Biomimetic Construction of Abiotic Membranes Through Acylation of Cationic Diamino Acids

TL;DR

Scientists created a new type of artificial membrane using simple chemical steps that mimic natural processes, enabling the formation of functional vesicles for synthetic biology.

Contribution

A modular and simple method to synthesize abiotic phospholipid analogs with choline-like cations on diamino acid backbones.

Findings

APAs self-assemble into vesicles under various conditions.

The method allows tuning of headgroup charge and composition.

APAs mimic key features of natural phospholipids.

Abstract

Compartmentalization is a defining feature of living systems and a cornerstone of bottom‐up synthetic biology. Although phospholipids dominate modern biological membranes, their de novo synthesis in the laboratory remains chemically demanding and offers limited headgroup diversity, thereby constraining the functional scope of artificial membranes. Here, we describe a simple and modular strategy to access choline‐mimetic abiotic phospholipid analogs (APAs) that recapitulate key structural and functional features of natural phospholipids. Diamino acids serve as minimal scaffolds onto which a trimethylammonium (TMA) headgroup is introduced in a single step through carboxylate conjugation with Girard's Reagent T (GRT), generating a permanent choline‐like cation. Subsequent N‐acylation under mild aqueous conditions with prebiotically relevant thioesters yields diacylated amino GRT (DAAG) amphiphiles with a classical phospholipid topology. These APAs spontaneously self‐assemble into well‐defined vesicular structures and allow systematic tuning of headgroup charge and composition. Our approach facilitates access to chemically versatile, TMA‐functionalized membranes that expand the toolkit for constructing functional artificial membranes in synthetic biology, protocell research, and molecular delivery.

Cells produce membrane‐forming phospholipids through complex enzymatic pathways. Here, we present the in situ synthesis of a novel class of abiotic phospholipid analogs (APAs) featuring a permanent choline‐mimetic cation on diamino acid backbones. These APAs spontaneously self‐assemble into robust vesicles across a broad range of conditions.© 2026 WILEY‐VCH GmbH

Figures

In situ formation of cationic phospholipid‐like membranes. (A) Chemical structures of diacylated amino acid (DAAA) and diacylated amino GRT (DAAG) amphiphiles, APAs that resemble natural phospholipids [DOPC: 1,2‐dioleoyl‐sn‐glycero‐3‐phosphocholine; DPPC: 1,2‐dipalmitoyl‐sn‐glycero‐3‐phosphocholine]. (B) Schematic representation of spontaneous vesicle assembly driven by in situ synthesis of DAAGs (DOAG and DPAG). The approach exploits dual imidazole‐promoted aminolysis ligation between a TMA‐containing diamino acid (1b) and an acyl thioester (oleoyl thioester 2 or 5 for DOAG, and palmitoyl thioester 3 for DPAG).

Synthesis of DOAG by dual imidazole‐promoted aminolysis ligation between 1b and MESNA oleoyl thioester 2. (A) Formation of DOAG at various time points (0, 15, 30, 45, 60, 75, 90, 105, and 120 min). HPLC (210 nm) traces monitoring the dual aminolysis reaction between 25 mM 1b (1 equiv) and 50 mM oleoyl thioester 2 (2 equiv) in the presence of imidazole (1.5 M). Retention times (tR) were verified by mass spectrometry. (B) Kinetic plot of DOAG formation during dual N‐acylation. Absorption at 210 nm was monitored, and the area under the peak for DOAG was plotted over time.

Characterization of DOAG vesicular structures. (A) Phase contrast microscopy images of DOAG vesicles. Scale bar denotes 10 µm. (B) Fluorescence microscopy image of DOAG vesicles stained with a 0.1 mol% of Texas Red DHPE. Scale bar denotes 10 µm. (C) Particle size distribution (left) determined by dynamic light scattering (DLS) and zeta potential values (right) obtained by electrophoretic light scattering (ELS) of DOAG nanoparticles generated using the microfluidic‐based platform TAMARA. The bar graph represents the mean particle size, while the green dot corresponds to the zeta potential. Error bars denote standard deviation from three independent measurements (n = 3). (D) Fluorescence microscopy images of DOAG vesicles encapsulating HPTS. Scale bar denotes 10 µm.

Biocompatibility of DOAG vesicular structures. (A) Phase‐contrast microscopy images showing the stability of DOAG vesicles in 1X PBS (pH 7.4) supplemented with varying concentrations of FBS (1, 10, and 50% v/v) after 2 and 24 h. (B) Viability of HEK 293T cells exposed to DOAG vesicles. The bar graph shows the mean viability values for each tested concentration. Error bars denote standard deviation from three independent measurements (n = 3). No statistically significant differences were observed between groups (one‐way ANOVA followed by Tukey's multiple comparisons test, p > 0.05).

Full text

Introduction

1

Compartmentalization is a defining feature of living systems and is widely considered to have played a critical role in the transition from inanimate matter to life [1]. Although a variety of physicochemical mechanisms can drive the de novo emergence of compartmentalized structures [2], phospholipids are particularly relevant candidates, as they are key constituents of evolved biological membranes and hold promise for the development of advanced technologies [3, 4]. In biological systems, phospholipids are synthesized through the enzymatic diacylation of glycerol‐3‐phosphate with acyl‐CoA‐activated fatty acids, producing key intermediates that undergo further modification to generate the lipid diversity characteristic of mature membranes [5, 6, 7]. In contrast, the de novo synthesis of phospholipids in the laboratory remains challenging, often resulting in low yields and the accumulation of monoacylated byproducts [8, 9, 10, 11, 12]. Moreover, the limited chemical diversity of canonical lipid headgroups imposes additional synthetic constraints and restricts the functional versatility of model membranes, particularly in applications requiring specific interactions or responsive behavior. Together, these challenges underscore the need for alternative strategies to streamline the preparation and assembly of synthetic amphiphilic molecules.

Structurally, phospholipid membranes are built on a simple principle: two hydrophobic chains linked to a polar headgroup are sufficient to drive the formation of closed compartments (Figure 1A) [13]. This minimal architecture underpins essential membrane functions, including permeability regulation and the spatial organization of biochemical reactions [14]. Reconstructing this logic using synthetic molecules has become a powerful strategy for probing membrane function and generating new mimetics with predictable behavior [15, 16, 17]. In this context, amino acid‐derived amphiphiles have emerged as appealing alternatives to canonical lipids [18]. They are synthetically accessible, modular, and inherently programmable [11, 19]. Within this family, diamino acids represent particularly compelling scaffolds: their two amine groups enable the formation of hydrophobic domains through acylation, while the carboxyl group can serve as a polar headgroup. For instance, we have recently demonstrated that diacylated amino acids (DAAAs) self‐assemble into vesicular structures capable of encapsulating small molecules, supporting their use in the bottom‐up construction of artificial membranes (Figure 1A) [20]. However, these systems remain at an early stage as mimics of natural bilayers, and the chemical versatility imparted by the carboxylate headgroups has yet to be exploited to recreate key features of canonical phospholipids and to enable new applications in synthetic biology.

