Arginine‐Rich Cell‐Penetrating Peptides Induce Lipid Rearrangements for Their Active Translocation across Laterally Heterogeneous Membranes

Abstract

Arginine‐rich cell‐penetrating peptides (CPPs) have emerged as valuable tools for the intracellular delivery of bioactive molecules, but their membrane perturbation during cell penetration is not fully understood. Here, nona‐arginine (R₉)‐mediated membrane reorganization that facilitates the translocation of peptides across laterally heterogeneous membranes is directly visualized. The electrostatic binding of cationic R₉ to anionic phosphatidylserine (PS)‐enriched domains on a freestanding lipid bilayer induces lateral lipid rearrangements; in particular, in real‐time it is observed that R₉ fluidizes PS‐rich liquid‐ordered (L_o) domains into liquid‐disordered (L_d) domains, resulting in the membrane permeabilization. The experiments with giant unilamellar vesicles (GUVs) confirm the preferential translocation of R₉ through L_d domains without pore formation, even when L_o domains are more negatively charged. Indeed, whenever R₉ comes into contact with negatively charged L_o domains, it dissolves the L_o domains first, promoting translocation across phase‐separated membranes. Collectively, the findings imply that arginine‐rich CPPs modulate lateral membrane heterogeneity, including membrane fluidization, as one of the fundamental processes for their effective cell penetration across densely packed lipid bilayers.

Introduction

The transport of active molecules (e.g., ions, metabolites, and drugs) across cell membranes has been significantly challenging due to their limited diffusion through phospholipid bilayers. To address this, cell-penetrating peptides (CPPs) have attracted great attention because of their ability to cross cell membranes with cargoes.^[1,2] In particular, arginine-rich peptides, including HIV-1 TAT peptide and polyarginines (R_n), are widely used for imaging and therapeutic applications.^[3] While energy-dependent endocytosis has been regarded as a major route of cellular internalization of various CPPs, growing evidence reveals that arginine-rich CPPs could be directly translocated across membranes in an energy-independent manner.^[4–8] The direct translocation of arginine-rich CPPs has been suggested to be related to their strong interactions with lipid headgroups. Specifically, the positively charged arginine prefers to bind to negatively charged lipids (e.g., phosphatidylserine (PS) and phosphatidylglycerol (PG)), and the guanidinium group of arginine even forms hydrogen bonds with phosphate, carboxylate or sulfate groups of lipids, causing structural alterations of peptide-bound membranes.^[9,10]

Multiple mechanisms have been proposed to explain how CPPs traverse cell membranes. One mechanism is that the binding of CPPs induces stable pores in lipid membranes for subsequent cell penetration of molecules. The strong interactions between arginine-rich CPPs and lipids are well known to significantly reduce the thermodynamic cost of membrane pore formation; both HIV-1 TAT^[11–13] and R_n^[14–16] have proven to induce pores in membranes, allowing the passage of small dyes or ions. The other mechanism suggests that when transient pores naturally occur with thermal fluctuations, CPPs utilize these pores in translocation.^[15,17] Based on the classical theory of pore formation,^[18] the probability of both stable and transient pore formation is mainly determined by two factors; the lateral tension of the membrane, which promotes pore expansion, and the line tension of the pore edge, which promotes pore closure. Accordingly, the translocation ability of CPPs has been found to be influenced by the mechanical properties of lipid membranes, relevant to lipid chain length and saturation, and cholesterol levels.^[19–22]

Another fundamental feature of cellular membranes is their lateral heterogeneity.^[23,24] Several studies have shown that the alterations in lateral membrane organization have a significant role in the functionality of diverse membrane-active peptides. Antimicrobial peptides (AMPs) have been suggested to disrupt bacterial biological functions by affecting the lateral mobility of membrane components and the distribution of membrane proteins.^[25–28] Phase separation induced by a few CPPs has also been reported, raising the possibility of peptide translocation via phase boundary defects^[29–31] or modulated membrane curvatures.^[32] While it is evident that the translocation ability of the CPPs could not be fully understood without membrane heterogeneity, the relationship between CPP translocation and dynamic membrane rearrangements in heterogeneous membranes remains poorly understood. Instead, penetration efficiencies have typically been evaluated by externally modulating overall membrane properties by changing membrane compositions.^{[7,12,15,33,34]}

