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SARS-CoV-2 FP1 Destabilizes Lipid Membranes and Facilitates Pore Formation

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Submitted:

13 December 2024

Posted:

16 December 2024

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Abstract

SARS-CoV-2 viral entry requires membrane fusion, facilitated by fusion peptides within its spike protein. While these predominantly hydrophobic peptides insert into target membranes, their precise mechanistic role in membrane fusion remains incompletely understood. Here, we investigate how FP1, the N-terminal fusion peptide sequence, modulates membrane stability and barrier function across various model membrane systems. Through a complementary suite of biophysical techniques—including electrophysiology, fluorescence spectroscopy, and atomic force microscopy—we demonstrate that FP1 significantly promotes pore formation and alters membrane mechanical properties. Our findings reveal that FP1 reduces the energy barrier for membrane defect formation and stimulates the appearance of stable conducting pores, with effects modulated by membrane composition and mechanical stress. The observed membrane-destabilizing activity suggests that beyond its anchoring function, FP1 may facilitate viral fusion by locally disrupting membrane integrity. These results provide mechanistic insights into SARS-CoV-2 membrane fusion mechanisms and highlight the complex interplay between fusion peptides and target membranes during viral entry.

Keywords: 
lipid bilayer
;  
fusion peptide
;  
SARS-CoV-2
;  
AFM
;  
membrane pore
Subject: 
Physical Sciences  -   Biophysics
Preprints on COVID-19 and SARS-CoV-2

1. Introduction

Coronaviruses, including severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), belong to the family of enveloped viruses, characterized by a viral capsid encased in a lipid bilayer acquired during assembly through budding from host cell membranes [1]. The lipid envelope renders SARS-CoV-2 infection contingent upon membrane fusion—a fundamental process in which the viral envelope merges with cellular membranes to create a continuous lipid bilayer [2,3]. This critical mechanism enables viral capsid disassembly and the subsequent release of viral RNA into the host cell cytoplasm.
Membrane fusion, the merging of two lipid bilayer-bounded compartments, is an energetically demanding process requiring extensive local reorganization of bilayer structure, proceeding through the formation of distinct metastable, non-lamellar intermediates [4,5,6]. The initial step involves the formation of a stalk—a characteristic hourglass-shaped structure where proximal membrane leaflets merge while distal leaflets remain separate [7,8]. To complete membrane fusion and establish a continuous aqueous connection between the two compartments, the stalk must transform into a fusion pore [6,9,10,11]. Both these transitions require overcoming substantial energy barriers.
In SARS-CoV-2, this energetically intensive process is mediated by the spike (S) protein, a trimeric class I fusion protein composed of two functional subunits: the receptor-binding S1 and the fusion-mediating S2. The S1 subunit contains the receptor-binding domain (RBD), which specifically recognizes and binds to angiotensin-converting enzyme 2 (ACE2) on host cells. Meanwhile, the S2 subunit houses the fusion machinery, including the fusion peptide (FP) rich in hydrophobic amino acids, heptad repeat regions (HR1 and HR2), and the transmembrane domain [12,13].
The fusion competence of the S protein requires proteolytic processing at two distinct sites: the S1/S2 boundary and the S2’ site. While S1/S2 cleavage occurs during protein biosynthesis, the critical S2’ site, located immediately upstream of the FP, must be cleaved during the entry process [14]. The S2’ proteolysis occurs at Ser816 in SARS-CoV-2, forming a mature N-terminus of the fusion peptide [15]. The cleavage can occur at the plasma membrane via the transmembrane serine protease TMPRSS2 or within endosomes by cathepsin L, corresponding to the two distinct entry pathways available to SARS-CoV-2 [12,16,17].
Upon receptor engagement and subsequent S2’ proteolytic activation, S1 dissociation triggers extensive conformational changes in S2, resulting in a dramatic transformation from the prefusion spike to the rod-shaped post-fusion S2 subunit. This rearrangement is thought to transition through an intermediate state that exposes the hydrophobic FP, which can readily insert into the hydrophobic core of the target membrane when in proximity [18,19,20].
The final refolding of S2 into its post-fusion state involves the trimerization of HR1 domains and their subsequent assembly with HR2 domains into a six-helix bundle, with the FP serving as an anchor in the target membrane [15,18,21]. This structural transformation generates considerable force, pulling the membrane-anchored FPs toward the viral transmembrane domain and bringing the opposing membranes into close proximity [12,22]. This mechanical action overcomes hydration repulsion, thereby initiating the fusion cascade [23].
The fusion peptide consists of a 41-amino acid sequence that begins at Ser816 [24,25] and comprises approximately equal-length sections FP1 and FP2, located at the N- and C-termini, respectively. This sequence features motif that are highly conserved among coronaviruses [26,27], making it potential antigens for monoclonal antibodies targeting the fusion peptides across all human-infecting coronaviruses [28]. Such antibodies have the capacity to neutralize viruses by inhibiting their fusion with host cell membranes, highlighting the fusion peptide as a promising candidate epitope for next-generation coronavirus vaccine development.
Despite its therapeutic potential, the precise role of the fusion peptide in the membrane fusion process remains incompletely understood. While this peptide likely acts as a molecular ‘hook’ to anchor the viral machinery to the target membrane, it also functions as a membrane-modulating agent [24,29,30,31,32]. Evidence indicates that the isolated SARS-CoV-2 fusion peptide deeply penetrates lipid bilayers upon membrane interaction [25,30,33]. Nuclear magnetic resonance (NMR) studies have revealed that FP, which exists in a dynamically disordered state in aqueous environments, undergoes significant structural changes upon membrane insertion. Membrane interaction induces a conformational rearrangement characterized by two amphipathic helices in the N-terminal half (FP1) and a single polar helix in the C-terminal half (FP2) [25,33]. FP1 resides within the hydrophobic core of the lipid bilayer, while FP2 interacts with the polar lipid heads [24,25]. Furthermore, isolated FP1 and FP2 demonstrate similar behaviors when studied independently [30].
The precise influence of the SARS-CoV-2 fusion peptide on lipid order is complex and context-dependent, with conflicting results reported in the literature. Some studies have observed a decrease in lipid order and an increase in membrane fluidity upon FP interaction [29,30], while others have reported an increase in lipid order in specific membrane regions [24]. These discrepancies underscore the sensitivity of FP-induced membrane changes to experimental conditions, including pH, calcium concentration, and specific lipid composition [30,34]. Nevertheless, it is consistently observed that FP1 insertion significantly alters lipid packing in the vicinity of the insertion site. Recent studies indicate that this insertion involves the incorporation of polar and charged amino acids into the hydrophobic core, resulting in localized membrane narrowing [33]. This wedging effect, combined with the introduction of charged and polar moieties, can potentially induce spontaneous membrane poration, thereby compromising membrane barrier function.
In the canonical model of membrane fusion, the integrity of merging compartments is preserved, maintaining their volume identity relative to the surrounding environment [23]. However, emerging studies suggest a fusion pathway in which holes may form in one of the fusing membranes, evolving into a fusion pore, including those promoted by viral fusion proteins [35,36]. Experimental evidence indicates that a pore can sometimes develop during the early stages of viral fusion in one of the membranes [37]. Molecular dynamics simulations further corroborate that fusion can occur alongside leakage in these initial stages [38]. The formation of pores can inhibit fusion as well [39]; whether a pore aids or hinders membrane merging depends on the depth and geometry of FP insertion [39,40]. Thus, pore formation can facilitate fusion by serving as an alternative pathway for hemifusion as well as the transition from stalk formation to fusion pore development, although it may also inhibit fusion if the metastable pore possesses lower energy than the fusion intermediates.
To further elucidate these mechanisms, we investigated how FP1 peptide influences membrane stability and organization across three distinct lipid compositions: pure dioleoyl phosphatidylcholine (DOPC) bilayers, DOPC:cholesterol (Chol) mixtures, and a cellular membrane-mimicking composition (CMM). The CMM contained DOPC, Chol, dioleoyl phosphatidylserine (DOPS), and dioleoyl phosphatidylethanolamine (DOPE) in molar ratios of 35.7:35:8.3:20.9, respectively [41]. Using AFM force spectroscopy, we monitored membrane rupture dynamics [42,43], while electrophysiology measurements and calcein leakage assays revealed FP1-induced changes in membrane permeability [44,45]. Our findings demonstrate that FP1 significantly reduces the activation barrier for conductive defect formation in lipid bilayers, with defect size and dynamics strongly dependent on membrane composition. These results suggest a possible role for controlled membrane destabilization in viral fusion, where precisely regulated pore defect formation may facilitate the fusion process.

