Protein coacervation-driven active forces power protocell dynamics

Domain: chemistry
Published: 2026-04-07
Editorial outcome: major_revision

Abstract
Protein coacervates formed by liquid-liquid phase separation are emerging as active force generators, independent of ATP-driven motors. Nevertheless, the coordination and force scaling of protein coacervates remain largely unexplored. Here, we engineer a temperature-responsive elastin-based protocell model displaying temperature-modulated contractility and attendant force harnessing. By leveraging the phase separation properties, we modulate the protocell dynamics associated with volume contraction and membrane budding. Crosslinking of the elastin-based membrane influences the contraction dynamics such that the accumulation of mechanical forces in the protocells results in the spontaneous expulsion of internally trapped protein liquid-liquid phase separation (LLPS) complexes. We use a simple mathematically model to show how protein coacervation can amplify small piconewton-scale forces to perform large-scale mechanical work, highlighting the mechanical potential of protein coacervation dynamics. Taken together, our results provide a model framework for harnessing protein coacervates-driven forces and offer a step to future applications in synthetic biology, biomaterials and next-generation soft robotics.

Introduction
Cells are extraordinary engines of force, driving the fundamental processes that sustain life, including muscle contraction1, cell migration2, cytokinesis3, exocytosis4 and tissue morphogenesis5. Central to this remarkable capability are the active forces generated by a range of mechanisms: cytoskeletal dynamics, cell-matrix adhesion, osmotic pressure, membrane tension, and protein phase separation1,6-9. These forces, often operating in the piconewton range, scale up and orchestrate the cellular activities. Among these mechanisms, protein liquid-liquid phase separation (LLPS), including protein condensation and coacervation, have recently emerged as a surprising source of substantial interfacial forces within living cells, functioning independently of ATP-fueled molecular motors9. Such LLPS-based forces, arising from processes like coalescence10,11, Ostwald ripening12, and interface wetting13, underscore the liquid-like nature of protein condensates, enabling them to flow, fuse, and dynamically assemble or disassemble in response to environmental cues. This capacity allows cells to harness these forces for the dynamic compartmentalization of intracellular structures, influencing critical processes such as cell signaling, endocytosis, and autophagy. Unraveling the molecular origins and chemical principles underlying large-scale protein LLPS dynamics should provide insights into how non-motor protein harness forces within living cells, and potentially revolutionize our understanding of the structure, organization, and function of living cells. Furthermore, employing and scaling-up such forces could be important for the biomimetic engineering of soft robots14,15 and synthetic cells16,17, and offer new frameworks for the utilization of protein LLPS-based forces in fields such as biomaterials and drug delivery 18,19.

The ability of protein coacervates to rapidly transition in response to local cytoplasmic conditions facilitates dynamical adaptation to changes in environmental conditions and regulation of various cellular processes. However, the chemical and biological complexities of how these protein coacervation-induced forces are coordinated and scaled to perform complex mechanical tasks remain largely elusive and challenging to unravel. As a step towards addressing these challenges, we present a simple protocell model based on the stabilization of water-in-oil microemulsion droplets with a self-coacervating protein (elastin)20,21 membrane capable of exhibiting temperature-modulated contractability and attendant force harnessing within controllable confined spaces. The protocell membrane consists of a closely packed array of amphiphilic elastin-polymer nanoconjugates that spontaneously assemble at the water/oil droplet interface. If required, the elastin-based membrane can be further stabilized by covalent crosslinking to modify the contractile behavior. By leveraging the phase separation properties of the membrane building blocks, we engineer the protocell dynamics associated with contraction, membrane budding and shape remodeling, and model the collective accumulation of force during large-scale phase fusion. Finally, we show how continuous protocell contraction results in the transient production of multiphase droplets and their subsequent force-induced expulsion into the surrounding environment.

Results
Temperature-driven contraction dynamics of elastin-based protocells
Model protocells capable of temperature-dependent contractability were constructed using elastin-stabilized water-in-oil microemulsion droplets. Covalent coupling of amino-functionalized elastin to mercaptothiazoline-terminated poly(N-isopropylacrylamide) (PNIPAAm20, Mn 8,800 g/mol) was used to prepare surface-active amphiphilic protein-polymer nanoconjugates. In this system, PNIPAAm primarily contributes amphiphilicity that promotes interfacial enrichment and stabilizes a water-oil interface (Fig. 1a). Although both elastin and PNIPAAm exhibit lower critical solution temperature (LCST)-type thermal transitions21,22, their functional contributions are distinct. Elastin alone forms simple coacervates (Supplementary Fig. 1a), whereas elastin–PNIPAAm conjugates display enhanced temperature responsiveness (Supplementary Fig. 1), producing smaller particle-like domains that assemble into an interconnected interfacial network. In contrast, BSA–PNIPAAm conjugates form only network-like aggregates without evident LLPS (Supplementary Fig. 1b–c), indicating that PNIPAAm alone is insufficient to drive phase separation. These observations support that elastin LLPS dominates the thermodynamic condensation process, while PNIPAAm modulates interfacial organization and temperature sensitivity.

Addition of the nanoconjugates to a dextran-containing aqueous/2-ethyl-1-hexanol mixture at a range of temperatures produced microemulsion suspensions comprising elastin-stabilized polysaccharide-containing water-in-oil droplets with a mean diameter of 14.32 ± 4.76 μm (Fig. 1a and Supplementary Fig. 2a-b). The emulsion droplets were stable when stored at 4°C but exhibited a temperature-dependent decrease in mean diameter above 20°C (Fig. 1b, Supplementary Fig. 2c). This diameter reduction was consistently observed by both confocal and wide-field imaging (Supplementary Fig. 2d), and is consistent with the thermos-responsive behavior of elastin and its nanoconjugates (Supplementary a Fig. 1). For example, the mean volume of the droplets decreased by 25% and 74% when incubated at 25°C or 33°C for 20 minutes. In contrast, water-in-oil droplets stabilized by a surface-active shell of bovine serum albumin (BSA)-PNIPAAm nanoconjugates showed only a weak response below 35°C23,24. This indicated that the droplet contractability was driven by liquid-liquid phase separation (LLPS) of the elastin-based conjugates due to the onset of enhanced hydrophobic interactions between the elastin molecules as the temperature was raised above 20°C. In general, the rate of volume contraction increased as the temperature was increased to 37°C.

Fluorescence microscopy was used to track the time-dependent localization of elastin and dextran during progressive contraction of the elastin-polymer-coated water droplets (Fig. 1c-f, Supplementary Movie 1). Over time, elastin, which was initially distributed both in the droplet interior and membrane, became enriched at the water/oil interface as the protocell diameter decreased (Fig. 1g). During this coacervation process, the inner elastin–Atto647 fluorescence decreased between 5 and 20 min due to the migration and accumulation of elastin molecules at the interface, resulting in reduced internal signal and a corresponding increase in interfacial fluorescence (Supplementary Movie 1). In contrast, dextran remained homogeneously distributed in the droplet interior where it became increasingly concentrated as the volume of the droplet decreased (Fig. 1h). Contraction of the protocell resulted in an increase in the concentration of the entrapped soluble elastin-polymer nanoconjugates such that LLPS occurred within the interior of the shrinking droplets to produce an encapsulated dispersion of elastin coacervates with diameters around 2.03 ± 0.31μm (n=10) (Fig. 1i-j). The size of the phase-separated coacervates within the confined environment was consistent with previous reports21. The entrapped coacervates became localized under the elastin-based membrane as the droplets continued to contract (Supplementary Movie 1). Similar results were observed with elastin-based protocells containing aqueous solutions of BSA (Supplementary Fig. 3, Supplementary Movie 2). By around 25 min, the interfacial elastin layer had dissipated, and elastin appeared as dispersed small coacervates in the surrounding phase rather than remaining concentrated at the interface (Supplementary Movie 1).

Quantitative analysis of temperature-dependent changes in protocell diameter and the fluorescence intensity of entrapped FITC-dextran was performed during gradual heating from room temperature to 37 °C. The contracting droplets became transiently enlarged and irregular in surface morphology after ca. 30 min due to the onset of solvent molecule exchange across the elastin-polymer interface (Fig. 2a-c and Supplementary Movie 3). Fluorescence imaging of the protocells undergoing transient enlargement indicated that the increase in volume was associated with the irreversible uptake of oil droplets into the protocells along with a non-compensating efflux of water and elastin-polymer nanoconjugates into the continuous oil phase (Fig. 2d-g, Supplementary Fig. 4a). The period of volume enlargement was ca. 5 min, after which the emulsion droplets underwent a second contraction stage during which the transferred oil droplets coalesced inside the protocell. Because the elastin-rich coacervate layer retained a small amount of bound water, elastin-coated oil-in-oil droplets were transiently formed and subsequently collapse during contraction (Fig. 2h-i, Supplementary Movie 4), consistent with the weak elastic character of uncrosslinked elastin coacervates25.

To gain insight into the dynamics and force accumulation in the elastin-based protocell system, we developed a biophysical model incorporating principles of thermodynamics26, pressure dynamics27, and liquid-liquid phase separation28. The model postulated that elastin phase separation at the droplet interface induces a pressure differential, which drives contraction forces within the protocell (Supplementary Fig. 5a). Model parameters were obtained by global fitting to experimentally measured diameter–time shrinkage profiles across multiple temperatures (Supplementary Fig. 5b–c). Using these parameters, the model predicted a pressure differential across the elastin-based interface (Supplementary Fig. 5d), corresponding to an integrated contractile force on the order of 10²–10³ pN at 37°C for protocells with a diameter of 25 µm. This magnitude of force is sufficient to initiate the inward contraction of the protocell (Supplementary Fig. 5e). As LLPS typically generates forces on an order of 10-1 pN9, the model suggests that these small forces are amplified by the microcompartmentalized structure by several hundred-fold. Moreover, the model simulated an increase in capillarity pressure as a function of time and decrease in droplet radius, eventually reaching a maximum pressure of ca. 0.1 Pa at 37°C (Supplementary Fig. 5f). Overall, the model captures the dynamics of elastin-driven phase separation and allows an estimation of the collective forces that influence the mechanical properties of the protocell membrane during contraction.

