Spontaneous formation of droplets from amino acid thioesters. We designed a monomer M that was capable of producing peptides and facilitating the self-assembly of molecules under aqueous conditions sufficiently mild to allow formation of droplets and self-reproduction of the building blocks of M (Fig. 2a). A monomer with a thioester and unprotected cysteine group at its C- and N-terminus, respectively, would be expected to polymerise spontaneously in water. Therefore, to provide M while reducing with dithiothreitol (DTT) in subsequent experiments, a disulfide precursor of M (Mpre) was synthesised (Figs. S1–S3). The C-terminus of M was capped with a benzyl mercaptan (BnSH) leaving group.
The reduction of the Mpre disulfide by DTT in water generated two M molecules, which then reacted spontaneously to yield peptides (Fig. 2a). To confirm the formation of droplets, we monitored turbidity as a function of time and recorded microscopic images of an aqueous solution containing Mpre and DTT (Figs. 2b, c). Five minutes after addition of Mpre (5 mg, 10 mM) and DTT (4 mg, 25 mM) to deionised water, the turbidity began to increase, and it continued to increase for 16 h (Figs. 2b and c [red line]). The increase in turbidity suggested the formation of molecular self-assemblies in solution. We therefore examined the solution under a differential interference contrast (DIC) microscope to confirm the formation of molecular self-assemblies induced by a series of reactions after addition of Mpre (Fig. 2d and Supplementary Movie 1). No molecular self-assemblies were observed within the first 5 min after mixing the Mpre and DTT reagents; however, micrometre-sized molecular self-assemblies appeared after 1 h. Twenty-seven hours after mixing, the spherical molecular self-assemblies had grown and were present in significant numbers. Furthermore, DIC microscopy revealed that fusion of the molecular self-assemblies (Supplementary Movie 2) had begun 1 h after addition of Mpre. The spherical shapes maintained by the fused assemblies suggested that the formed aggregates were droplets.
To confirm that the reduction of Mpre induced the development of turbidity in the solution containing both Mpre and DTT, the solution was observed in real time with a microscope, and the turbidity of a sample of the solution dispensed every hour from the initial solution was measured. When Mpre was dissolved in deionised water in the absence of DTT, no molecular assemblies larger than 1 µm were observed under a DIC microscope, although smaller assemblies resulted in the development of slight turbidity (Figs. 2c [green line] and S4a). Moreover, the turbidity was close to zero when DTT was dissolved in water because DTT is soluble in water (Fig. 2c [blue line]). In contrast to the dispersed reaction solution after addition of thioesterified cystine Mpre and DTT, no turbidity was observed when cystine was mixed with cysteine dihydrochloride, BnSH, and DTT (Figs. S4b, c) because the molecular self-assemblies did not disperse, and the aggregates precipitated in a similar way with the addition of only cystine to water (Fig. S4d). These results confirmed that droplet formation resulted from thioester-induced reactions. To clarify the contribution of the cysteine moiety in the monomers to droplet formation, we also synthesised thioesterified glycine (Gly-SBn) (Methods, Fig. S5) by a protocol similar to the synthesis of Mpre (Fig. S1). There was no turbidity in an aqueous solution containing Gly-SBn and DTT either 5 min or 24 h from its preparation (Fig. S6). These results indicated that the reduction of the disulfide precursor with DTT and the subsequent chemical reaction at the thioester site and the cysteine side chain of Mpre were essential for spontaneous droplet formation. The pH range for droplet formation was at least 3–11 (Fig. S7).
Oligomerisation-induced self-assembly of liquid–liquid, phase-separated droplets via autocatalysis. To verify that oligomerisation was induced at the thioester site and the cysteine side chain of M, we allowed mixing of Mpre (10 mM) with DTT (25 mM) to reduce Mpre and oligomerise the generated M in deionised water. The product was separated from the droplet dispersion. The reaction solution was lyophilised to remove water and BnSH, and the white powder residue was washed with acetonitrile to remove unreacted and oxidised DTT (Figs. 3a, S8). The obtained powder was analysed by proton nuclear magnetic resonance (1H NMR) (Fig. S9a) and electrospray ionisation mass spectrometry with a time-of-flight mass spectrometer (ESI-TOF-MS). Comparison of oligopeptide spectra with those of Mpre (Fig. S9b) showed that the peak of the benzene ring (peak a in Fig. S9b) and the peaks near the disulfide (peaks b and c in Fig. S9b) had almost disappeared, whereas an amide proton (d) was newly detected. The mean degree of polymerisation of the obtained powder was estimated from the ratio of the areas of the peaks at 8.8 ppm and 4.8 ppm in Fig. S9a to be 4.1. Each peak was assigned to the proton of terminal amine groups and amide bonds, respectively. In addition, the mass-to-charge ratios observed in the ESI-TOF-mass spectra revealed degrees of polymerisation of 2 to at least 4 in the reaction solution 24 h after mixing Mpre and DTT (Fig. S10). The intensities of the monomer and dimer decreased from their initial values, whereas the intensity of the trimers and tetramers tended to increase. These results indicated that Mpre was reduced by DTT to yield M, which then formed at least di-, tri-, and tetrapeptides.