2. Results

2.1. Circular Dichroism Analysis of FP1 Peptide: Structural Changes and Membrane Interaction

We synthesized the FP1 peptide (SFIEDLLFNKVTLADAGFIK), corresponding to positions 816-835 of the SARS-CoV-2 spike protein S (Figure 1a). To verify the peptide’s ability to bind to lipid membranes and to assess changes in its secondary structure during this process, we measured the peptide’s circular dichroism (CD) spectrum in the UV range (195-250 nm). The resulting data were analyzed using the BeStSel web server to predict secondary structure composition and folding patterns [46].
Figure 1b shows CD spectra of FP1 measured under different environmental conditions, along with their corresponding BeStSel algorithm fits used to calculate peptide structure. In low calcium and low ionic strength conditions (10 mM PBS buffer, pH 7.0), FP1 exhibited a CD spectrum with a negative extremum at approximately 200 nm, characteristic of an unstructured state [47]. BeStSel analysis revealed minimal structuring, with only 8% alpha-helical content (Figure 1B, inset).
A markedly different dichroic signature was observed when FP1 was dissolved in methanol at the same bulk concentration (25 µM). The emergence of two negative extrema at 208 nm and 222 nm indicated substantial helical secondary structure formation [48]. BeStSel analysis confirmed significantly increased structuring in methanol compared to water, with alpha-helical content exceeding 55%. This enhanced structuring is consistent with the peptide’s high content of hydrophobic amino acid residues, which prefer the less polar methanol environment.
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Figure 1. (a) Primary sequence and helical wheel projection of FP1 peptide generated using HeliQuest software [49]. (b) Circular dichroism (CD) spectra of FP1 peptide (25 µM) under different conditions: in 10 mM PBS buffer, pH 7.4 (grey curve); in the presence of DOPC liposomes at peptide:lipid molar ratio 1:1000 (blue curve); in methanol (red curve). Dashed lines represent BeStSel fitting curves used to determine the α-helical content in each condition [46]. The resulting α-helical content values are shown in the inset.
Figure 1. (a) Primary sequence and helical wheel projection of FP1 peptide generated using HeliQuest software [49]. (b) Circular dichroism (CD) spectra of FP1 peptide (25 µM) under different conditions: in 10 mM PBS buffer, pH 7.4 (grey curve); in the presence of DOPC liposomes at peptide:lipid molar ratio 1:1000 (blue curve); in methanol (red curve). Dashed lines represent BeStSel fitting curves used to determine the α-helical content in each condition [46]. The resulting α-helical content values are shown in the inset.
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The CD spectrum of FP1 in phosphate buffer following 1h incubation with DOPC small unilamellar vesicles (SUVs, 100 nm diameter) at peptide:lipid molar ratio 1:1000 showed an intermediate profile. BeStSel analysis indicated an increase in alpha-helical content to 27%, confirming peptide-membrane interaction and insertion into the hydrophobic region of the lipid bilayer. Notably, the inclusion of either negatively charged DOPS or cholesterol in the SUV formulation had no considerable effect on the CD spectra (data not shown), suggesting that the peptide’s secondary structure upon membrane binding is primarily determined by its insertion into the hydrophobic core rather than specific lipid interactions. This structural reorganization upon membrane binding typically affects local membrane properties and may alter membrane permeability, which we investigate in subsequent sections.