In situ formation of cationic phospholipid‐like membranes. (A) Chemical structures of diacylated amino acid (DAAA) and diacylated amino GRT (DAAG) amphiphiles, APAs that resemble natural phospholipids [DOPC: 1,2‐dioleoyl‐sn‐glycero‐3‐phosphocholine; DPPC: 1,2‐dipalmitoyl‐sn‐glycero‐3‐phosphocholine]. (B) Schematic representation of spontaneous vesicle assembly driven by in situ synthesis of DAAGs (DOAG and DPAG). The approach exploits dual imidazole‐promoted aminolysis ligation between a TMA‐containing diamino acid (1b) and an acyl thioester (oleoyl thioester 2 or 5 for DOAG, and palmitoyl thioester 3 for DPAG).

A key challenge in designing abiotic phospholipid analogs (APAs) is the incorporation of a trimethylammonium (TMA) headgroup, as found in choline (Figure 1A). First described by Adolf Strecker in 1862, choline serves as the biosynthetic precursor of phosphatidylcholine – the major phospholipid in eukaryotic cell membranes – via the Kennedy pathway [21]. The permanent positive charge of the TMA moiety plays a central role in membrane architecture, influencing lipid packing, hydration, and interactions with proteins and anionic biomolecules [22]. Beyond structural significance, TMA‐functionalized lipids are widely used as cationic vectors in biomedical applications, particularly in nucleic acid delivery, where strong electrostatic interactions enhance cellular uptake and endosomal escape [23, 24]. Despite this functional importance, synthetic access to TMA‐bearing phospholipids remains limited. Current strategies typically require multistep procedures or harsh conditions, which constrain headgroup diversity and hinder the development of chemically versatile analogs suitable for synthetic biology and therapeutic applications [24, 25, 26].

Here, we present a straightforward, one‐step strategy to access TMA‐containing APAs, in which the cationic functionality is introduced via the carboxylate group of a diamino acid using Girard's Reagent T (GRT) (Figure 1, Scheme S1). Widely employed in bioconjugation and mass spectrometry workflows, GRT provides a preinstalled TMA moiety that mimics choline [27]. The amino acid's diamine functionality subsequently enables efficient dual N‐acylation in aqueous media using prebiotically relevant thioesters as acyl donors (Figure 1B) [28]. This approach yields a family of diacylated amino GRT (DAAG) amphiphilic molecules that preserve the classical phospholipid topology while offering a simple, modular, and water‐compatible synthetic route (Figure 1). The resulting APAs self‐assemble into well‐defined vesicles, exhibit characteristic aggregation behavior, and provide a versatile platform for tuning headgroup charge and composition without perturbing the hydrophobic core.

Results and Discussion

2

Preparation of TMA‐Containing Diamino Acid Head Groups

2.1

Diaminocarboxylic acids are widely distributed in nature. Although they are nonproteinogenic, these compounds are of particular interest in origins‐of‐life research, as several members of this family have been identified in the Murchison meteorite, prompting hypotheses regarding their role as exogenous prebiotic building blocks in early chemical evolution [29, 30]. Among them, L‐2,3‐diaminopropionic acid (L‐Dap‐OH) was selected as the central scaffold for the design of the proposed DAAGs, owing to its ability to mimic the backbone geometry of sn‐glycerol‐3‐phosphate (Figure 1A). To prevent undesired inter‐ and intramolecular reactions between the free amino groups and the carboxylic acid, L‐Dap‐OH was initially employed in its dual N‐Boc‐protected form [Boc‐Dap(Boc)‐OH]. This protected amino acid was activated with 2‐(7‐Aza‐1H‐benzotriazole‐1‐yl)−1,1,3,3‐tetramethyluronium hexafluorophosphate (HATU) in the presence of N, N‐diisopropylethylamine (DIPEA), and subsequently reacted with GRT to afford its TMA‐functionalized version [Boc‐L‐Dap(Boc)‐GRT, 1a] (Scheme S1). The resulting compound was analyzed by high‐performance liquid chromatography (HPLC) and fully characterized by ^1^H‐ and ^13^C‐nuclear magnetic resonance (NMR) spectroscopy and high‐resolution mass spectrometry (HRMS) (see Supporting Information). After purification, deprotection under acidic conditions yielded the free L‐Dap‐GRT (1b) (Scheme S1), which was used directly without further purification.

Dual N‐Acylation of TMA‐Functionalized Diamino Acids with Acyl Thioesters

2.2

Direct aminolysis was selected as the acylation pathway for L‐Dap‐GRT (1b) (Figure 1B). Despite its inherently slow kinetics, imidazole‐mediated direct aminolysis has proven to be a powerful tool in synthetic biology [28]. Notably, it has been applied to the acylation of both amino acids and glycerolphosphocholines, yielding amphiphilic structures that self‐assemble in situ into membrane‐based synthetic cells [20, 28]. Thioesters were chosen as N‐acyl donors because they are widely recognized to have played key roles in prebiotic biochemistry and constitute a mimicking platform for natural phospholipid synthesis [31, 32, 33]. To evaluate the effectiveness of this strategy for acylating the TMA‐functionalized diamino acid headgroup, the reactivity of 1b toward water‐soluble thioesters (2, 3, and 5) was investigated. Thioesters bearing different leaving groups (MESNA vs. thiocholine), as well as fatty acyl chains with varying degrees of length and unsaturation (palmitoyl vs. oleoyl), were examined (Figure 1B). Initially, 1b was reacted with oleoyl sodium 2‐mercaptoethanesulfonate (MESNA) (2) in aqueous imidazole (Figure 2A, Scheme S6). The formation of DOAG was monitored over time using HPLC (Figures 2, S1), and the final product was characterized by ^1^H‐ and ^13^C‐NMR spectroscopy and HRMS (see Supporting Information, and Figure S1). Reaction progress was followed by liquid chromatography for up to 120 min, at which point DOAG formation reached a plateau (Figure 2B). Interestingly, the monoacylated intermediate (MOAG) was not detected during the reaction (Figure 2, Scheme S6). We attribute this to hydrophobic interactions between the alkyl chains of MOAG and 2, which likely bring the molecules into close proximity, possibly within mixed micelles, facilitating rapid second acylation. Consistent with previous findings [10, 20], our data suggest that MOAG formation constitutes the rate‐limiting step, as no detectable monoacylated intermediates are observed at any time point, and once formed, MOAG is rapidly converted into DOAG. The robustness of the approach was further evaluated by reacting 1b with a palmitoyl thioester bearing MESNA as the leaving group (3), which afforded the corresponding DPAG with high conversion, as evidenced by HPLC analysis of the crude reaction, in which DPAG was the predominant species (Scheme S7, Figure S1). Notably, replacing MESNA with choline (5) as the leaving group (Scheme S8) resulted in altered reaction kinetics of DOAG and an ≈11% decrease in conversion (53% yield when using MESNA versus 42% yield when using thiocholine). We attribute this effect to electrostatic repulsion between the positively charged choline‐containing micelles of 5 (zeta potential = +52 ± 4 mV) (Figure S2) and the TMA functionality of 1b. In contrast, reactions involving MESNA are expected to be favored due to the lower steric hindrance of the leaving group and attractive electrostatic interactions between the negatively charged sulfonate groups of MESNA thiooleate (2) micelles (zeta potential = −53 ± 2 mV) (Figure S2) and the positive charge of 1b.

