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

Domain: biology
Published: 2026-03-19
Editorial outcome: major_revision

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.

Introduction
Biomolecular condensates, also known as membraneless organelles, play a critical role
in diverse cellular functions 1. 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 emerge 4. Classic examples
include nucleoli 5 , Cajal bodies 6, and stress granules 7. 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 kinetics 14. 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
responses 16, cytoskeletal assembly 17, presynaptic contact zones 18, RNP granules in
fungi 19, and endocytosis 20. 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 behavior. 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 behavior
resembles a prewetting-like transition, where membrane attraction drives α-Syn
condensation on the membrane surface below the bulk saturation threshold 21. 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 Figure 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 hours (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 Figure 1b-c).

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 hours (Fig. 1b-f). While the physiological bulk concentration of α-Syn is
estimated at ~22 µM 22, it is known that nanomolar to low micromolar concentrations can
suffice for surface-induced aggregation in vitro 23. No detectable binding of α-Syn was
observed on TBE, DOPC, or DOPC:DOPS (4:1) membranes over the time-course of 25
hours (Fig. 1b, 1c, 1f & Supplementary Figure 2), consistent with previous reports 24.
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 Figure 3). Intriguingly, condensation occurred
only on the DOPC:DOPS (6:4) membrane, emerging at around 24 hours. 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 100nM and 2
µM in the absence of membrane surface and crowding in line with previous reports
(Supplementary Figure 4a-b). Membrane surface (PC:PS 6:4) also did not show any
condensation over the time course of 26 hours in the absence of α-Syn (Supplementary
Figure 4c). No condensation of α-Syn was observed neither in the presence of ~ 1µM
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 Figure 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 Figure 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 Figure 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 hours (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 10nM, 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 CaCl which is expected to neutralize
2
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 Figure 9 a-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 hours;
however, strong and stable binding became evident only after approximately 18 hours
(Supplementary Figure 10a). Condensate formation began after ~23 hours 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 Figure 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
Figure 11). Strikingly, around 26 hours, we observed fibrillar structures emerging from
mature α-Syn droplets tubulating the membrane surface (Fig. 2d & Supplementary Figure
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.

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 analyzing 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 - 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 Figure 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.

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 hour), 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).

To examine membrane mechanics, we used micropipette aspiration of giant unilamellar
vesicles (GUVs). In the absence of protein, the protrusion length (L ) of aspirated GUVs
p
remained stable for over 10 minutes, establishing baseline tension (Fig. 4c,
Supplementary video 3). However, after injection of 100 nM monomeric α-Syn, a reduction
in L and expansion of the spherical region of the GUVs were observed, indicating a rapid
p
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-leaflet 26. The observed reduction in
membrane tension indicates protein induced compensatory response to curvature stress.
Within ~15 seconds, 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 = 6.79 μM) compared to early-stage binding (K = 12.5 μM) (Fig. 4e), suggesting
d d
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 layer 27. 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.

Zeta potential, ζ = ψ e−κd (1)
0
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.

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 ) exceeds a critical threshold (J ). Similarly,
m cm
protein condensation into bulk liquid-like phases requires that the protein-protein
interaction strength (J ) exceeds a critical value (J ), provided the protein concentration
p cp
surpasses the saturation limit. Importantly, even when bulk phase separation is not
favored, 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 surface 29. To estimate the PS induced changes in membrane interfacial
potential we used voltage-sensitive probes, such as ANSEPPS (4-[2-[6-(dioctylamino)-2-
naphthalenyl]ethenyl]-1-(3-sulfopropyl)-pyridinium). Ratio of ANEPPS fluorescence
intensity at 530nm/440nm (R) is related to voltage change (ΔV ) as follows:
m
R = R
o
+ S ΔV
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 hours (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.

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 31,32. 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 Figure 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 Figure 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 MgCl , 100 μM glutamine (Glutamate derivative that does not act as a
2
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 Figure 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.

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 same 59. Avidin-coated chamber slides were purchased from Sigma-Aldrich.

Protein expression and purification
α-Synuclein was expressed and purified as described earlier 60. 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 uM Tris-HCl, 10 uM EDTA, 150 uM 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 15000 g for 10 min at four °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 15000 g for 15 mins at 4 °C and
then washed with 1 mL of 50% ammonium sulfate solution. The resulting pellet was then
resuspended in 900μL 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 minutes
for 120 hours. To accelerate aggregation and improve reproducibility, we added glass
beads of homogeneous size and spherical shape, which were selected by visual
inspection61,62. Glass beads (#LA715A, Himedia) were dipped in MQ water for ten
minutes 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-minute water
bath treatment, it was vortexed for 5 minutes. 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 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 hours. 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 hours 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 minutes, 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 hours. The dried lipid film was then rehydrated in
PBS buffer (pH 7.4, 300 ± 5 mOsm). The mixture was then sonicated for 10 mins at 50%
amplitude with a 10 sec 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
minutes 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 (561nm) and Alexa-488(488nm). 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-labeled
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 seconds, followed every few seconds for
2-3 minutes 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 (𝜏 ), which is the recovery half-time. The diffusion
1
2
coefficient (D) was calculated using the Soumpasis equation for 2D diffusion:
r
r2
D = 0.224 (1)
r
τ1
2
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-hour dark incubation, the fluorescence anisotropy was measured every 4 hours till 72
hours 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 = IVV−GIVH (2)
IVV+2GIVH
where 𝐼 and 𝐼 are the fluorescence intensities of the vertical and horizontal
𝑉𝑉 𝑉𝐻
components, respectively. The grating correction factor G (= IHV ) corrects for any
IHH
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 mins 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 ) was reached. The isotherm data were
c
further analyzed to calculate the compressibility modulus (𝐶−1), which is defined as
s
𝐶−1 = −A dπ (3)
s
dA
Where A is the molecular area, and dπ Represents the derivative of surface pressure
dA
concerning the molecular area.