2.2. FP1-Induced Membrane Permeabilization: Effects of Lipid Composition

To assess the impact of FP1 incorporation on lipid bilayer permeability, we employed a standard calcein leakage assay. SUVs with diameters of 100 nm and varying lipid compositions were prepared and loaded with 20 mM calcein, resulting in near-complete self-quenching. Membrane permeabilization leads to calcein release into the external solution, increasing total fluorescence due to dilution and subsequent dequenching of the die. We observed slow leakage kinetics in liposomes composed of DOPC. The addition of negatively charged DOPS did not significantly affect calcein leakage, implying that hydrophobic interactions predominantly govern FP1’s affinity for the membranes. In contrast, no calcein release was detected in SUVs containing cholesterol, suggesting that cholesterol reduces membrane disruption caused by FP1.
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Figure 2. Calcein leakage from small unilamellar vesicles (SUVs) of varying compositions, as indicated in the figure, induced by the addition of FP1 (25 µM) at time t = 0.
Figure 2. Calcein leakage from small unilamellar vesicles (SUVs) of varying compositions, as indicated in the figure, induced by the addition of FP1 (25 µM) at time t = 0.
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The observed membrane rupture likely results from structural distortions in the lipid bilayer associated with FP1 incorporation, indicating that FP1 enhances the likelihood of pore formation in the vesicle membrane—an effect mitigated by cholesterol. Given that cholesterol has minimal effect on membrane rigidity in DOPC bilayers [50], its protective role could arise through either indirect membrane stabilization or direct peptide interaction. The indirect mechanisms may include: (i) cholesterol’s negative spontaneous curvature counteracting local membrane deformations, and (ii) rapid cholesterol flip-flop between membrane leaflets compensating for the area imbalance between external and internal lipid leaflets created by FP1. Alternatively, cholesterol might directly modulate FP1 activity through changes in the membrane’s lateral pressure profile, affecting peptide penetration depth and orientation. To distinguish between these possibilities, we employed atomic force microscopy (AFM) to measure the force required to rupture supported lipid bilayers in the presence and absence of FP1.