In this work, we demonstrate the dissolution of ordered lipid domains in heterogeneous membranes upon the binding of nona-arginine (R₉), a representative arginine-rich CPP,^[35,36] that can facilitate peptide translocation across the membrane. Using planar freestanding lipid membranes,^[37] we successfully visualized the lateral rearrangements induced by R₉ in real-time. In particular, we found that the lipid rearrangements could lead the liquid-ordered (L_o) phase to fluidize into the liquid-disordered (L_d) phase, thereby lowering the phase transition temperature of the lipid bilayers. Penetration analysis of R₉ into phase-separated giant unilamellar vesicles (GUVs) further confirmed that the dissolving of the L_o domains accelerates the direct translocation of R₉. These findings suggest that peptide-induced membrane remodeling may accelerate membrane translocation of CPPs by altering the fluidity of tightly packed lipid membranes.

Results

Experimental Setup for Real-Time Monitoring of Lateral Membrane Reorganization

To visualize the dynamic membrane reorganization induced by nona-arginine (R₉), we used a freestanding planar lipid membrane array in a transmission electron microscopy (TEM) grid (Figure 1a; Figure S1, Supporting Information).^[37,38] We prepared simplified lipid mixtures composed of phosphatidylcholine (PC), phosphatidylserine (PS), and cholesterol, mimicking mammalian plasma membranes. The preparation of phase-separated membranes began with the mixing of unsaturated phospholipid (DOPC) and saturated phospholipid (DPPC), both of which are neutral in charge, and the resulting DOPC/DPPC/cholesterol membrane underwent phase separation into a bright L_d phase and a dark L_o phase at 24 °C with 0.5 mol% of Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (TR-DHPE) (Figure 1b). We note that the L_d phase is enriched in unsaturated phospholipids and fluorescent TR-DHPE, and the L_o phase is enriched in saturated phospholipids and cholesterol. Either the L_d or L_o phase can be more negatively charged to strongly interact with the positively charged R₉; by introducing PS, an anionic phospholipid, in a saturated or unsaturated form (i.e., DPPS or DOPS), we could determine which phase preferentially interacted with the cationic peptide. For example, upon the inclusion of DPPS, the fluorescence of FITC-labeled R₉ (FITC-R₉) was predominately found in the L_o phase, where anionic saturated phospholipids were more abundant (Figure 1b). Even with GUVs, we successfully visualized the phase-preferential R₉ binding (Figure S2, Supporting Information), and the R₉ preferred binding to the L_d or L_o phase with 10 mol% DOPS or DPPS, respectively, whereas the binding did not occur on the neutral membrane of DOPC/DPPC/cholesterol.

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R₉-Induced Lipid Rearrangements on Phase-Separated Membranes

On heterogeneous membranes, the binding of cationic R₉ induced lateral rearrangements of anionic lipids (Figure 2). The phase behaviors of freestanding lipid bilayers were monitored in real-time by introducing 50 µm R₉ to the membranes while varying the membrane compositions. On neutral DOPC/DPPC/cholesterol membranes, the addition of positively charged R₉ had no effect due to the absence of negatively charged lipids (Figure 2a). However, when cationic DPPS was included in the heterogeneous membranes, two significant changes in phase behavior occurred (Figure 2b; Movie S1, Supporting Information). First, immediately upon the addition of R₉, small L_o domains were formed within the pre-existing L_d domains as a result of R₉ binding (Figure S3, Supporting Information). It is known that electrostatic interactions between anionic lipids and cationic peptides can induce lateral segregation of anionic lipids,^{[28,30,39,40]} and we also confirmed that initially homogeneous DOPC/DPPS membranes underwent phase separation due to the DPPS clustering upon the addition of R₉ (Figure S3, Supporting Information), making FITC-R₉ highly concentrated on the surface of newly formed L_o domains (Figure S4, Supporting Information). Second, over several tens of minutes, the total area of L_o domains decreased dramatically as a result of domain dissolution, that is, phase mixing. On membranes containing 10 mol% DPPS, for example, L_o domains decreased to 49.5 ± 31.4% (n = 4) of the initial area after 30 min (Figure 2d). Similarly, with 4 µm of HIV-1 TAT, another representative arginine-rich CPP, the L_o domains of membranes containing 16 mol% DMPG were reduced to 53.7 ± 7.1% (n = 10) of the initial area after 1 h (Figure S5, Supporting Information). We note that there were no alterations in the phase behavior with membranes containing DOPS (Figure S6, Supporting Information). It was anticipated that these membranes would undergo lateral rearrangements in a similar manner; however, the inability of unsaturated lipid DOPS to form dark ordered phase domains made their observation technically challenging through domain visualization based on dye partitioning.^[41,42]

