Self-replicators from dynamic molecular networks: selection, competition and subsystem
coupling
Komáromy, Dávid
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Chapter 3
Effector-triggered self-replication in coupled
subsystems
In living systems processes like genome duplication and cell division are carefully
synchronized through sub-system coupling. If we are to create life de-novo, similar control
over essential processes such as self-replication need to be developed. Here we report that
coupling two dynamic combinatorial subsystems, featuring two separate building blocks,
enables effector-mediated control over the onset of self-replication. The subsystem based
on the first building block shows only self-replication, whereas that based on the second
one is solely responsive toward a specific external effector molecule. Mixing the subsystems
arrests replication until the effector molecule is added, resulting in the formation of a
host-effector complex and the liberation of the building block that subsequently engages in
self-replication. The self-replication processes show the same characteristics at the molecular
as well as the supramolecular level, independent of subsystem coupling, demonstrating the
modularity of the approach. The onset, rate and extent of self-replication is controlled by
the amount of effector present.
This chapter has been published:
D. Komáromy, M. Tezcan, G. Schaeffer, I. Marić, S. Otto Angew. Chem. Int. Ed. 2017, 56,
14658-14662.
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3.1. Introduction
Minimal living systems are generally believed to feature three main characteristics: replication, compartmentalization and metabolism.[1,2] Efforts towards the construction of fully synthetic living
systems have focused mostly on isolated subsystems capable of self-replication[3–6] or compartment
formation.[7–9] The functional integration of these subsystems[10] represents an arduous challenge and
new concepts for achieving subsystem coupling are badly needed. In this context, systems of replicators for which the onset and extent of replication can be triggered and controlled by effector molecules are highly relevant research targets. However, triggered self-replication has received only little attention and currently triggers are limited to light[11,12] and stabilization of the replicator by binding.[13]
Scheme 3. 1. Dynamic Combinatorial Library made from 1 and 2 and Guest 3.
We envisaged that triggered replication systems may also be achieved using a conceptually different modular approach, by coupling two subsystems: one devoted to replication and another to effector recognition. The coupling of these two chemical subsystems may be realized through dynamic processes (i.e. chemical equilibria) which can interfere with each other via common components; i.e. dynamic
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molecular networks or dynamic combinatorial libraries.[14] In such networks building blocks, appended
with functional groups capable of reversible covalent bond formation, react with each other to form a diverse collection of oligomeric compounds, a dynamic combinatorial library (DCL). The members of such libraries constantly exchange building blocks via the formation and breakage of the reversible covalent bonds, i.e. the distribution of the members is governed by a large number of interconnected equilibria. Consequently, perturbing one equilibrium in one part of the network in principle affects the distribution of all DCL members, including those that are only indirectly connected to the perturbed dynamic process. Coupling of thermodynamically controlled subsystems in the form of such signal-cascading networks has advanced experimental[15] and theoretical[16] underpinnings.
Individually, many dynamic combinatorial systems that feature effector recognition through host-guest chemistry have been reported.[17–19] In such systems guest addition leads to the selective
amplification of those library members that bind the guest. More recently, also dynamic combinatorial systems showing self-replication have been developed.[20–30] In such systems binding of specific library
members to copies of themselves leads to their proliferation in an often autocatalytic manner. However, no functional integration of guest recognition and replication subsystems has yet been reported.
We now show how an effector molecule can trigger self-replication in a DCL which is composed of two building blocks: the subsystem constructed from the first serves as a platform for self-replication, whereas the second building block is responsible solely for effector recognition.
3.2. Results and Discussion
Recently we reported that in disulfide-based[31] dynamic combinatorial libraries of dithiol building
block 1 (Scheme 3. 1) stirring induces the emergence of a self-replicator (see previous chapter) in the form of the cyclic hexamer (16), self-assembling first into nanoribbons, then transforming into
nanoplatelets.[32] We set out to couple this self-replicating system to a host-guest system also established
previously[33] in which spermine guest (3) is able to amplify the formation of a cyclic tetramer host (2 4) in
a DCL made from building block 2. We reasoned that, upon oxidation of a mixture of thiols 1 and 2, a DCL of various cyclic disulfide oligomers should be formed in which assembly nucleation, and thereby replicator emergence, is hampered by the low concentration of the replicator in the mixture. Upon addition of effector 3, the cyclic tetramer 24 should be amplified, leading to the depletion of building block
2 from the DCL. This, in turn allows the library to re-equilibrate, leaving behind 1-only oligomers, besides
the 24·3 complex. Nucleation of the 16 replicator should now be more likely and the continued stirring of
this re-equilibrated library would lead to the emergence of self-replicator 16 through a growth-breakage
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Figure 3. 1.UPLC traces of a DCL composed of equimolar amounts of 1 and 2 ([1] = [2] = 2.0 mM) in aqueous borate buffer (50 mM, pH 8.2) (A) after 2 days of stirring and (B) subsequent addition of 0.50 equivalents (1.0 mM) of 3 and 1 day in the absence of agitation. (C) After 2 days of stirring, followed by addition of 0.5 equivalents (1.0 mM) of 3 and 1 day of stirring.