Micropipette aspiration
All the measurements were conducted using an Olympus IX83 epifluorescence
microscope equipped with Hoffman modulation optics and a 100X 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 analyze 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=0,1. The x- and y-
i
directions have periodic boundaries. In the z-direction the membrane is present at z=L,
and is modeled as a two-dimensional Ising system. The two states s=±1 represent the
i
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 𝜙 fraction of the s=+1 spins
t
and extend L sites into the bulk. The total Hamiltonian for the system is given as
t
H = −J ∑ s s −J ∑ n n −J ∑ n −μ∑ n (4)
m 〈i,j〉 i j p 〈i,j〉 i j t i∈tether i i∈bulk i
∈mem ∈bulk
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 . The proteins and tethers interact via the coupling constant J which acts
p t,
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 T. The
B
calculated values for the measured parameters are presented in Table 1.

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, defined 65 as
C = ∑N (𝜙 −𝜙 ) (5)
s n=1 n b
Where 𝜙 is the bulk volume fraction far away from the surface and can be calculated as.
b
𝜙 = 1 ∑M 𝜙 (6)
b M i=1 i

Simulation parameters
Table 1: Mathematical Simulation parameters
Parameter used Range/Value
Lattice size, L 20
Tether length, L 0-10
t
Tether density, 𝜙 0.005-0.6
t
Fraction of up spins, 𝜙 0.1-0.8
m
Protein-protein interaction strength, J 0.6-2.0
b
/k T
B
Lipid-lipid interaction strength, J /k T 0.3-0.6
m B
Tether-protein interaction strength, J /k T 0.0-3.0
t B
Bulk chemical potential, -μ /k
B
T 1.5-6.5

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 hours, 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 0.75mg/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 1µg/ml of Doxycycline, 24 hours after plating. Media was replaced with fresh media
containing 1µg/ml of Doxycycline to which 70µM, 100µM and 150µM of exogenous
Phosphatidyl Serine (DOPS), were added respectively for each condition. After 24 hours
of induction and treatment with 70µM, 100µM and 150µM 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 1x PBS thrice and then fixed using 4%
Paraformaldehyde (PFA) for 20 minutes at room temperature. Following fixation, cells
were washed gently with 1x PBS thrice, and then the cells were permeabilized with 0.1%
Triton X-100 for 10 minutes and blocked using 1% BSA for 45 minutes. The primary
antibodies, rabbit MAP2 (1:150, Cell Signaling 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 Signaling Technology, #4412S), anti-
mouse Alexa Fluor 555 (1:1000, Cell Signaling 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 minutes in a humidified chamber in the dark. Coverslips were mounted using DAPI-
containing mounting media, and images were captured using 100x 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 minutes. 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 minutes 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 (0.5µg/well) and Lipofectamine™
2000 (Invitrogen) were each diluted separately in Minimal Essential Medium (Opti-
MEM™, Gibco) and incubated for 5 minutes 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 minutes 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
hours. After incubation, the medium was replaced with fresh, complete Neurobasal-A.
Cells were treated after 36 hours 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 hours 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 minutes at 37°C. After incubation, the cells were
incubated at room temperature for an additional 30 minutes. The dye was removed, and
assay buffer was added during recording. Fluorescence intensity was recorded at 33
frames per second (fps) for 2 minutes 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 hours. Following
treatment, cells were gently washed three times with 1× PBS and incubated with 10 µM
Di-8-ANEPPS for 15 minutes 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 580-620nm66-68. 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 plotted66-68.