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Despite the early temporal clustering of DPPS-rich ordered domains within the L_d phase, DPPS-rich L_o domains, including newly formed ones, eventually dissolved over time. The observed phase mixing suggests that the binding of cationic R₉ to anionic L_o domains would ultimately decrease the miscibility temperature (T_m) of lipid phase separation. To analyze the temperature-dependent phase separation behavior of laterally heterogeneous membranes, the temperature of an imaging chamber was increased at a rate of ≈0.4 °C min⁻¹, and the number of phase-separated membranes was counted every 2 °C. By linearly fitting the collected data, the apparent T_m was determined as the temperature at which 50% of the total membranes undergo phase separation (Figure 2e). For example, the T_m of bare DOPC/DPPC/DPPS/cholesterol membranes was measured to be 30.1 °C, but 30 min incubation with 50 µm R₉ lowered the T_m to 28.2 °C. This decrease in T_m indicates that the cationic R₉ fluidizes the anionic lipid membranes, leading to the observed L_o domain dissolution by phase mixing.

Electrostatic Interactions of R₉ with Heterogeneous Membranes in Phase Mixing

As the dissolution of L_o domains is likely due to the electrostatic interactions between the cationic R₉ and the anionic DPPS in the DPPS-containing heterogeneous membranes, we next investigated the effect of DPPS and R₉ concentration, respectively. We first varied the molar content of DPPS in the phase-separated membranes, and the total area of L_o domains was measured over time (Figure 2b–d). While 10 mol% DPPS resulted in a decrease of ≈50.5% in the area of the L_o domains (Figure 2b), 5 mol% DPPS did not cause such a significant change (Figure 2c). It should be noted that we could not investigate the effect of higher mol% DPPS; as the percentage of DPPS increased, the resulting lipid bilayer became increasingly unstable, impeding the formation of freestanding planar lipid membranes in a TEM grid. However, the effect of R₉ concentrations on the extent of L_o domain dissolution was well evaluated when the DPPS content was up to 10 mol%. When the R₉ concentration was gradually changed from 5 to 50 µm for 180 min, the decrease in the total area of L_o domains became more significant at higher R₉ concentrations (Figure 2f). Although we only observed the membranes for 30 min at each concentration, phase mixing occurred slowly and continuously even at low R₉ concentrations, and were able to identify the trend of R₉ concentration-dependent dissolution of the L_o domains. Note that in our experiment system, the lipid-to-peptide ratio is expected to be ≈1:10 at 50 µm R₉, potentially leading to dramatic phase mixing at this high peptide concentration.

The real-time observations indicated that the rate of phase mixing would be accelerated with higher levels of anionic lipids and cationic peptides. When there were stronger interactions between the anionic lipids and the cationic peptides, the T_m of phase-separated membranes would decrease further, leading to more domain dissolution by phase mixing. Indeed, the dissolution of L_o domains on heterogeneous membranes became faster with increasing concentrations of anionic DPPS and cationic R₉ (Figure 2d–f); even at low concentrations of DPPS and R₉, the membrane reorganization would occur, although with a limited extent of phase mixing. Thus, we concluded that the presence of R₉ would induce membrane reorganization and subsequent fluidization, allowing us to speculate that arginine-rich CPP-mediated lipid rearrangements would be directly related to its active translocation across laterally heterogeneous membranes.

Investigating Penetration of R₉ into Homogeneous and Heterogeneous GUVs

The energy-independent cell penetration of arginine-rich CPPs is generally explained by the aforementioned two mechanisms, with the critical difference being whether or not the peptide is directly involved in pore formation at plasma membranes. Using PS-containing heterogeneous membranes, we explored the possibility of R₉-induced pore formation and its membrane penetration. To evaluate the pore-forming ability of R₉, we encapsulated calcein, a water-soluble fluorescent dye (≈6.5 Å in radius), into DOPC/DPPC/DPPS/cholesterol or DOPC/DOPS/DPPC/cholesterol GUVs, and observed the dye leakage from the interior to the exterior of the GUVs in real-time (Figure 3). Because the GUVs were unstable at high R₉ concentrations (>50 µm), our experimental setup was limited to 10 µm R₉, making our leakage measurements reliable and reproducible. It should be noted that this concentration of peptide was still capable of inducing a phase transition from L_o to L_d phase. Over 30 min, the addition of R₉ did not cause notable fluorescence changes inside the GUVs (Figure 3a), indicating that there were no large pores for the calcein to pass through. Moreover, there was no big difference in dye leakage between DPPS- and DOPS-containing membranes, confirming that phase-preferential R₉ binding would be irrelevant to pore formation as well (Figure 3b).