Crosslinked-enhanced membrane dynamics in elastin-based protocells
We employed three chemical crosslinkers with different chain lengths – glutaraldehyde, DTSSP and PEG-bis(N-succinimidyl succinate (Mn 2,000 g/mol) – to elucidate the influence of membrane dynamics on the temperature-dependent contractibility of elastin-stabilized water-in-oil emulsion droplets (Fig. 3a). In each case, time-dependent fluorescence microscopy was used to track the structural changes accompanying droplet contraction at 37°C. Crosslinking the elastin-PNIPAAm nanoconjugates at the water droplet/oil interface with glutaraldehyde gave rise to significant morphological changes during contraction, transitioning the protocells from a relatively smooth and spherical form to a more compacted and irregular shape (Fig. 3b, Supplementary Movie 5). Time-resolved imaging confirms that, upon glutaraldehyde crosslinking, contour irregularities nucleate locally early during contraction and then subsequently amplify (Supplementary Fig. 6a). Such behavior likely arises from the formation of short, rigid covalent crosslinks, which restrict the elastin membrane’s ability to uniformly redistribute strain and accommodate rapid surface-area changes during contraction. In contrast, the protocells remained spherical under contraction when crosslinked with the longer molecular connectors of DTSSP or PEG (Fig. 3c-d), indicating that the membrane dynamics were adaptive to the changes in volume. Moreover, compared with non-crosslinked droplets, DTSSP-crosslinked elastin membranes became progressively thicker and more condensed as the droplets decreased in size, and exhibited approximately 2.7-fold slower contraction rates to reach 50% of their original volume demonstrated in Fig. 1b (Fig. 3e and Supplementary Movie 6).

Notably, no process of transient volume enlargement was observed for the glutaraldehyde- or DTSSP-crosslinked protocells, indicating that unlike the non-crosslinked droplets there was minimal transfer of oil, water and elastin across the water droplet-oil interface during contraction. In contrast, transient enlargement was observed when the protocells were crosslinked with PEG (Fig. 3d, Supplementary Movie 7), indicating that the crosslinked membrane remained sufficiently flexible to support the uptake of oil molecules. Consequently, localized microphase separation of elastin in the PEG-crosslinked membrane was still apparent during contraction and gave rise to the formation of coacervated domains at the water droplet/oil interface (Fig 3f, Supplementary Movie 8). Notably, although PEG-crosslinked elastin membranes retain some flexibility, they exhibit limited fluidity (Supplementary Fig. 6b–c), in contrast to lipid membranes.

Increasing the degree of PEG crosslinking resulted in a progressive increase in the size and fluorescence density of the microphase-separated regions as the droplets contracted such that distinct irregular patches of condensed elastin-PNIPAAm nanoconjugates became embedded in the protocell membrane (Fig. 3g, Supplementary Fig. 6d. Supplementary Movie 9). Surprisingly, the elastin coacervate particles were radially ejected from the protocell surface along linear trajectories under high levels of contraction (Fig. 3h-i, Supplementary Movie 10). The rapid ejection of particles without filament or bead-on-a-string morphologies indicates that this process is probably driven by the sudden release of elastic stress stored in the contracting elastin-rich membrane29,30. The propelled particles traveled linearly for 71.74 ± 16.69 µm within 110 seconds with a median speed of 0.82 ± 0.16 µm/s (Fig. 3j-k), which was approximately 2.65 times higher than the speed of free-diffusing particles (0.31 ± 0.13µm/s) under similar environmental conditions. Analyses of single ejected particles revealed a rapid initial acceleration, with initial peak velocities of ca. 0.93 µm/s, followed by gradual deceleration (Fig. 3l-m). Using Stokes’ law together with the measured median velocity, the viscous drag force was calculated to be F drag ≈0.31±0.06 pN. Under low–Reynolds-number conditions, this drag balances the active launching force, such that F launch ≃ F drag.

Taken together, the above observations indicate that crosslinking of the elastin-based membrane not only influences the dynamics associated with protocell contraction but also generates mechanical forces capable of launching and rapidly distributing the internal contents of the protocell into the environment, providing insights into the nature of force generation during phase separation at water droplet/oil interfaces.

Force-mediated release of nested coacervates
Given the above observations, we designed a thermally mediated self-driving protocell system for the contraction-induced release of encapsulated LLPS complexes (Fig. 4a). The latter were endogenously produced by encapsulating encapsulated dextran and PEG3300 within the elastin-based protocells and inducing protocell contraction at 37 °C. As the dextran and PEG levels were initially below the threshold for segregative LLPS, thermally induced elastin-mediated contraction initiated coacervation within the protocells by volume-dependent increases in concentration. Consequently, subsequent demixing of dextran and PEG resulted in the endogenous formation of elastin/dextran coacervate microdroplets dispersed within a PEG-rich continuous phase within the protocells (Fig.4b-c, Supplementary Movie 11).

Undertaking the above procedure in non-crosslinked elastin-stabilized water-in-oil droplets gave rise to partial fusion of the trapped elastin/dextran coacervate microdroplets as the contraction increased but without expulsion from the protocells due to the inherent flexibility in the absence of membrane crosslinks (Fig.4d, Supplementary Fig. 7a). In contrast, replacing PEG3300 with a PEG crosslinker gave rise to contractive protocells capable of a two-stage process involving the induction and ejection of the phase-separated elastin/dextran droplets at 37 °C (Fig. 4e). In the first stage, the protocells underwent enlargement-contraction and corresponding increases in internal concentration to initiate dextran/PEG demixing and elastin/dextran phase separation (Supplementary Movie 12). This was followed by an influx of oil droplets, movement and partial coalescence of the LLPS particles at the membrane to generate a densely packed shell of discrete elastin/dextran satellite particles that were spontaneously ejected (Fig. 4f-h, Supplementary Movie 13). During the second stage, the satellite particles on the interface fused with each other, gradually formed larger coacervates with diameters increasing from 0.73 ± 0.06 µm to 4.96 ± 0.71 µm over 16 min (Fig. 4f, Supplementary Fig. 7b). Notably, protocell contraction drove particle release during the initial 50 s, with a measured ejection frequency of 0.91 ± 0.14 s⁻¹ on a single confocal plane (Fig. 4g, Supplementary Fig. 7c-e).

Methods
Amine-modified elastin (elastin-NH2) was synthesized according to a previously described method24. Briefly, 1,6-hexanediamine (HMDA, sigma, H11696) was covalently conjugated to aspartic and glutamic acid residues on the surface of elastin via carbodiimide activation. Typically, an aqueous solution of HDMA (500 mg, 4.3 mmol, 5 mL) was adjusted to pH 6.5 using 5 M HCl and added dropwise to a 5 mL stirred solution of elastin from bovine neck ligament (100 mg, 20 mg/ml, sigma, E6527). The pH was readjusted if necessary to 6.5 with 0.1 M HCl. The coupling reaction was initiated by adding 50 mg N-(3-dimethylaminopropyl)-N’-ethylcarbodiimide hydrochloride (EDC, sigma, E7750) to the stirred solution. The pH value was maintained at 6.5 by adding aliquots of 0.1 M HCl, and another 50 mg of EDC were added after 4 h. The solution was then stirred overnight. The elastin-NH2 was then dialyzed (Medicell, dialyzing tubing, MWCO 12-14 kDa) for 2 days against Milli-Q water, then filtered (Sartorius Ministart NML syringe filters, 45 μm) to remove any precipitate and the supernatant was freeze-dried.

Fluorescence labelling of elastin. 1) FITC labelling: 10 μL of 2 mg/mL Fluorescein isothiocyanate isomer I (FITC, sigma, F4274) solution in DMSO was added dropwise to elastin-NH2 (2 mg/mL in 0.1 M carbonate buffer with pH 9.0), with continuous mixing to avoid high local DMSO concentrations and thereby prevent protein denaturation. The mixture was stirred at room temperature for 5 hours, then dialyzed (Medicell, dialyzing tubing, MWCO 12-14 kDa) for 2 days against Milli-Q water and dialyzed for 1 days against PBS buffer (pH 8.0). Then protein solution was centrifuged to remove any precipitate and the supernatant was concentrated to 2 mg/ml. Based on UV-vis spectroscopy measurements, there were on average 1-2 dye molecules per protein.
2) Alexa Fluor 647 labelling: 50 μL Alexa Fluor 647 Succinimidyl Esters (NHS esters, thermos fisher, A20006) solution in DMSO (2 mg/mL) were added dropwise to 1 mL elastin-NH2 solution (2 mg/mL in 0.1M carbonate buffer with pH 9.0). The mixture was stirred at room temperature for 1 h, then dialyzed (Medicell, dialyzing tubing, MWCO 12-14 kDa) for 2 days against Milli-Q water and dialyzed for 1 days against PBS buffer (pH 8.0) at 4 °C. Then protein solution was centrifuged to remove any precipitate and the supernatant was concentrated to 2 mg/mL.