2.3. Quantitative Analysis of FP1-Induced Membrane Destabilization via AFM Force Spectroscopy

In our study, we employed atomic force microscopy (AFM) to evaluate the impact of FP1 on the mechanical properties of supported lipid bilayers on atomically flat mica surfaces (Figure 3a, b). Using force spectroscopy, we recorded force-distance curves during membrane puncture to measure the bilayer’s response to applied stress. To assess the effect of cholesterol, we compared the responses of DOPC bilayers with those of CMM membranes, which serve as a model for cholesterol-containing membranes.
When the AFM tip applies a point force (F) to the membrane surface, the membrane initially exhibits linear-elastic behavior, with local deformation proportional to the applied force. However, as F increases, the induced stress eventually exceeds the activation barrier for critical defect formation, leading to sudden membrane rupture. This manifests as an abrupt cantilever displacement equal to the membrane thickness, h, at which membrane rupture occurs, as the tip contacts the underlying substrate.
To ensure consistent loading conditions, all measurements were performed at a fixed cantilever approach speed of 1 μm/s. The resulting force-displacement curves yield two critical parameters: breakthrough force, f, which reflects the energy barrier for critical defect formation, and h (Figure 3c). Prior to FP1 addition, DOPC and CMM bilayers exhibited mean breakthrough forces of <f₀> = 2.9 ± 0.2 nN and 5.5 ± 0.1 nN, with h = 4.7±0.3 nm and 4.0±0.3 nm, respectively. All values are in good agreement with the published ones for lipid bilayers, where similar conditions for breakthrough force measurement were employed [43,51].
Incubation with FP1 (2 μM) significantly altered membrane properties. The root mean square (RMS) surface roughness increased from 0.1 nm to 0.2 nm (Figure 3a, b), indicating successful peptide incorporation into the bilayer. Following FP1 insertion, both compositions showed comparable relative reductions in breakthrough force (Figure 3d, f). Interestingly, while DOPC bilayer showed a considerable reduction of thickness at which rupture occurs to h = 3.6±0.5, the threshold thickness of CMM bilayers remained unchanged (Figure 3e). Analysis of force-distance curves for DOPC bilayers before and after FP1 addition reveals bilayer thinning induced by FP1, as evidenced by the leftward shift of the linear force-displacement region, while FP1 insertion in CMM appears to have no considerable impact on its thickness.
Membrane rupture represents a stochastic process occurring through critical defect formation [52]. The probability of defect formation depends on its activation energy ∆ U , which is the function of applied force F. As the force increases linearly with time, t according to F = K v t , where K – is the cantilever spring constant, and v is the approach velocity. According to the molecular model developed in [53], the mean breakthrough force <f> exhibits a logarithmic dependence on loading rate:
f = α f T l n K v l n 2 k 0 α f T + 1
where fT – is a “thermal” force, k0 is the rate of spontaneous formation of a pore in a membrane when F=0, and α – is geometrical factor equal 0.5 when the cantilever tip radius (R) is much larger than the distance between adjacent lipid molecules. In this model, pressure increases the energy of lipid molecules within a finite activation volume V beneath the tip, thereby enhancing the probability of molecular displacement to adjacent pressureless membrane regions, which causes a pore. The thermal force f T = 4 π h R k B T V represents the force required to compress the activation volume by an amount equivalent to thermal energy kBT.
We investigated the impact of FP1 on the fundamental parameters of membrane stability by measuring breakthrough forces at multiple loading rates. By varying the cantilever approach speed from 0.1 to 3.0 μm/s, we obtained the dependence of breakthrough force on loading rate for DOPC bilayers before and after FP1 exposure. This allowed us to determine how FP1 affects both V and k₀. Figure 4a shows the obtained <f(v)> dependencies plotted on a logarithmic scale. Linear approximation of these curves, measured using cantilevers with spring constant K = 0.12 N/m and tip radius R = 10 nm, enabled determination of FP1-induced changes in k₀ and V (Figure 4b).
Following FP1 incorporation into the lipid bilayer, we observed a ~40% decrease in activation volume V and a significant increase in the rate of pore formation k₀, which rose by several orders of magnitude from k 0 = 3.3 · 10 − 5 Hz to   k 0 = 2.2 · 10 − 2 Hz once the fusion peptide has been incorporated in the lipid bilayer. Importantly, both V and k₀ values were in good agreement with published data [43,54], where measurements were conducted for similar lipid compositions under comparable conditions. Notably, taking into account the reduction in bilayer thickness caused by FP1, the calculated decrease in the area of the activation zone (S = V/h) is less than 25%. This suggests that the dramatic increase in the rate of spontaneous pore formation is the primary factor driving the observed reduction in breakthrough force.
Our AFM studies confirmed that FP1 significantly enhances the rate of spontaneous pore formation, consistent with a substantial reduction in the activation energy barrier for pore nucleation. To further explore the dynamics of FP1-induced membrane destabilization, we employed bilayer lipid membranes (BLMs), enabling real-time observation of pore formation and evolution. These complementary approaches underscore the central role of FP1 in inducing rapid structural disruption, further elucidating its membrane-destabilizing mechanisms.

2.4. Electrophysiological Characterization of FP1-Induced Membrane Defects

BLM experiments reveal FP1’s ability to induce conducting defects and pores in lipid bilayers under mechanical tension. All measurements were performed under physiological conditions (150 mM NaCl, low Ca²⁺, pH 7.0) with an applied voltage bias of 50 mV. FP1 was added at a concentration of 2 μM to one side of the BLM. We observed several distinct patterns of FP1-induced conducting events (Figure 4a): i) rupture: instant current increase due to membrane short-circuit; ii) spike: momentary charge passage through transient defects; iii) erratic: continuous current through dynamic membrane defects; iv) multi-level: multiple dynamic conducting defects; v) step: stable conducting defects or pores; and vi) channel-like: pores with defined diameter oscillating between open and closed states.
FP1 induced conducting defects more frequently in DOPC membranes compared to CMM composition, with CMM membranes exhibiting significantly more complex dynamic behavior (Figure 5b). While FP1 addition to DOPC membranes primarily led to spikes and erratic conductance increases culminating in membrane rupture, CMM membranes displayed stable conductance changes characterized by steps and channel-like behavior, suggesting the formation of metastable defects of specific sizes.
The stability difference is quantitatively reflected by event timing and membrane survival statistics. CMM bilayers maintained integrity for longer periods during FP1 exposure, with a mean time to first event of 10 ± 8 min compared to 4 ± 3 min for DOPC (p<0.05) (Figure 5c). Similarly, the mean time until complete rupture was extended for CMM (14 ± 12 min vs 9 ± 10 min), alongside a lower probability of membrane rupture during the observation period of 30 minutes (CMM: 57% vs DOPC: 74%).
Analysis of current jumps during pore formation (Step and Channel-like events, Figure 5a) provided insights into pore dimensions. Using cylindrical approximation for the forming pores, we found a bimodal size distribution in CMM membranes, with a prominent peak around 0.9 nm and a second population in the range of 1.4-2.3 nm (Figure 5d). These dimensions align with theoretical predictions for quasistable pore formation [55]. Notably, only CMM membranes demonstrated reproducible channel-like behavior with stable pores. In contrast, DOPC membranes exhibited gradual, long-term conductance increases, indicating the development of large, unstable conducting defects that rapidly progressed to complete rupture.