Data Analysis
Calcium Imaging Analysis
Calcium imaging data were analyzed 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 analyzed 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 analyze 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).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This paper sets out to investigate how the interfacial potential of lipid membranes governs the condensation and fibrillation of α-synuclein (α-Syn)—a key protein implicated in Parkinson’s disease (PD). The authors demonstrate that specific membrane compositions facilitate α-Syn condensation, as the changing of lipid composition will later the interfacial potential of lipid membrane. Furthermore, they show that membrane surfaces act as nucleation sites, where α-Syn initially forms liquid-like condensates that later transition into fibrils. This process involves a prewetting-like transition, supported by lattice gas simulations. In cellular experiments, the authors find that perturbing the membrane interfacial potential—either by adding PS liposomes or KCl—alters intracellular condensate formation by modulating membrane potential, consistent with in vitro results. The study focuses on the interplay between lipid membrane potential and α-Syn condensation, a topic of significant importance. I would love to congratulate the authors for this discovery. While the article reports intriguing phenomena and this new concept raised by the authors is very important and tends to be general, several key questions remain to be addressed. Specifically, more evidence should be provided to prove that the membrane interfacial potential is the most critical determinant in facilitating α-Syn condensation. After the authors conducted the following key controls and critical experiments, this paper can be published.\nComments:\n1. Controls missing for lipid compositions in Fig. 1a. In Figure 1a, while the authors examined α-Syn aggregation in the presence of PC, PS, and mixed PC:PS vesicles, the experimental design lacks rigorous control groups. Specifically, the PC and PS lipids were each only tested at a single concentration. However, for the PC:PS mixtures (6:4 and 4:1), it would be critical to include controls with corresponding individual PC and PS concentrations to distinguish specific synergistic effects from simple additive effects of each lipid.\n2. Tuning the stoichiometry of lipid may lead to the phase separation of lipids, which could be the driving force of α-Syn condensation by providing nucleation sites. Or in other words, changing the stoichiometry of lipids may not be as simple as tuning the stoichiometry or two miscible small molecules. Could you discuss regarding this matter with experimental evidence, as the formation of lipid domains may also provide nucleation site. The title claimed that the interfacial potential of membrane is the determinant of α-Syn condensation, therefore ruling our other possibilities would be necessary.\n3. For Figure 1b–f, please provide separate single-channel images (Rhodamine-PE in red, α-Syn Hilyte-488 in green) alongside the merged images to clearly visualize co-localization or exclusion.\nAdditionally, the current panel labeling appears inconsistent: Figure 1e and 1f both contain α-Syn and should be sequentially labeled following 1d to maintain logical flow.\n4. The lipid area visualized via Rho-PE in Figure 1d (PC:PS 6:4) is markedly smaller than in Figures 1b, c, e, and f. Could this indicate partial detachment or fewer intact bilayers, potentially allowing more free Rho-PE in solution? This is important because if excess free Rho-PE is present, it could potentially facilitate α-Syn condensation independent of the supported membrane.\n5. Base on the previous question, the critical concentration for the condensation of α-Syn is 20 times lower in the presence of lipid membrane (PC:PS=6:4). Could author provide more experimental evidence showing that this facilitated condensation is not due to that PC or PS were crosslinking the α-Syn in the form of coacervation, rather than due to the change of interfacial potential?\n6. In Figure 2a, α-Syn condensates appear to colocalize with Rho-PE. Could this imply that free Rho-PE micelles or aggregates are directly participating in condensate formation? If so, this would complicate the interpretation that membrane stoichiometry and interfacial potential are the primary determinants. Please provide more experimental data regarding this issue (e.g., with mass spectroscopy of α-Syn condensates).\n7. Figure 5a uses zeta potential as a surrogate for interfacial potential. While zeta potential (ζ) reflects the electrical environment near the shear plane, it does not directly equal the interfacial potential (ψ₀) at the membrane surface.\n(1) Could the authors discuss more mathematically whether higher ζ guarantees higher ψ₀ under their experimental ionic conditions?\n(2) Additionally, could Stark-effect-based voltage-sensitive probes, such as ANEPPS, be employed to directly estimate interfacial potential or changes in local field strength more precisely?\n(3) The section on α-Syn reducing membrane tension and compressibility is intriguing but currently feels disconnected from the central condensation mechanism. Could the authors elaborate mechanistically how reduced membrane tension promotes or regulates α-Syn surface condensation?\n8. The cellular experiment for Figure 6 is interesting. Please provide a schematic illustration to better demonstrate the phenomenon and the underlying mechanism. A question regarding condensation is that the strategy the author used to tune membrane potential is adding salts or osmolytes, which in turn should also affect the condensation threshold of alpha-syn, so some control experiments are needed to understand the interplay. Further, the alterations of membrane potential are not experimentally demonstrated.", "reviewer_2": "Shaikh et al. aimed to test how lipid membranes affect alpha-synuclein's (aSyn) phase separation. aSyn is known to phase separate at very high concentrations and/or with help of crowding agents. Here they show that membrane interaction promotes aSyn condensation at physiological concentrations without crowding agents. Furthermore, these condensates can nucleate fibril formation. Biophysical aspects of the paper suggest a possibly interesting cascade of aSyn nucleation on membranes which has been suggested before, but has remained mechanistically/biophysically unclear.