Membrane interfacial potential governs surface condensation and fibrillation of α-Synuclein in neurons

Abstract

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) are essential for cellular organization. α-Synuclein, an amyloidogenic protein linked to Parkinson’s Disease (PD), undergoes phase separation at high concentrations, but the influence of lipid membranes on this process remains unclear. Here, combining in vitro reconstitution, cell biology, and simulations, we show that membranous interfaces promote α-Synuclein condensation at physiologically relevant sub-critical concentrations ( ~ 10 nM) without crowding agents. Notably, condensation occurs only on membranes with a specific stoichiometry of lipids, underscoring the role of interfacial potential. These condensates serve as nucleation sites for fibril formation, leading to membrane deformation and rupture. A lattice gas model reveals this behavior as a prewetting-like transition, where an attractive membrane induces local phase separation below the bulk saturation concentration. Indeed altering interfacial potential by lipid composition and membrane depolarization not only drastically changes α-Synuclein puncta size and number but also triggers their release from neurons. These findings reveal the crucial role of lipid membrane interfaces in regulating α-Synuclein condensation, aggregation and release, shedding light on a potential mechanism of their cell-to-cell propagation during neurodegeneration.

Authors demonstrate that membrane interfacial potential governs the condensation of α-Synuclein at physiologically relevant concentrations in neurons. These condensates drive fibril formation, which in turn can deform and rupture the membrane.

Introduction

Biomolecular condensates, also known as membraneless organelles, play a critical role in diverse cellular functions¹. These condensates form via phase separation of proteins, nucleic acids, and other biomolecules through weak, multivalent interactions^2,3. When the bulk concentration of such biomolecules exceeds a threshold—known as the saturation concentration—phase-separated condensates emerge⁴. Classic examples include nucleoli⁵, Cajal bodies⁶, and stress granules⁷. A unifying feature of many phase-separating proteins is the presence of intrinsically disordered regions (IDRs) and low-complexity domains (LCDs)^8,9—a characteristic also found in amyloidogenic proteins^10–12. α-Synuclein (α-Syn), an amyloidogenic protein associated with Parkinson’s disease (PD), has been shown to undergo phase separation at high concentrations (~200 µM) and under crowding conditions with polyethylene glycol (PEG)¹³.

Lipid chemical properties—such as headgroup charge and degree of unsaturation—are known to influence α-Syn aggregation kinetics¹⁴. Soluble α-Syn encounters several intracellular membranes, including the inner leaflet of the plasma membrane, synaptic vesicles¹⁵, and organellar membranes. Recent studies reveal that membranous interfaces not only recruit but also regulate biomolecular condensates involved in immune responses¹⁶, cytoskeletal assembly¹⁷, presynaptic contact zones¹⁸, RNP granules in fungi¹⁹, and endocytosis²⁰. However, how membranous interfaces influence α-Syn condensate formation and their transition to fibrils—particularly under physiologically relevant protein concentrations—remains unknown.

In this study, we combine in vitro reconstitution, numerical simulation and live/fixed cell imaging of primary neurons, to explore how membrane composition and interfacial properties regulate α-Syn behaviour. We show that membrane surfaces composed of phosphatidylcholine:phosphatidylserine (PC:PS) at a 6:4 molar ratio promote α-Syn condensation at nanomolar concentrations (~10 nM), even in the absence of crowding agents. This finding underscores the critical role of membrane interfacial potential in driving condensation. These membrane-associated condensates act as nucleation sites for amyloid fibril formation. Using a lattice gas model, we demonstrate that this behaviour resembles a prewetting-like transition, where membrane attraction drives α-Syn condensation on the membrane surface below the bulk saturation threshold²¹. During the formation of α-Syn condensates, membranes undergo increased rigidity. Eventually, as the condensates mature and reach a critical threshold, they rupture - releasing fibrils that cause pronounced membrane deformation. Perturbation of membrane surface charge by external addition of PS liposomes reduced the size of pre-induced α-Syn puncta, likely by inhibiting PS synthase activity and disrupting interfacial potential. On the contrary, neuronal membrane depolarization via depolarizing agents led to ~10-fold increase in α-Syn puncta as well as elevated extracellular release in primary neurons. Together, our findings reveal that membrane interfacial potential is a key regulator of α-Syn condensate formation and fibril nucleation. These condensates, in turn, can remodel and deform membranes, thus promoting fibril propagation, potentially contributing to the pathogenesis of neurodegenerative diseases such as Parkinson’s.

Results

Membrane composition dictates surface-mediated condensation of α-Syn at nanomolar concentrations

α-Synuclein (α-Syn) exists in both soluble and membrane-bound forms. Notably, recent studies have demonstrated that the soluble form can undergo liquid-liquid phase separation (LLPS)¹³. However, the role of membrane surfaces in modulating the phase separation behaviour of α-Syn remains poorly understood. To address this gap, we reconstituted purified α-Syn (Supplementary Fig. 1a) onto supported lipid bilayers that mimic the compositional heterogeneity of neuronal membranes. We first examined α-Syn (50 µM) binding and aggregation kinetics in the presence of membrane vesicles composed of total brain extract (TBE) as well as minimal model membranes made from di-oleoyl-phosphatidylcholine (DOPC) and di-oleoyl-phosphatidylserine (DOPS) at varying molar ratios (DOPC:DOPS = 1:0, 0:1, 6:4, 4:6, 4:1; lipid concentration 200 µM). The Thioflavin T fluorescence signal for purified α-Syn and α-Syn in the presence of DOPC:DOPS (6:4) membrane displayed similar nucleation kinetics, where fibrillation was initiated around 20 h (Fig. 1a). However, the elongation phase of α-Syn appeared slower and thus attained the plateau phase late in the presence of DOPC:DOPS (6:4) membrane. In contrast, TBE, DOPC, DOPS and DOPC:DOPS at 4:1 or 4:6 ratios exhibited significantly slower nucleation kinetics as well as elongation, as reflected in the ThT fluorescence (Fig. 1a). This suggests that the specific stoichiometry of DOPC/DOPS provides a unique lipid membrane environment that promotes nucleation synergistically. Similar ThT fluorescence kinetics were observed for individual DOPC, DOPS and combinations of DOPC:DOPS (1:0, 0:1, 6:4, 4:6, 4:1) at lipid concentrations of 100 μM and 300 μM indicating that membranes ability to modulate α-Syn is independent of lipid concentration (Supplementary Fig. 1b, c).

Fig. 1. Lipid specificity and membrane-induced condensation of α-Syn at nanomolar concentrations.

a ThT aggregation kinetics of 50 μM α-Syn with or without 200 μM LUVs under shaking conditions (n = 3). b–h SLBs (Rhodamine-PE, red) incubated with 100 nM Hilyte-488-α-Syn (green) for 24 h (n = 3; scale bar, 25 μm). b Binding varied with composition: weak on total brain extract (TBE). c delayed on DOPC (d) strong binding and condensates on PC:PS (6:4). e micellar structures on PC:PS (4:6). f No binding on PC:PS (4:1). g No condensates with free Rhodamine-PE or (h) PC:PS (6:4) lipids alone (n = 7). i No condensates on PC:PS (4:6) without protein (n = 3). j Proposed mechanism: α-Syn exists in bulk, adsorbed, or condensed membrane phases. Below a threshold, only adsorption occurs; above it, membrane condensates form, and at higher concentrations, bulk LLPS droplets appear. Confocal images show membrane condensates at 27 h (5–100 nM; scale bar, 10 μm; contrast adjusted for visibility at 5 nM). k Condensate fluorescence intensity increases with protein concentration (25 nM–2 μM), saturating above 100 nM (means ± SD; 159 droplets; ****P < 0.0001, two-tailed, multiple t-tests; n = 3). Relevant source data are provided as a Source Data file.