3. Discussion

Our results reveal that SARS-CoV-2 FP1 functions not merely as a membrane anchor but as an active membrane-modulating agent, with its effects finely regulated by lipid composition. Upon membrane binding, FP1 undergoes a significant structural reorganization, transitioning from an unstructured state in solution to a partially α-helical conformation, as demonstrated by our CD measurements (Figure 1b). This structural transition aligns with observations from previous studies using lysolipid micelles [56] and small unilamellar vesicles [24] of varying lipid compositions, suggesting that FP1’s interaction with membranes is primarily driven by hydrophobic rather than electrostatic forces. The high content of non-polar amino acid residues in FP1 facilitates its insertion into the hydrophobic core of the lipid bilayer. Furthermore, its partial α-helical formation in methanol (Figure 1b), a relatively non-polar solvent, underscores the importance of hydrophobic environments in driving FP1’s conformational changes. This behavior is consistent with the properties of other amphipathic peptides, which often undergo conformational changes upon membrane interaction [57].
We further demonstrated that FP1 insertion into unilamellar vesicles leads to membrane-destabilizing effects, compromising barrier function of the membranes. Calcein leakage assays showed that FP1 induces significant membrane permeabilization. This effect is consistent with earlier reports of FP1 activity [56] and has also been observed for other viral fusion peptides, such as those from influenza and HIV [58,59]. Notably, FP1-driven membrane rupturing has been linked to viral fusion catalysis [30], where it is proposed to represent an alternative molecular mechanism to hemifusion through the formation of a “π-shaped” structure observed for hemagglutinin-induced membrane fusion [36,60]. Unlike the canonical stalk-hemifusion mechanism, the “π-shaped” structure involves transient pore formation within the host membrane as an intermediate step, potentially facilitating membrane merging by increasing connectivity between proximal monolayers (Figure 6 a).
AFM force spectroscopy provided quantitative insights into the mechanical changes induced by FP1 in supported lipid bilayers. Our measurements directly demonstrated that FP1 significantly reduces the point force required to pierce the supported lipid bilayers on mica. According to the molecular model developed in [53], when breakdown conditions (tip radius R, cantilever spring constant K, and approaching velocity v) are maintained constant, a reduction in breakthrough force (Eq.1) indicates either a decrease in thermal force, fT or an increase in the rate of critical defect spontaneous formation, k0. Our data convincingly demonstrated that the increase in k0 is responsible for the decrease in membrane piercing strength, implying a substantial reduction in the activation barrier for critical pore formation. This finding is consistent with our observations of increased membrane permeabilization in calcein leakage experiments.
This mechanism aligns with studies on other amphipathic peptides, such as magainin H2 [61] and the M2AH peptide from influenza A M2 protein [62], which also showed reduced rupture forces upon membrane interaction, indicating weakened mechanical stability. In the case of the HIV-1 fusion inhibitor T-1249 [63], a significant reduction in rupture force was observed, accompanied by increased membrane fluidity and roughness, further supporting a shared mechanism of membrane destabilization among amphipathic peptides, including FP1.
While rich with cholesterol CMM bilayer exhibited higher overall breakthrough forces, consistent with their enhanced mechanical stability, FP1 incorporation still led to a significantly much stronger relative reduction in force than for DOPC (Figure 3f). This observation suggests that FP1 lowers the activation barrier for pore formation even in cholesterol-rich membranes, albeit to a lesser absolute value than in pure DOPC bilayers. This interpretation is supported by our planar membrane experiments, where we observed conductive defects in cholesterol-containing bilayers despite their increased stability. Thus, while cholesterol enhances overall membrane integrity, it does not prevent FP1-induced destabilization but instead delays and modulates the process. The diversity in rupture force distributions observed via AFM further suggests distinct mechanisms of disruption, including the potential formation of metastable pores in cholesterol-containing systems.
Importantly, FP1 may also catalyze fusion through an alternative mechanism involving fusion pore formation in the vicinity of hemifusion structures (Figure 6b). Fusion involves a progression from stalk formation to hemifusion and, ultimately, to the formation of a full fusion pore. Hemifusion structure is a high-energy intermediate displaying significant elastic stress in the lipid bilayer, particularly in the hemifusion diaphragm, trilamellar rim and its nearby regions. The geometric constraints of the post-fusion state place FP1 in precisely this stressed environment, allowing it to interact with and destabilize the highly strained lipids in this region (Figure 6b). We speculate that cholesterol may play a pivotal role in regulating the threshold tension or elastic stress at which membrane destabilization could occur, facilitating pore formation at late stages when the hemifusion structure is already formed. This regulation helps ensure that under conditions of high elastic stress—when FP1 is likely situated nearby due to geometric constraints—membrane destabilization and pore formation can proceed efficiently.
Our findings suggest that FP1 induces localized structural defects that are particularly effective in areas subject to high elastic stress, such as those surrounding hemifusion diaphragms or stalks, thereby facilitating the emergence of metastable pores that bridge the final stage of viral fusion. This mechanism aligns with our observations of FP1 disrupting membranes under tension, such as in planar bilayers. The high lateral tension in these bilayers mimics the elastic strain present in hemifusion diaphragms during the fusion process. Conductive defects observed in cholesterol-containing planar membranes further suggest that FP1-mediated destabilization persists even in challenging membrane environments, highlighting its functional versatility in catalyzing fusion.
The apparent discrepancy between the cholesterol-mediated protection against calcein leakage in vesicle experiments and the observed membrane destabilization in planar bilayers and the reduction of rupture forces by FP1 might be explained by the different membrane conditions. The ability of cholesterol to rapidly flip-flop between monolayers and mitigate the elastic stress associated with FP1 incorporation may be less effective in planar bilayers, as well as in the hemifusion structure, where lateral tension is set by external reservoir of lipid molecules [64]. In vesicles, where number of lipid molecules is fixed the rapid cholesterol redistribution between monolayers prevents significant area asymmetry and the buildup of elastic stress, leading to greater resistance against FP1 induced rupture.