Synthesis of DOAG by dual imidazole‐promoted aminolysis ligation between 1b and MESNA oleoyl thioester 2. (A) Formation of DOAG at various time points (0, 15, 30, 45, 60, 75, 90, 105, and 120 min). HPLC (210 nm) traces monitoring the dual aminolysis reaction between 25 mM 1b (1 equiv) and 50 mM oleoyl thioester 2 (2 equiv) in the presence of imidazole (1.5 M). Retention times (tR) were verified by mass spectrometry. (B) Kinetic plot of DOAG formation during dual N‐acylation. Absorption at 210 nm was monitored, and the area under the peak for DOAG was plotted over time.

Characterization of the DAAG Vesicular Structures

2.3

The ability of both DOAG and DPAG to self‐assemble into artificial membranes in aqueous environments was assessed by microscopy studies on purified DAAG samples hydrated at 37°C in Milli‐Q water containing 5 mM of the corresponding amphiphile (Figures 3, S3). DOAG readily formed vesicles, as observed by phase‐contrast microscopy (Figure 3A). In contrast, hydration of DPAG under identical conditions resulted in irregular aggregation, giving rise to fibrous and amorphous structures (Figure S3A). This lack of well‐defined assembly was attributed to the higher phase‐transition temperatures expected for palmitate‐derived amphiphiles compared to their oleoyl counterparts. Interestingly, when DPAG was hydrated at 50°C, spherical vesicles were observed (Figure S3B). However, such elevated temperatures are incompatible with many biochemical processes, limiting practical applicability. Subsequent analyses therefore, focused on DOAG. To further understand how the TMA group affects membrane packing, DOAG nanoparticles were synthesized using TAMARA, a microfluidic‐based system. The size of the generated particles (69 ± 2 nm) (Figures 3C, S4) was similar to that formed from the analogous diacylated amino acid (DOAA: 62 ± 5 nm) (Figure S4). Successful charge inversion was confirmed by comparing the zeta potential values of DOAG vesicles (+33 ± 3 mV) (Figures 3C, S4) with those reported for vesicles formed from diacylated amino acids (DOAA: −72 ± 3 mV) (Figure S4) [20]. This pronounced shift in surface charge not only validates the effectiveness of the proposed strategy for generating cationic abiotic membranes but also highlights the potential of these vesicles as nanoreactors and delivery vehicles capable of establishing electrostatic interactions with relevant cargo, such as nucleic acids. Finally, the formation of DOAG vesicles was corroborated by fluorescence microscopy using the apolar membrane dye 1,2‐dihexadecanoyl‐sn‐glycero‐3‐phosphoethanolamine, triethylammonium salt (Texas Red DHPE) (Figure 3B).

Characterization of DOAG vesicular structures. (A) Phase contrast microscopy images of DOAG vesicles. Scale bar denotes 10 µm. (B) Fluorescence microscopy image of DOAG vesicles stained with a 0.1 mol% of Texas Red DHPE. Scale bar denotes 10 µm. (C) Particle size distribution (left) determined by dynamic light scattering (DLS) and zeta potential values (right) obtained by electrophoretic light scattering (ELS) of DOAG nanoparticles generated using the microfluidic‐based platform TAMARA. The bar graph represents the mean particle size, while the green dot corresponds to the zeta potential. Error bars denote standard deviation from three independent measurements (n = 3). (D) Fluorescence microscopy images of DOAG vesicles encapsulating HPTS. Scale bar denotes 10 µm.

We next evaluated the capacity of DOAG vesicles to encapsulate hydrophilic molecules, using 8‐hydroxypyrene‐1,3,6‐trisulfonic acid (HPTS) as a model probe (Figure 3D). Fluorescence imaging revealed a clear distribution of HPTS within the vesicles, corroborating their ability to trap and retain small solutes. To further investigate vesicle permeability, we monitored the time‐dependent fluorescence of externally added HPTS to prehydrated DOAG vesicles (Figure S5). Initial images showed HPTS localized along the membrane, likely due to attractive interactions between the sulfonate groups of HPTS and the cationic TMA moieties on the DOAG vesicle surface. Interestingly, after 2 h of incubation at 25°C, the images revealed a heterogeneous fluorescence distribution, with the formation of HPTS‐rich domains along the membrane. This is likely due to the generation of fluid regions via associative phase separation of HPTS at the highly cationic surfaces. After 24 h of incubation, fluorescence levels became indistinguishable between the vesicle interior and the surrounding medium, indicating high permeability and a preferential localization of HPTS within the bilayer. Consistent with the permeability profiles shown in Figure S5, a gradual decrease in the fluorescence of HPTS‐encapsulating vesicles was observed over time, further confirming the semipermeable nature of these membranes. Although further studies are needed, these observations suggest that the incorporation of TMA functionality in amino acid‐based amphiphiles significantly alters membrane organization, shifting from the relatively impermeable vesicles formed by carboxylate‐terminal amphiphiles to more permeable vesicles upon cationic functionalization [20].

Additional studies were conducted to evaluate the stability of DOAG vesicles under varying physicochemical conditions (see Supporting Information). While DOAA vesicles are expected to exhibit pH‐dependent aggregation due to protonation of carboxylate anions in acidic environments (pKa ∼ 4–5), the stability of DOAG vesicles was assessed across a range of pH values (Figure S6). As anticipated, no vesicle disruption was observed, confirming that DOAG compartments remain stable under both acidic and basic conditions. The thermal stability of DOAG vesicles was also investigated over time (Figure S7). Vesicular structures remained intact at 4 and 25°C, with only minor aggregation detected at 45°C, demonstrating the robustness of DOAG assemblies under both physiologically relevant and elevated temperatures.