\nIn cell culture experiments (Fig. 6), the results are difficult to interpret for multiple reasons: \"aggregate\" and \"puncta\" are used interchangeably. The authors should keep to \"puncta\" in panels (a-b), since puncta are being observed and measured.\nPanel (c) shows blebbing however, possibly due to overexpression+depolarization. Large size of the GFP tag and overexpression make this experiment difficult to interpret. Panel (d) may or may not have any change in aSyn colocalization with synapsin-1 between the different conditions. Recruitment of aSyn to presynapses is not clear. In panel (d & g), the extracellular puncta can be in the presynaptic and/or axonal compartments (where natively expressed aSyn actually localizes), since MAP2 localizes to dendritosomatic compartments, which do not have aSyn unless overexpressed."}, "author_response": "RESPONSE: We thank the reviewer for acknowledging the novelty and importance of the findings and positive assessment of our work. We also thank the reviewer for excellent constructive comments that have immensely strengthened the work.\nWe would like to point out that in order to accommodate new experimental data that is more important we have now split the earlier Fig 6 into two figures (now Fig 6 and Fig 7) and moved some aspects to the supplementary file as detailed below.\nComments:\n1. Controls missing for lipid compositions in Fig. 1a. In Figure 1a, while the authors examined α-Syn aggregation in the presence of PC, PS, and mixed PC:PS vesicles, the experimental design lacks rigorous control groups. Specifically, the PC and PS lipids were each only tested at a single concentration. However, for the PC:PS mixtures (6:4 and 4:1), it would be critical to include controls with corresponding individual PC and PS concentrations to distinguish specific synergistic eUects from simple additive eUects of each lipid.\nRESPONSE: We thank the reviewer for raising this excellent point. As suggested we have now performed new sets of ThT experiments to take into account the diUerent concentrations of PC and PS, both individually (i.e, PC, PS both at 100 μM and 300 μM, in addition to the previously used 200 μM) as well as in mixtures (i.e, PC:PS 6:4/4:6/4:1 at 100 μM and 300 μM, in addition to the previously used 200 μM). To make the results comparable, we performed all the experiments again. This is important to take care of the instrumental oUsets for ThT intensities that vary 5-10% everytime. 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 hours (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). As rightly pointed out by the reviewer the specific stoichiometry of DOPC/DOPS indeed provides a unique lipid membrane environment that promotes nucleation synergistically. Similar kinetic behaviour is seen for all the chosen concentrations suggesting that the lipid concentration has no bearing on the behaviour of the mixture (Suppl Fig. 1b-c).\nWe have now incorporated new sets of ThT experiments in Fig 1a and Supplementary Fig 1b-c. (Page No. 5, Line Nos 94-106).\nPlease note that the current ThT kinetics data showed some diUerences compared to our earlier observations with regard to PC:PS 6:4 behaviour that could be attributed to instrumental error. Earlier we saw an anomalous behavior as in increase in the intensity followed by a decrease and our interpretation was perhaps it is due to settling of droplets or mature fibrils. However, upon using beads during the experiment to avoid the settling down of the protein we do not see such behaviour anymore. We have ensured the correctness of the ThT kinetics data by rigorously repeating the ThT experiments multiple times.\n2. Tuning the stoichiometry of lipid may lead to the phase separation of lipids, which could be the driving force of α-Syn condensation by providing nucleation sites. Or in other words, changing the stoichiometry of lipids may not be as simple as tuning the stoichiometry or two miscible small molecules. Could you discuss regarding this matter with experimental evidence, as the formation of lipid domains may also provide nucleation site. The title claimed that the interfacial potential of membrane is the determinant of α-Syn condensation, therefore ruling our other possibilities would be necessary.\nRESPONSE: We thank the reviewer for this insightful comment. We have now performed new set of experiments with ternary membrane conditions containing (PC/SM/Cholesterol) in diUerent stoichiometries. We chose the stoichiometries of (5:5:0, 4:4:2 and 3:3:4) that enable diUerent degrees of phase separation (i.e, l and l d o phases) that are well characterized on phase diagrams. As evident we did not observe the condensation of α-Syn in any of the three conditions (Supplementary Fig. 8). Though the GUVs used to prepare the SLBs showed distinct phase separated domains but upon the adsorption and formation of the SLBs such phase separated domains were not visible microscopically. Some amount of the protein was seen binding to the defects in the SLBs that appear very diUerent from a typical condensate. Though no phase separated domains are microscopically visible on the SLBs, sub-micron scale regions of the lipid membrane representing diUerent order cannot be totally ruled out. We think phase separating boundaries ideally should increase the tension in the membrane that would probably hinder condensation.\nWe have now included this new experimental data in the supporting information and we have also discussed this important aspect in results section (Page No. 6, Line No. 130-135).