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Without the formation of large pores, arginine-rich CPPs can be internalized into PS-containing GUVs, but at different rates for different phases of lipid membranes (Figure 4). To determine which phase is more helpful for the penetration of R₉, we prepared anionic L_d or L_o phase GUVs, which can be represented by DOPC/DOPS/cholesterol or DPPC/DPPS/cholesterol, respectively. To visualize and quantify the penetration of R₉, we used FITC as a fluorophore conjugated to R₉. We note that due to its anionic nature under the experimental pH, the physicochemical properties of the peptide can be altered, potentially affecting its ability to penetrate the membrane.^[43,44] Into the L_d phase GUVs, the penetration of FITC-R₉ was extremely rapid even at 5 µm FITC-R₉, so that the fraction of R₉-internalized GUVs reached 100% in just 10 min (Figure 4a, empty red diamond); note that we extrapolated the FITC-R₉ concentrations inside GUVs from their fluorescence intensities using confocal fluorescence microscopy images, such that GUVs with >0.2 µm FITC-R₉ were regarded as the R₉-internalized GUVs (Figure S7, Supporting Information). On the other hand, the entry of FITC-R₉ into the L_o phase GUVs occurred less frequently than that into the L_d phase GUVs (empty black diamond). In particular, when comparing the quantities of FITC-R₉ between the two different GUVs, the L_d phase GUVs composed of unsaturated lipids (Figure 4b) exhibited a higher accumulation of FITC-R₉ in their lumens compared to the L_o phase GUVs composed of saturated lipids (Figure 4c), which is in good agreement with previous studies showing a rapid penetration of arginine-rich CPPs through the membranes composed of unsaturated lipids.^[15,16] Higher concentrations of FITC-R₉ accelerated its accumulation in both the L_d and L_o phase GUVs (Figure S8a,b, Supporting Information), and the absence of anionic lipids caused neither calcein leakage nor FITC-R₉ penetration even at high R₉ concentrations (≈50 µm) (Figure S9, Supporting Information).

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As we have previously demonstrated in planar membranes, the presence of arginine-rich CPPs in phase-separated GUVs can cause a further phase transition from L_o to L_d phase, facilitating their translocation across more fluidized membranes. Similar to the single-phase GUVs, PS-containing phase-separated GUVs allowed consistent penetration of FITC-R₉, and the presence of anionic L_d domains was more effective for membrane penetration than that of anionic L_o domains; at 5 µm FITC-R₉, 97.1% DOPS-containing GUVs were determined as R₉-internalized GUVs within 10 min (Figure 4a, solid red square), and the DPPS-containing GUVs displayed 74.8% during the same period (solid black square). However, compared to the DPPS-containing homogeneous GUVs (empty black diamond) with the same content of PS and cholesterol (10 and 30 mol%, respectively), the DPPS-containing heterogeneous GUVs could accelerate the penetration of R₉, attributed to the dissolution of L_o domains into L_d ones upon the phase-preferential R₉ binding. Lipid packing defects at the domain boundary of phase-separated membranes may also have a supplementary effect on enhancing R₉ internalization.^[45] Even the time-dependent R₉ accumulations in the DPPS-containing heterogeneous GUVs (Figure 4e) were much larger than those in the DPPS-containing homogeneous GUVs at 5 µm FITC-R₉ (Figure 4c). Similar trends were observed at higher R₉ concentrations (Figure S8b–d, Supporting Information). Although the time-dependent R₉ accumulations were smaller in the DOPS-containing heterogeneous GUVs (Figure 4d; Figure S8c, Supporting Information) than those in the DOPS-containing homogeneous GUVs (Figure 4c; Figure S8a, Supporting Information), these would be due to the different area fraction of L_d domains relevant to the total amount of unsaturated lipids.