PNIPAAm-conjugated elastin building block. PNIPAAm polymer (Mn = 8,800 g/mol) was synthesized according to published approach24. Briefly, 4,4′-azobis(4-cyanovaleric acid) (ACVA) (4.01 g, sigma, 1159) and 2-mercaptothiazoline (4.50 g, Sigma, M6204) were dissolved in 1,4-dioxane (200 mL, sigma, 360481) and degassed with nitrogen for 30 min. N,N′-dicyclohexylcarbodiimide (6.82 g, sigma, D80002) and 4-(dimethylamino)pyridine (0.10 g, sigma, 107700) in 1,4-dioxane (100 mL, sigma, 36048) were added, and the mixture was stirred at room temperature for 20 h. After filtration and concentration, the product was precipitated in cold diethyl ether to yield the mercaptothiazoline activated 4,4’-azobis(4-cyanovaleric acid) (ACVA-ACPM). Then 3g ACVA-ACPM was then reacted with bis(propylsulfanylthiocarbonyl) disulfide (1.37 g, sigma, 723126) in ethyl acetate (50 mL, sigma, 3 19902) under reflux for 18 h to generate the mercaptothiazoline-activated RAFT chain-transfer agent, which was purified by silica gel chromatography (ethyl acetate/hexane, 1:1 v/v). For polymerization, the RAFT agent (18.9 mg), 2,2′-azobisisobutyronitrile (1.7 mg, sigma, 441090), and N-isopropylacrylamide (7.5 mmol, sigma, 731129) were dissolved in acetonitrile (8 mL, sigma, 34851) in a sealed round-bottom flask. The solution was degassed by four freeze–pump–thaw cycles and polymerized at 60 °C for 8 h. The polymer was isolated by three precipitation cycles in diethyl ether/hexane (2:1 v/v) and dried under vacuum. Elastin-PNIPAAm conjugates was synthesized based on reaction of the PNIPAAm and amino group of proteins. Briefly, a 500 μL aqueous solution of the polymer (2 mg/mL) was added dropwise to 500 μL fluorescence-labelled elastin-NH2 (2 mg/mL in PBS, pH8.0). The mixed solution was stirred for 12 h at 4 °C, then purified by using a centrifugal tube (Millipore, Amicon Ultra, MWCO 100 kDa) to remove any unreacted PNIPAAm and salts. The solution was further concentrated to 5 mg/mL. The conjugates can be shortly stored at 4 °C.

Elastin conjugation level quantification
Primary amine content was quantified using a TNBSA (2,4,6-Trinitrobenzene Sulfonic Acid) assay31. Briefly, samples (elastin, elastin-NH2, elastin-NH2-FITC and elastin-NH2-FITC-PNIPAAm) were reacted with TNBSA in bicarbonate buffer (alkaline pH) at 37 °C for a fixed incubation time. The reaction product was quantified by absorbance at 335 nm and compared to a glycine calibration curve. Primary amine content was reported as µmol NH2 per mg protein. Unmodified elastin exhibited a primary amine content of 0.293± 0.001 µmol NH2 per mg (n=3). After amination (elastin–NH2), the primary amine content was 1.007 ± 0.005 µmol/mg (n=3). Following FITC labeling (elastin–NH2–FITC), the accessible primary amine content decreased to 0.666 ± 0.001 µmol/mg (n=3). After PNIPAAm conjugation (elastin–NH2–FITC–PNIPAAm), the primary amine content further decreased to 0.528 ± 0.002 µmol/mg (n=3). Because PNIPAAm conjugation proceeds via reaction with elastin primary amines, the decrease in TNBSA-accessible amines during this final step provides a quantitative proxy for PNIPAAm conjugation extent. Unless otherwise specified, elastin–NH2–FITC–PNIPAAm samples used throughout this study correspond to a PNIPAAm conjugation level of approximately 20.7 %, calculated from the fractional decrease in primary amines relative to elastin–NH2–FITC immediately prior to PNIPAAm coupling.

Thermo-response of elastin protein and elastin-PNIPAAm conjugates.
Elastin-NH2-FITC and elastin-NH2-FITC-PNIPAAm were each prepared at 0.8 mg/mL and stored on ice. Phase separation was triggered by adding 15 µL of 5 M NaCl to 50 µL of protein solution, yielding a final concentration of 0.6 mg/mL protein and 1.15 M NaCl. Samples were incubated for 15 min at 4, 15, 20, 25, 30, 33, or 37 °C using a metal bath. Phase separation was subsequently monitored by confocal microscopy (Leica SP8).

Preparation of elastin protocells. To test the dynamics of elastin protocells, the interfacial elastin proteins were either non-crosslinked or crosslinked.
1) Non-crosslinked: The inner aqueous solution of protocell was mixed with 5 μL of a 4.0 mg/mL elastin-PNIPAAm conjugate solution (no labelling, labelled with FITC or Alex Fluor 647) with 1μL 0.1 M NaHCO3 buffer. To encapsulate other contents, 1μL NaHCO3 buffer can be replaced with equal volume of Dextran-FITC or BSA-FITC. Unless otherwise noted the concentration of BSA-FITC and Dextran-FITC (70 kDa, sigma, 90718) was 2 mg/mL. Elastin-PNIPAAm stabilized w/o droplets were prepared by 6 μL elastin-PNIPAAm aqueous solution with 0.1 mL of 2-ethyl-1-hexanol (sigma, 538051) (aqueous/oil volume fraction, 0.06). The mixture was gently hand-shaken for 30 s to produce a protein-polymer stabilized water-in-oil emulsion. The water-in-oil emulsion should then be stored on ice or 4 °C before the microscopy imaging.
2) Crosslinked: Unless otherwise noted elastin protocells were crosslinked with PEG crosslinker (O,O’-Bis[2-(N-Succinimidyl-succinylamino) ethyl]polyethylene glycol), average Mn 2000, sigma, 713783). Briefly, 1μL 0.2 mg/μL PEG cross-linker (in 50 mM carbonate buffer, pH 8.5) was mixed with 5 μL of a 4.0 mg/mL elastin-PNIPAAm conjugate solution (no labelling, labelled with FITC or Alex Fluor 647), immediately followed by the addition of 0.1 mL of 2-ethyl-1-hexanol (aqueous/oil volume fraction, 0.06). The mixture was gently hand-shaken for 30 s to produce a protein-polymer stabilized water-in-oil emulsion. The water-in-oil emulsion should then be crosslinked on ice or 4 °C for at least 5h before the microscopy imaging. Contents such as Dextran-FITC (70 kDa) can be encapsulated by following the above non-crosslinked protocol.
Different types of crosslinkers were used as well, such as glutaraldehyde (sigma, 340855) and 3,3´-dithiobis[sulfosuccinimidylpropionate](DTSSP, sigma, 803200). For glutaraldehyde crosslinker, a 23 % solution was prepared first. In the step of w/o droplet preparation, 1 μL 23 % glutaraldehyde was used. The reaction between glutaraldehyde with protein was fast. Thus, the crosslinking was performed at 37 °C for 2-3 min and stored on ice or 4 °C before the microscopy imaging. Regarding the DTSSP crosslinker, a stock solution of DTSSP with 50 mM concentration was prepared in DMSO. In the step of w/o droplet preparation, 1 μL 50 mM DSP was added. The water-in-oil emulsion should then be crosslinked on ice or 4 °C for at least 2 h before the microscopy imaging.

Thermal dynamics of elastin protocells. All samples were imaged in silicone isolator chambers (9 mm diameter, 1 mm depth; Thermo Fisher Scientific) adhered to glass coverslips (24 × 24 mm, thickness #1.5; Thermo Fisher Scientific). The coverslips were pretreated one day prior with trimethoxy-[3-(2-methoxyethoxy)propyl]silane (sigma, 551635) (0.05% v/v in toluene) to enhance surface compatibility. Dynamic measurements of elastin protocells were conducted using a confocal microscope equipped with a heating chamber. Unless otherwise specified, the chamber was pre-set to a target temperature of 37 °C before imaging. Due to the large, open design of the chamber, stabilization of the temperature required additional time. During this period, a calibration coverslip of the same thickness as the sample was used to pre-focus the microscope at the correct z-position, minimizing delays in finding the focus during actual measurements. Once the target temperature was achieved, the sample chamber was mounted on the microscope, and imaging commenced immediately. Temperature-dependent dynamics shown in Supplementary Fig. 2 were measured sequentially in the heating chamber. Measurements began at 20 °C (room temperature) and progressed incrementally to 25 °C, 33 °C, 35 °C, and 37 °C. A fresh sample was used for each temperature to ensure reproducibility and avoid thermal history effects.