To investigate the underlying mechanism, we performed long-duration time-lapse imaging of 100 nM Hilyte-labelled α-Syn binding to supported bilayers formed on cleaned glass coverslips. These experiments were conducted under evaporation-free conditions for up to 30 h (Fig. 1b–f). While the physiological bulk concentration of α-Syn is estimated at ~22 µM²², it is known that nanomolar to low micromolar concentrations can suffice for surface-induced aggregation in vitro²³. No detectable binding of α-Syn was observed on TBE, DOPC, or DOPC:DOPS (4:1) membranes over the time-course of 25 h (Fig. 1b, c, f & Supplementary Fig. 2), consistent with previous reports²⁴. However, α-Syn bound readily to DOPC:DOPS bilayers at both 6:4 and 4:6 ratios within a few hours (Fig. 1d, e & Supplementary Fig. 3). Intriguingly, condensation occurred only on the DOPC:DOPS (6:4) membrane, emerging at around 24 h. Most notably, only 100 nM α-Syn was sufficient to drive condensation in the presence of this specific membrane composition (Fig. 1d). α-Syn did not undergo condensation at 100 nM and 2 µM in the absence of membrane surface and crowding in line with previous reports (Supplementary Fig. 4a, b). Membrane surface (PC:PS 6:4) also did not show any condensation over the time course of 26 h in the absence of α-Syn (Supplementary Fig. 4c). No condensation of α-Syn was observed neither in the presence of ~ 1 µM free Rho-PE lipids in solution ruling out the probe’s ability to drive condensation (Fig. 1g), nor in the presence of free-floating DOPC:DOPS (6:4 or 4:6) lipids (Fig. 1h–I and Supplementary Figs. 5, 6), indicating the importance of the membrane surface. In addition, we negated the possibility of Rho-PE micelles or aggregates participating in α-Syn condensation by imaging unlabelled SLBs (PC:PS 6:4) with α-Syn and still observed condensation (Supplementary Fig. 7). We next investigated whether phase-separated membranes could contribute to the observed phenomena of α-Syn condensation. No condensation was seen on membranes reconstituted from phase-separated GUVs (i.e, PC/SM/Chol of 5:5:0, 4:4:2, 3:3:4). Though no phase-separated domains are microscopically visible on the SLBs, sub-micron scale regions of the lipid membrane representing different order cannot be totally ruled out (Supplementary Fig. 8).

To further probe this membrane-facilitated condensation, we examined the threshold α-Syn concentration required for condensation on the DOPC:DOPS (6:4) surface monitored over ~27 h (Fig. 1j). Surprisingly, the DOPC:DOPS (6:4) membrane appears to facilitate a surface condensation, whereby α-Syn formed condensates at concentrations as low as 10 nM, an order of magnitude below its saturation concentration for bulk LLPS (~200 µM) (Fig. 1j)¹³. No condensation was observed at 5 nM α-Syn (Fig. 1j). Notably, in classical liquid-liquid phase separation, the concentrations of the dilute and droplet phases remain constant even when the total concentration of the system is increased. In contrast, we observed that the intensity of the condensates increased with higher total concentrations, indicating a deviation from typical phase separation behaviour (Fig. 1j–k). These results indicate that surface condensation of α-Syn is highly sensitive to lipid composition. Such surface-mediated condensation is qualitatively distinct from liquid-liquid phase separation, driven by the interplay of protein-protein and protein-membrane interactions. Next, we wondered whether any specific crosslinking between lipids and α-Syn could drive or participate in condensation. To test this, we performed the same experiment in the presence of 50 mM and 100 mM CaCl₂, which is expected to neutralize the negative charges of the PS. Crosslinking would be insensitive to ionic screening by Ca²⁺ and facilitate condensation driven by specific, high-affinity interaction of lipids and α-Syn. As evident from the time-lapse micrograph the presence of Ca²⁺ completely abolished α-Syn condensation (Supplementary Fig. 9a, b). This indicates that the observed phenomena is sensitive to electrostatic screening and not specific binding.

Membrane surface-induced condensation of α-Syn drives transition to fibrillar aggregates

We next captured the dynamics of α-Syn condensate growth on the PC:PS (6:4) membrane surface. Initial membrane binding of α-Syn was observed around 4 h; however, strong and stable binding became evident only after ~18 h (Supplementary Fig. 10a). Condensate formation began after ~23 h and progressively increased in size (Fig. 2a, Supplementary video 1). Widespread droplet growth across the entire field of view was observed, likely due to the accumulation of numerous lipid membrane patches and surface-associated debris over extended imaging durations (Fig. 2a, saturated red channel). Since no α-Syn binding was detected on DOPC-only membranes (Fig. 1c), we conclude that phosphatidylserine (PS) in the membrane plays a crucial role in promoting α-Syn binding and facilitating intermolecular interactions that lead to condensate formation. We further extracted the spatial dynamics over time and show that α-Syn initially forms a thin layer on the membrane surface, followed by the appearance of discrete condensates as demonstrated in kymographs (Fig. 2b). The surface condensation process appears to proceed through homogeneous mixing of α-Syn with PS lipids, followed by the emergence of an outer protein-rich layer (Fig. 2b, merge & Supplementary Fig. 11). Large condensates grew over time through Ostwald ripening, a process where smaller condensates dissolve and their material is transferred to larger ones due to surface tension effects (Fig. 2c & Supplementary video 2)²⁵. Three-dimensional analysis of the condensates revealed an acute contact angle (Supplementary Fig. 11). Strikingly, around 26 h, we observed fibrillar structures emerging from mature α-Syn droplets tubulating the membrane surface (Fig. 2d & Supplementary Fig. 10), providing direct visual evidence of the dynamics of fibril release from membrane-bound condensates. This strongly suggests that surface condensation may serve as a precursor to fibril formation. We conclude that α-Syn condensates formed on the membrane surface exhibit liquid-like properties, including diffusion, coalescence, and fusion – that are hallmarks of dynamic condensates. These structures likely act as intermediate states, wherein α-Syn initially forms liquid-like condensates that later transition into solid-like fibrillar aggregates. This highlights the dynamic nature of α-Syn aggregation and underscores a potential role for membrane-assisted biomolecular condensation as a transient intermediate in the amyloid formation pathway.

Fig. 2. Dynamics of membrane surface-induced α-Syn condensation and fibrillation.

a Confocal time-lapse images show the formation of α-Syn droplets on a PC:PS (6:4) SLB. 100 nM α-Syn tagged with Hilyte-488 was incubated with Rhodamine-PE labelled SLB. (n = 7 independent experiments) (Scale bar, 25 μm.). b Kymograph reveals the temporal changes of α-Syn droplets, capturing their nucleation and progressive growth. c Time-lapse images of a fusion event of droplets after 23 h post-incubation of α-Syn tagged Hilyte-488 with Rhodamine-PE labelled SLB. Kymograph shows the temporal progression of the fusion event. (Scale bar, 5 μm). d Confocal microscopy images show the condensates after 26 h of incubation, and insets show fibril-like structures emanating from the droplets. (n = 7; Scale bar, 20 μm). Relevant source data are provided as a Source Data file.

Maturation of α-Syn condensates involves liquid-to-solid transition through diffusion-limited growth

We next tracked the evolution of α-Syn condensate populations by analysing changes in their size distribution over time. A broad range of condensate areas was observed, spanning from 6 to 65 μm². While the majority of condensates fell within the 5–30 μm² range, a smaller subset expanded to larger areas between 30 and 65 μm² (Fig. 3a). Further analysis revealed that the average condensate radius grew over time following a power-law scaling relationship, specifically with a time exponent of 1/3. This behaviour is characteristic of diffusion-limited Ostwald ripening (Fig. 3b). The corresponding fit also indicates that droplet volume increases linearly with time, shedding light on the dynamics and uniformity of condensate growth. To assess the internal mobility of both lipids and protein within the condensates, we performed fluorescence recovery after photobleaching (FRAP) on Rh-PE-labelled lipids and Hilyte-488-labelled α-Syn at three distinct stages of condensate maturation (i.e., 23 h 30 min, 24 h 30 min, and 25 h 30 min) (Fig. 3c–f). Mature α-Syn condensates exhibited markedly slower fluorescence recovery compared to newly formed ones (Fig. 3d), indicating a progressive reduction in mobility over time. A similar, though less pronounced, trend was observed for lipid mobility (Fig. 3f). Almost ~65% recovery was observed for membranes without bound α-Syn indicating the fluidity of the membrane (Supplementary Fig. 12). Together, these findings suggest that α-Syn condensates undergo a liquid-to-solid transition during maturation. This gradual loss of mobility and increasing rigidity likely represents a key intermediate state in α-Syn aggregation, potentially linking phase separation to the formation of pathogenic fibrillar assemblies.

