burst Dynamics of Dextran-laden GUVs. (AVI) S4.3. Supplementary Video S4.3. Swell-Burst Dynamics of GUVs Containing Mixture of PEG and Dextran Crowding Agents. (AVI) H
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Article
Journal of the American Chemical Society (21) Terasawa, H.; Nishimura, K.; Suzuki, H.; Matsuura, T.; Yomo, T. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 5942. (22) Chen, I. A.; Roberts, R. W.; Szostak, J. W. Science 2004, 305, 1474. (23) Macdonald, R. I. Biochemistry 1985, 24, 4058. (24) Lehtonen, J. Y. A.; Kinnunen, P. K. J. Biophys. J. 1995, 68, 525. (25) (a) Koslov, M. M.; Markin, V. S. J. Theor. Biol. 1984, 109, 17. (b) Oglecka, K.; Rangamani, P.; Liedberg, B.; Kraut, R. S.; Parikh, A. N. eLife 2014, 3, 18. (c) Chabanon, M.; Ho, J. C. S.; Liedberg, B.; Parikh, A. N.; Rangamani, P. Biophys. J. 2017, 112, 1682. (26) Zade-Oppen, A. M. M. J. Microsc. 1998, 192, 54. (27) Morales-Penningston, N. F.; Wu, J.; Farkas, E. R.; Goh, S. L.; Konyakhina, T. M.; Zheng, J. Y.; Webb, W. W.; Feigenson, G. W. Biochim. Biophys. Acta, Biomembr. 2010, 1798, 1324. (28) Dominak, L. M.; Keating, C. D. Langmuir 2008, 24, 13565. (29) Veatch, S. L.; Keller, S. L. Phys. Rev. Lett. 2005, 94, 148101. (30) Baumgart, T.; Hunt, G.; Farkas, E. R.; Webb, W. W.; Feigenson, G. W. Biochim. Biophys. Acta, Biomembr. 2007, 1768, 2182. (31) Ioan, C. E.; Aberle, T.; Burchard, W. Macromolecules 2000, 33, 5730. (32) Israelachvili, J. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 8378. (33) Sandre, O.; Moreaux, L.; Brochard-Wyart, F. P. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 10591. (34) Haynes, C. A.; Beynon, R. A.; King, R. S.; Blanch, H. W.; Prausnitz, J. M. J. Phys. Chem. 1989, 93, 5612. (35) Karatekin, E.; Sandre, O.; Guitouni, H.; Borghi, N.; Puech, P. H.; Brochard-Wyart, F. Biophys. J. 2003, 84, 1734. (36) Portet, T.; Dimova, R. Biophys. J. 2010, 99, 3264. (37) Peterlin, P.; Arrigler, V. Colloids Surf., B 2008, 64, 77. (38) Hamada, T.; Kishimoto, Y.; Nagasaki, T.; Takagi, M. Soft Matter 2011, 7, 9061. (39) Gonzalez-Tello, P.; Camacho, F.; Blazquez, G. J. Chem. Eng. Data 1994, 39, 611. (40) Brochard-Wyart, F.; de Gennes, P. G.; Sandre, O. Phys. A 2000, 278, 32. (41) Happel, J.; Brenner, H. Low Reynolds Number Hydrodynamics: With Special Applications to Particulate Media; Springer: Dordrecht, 1983. (42) (a) Levin, Y.; Idiart, M. A. Phys. A 2004, 331, 571. (b) BrochardWyart, F.; de Gennes, P. G.; Sandre, O. Phys. A 2000, 278, 32. (43) Kozer, N.; Kuttner, Y. Y.; Haran, G.; Schreiber, G. Biophys. J. 2007, 92, 2139. (44) Armstrong, J. K.; Wenby, R. B.; Meiselman, H. J.; Fisher, T. C. Biophys. J. 2004, 87, 4259. (45) (a) Pereira de Souza, T.; Steiniger, F.; Stano, P.; Fahr, A.; Luisi, P. L. ChemBioChem 2011, 12, 2325. (b) Luisi, P. L.; Allegretti, M.; Pereira de Souza, T.; Steiniger, F.; Fahr, A.; Stano, P. ChemBioChem 2010, 11, 1989. (46) (a) Gnutt, D.; Gao, M.; Brylski, O.; Heyden, M.; Ebbinghaus, S. Angew. Chem., Int. Ed. 2015, 54, 2548. (b) Groen, J.; Foschepoth, D.; te Brinke, E.; Boersma, A. J.; Imamura, H.; Rivas, G.; Heus, H. A.; Huck, W. T. S. J. Am. Chem. Soc. 2015, 137, 13041. (47) (a) Hyman, A. A.; Weber, C. A.; Juelicher, F. Annu. Rev. Cell Dev. Biol. 2014, 30, 39. (b) Brangwynne, C. P.; Tompa, P.; Pappu, R. V. Nat. Phys. 2015, 11, 899. (48) Angelova, M. I.; Dimitrov, D. S. Faraday Discuss. Chem. Soc. 1986, 81, 303.
I
DOI: 10.1021/jacs.7b10192 J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