4. Materials and Methods

Materials. 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-sn-glycero-3-phospho-L-serine (DOPS) and cholesterol were purchased from Avanti Polar Lipids (Alabaster/AL, US), stored at -80 °C and used without further purification. Calcein disodium salt was from Serva (Germany). Phosphate buffer saline (PBS) tablets from Sigma-Aldrich were used to produce 1x PBS solutions containing 2.7 mM KCl, 137 mM NaCl, pH = 7.40 in 200 ml of distilled water.
Fusion peptide (FP1) synthesis. The SFIEDLLFNKVTLADAGFIK peptide was synthesized via solid-phase peptide synthesis using a PurePep® Chorus automated peptide synthesizer (Protein Technologies, Inc.). The synthesis employed N-(9-Fluorenylmethoxycarbonyl)-L-isoleucine 2-Chlorotritul ester resin as the solid phase and Fmoc-protected amino acid derivatives. Amino acid coupling was performed using N,N’-Diisopropylcarbodiimide (DIC) with Ethyl cyano(hydroxyimino)acetate (Oxyma Pure) as an activator, using a 15-fold excess of Fmoc-amino acids relative to carrier capacity.
Fmoc deprotection was achieved using 20% 4-methylpiperidine in N,N-dimethylformamide. After each coupling step, a capping procedure was performed using 5% propionic anhydride in DMF for 10 minutes. All coupling, capping, and Fmoc-deprotection steps were conducted at 65°С.
The peptide was cleaved from the resin and simultaneously deprotected using a mixture of trifluoroacetic acid:3,6-dioxa-1,8-octandithiol:triisopropylsilane:anisole:water (183:5:2:5:5 v/v) for 3 hours (5 mL per 100 mg resin). The filtered solution was precipitated with cold diethyl ether, cooled at -20˚C for 30 minutes, and centrifuged (7000 rpm, 10 minutes). The precipitate was washed twice with diethyl ether (2 × 25 mL), dissolved in 1 mL of 5% acetonitrile, and lyophilized.
The crude peptide was purified by HPLC using a gradient of 2-100% acetonitrile in water over 25 minutes (retention time: 12.5 min). Final yield: 25 mg (31%). MALDI-MS analysis: calculated m/z = 2240.2094, found m/z = 2241.2153.
Small unilamellar vesicle (SUV) preparation. Small unilamellar vesicles (SUVs) were prepared using a combination of sonication and extrusion methods. Lipids (compositions detailed in Table 1) dissolved in chloroform were mixed at the desired molar ratios. The organic solvent was removed under an argon stream followed by vacuum drying for 1 hour. The resulting lipid films were hydrated with buffer solutions (specified in subsequent experimental sections), subjected to 10 freeze-thaw cycles (liquid nitrogen and hot water bath), and then processed further.
For sonication (used in AFM experiments), suspensions were sonicated for 3 minutes (10 seconds on, 10 seconds off) at 50% power using a probe-type sonicator. For extrusion (used in calcein leakage and circular dichroism experiments), suspensions were extruded 19 times through 200 nm polycarbonate membranes (Millipore, Darmstadt, Germany) using a mini extruder (Avanti, Alabaster, US) after the freeze-thaw cycles.
Planar lipid bilayers. Microfluidic slides coated with Teflon film from Ionovation (Compact, Osnabrück, Germany) were employed for the formation of bilayer lipid membranes (BLMs). The lipid mixtures used for the bilayers were prepared in n-decane at a total concentration of 15 mg/mL. BLMs were created using the painting technique with flexible plastic capillaries made from pipette tips, allowing for careful maintenance of the aperture within the Teflon film without damage.
To replace the buffer in the chamber, 20 µL of a new solution was pipetted into the chamber simultaneously while aspirating 20 µL of the previous buffer. This process was repeated ten times, effectively replacing the buffer in the 100 µL compartments with excess new solution. The buffer composition for the BLM assembly consisted of 5 mM HEPES, 150 mM KCl, and 2 mM EDTA, adjusted to a pH of 7.40.
The criteria for initiating membrane work included the presence of a sharp meniscus-membrane border and a membrane capacitance of at least 40 pF. The HEKA EPC10USB amplifier was employed to measure membrane capacitance, apply a constant potential of 50 mV across the membrane, and record the resulting current. Ag/AgCl electrodes (Ionovation GmbH) with 2 M KCl/agarose bridges (1.5% w/v agarose) were used to apply the transmembrane voltage. Data were sampled at a frequency of 10 kHz, with a low-pass filter cutoff set at 5 kHz. The gain setting was configured to 0.2 mV/nA.
Measurements were conducted using a custom HEKA Patchmaster software protocol, which recorded one-minute traces of conductance interspersed with brief capacitance and current measurements. This procedure enabled periodic self-checks to rule out scenarios of membrane overflow (if capacitance became too low) or membrane disruption (if the circuit was shorted). Pore diameters were estimated by modeling the pores as cylindrical electrolytic conductors with a length equal to the bilayer thickness (4 nm) and a radius (r). The specific resistance (ρ) of the buffer solution was utilized to calculate resistance R = ρ h π r 2 + ρ 2 r applying the observed pore current with a known applied potential.
Calcein leakage assay. Calcein was dissolved in phosphate-buffered saline (PBS; 10 mM phosphate buffer, 2.7 mM KCl, 137 mM NaCl, pH 7.40; PBS tablets from Sigma) at a self-quenching concentration of 40 mM. Extruded liposomes were then separated from unencapsulated calcein using gel permeation chromatography on a Sephadex G-75 column with PBS as the elution buffer. Fluorescence measurements were conducted using a Hitachi F-2500 spectrofluorometer (Japan) in time-scan mode, with an excitation wavelength of 495 nm and emission recorded at 510 nm. A baseline fluorescence (I₀) was established by recording fluorescence for 1 minute with the liposome sample, followed by the addition of the peptide, after which the fluorescence (Iₜ) was recorded. To determine maximum fluorescence (Iₘₐₓ), 10 µL of 20% (w/w) Triton X-100 in water was added, and the sample fluorescence was recorded for an additional 1 minute. After the addition of each component, the sample was thoroughly mixed in the cuvette by gently pipetting half of the sample volume back and forth at least 10 times. The percentage of calcein leakage was calculated using the equation:
L e a k a g e % = I m a x − I t I m a x − I 0
Circular dichroism. Circular dichroism (CD) measurements were performed using a Jasco J-1500 CD spectrometer with quartz cuvettes of 2 mm path length. The spectra were recorded in the wavelength range of 190–250 nm. PBS was used as the buffer. For measurements involving peptides in the presence of liposomes, the peptide was added to liposomes suspended in PBS, and the sample was incubated for 60 minutes while being vortexed to ensure complete redistribution of the peptide into the lipid bilayers prior to analysis.
Atomic Force Microscopy (AFM) Imaging and Force Spectroscopy. Lipid bilayers were prepared on freshly cleaved mica sheets using the liposome fusion method. To create the liquid chamber, a circular section was removed from the center of the bottom of a Petri dish, and a coverslip was glued to the underside. A small piece of mica (0.6 cm x 0.6 cm) was adhered to the center of the coverslip, and its surface layer was freshly cleaved using adhesive tape immediately before depositing the sonicated liposomes onto the mica.
AFM measurements were performed using an NTEGRA Prima atomic force microscope (Russia). For all experiments, silicon cantilevers (CSG-10) with a nominal spring constant of 0.12 N/m were used and calibrated prior to measurements using the thermal noise method available in the software. Sonicated liposomes, at a concentration of 0.5 mg/mL, were applied to the cleaved mica surface and incubated for 30 minutes to form the lipid bilayer. After incubation, excess liposomes were removed by flushing the surface and gently washing off unbound vesicles. Before data acquisition, the system was equilibrated until the deflection signal stabilized.
Force spectroscopy measurements were performed to confirm bilayer formation using a fixed loading rate of 1 µm/s. For dynamic force spectroscopy experiments, the loading rate was varied between 0.1 and 3 µm/s. AFM imaging was carried out in tapping mode with a scanning rate of 1 Hz. All experiments were conducted in a buffer solution containing 150 mM KCl, 5 mM HEPES, at pH 7.4.
Data analysis. AFM images were treated with Gwyddion software. The force-distance curves were analyzed with Image Analysis. The data were plotted with Origin Lab.