Microscopy and imaging analysis. All images were acquired using a Leica SP8 confocal laser scanning microscope equipped with a 63× HC PL APO CS2 objective. Fluorescence excitation was performed with a 65 mW Ar laser for FITC and a 10 mW Red He/Ne (633 nm) laser for Alexa Fluor 647. Images were captured with a pinhole size of 1 Airy unit for both channels and a resolution of 1024 × 1024 pixels. Representative micrographs and intensity profiles were generated from at least four independently repeated experiments, as detailed in the respective figure captions. Image processing and analysis were conducted using Fiji software (version v1.54f). Adjustments to brightness and contrast were applied homogeneously across all displayed images to ensure consistent presentation. For line intensity plots, Gaussian smoothing with a pixel width of 2 was applied in Fiji to reduce noise. Particle tracking was performed using the Fiji plugin TrackMate (version 3.4.2)32 following the official manual. Parameters such as traveling speed, max distance traveled, and radial displacement were analyzed within the TrackMate framework. Under low-Reynolds-number conditions, particle motion is governed by a balance between the active launching force and viscous drag. To quantify the viscous drag acting on each particle, we used the measured steady-state velocities and Stokes’ law, F drag =6πηr p v, where 𝑟 is particles radius, 𝑣 is velocity, 𝜂 is viscosity of medium. Because inertial contributions are negligible in this regime, the viscous drag force directly balances the capillary- or elasticity-derived launching force. Thus, the active force driving particle motion at steady state is approximated as: F launch ≃ F drag =6πηr p v. This estimation assumes a particle diameter of 5 μm and 2-ethyl-1-hexanol viscosity 𝜂 of 8 mPa·s at 37°C.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Reviewer #1 (Remarks to the Author):\nSummary: This work uses modified proteins (elastin-Nh2) forming water-in-oil microemulsion\ndroplets and shows the temperature effects on the droplet, with modelling to understand the\nunderlying principles. The experiments are amazing! And visuals stunning! But I have some\nconcerns about this work, especially about the model which has some overextended claims and\npotentially erroneous calculation of zetaNewtons– my estimate shows their numbers could be\n7 orders of magnitude off (however, i will admit, i did a quick and dirty back of the envelop\ncalculation - so i could also be wrong - but clarification is definitely needed!). However, if i’m\nright, then the core claim about force `amplification’ are weakly validated. Nonetheless, the\nexocytosis-like expulsion is super cool!\nReviewer #2 (Remarks to the Author):\nIn the present paper Jia et al. present a new approach introducing a contractile, elastin-based\nprotocell system in which liquid–liquid phase separation is key in the contraction observed.\nThis work highlights that condensation is able to develop forces at the piconewton scale that\ncan be amplified and eventually lead to observable expelling of components from inside the\nprotocell. Several cascade processes including temperature-induced LLPS, interfacial\nadsorption of the condensates and possible crosslinking as well as interfacial solvent transport\nare well understood and leveraged to explain and control the active forces generated. The\nauthors also provide a convincing biophysical model incorporating principles of elasticity,\nthermodynamics, pressure dynamics to explain the collective effects of the LLPS on the\ngeneration of active forces. This study represents an big improvement in the understanding of\nprotocells dynamics and provides design principles for more functional protocells.\nConsequently, I strongly support the publication of this study in Nature Communications after\nthe minor comments cited below are addressed.\nReviewer #3 (Remarks to the Author):\nThe manuscript presents a novel cell model based on elastin that exhibits temperature-\nresponsive behavior. The study systematically investigates the contractile dynamics, force\ngeneration, and cargo release driven by protein liquid-liquid phase separation. By integrating\nexperimental approaches with theoretical modeling, the research elucidates how\ncompartmentalized structures amplify piconewton-scale forces from phase separation to\nperform large-scale mechanical work. The topic is highly innovative and sits at the intersection\nof synthetic biology, biophysics, and soft matter research. The experimental design and model\nconstruction in this manuscript are both sound and appropriate. However, some issues related\nto mechanistic explanation and data presentation need to be addressed before the manuscript\ncan meet the standards for publication in Nature Communications.", "reviewer_2": "In the present paper Jia et al. present a new approach introducing a contractile, elastin-based protocell system in which liquid–liquid phase separation is key in the contraction observed. This work highlights that condensation is able to develop forces at the piconewton scale that can be amplified and eventually lead to observable expelling of components from inside the protocell. Several cascade processes including temperature-induced LLPS, interfacial adsorption of the condensates and possible crosslinking as well as interfacial solvent transport are well understood and leveraged to explain and control the active forces generated. The authors also provide a convincing biophysical model incorporating principles of elasticity, thermodynamics, pressure dynamics to explain the collective effects of the LLPS on the generation of active forces. This study represents an big improvement in the understanding of protocells dynamics and provides design principles for more functional protocells. Consequently, I strongly support the publication of this study in Nature Communications after the minor comments cited below are addressed.\nSpecific comments:\nFigure 1:\n  The authors should clarify the interest of conjugating pNIPAM to elastin. Is it to allow for the thermosensitive LLPS of elastin, or to increase surface activity? Or both?\n  Elastin-like polypeptides show intrinsic LCST properties, is it also the case for natural elastin? If yes, how does it impact the thermoresponsive behavior of the system? In this direction, the comparison with BSA-pNIPAM conjugates is interesting however its is not clear what should be understood from it. Please could you clarify this point.\n  In Figure 1c, after 30 min, the elastin does not look to be anymore especially concentrated at the interface? Can the authors clarify this point?\n  “Contraction of the protocell resulted in an increase in the concentration of the entrapped soluble elastin-polymer nanoconjugates such that LLPS occurred within the interior of the shrinking droplets to produce an encapsulated dispersion of self-coacervated protein molecules, 2.03 ± 0.31μm (n=10) in size (Fig. 1i-j).”\nThis part is unclear to me. What is this 2 µm related to? The layer thickness? How can the authors state that the elastin-rich drops (also called “aggregates” just after to be clarified too) do not merge in an homogeneous membrane contrarily to the scheme proposed in Figure 1a.\nFigure 2:\n  Can the authors comment on the residual water content of the elastin-rich layer around the oil-in-oil droplets? Why does the shrinking doesn’t eventually lead to only elastin-rich droplets in oil continuous phase ? Can the authors subsequently comment on the elasticity of the elastin-rich phase?", "reviewer_3": "The manuscript presents a novel cell model based on elastin that exhibits temperature-responsive behavior. The study systematically investigates the contractile dynamics, force generation, and cargo release driven by protein liquid-liquid phase separation. By integrating experimental approaches with theoretical modeling, the research elucidates how compartmentalized structures amplify piconewton-scale forces from phase separation to perform large-scale mechanical work. The topic is highly innovative and sits at the intersection of synthetic biology, biophysics, and soft matter research. The experimental design and model construction in this manuscript are both sound and appropriate. However, some issues related to mechanistic explanation and data presentation need to be addressed before the manuscript can meet the standards for publication in Nature Communications.\n1. The authors observed that elastin coacervate particles were radially ejected from the protocell surface along linear trajectories under high levels of contraction, creating a striking visual impact. It would be beneficial for the authors to delve deeper into the physical origin of this phenomenon. Specifically, an analysis to determine whether this ejection results from the sudden release of accumulated elastic stress within the membrane or from Plateau-Rayleigh instability would significantly strengthen the manuscript.\n2. The manuscript provided limited direct measurement of the active force. Specifically, the model predicts a force of approximately 123 pN, whereas the experimental estimate is around 5.0 zN, a discrepancy of over ten orders of magnitude. This vast difference is concerning. The paper currently lacks more direct and compelling experimental evidence to substantiate the existence and precise magnitude of these forces. For instance, techniques such as membrane deformation could provide more robust validation. Additionally, the reliability of the current estimation methods warrants further in-depth discussion.\n3. The authors' explanation that the irregular contraction is due to short and rigid crosslinks appears oversimplified and requires further substantiation. Specifically, additional experimental evidence should be provided to support this claim.\n4. In the modeling section, the derivation or determination of key parameters in the model (such as κ, α, Tmelt) from experimental data is not clearly explained. The statement “The simulations accurately reproduced the experimental trends” lacks quantitative validation, such as a fit coefficient (e.g., R²) to demonstrate the degree of correspondence between the model predictions and experimental observations. Moreover, there is an inconsistency in the notation of the parameter α. In equations (2) and (4), the parameter α is used but has a different meaning compared to α in equation (1).\n5. Several conclusions are drawn based on qualitative “observations” from microscope images, which lack robust quantitative support. For instance, the formation of “distinct irregular patches” depicted in Fig. 3 and the fusion and ejection frequency of phase-separated droplets illustrated in Fig. 4 would benefit from quantitative data to substantiate their generalizability. Similarly, when discussing the “slower rates of contraction” on Page 7, the manuscript would be significantly strengthened by incorporating specific quantitative comparisons. For example, citing the time required for the structure to shrink to 50% of its original volume would provide a clear, measurable metric to support the conclusion. This approach would enhance the credibility and persuasiveness of the findings.\n6. The methodology for measuring protocell diameters in Fig. 2b needs clarification, specifically regarding the sample size for statistics and whether the values represent a mean or a measurement from the largest protocell.\n7. The chemical structures of the crosslinkers in Fig. 3a is obscured and needs to be revised for clarity.\n8. To improve clarity, Fig. 1c should be revised to explicitly highlight the variations resulting from time and temperature respectively. The current presentation could potentially mislead readers.