To clarify the contribution of the generated oligopeptide to droplet formation, products except biproducts (BnSH and oxidized DTT), were purified from the solution 24 h after mixing Mpre and DTT, and its ability to form a droplet was investigated (Table in Fig. 3a). The solutions that contained no oligopeptides or BnSH did not become turbid, whereas those containing both oligopeptides and BnSH were dispersed, and the formation of spherical molecular assemblies in them was apparent under a DIC microscope (Fig. S11). The indication that oligomerisation-induced self-assembly had occurred in these mixtures strongly suggested that the association of oligopeptides and BnSH components was essential for LLPS-droplet formation. We therefore concluded that the droplets were formed by associative LLPS32. The fact that the terminus of the peptide has an ammonium cation and that BnSH has a benzene ring suggests that the droplet may be an associative LLPS caused by cation–π interactions. Indeed, it was reported that LLPS droplets in vivo were formed due to cation-π interactions between lysine residues with an ammonium cation and other amino acids residues with an aromatic ring in the protein side chain33.
The Mpre residual proportion, i.e., the proportion of the primary amines that was not involved in peptide formation, was determined from the amount of primary amine Mpre that was consumed (Fig. 3b), which was estimated by the fluorescamine method (Fig. S12a). The rate of formation of droplets was calculated from the changes of the areas of the peaks corresponding to benzene ring protons in the 1H NMR spectrum of the solution (Fig. S12b). The decrease of the Mpre residual proportion (Fig. S12c) was consistent with the observed increase of the droplet formation rate (Fig. 3b). The curve of the droplet formation rate was sigmoidal and was fit to the autocatalytic reaction equation using the Levenberg–Marquardt method on the assumption that the reaction was autocatalytic (Fig. S13). The autocatalytic nature of the peptide synthesis was confirmed by the observation that the shape of the curve of the Mpre residual proportion also became sigmoidal (Fig. 3c) when the amount of DTT added was decreased to reduce the reaction rate. However, the curve of the droplet formation rate was sigmoidal even though the amount of added DTT did not change because the droplet formation rate was controlled by the rate of decomposition of Mpre, and it increased at a slower rate than the rate of decrease of the Mpre residual proportion. These results indicated that LLPS droplets were formed autocatalytically, and a hypothesis that the droplets themselves served as sites of peptide generation.
Recursive self-reproduction of LLPS droplets. To demonstrate the continuous growth of the droplets upon serial additions of Mpre and DTT, we measured the changes in the size distribution of the droplets that formed after each addition. Figure 4a shows the predicted size distributions of LLPS droplets after repeated additions of Mpre and DTT. After the first addition of Mpre, the droplet size distribution was expected to shift fully to the right as the droplets grew. This expectation was confirmed by the continuous increase in the size of the LLPS droplets revealed by the droplet size analysis (Fig. 4b). This result indicated that nanometre-sized molecular aggregates were formed during the first five minutes after mixing of the Mpre and DTT. After five minutes, they grew or fused to become large enough to be observed with a microscope.
Upon subsequent addition of Mpre and DTT into the dispersion containing the LLPS droplets, the droplet size distribution was expected to change into one of two patterns, depending on the region of oligomerisation in the droplets (Fig. 4a, right). Two possible cases were considered. In the first case, if new droplets formed spontaneously in the solution as oligomerisation proceeded, then a new peak at a smaller size would appear in the corresponding distribution (Fig. 4a, upper right). In the second case, if oligomerisation occurred inside or at the interface of the LLPS droplets, the pre-existing LLPS droplets would grow larger, and no new LLPS droplets would be generated because no oligopeptides would be available in the solution. No separate peak would therefore appear in the size distribution (Fig. 4a, lower right). In the second case, some oligopeptides would also be generated outside and then incorporated into the existing LLPS droplets. To identify the actual oligomerisation site, we added equal volumes of Mpre and DTT to 1 mL of LLPS droplet dispersion 24 h after the first addition of Mpre (10 mM) and DTT (25 mM), and we then monitored the temporal evolution of the size distribution of the LLPS droplets (Fig. 4c). The sizes of the existing LLPS droplets increased with time, and no additional peak corresponding to newly formed LLPS droplets was detected. This result strongly supported the hypothesis that oligomerisation occurred inside or at the interface of the LLPS droplets: that is, these findings pointed to autocatalytic self-reproduction of the LLPS droplets due to the ability of LLPS droplets to serve as active sites for oligopeptide generation.