Fig. 3. α-Syn condensates undergo liquid-like to solid-like transition.

a The histogram shows the area and number of droplets at 24.40-h time points in the presence of 100 nm α-Syn. b Temporal evolution of Droplet Radius (means ± SD, n = 40 droplets). c Confocal microscopy images capture α-Syn aggregates during the FRAP experiment in the α-Syn imaging channel (Scale bar, 5 μm). d FRAP recovery dynamics were analysed to determine their dependence on the maturation stage of α-Syn condensates, as monitored in the α-Syn imaging channel. (means ± SD, n = 3). e Confocal microscopy images also depict the ageing of α-Syn aggregates during the FRAP experiment in the lipid imaging channel (Scale bar, 5 μm). f The FRAP recovery behaviour was also evaluated in relation to the maturation stage of α-Syn condensates, as observed in the lipid imaging channel. (means ± SD, n = 3). Relevant source data are provided as a Source Data file.

α-Syn binding reduces membrane tension and compressibility

We next investigated how the membrane is affected during the stages of α-Syn binding and subsequent fibrillar growth. To probe early molecular interactions (within 1 h), we employed Langmuir monolayer studies using surface pressure–area (π–A) isotherms to assess the thermodynamic and physicochemical changes in model membranes upon α-Syn interaction. The π–A isotherms for PC:PS (6:4) membranes exhibited a rightward shift upon α-Syn incorporation, indicating an increase in molecular area and suggesting lipid expulsion from the monolayer (Fig. 4a). A distinct plateau between 28–32 mN/m was observed, corresponding to a transition from liquid-expanded (LE) to liquid-condensed (LC) states. To quantify membrane elasticity, we calculated the compressibility modulus (Cs⁻¹), which reflects the in-plane rigidity of the monolayer. In the surface pressure range of 25–30 mN/m—relevant to bilayer mechanics—α-Syn binding caused Cs⁻¹ to rise from ~90 mN/m to ~180 mN/m, indicating increased membrane elasticity and reduced compressibility as the membrane entered a more ordered LC phase (Fig. 4b).

Fig. 4. The binding of α-Syn decreases the membrane tension and compressibility modulus.

a Surface pressure (π)-mean molecular area isotherm of the PC:PS (6:4) membrane was measured in the absence (blue) and presence (grey) of α-Syn. b The compressibility modulus (C_s⁻¹)-surface pressure(π) curves for the control membrane model and in the presence of α-Syn are presented in the graph. The data follows the same labelling order and colour scheme as stated in (a). All monolayer experiments were conducted on a PBS subphase (pH 7.4) at 25 °C, with each isotherm representing the mean of three independent replicates. c Representative time-lapse Epi-fluorescence images of a GUV mimicking PC:PS (6:4) composition labelled with 0.1% Rhodamine-PE (red). Aspirated GUVs with stable protrusion length were observed. Upon injection of protein, change in spherical geometry and protrusion length was monitored over time. (n = 3 independent experiments) (Scale bar, 10 μm). d Plot for time vs protrusion length (ΔLP) and the vesicle radius (R_v). Data points are shown for a GUV (n = 3 independent experiments). e α-Syn binding to PC:PS (6:4) membrane intensifies over time. Over 80 h, α-Syn exhibited a gradual yet striking increase in binding to PC:PS (6:4) membranes. While initial interactions were minimal, by 80 h, the binding had surged significantly, highlighting a time-dependent strengthening of membrane association (means ± SD, n = 3 independent experiments). f The addition of α-Syn caused a significant increase in the fluorescence anisotropy of TMA-DPH-labelled PC:PS (6:4) membranes, which implies that α-Syn reduces the membrane fluidity (means ± SD, n = 3). Relevant source data are provided as a Source Data file.

To examine membrane mechanics, we used micropipette aspiration of giant unilamellar vesicles (GUVs). In the absence of protein, the protrusion length (L_p) of aspirated GUVs remained stable for over 10 min, establishing baseline tension (Fig. 4c, Supplementary video 3). However, after injection of 100 nM monomeric α-Syn, a reduction in L_p and expansion of the spherical region of the GUVs were observed, indicating a rapid drop in membrane tension. The binding of α-Syn monomers leads to the insertion of amphipathic helix of α-Syn monomers induces area expansion while imposing a negative curvature stress due to expansion of the bound outer-leaflet²⁶. The observed reduction in membrane tension indicates protein induced compensatory response to curvature stress. Within ~15 s, GUVs relaxed and exited the pipette, undergoing notable morphological rearrangements (Fig. 4c, d, Supplementary video 4). We then explored whether α-Syn’s membrane affinity changes over the course of aggregation. Steady-state fluorescence spectroscopy revealed a significantly stronger binding at later fibrillar stages (K_d = 6.79 μM) compared to early-stage binding (K_d = 12.5 μM) (Fig. 4e), suggesting increased membrane affinity with aggregation progression. Finally, we assessed changes in membrane order using fluorescence anisotropy with the TMA-DPH probe, which reports on interfacial lipid packing. Upon α-Syn binding, the PC:PS (6:4) membrane exhibited increased fluorescence anisotropy, indicating enhanced ordering at the membrane interface. This ordering reduces entropic fluctuations, thereby facilitating stable protein-protein interactions (as evident from slow fluorescence recovery from the vicinity) necessary for the slow nucleation of condensates. In contrast, membranes showed a progressive decrease in anisotropy over time, consistent with increasing disorder in the absence of the protein (Fig. 4f). Together, these findings demonstrate that α-Syn binding and fibrillation actively remodel membrane structure and mechanics that in turn provide a conducive environment for α-Syn condensation.

Interfacial potential drives lipid stoichiometry-dependent surface condensation of α-Syn

The interfacial potential of the cell membrane-arising from lipid composition and ion adsorption - is a critical factor essential for both intra- and intercellular interactions. We hypothesized that the lipid stoichiometry-specific binding and condensation of α-Synuclein (α-Syn) at nanomolar concentrations is governed by this interfacial potential. To test this, we measured the zeta potential of various membrane surfaces with differing phosphatidylcholine (PC) and phosphatidylserine (PS) ratios. Zeta potential, which reflects the electric potential at the shear plane (~1–10 nm from the surface), serves as an effective proxy for interfacial potential and is determined by surface charge and the structure of the electric double layer²⁷. Our measurements revealed that α-Syn binding occurred on membranes with zeta potentials in the range of −12 mV to −20 mV, with condensation observed only at ~−20 mV (Fig. 5a), suggesting a threshold interfacial potential is necessary to drive surface condensation. While we use zeta potential as a practical indicator of the interfacial electrical environment, it is important to note its distinction from the true interfacial potential (ψ₀). Gouy-Chapman theory suggests that ψ₀ (membrane potential) is directly related to σ (surface charge density); thus, a higher negative surface charge density (σ) should result in more negative ψ₀.

$$\mathrm{Zeta\; potential},\mathrm{\zeta }=\mathrm{\psi }0{\mathrm{e}}^{-\mathrm{\kappa }\mathrm{d}}$$ 1

where, d = distance from the membrane surface to the shear plane (~0.2–0.5 nm for lipids), κ is the ionic strength of the buffer. For the chosen experimental conditions involving supported bilayers, both the shear plane position (d) and ionic strength of the environment (k) are fixed. Thus, we expect the trends in ζ to reliably reflect the trends in ψ₀.

Fig. 5. Interfacial potential drives surface condensation of α-Syn.

a Changes in zeta potential were evaluated across membranes containing neutral and negatively charged compositions (means ± SD, n = 3 independent experiments). b Model schematic representing the interaction of α-Syn with the membrane. The membrane is composed of two kinds of lipids (DOPC and DOPS) and is represented via a 2D Ising model (lipid-lipid interaction J_m). α-synuclein proteins are represented using a 3D lattice-gas model (protein-protein interaction J_p). The proteins interact with the membrane via tethers (protein-tether interaction J_t). c Here, we show the excess density, which quantifies the degree of surface condensation, as a function of the bulk chemical potential. We see a sudden jump in the excess density, which is characteristic of a prewetting transition. Bulk phase separation occurs at μ = −3.6. (Figure parameters: J_p = 1.2, J_m = 0.5, J_t = 1.5, L_t = 5, ϕ_m = 0.5, ϕ_t = 0.2). d Excess density for various tether lengths. Surface condensation occurs only above a certain threshold tether length (L_t = 3 in this case), emphasizing the role of the range of protein-membrane interaction. (Figure parameters: J_p = 1.5, J_m = 0.5, J_t = 1.5, μ = −4.6, ϕ_m = 0.5, ϕ_t = 0.2). e Simulation snapshots for various tether lengths, L_t = 2,4,6,10 (all other parameters same as in (d)). f Here, we show the excess density as a function of the protein-tether interaction strength. Similar to the tether length, there is a threshold interaction strength below which surface condensation is negligible. (Figure parameters: J_p = 1.2, J_m = 0.5, L_t = 5, μ = −4.3, ϕ_m = 0.5, ϕ_t = 0.2). Relevant source data are provided as a Source Data file.