\n9. The manuscript exhibits several issues related to writing and presentation, which are primarily reflected in the following aspects: incorrect unit formats; disorganized figure and text citations; failure to define necessary concepts or redundant definitions; inconsistent reference formats."}, "author_response": "We sincerely thank the reviewer for the positive assessment of the experimental design, visual quality, and the novelty of the exocytosis-like expulsion phenomenon, as well as for the careful and constructive evaluation of our work. We have thoroughly revised the manuscript to address all concerns, including correcting force estimations, refining the physical basis of the model, and explicitly acknowledging model limitations. Specifically, we identified and corrected the error in the force calculation and accordingly revised the force values and interpretations. These clarifications and corrections ensure that the mechanical conclusions are physically grounded and transparently supported by the data. Together, these revisions substantially improve the rigor, transparency, and interpretability of the study while preserving the core experimental findings.\nMajor concern:\n1. The term “Programmbale” in the paper doesn’t demonstrate sufficient and precise programmable capability beyond the expansion/shrinkage? For example, can the water-in-oil droplet be programmed to a certain size, or programmed at a certain shrinkage rate, or could be re-programmed after once? I agree that nested coacervate formation and cargo release are really cool! . but could you demonstrate how you are going to program it?\nResponse: We thank the reviewer for this insightful comment. In our current study, we use the term “programmable” to describe the tunable dynamic behaviors of the protocells that can be modulated by varying temperature or using different crosslinkers. At present, our approach allows control over distinct dynamic responses rather than precise programming of parameters such as final size, shrinkage rate, or reversibility. We agree that these aspects would represent true programmable capabilities, and we are actively exploring strategies to achieve them in future work. To avoid potential misunderstanding, we have revised the manuscript to remove the term “programmable” and clarify our description accordingly.\nPage 1, line 23-24: Here, we engineer a temperature-responsive elastin-based protocell model displaying temperature-modulated contractility and attendant force harnessing.\nPage 2, line 27-31: As a step towards addressing these challenges, we present a simple protocell model based on the stabilization of water-in-oil microemulsion droplets with a self- coacervating protein (elastin)20,21 membrane capable of exhibiting temperature-modulated contractability and attendant force harnessing within controllable confined spaces.\n2. “Temperature-responsiveness” is from the protein Amine-modified elastin (elastin-NH ).\n2\nin-oil droplet? Additionally, could specify the temperature limitation for this? The temperatures\nyou choose (20 °C, 25 °C, 33 °C, 35 °C, and 37 °C) are discrete data points. Could you explain\nfig1b why the amine-modified elastin( not the protocell) changes from 20 °C to 40 °C?\nResponse: We thank the reviewer for these valuable comments. We have performed additional experiments to characterize the temperature-dependent coacervation of elastin-NH₂ in solution, without forming water-in-oil droplets, and have clarified the corresponding temperature range in the revised manuscript (Extended Data Fig. 1d). The results are in according with Fig. 1d.\nElastin-NH₂ exhibits typical lower critical solution temperature (LCST) behavior: phase separation is initiated above a threshold temperature, driven by enhanced hydrophobic interactions among elastin segments (Biomacromolecules 2008, 9, 1739–1744). The extent and rate of coacervation increase progressively from 20 °C to 40 °C, which explains the trend observed in Fig. 1b. We have now specified these observations and the temperature limitation in the revised version. (Page 3, line 13-15; Page 3, line 21-26)\nd-e) Thermal response of Elastin -NH₂ and Elastin-NH₂-PNIPAAm at different temperature.\nScale bar, 10 µm.\nPage 3, line 13-15: Elastin–pNIPAAm conjugates exhibited temperature responsiveness\n(Extended Data Fig. 1), but instead of forming simple coacervates as elastin (Extended Data\nFig. 1a), they generated smaller particle-like domains that assemble into a network.\nPage 3, line 21-26: The emulsion droplets were stable when stored at 4°C but exhibited a\ntemperature-dependent decrease in mean diameter above 20°C (Fig. 1b, Extended Data Fig.\n2c). This diameter reduction was consistently observed by both confocal and wide-field\nimaging (Extended Data Fig. 2d), and is consistent with the thermos-responsive behavior of\nelastin and its nanoconjugates (Extended Data Fig. 1).\n3. Could you explain for 1g why the fluorescence intensity of inner Elastin-atto-647 drops between 5-20min?\nResponse: We thank the reviewer for this helpful comment. The decrease in inner Elastin-\nAtto-647 fluorescence between 5 and 20 min reflects the ongoing coacervation process. As\ntemperature-induced phase separation proceeds, elastin molecules migrate and associate with\nthe water–oil interface, leading to reduced inner fluorescence and enhanced interfacial signal.\nWe have clarified this explanation in the revised manuscript (Page 4, line 5-8).\nPage 4, line 5-8: During this coacervation process, the inner elastin–Atto647 fluorescence\ndecreased between 5 and 20 min due to the migration and accumulation of elastin molecules\nat the interface, resulting in reduced internal signal and a corresponding increase in interfacial\nfluorescence (Movie S1).\n4. They are astonishing videos (video 1 and 2) but I wonder whether there is z-direction movement (protocells drifting in the z-direction) that influences the size of protocells, and makes them seem smaller. Suggested experiment: Could you use 3D confocal imaging to ensure the protocell size changes and it’s not influenced by the z-direction motion?\nResponse: We thank the reviewer for this thoughtful suggestion. Indeed, the elastin-based\nprotocells physically shrink upon temperature stimulation. While 3D confocal (z-stack)\nimaging would more precisely capture the volumetric changes, it is technically challenging to\nperform high-resolution 3D imaging during rapid dynamic processes. To better address this\nconcern, we have included additional wide-field imaging results (Extended Data Fig. 1d),\nwhich confirm that the observed size reduction is due to genuine shrinkage rather than z-\ndirectional movement. We have clarified this point in the revised manuscript (Page 3, line 23-\n26).\nPage 3, line 23-26: This diameter reduction was consistently observed by both confocal and\nwide-field imaging (Extended Data Fig. 2d), and is consistent with the thermos-responsive\nbehavior of elastin and its nanoconjugates (Extended Data Fig. 1).\n5. For the method, using modified proteins as bio-surfactants to create protocells is really\ncreative, but will DMSO and other strong solvents in your method denature/ impact the protein?\nResponse: We thank the reviewer for this important comment. While high concentrations of\nDMSO or other organic solvents can denature proteins, in our experiments only a very low\namount of DMSO was used—well below levels known to disrupt protein structure. No\nprecipitation or loss of functionality was observed during sample preparation, and the modified\nelastin retained its temperature responsiveness, indicating preserved structural integrity. We\nhave clarified this point in the Methods section (Page 17, line 15-18).\nPage 17, line 15-18: Fluorescein isothiocyanate isomer I (FITC, sigma, F4274) solution in\nDMSO was added dropwise to elastin-NH (2 mg/mL in 0.1 M carbonate buffer with pH 9.0),\n2\nwith continuous mixing to avoid high local DMSO concentrations and thereby prevent protein\ndenaturation.\nFor the modelling:\n6. I have concerns about the oversimplification of the linear relationship in Eq.1, where\nshrinkage rate dR/dt is linearly correlated to pressure differential. Regarding the equation 8 and\n9, osmotic pressure, shouldn't they be coupled to equation 1 to explain the shrinkage? For\nextended data figure 3b, it appears more like data-fitting than a physical model.\nResponse: We thank the reviewer for this very constructive comment. We have clarified the\nphysical basis of Eq. 1 and its coupling to the capillary pressure contributions (Eq. 8), and we\nhave revised the theoretical section accordingly (SI-Page 2, line 8-14). In the updated version,\nnew Eq. 1 is derived within a low–Reynolds-number framework by relating the radial\nshrinkage velocity of the vesicle to the net normal stress at the interface, which includes\npressure, capillary, and LLPS-induced contributions.\nRegarding Extended Data Fig. 3b, our intention was to perform a global fit of the model to\nthe experimental shrinkage data in order to extract effective parameters that are difficult to\nobtain directly from experiments. We have now explicitly stated this purpose in the figure\nlegend.\nSI-Page 2, line 8-14:\ndR(t) 2γ(T)\n= −κR(t)[(P −P )+ +αΘ(T−T )] (Eq. 1)\nin out c\ndt R(t)\nWhere γ(T)is temperature-dependent interfacial tension, described as γ(T) = γ [1−\n0\nβ (T−T )], where β is the temperature coefficient of surface energy.\nγ 0 γ\nThis equation describes the rate of vesicle shrinkage under the combined influence of internal\nand external pressures, osmotic forces, interfacial tension, and temperature-dependent phase\nseparation effects. The shrinking behavior of the vesicle captures essential dynamics of the\nsystem, including pressure balance, temperature-dependent interfacial tension and LLPS.\nExtended Data Figure 3 b) Global fitting of the protocell radius changes over time…\n7. Also no uncertainty analysis or parameter assessment is provided - - so the peak force of\n123pN is very model/parameter choice dependent and not anchored in their data – need some\njustification/explanation/comments here...\nResponse: We thank the reviewer for this important point. In the revised manuscript we have\n(i) clarified how the model parameters are constrained by the experimental data, (ii) performed\na parameter sensitivity analysis, and (iii) softened the reported force value to emphasize its\napproximate character. Specifically, we now obtain the parameter set by global fitting the\nvesicle shrinkage trajectories at different temperature (SI-Theoretic section, 7) Global fitting\nand mechanical parameter extraction). To assess how parameter uncertainty propagates to the\npredicted force, we systematically varied each parameter within ±20% within the\nexperimentally estimated error range and recalculated the simulated force–time curves\n(Extended Data Fig. 5). Across all parameter sets compatible with the data, the force remained\nwithin 102–103 pN. We therefore now report the peak force as approximately 103 pN rather\nthan a single value and explicitly state that this number represents the typical magnitude\npredicted for parameter sets that fit the experimental data. We have now clarified this in the\nmain text (Page 6, line 31- Page 7, line 2).\nPage 6, line 31- Page 7, line 2: Using these parameters, the model predicted a pressure\ndifferential across the elastin-based interface (Extended Data Fig. 5d), corresponding to a\nforce on the order of 10²–10³ pN at 37°C for protocells with a diameter of 25 µm. This\nmagnitude of force is sufficient to initiate the inward contraction of the protocell (Extended\nData Fig. 5e).