Biocompatibility of DAAG Vesicles

2.4

Cationic functionality is known to influence the behavior of self‐assembled structures in biological environments by promoting electrostatic interactions with negatively charged biomolecules. To assess the stability and compatibility of DOAG vesicles under biologically relevant conditions, their performance was evaluated in media supplemented with fetal bovine serum (FBS). While divalent ions present in serum, such as Ca^2+^ and Mg^2+^, can perturb lipid packing [34], potentially leading to vesicle aggregation or collapse, negatively charged serum proteins – particularly albumin – can adsorb onto cationic surfaces and trigger opsonization, thereby contributing to vesicle destabilization and clearance in delivery applications [35]. DOAG vesicles were incubated with increasing concentrations of FBS (1, 10, and 50%; v/v) and monitored for colloidal stability (Figure 4A). Vesicle integrity was retained under all tested conditions, with only minor aggregation observed at the highest serum concentration, suggesting surface interactions without structural disruption. To evaluate whether these interactions affect the biocompatibility of DOAG vesicles, in vitro cytotoxicity was assessed in HEK 293T cells exposed to increasing concentrations of DOAG for 24 h using the CCK‐8 assay. Cell viability remained above 85% relative to untreated controls across all concentrations tested (Figure 4B). These results indicate that the presence of a permanently charged TMA headgroup in DOAG does not compromise membrane stability or induce cytotoxic effects, thereby underscoring its suitability for gene or drug delivery applications.

Biocompatibility of DOAG vesicular structures. (A) Phase‐contrast microscopy images showing the stability of DOAG vesicles in 1X PBS (pH 7.4) supplemented with varying concentrations of FBS (1, 10, and 50% v/v) after 2 and 24 h. (B) Viability of HEK 293T cells exposed to DOAG vesicles. The bar graph shows the mean viability values for each tested concentration. Error bars denote standard deviation from three independent measurements (n = 3). No statistically significant differences were observed between groups (one‐way ANOVA followed by Tukey's multiple comparisons test, p > 0.05).

Experimental Section

3

Material and Methods

3.1

All reagents commercially supplied were used without further purification. Oleic acid, sodium 2‐mercaptoethanesulfonate (MESNA), 1‐(3‐dimethylaminopropyl)‐3‐ethylcarbodiimide hydrochloride (EDC·HCl), (2S)−2,3‐bis(tert‐butoxycarbonylamino)propanoic acid N‐cyclohexylcyclohexanamine salt [Boc‐L‐Dap(Boc)‐OH·DCHA], imidazole (Im), 2‐(acetylthio)‐N, N,N‐trimethylethanaminium chloride, palmitic acid, and sodium 8‐hydroxypyrene‐1,3,6‐trisulfonate (HPTS) were purchased from BLD Pharmatech (China). GRT was purchased from Angene (China). 2‐(7‐Aza‐1H‐benzotriazole‐1‐yl)−1,1,3,3‐tetramethyluronium hexafluorophosphate (HATU) and 4‐dimethylaminopyridine (DMAP) were purchased from Fluorochem (UK). 1,2‐dihexadecanoyl‐sn‐glycero‐3‐phosphoethanolamine, triethylammonium salt (Texas Red DHPE) was obtained from ThermoFisher. Anhydrous dichloromethane (CH_2_Cl_2_), trifluoroacetic acid (TFA), and formic acid (FA) were purchased from Sigma–Aldrich (USA). Deuterated solvents were purchased from Deutero GmbH (Germany). HEK 293T cells (RRID: CVCL_0063) were kindly donated by Dr. Ana Rey Rico and Dr. Juan Antonio Fafián Labora (CICA, UDC). HPLC analysis was carried out on an Agilent 1260 Infinity II LC system (Agilent Technologies, USA) using an Eclipse Plus C8 analytical column with Phase A/Phase B gradients [Phase A: H_2_O with 0.1% formic acid; Phase B: MeOH with 0.1% formic acid]. HPLC purification was carried out on an Agilent 1260 Infinity II LC system (Agilent Technologies, USA) using a Zorbax SBC18 semipreparative column with Phase A/Phase B gradients [Phase A: H_2_O with 0.1% formic acid; Phase B: MeOH with 0.1% formic acid]. Proton (^1^H‐NMR) spectra were recorded on a Bruker Avance AVA‐500 spectrometer (500 MHz) and were referenced relative to residual proton resonances in CD_3_OD (at 4.87 or 3.31 ppm) or d_6_‐DMSO (at 2.50 ppm). Chemical shifts were reported in parts per million (ppm, δ) relative to tetramethylsilane (δ 0.00). ^1^H NMR splitting patterns are assigned as singlet (s), doublet (d), triplet (t), quartet (q), or pentuplet (p). All first‐order splitting patterns were designated based on the appearance of the multiplet. Splitting patterns that could not be readily interpreted are designated as multiplet (m) or broad (br). Coupling constants are indicated in Hz. Carbon (^13^C‐NMR) spectra were recorded on a Bruker Avance AVA‐500 spectrometer (500 MHz) and were referenced relative to residual carbon resonances in CD_3_OD (at 49.15 ppm) or d_6_‐DMSO (at 39.51 ppm). Electrospray ionization spectra (ESI‐Orbitrap‐MS) were obtained using a Thermo MAT95XP. Phase‐contrast microscopy images were obtained from an Olympus microscope model BX53, using a 100x oil immersion objective and a phase contrast condenser (PH3). Images were processed using Fiji. Dynamic light scattering (DLS) and electrophoretic light scattering (ELS) were measured using a Malvern Zeta‐Sizer (Malvern Instruments, UK). Microfluidic experiments were performed using the TAMARA platform (Inside Therapeutics, France). Plates were analyzed using a Synergy HTX plate reader (BioTek, Winooski, USA).