\n3. For Figure 1b–f, please provide separate single-channel images (Rhodamine-PE in red, α-Syn Hilyte-488 in green) alongside the merged images to clearly visualize co-localization or exclusion.\nAdditionally, the current panel labeling appears inconsistent: Figure 1e and 1f both contain α-Syn and should be sequentially labeled following 1d to maintain logical flow.\nRESPONSE: We have now incorporated single-channel images for the entire panel as suggested by the reviewer in Fig 1. We have also rectified the sequential labelling to make it consistent (Page No. 5-6, Line No. 107-123 as well as Fig 1 legends).\n4. The lipid area visualized via Rho-PE in Figure 1d (PC:PS 6:4) is markedly smaller than in Figures 1b, c, e, and f. Could this indicate partial detachment or fewer intact bilayers, potentially allowing more free Rho-PE in solution? This is important because if excess free Rho-PE is present, it could potentially facilitate α-Syn condensation independent of the supported membrane.\nRESPONSE: This is an interesting point. We do think that in this particular condition (PC:PS 6:4) there were few intact bilayers giving rise to a smaller membrane island which happens many a times as the SLB formation and size is beyond our control. We did not crop the field of view, as the droplets were also observed outside the island area. We think the surrounding area is likely to contain the earliest adsorbed remnants of burst-out vesicles, possibly micellar suspensions containing PC/PS and Rh-PE lipids. To rule out reviewers concern that excess Rh-PE alone could be driving α-Syn condensation independent of the supported membrane, we performed a new experiment monitoring if Rho-PE alone can drive α-Syn condensation? As shown in the new figure panel (Fig 1g), no condensation was observed. Likewise, free lipid suspension of PC/PS 6:4 and 4:6 (doped with Rho-PE, Fig 1h) also did not induce α-Syn condensation suggesting that the surface is essential for α-Syn condensation for lower physiological concentration.\nWe have now included this new experimental data as Fig 1g and we have also discussed this important aspect in results section (Page No. 6, Line No. 123-125).\n5. Base on the previous question, the critical concentration for the condensation of α-Syn is 20 times lower in the presence of lipid membrane (PC:PS=6:4). Could author provide more experimental evidence showing that this facilitated condensation is not due to that PC or PS were crosslinking the α-Syn in the form of coacervation, rather than due to the change of interfacial potential?\nRESPONSE: We thank the reviewer for this very interesting point. To rule out PC or PS crosslinking α-Syn, we performed the same experiment in the presence of 50 mM and 100 mM CaCl which is expected to neutralize the negative charges of the PS. 2 Crosslinking would be insensitive to ionic screening by Ca2+ and facilitate condensation driven by specific, high-aUinity 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.\nWe have now included this new experimental data in the supporting information and we have also discussed this important aspect in results and discussion (Page No. 7, Line No. 149-157).\n6. In Figure 2a, α-Syn condensates appear to colocalize with Rho-PE. Could this imply that free Rho-PE micelles or aggregates are directly participating in condensate formation? If so, this would complicate the interpretation that membrane stoichiometry and interfacial potential are the primary determinants. Please provide more experimental data regarding this issue (e.g., with mass spectroscopy of α-Syn condensates).\nRESPONSE: To rule out the direct participation of Rho-PE in the condensation process, we performed another new experiment where we observe α-Syn condensation on unlabelled PC/PS 6:4 observed without the doped Rho-PE. The bilayer islands were scanned and visualized after the initial binding of the labelled α-Syn-HiLyte 488. This confirms that PC/PS in the absence of Rho-PE are suUicient to drive condensation and whatever Rho-PE signal is obtained in the doped bilayers is because of the passive incorporation of the probe.\nWe have now discussed this aspect in results and included the new experimental data as Supplementary Figure 7 (Page No. 6, Line No. 127-130).\n7. Figure 5a uses zeta potential as a surrogate for interfacial potential. While zeta potential (ζ) reflects the electrical environment near the shear plane, it does not directly equal the interfacial potential (ψ₀) at the membrane surface.\n(1) Could the authors discuss more mathematically whether higher ζ guarantees higher ψ₀ under their experimental ionic conditions?\nRESPONSE: We thank the reviewer for this excellent comment. 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.\nFurther, Zeta potential ζ = ψ e−κd 0 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.\nFor 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 unless there is a significant change in ionic strength of the environment which is not the case.\nWe have now discussed this aspect in results section (Page No. 10,11, Line No. 272-284).\n(2) Additionally, could Stark-eUect-based voltage-sensitive probes, such as ANEPPS, be employed to directly estimate interfacial potential or changes in local field strength more precisely?\nRESPONSE: We thank the reviewer for this excellent suggestion. We have now included new ratiometric imaging of ANEPPS fluorescence emission 530nm/440nm under changing lipid environment to provide a reasonably precise estimate of membrane interfacial potential. Ratio of ANEPPS fluorescence intensity at 530nm/440nm (R) is related to voltage change (ΔV ) as follows: m R = R + S ΔV o m where, R is the ratio at the resting membrane potential (or control condition) S is the o 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.