\n8. The authors say “Estimating the acceleration of the particles gave a maximum net force\nacting in the radial direction of ca. 5.0 zeptonewtons. “I had to do a double check as in my\nentire career i’ve never seen the word or number of zeptonewtons (10^-21!!!)... are the authors\nentirely sure about this? For example, for a 2.5um particle with a velocity of 0.8 um/s, for\nwater-like viscosity (1 mPa), you get force of 40fN – this is 7 orders of magnitude larger than\n5zN – this makes me very nervous about the modeling sanity checks! My suspicions is that the\nauthors use F=ma instead of Stokes law, F=6\\pi\\eta R v.... Which is teh right choice at low Re.\nResponse: We thank the reviewer for this important and insightful comment. We agree that\nthe previously reported value of ~5 zeptonewtons was unrealistically small. Upon re-examining\nour analysis, we confirmed that this discrepancy resulted from using the inertial term F=ma\nrather than the appropriate viscous drag expression F=6πηRv, which indeed governs particle\nmotion in the low Reynolds number regime relevant to our system. We have now recalculated\nthe forces using Stokes’ law. The results are consistent with the reviewer’s estimate. We have\ncorrected the force values in the revised manuscript (Page 10, line 16-18) and clarified the\nanalysis in the method section (Page 20, line 5-13).\nPage 10, line 16-18: Using Stokes’ law together with the measured median velocity, the\nviscous drag force was calculated to be F ≈0.31±0.06 pN. Under low–Reynolds-number\ndrag\nconditions, this drag balances the active launching force, such that F ≃F .\nlaunch drag\nPage 20, line 5-13: Under low-Reynolds-number conditions, particle motion is governed by a\nbalance between the active launching force and viscous drag. To quantify the viscous drag\nacting on each particle, we used the measured steady-state velocities and Stokes’ law,\nF =6πηr v., where 𝑟 is particles radius, 𝑣 𝑖𝑠 𝑣𝑒𝑙𝑜𝑐𝑖𝑡𝑦 , 𝜂 𝑖𝑠 𝑣𝑖𝑠𝑐𝑜𝑠𝑖𝑡𝑦 𝑜𝑓 𝑚𝑒𝑑𝑖𝑢𝑚 .\ndrag p 𝑝\nBecause inertial contributions are negligible in this regime, the viscous drag force directly\nbalances the capillary- or elasticity-derived launching force. Thus, the active force driving\nparticle motion at steady state is approximated as: F ≃ F =6πηr v. This estimation\nlaunch drag p\nassumes a particle diameter of 5 μm and 2-ethyl-1-hexanol viscosity 𝜂 of 8 mPa·s at 37 °C.\n9. The interfacial tension of .033 pn/um is estimated – however wont this be a function of\ntemperature / crosslink density? – is there a way to calibrate/cross-validate this?\nResponse: We appreciate the reviewer’s concern regarding the potential dependence of\ninterfacial tension on temperature and crosslink density. We agree that the interfacial tension\nof elastin-based protocells is expected to vary with temperature and molecular organization. In\nthe main text, the reported value of 0.033 pN μm⁻¹ corresponds specifically to the interfacial\ntension at 37 °C, and we have now clarified this point explicitly in the revised manuscript (Page\n7, line 5–7). To account for temperature effects, we have incorporated an explicit temperature\ndependence of the interfacial tension into the revised model. Calibration against shrinkage\nkinetics measured at different temperatures yields a consistent and physically grounded\nestimate of γ(T), as shown in Extended Data Fig. 5c, thereby providing cross-validation of the\ntemperature dependence. Regarding crosslink density, we agree that chemical crosslinking can\nsignificantly influence elastin stiffness, hydration, and interfacial properties. However, in the\npresent model system, no chemical crosslinkers are introduced, and the observed shrinkage\ndynamics are dominated by liquid–liquid phase separation of elastin rather than crosslink-\ndriven elasticity. We have clarified in the revised text that crosslinking is not a contributing\nfactor in this specific model (SI-Page1, line 21–23; Page 5, line 3–7).\nPage 7, line 5–7: Moreover, the model simulated an increase in membrane tension as a\nfunction of time and decrease in droplet radius, eventually reaching a maximum tension of ca.\n0.05 pN/µm at 37°C (Extended Data Fig. 5f)\nSI-Page1, line 21–23: Our model simulates the dynamics of an elastin-based protocell system\n(non-crosslinked), where vesicles undergo shrinkage due to phase separation triggered by\ntemperature changes.\nPage 5, line 3–7: This simplification highlights natural directions for future work, including\nexplicit incorporation of solvent and solute transport, partitioning between aqueous and oil\nphases, crosslink density–dependent elasticity and interfacial flows driven by surface-tension\ngradients (e.g., Marangoni effects), which are likely to underlie more complex behaviors such\nas transient swelling, oil engulfment, and non-monotonic size changes.\n10. For the modelling and volume shrinkage, I'd like to know where the solution inside of\ndroplets goes. There should be some mass-conservative equation. If it diffuses outward, could\nyou explain what facilitates such rapid diffusion?\nResponse: We thank the reviewer for this insightful comment. Our mechanical model focuses\non predicting membrane tension and forces from the experimentally measured shrinkage rate.\nFor this reason, we assume that all solute mass remains within or on the protocell, and only\nwater volume changes—consistent with our experimental observations of rapid water efflux\nand occasional late-stage oil influx.\nWe agree that a complete mass-balance model, including coupled solvent fluxes and solute\nexchange, would be valuable for understanding transport mechanisms such as water\npermeation or droplet engulfment. However, these processes involve temperature-dependent\nchanges in membrane permeability and oil–water partition interactions that lie beyond the\nscope of the present study. We have clarified this point in the revised manuscript, point out the\nlimitation of our model, and outlined such transport modeling as an important direction for\nfuture work (SI-Page 4, line 25- Page 5 line).\nSI-Page 4, line 25- Page 5 line: 8) Model limitations and future opportunities\nThe theoretical framework presented here is intentionally minimal and designed to isolate the\ndominant mechanical contributions governing protocell contraction under temperature-\ntriggered phase separation. By focusing on an effective interfacial stress balance inferred from\nexperimentally measured shrinkage kinetics, the model enables quantitative estimation of\nmembrane tension and force generation without introducing poorly constrained transport\nparameters. As a result, osmotic pressure and explicit mass-conservation equations are not\nincluded; instead, volume changes are treated phenomenologically through the observed\nradius evolution, consistent with experimental observations of rapid water efflux and episodic\noil uptake. This simplification highlights natural directions for future work, including explicit\nincorporation of solvent and solute transport, partitioning between aqueous and oil phases,\ncrosslink density–dependent elasticity and interfacial flows driven by surface-tension gradients\n(e.g., Marangoni effects), which are likely to underlie more complex behaviors such as\ntransient swelling, oil engulfment, and non-monotonic size changes. In this context, the present\nmodel serves as a first-order mechanical backbone that captures the dominant stress-driven\ncontraction regime and provides a foundation for more comprehensive, transport-resolved\ndescriptions of protocell dynamics.\n11. For equation 2, could you justify the “ theta(T-Tmelt)” term, melting temperature (Tm) is\na solid-liquid transition temperature, not directly the temperature of liquid-liquid phase\nseparation.And you set T_melt to ~ 18.19 Celsius in Extended figure 3b, But for the figure 1b,\nfor 4 Celcius, the size also shrinks as well? Could you elaborate more on (T-T_melt) ?\nResponse: We appreciate the reviewer’s careful reading and thank for pointing out the\nconfusion regarding the use of the term “melting temperature.” In our revised model, the\nHeaviside term Θ(𝑇−𝑇 ) no longer refers to a melting transition but instead represents the onset\n𝑐\ntemperature for liquid–liquid phase separation (LLPS) of elastin. We have replaced the symbol\n𝑇 with 𝑇 throughout the manuscript to accurately reflect this LLPS activation threshold.\nmelt 𝑐\nThe purpose of this term is to switch on the LLPS-related pressure contribution only at\ntemperatures above the LLPS threshold. Below 𝑇 such as 4 °C, vesicle shrinkage can still\n𝑐\noccur due to temperature-dependent changes in interfacial tension and intrinsic elasticity,\nwhich are captured by the other terms in our model. We have clarified the terminology, updated\nthe notation, and explicitly explained the physical meaning of Θ(T−T ) in the revised\nc\nmanuscript (SI-Page 1, line 36- Page 2, line 3).\nSI-Page 1, line 36- Page 2, line 3: Phase separation effects were introduced to account for\ntemperature-dependent behavior. When the temperature (𝑇) exceeds a critical threshold (𝑇),\nc\nLLPS exerts an additional pressure on the vesicle’s interior, promoting further shrinkage. This\neffect is represented by a 𝛼𝛩(𝑇−𝑇 ), where 𝛼 is the LLPS coefficient and 𝛩 is the Heaviside\n𝑐\nstep function, and is that activates this term only when 𝑇 is above 𝑇 3.\nc\n12. Eq.1 and Eq.4 creates a circular definition for P_in if P_in was supposed to be\nthe driving in the dynamics in the first place.\nResponse: We thank the reviewer for raising this point. There is no circularity between Eq. 1\nand Eq. 4, as they play distinct and sequential roles in the analysis. Eq. 1 is used as a forward\ndynamical model to fit the experimentally measured diameter–time trajectories. In this step,\nP appears as an effective, time-averaged internal pressure parameter that captures the net\nin\ninward stress driving contraction and is determined through global fitting. Eq. 4 is not used as\nan input to the dynamics. Instead, after all parameters are obtained from Eq. 1, Eq. 4 is applied\nto reconstruct the time-dependent internal pressure P (t) by inserting the experimentally\nin\nmeasured radius R(t) and its time derivative. Thus, Eq. 4 serves as a diagnostic relation and\ndoes not feed back into Eq. 1.\nWe have clarified this workflow in the revised manuscript to explicitly distinguish between\nthe fitted effective pressure parameter and the reconstructed instantaneous internal pressure\n(SI-Theoretic section: 7) Global fitting and mechanical parameter extraction).\n13. In the paper, “undergoing transient enlargement indicated that the increase in volume was\nassociated with the irreversible uptake of oil droplets into the protocells along with a non-\ncompensating efflux of water”And also could you elaborate more why/ what triggers the\n“bigger → the latter smaller” transitions in the late stage ? If it's an oil influx (btw, figure 1e\nseems too noisy to recognize the oil drop position to support the oil influx) what triggers the\nsudden oil influx? Why is fig2b at 37 Celsius different from figure 1b after 30 mins? What\nhappens to protocells at other temperatures after 30mins in figure 1b?\nResponse: We thank the reviewer for this constructive comment. We propose that the transient\nenlargement followed by renewed shrinkage is associated with LLPS-driven reorganization of\nthe elastin-rich interfacial membrane (Fig. 2c). During this stage, the interface can temporarily\nadopt a heterogeneous and irregular morphology, which may permit transient oil influx and\nlead to a short-lived increase in protocell volume. As the interfacial layer reorganizes into a\nmore continuous and mechanically coherent membrane, further oil influx is suppressed and\ncontraction resumes.