5. Conclusions

In conclusion, the combination of AFM force spectroscopy, calcein leakage assays, and planar bilayer experiments provides a comprehensive picture of FP1-mediated membrane destabilization. Our findings not only demonstrate a reduction in the activation barrier for pore formation in DOPC membranes, but also show that this mechanism extends to cholesterol-containing membranes, where FP1 facilitates intermediate pore formation and fusion pore catalysis. The similarities observed between FP1 and other amphipathic peptides suggest shared membrane-destabilizing mechanisms. Furthermore, by potentially regulating the threshold tension or elastic stress for membrane destabilization, cholesterol may play a critical role in ensuring successful viral entry at the appropriate stages of fusion. This underscores FP1’s critical versatility in viral fusion processes and presents opportunities for therapeutic intervention by targeting fusion peptide-membrane interactions.

Author Contributions

Conceptualization, M.S., R.P. and P.B.; methodology, M.S. and R.P.; validation, M.S., R.P., R.M. and P.B.; formal analysis, M.S., R.P., T.L. and E.V.; investigation, M.S., R.P., T.L. and E.V; resources, G.K., O.F. and P.B.; data curation, M.S., R.P., T.L. and E.V.; writing—original draft preparation, M.S., R.P., R.M. and P.B.; writing—review and editing, M.S., R.P., R.M. and P.B.; visualization, M.S. and R.P.; supervision, P.B.; project administration, P.B.; funding acquisition, P.B. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by Russian Science Foundation, grant number 22-15-00265.