\nThe ratiometric imaging of HT22 cells results in a significant increase in the R upon addition of PS vesicles indicating a change in local field strength (Figure 6a-b). This confirms that indeed altering lipid composition induces change in local electric field strength or potential.\nWe have now added these new data and discussed in the results section (Page No 11,12, Line No. 312-322, 330-333)\n(3) The section on α-Syn reducing membrane tension and compressibility is intriguing but currently feels disconnected from the central condensation mechanism. Could the authors elaborate mechanistically how reduced membrane tension promotes or regulates α-Syn surface condensation?\nRESPONSE: We thank the reviewer for pointing this out. We have now elaborated the plausible mechanistic aspects of how these parameters facilitating condensation. 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-leaflet. The observed reduction in membrane tension (ordering of lipids) is a protein induced compensatory response to curvature stress. This ordered platform 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.\nWe have now incorporated this aspect in the Figure 4 results section (Page No. 9, Line No. 237-241; Page No. 10, Line No. 251-257).\n8. The cellular experiment for Figure 6 is interesting. Please provide a schematic illustration to better demonstrate the phenomenon and the underlying mechanism.\nRESPONSE: We have now included a new schematic model capturing the concept and mechanism in Fig. 8 depicting how perturbations of interfacial potential can govern the condensation and subsequent fibril formation of α-Syn.\nA question regarding condensation is that the strategy the author used to tune membrane potential is adding salts or osmolytes, which in turn should also aUect the condensation threshold of alpha-syn, so some control experiments are needed to understand the interplay.\nRESPONSE: As per reviewer suggestion we have now performed a series of control experiments to rule out the role of salts/osmolytes on α-Syn condensation. First, we treated the primary neurons with 10 mM MgCl and did not see any condensation, ruling 2 out the possibility of any salt causing the condensation (Fig. 7b-c; Supplementary Figure. 15). Next we treated the cultures with 100uM Glutamine (Glutamate derivative that does not act as a neurotransmitter) and we did not see any condensation. Further, we treated the cultures with GABA (an inhibitory neurotransmitter known to hyperpolarize neuronal membranes) and we did not see any α-Syn condensation (Fig. 7b-c; Supplementary Figure. 15). This proves that the depolarizing membrane potential is essential for α-Syn condensation to be triggered.\nWe have now added these new observations to results section (Page No. 13, Line No. 364-370)\nFurther, the alterations of membrane potential are not experimentally demonstrated.\nRESPONSE:\nAs suggested by the reviewer we have experimentally quantified the change in the electric field strength or voltage via ratiometric imaging of ANEPPS on primary neurons. We found that application of 25 mM KCl caused increase in the R value by 2-3 fold indicating that the neuronal membrane is getting depolarized (Fig. 7d-e). The details are same as the response to point 7 (2).\nWe have now added these new observations to results section (Page No. 13, Line No. 362-364)\nReviewer #2 (Remarks to the Author):\nShaikh et al. aimed to test how lipid membranes aUect alpha-synuclein's (aSyn) phase separation. aSyn is known to phase separate at very high concentrations and/or with help of crowding agents. Here they show that membrane interaction promotes aSyn condensation at physiological concentrations without crowding agents. Furthermore, these condensates can nucleate fibril formation. Biophysical aspects of the paper suggest a possibly interesting cascade of aSyn nucleation on membranes which has been suggested before, but has remained mechanistically/biophysically unclear.\nWe thank the reviewer for finding the work interesting and providing excellent comments that have immensely strengthened the work. We would like to point out that in order to accommodate new experimental data that is more important we have now split the earlier Fig 6 into two figures (now Fig 6 and Fig 7) and moved some aspects to the supplementary file as detailed below.\nIn cell culture experiments (Fig. 6), the results are diUicult to interpret for multiple reasons: \"aggregate\" and \"puncta\" are used interchangeably. The authors should keep to \"puncta\" in panels (a-b), since puncta are being observed and measured.\nRESPONSE: As per the suggestion of the reviewer we have now carefully reviewed the manuscript and ensured that we consistently use the term “puncta” throughout results section describing cellular experiments in Fig 6 and Fig 7.\nPanel (c) shows blebbing however, possibly due to overexpression+depolarization. Large size of the GFP tag and overexpression make this experiment diUicult to interpret.\nRESPONSE: We thank the reviewer for raising this point. We have now repeated the key experiments under conditions of endogenous aSyn expression, as is known to occur in primary cortical neurons (Rideout et al., 2002; Clough et al., 2011). Our results remain consistent, both 25 mM and 50 mM KCl elicit a pronounced increase in the number and size of aSyn puncta (Fig. 7 f-h). This critical replication demonstrates that our original observations are robust and not a consequence of aSyn overexpression or the presence of a GFP tag.\nWe have now added these new observations to results section (Page No. 13, Line No. 370-378)\nPanel (d) may or may not have any change in aSyn colocalization with synapsin-1 between the diUerent conditions. Recruitment of aSyn to presynapses is not clear.