Synthesis of TMA‐Functionalized Diamino Acids

3.2

Boc‐L‐Dap(Boc)‐GRT (1a)

3.2.1

To a solution of Boc‐L‐Dap(Boc)‐OH·DCHA (75.0 mg, 0.247 mmol) in DMF (2 mL) was added HATU (140.7 mg, 0.37 mmol) and DIPEA (107.4 µL, 0.617 mmol). After 10 min, Girard's Reagent T (GRT, 55.6 mg, 0.37 mmol) was added, and the reaction mixture was stirred at rt for 18 h. The mixture was then gradually poured into diethyl ether (10 mL) at 0°C. The resulting pale‐yellow precipitate was collected, dried, and analyzed by HPLC, confirming product formation [t_R_ = 2.3 min (50‐95% Phase B in Phase A, 0–1 min; 95% Phase B in Phase A, 1–8 min; 95‐100% Phase B in Phase A, 8–10 min; 100% Phase B, 10–13 min; 100‐50% Phase B in Phase A, 13–15 min)]. The crude product was further purified by HPLC [t_R_ = 3.9 min (50% Phase B in Phase A, 0–1 min; 5‐95% Phase B in Phase A, 1–3 min; 95% Phase B in Phase A, 3–15 min; 95‐50% Phase B in Phase A, 15–17 min; 50% Phase B in Phase A, 17–20 min)], affording 44.8 mg of 1a as a white solid (40%). ^1^H‐NMR (CD_3_OD, 500.13 MHz, δ): 4.28 (t, J = 5.1 Hz, 1H, 1 × CH); 4.16 (s, 2H, 1 × CH_2_); 3.50‐3.41 (m, 2H, 1 × CH_2_); 3.34 (s, 9H, 3 × CH_3_); 1.44 (s, 18H, 3 × CH_3_). ^13^C‐NMR (CD_3_OD, 125.77 MHz, δ): 170.67; 167.92; 162.29; 157.15; 156.28 79.55; 79.15; 63.17; 53.56; 41.56; 27.32. HRMS (ESI) calculated for C_18_H_36_N_5_O_6_ [M‐Cl]^+^: 418.27, found: 418.93.

L‐Dap‐GRT (1b)

3.2.2

To a solution of 1a (22.3 mg, 0,049 mmol), in CH_2_Cl_2_ (500 uL) was added TFA (150 uL) dropwise at 0°C under stirring. The mixture was then stirred at rt for 30 min. Afterwards, the solvent and TFA were removed under a steam of N_2_, yielfding 10.5 mg of product 1b (>99%), which was used directly without further purification. HRMS (ESI) calculated for C_6_H_20_N_5_O_2_ [M‐TFA]^+^: 218.17, found: 218.93.

Synthesis of Acyl Thioesters

3.3

MESNA Thiooleate (2)

3.3.1

The procedure was adapted from previously reported literature, following the Scheme S2 [20, 28]. Oleic acid (189.2 mg, 670.0 μmol) was dissolved in CH_2_Cl_2_ (5 mL) and stirred at 0°C for 10 min, after which DMAP (7.4 mg, 60.6 μmol) and EDC·HCl (128.4 mg, 670.0 μmol) were added sequentially. After stirring for an additional 10 min at 0°C, sodium 2‐mercaptoethanesulfonate (MESNA, 100.0 mg, 609.1 μmol) was added. The reaction mixture was stirred at rt for 5 h an then extracted with H_2_O (2 x 3 mL). The combined aqueous phases were washed with EtOAc (3 mL), and the solvent was removed under reduced pressure. The resulting residue was washed with CH_3_CN (5 mL) and filtered to afford 100.1 mg of 2 as a white solid (44%). ^1^H‐NMR (d_6_‐DMSO, 500.13 MHz, δ): 5.36‐5.16 (m, 2H, 2×CH); 3.05‐2.85 (m, 2H, 1 × CH_2_); 2.60‐2.51 (m, 4H, 2×CH_2_); 1.98‐1.94 (m, 4H, 2 × CH_2_); 1.48 (q, J = 7.4 Hz, 2H, 2 × CH_2_); 1.34‐1.08 (m, 24H, 12 × CH_2_); 0.85 (t, J = 5.7 Hz, 3H, 1 × CH_3_). ^13^C‐NMR (d_6_‐DMSO, 125.77 MHz, δ): 198.7; 129.8; 129.7; 51.0; 43.4; 31.4; 29.2; 29.1; 28.9; 28.8; 28.7; 28.6; 28.5; 28.3; 26.7; 26.6; 25.1; 24.4; 22.2; 14.1. HRMS (ESI) calculated for C_20_H_37_O_4_S_2_ [M‐Na]^−^: 405.2, found 405.2.

MESNA Thiopalmitate (3)

3.3.2

The procedure was adapted from previous literature, following the Scheme S3 [20, 28]. Palmitic acid (171.8 mg, 670.0 µmol) was dissolved in CH_2_Cl_2_ (5 mL) and stirred at 0°C for 10 min, after which DMAP (7.4 mg, 60.6 µmol) and EDC·HCl (128.4 mg, 670.0 µmol) were added sequentially. After stirring for an additional 10 min at 0°C, MESNA (100.0 mg, 609.1 µmol) was added. The reaction mixture was stirred at rt for 5 h and then extracted with H_2_O (2 × 3 mL). The combined aqueous phases were washed with EtOAc (3 mL), and the solvent was removed under reduced pressure. The resulting residue was washed with CH_3_CN (5 mL) and filtered to afford 179.8 mg of 3 as a white solid (73%). ^1^H‐NMR (d_6_‐DMSO, 500.13 MHz, δ): 3.05–3.00 (m, 2H, 1 × CH_2_); 2.61–2.56 (m, 2H, 1 × CH_2_); 2.53 (t, 2H, 1 × CH_2_); 1.52 (m, 2H, 1 × CH_2_); 1.23 (s, 24H, 12 × CH_2_); 0.88 (t, J = 7.0 Hz, 3H, 1 × CH_3_). ^13^C‐NMR (d_6_‐DMSO, 125.77 MHz, δ): 199.13; 51.59; 43.77; 31.75; 29.41; 29.29; 29.19; 29.10; 28.71; 25.51; 24.80; 22.57; 14.40. MS (ESI) [m/z (%)]: 379 ([M‐Na]^+^, 26). HRMS (ESI) calculated for C_18_H_35_O_4_S_2_ [M‐Na]^−^: 379.1982, found 379.1983.

Thiocholine Chloride (4)

3.3.3

The procedure was adapted from previous literature, following the Scheme S4 [11]. 2‐(acetylthio)‐N, N,N‐trimethylethanaminium chloride (500.0 mg, 2.50 mmol) was dissolved in a 6 M HCl aqueous solution (5 mL). The reaction mixture was stirred at 85°C for 2 h. After this time, the solvent was removed under reduced pressure, affording 320.0 mg of 4 as a white powder (75%). ^1^H‐NMR (D_2_O, 500.13 MHz, δ): 3.52–3.40 (m, 2H, 1 × CH_2_); 3.06 (s, 9H, 3 × CH_3_); 2.93–2.81 (m, 2H, 1 × CH_2_). ^13^C‐NMR (D_2_O, 125.77 MHz, δ): 67.99; 52.95; 16.62. MS (ESI) [m/z (%)]: 120 ([M‐Cl]^+^, 100). HRMS (ESI) calculated for C_5_H_14_NS [M‐Cl]^+^: 120.0841, found: 120.0838.