Data Availability Statement

The original contributions presented in this study are included in the article/supplementary material. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 3. AFM analysis of FP1 effects on breakthrough force of lipid bilayers (a) Height image topography of lipid bilayer without FP1 and (b) after FP1 incubation with corresponding height profiles along the navy lines. Scale bar: 500 nm. (c) Representative approach force-distance curves obtained for the studied lipid bilayers before and after FP1 addition. (d) Rupture force distribution histograms for lipid bilayers before and after FP1 incubation. (e) Relative rupture force of DOPC and CMM lipid bilayers after FP1 exposure, normalized to untreated bilayers. (f) ox-plot comparison of rupture distances for studied lipid bilayers before and after FP1 incubation. Statistical significance: # – p<0.005; *** – p<0.001.
Figure 3. AFM analysis of FP1 effects on breakthrough force of lipid bilayers (a) Height image topography of lipid bilayer without FP1 and (b) after FP1 incubation with corresponding height profiles along the navy lines. Scale bar: 500 nm. (c) Representative approach force-distance curves obtained for the studied lipid bilayers before and after FP1 addition. (d) Rupture force distribution histograms for lipid bilayers before and after FP1 incubation. (e) Relative rupture force of DOPC and CMM lipid bilayers after FP1 exposure, normalized to untreated bilayers. (f) ox-plot comparison of rupture distances for studied lipid bilayers before and after FP1 incubation. Statistical significance: # – p<0.005; *** – p<0.001.
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Figure 4. Influence of FP1 on Membrane Defect Formation (a) Dependence of the mean rupture force on the loading rate and (b) Rate of spontaneous pore formation (k₀) and activation volume (V) for pure DOPC bilayers versus DOPC bilayers exposed to FP1.
Figure 4. Influence of FP1 on Membrane Defect Formation (a) Dependence of the mean rupture force on the loading rate and (b) Rate of spontaneous pore formation (k₀) and activation volume (V) for pure DOPC bilayers versus DOPC bilayers exposed to FP1.
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Figure 5. FP1-induced conductance patterns and membrane defects (a) Representative current traces showing characteristic defect types induced by FP1 during planar bilayer lipid membrane (BLM) permeation. (b) Average time until the first conductance event caused by FP1 in DOPC and CMM membranes, showing significantly longer latency in CMM membranes (* p<0.05). (c) Distribution of membrane defect types present in traces for DOPC and CMM membranes. (d) Pore size distribution calculated from step and channel-like current patterns obtained with CMM membranes. Normal distribution peaks correspond to 0.9 and 1.6 nm.
Figure 5. FP1-induced conductance patterns and membrane defects (a) Representative current traces showing characteristic defect types induced by FP1 during planar bilayer lipid membrane (BLM) permeation. (b) Average time until the first conductance event caused by FP1 in DOPC and CMM membranes, showing significantly longer latency in CMM membranes (* p<0.05). (c) Distribution of membrane defect types present in traces for DOPC and CMM membranes. (d) Pore size distribution calculated from step and channel-like current patterns obtained with CMM membranes. Normal distribution peaks correspond to 0.9 and 1.6 nm.
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Figure 6. Mechanistic illustration of FP1-mediated membrane perturbation and its possible role in viral fusion (a) catalysis of hemifusion diaphragm formation: (i) FP1 inserts into the target membrane, locally perturbing its structure. (ii) This perturbation induces rupture of the target membrane, leading to local destabilization. (iii) Membrane reconnection between the viral and target membranes subsequently forms a hemifusion diaphragm. (b) catalyzing fusion pore formation: (i) FP1, positioned laterally to the hemifusion diaphragm, further destabilizes the surrounding lipid bilayer, concentrating its effect at the trilamellar junction. (ii) This destabilization triggers pore formation at the junction. (iii) The hemifusion diaphragm collapses into the membrane, resulting in the formation of a fusion pore.
Figure 6. Mechanistic illustration of FP1-mediated membrane perturbation and its possible role in viral fusion (a) catalysis of hemifusion diaphragm formation: (i) FP1 inserts into the target membrane, locally perturbing its structure. (ii) This perturbation induces rupture of the target membrane, leading to local destabilization. (iii) Membrane reconnection between the viral and target membranes subsequently forms a hemifusion diaphragm. (b) catalyzing fusion pore formation: (i) FP1, positioned laterally to the hemifusion diaphragm, further destabilizes the surrounding lipid bilayer, concentrating its effect at the trilamellar junction. (ii) This destabilization triggers pore formation at the junction. (iii) The hemifusion diaphragm collapses into the membrane, resulting in the formation of a fusion pore.
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Table 1. Lipid compositions used in this work.
Table 1. Lipid compositions used in this work.
Composition Molar ratios Abbreviation
DOPC 100 -
DOPC-Chol 70-30 -
DOPC-DOPS 90-10 -
DOPC:Chol:DOPE:DOPS 35.7:35:20.9:8.3 [41] CMM
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Copyright: This open access article is published under a Creative Commons CC BY 4.0 license, which permit the free download, distribution, and reuse, provided that the author and preprint are cited in any reuse.

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