To further investigate the role of interfacial potential and emergence of surface condensates, we employed a lattice-gas model, adapted from Rouches et al.²⁸ in which proteins interact with a membrane represented as a 2D Ising model tethered to a 3D lattice (Fig. 5b; see Methods). The model membrane consists of DOPC and DOPS, where α-Syn, with its positively charged N-terminal region, preferentially associates with the negatively charged PS lipids. In this framework, lipid phase separation occurs when the interaction strength between like lipids (J_m) exceeds a critical threshold (J_cm). Similarly, protein condensation into bulk liquid-like phases requires that the protein-protein interaction strength (J_p) exceeds a critical value (J_cp), provided the protein concentration surpasses the saturation limit. Importantly, even when bulk phase separation is not favoured, membrane interactions can induce surface condensation via a pre-wetting transition, characterized by a sharp increase in protein surface density indicating the formation of a thick protein layer on the membrane (Fig. 5c). The model also predicts that increasing the protein concentration or the tether length representing the spatial range of membrane interaction—enhances surface condensation (Fig. 5c, d). Notably, below a critical tether length, surface condensation does not occur (Fig. 5e), emphasizing the role of membrane proximity and coupling strength. Additionally, a threshold interaction strength between the protein and the membrane tether is required; below this value, condensation is absent (Fig. 5f). Together, these findings underscore that membrane interfacial potential is a key driver of α-Syn condensation at concentrations below bulk saturation concentrations.

Perturbing membrane interfacial potential via lipids in hippocampal neuronal cells alters preformed α-Syn condensate size

Membrane interfacial potential can be modulated by altering the turnover of charged lipids at the cell surface²⁹. To estimate the PS-induced changes in membrane interfacial potential we used voltage-sensitive probes, such as ANEPPS (4-[2-[6-(dioctylamino)-2-naphthalenyl]ethenyl]-1-(3-sulfopropyl)-pyridinium). Ratio of ANEPPS fluorescence intensity at 530 nm/440 nm (R) is related to voltage change (ΔV_m) as follows:

$$\mathrm{R}={\mathrm{R}}_{\mathrm{o}}+\mathrm{S}{\Delta V}_{\mathrm{m}}$$ 2

where, Ro is the ratio at the resting membrane potential (or control condition), S is the sensitivity factor of the dye. Thus, any change in the ratio R reflects corresponding change in the electric field strength that reasonably captures interfacial potential. Indeed ~two-fold increase in R the was observed suggesting PS treatment induces changes in local electric field strength reflecting altered potential (Fig. 6a, b). To test whether such perturbations influence the morphology of pre-formed α-Syn (α-Syn) puncta, we investigated the effect of modulating phosphatidylserine (PS) mediated interfacial potential changes on α-Syn aggregation in doxycycline-inducible HT22 mouse hippocampal neuronal cells. Cells were treated with liposomes composed of PS at increasing concentrations (70 μM, 100 μM, and 150 μM) for 24 h (Fig. 6c). Immunofluorescence imaging using an anti-Myc antibody to detect α-Syn resulted in increase in the puncta size followed by a decrease observed at 150 μM PS (Fig. 6d). The observed reduction in the puncta size can be attributed to the destabilization of the preformed puncta as a result of depolarization. This also suggests that interfacial potential changes as a result of local lipid alteration can modulate membrane-protein interactions and perturb surface-associated α-Syn condensation. These findings suggest that perturbing interfacial potential - specifically, through external PS treatment - can significantly alter α-Syn aggregation dynamics by modulating the membrane’s biophysical landscape.

Fig. 6. Lipid modulation alters interfacial potential modulate α-Syn puncta in hippocampal HT22 cells.

a Representative DI-8-ANEPPS fluorescence images showing membrane potential changes in untreated and 150 μM PS-treated cells. (Scale bar, 5 μm.). b Quantification of membrane potential using Di8-ANEPPS reveals that 150 μM PS-treated cells alter membrane potential in primary neurons (mean ± SEM; n = 3; ns (non-significant) P > 0.05, *P < 0.05, ***P < 0.001; two-tailed, multiple t-tests). P values for PS treated was <0.0001. c HT22 cells expressing α-Syn under doxycycline induction were treated with increasing doses of Phosphatidyl Serine (PS) (70, 100 and 150 μM) for 24 h. Immunofluorescence images, using Myc-tagged antibodies to visualise α-Syn, are shown. (Scale bar, 5 μm.). d Quantification of α-Syn puncta size after 24 h of PS treatment (mean ± SD, n = 6; ns (non-significant) P > 0.05, *P < 0.05, ***P < 0.001; two-tailed, multiple t-tests). P values for 70 μM, 100 μM and 150 μM treated were 0.0113, 0.2882, and <0.0001, respectively. A statistically significant reduction in puncta size was observed at 150 μM. Relevant source data are provided as a Source Data file.

Membrane depolarization drives α-Syn condensate formation and extracellular secretion in primary cortical neurons

Membrane interfacial potential can also be modulated by altering ionic activity across the plasma membrane. To examine how such changes affect α-Syn (α-Syn) condensate formation, we expressed GFP-tagged α-Syn in rat primary cortical neurons and experimentally altered their membrane potential. Primary neurons are ideal for this purpose, as they undergo dynamic membrane potential changes during synaptic activity. Previous studies have linked neuronal depolarization with increased extracellular secretion of α-Syn, although the mechanisms remain unclear^30–32. To investigate whether membrane depolarization modulates α-Syn interaction with neuronal membranes, we treated neurons with depolarizing agents—25 mM and 50 mM KCl, as well as glutamate. Upon depolarization, α-Syn formed punctate structures within neurons (Fig. 7a). Quantification revealed a ~4-5 fold increase in the number of intracellular α-Syn puncta in depolarized neurons compared to controls at resting potential (Fig. 7b). Interestingly, while 25 mM KCl stimulation significantly increased intracellular puncta, no further increase was observed at 50 mM KCl. However, extracellular α-Syn puncta increased in both depolarization conditions and positively correlated with the extent of depolarization (Fig. 7c and Supplementary Fig. 13). This suggests that stronger depolarization may promote the growth and release of α-Syn condensates, possibly by enhancing their membrane interaction and subsequent secretion (Fig. 7a–c). Depolarization was confirmed through calcium imaging in the same neuron-astrocyte co-cultures used for α-Syn puncta analysis (Supplementary Fig. 14; Supplementary Video 5–7). Further, using a potentiometric dye, di-8-ANEPPS, we corroborated the change in electric field strength reflecting the depolarization of neuronal membrane potential upon treatment with KCl (Fig. 7d, e). No α-Syn puncta was observed upon treatment with 10 mM MgCl₂, 100 μM glutamine (Glutamate derivative that does not act as a neurotransmitter), 100 μM GABA (an inhibitory neurotransmitter known to hyperpolarize neuronal membranes) suggesting that condensation of α-Syn was not influenced by the presence of salt or osmolytes and was specifically because of membrane depolarization (Fig. 7b, c, Supplementary Fig. 15). To confirm that the observed α-Synuclein (α-Syn) puncta were not an artifact of protein overexpression, we performed immunostaining for endogenous α-Syn. Under depolarizing conditions (25 mM, 50 mM KCl), we identified similar punctate structures, confirming their formation in a physiologically relevant context. These endogenous α-Syn puncta were distributed throughout all neuronal compartments visualised by dual staining with MAP2, a marker of somatodendritic compartment and beta III tubulin, a marker of the whole neuron (Fig. 7f). Notably, a substantial proportion of the α-Syn puncta was consistently localized within the axon under all conditions tested and manifested similar increase upon depolarization (Fig. 7f–h). Additionally, the endogenous α-Syn puncta demonstrated significant colocalization with the presynaptic marker Synapsin1, exhibiting a similar increase in degree of colocalization upon depolarization with 25 mM KCl in line with the overexpression data (Fig. 7i, j, Supplementary 16). Taken together, these findings suggest that neuronal membrane depolarization—common in neurodegenerative conditions like Parkinson’s disease and dementia with Lewy bodies—can promote α-Syn condensation and facilitate its membrane association and extracellular release, potentially contributing to its pathological spread.