The LLPS droplets formed in the current study self-reproduced recursively while they were continuously nourished by consumption of Mpre and were extruded as a means of periodic dilution to induce shearing (Fig. 4d). In particular, the LLPS droplets grew and fused by autocatalytic self-reproduction and then divided upon addition of Mpre and DTT (Fig. 4e). To quantitatively evaluate the recursive growth and division of the LLPS droplets, we analysed the temporal evolution of the average diameter of LLPS droplets in the dispersion. We monitored the droplet population over six periods: the initial droplet-formation period and five cycles of Mpre addition to the existing, uniformly sized droplets (nutrient, white triangles in Fig. 4e) and extrusion using a syringe (shear, black triangles in Fig. 4e). During each cycle, we observed an increase in the size of the LLPS droplets stimulated by addition of Mpre and DTT that was followed by a decrease in size upon extrusion. From the second to the sixth period, the particle size at the beginning and end of a cycle was almost the same, and the mode of particle size development also remained approximately unchanged. Referencing the time-course analysis of the droplet size in Fig. 4e, a significant correlation exceeding the 95% confidence interval (light blue zone in Fig. S14a) was found at the 33rd lag, which corresponds to the time immediately after the nutrient was taken in. This result clearly indicated that there was a high autocorrelation between particle size changes in every cycle. Use of DIC microscopy also revealed similar recursive patterns of LLPS droplet diameters (Figs. S14b, c). The consistency up to 3 h after mixing Mpre and DTT between the increase in average droplet size (Fig. 4e) and the rate of droplet formation to the one third (Fig. 3b) (correlation coefficient = 0.96) strongly suggested that the increase in droplet size mainly depended on the chemical reaction at the initial stage. However, the fact that the correlation coefficient between the two experiments more than 3 hours after mixing decreases extremely to 0.068 suggested that the droplets generated by the reaction grew by the fusion dominantly (Fig. S15). However, the fact that no significant increase in particle size was observed when only water was added to the extruded droplet dispersion (Fig. S16) indicated that the increase in particle size at the initial stage was not due to the fusion of droplets after extrusion but instead was an effect of the reaction. The fact that almost the same size of the droplets reached a steady state during every period therefore meant that the number density of droplets, despite the effect of dilution, was kept constant by the addition of precursors. These results demonstrated that LLPS droplets underwent a recursive growth–division process, that is proliferation, in response to the external stimuli of nutrient addition and extrusion.
Nucleic acid/lipid concentration in droplets. In the origin of life, a simpler prebiotic polymer34,35 might have provided a scaffold for peptide-droplets formation before the synthesis of the common major components of current organisms, i.e. nucleic acids, lipids and proteins. However, to evolve into the ancestors of all modern organisms, proliferating peptide-CDs require to cooperate with these major components36–38. The absence of such a CD up to the present has led to a gap between the three major scenarios—the “RNA world”39, “lipid world”40, and “protein world”41—each of which envisions that a self-reproducing system of the corresponding molecules has evolved into a proliferating protocell via interactions with other molecules. We therefore tested the ability of the droplets created in this study to serve as active sites to incorporate and concentrate fluorescence-tagged nucleic acids and lipids into a droplet (Fig. 5a). Twenty-four hours after mixing Mpre and DTT, 6-carboxy-tetramethylrhodamine (TAMRA)-tagged RNA (TAMRA-RNA) and boron-dipyrromethene (BODIPY)-tagged phospholipid (BODIPY-lipid) solutions were added to the droplet dispersion. The droplets were then observed with a confocal laser scanning (CLS) microscope. No fluorescence emission from the droplets was observed two minutes after addition of the RNA and lipid solutions. Thirty minutes after addition, however, the molecular assembly gradually began to emit fluorescence derived from TAMRA and BODIPY, and that fluorescence continued for 360 min (Fig. 5b). Line profiles of the fluorescence intensity (Fig. 5c–f) were generated from each fluorescence channel image of the LLPS droplets (Fig. 5b, bottom row). The time-course of the line profiles confirmed the gradual incorporation of RNA oligomers and phospholipids into the LLPS droplets after simultaneous addition of these molecules (Figs. 5c, e). In the case of RNA, the maximum fluorescence was detected near the inner boundaries of the droplet, whereas the fluorescence peaked at the centre of the droplet in the case of the lipids. A comparison of the CLS microscopy images of the LLPS droplets during independent incorporation of the RNA oligomers or