Dynamics of R₉ Binding and Its Penetration into Phase-Separated GUVs

To further understand the arginine-rich CPP-mediated lipid rearrangements and its active translocation across laterally heterogeneous membranes, we visualized the R₉ binding and its penetration in real-time in the coexistence of L_d and L_o phase domains (Figure 5). Using FITC-R₉ (green) and TR-DHPE (red), we readily assess peptide binding at L_d domains with strong red fluorescence and L_o domains with weak red fluorescence; the phase-preferential R₉ binding and subsequent R₉ accumulation were evaluated with the intensity of FITC fluorescence at the rim of each phase and in the lumen of GUVs composed of DOPC/DPPC/DPPS/cholesterol (Figure 5a). In measuring the rim intensities, a straight line was drawn through three points: the L_o phase region, the GUV center, and the L_d phase region (Figure S10, Supporting Information). The first and second highest peaks in the TR-DHPE intensity profile corresponded to the L_d and L_o phase regions, respectively. The FITC intensity at each peak represented the concentration of FITC-R₉ at each phase domain, and we obtained the average values from 5 distinct lines.

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As soon as the DPPS-containing heterogeneous GUVs were exposed to FITC-R₉, their outer surfaces were fully covered with the bound FITC-R₉, with a larger amount bound to the L_o domain than at the L_d domain (point (1) in Figure 5). For the following 200 s (from point (1) to (2)), the normalized intensity of L_o phase rim (solid black square) was maintained at ≈0.5, while that of L_d phase rim (solid red square) slowly increased, inferring that the FITC-R₉ would not pass through the L_o domain but through the L_d domain. However, at point (2), the intensity of L_o phase rim increased with the time delay; presumably, the FITC-R₉ that entered the GUV through the L_d domain would be bound to the inner surface of the L_o domain rather than being free in the lumen, as there was no significant increase of accumulated FITC-R₉ (empty green square). Nevertheless, a certain fraction of the peptide did not adhere to the inner membrane surface and instead remained within the lumen. At point (3), the intensity of L_d phase rim reached a plateau with no further increase, but that of GUV lumen began to increase rapidly, indicating continuous penetration of FITC-R₉ and its subsequent accumulation in the GUV. As the concentration of FITC-R₉ inside the GUV became sufficiently high, at point (4) the intensity of L_o phase rim also reached a plateau, suggesting that even the inner surface of the L_o domain would be completely occupied with the internalized FITC-R₉. However, the intensity of the GUV lumen consistently increased with FITC-R₉ accumulation over time (from point (4) to (5)).

The preferential penetration of R₉ through the L_d domain suggests that when L_d and L_o domains coexist in phase-separated membranes, translocation of arginine-rich CPPs would favor the thinner and less rigid domains; in other words, the ordered domains have a higher concentration threshold for peptide penetration. Based on the previously reported phase diagrams,^[46–48] we estimated that with our membrane composition (10 mol% DPPS), the L_d and L_o phase domains would contain ≈7–13 mol% DPPS, respectively. Because of lower charge density, the L_d domain was less covered with the FITC-R₉ than the L_o domain, but the initial increase in the intensity of L_d phase rim validated that the membrane flexibility would be more crucial for the penetration of FITC-R₉. As the penetration of R₉ across the less anionic L_d domains prevails over that across the more anionic L_o domains, it is evident that the R₉-induced domain dissolution from a more ordered state to a less ordered state would actively lead to their easier translocation across laterally heterogeneous membranes. This implies that the dissolution of the L_o phase into the L_d phase induced by R₉ plays a substantial role in facilitating the penetration of the peptide.

Conclusion

In exploring how arginine-rich CPPs traverse laterally heterogeneous cell membranes, we directly visualized in real-time the dynamic reorganization of membranes with coexisting L_d and L_o phases mediated by R₉ binding and its penetration. By analyzing the fluorescence intensities of FITC-R₉ at the surface and in the lumen of phase-separated GUVs, we observed that R₉ translocated through the disordered domains rather than the ordered domains. This result is consistent with previous findings that the penetration of R₉ was readily observed when the membrane was mechanically weak and deformable, but became rare when it was not.^[15,45,49] However, we also demonstrated that R₉ eventually dissolved the L_o domains containing anionic lipids, suggesting that the arginine-rich CPPs actively alter the spatial and temporal distribution of membrane components. Presumably, in the heterogeneous GUVs, additional R₉ would lead to lateral expansion of ordered domains to form energetically unfavorable voids, and hydrocarbon chains of neighboring lipids would rearrange into a more disordered conformation to remove the free volumes in the bilayer.^[50,51] Owing to the R₉-induced membrane reorganization_, the heterogeneous membranes would become fluidized, thereby facilitating subsequent translocation of R₉ across otherwise highly ordered, impermeable membranes (Figure 6).