\nWe acknowledge that oil droplets are difficult to observe in Fig. 1e due to the rapid dynamics\nand imaging noise. Fig. 2e–f, obtained under a slower temperature ramp, more clearly\nillustrates oil influx events during this transient enlargement phase. The apparent difference\nbetween Fig. 1b and Fig. 2b at 37 °C arises from the different temperature protocols: Fig. 2b\nemploys a gradual temperature increase, resulting in slower overall dynamics, such that the\nstate at 30 min is not directly comparable to Fig. 1b. These points have now been clarified in\nthe revised manuscript and figure captions (Page 6, line 12-14; Page 8, line 4-6).\nPage 6, line 12-14: Quantitative analysis of temperature-dependent changes in protocell\ndiameter and fluorescence intensity of entrapped FITC-dextran during gradual heating from\nroom temperature to 37 °C…\nPage 8, line 4-6: b) Time course of protocell diameter and associated fluorescence intensity of\nprotocell contraction and enlargement induced by gradual heating from room temperature to\n37 °C. Data are shown as mean ± s.d.; n = 3.\n14. I will note that some of the most striking behaviors such as the transient swelling, influx,\nwater efflux are transport driven -such as permeation, solvent partitioning and potentially even\nmarangoni flows. I respect the authors attempt at using a simple model, and indeed there is\nvalue in simple models, but acknowledgment that this simple model overlooks a lot of key\ntransport ideas relevant to their experiments is key - for posterity. Also it limits any\n`explanatory power’ or predictive power of their model - which should be acknowledged.\nResponse: We thank the reviewer for these insightful comments. Our current model was\nintentionally formulated as a minimal, simplified description to capture the shrinkage kinetics,\nrather than to account for the full complexity of interfacial transport phenomena.\nWe recognize that by neglecting explicit solvent transport, interfacial diffusion, and surface-\ntension-gradient–driven flows, the model has limited explanatory and predictive power with\nrespect to these richer dynamical behaviors. To address this concern, we have added a\ndedicated paragraph in the model section explicitly outlining these limitations and clarifying\nthat the model should be interpreted as a phenomenological framework rather than a\ncomprehensive physical description. We also note that incorporating these transport\nmechanisms will be an important direction for future theoretical development aimed at\nimproving predictive capability(Page 14, line 17-21; SI-Page 4, line 25- Page 5 line).\nPage 14, line 17-21: In the present study, these forces are investigated and estimated through\na combination of experiments and a simplified physical model. Further development of\nquantitative mechanical characterization approaches will be required to more fully resolve the\nnon-homogeneous elasticity and spontaneous force generation in future work.\nSI-Page 4, line 25- Page 5 line: 8) Model limitations and future opportunities\nThe theoretical framework presented here is intentionally minimal and designed to isolate the\ndominant mechanical contributions governing protocell contraction under temperature-\ntriggered phase separation. By focusing on an effective interfacial stress balance inferred from\nexperimentally measured shrinkage kinetics, the model enables quantitative estimation of\nmembrane tension and force generation without introducing poorly constrained transport\nparameters. As a result, osmotic pressure and explicit mass-conservation equations are not\nincluded; instead, volume changes are treated phenomenologically through the observed\nradius evolution, consistent with experimental observations of rapid water efflux and episodic\noil uptake. This simplification highlights natural directions for future work, including explicit\nincorporation of solvent and solute transport, partitioning between aqueous and oil phases,\ncrosslink density–dependent elasticity and interfacial flows driven by surface-tension gradients\n(e.g., Marangoni effects), which are likely to underlie more complex behaviors such as\ntransient swelling, oil engulfment, and non-monotonic size changes. In this context, the present\nmodel serves as a first-order mechanical backbone that captures the dominant stress-driven\ncontraction regime and provides a foundation for more comprehensive, transport-resolved\ndescriptions of protocell dynamics.\nMinor:\n1. “LLPS complexes” appears in the abstract for the first time without explanation. Could you\nwrite the “protein liquid-liquid phase separation (LLPS)” in the abstract for the readers’ benefit?\nResponse: We have revised the abstract to introduce the term in full as “protein liquid–liquid\nphase separation (LLPS)” for clarity to readers encountering it for the first time.\nPage 1, line 26-29: Crosslinking of the elastin-based membrane influences the contraction\ndynamics such that the accumulation of mechanical forces in the protocells results in the\nspontaneous expulsion of internally trapped protein liquid-liquid phase separation (LLPS)\ncomplexes.\n2. Figure 2h , could you specify the temperature in degrees Celsius? or specify it in the caption ?\nIn figure 4a, for content condensing T Celcius , and T Celsius compression, could you specify\nwhat exact temperatures?\nResponse: We have now specified all temperatures in degrees Celsius in the figure panels.\nPage 9, line 3-4: i) Time-lapse series of the condensation and fusion of oil droplets within\nprotocells encapsulated with PEG-3300 at 37 °C.\nPage 13, line 10-11: a) Schematic illustration of nested coacervate formation within the\nprotocell, driven by thermally induced co-coacervation of elastin and dextran at 37 °C.\n3. Figure 3a: The chemical structure is overlapped by the arrow.\nResponse: We thank the reviewer for this pointing out. We have adjusted Figure 3a to ensure\nthat the chemical structure is clearly visible and no longer overlapped by the arrow.\n4. Figure 3k and video “645821_0_video_750257_t0vk4v”: the tracking is not continuous.\nSome trajectories are zig-zagged, possibly because the frame rate for tracking is too small?\nCould you improve the tracking and verify the tracking and the following analysis?\nResponse: We thank the reviewer for this thoughtful comment. The occasional non-continuous\nor zig-zag trajectories are not caused by an insufficient frame rate but instead reflect the highly\ndynamic and complex behavior of particles during fission events, where rapid changes in\nparticle shape, intensity, and motion make continuous identity assignment challenging. The\nimaging was already performed at a high temporal resolution, and further increasing the frame\nrate would require compromising spatial resolution and signal-to-noise, which would in fact\ncomplicate tracking. We carefully re-examined the tracking and verified that the quantitative\ntrends remain robust.\n5. Could you quantify the diffusion of debris after the breakout (~ 300s) in the video\n“645821_0_video_750256_t0dk4v”? The movement appears too fast to be simple diffusion?\nCould you quantify and verify the rate?\nResponse: We thank the reviewer for this comment. The quantitative tracking of the debris\ntrajectories, including their paths and velocities after the breakout event, is provided in Fig. 3i–\nm. These data explicitly quantify the post-breakout dynamics and demonstrate that the motion\nis fast and directed, rather than diffusive. To avoid any potential confusion, we have now\nclarified this point in the main text and more explicitly referenced Fig. 3i–m when discussing\nthe post-breakout behavior (Page 10, line 11-14.\nPage 10, line 11-14: The propelled particles traveled linearly for 71.74 ± 16.69 µm within 110\nseconds with a median speed of 0.82 ± 0.16 µm/s (Fig. 3j-k), which was approximately 2.65\ntimes higher than the speed of free-diffusing particles (0.31 ± 0.13µm/s) under similar\nenvironmental conditions.\n6. Typo of ratical launch > radial\nResponse: We have corrected the typo.\nPage 10, line 6-8: Surprisingly, the elastin coacervate particles were radially ejected from the\nprotocell surface along linear trajectories under high levels of contraction\nPage 20, line 2-5: Parameters such as traveling speed, max distance traveled, and radial\ndisplacement were analyzed within the TrackMate framework.\n7. Typo of lengths> lengths\nResponse: We have corrected the typo.\nPage 11, line 2-3: a) Protein crosslinkers with different lengths."}, {"round": 2, "reviewer_comments": {"reviewer_1": "Reviewer #1 (Remarks to the Author):\nSummary: This work uses modified proteins (elastin-Nh2) forming water-in-oil microemulsion\ndroplets and shows the temperature effects on the droplet, with modelling to understand the\nunderlying principles. The experiments are amazing! And visuals stunning! But I have some\nconcerns about this work, especially about the model which has some overextended claims and\npotentially erroneous calculation of zetaNewtons– my estimate shows their numbers could be\n7 orders of magnitude off (however, i will admit, i did a quick and dirty back of the envelop\ncalculation - so i could also be wrong - but clarification is definitely needed!). However, if i’m\nright, then the core claim about force `amplification’ are weakly validated. Nonetheless, the\nexocytosis-like expulsion is super cool!\nReviewer #2 (Remarks to the Author):\nIn the present paper Jia et al. present a new approach introducing a contractile, elastin-based\nprotocell system in which liquid–liquid phase separation is key in the contraction observed.\nThis work highlights that condensation is able to develop forces at the piconewton scale that\ncan be amplified and eventually lead to observable expelling of components from inside the\nprotocell. Several cascade processes including temperature-induced LLPS, interfacial\nadsorption of the condensates and possible crosslinking as well as interfacial solvent transport\nare well understood and leveraged to explain and control the active forces generated. The\nauthors also provide a convincing biophysical model incorporating principles of elasticity,\nthermodynamics, pressure dynamics to explain the collective effects of the LLPS on the\ngeneration of active forces. This study represents an big improvement in the understanding of\nprotocells dynamics and provides design principles for more functional protocells.\nConsequently, I strongly support the publication of this study in Nature Communications after\nthe minor comments cited below are addressed.\nReviewer #3 (Remarks to the Author):\nThe manuscript presents a novel cell model based on elastin that exhibits temperature-\nresponsive behavior. The study systematically investigates the contractile dynamics, force\ngeneration, and cargo release driven by protein liquid-liquid phase separation. By integrating\nexperimental approaches with theoretical modeling, the research elucidates how\ncompartmentalized structures amplify piconewton-scale forces from phase separation to\nperform large-scale mechanical work. The topic is highly innovative and sits at the intersection\nof synthetic biology, biophysics, and soft matter research. The experimental design and model\nconstruction in this manuscript are both sound and appropriate. However, some issues related\nto mechanistic explanation and data presentation need to be addressed before the manuscript\ncan meet the standards for publication in Nature Communications.", "reviewer_2": "The authors provided convincing response to my comments.