Fig. 7. Membrane depolarisation triggers α-synuclein condensation and secretion from primary cortical neurons.

a Representative confocal images of α-Syn puncta distribution in neurons under resting conditions or depolarised by glutamate/glycine (10:1), 25 mM or 50 mM KCl. Insets show increased puncta upon depolarisation. White and blue arrows show the intracellular and extracellular puncta, respectively. b Quantification of intracellular puncta: significant increase with glutamate, 25 mM, and 50 mM KCl, but not with glutamine, GABA, or MgCl₂ (mean ± SEM; n = 3; ns P > 0.05, *P < 0.05, ***P < 0.001; two tailed, multiple t-tests). P values for Gly+Glu, KCl 25 mM, KCl 50 mM, GABA, Glutamate, MgCl₂ were <0.0001, <0.0001, <0.0001, 0.4425, 0.5731, 0.3080, respectively. c Extracellular puncta quantification: increase only with 50 mM KCl (mean ± SEM; n = 3; ns P > 0.05, *P < 0.05; two tailed, multiple t-tests). P values for Gly+Glu, KCl 25 mM, KCl 50 mM, GABA, Glutamate, MgCl₂ were 0.0829, 0.0699, 0.0006, 0.0568, 0.0975, 0.0568, respectively. d Representative DI-8-ANEPPS fluorescence images showing membrane potential changes with 25 mM KCl (scale bar, 2 μm). e Quantification of membrane potential by DI-8-ANEPPS confirms depolarization by 25 mM KCl (mean ± SEM; n = 3; *P < 0.05; two tailed, Student’s t test). P values for 25 mM KCl 0.002. f Endogenous α-Syn puncta (white: intracellular; blue: extracellular) increase upon KCl treatment. g Total α-Syn puncta per neuron increase with glutamate, 25 mM, and 50 mM KCl (mean ± SEM; n = 3; *P < 0.05, ***P < 0.001; two tailed, multiple t-tests). P values for 25 mM KCl and 50 mM KCl were 0.0011 and <0.0001, respectively. h Axonal α-Syn puncta increase with 25 mM and 50 mM KCl (mean ± SEM; n = 3; *P < 0.05, ***P < 0.001; two tailed, multiple t-tests). P values for 25 mM KCl and 50 mM KCl were <0.0001 and 0.0008, respectively. i Representative images showing that endogenous α-Syn colocalizes with Synapsin-1, enhanced by 25 mM KCl. j Manders coefficient quantification shows increased colocalization with Synapsin-1 upon 25 mM KCl treatment (mean ± SEM; n = 3; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001; one-way ANOVA with Sidak’s test). Relevant source data are provided as a Source Data file.

Discussion

Aggregation of α-Syn is implicated in neurodegeneration; however, the dynamics of the early events that govern this process remain unclear. The lipidome of a human brain is spatiotemporally highly heterogeneous³³. Elevated levels of Phosphatidylserines are reported in neuronal membranes in Synucleopathies³⁴. This prompted us to investigate the role of altering lipid environment on the α-Syn aggregation. α-Syn is known to form liquid-like droplets at high critical concentrations (~200 μM) in the presence of molecular crowders and low pH¹³. In sharp contrast, here we discover that a lipid membrane surface with a defined interfacial potential is sufficient to drive α-Syn condensation even at sub-critical concentrations seen endogenously in primary neurons (10–100 nM), eventually leading to fibrillation. We demonstrate that indeed the stoichiometry of the lipid components of the membrane plays an important role in the binding of α-Syn (Fig. 1). Condensation of α-Syn is only induced in the presence of a membrane surface with specific stoichiometry of PC:PS (6:4) but not free suspended lipids of the same compositional stoichiometry (Fig.1). These observations are consistent with a prewetting-like transition driven by favourable interactions of α-Syn with negatively charged membrane surface. Upon reaching a critical concentration on the membrane surface, α-Syn undergoes a transition from a thin adsorbed layer to a thick condensed layer forming macroscopic gel-like droplets that grow over time (Figs. 2 and 3). Indeed, it has been shown that changes in the conformational dynamics of disordered regions of intrinsically disordered protein such as α-Syn result in the formation of gel-like structures^7,35,36.

Various intracellular surfaces have been recognized for their role in promoting biomolecular condensation and potentially regulating this process^19,37. Originally predicted by Cahn, prewetting and wetting transition plays a crucial role in biomolecular condensation on cellular surfaces, including membranes^21,38,39, microtubules⁴⁰, actin^41,42, and DNA⁴³. Another recent study demonstrated that N Wasp undergoes prewetting transition to form surface condensates⁴⁴ on lipid membranes. One intriguing aspect of the prewetting transition is that α-Syn can undergo phase separation at sub-critical concentrations of ~10 nM near a membrane surface. In contrast, bulk phase separation of α-Syn requires a much higher concentration of ~200–500 μM, which far exceeds the physiological levels observed across different biological systems.

By reducing membrane tension and compressibility, α-Syn binding facilitates condensate formation on the membrane surface, governed by a narrow range of interfacial potential regulated by PS (Figs. 4 and 5). PS is indeed one of the major functionally critical lipids in cell membranes and known to regulate membrane surface charge and protein colocalization^29,45. Interestingly, altering cellular PS levels by providing PS liposomes externally perturbed the size of the α-Syn puncta in the cells (Fig. 6). Spatio-temporal differences and modulation of PS levels in the membrane are likely to affect the interfacial potential that can have broad impact on transmembrane potential and membrane protein function including ion channel activity²⁹. Interestingly, neuronal hyperactivity leads to translocation of α-Syn from intracellular to the extracellular side; however, the trigger remains unknown^31,46–48 (Fig. 7). We propose that both lipid composition and electrical activity across neuronal membranes modulate interfacial potential, thereby promoting α-Syn binding, and subsequent fibril formation (Fig. 8). Such condensation has also been reported to spatially coordinate microtubule nucleation and branching^49,50. Very recently electrochemical potential of biomolecular condensates is shown to regulate their physicochemical activities⁵¹. Electrochemical properties can make the condensates chemically active^52–55 and facilitate numerous cellular functions such as translation, stress response and cell-to-cell communication^56–58. Condensates that function as the nucleation centres for fibrils would facilitate deformation owing to the higher affinity for membrane surface and thus help propagation across the membrane into the extracellular side. However, the exocytic trafficking of α-Syn puncta cannot be ruled out and thus mechanism of extracellular release would require further investigation. Taken together, our results offer fundamental insights into how membrane surface and interfacial potential drive α-Syn condensation in the sub-critical nanomolar concentration range, followed by fibril-mediated membrane deformation - a process crucial to the progression of neurodegeneration.

Fig. 8. A schematic of the proposed model illustrating membrane surface-driven condensation of α-Syn and subsequent fibrillation.

a Changes in membrane interfacial potential, visualized here as a heat map, regulate the surface condensation of α-Synuclein within a narrow range. b Surface condensation of α-Synuclein begins with membrane prewetting, followed by droplet growth. These condensates serve as a hub for the subsequent release of fibrils. c Surface mediated condensation could be potentially one of the factors driving the extracellular release.

Methods

Ethical statement

All animal experiments were conducted in accordance with the guidelines of the Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA) (Registration No. 1634/GO/ReRcBiBt/S/12/CPCSEA; Date of Registration: 16.05.2023). The experimental protocols were approved by the Institutional Animal Ethics Committee (IAEC) of the National Institute of Science Education and Research (NISER), Bhubaneswar, India, and all procedures were performed in compliance with institutional and national ethical guidelines (Ethical Approval No. NISER/SBS/AH-322).

Animals

Sprague-Dawley rat pups were used for all experiments. Pups of either sex (male and female) at postnatal day 0 or 1 (P0–P1) were included. A total of 9–10 pups were used per experiment. Sex was not considered as a biological variable in this study, as experiments were performed exclusively using neonatal (P0–P1) Sprague-Dawley rat pups.