\nRESPONSE: We thank the reviewer for pointing this out. We may not have been clear previously although we had included the data addressing this aspect in the previous supplementary figure 20. By colocalization analysis using Manders' coeUicient (details in method section), we have shown that area of aSyn puncta colocalizing with Synapsin I puncta is greater under depolarizing conditions (now Supplementary Fig. 16). However we agree with the reviewer that without delineating the axonal compartment it may be diUicult to interpret the presynaptic localization. Therefore, to improve visualization of the aSyn puncta, we have now stained the neurons with antibody against βIII tubulin and synapsin I to locate the presynapse more accurately and then shown the endogenous aSyn puncta (Fig. 7i,j).\nWe have now added these new observations to results section (Page No. 13-14, Line No. 378-382)\nIn panel (d & g), the extracellular puncta can be in the presynaptic and/or axonal compartments (where natively expressed aSyn actually localizes), since MAP2 localizes to dendritosomatic compartments, which do not have aSyn unless overexpressed.\nRESPONSE: We thank the reviewer for their insightful comment regarding the subcellular localization of α-synuclein (aSyn). We agree that aSyn is primarily presynaptic in mature neurons; however, its localization is dynamic. We would like to clarify two key points:\nContext of Localization: In young cultured neurons, aSyn is initially expressed in the soma before traUicking to the presynapse, a slow process that explains its somatodendritic presence in our system which consists of primary neurons from 1 week old cortical cultures (Burré et al, Cold Spring Harb Perspect Med. 2018; Murphy et al., J Neurosci. 2000). Furthermore, a redistribution of endogenous/native aSyn back to the somatodendritic compartment is a recognized feature of both aging and pathology, as shown in human brain studies (Calabresi et al., Cell Death Dis, 2023; Chu et al., Neurobiol Dis. 2007; Benskey et al., J Neurochem. 2016; Marui et al., J. Neurol. Sci., 2002). Our model of chronic depolarization (25-50 mM KCl) is designed to mimic a pathophysiological state, specifically, increased neuronal excitability (Chen et al., npj Parkinsons Dis.2025) which has recently been shown to be associated with α-synuclein aggregation. Therefore, the localization we observe is consistent with these established mechanisms.\nEvidence for Axonal/Presynaptic aSyn: To directly address the reviewer's concern, we have now performed additional co-staining experiments. As shown in our new Figure 7f-j, we detect endogenous/native aSyn puncta clearly localized within the axonal compartment, and a significant portion of this signal colocalizes with the presynaptic marker Synapsin-I (Figure 7i-j) . This confirms that aSyn is also present in its canonical location in our model.\nFinally, the persistence of extracellular aSyn puncta under depolarizing conditions (50 mM KCl) indicates that depolarization is aiding release of α-Syn condensates (Figure 7f).\nIn summary, our data capture aSyn in a dynamic state, with the presence of endogenous aSyn in axonal/presynaptic compartments and in the extracellular milieu upon membrane depolarization.\nWe have now added these new observations to results section (Page No. 13-14, Line No. 370-382)"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The author has greatly improved the manuscript in response to the reviewers' comments. However, some final issues require attention before the article can be accepted.\n1. In Figure 1, the authors used glass beads to accelerate aggregation and improve reproducibility. Please provide references to show that this is a standard protocol. Additionally, please provide the catalog number of the glass beads used and state whether they were properly washed or not.\n2. The author used Di-8-ANEPPS to measure membrane potential. However, standard ratiometric measurement with Di-8-ANEPPS is typically performed by comparing the probe's emission at different excitation wavelengths (~420 nm and ~520 nm), not by comparing the emission ratio as was done by the author. Please provide references supporting your experimental design, or redo the corresponding experiments.\n3. Please provide a more detailed explanation in the legend for Figure 8.", "reviewer_2": "Shaikh et al. have addressed most of this reviewer's concerns adequately. A key cell biological concern remains. The evidence shown of the release of alpha synuclein \"puncta\" is not convincing, for the following reasons: 1. No extracellular puncta are visible in Fig. 7a, but these are quantitated in Fig. 7c, 2. The supposed extracellular puncta (blue arrow in Fig. 7f of in supplementary Fig. 13) are shown with BetaIIItubulin and Map2 co-staining, and without any synaptic co-staining, and as mentioned in the previous review, these could be synapses, 2. While it is difficult to accurately gauge the molecular size by puncta size alone, the puncta shown outside of Map2 staining in Supplementary Fig. 13 and outside of Map2+BetaIIItubulin staining in Fig. 7f are very large (in microns). If these are extracellular, and released directly from cytosol (as shown in Fig. 8 schematic), and not by membrane-membrane fusion, their exit would come with large breaches of neuronal plasma membrane. The study has multiple points of novelty and strength without emphasizing the release of large synuclein puncta from living primary neurons."}, "author_response": "Point-By-Point Response To Reviewer Comments (NCOMMS-25-45391A)\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThe author has greatly improved the manuscript in response to the reviewers' comments. However, some final issues require attention before the article can be accepted.\nResponse: We thank the reviewer for finding the revised version greatly improved.\n1. In Figure 1, the authors used glass beads to accelerate aggregation and improve reproducibility. Please provide references to show that this is a standard protocol. Additionally, please provide the catalog number of the glass beads used and state whether they were properly washed or not.\nResponse: We have now added the catalogue number of the glass beads and the following two citations supporting the use of glass beads for ThT kinetics in the methods section (Page No.18, Line No. 493-495).