Choline Thiooleate (5)

3.3.4

The procedure was adapted from previous literature, following the Scheme S5 [11]. EDC·HCl (351.8 mg, 1.84 mmol) and DMAP (20.4 mg, 0.17 mmol) were dissolved in CH_2_Cl_2_ (13 mL), and oleic acid (518.4 mg, 1.84 mmol) was added. The mixture was stirred at −78°C under an argon atmosphere. After 30 min, thiocholine chloride (4, 258.7 mg, 1.67 mmol) was added, and the reaction was allowed to warm to rt and stirred for 18 h. Afterward, the mixture was concentrated under reduced pressure, and the crude product was purified by HPLC [t_R_ = 10.37 min (50 min Phase B in Phase A, 0–5 min; 95% Phase B in Phase A, 5–6 min; 100% Phase B in Phase A, 6–8 min; 100% Phase B in Phase A, 8–10 min; 50% Phase B in Phase A, 10–11 min; 50% Phase B in Phase A, 11–15 min)], affording 300.0 mg of 5 as a pale yellow oil (43%). ^1^H‐NMR (CD_3_OD, 500.13 MHz, δ): 5.24–5.18 (dt, J 1 = 2.6 Hz, J 2 = 5.6 Hz, 2H, 2 × CH); 3.29–3.19 (m, 2H, 1 × CH_2_); 3.06 (s, 9H, 3 × CH_3_); 3.05–2.99 (m, 2H, 1 × CH_2_); 2.46 (t, J = 7.3 Hz, 2H, 1 × CH_2_); 1.93–1.83 (m, 2H, 2 × CH_2_); 1.51(p, J = 7.4 Hz, 2H, 1 × CH_2_); 1.26–1.1 (ds, 20H, 10 × CH_2_); 0.75 (t, J = 6.1 Hz, 3H, 1 × CH_3_). ^13^C‐NMR (CD_3_OD, 125.77 MHz, δ): 198.01; 129.57; 129.34; 64.67; 55.10; 52.21; 43.27; 41.99; 35.93; 34.48; 31.72; 29.44; 29.38; 29.23; 29.06; 28.95; 28.87; 28.72; 28.53; 26.73; 25.66; 25.12; 22.37; 21.18. MS (ESI) [m/z (%)]: 384 ([M‐Cl]^+^, 100). HRMS (ESI) calculated for C_23_H_46_NOS [M‐Cl]^+^: 384.3300, found: 384.3279.

In Situ Synthesis of DAAGs

3.4

DOAG

3.4.1

Synthesis by Dual N‐Acylation of 1b With MESNA Thiooleate (2), Following Scheme S6

3.4.1.1

A solution of 1b (6.3 mg, 28.8 µmol), MESNA thiooleate (2, 24.8 mg, 57.8 µmol), and imidazole (143.0 mg, 2.10 mmol; Final concentration: 1.5 M) in H_2_O was stirred for 2 h. Reaction progress was monitored by HPLC, confirming product formation [t_R_ = 4.3 min (50‐95% Phase B in Phase A, 0–1 min; 95% Phase B in Phase A, 1–8 min; 95‐100% Phase B in Phase A, 8–10 min; 100% Phase B, 10–13 min; 100‐50% Phase B in Phase A, 13–15 min)] (Figure S1). The crude reaction mixture was subsequently purified by HPLC [t_R_ = 9.15 min (50‐95% Phase B in Phase A, 0–3 min; 95% Phase B in Phase A, 3–5 min; 95‐100% Phase B in Phase A, 3–5 min; 100% Phase B, 5–25 min; 100‐50% Phase B in Phase A, 25–27 min; 50% Phase B in Phase A, 27–30 min)] to afford 11.4 mg of DOAG as a colorless oil (53%). ^1^H‐NMR (CDCl_3_, 500.13 MHz, δ): 7.95 (s, 1H, 1 × NH); 7.47 (s, 1H, 1 × NH); 5.38–5.28 (m, 4H, 4 × CH); 3.75 (s, 2H, 1 × CH_2_); 3.71–3.52 (m, 2H, 1 × CH_2_); 3.34 (s, 9H, 3 × CH_3_); 2.25–2.15 (m, 4H, 2 × CH_2_); 2.05–1.92 (m, 8H, 4 × CH_2_); 1.60–1.50 (m, 4H, 2 × CH_2_); 1.26 (s, 48H, 24 × CH_2_); 0.86 (t, 6H, 2 × CH_3_). ^13^C‐NMR (CDCl_3_ 125.77 MHz, δ): 176.23; 175.14; 169.84; 162.03; 129.95; 129.62; 63.68; 54.77; 50.75; 41.07; 36.29; 31.83; 29.79; 29.69; 29.54; 29.32; 29.25; 27.24; 25.66; 25.54; 22.69. MS (ESI) [m/z (%)]: 746 ([M‐FA]^+^, 100). HRMS (ESI) calculated for C_44_H_84_N_5_O_4_ [M‐FA]^+^: 746.6518, found: 746.6957.

3.4.2

Synthesis by Dual N‐Acylation of 1b With Choline Thiooleate (5), Following Scheme S8

3.4.2.1

A solution of 1b (6.5 mg, 29.8 µmol), choline thiooleate (5, 31.2 mg, 74.5 µmol), and imidazole (122.5 mg, 1.80 mmol; Final concentration: 1.5 M) in H_2_O was stirred for 2 h. Reaction progress was monitored by HPLC, confirming product formation [t_R_ = 4.2 min (50‐95% Phase B in Phase A, 0–1 min; 95% Phase B in Phase A, 1–8 min; 95‐100% Phase B in Phase A, 8–10 min; 100% Phase B, 10–13 min; 100‐50% Phase B in Phase A, 13–15 min)]. The crude reaction mixture was subsequently purified by HPLC [t_R_ = 9.15 min (50‐95% Phase B in Phase A, 0–3 min; 95% Phase B in Phase A, 3–5 min; 95‐100% Phase B in Phase A, 3–5 min; 100% Phase B, 5–25 min; 100‐50% Phase B in Phase A, 25–27 min; 50% Phase B in Phase A, 27–30 min)] to afford 9.3 mg of DOAG as a colorless oil (42%).