\nThus, I support the publication of the manuscript in Nature Communications as it is.\nCongratulations.", "reviewer_3": "The authors have provided detailed responses to the reviewers' comments and have conducted additional experiments to further substantiate their claims. These revisions have effectively addressed the concerns raised. I am satisfied with the improvements made and recommend that this manuscript be accepted for publication."}, "author_response": "The authors made the necessary corrections to the paper, and the manuscript reads significantly better. However, 3 points require clarification, which will help the readers to understand and follow the work presented in this paper:\nResponse: We sincerely thank the reviewer for the positive assessment of our revisions and for recognizing the improvements in clarity. We appreciate the opportunity to further refine the manuscript and address the remaining points. Below, we provide detailed responses to each of the three issues raised, with corresponding revisions incorporated into the main text and Supporting Information to improve clarity and accessibility for readers.\n1- The manuscript attributes droplet contractility to LLPS of the elastin-based conjugates and uses BSA–PNIPAAm as a control. This control supports the point that PNIPAAm alone is not sufficient, but it does not address what PNIPAAm contributes in the elastin–PNIPAAm conjugate, especially at 37 °C where PNIPAAm collapse could plausibly act as a switch that sharpens or gates the response. Please add a clear mechanistic statement that separates PNIPAAm’s role in providing amphiphilicity and interfacial assembly from any role of PNIPAAm’s LCST-type transition in modulating the onset/kinetics of elastin LLPS and contraction under the experimental conditions. If feasible, a more discriminating control would be elastin conjugated to a non-thermoresponsive polymer of similar size (e.g., PAAm or PEG; PAA may introduce additional charge/pH effects), to decouple elastin LLPS from PNIPAAm LCST effects. If that experiment is not feasible, please discuss this limitation explicitly. It would also help to cite a standard reference for PNIPAAm LCST behavior (e.g., Heskins and Guillet, J. Macromol. Sci. Chem., 1968) and note that LCST can shift with salt and conjugation.\nThis discussion will also support the liquid-liquid phase transition dynamics observed in this work.\nResponse: We thank the reviewer for this insightful comment. In the revised manuscript, we have added a dedicated mechanistic clarification paragraph (Page 3, lines 13–23. Briefly, we now state explicitly that elastin LLPS provides the dominant thermodynamic driving force for droplet condensation and contractility, whereas PNIPAAm primarily contributes amphiphilicity that promotes interfacial enrichment and stabilizes a mechanically active interface. We further clarify that PNIPAAm’s LCST-type transition may promote the temperature response and potentially influence the kinetics of interfacial densification, but that our current data indicate it is not sufficient on its own to generate contractility, as demonstrated by the BSA–PNIPAAm control.\nRegarding the suggested discriminating control, we agree that conjugating elastin to a non-thermoresponsive polymer (e.g., PEG or PAAm) would provide a more rigorous decoupling of elastin LLPS from PNIPAAm LCST effects. However, additional polymer modification would be required to enable efficient protein conjugation, and this was not feasible within the scope of the present study. We now acknowledge this limitation explicitly in the revised manuscript (Page 14, lines 20–23). To further support the discussion of PNIPAAm thermoresponsiveness, we have cited the foundational work of Heskins and Guillet (J. Macromol. Sci. Chem., 1968) describing LCST behavior and its environmental sensitivity (Page 3, lines 13–23).\nPage 3, lines 13–23: In this system, PNIPAAm primarily contributes amphiphilicity that promotes interfacial enrichment and stabilizes a water-oil interface (Fig. 1a). Although both elastin and PNIPAAm exhibit lower critical solution temperature (LCST)-type thermal transitions1,2, their functional contributions are distinct. Elastin alone forms simple coacervates (Supplementary Fig. 1a), whereas elastin–PNIPAAm conjugates display enhanced temperature responsiveness (Supplementary Fig. 1), producing smaller particle-like domains that assemble into an interconnected interfacial network. In contrast, BSA–PNIPAAm conjugates form only network-like aggregates without evident LLPS (Supplementary Fig. 1b–c), indicating that PNIPAAm alone is insufficient to drive phase separation. These observations support that elastin LLPS dominates the thermodynamic condensation process, while PNIPAAm modulates interfacial organization and temperature sensitivity.\nPage 14, lines 20–23: While our results support a dominant role of elastin LLPS in driving contractility, a more stringent mechanistic decoupling—such as conjugating elastin to a non-thermoresponsive polymer to fully isolate LCST contributions—was beyond the scope of this work and remains an important direction for future investigation.\n2- This material system is based on elastin, which exhibits liquid-liquid phase transitions. The kinetics of these phase transitions are highly reliant on the source, chemistry, and sequence of elastin. A clarifying statement on the origin, chemistry, and sequence of the elastin used in this system will help readers better understand it. For instance, if it is naturally extracted from an elastin source or purchased from a chemical company, please provide the source or the company. If it is recombinant elastin, providing the recombinant elastin sequence can certainly improve the manuscript's clarity. In either case, please report the degree of amination (or other modification) and the PNIPAAm conjugation level (or a proxy), since these parameters can influence both LLPS and temperature response…\nResponse: We thank the reviewer for these constructive suggestions. In the revised Methods section, we have substantially expanded the description of the elastin component. Specifically, we now clearly state the origin of the elastin material, including source and catalog number (Page 15, line 5-7). We have quantified the degree of amination using a TNBSA assay, and the full assay procedure and calculation details are now provided in the Methods (Page 16, line 8-24). In addition, we have included a clarifying statement reporting both the degree of amination and the PNIPAAm conjugation level used throughout this study (Page 16, line 8-24). We believe these revisions significantly enhance the chemical transparency, reproducibility, and overall rigor of the manuscript.\nPage 15, line 5-7: Typically, an aqueous solution of HDMA (500 mg, 4.3 mmol, 5 mL) was adjusted to pH 6.5 using 5 M HCl and added dropwise to a 5 mL stirred solution of elastin from bovine neck ligament (100 mg, 20 mg/ml, sigma, E6527).\nPage 16, line 8-24: Elastin conjugation level quantification\nPrimary amine content was quantified using a TNBSA (2,4,6-Trinitrobenzene Sulfonic Acid) assay3. Briefly, samples (elastin, elastin-NH2, elastin-NH₂-FITC and elastin-NH₂-FITC-PNIPAAm) were reacted with TNBSA in bicarbonate buffer (alkaline pH) at 37 °C for a fixed incubation time. The reaction product was quantified by absorbance at 335 nm and compared to a glycine calibration curve. Primary amine content was reported as µmol NH per mg protein. Unmodified elastin exhibited a primary amine content of 0.293± 0.001 µmol NH₂ per mg (n=3). After amination (elastin–NH₂), the primary amine content was 1.007 ± 0.005 µmol/mg (n=3). Following FITC labeling (elastin–NH₂–FITC), the accessible primary amine content decreased to 0.666 ± 0.001 µmol/mg (n=3). After PNIPAAm conjugation (elastin–NH₂–FITC–PNIPAAm), the primary amine content further decreased to 0.528 ± 0.002 µmol/mg (n=3). Because PNIPAAm conjugation proceeds via reaction with elastin primary amines, the decrease in TNBSA-accessible amines during this final step provides a quantitative proxy for PNIPAAm conjugation extent. Unless otherwise specified, elastin–NH₂–FITC–PNIPAAm samples used throughout this study correspond to a PNIPAAm conjugation level of approximately 20.7 %, calculated from the fractional decrease in primary amines relative to elastin–NH₂–FITC immediately prior to PNIPAAm coupling.\n3- The model and the associated plots mix interfacial tension, capillary pressure, and membrane tension in a way that is hard to follow. In the SI model, the membrane tension expression (τ = γ/R) has dimensions of pressure, but Extended Data Fig. 5f reports τ in pN/µm and the main text compares it to literature interfacial tensions. It read pretty inconsistent and confusing to me… Please correct the definitions and units and explicitly distinguish interfacial tension γ (N/m), Laplace pressure 2γ/R (Pa), and any membrane-tension term used in the model. Related, the “force” quantity computed from pressure times total surface area should be defined carefully, since it is not a net force on a closed sphere. A short clarification of what physical quantity is being reported and how it should be interpreted would remove ambiguity….\nResponse: We thank the reviewer for the valuable suggestions. In the revised model, we now explicitly distinguish only between interfacial tension (units N m⁻¹) and the corresponding Laplace pressure (units Pa). These quantities are treated separately and reported with consistent units throughout the manuscript. In addition, to avoid ambiguity, we have removed the membrane tension term and related Extended Data Fig. 5f entirely. This revision does not affect the model predictions and overall conclusions regarding LLPS-driven pressure generation and force amplification. Instead, the revised Supplementary Fig. 5f now presents the Laplace pressure (Page 7, lines 5–7).\nRegarding the force calculation, we define the reported quantity as the integrated contractile force, obtained by integrating the pressure-induced normal stress over the droplet surface. This clarification has been incorporated into both the main text (Page 6, line 32–Page 7, line2) and the SI (SI-Page 3, lines 6–12).\nPage 7, lines 5–7: Moreover, the model simulated an increase in capillarity pressure as a function of time and decrease in droplet radius, eventually reaching a maximum pressure of ca. 0.1 Pa at 37°C (Supplementary Fig. 5f).\nPage 6, lines 32–Page 7, line2: Using these parameters, the model predicted a pressure differential across the elastin-based interface (Supplementary Fig. 5d), corresponding to an integrated contractile force on the order of 10²–10³ pN at 37°C for protocells with a diameter of 25 µm.\nSI-Page 3, lines 6–12: This equation defines the integrated contractile force associated with the internal pressure, obtained by integrating the pressure-induced normal stress over the total surface area of the vesicle. For a closed spherical vesicle under uniform pressure, this quantity does represent a scalar measure of the cumulative normal stress acting on the membrane. Because the surface area scales with R(t)2, changes in vesicle radius directly modulate the magnitude of the integrated contractile force. As the vesicle contracts, the increasing pressure differential results in a corresponding increase in the total contractile load borne by the membrane."}, {"round": 3, "reviewer_comments": {"reviewer_1": "looks great - thanks!"}, "author_response": ""}]

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