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The finding of this study, demonstrating that the dissolution of ordered membrane domains promotes the penetration of CPPs, is in accordance with established explanations underlying membrane permeability. Membrane thickness is an important factor in determining permeability across the membrane; the disordering of membrane lipids typically leads to a decrease in membrane thickness,^[52] subsequently lowering the energy barrier for peptide penetration.^[49,53] In particular, the transition of the membrane from L_o to L_d phase dramatically improves the permeability of small molecules through the membrane,^[54,55] which is consistent with our results with R₉. Furthermore, an increase in the lipid bilayer fluidity makes the bilayer more susceptible to further membrane reconstructions, such as alterations in membrane curvature^[16,32,56] and redistribution of membrane components,^{[13,29,32,57]} which have been proposed as a potential mechanism for CPP translocation.

In a cell, negatively charged lipids are not uniformly distributed throughout the plasma membrane. For instance, PS is predominantly located at the inner leaflet of the eukaryotic plasma membrane.^[58] This asymmetry is disrupted in cancer cells, as well as other pathological and apoptotic cells.^[59,60] Consequently, PS has emerged as a promising target for anticancer therapeutics,^[61] including arginine-rich CPPs.^[62,63] Additionally, a considerable fraction of PS is found to co-segregate with cholesterol to form distinct microdomains called lipid rafts,^[64–67] and the lipid rafts are generally accepted to exist as L_o domains.^[68,69] Accordingly, it is typically regarded that penetration of CPPs would be significantly challenging through the highly ordered lipid rafts; previous studies have reported that increasing the structural order of membranes, particularly with increasing cholesterol content, suppresses the translocation of CPPs.^[15,33,34] To increase the accumulation of CPPs within cells or vesicles, perturbation of lipid packing, such as cholesterol extraction and the addition of membrane-disrupting molecules, has been suggested.^[57,70–72] In this regard, our current findings expand previous works by demonstrating that CPPs themselves can locally loosen the packing of pathological cell membranes through lateral membrane reorganization prior to their active translocation. We believe that this complements existing model membrane studies that have assessed the translocation of CPPs only with overall membrane properties,^[73] by elucidating how locally concentrated CPPs reconstitute heterogeneous lipid membranes, thereby providing insight into highly effective drug delivery.

Experimental Section

Materials

1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dioleoyl-sn-glycero-3-phospho-L-serine (DOPS), 1,2-dipalmitoyl-sn-glycero-3-phospho-L-serine (DPPS), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dimyristoyl-sn-glycero-3-phospho-(1′-rac-glycerol) (DMPG), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl) (biotin-cap-PE), and cholesterol were purchased from Avanti Polar Lipids. Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (Texas Red-DHPE) was purchased from Thermo Fischer Scientific. Other chemicals, including hexadecane, silicone oil AR20, chloroform, ethanol, sodium chloride (NaCl), sodium tetraborate, 4-2(2-hydroxymethyl)−1-piparazineethanesulfonic acid (HEPES), 1-dodecanethiol, poly-L-lysine (PLL) hydrobromide, biotinylated bovine serum albumin (biotin-BSA), streptavidin, glucose, calcein, methyl-β-cyclodextrin (MβCD), Fmoc-protected arginine, 1-Hydroxybenzotriazole hydrate (HOBT), N,N,N′,N′-Tetramethyl-O-(1H-benzotriazol-1-yl)uronium hexafluorophosphate (HBTU), Fmoc-Arg(pbf)─OH, dimethylformamide (DMF), N,N-diisopropylcarbodiimide (DIC), trifluoroacetic acid (TFA), triisopropylsilane (TIS), 3-Aminobutanoic acid (Abu), and 5,6-carboxyfluorescein (FITC) were obtained from Sigma Aldrich. Amine-reactive polyethylene glycol (PEG), mPEG–succinimidyl valerate, (MW 5000), was purchased from Laysan Bio. Rink amide MBHA resin (0.45 mmol g⁻¹) was purchased from Bead Tech. TAT (47-57) peptide (HIV-1 TAT) was purchased from AnaSpec. All the lipids were dissolved in chloroform or the chloroform/methanol (2:1 v/v), and stored at −20 °C. NaCl and HEPES were dissolved in distilled water to make the aqueous buffer containing 100 mm NaCl and 10 mm HEPES, followed by titration to pH 7.4 with 1 m NaOH solution, and stored at 4 °C.