Materials

1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2 dioleoyl- sn-glycero-3-phosphoethanolamine (DOPE), L-α-phosphatidylinositol (liver PI), 1,2-dioleoyl-sn-glycero-3-phospho-L-serine (DOPS), Porcine Total Brain Extract (Avanti Polar Lipids),1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N- (lissamine rhodamine B sulfonyl) (Rhod PE), and cholesterol were purchased from (Avanti Polar Lipids). Rhod PE is known to partition into liquid-disordered phases preferentially and is thus used to visualise the same⁵⁹. Avidin-coated chamber slides were purchased from Sigma-Aldrich.

Protein expression and purification

α-Synuclein was expressed and purified as described earlier⁶⁰. Briefly, α-Syn was first expressed in E. coli using plasmid pT7-7 in BL21(DE-3)-competent cells in LB medium at 37 °C until O.D. reached 0.6–0.8. They were supplemented with ampicillin to maintain plasmid selection. IPTG induction-initiated protein expression, followed by a four-hour incubation at 37 °C. Cells were harvested by centrifugation and resuspended in a lysis buffer containing (50 μM Tris-HCl, 10 μM EDTA, 150 μM NaCl and PMSF, pH 7.4). Cell disruption was achieved through freeze-thaw cycles and sonication for around two hours. After boiling and centrifugation, the supernatant was treated with streptomycin sulfate (136 μL of 10% solution/mL supernatant) and glacial acetic acid (228 μL/mL supernatant) was added to remove nucleic acids. Followed by 15,000 × g for 10 min at 4 °C, Ammonium sulfate (up to 50% saturation, as calculated using the online Ammonium sulphate Calculator from Encor Biotechnology Inc.) precipitation was achieved. The precipitated protein was collected using centrifugation at 15,000 × g for 15 min at 4 °C and then washed with 1 mL of 50% ammonium sulfate solution. The resulting pellet was then resuspended in 900 μL of 100 mM ammonium acetate, forming a cloudy solution followed by an equal volume of ethanol at room temperature. Repetition of ethanol precipitation followed by dialysis at 4 °C in dialysis buffer. After the dialysis, the protein was purified using ion-exchange chromatography; the protein was loaded onto a HiTrap Q FF anion exchange column (GE Healthcare, Uppsala, Sweden). The α-Synuclein was eluted at ~300 mM NaCl with a salt gradient from 0 mM to 1000 mM NaCl. The purification was verified by SDS-PAGE electrophoresis. Protein concentration was determined spectrophotometrically at 275 nm using a calculated extinction coefficient of 5600 M⁻¹ cm⁻¹.

Thioflavin T assay for the measurement of fibrillation kinetics of α-Synuclein

To monitor the aggregation, an assay of the different α-Syn concentrations was used in phosphate buffer (pH 7.4) incubated separately with 20 μM ThT working concentration to monitor the aggregation. The lipid specificity of α-Syn was screened by incubating the protein with LUVs of different lipids for amyloid aggregation. The fluorescence measurements were performed on a Clariostar microplate reader on a Nunc black plate (Thermo Fisher Scientific). The fluorescence emission spectra were recorded on 448/482 nm excitation/emission filters. The recordings in triplicate were recorded every 15 min for 120 h. To accelerate aggregation and improve reproducibility, we added glass beads of homogeneous size and spherical shape, which were selected by visual inspection^61,62. Glass beads (#LA715A, Himedia) were dipped in MQ water for 10 min and then dried, and again re-dipped in 100% IPA, then the beads were dried and added to the wells for the experiment.

Preparation of large unilamellar vesicles

A 1 mM lipid stock was prepared and dried under nitrogen gas for each membrane condition. Then, it was vacuumed for an hour to remove any remaining residual solvent. The lipid film was then rehydrated with 1 mL of PBS buffer. Following a 15-min water bath treatment, it was vortexed for 5 min. To form Large Unilamellar vesicles (LUVs), the lipid suspension was extruded using an Avanti polar extruder. Dynamic light scattering (DLS) analysis confirmed that the average size of LUV was around 130 nm.

Preparation of giant unilamellar vesicles

Giant Unilamellar Vesicles (GUVs) composed of 60% DOPC, 40% DOPS and 0.1% Rhodamine-PE were generated using the electroformation method as described in refs. ^63,64. To optimize GUV yield, 15 µL of a 5 mg/mL lipid mixture was uniformly spread onto indium tin oxide (ITO)-coated conductive glass slides (Nanion Technologies, GmbH) and dried under vacuum for at least 2 h. The dried lipid film was then rehydrated in PBS buffer (pH 7.4, 300 ± 5 mOsm), and electroformation was performed by applying a 2 V, 10 Hz sine wave for 2 h at 60 °C.

Preparation of supported lipid bilayers (SLBs)

The GUVs, diluted in buffer, were burst onto a plasma-cleaned coverslip at the base of a flow chamber (ibidi sticky-Slide VI 0.4) to form the supported lipid bilayers. Following that, casein (1 mg/ml) passivation was performed for 20 min, and the sample was washed with phosphate-buffered saline before the experiment.

Preparation of free Rhodamine-PE

To form free Rhod-PE mixtures, 0.1% Rhodamine-PE without membrane composition was dried under vacuum for at least 2 h. The dried lipid film was then rehydrated in PBS buffer (pH 7.4, 300 ± 5 mOsm). The mixture was then sonicated for 10 min at 50% amplitude with a 10 s on/off cycle.

Confocal fluorescence microscopy

An 8-well chamber slide from Ibidi was used for incubating the SLBs and α-Syn labelled with Alexa-488. For GUV imaging, the slide was coated with 10 μL of 1 mg/mL streptavidin, followed by 200 μL of biotinylated GUVs, which were incubated for 30 min to allow immobilization. Then, the 5 μM Alexa-488 α-Syn was added to the GUV solution. The chamber was sealed with the Opti-Seal to prevent evaporation during the long imaging and incubation time. Imaging was conducted on a Leica TCS SP8 using the appropriate lasers for Rhodamine-PE (561 nm) and Alexa-488(488 nm). Identical laser power and gain settings were used during all the experiments. Image processing was done using Fiji.

Fluorescence recovery after photobleaching

Fluorescence recovery after photobleaching (FRAP) measurements were performed on supported lipid bilayers (SLBs) doped with 0.1% rhodamine-labelled phosphatidylethanolamine(rhodamine-PE) and incubated with 100 Nm α-Syn doped with 10% Hilyte-488 α-Syn. Pre-bleach images were acquired with a low-intensity laser to establish the baseline. A specific region of interest (ROI) with a radius of 5 μm was photobleached using full laser intensity for 30 s, followed every few seconds for 2–3 min to monitor the fluorescence recovery. The experiment was repeated three times for each condition, and the resulting recovery curves were normalized to account for the variations in initial fluorescence intensity. The normalized curves were used to determine the half-recovery time (${\tau }_{\frac{1}{2}}$), which is the recovery half-time. The diffusion coefficient (D_r) was calculated using the Soumpasis equation for 2D diffusion:

$${\mathrm{D}}_{\mathrm{r}}=0.224\frac{{\mathrm{r}}^2}{{\mathrm{\tau }}_{\frac{1}{2}}}$$ 3

Where r is the radius of the bleached ROI (5 μm), and 0.224 is a numerically determined constant.

Fluorescence spectrophotometric assay

The fluorescence spectroscopy experiment investigated the impact of α-Syn on lipid membrane fluidity. The Fluorescence spectroscopy experiments were conducted on previously prepared LUVs. TMA-DPH was dissolved in DMSO to reach the final concentration of 2 mM. The LUVs were then incubated with 5 μM and 500 μM TMA-DPH, bringing the total volume of the LUV mixture, α-Syn and TMA-DPH to 1 mL. Followed by 2-h dark incubation, the fluorescence anisotropy was measured every 4 h till 72 h using FLS 1000 (Edinberg Instruments, UK) with excitation at 360 nm and emission at 430 nm. All the fluorescence spectra were recorded using a quartz cuvette with a fixed 1 cm path length. Anisotropy(r) was calculated automatically by the instrument using the equation:

$$r=\frac{I_{VV}-{GI}_{VH}}{I_{VV}+{2GI}_{VH}}$$ 4

where $I_{VV}$ and $I_{VH}$ are the fluorescence intensities of the vertical and horizontal components, respectively. The grating correction factor G ( = $\frac{{\mathrm{I}}_{\mathrm{HV}}}{{\mathrm{I}}_{\mathrm{HH}}}$) corrects for any wavelength-dependent polarizer distortion. All the experiments were repeated multiple times to ensure reproducibility and statistical significance of the results.