lipids with the corresponding line profiles (Figs. 5d, f) revealed positions of fluorescence intensity peaks similar to the peak positions when TAMRA-RNA (Figs. 5d, g) or BODIPY-lipid (Figs. 5f, h) was added separately to the different droplet dispersions. To show that this experiment was not affected by the fluorophore itself, we confirmed that the results were similar for experiments labelled with different fluorophores. Similar distributions of fluorescence intensity were detected for RNA, phospholipids, and DNA labelled with alternative fluorescent probes. Fluorescence derived from fluorescein isothiocyanate-tagged RNA (FITC-RNA) or DNA and TAMRA-tagged DNA (TAMRA-DNA) were detected along the inner boundary of the droplet (Figs. S18a–c), and the fluorescence of Texas Red–tagged phospholipids (Texas Red–lipid) was detected around the centre of the droplet (Fig. S18d). The interactions between nucleic acids, lipids, and peptides generated spatial heterogeneity between the RNA and lipids in the same LLPS droplet. These results implied that the droplets were heterogeneous, with a gradual hydrophilic boundary and a hydrophobic centre that were composed mainly of oligopeptides and BnSH, respectively. Raman microspectrometry revealed that the droplets were composed of a relatively hydrophobic central part with a relatively large Bn/water ratio and of a relatively hydrophilic peripheral part with a relatively small Bn/water ratio (Fig. S19).
When only phospholipids were added, their surfactant effect resulted in a reduction in the size of the droplets (Figs. 5h, f); however, no significant decrease of droplet size was observed upon addition of RNA and lipid (Figs. 5b, c, e) or of RNA (Figs. 5d, g). This result suggested that the incorporation of nucleic acids enabled the proliferating droplet to maintain its size despite the perturbation associated with lipid addition. To confirm that the self-maintenance ability associated with the incorporation of nucleic acids was ubiquitous, we mixed the droplet dispersion with a TAMRA-RNA solution and BODIPY-lipid dispersion or with RNase-free water, followed by a 24-h incubation and fluorescence-activated cell sorting (FACS) analysis. Population analysis of the FACS data after addition of either fluorescence-labelled RNA or fluorescence-labelled lipids indicated that the fluorescence intensity, which corresponded to the amount of RNA or lipids incorporated by the droplet, increased in both cases, and that the width value of forward scattering pulse, which corresponds to the droplet size, decreased only when lipid was added (Fig. S20). These results strongly suggested that RNA suppressed any reduction of particle size upon lipid addition. Collectively, these results indicated that the localisation of RNA oligomers near the interface of the droplets contributed to the self-maintenance of the droplets. Raman spectroscopy revealed that the hydrophobic ratio was higher in the centre of the droplet than in the periphery of the droplet that contained the non-fluorescent labelled nucleic acids and lipids (Fig. S21). Compared with the droplets without addition of biomolecules (Fig. S19), it was suggested that the water inside the droplets was replaced by hydrophilic DNA and amphiphilic phospholipids. The heterogeneity of the internal structure of the droplet led to the self-maintenance of the droplet, because the nucleic acid was localised in the peripheral part of the droplet, where it contributed to the undercoat structure. The possibility that this localisation was caused by size exclusion as well as by the hydrophilic/hydrophobic balance of inner molecules cannot be dismissed.
To evaluate the effects of oligopeptides on the incorporation and concentration of hydrophilic RNA oligomers and amphiphilic (rather than hydrophobic) phospholipids into the BnSH phases that could be regarded as a main component of the droplet centre, we measured the decrease in the fluorescence intensities of aqueous TAMRA-RNA or BODIPY-lipid solutions layered for 24 h over the BnSH solution via high-sensitivity fluorescence spectroscopy with photon-counting detectors. The fluorescence intensity in the aqueous phase decreased in both the TAMRA-RNA solution and BODIPY-lipid solution (Fig. S22a), but there was a difference between the two in the magnitude of the decrease of fluorescence in the presence of the oligopeptides (Figs. S22a, c). In the case of the TAMRA-RNA solution, the fluorescence intensity in the presence of the oligopeptides was less than one-third the intensity in the absence of the oligopeptides, whereas in the case of the BODIPY-lipid solution, there was little difference in the fluorescence intensity in the presence or absence of the oligopeptides. A similar result was obtained when the fluorescence moiety was changed (Figs. S22b, d). These results indicated that RNA oligomers and phospholipids were not only incorporated but also concentrated in the BnSH phases, and that oligopeptides could further enhance the RNA enrichment of the droplets because of the high permeability of the membraneless structure of the LLPS droplets.