Peptide Synthesis

All peptides were synthesized with the solid-phase peptide synthesis method. Rink amide MBHA resins (100 mg, 45 µmoL) were swollen with DMF (2 mL) in a 5 mL fritted syringe overnight. The Fmoc protecting group was removed by treatment with 20% piperidine in DMF (2 × 10 min). On the resin, arginine residues were added by a repeated standard route. In detail, for amino-functionalized resin conjugation, Fmoc-Arg(pbf)─OH (5 equiv.) were treated with 1 m HOBT (5 equiv.), 1 m HBTU (5 equiv.), and 1 m DIC (10 equiv.) in DMF. After shaking for 2 h, the reaction was finished and mixture was drained out and the resin was washed with DMF (7×). Unless noted, this washing step was repeated at each reaction step. This process was repeated until desired sequences of nona-arginine (R₉) were obtained. For cleavage, the beads were treated with 1 mL of a cleavage cocktail (95% TFA, 2.5% TIS, and 2.5% distilled water) for 2 h. To synthesize FITC-labeled R₉ (FITC-R₉), the N-terminus of R₉ was coupled with Abu linker by standard Fmoc chemistry. After deprotection, the amine group of the Abu was coupled with FITC (10 equiv.) using the peptide coupling chemistry. For cleavage, the beads were treated with 1 mL of a cleavage cocktail (95% TFA, 2.5% TIS, and 2.5% distilled water) for 2 h.

After reactions were completed, the crude products were purified by high performance liquid chromatography (HPLC), and their purity and identity were analyzed by liquid chromatography-mass spectrometry (LC-MS) and matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS). Analytical HPLC and LC-MS characterization were performed on an Agilent system with a C18 reversed-phase HPLC column (Agilent, 3.5 µm, 4.6 mm × 150 mm). A gradient elution of 10%–100% B in 7 min (keep 100% B till 13 min) was used at a flow rate of 0.7 mL mi⁻¹n (solvent A: distilled water, 0.01% TFA; solvent B: acetonitrile, 0.01% TFA). With a C18 reversed-phase column (Agilent, 5 µm, 21.2 mm × 150 mm) changing solvent composition with a linear gradient of 100% solvent A in 5 min followed by 100% solvent B in 65 min. MALDI-TOF MS was performed on 4700 Proteomics Analyzer (Applied Biosystems) and Autoflex speed LRF (Bruker) using α-cyano-4-hydroxycinnamic acid and 2,5-dihydroxybenzoic acid as matrices. Small aliquots of the obtained R₉ and FITC-R₉ were flash-frozen using liquid nitrogen and stored at −20 °C. Peptides for long-term storage were freeze-dried.

Preparation of Planar Lipid Membrane Imaging

The imaging chamber was assembled with a bottom coverslip and side slide glasses. PLL–PEG was then used to passivate the glass coverslips. PLL-PEG was prepared by conjugation of amine-reactive PEG with PLL at a molar ratio of 1:5. The reaction was performed in 50 mm sodium tetraborate pH 8.5 solution. The mixture was continuously stirred overnight at room temperature, buffer exchanged into 10 mm HEPES, 100 mm NaCl, pH 7.4 using a D-Tube Dialyzer (MWCO 6–8 kDa, Novagen), and stored at 4 °C. The 200 µL of 0.5% PLL-PEG solution was added to each imaging chamber. After 20 min of incubation, the chamber was rinsed with 10 mm HEPES, 100 NaCl, and pH 7.4 buffer.

Freestanding Planar Lipid Membranes

Freestanding planar lipid membranes were prepared using an ultra-stable freestanding planar lipid membrane array, as described in previous studies.^[37,38] First, the lipids dissolved in chloroform were mixed in a glass vial in the desired molar ratio and dried under a gentle N₂ stream. The dried lipids were dissolved in a mixture of hexadecane and silicone oil (1:1 v/v) to obtain a lipid oil solution with a total lipid concentration of 5 mm. Each oil was filtered through a 0.2 µm syringe filter (Whatman) to remove impurities before use. The lipid oil solution was sonicated for 1 h and used for experiments within 5 h.