DPAG

3.4.3

Synthesis by Dual N‐Acylation of 1b With MESNA Palmitate (3), Following Scheme S7

3.4.3.1

A solution of 1b (6.5 mg, 29.8 µmol), MESNA thiopalmitate (3, 45.4 mg, 74.5 µmol), and imidazole (122.5 mg, 1.80 mmol; final concentration 1.5 M) in H_2_O was stirred for 2 h. Reaction progress was monitored by HPLC, confirming product formation [t_R_ = 2.9 min (50‐95% Phase B in Phase A, 0–1 min; 95% Phase B in Phase A, 1–8 min; 95‐100% Phase B in Phase A, 8–10 min; 100% Phase B, 10–13 min; 100‐50% Phase B in Phase A, 13–15 min)] (Figure S1). The crude reaction mixture was subsequently purified by HPLC [t_R_ = 8.5 min (50‐95% Phase B in Phase A, 0–3 min; 95% Phase B in Phase A, 3–5 min; 95‐100% Phase B in Phase A, 3–5 min; 100% Phase B, 5–25 min; 100‐50% Phase B in Phase A, 25–27 min; 50% Phase B in Phase A, 27–30 min)] to afford 9.0 mg of DPAG as a colorless oil (44%). ^1^H‐NMR (CDCl_3_, 500.13 MHz, δ): 7.51 (s, 1H, 1 × NH); 4.90–4.78 (m, 1H, 1 × CH); 4.32 (s, 2H, 1 × CH_2_); 3.48 (s, 9H, 3 × CH_3_); 2.34 (t, J = 7.5 Hz, 9H, 3 × CH_3_); 1.67–1.58 (m, 4H, 2 × CH_2_); 1.25 (s, 48H, 24 × CH_2_); 0.87 (t, J = 7.4 Hz, 6H, 2 × CH_3_). ^13^C‐NMR (CDCl_3_, 125.77 MHz, δ): 177.80; 174.36; 167.81; 157.84; 76.03; 70.02; 36.25; 33.77; 31.94; 29.69; 29.60; 29.45; 29.37; 29.25; 29.09; 28.10. MS (ESI) [m/z (%)]: 694 ([M‐FA]^+^, 100). HRMS (ESI) calculated for C_40_H_80_N_5_O_4_ [M‐FA]^+^: 694.6205, found: 694.6656.

Vesicle Formation

3.5

Hydration Method

3.5.1

15 µL of a 50 mM solution of DAAG (DOAG or DPAG) in a 1:1 mixture of MeOH:CHCl_3_ was added to a 1 mL glass vial, and the solvent was evaporated under a gentle N_2_ stream to form a thin lipid film. The resulting film was hydrated with 100 µL of Milli‐Q water at either 37 or 50°C. The samples were subsequently analyzed by phase‐contrast and fluorescence microscopy. Vesicles were stained with 0.1 mol% Texas Red DHPE (50 µM in EtOH) (Figures 3A,B, S3).

Microfluidics

3.5.2

Vesicles were prepared by mixing an ethanolic solution of DOAG (Final concentration: 10 mg/mL) with Milli‐Q water at a 1:3 volume ratio and a total flow rate of 4 mL/min. Vesicular nanostructures were generated using TAMARA, a microfluidic‐based nanoparticle formulation system. The resulting DOAG nanoparticles (Final concentration of DOAG: 2.5 mg/mL) were collected in Eppendorf tubes. Vesicle size was characterized by DLS (Figures 3C, S4), and surface charge was determined by zeta potential analysis (Figures 3C, S4).

Encapsulation of Fluorescent Dyes

3.6

Inverse‐Emulsion Method

3.6.1

30 μL of a 20 mM solution of DOAG in MeOH/CHCl_3_ (1:1) was added to a 1 mL glass vial, and dried under a gentle N_2_ stream for 15 min to form a lipid film. Subsequently, 100 μL of mineral oil was added, and the vial was flushed with N_2_ to displace the air above the mineral oil. The resulting mixture was sonicated with heating (∼ 55°C) for 1 h. The oil phase was then transferred to a 1 mL Eppendorf tube, and 10 μL of the upper aqueous buffer (50 μM HPTS + 200 mM sucrose in 100 mM HEPES buffer, pH 7.5) was added. The mixture was gently flicked and vortexed until a cloudy emulsion formed. The corresponding emulsion was carefully layered onto 100 μL of the lower aqueous buffer (200 mM glucose in 100 mM HEPES buffer, pH 7.5) in a 1 ml Eppendorf tube, allowing it to float on top. After standing for 10 min, the sample was centrifuged at 9,000–10,000 rcf for 10 min. The lower aqueous phase was then collected by either aspirating the oil layer or withdrawing the sample using a syringe and needle. The recovered sample contained DOAG vesicles encapsulating HPTS, which were subsequently imaged by fluorescence microscopy (Figure 3D).

Permeability Study

3.7

15 µL of a 50 mM DOAG solution in a 1:1 mixture of MeOH:CHCl_3_ was added to a 1 mL glass vial, and the solvent was evaporated under a gentle N_2_ stream to obtain a lipid film. The film was then hydrated with 100 µL of 1 mM phosphate‐buffered saline (PBS) (pH 7.4) for 1 h at rt. The final lipid concentration was adjusted to 50 µM by diluting the sample with 1 mM PBS buffer (pH 7.4) containing 50 µM HPTS. The resulting samples were analyzed by fluorescence microscopy (Figure S5).

pH Stability

3.8

10 µL of a DOAG solution prepared according to the general hydration method were diluted with 10 µL of buffers at different pH values: 50 mM sodium acetate (AcONa) at pH 4.0 and 5.0, 50 mM phosphate buffer (PB) at pH 6.0, 7.0, and 8.0, and 50 mM bicine at pH 9.0 and 10.0. The resulting mixtures were monitored using phase‐contrast microscopy after 2 and 24 h of incubation at rt to assess vesicle morphology (Figure S6).

Temperature Stability

3.9

10 µL of a DOAG solution prepared according to the general hydration method was diluted with 5 µL of Milli‐Q water. The resulting mixtures were incubated at 4, 25, and 45°C under agitation (100 rpm) using a Thermo Scientific Digital Heating Shaking Drybath for 2 and 24 h. The samples were subsequently analyzed by phase‐contrast microscopy to assess vesicle morphology (Figure S7).

Serum Stability

3.10

10 µL of a DOAG solution prepared according to the general hydration method was mixed with an equal volume of 1X PBS (pH 7.4) containing increasing concentrations of fetal bovine serum (FBS; 1, 10, and 50% v/v). Samples were incubated under orbital agitation for 2 and 24 h using a Thermo Scientific Digital Heating Shaking Drybath, and subsequently analyzed by phase‐contrast microscopy to assess vesicle morphology (Figure 4).