\n1. Potent α-Synuclein Aggregation Inhibitors, Identified by High-Throughput Screening, Mainly Target the Monomeric State.\nKurnik, Martin et al., Cell Chemical Biology, 25, 11, 1389-1402.e9 (2018)\n10.1016/j.chembiol.2018.08.005\n2. α-Synuclein fibrils enhance HIV-1 infection of human T cells, macrophages and microglia.\nOlari, LR. et al., Nat Commun 16, 813 (2025)\nhttps://doi.org/10.1038/s41467-025-56099-z\n2. The author used Di-8-ANEPPS to measure membrane potential. However, standard ratiometric measurement with Di-8-ANEPPS is typically performed by comparing the probe's emission at different excitation wavelengths (~420 nm and ~520 nm), not by comparing the emission ratio as was done by the author. Please provide references supporting your experimental design, or redo the corresponding experiments.\nResponse: We apologize for missing out on citations on this aspect. We used the following 3 citations that demonstrated ratiometric imaging to quantify potential changes. All the three studies used the excitation wavelengths of ~ 420 nm and ~ 520 nm to excite the samples and measured their emission intensity ratio R440/530 at a far red wavelength of ~ 580-620 nm. In our experiments we also used the excitation wavelength of 440nm and 530nm and measured their emission intensity in the far red ~ 580-620nm as detailed in the method section.\n1. Influence of Anions and Cations on the Dipole Potential of Phosphatidylcholine Vesicles: A Basis for the Hofmeister Effect.\nClarke, et al. Biophysical Journal, 5, 76, (1999)\nhttps://doi.org/10.1016/S0006-3495(99)77414-X\n2. Optical detection of membrane dipole potential: avoidance of fluidity and dye-induced effects.\nClarke RJ, et al. Biochim Biophys Acta, Jan 31;1323(2):223-39.,(1997)\nhttps://doi.org/10.1016/s0005-2736(96)00188-5.\n3. Effects of the Membrane Dipole Potential on the Interaction of Saquinavir with Phospholipid Membranes and Plasma Membrane Receptors of Caco-2 Cells.\nAsawakarn, Tanong et al. Journal of Biological Chemistry, 42, 276 (2021).\nhttps://doi.org/10.1074/jbc.M103269200.\n(Page No. 26-27, Line No. 731-741).\n3. Please provide a more detailed explanation in the legend for Figure 8.\nResponse: We have elaborated the details of the mechanism in the Fig 8 legend.\n(Page No. 42, Line No. 1107-1113).\nReviewer #2 (Remarks to the Author):\nShaikh et al. have addressed most of this reviewer's concerns adequately. A key cell biological concern remains. The evidence shown of the release of alpha synuclein \"puncta\" is not convincing, for the following reasons: 1. No extracellular puncta are visible in Fig. 7a, but these are quantitated in Fig. 7c,\nResponse: We thank the reviewer for pointing this out and we agree with reviewers concern. It is important to note that not every field of view showed both intracellular and extracellular puncta and there is some heterogeneity. And previously we were focussing more on showing the intracellular puncta. We have now provided representative images that capture both the intracellular and extracellular puncta in Fig 7a and edited the corresponding Fig 7 legend.\n2. The supposed extracellular puncta (blue arrow in Fig. 7f of in supplementary Fig. 13) are shown with BetaIIItubulin and Map2 co-staining, and without any synaptic co-staining, and as mentioned in the previous review, these could be synapses,\nResponse: We agree with the reviewer, however, we would like to point out that Fig 7i shows the presence of extracellular puncta in neuronal cultures co-stained with Beta III tubulin and Synapsin 1 marker. We have further added blue arrows to Fig 7i marking the extracellular puncta for better visualization and edited the corresponding Fig 7i legend.\n2. While it is difficult to accurately gauge the molecular size by puncta size alone, the puncta shown outside of Map2 staining in Supplementary Fig. 13 and outside of Map2+BetaIIItubulin staining in Fig. 7f are very large (in microns). If these are extracellular, and released directly from cytosol (as shown in Fig. 8 schematic), and not by membrane-membrane fusion, their exit would come with large breaches of neuronal plasma membrane. The study has multiple points of novelty and strength without emphasizing the release of large synuclein puncta from living primary neurons.\nResponse: These are interesting points raised by the reviewer and technically limiting. We think the heterogeneity in the size of puncta, be it intracellular or extracellular, arises as the dynamics of puncta growth are limited by the local concentration of the protein as well as the neighbouring punctas that can cluster together to form a larger punctas. Our cell based experiments are limited by the static visualization of punctas capturing various punctas at different states of their dynamic growth.\nWe thank the reviewer for this second point regarding the possibility of the membrane-membrane fusion. We agree with the reviewer that conventional vesicle driven exocytic extracellular secretion of punctas cannot be ruled out. We demonstrate that the potential changes at the membrane surface can regulate surface condensation of the Synuclein that can be potentially one of the routes for its extracellular release via direct deformation of the membrane. But we agree with the reviewer and have now toned down the aspect of the extracellular puncta release in the discussion clearly stating that while interfacial potential could be a driving factor for condensation and potentially play a role in extracellular release further work would be needed to decipher the mechanism of exit and also edited the schematic accordingly showing no rupture of the membrane.\n(Page No. 15, Line No. 442-444 and Fig 8 schematic)."}, {"round": 3, "reviewer_comments": {"reviewer_1": "This can be published now.", "reviewer_2": "The authors have addressed this reviewers' concerns adquately."}, "author_response": ""}]

Decision letter