A total of 1 mL of the aqueous buffer was injected into the PLL-PEG passivated imaging chamber, and 2 µL of lipid oil solution was gently dropped and spread on the air–aqueous buffer interface. ≈3 min after dropping the lipid oil solution, a hexagonal TEM grid (G200HEX, G150HEX, G100HEX, Gilder Grids) treated with 1-dodecanethiol for hydrophobic coating was gently placed on the air-oil interface using tweezers. For the next 2 min, the hydrophobic surface of the TEM grid was wetted with the lipid oil solution, and the holes of the grid were filled with thin oil films. The grid was submerged into the aqueous buffer using a syringe needle to position it on the PLL-PEG-coated glass surface. As the oil drained, the thickness of the oil film decreased sufficiently for zipping of the two lipid monolayers, which resulted in the formation of the planar freestanding lipid bilayers. The lipid membrane array was incubated at 45 °C for at least 30 min and cooled to 24 °C. ≈1 h after cooling, a cholesterol-methyl-β-cyclodextrin solution (4 mm, 20:1 mol%) was injected into the aqueous buffer at a concentration of 60 mm to increase the cholesterol concentration in the lipid membranes to ≈30 mol%. After ≈1 h of incubation, the prepared R₉ or FITC-R₉ solution was injected into the aqueous solution to the desired concentration. Experiments were done at 24 °C. The lipid membranes were directly visualized using an inverted fluorescence microscopy (IX73, Olympus) with an iXon EMCCD camera (X-6880, Andor Solis). Images were analyzed using the open-source program, Fiji.^[74]

GUV Preparation

The lipids dissolved in chloroform were mixed to a desired ratio at a total concentration of 1 mg mL⁻¹. For all GUV experiments, 2 mol% of biotin-cap-PE was included for vesicle tethering on the glass surface. GUVs were prepared based on the electroformation method.^[75] Briefly, the lipid mixture in chloroform was spread on indium-tin-oxide coated glass slides (30 Ω, Omniscience) and dried under vacuum for at least 2 h. Electroporation was performed at 55 °C in 200 mOsm L⁻¹ glucose solution or 200 mOsm L⁻¹ glucose solution with 10 µm calcein. The voltage was increased from 50 to 1400 mVpp for the first 30 min at 10 Hz, and then held at 1400 mVpp for 120 min. For the last 30 min, voltage and frequency were adjusted to 2200 mVpp and 5 Hz, respectively.

Coating of Glass Surface for GUV Tethering

GUVs were tethered on a glass coverslip using the strong interaction between biotin and streptavidin. The imaging chamber was assembled with a coverslip and 2 mm thick silicon gaskets with 3 mm diameter holes. Prior to chamber assembly, gaskets, and coverslips were cleaned with 2 vol% Micro-90 concentrated cleaning solution (International Products Corporation), rinsed thoroughly with water, and dried under a nitrogen stream. 10 µL of 1 mg mL⁻¹ biotin-BSA solution was added to the chamber and incubated for at least 30 min. Excess biotin-BSA was washed with 10 mm HEPES, 100 mm NaCl, and pH 7.4 buffer by gentle pipetting. Next, 0.2 mg mL⁻¹ streptavidin was added to the chamber, incubated for at least 30 min, and then rinsed with the same buffer.

Peptide Internalization Assay and Fluorescence Leakage Assay with GUVs

GUVs were observed using a confocal laser scanning microscope (LSM880, Carl Zeiss) in the KAIST Analysis Center for Research Advancement (KARA). To investigate R₉-induced pore formation of GUVs, calcein-encapsulated GUVs were tethered on a glass coverslip and R₉ solution was added to the desired concentration. To investigate the internalization of fluorescence-labeled peptides into GUVs, calcein-free GUVs were tethered on a glass coverslip and FITC-R₉ solution was added to the desired concentration. GUVs were exposed to the laser only when imaging to minimize the photobleaching of the fluorescence probes. Imaging was done at 20 °C and images were analyzed using the open source program, Fiji.^[74]

Statistical Analysis

The normalized area and normalized calcein fluorescence intensities inside the GUVs values were obtained by normalizing to the initial values before peptide addition. The FITC fluorescence intensities at GUV rims were normalized to the final intensity of each rim, and the FITC intensities of GUV lumen were normalized to the bulk intensity outside the GUV. The data shown in the graphs were described as mean ± SD of at least two independent experiments. The number of samples for each analysis was specified in the figure legends.

Acknowledgements

This research was supported by the Korea Health Technology R&D Project grant through the Korea Health Industry Development Institute (KHIDI) funded by the Ministry of Health and Welfare, Republic of Korea (HP20C000601), the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by Ministry of Science and ICT (NRF-2021R1A2C2009859, RS-2023-00276535).

Conflict of Interest

The authors declare no conflict of interest.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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