Cell Studies

3.11

Cell Culture

3.11.1

HEK 293T cells were cultured at 37°C in a 5% CO_2_ atmosphere in high‐glucose DMEM supplemented with 10% FBS, 50 U/mL penicillin, and 50 U/mL streptomycin, using 75 cm^2^ cell culture flasks. This modified DMEM is referred to as ‘medium’ throughout the text.

Cell Viability

3.11.2

A stock solution of DOAG (500 µg/mL) was prepared using the general hydration method, with Opti‐MEM as the hydration medium. The resulting sample was diluted with medium to final concentrations ranging from 5 to 250 µg/mL. HEK 293T cells were seeded at 25,000 cells/well in 96‐well plates (Corning Costar, Thermo Scientific) and incubated at 37°C with 5% CO_2_ for 24 h. The medium was then replaced with 50 µL of the DOAG solutions, and cells were incubated for an additional 24 h. Cell viability was assessed using a colorimetric CCK‐8 assay according to the manufacturer's protocol. After 2 h of incubation with a 6% solution of CCK‐8 in fresh medium, absorbance at 450 nm was measured using a Synergy HTX plate reader. Cell viability was calculated as follows:

$$[eqn]$$

Statistics

3.12

All statistical analyses were performed using GraphPad Prism 10. The primary objective was to assess the tolerability of increasing DOAG concentrations in HEK293T cells using a CCK‐8 colorimetric viability assay. Background absorbance from the CCK‐8 reagent in cell‐free medium was subtracted from all measurements, and values were normalized to untreated control cells. Data are presented as mean ± standard deviation. Statistical comparisons were performed using one‐way analysis of variance (ANOVA) followed by Tukey's multiple comparisons test. No statistically significant differences were observed between groups (all adjusted p > 0.05).

Differences in hydrodynamic radius between DOAG and DOAA vesicles were analyzed using an unpaired two‐tailed Student's t‐test (Figure S4). Data are presented as mean ± standard deviation.

Conclusion

4

In summary, we have demonstrated that key architectural features of biological phospholipids, such as permanent headgroup charge, diacyl topology, and vesicle‐forming ability, can be reconstructed using minimal amino acid‐based systems. By combining a TMA‐functionalized diamino acid headgroup with aqueous, thioester‐driven acylation, we establish a chemically feasible route to membrane‐forming cationic, diacyl amphiphiles that bridge prebiotic relevance and contemporary functional requirements. Importantly, this strategy enables systematic variation of headgroup chemistry without compromising bilayer integrity, providing a level of modularity difficult to achieve with natural phospholipids. Beyond generating novel choline‐mimetic membranes, these results suggest that membrane function can emerge from simplified chemical designs, supporting the use of synthetic amphiphiles both as models for early compartmentalization and as versatile platforms for synthetic biology and molecular delivery.

Author Contributions

Federica A. Souto‐Trinei: conceptualization (lead), data curation (supporting), formal analysis (lead), investigation (lead), methodology (lead), writing – original draft (lead), writing – review & editing (lead). Bruno Delgado Gonzalez: conceptualization (lead), data curation (supporting), formal analysis (lead), investigation (lead), methodology (lead), writing – original draft (lead), writing – review & editing (lead). Roberto J. Brea: conceptualization (lead), data curation (lead), formal analysis (lead), funding acquisition (lead), investigation (lead), methodology (lead), project administration (lead), resources (lead), supervision (lead), validation (lead), writing original draft (lead), writing – review & editing (lead).

Supporting Information

Additional supporting information can be found online in the Supporting Information section. The authors have incorporated additional data in the Supporting Information, including Supplementary Schemes S1‐S8, Supplementary Figures S1‐S7, and NMR and MS spectra. Supporting Scheme S1: Synthesis of L‐Dap‐GRT (1b). Supporting Scheme S2: Synthesis of MESNA thiooleate (2). Supporting Scheme S3: Synthesis of MESNA thiopalmitate (3). Supporting Scheme S4: Synthesis of thiocholine chloride (4). Supporting Scheme S5: Synthesis of choline thiooleate (5). Supporting Scheme S6: Synthesis of DOAG by dual N‐acylation of L‐Dap‐GRT (1b) with MESNA thiooleate (2). Monooleoylated intermediate (MOAG) is also shown. Supporting Scheme S7: Synthesis of DPAG by dual N‐acylation of L‐Dap‐GRT (1b) with MESNA thiopalmitate (3). Monopalmitoylated intermediate (MPAG) is also shown. Supporting Scheme S8: Synthesis of DOAG by dual N‐acylation of L‐Dap‐GRT (1b) with choline thiooleate (5). Monooleoylated intermediate (MOAG) is also shown. Supporting Fig. S1: HPLC (210 nm) traces corresponding to purified DOAG (black) and DPAG (blue). Retention times (tR) were verified by mass spectrometry. Supporting Fig. S2: Comparison of particle size distribution (left) obtained by DLS and zeta potential values (right) measured by ELS for MESNA thioleate (2) and choline thioleate (5). Bar graphs represent mean particle size and blue (for 2) and red (for 5) dots represent corresponding zeta potential. Error bars denote standard deviation from three independent measurements (n = 3). Supporting Fig. S3: Phase‐contrast (left) and fluorescence (right) microscopy images of DPAG films hydrated at 37°C (A) and 50°C (B) for 1 h. Samples were stained using 0.1 mol% Texas Red DHPE. Scale bars denote 10 µm. Supporting Fig. S4: Comparison of particle size distribution (left) obtained by DLS and zeta potential values (right) measured by ELS for DOAG and DOAA. Bar graphs represent mean particle size, and dots represent the corresponding mean zeta potential. Error bars denote standard deviation from three independent measurements (n = 3). Differences in hydrodynamic radius between DOAG and DAAG vesicles were analyzed using an unpaired two‐tailed Student's t test, showing no statistical difference (p > 0.05). Supporting Fig. S5: Fluorescence microscopy time‐lapse images of DOAG vesicles recorded at 0, 2, and 24 h in 1 mM PBS buffer (pH 7.4) at 37°C following external addition of HPTS (50 µM). Scale bars denote 10 µm. Supporting Fig. S6: Representative phase‐contrast microscopy time‐lapse images of DOAG vesicles recorded at 2 and 24 h across a pH range of 4‐10. Scale bars denote 10 µm. Supporting Fig. S7: Representative phase‐contrast microscopy time‐lapse images of DOAG vesicles recorded at 2 and 24 h across a temperature range of 4‐45°C. Scale bar denotes 10 µm.

Funding

This work was supported by the Agencia Estatal de Investigación (PID2021‐128113NA‐I100, RYC2020‐030065‐I); Consellería de Cultura, Educación e Ordenación Universitaria, Xunta de Galicia (ED431F 2024/07, ED431B 2023/60).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supplementary Material