Langmuir lipid monolayer experiments

Lipid monolayers were conducted using a KSV NIMA Langmuir balance equipped with two barriers for compression and a Wilhelmy microbalance with filter paper as a surface pressure sensor. The setup was enclosed in a transparent glove box to maintain a controlled environment. Before each experiment, the trough was meticulously cleaned with methanol, ethanol, and ultrapure water to eliminate surface impurities. Monolayers composed of DOPC:DOPS (6:4) were prepared by spreading a lipid/chloroform solution (1 mg/ml) dropwise, followed by a 15-mins equilibration period to allow for chloroform evaporation. The subphase consisted of phosphate-buffered saline (PBS) maintained at a constant temperature of 25 °C. The monolayer was left undisturbed for 15 min to relax the monolayer to 0 mN/m. Subsequently, a final working concentration of 100 nM α-Syn was injected into the monolayer. The solution was gently stirred using a magnetic stirrer to ensure distribution for isotherm assays. The compression was applied at a uniform 1 mm/min speed until the collapse pressure(π_c) was reached. The isotherm data were further analysed to calculate the compressibility modulus ($C_s^{-1}$), which is defined as

$$C_s^{-1}=-A\frac{\mathrm{d}\mathrm{\pi }}{dA}$$ 5

Where A is the molecular area, and $\frac{\mathrm{d}\mathrm{\pi }}{\mathrm{dA}}$ represents the derivative of surface pressure concerning the molecular area.

Micropipette aspiration

All the measurements were conducted using an Olympus IX83 epifluorescence microscope equipped with Hoffman modulation optics and a 100× objective lens for high-resolution imaging. Three-axis hydraulic micromanipulators were employed to position the micropipette accurately. The micropipettes were pulled from 10 μm diameter glass capillaries using a Sutter Instruments P-87 puller and a Narishige Microscope MF-900. 100 μL of GUVs was added to a custom-made open coverslip chamber. The desired suction pressure was applied to aspirate a GUV through the micropipette tip, forming an inner protrusion. Video recordings were taken over time to analyse the changes in vesicle geometry upon the addition of α-Syn.

Model for surface condensation of α-synuclein on a lipid bilayer membrane

Following the study by Rouches et al.²⁸, we model the α-synuclein proteins as particles on a square lattice of size LxLxL. The state at each site on the lattice is represented by 0 if it is empty or 1 if a protein molecule is present, denoted as n_i = 0,1. The x- and y-directions have periodic boundaries. In the z-direction the membrane is present at z = L, and is modelled as a two-dimensional Ising system. The two states s_i = ±1 represent the two kinds of lipids, DOPC and DOPS. α-Syn, with its abundance of positively charged residues, has a greater affinity towards PS, which is negatively charged. This interaction with the membrane is introduced via tethers, which sit on ${\varphi }_t$ fraction of the s = +1 spins and extend L_t sites into the bulk. The total Hamiltonian for the system is given as

$$H=-J_m\sum _{\begin{array}{c}\hfill ⟨i,j⟩\hfill \\ \hfill \in mem\hfill \end{array}}{s}_i{s}_j-J_p\sum _{\begin{array}{c}\hfill ⟨i,j⟩\hfill \\ \hfill \in bulk\hfill \end{array}}{n}_i{n}_j-J_t\sum _{i\in tether}{n}_i-\mu \sum _{i\in bulk}{n}_i$$ 6

Here the first term sums over all the nearest-neighbour pairs on the membrane, representing the interaction between lipids. The second term sums over all the nearest-neighbour pairs in the bulk, accounting for protein-protein interaction via the coupling parameter J_p. The proteins and tethers interact via the coupling constant J_t, which acts when a protein and tether occupy the same site. This is represented in the third term. Finally, the bulk chemical potential μ controls the overall concentration of proteins in the system. The chemical potential and all interaction strengths are in units of k_BT. The calculated values for the measured parameters are presented in Table 1.

Table 1.

Mathematical simulation parameters

Parameter used	Range/Value
Lattice size, L	20
Tether length, Lt	0–10
Tether density, ${\varphi }_t$	0.005–0.6
Fraction of up spins, ${\varphi }_m$	0.1–0.8
Protein-protein interaction strength, Jb /kBT	0.6–2.0
Lipid-lipid interaction strength, Jm /kBT	0.3–0.6
Tether-protein interaction strength, Jt /kBT	0.0–3.0
Bulk chemical potential, -μ /kBT	1.5–6.5

We use Monte Carlo simulation following the Metropolis algorithm to allow the system to equilibrate. The system is evolved to equilibrium for 10⁴–10⁵ Monte Carlo sweeps, and after equilibration, the final 5000 sweeps are used for data collection.

Each sweep of the system consists of the following moves:

Membrane lipids are simulated using conserved order-parameter (Kawasaki) dynamics. Lipids with tethers attached to it are not considered for this exchange. The tethers are randomly displaced to neighbouring sites containing s = +1 spins.

Proteins in the bulk can undergo displacement. Additionally, proteins can be added or removed, as we simulate the system in a grand-canonical ensemble.

The degree of surface condensation of the protein on the membrane is quantified by the excess density, defined⁶⁵ as

$$C_s=\sum _{n=1}^N\left({\varphi }_n-{\varphi }_b\right)$$ 7

Where ${\varphi }_b$ is the bulk volume fraction far away from the surface and can be calculated as.

$${\varphi }_b=\frac{1}{M}\sum _{i=1}^M{\varphi }_i$$ 8

Simulation parameters

Generation of stable cell line

Mouse hippocampal neuronal HT22 cells were transfected with 1 µg/mL of pINDUCER20-α-Syn plasmid DNA (Addgene) using Metafectene Pro (Biontex, lot no. RKP205/RK081621) following the manufacturer’s protocol. After 48 h, the cells were transferred to 100 mm culture dishes and subjected to antibiotic selection using 7.5 mg/mL G418 (Geneticin, G-418 sulfate; GOLDBIO, G-418-25) containing DMEM medium. Selection was continued until non-transfected cells were dead, which was for 14 days in our case.

Following selection, 24 single-cell-derived colonies were isolated and seeded into 48-well plates. Upon reaching ~80% confluency, cells were sequentially passage into 24-well, 12-well, and finally 6-well plates. Each colony was then screened for α-Syn expression via immunofluorescence using an anti-myc tag antibody, as the α-Syn construct included a myc tag. Colonies testing positive for myc-tagged α-Syn expression were selected and cryopreserved at –80 °C.

Mammalian cell culture and treatment with monomeric α-Syn

Mouse Hippocampal Neuronal HT22 stable cells expressing α-Syn were grown in DMEM Glutamax, supplemented with 10% FBS, 1% antibiotic-antimycotic, along with 0.75 mg/ml of G418 at 37 °C, 5% CO₂ in a humidified incubator. Early Passage cells (1–10) were used for all the experiments. Mouse HT22 doxycycline inducible stable cells were induced with 1 µg/ml of Doxycycline, 24 h after plating. Media was replaced with fresh media containing 1 µg/ml of Doxycycline to which 70 µM, 100 µM and 150 µM of exogenous Phosphatidyl Serine (DOPS), were added respectively for each condition. After 24 h of induction and treatment with 70 µM, 100 µM and 150 µM the cells were harvested and experiments such as Immunofluorescence Imaging were performed.

Immunofluorescence imaging

2*10⁴ cells were seeded in each 15 mm coverslip, and the cells were treated with exogenous Phosphatidyl Serine (DOPS) by the above-discussed protocol. After treatment, cells were gently washed with 1× PBS thrice and then fixed using 4% Paraformaldehyde (PFA) for 20 min at room temperature. Following fixation, cells were washed gently with 1× PBS thrice, and then the cells were permeabilized with 0.1% x Triton X-100 for 10 min and blocked using 1% BSA for 45 min. The primary antibodies, rabbit MAP2 (1:150, Cell Signalling Technology, #8707S), Guinea pig Synapsin1 (1:300, Synaptic Systems, #106104), chicken βIII-tubulin (1:200, Abcam, #AB41489), mouse α-synuclein (1:100, Invitrogen, #328100) and Rabbit anti-Myc of dilution (1:1000, Invitrogen, #PA1-981) were diluted in in the blocking solution (1% BSA) and incubated overnight at 4 °C. The next day, cells were washed once with PBS and twice with PBS. Secondary antibodies, anti-rabbit Alexa Fluor 568 (1:1000, Invitrogen, #A11011), anti-rabbit Alexa Fluor 488 (1:1000, Cell Signalling Technology, #4412S), anti-mouse Alexa Fluor 555 (1:1000, Cell Signalling Technology, #4409S), anti-Guinea pig Alexa Fluor 647 (1:1000, Invitrogen, #A21450) and anti-chicken 647 (1:1000, Invitrogen, #A21449) were added in the blocking solution (1% BSA). Alexa Fluor 594-conjugated goat Anti-Rabbit (1:1000, Invitrogen, #A-11012) was used, and cells were incubated for 45 min in a humidified chamber in the dark. Coverslips were mounted using DAPI-containing mounting media, and images were captured using 100× oil objectives of the Olympus IX83 Confocal Microscope.

Primary neuron-astrocyte co-culture

Cortical neurons and astrocytes were cultured from P0-P1 Sprague-Dawley rat pups. The brains were removed and placed in ice-cold calcium- and magnesium-free Hanks’ Balanced Salt Solution (HBSS) containing 10 mM glucose and HEPES (Sigma). The cortical tissue was separated and digested with 0.25% trypsin-EDTA (Gibco) and 150 units/ml DNAse (Sigma) at 37 °C for 15 min. After digestion, trypsin activity was stopped with 10% fetal bovine serum (FBS) (Gibco). The cells were dissociated mechanically and then centrifuged at 1000 rpm for 5 min at 4 °C. The resulting cell pellet was resuspended in culture medium composed of neurobasal-A (Gibco), 10% FBS (Gibco), 1% Glutamax (Gibco), 1% Anti-anti (Gibco), and 2% N2 supplement (Gibco). Cells were plated onto coverslips pre-coated with 0.1 mg/ml Poly-D-lysine. The cultures were maintained at 37 °C in a 5% CO₂ incubator with controlled humidity. The culture medium was replaced with fresh medium on day 4, and 5 µM AraC (Sigma) was added during each subsequent medium change, performed every 3 days.

Transfection of primary cultures with EGFP-tagged α-synuclein plasmid

On day 9 in vitro (DIV9), primary neuron–astrocyte co-cultures were transfected with an EGFP-tagged α-synuclein plasmid. The plasmid DNA (0.5 µg/well) and Lipofectamine™ 2000 (Invitrogen) were each diluted separately in Minimal Essential Medium (Opti-MEM™, Gibco) and incubated for 5 min at room temperature. The diluted DNA and Lipofectamine solutions were then combined at a 2:3 ratio (µg DNA:µL Lipofectamine) and incubated for 15 min at room temperature. Prior to transfection, the complete Neurobasal-A medium was removed from the cultures. The DNA-Lipofectamine complexes were added dropwise to the cells and incubated at 37 °C with 5% CO₂ for 4 h. After incubation, the medium was replaced with fresh, complete Neurobasal-A. Cells were treated after 36 h of transfection.

Induction of depolarization in transfected primary cells

To induce depolarization, primary cells were treated with HBSS containing 100 µM glutamate (Sigma) and 10 µM glycine (Tocris). Moreover, cells were treated with different concentrations of KCl (25 mM and 50 mM) (Fisher Scientific) in transfected cells to evaluate the exponential effects of depolarization. To ensure that the observed responses were due to depolarization rather than changes in osmolarity, control treatments were performed using 100 µM glutamine, 100 µM GABA, and 10 mM MgCl₂.

Calcium imaging with Fluo-4 Direct™ in primary neuron-astrocyte co-culture

For the calcium assay, the loading solution was prepared by adding 10 mL of Fluo-4 Direct™ calcium assay buffer and 200 μL of 250 mM probenecid stock solution to one bottle of 2X Fluo-4 Direct™ calcium reagent (Invitrogen). The solution was mixed 1:1 with culture media to achieve a final 1X Fluo-4 Direct™ calcium reagent concentration. After 24 h of treatment, the co-cultures were loaded with the 1X Fluo-4 Direct™ calcium reagent. The reagent was added to the glass-bottom dish containing cells, and the cells were incubated with the solution for 30 min at 37 °C. After incubation, the cells were incubated at room temperature for an additional 30 min. The dye was removed, and assay buffer was added during recording. Fluorescence intensity was recorded at 33 frames per second (fps) for 2 min on a Zeiss fluorescence microscope with an Axiocam 702 mono fast camera.

Measurement of membrane potential using di-8-ANEPPS

2 × 10⁵ cells (HT-22 and primary neuronal cells) were seeded into each 35-mm glass-bottom dish. Cells were treated with exogenous phosphatidylserine (DOPS) according to a previously described protocol, in the presence of 25 mM KCl, for 24 h. Following treatment, cells were gently washed three times with 1× PBS and incubated with 10 µM Di-8-ANEPPS for 15 min at 37 °C. After staining, cells were washed three times with 1× PBS and imaged using a 100× oil-immersion objective on an Olympus IX83 confocal microscope. Fluorescence intensity was recorded upon excitation at 440 nm and 530 nm and their emission was collected at 580–620 nm^66–68. For each cell, fluorescence intensities obtained at the two excitation wavelengths were quantified, and the ratio of fluorescence intensity measured upon 440 nm excitation to that measured upon 530 nm excitation (440/530) was calculated and plotted^66–68.

Data analysis

Calcium imaging analysis

Calcium imaging data were analysed using FIJI (ImageJ) and custom Python scripts. Regions of interest (ROIs) were manually selected for individual neurons. Fluorescence intensity (F) was recorded over time for each ROI. Baseline fluorescence (F₀) was calculated as the mean of the lowest 10 fluorescence values for each neuron. Changes in fluorescence (ΔF/F₀) were calculated using the formula: ΔF/F₀ = (F − F₀)/F₀. Peaks in calcium activity were detected using a peak-finding algorithm with parameters for minimum prominence, height, distance between peaks, and width. The data were plotted and statistically analysed for comparisons across experimental groups.

Colocalization analysis

Colocalization between α-synuclein (α-syn) and Synapsin-1 was quantified using Manders’ coefficient. Images were first thresholded to isolate α-syn puncta. Particle analysis was performed in FIJI to identify and count individual α-syn puncta. Colocalization was then assessed using the Coloc2 plugin. Mander’s coefficient was calculated to measure the fraction of α-syn puncta overlapping with Synapsin-1. This Manders’ value was multiplied by the total area of α-syn puncta. This gave the absolute colocalization area of α-syn with Synapsin-1. The resulting colocalized areas were plotted for comparison across experimental groups.

Quantification & statistical analysis

ImageJ software was used to analyse and process the images obtained from the experiments. A uniform detection setting was applied consistently across all the experiments to ensure comparability. The size of the condensates was manually quantified using ImageJ. Statistical analysis was performed using one-way ANOVA in SigmaPlot, with p-values less than 0.05 considered statistically significant for all the experiments. Some graphs were plotted using Origin Pro software and GraphPad (Prism).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

M.S., S.C. and S.G. conceptualized the work; M.S., S.C., S.G. and B.S.S. designed the research; J.S., A.N., T.M., A.S.M., K.B., G.M. and A.J.B. performed the research; M.S., S.C., S.G., B.S.S., J.S., A.N., T.M., A.S.M., K.B., G.M., A.J.B. and N.A.K. analysed and interpreted the data; M.S., S.C., S.G., J.S. and A.N. wrote the paper with contributions from all other co-authors.

Peer review

Peer review information

Nature Communications thanks the anonymous reviewers for their contribution to the peer review of this work. A peer review file is available.

Data availability

All data supporting the findings of this study are available within the main text and the Supplementary Information. Source data are provided with this paper.

Code availability

All code files have been deposited in the Zenodo repository: Here's the link: [10.5281/zenodo.18759321], and are publicly available.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-70840-2.