Indeed, stirring building block 1 and 2 ([1] = [2] = 2.0 mM) in aqueous borate buffer (50 mM, pH 8.2) gave a DCL composed mainly of mixed trimers and tetramers, alongside larger macrocycles, but essentially devoid of replicators (Figure 3. 1A). However, addition of 0.5 equivalent of 3 (relative to 1) and subsequent stirring for 1 day induced a dramatic change in the library composition which was now dominated by self-replicator 16 and spermine binder 24 (Figure 3. 1C). Interestingly, when the same
experiment was repeated, but the mixture was not agitated following the addition of effector 3 the most abundant 1-containing species were 13 and 14 (Figure 3. 1B), which are also the main products in a
non-agitated DCL prepared from 1 only (see previous chapter). These results show that upon addition of 3, the host-guest complex 24.3 is formed first, accompanied by the buildup of a range of 1-only oligomers.
It also shows that in the coupled subsystems the emergence of the replicator is promoted by mechanical agitation, as is the case for the separate replication subsystem.
In order to prove that self-replication is indeed a consequence of subsystem coupling and only happens when both stimuli (addition of 3 and stirring) are present, control experiments were performed.
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First, in the absence of 3, mechanical agitation of a DCL made from equimolar amounts of 1 and 2 showed only trace amounts of the replicator, even upon 4 months incubation (see Figure S3. 4). Interestingly, when 1 was applied in excess, stirring led to a DCL featuring mixed macrocycles, as well as self-replicator 16 (see Figure S3. 2). Second, in the absence of mechanical agitation, addition of 3
resulted in a DCL featuring 24 as the main 2-containing species, whereas the relative amounts of other
oligomers featuring 2 decreased substantially (Figure 3. 1B). Furthermore, the presence of 3 did not have any impact on the behavior of the DCL made from 1: mechanical agitation resulted in the formation of 16, whereas in the absence of stirring, trimers, tetramers and larger macrocycles formed (see Figure
S3. 1).
Figure 3. 2. Time evolution of a DCL composed of equimolar amounts of 1 and 2 ([1] = [2] = 2.0 mM) in aqueous borate buffer (50 mM, pH 8.2) after the addition of 3 (0.9 equivalents) at t = 0 h.
In order to provide further mechanistic insight into the triggered replication process, the time evolution of the stirred system following addition of 3 was monitored (Figure 3. 2). First, 24 appeared almost
instantaneously after the addition of 3, showing that its formation was the first event in the reaction cascade. The concentration of 24 reached its equilibrium value at a time scale (<12 hours; see also
Figure S3. 6) that is one order of magnitude smaller than timescale of the replication of 16 (140 hours).
Thus, the addition of 3 is indeed a trigger. Second, simultaneously with the appearance of 24, the relative
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formation of the self-replicator 16. This result shows that the formation of 16 proceeded via intermediates
13 and 14. Third, the increase in the relative amount of 16 showed a lag phase, indicative of a slow initial
nucleation event, followed by a rapid increase, most likely arising from an autocatalytic breakage-elongation mechanism as described before.[28]
Figure 3. 3. Example of the time evolution of a DCL composed of equimolar amounts of 1 and 2 (2.0 mM each) in aqueous borate buffer (50 mM, pH 8.2), upon addition of increasing amounts (0.3, 0.6 and 0.9 mM) of 3. Two repeats of this experiment are shown in the SI.
We then explored whether the trigger allows also quantitative control over the onset, rate and extent of replication. Thus, we prepared libraries with the same initial composition as described before ([1] = [2] = 2.0 mM), added different amounts of effector 3 (0.3, 0.6 and 0.9 mM) and monitored the time evolution of the system with UPLC. This set of experiments was carried out in three repeats and the overall trends were reproducible (see Figure S3. 9). One of the three datasets is shown in Figure 3. 3, revealing that the lag phase, the replication rate and the extent of replication all depend strongly on the amount of effector present. These findings can be interpreted as follows: The higher the effector concentration, the larger the amount of 2sequestered as 24-effector complex, and consequently, the larger the amount of
1-only containing macrocycles accessible for replication, leading to a higher final replicator
concentration. Furthermore, an increasing concentration of 1-only containing macrocycles increases the chance of nucleation of assemblies of 16, hence the shortening of the lag phase prior to replication.
Finally, the higher rate of replication at higher effector concentrations reflects the higher achievable concentration of the autocatalytic replicator. Another factor that contributes to the higher replication rate
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induced by 3 is the fact that fiber growth is faster when the composition of the trimer and tetramer macrocycles is biased towards those rich in 1. This is corroborated by the observation that, when the distribution of the two building blocks over the trimer and tetramer rings is 1:1, no significant replicator growth occurs in the course of 30 hours even upon seeding with as much as 60% pre-formed replicator (see Figure S3. 8).
Figure 3. 4. Negative stain transmission electron micrographs of a sample (A) containing 16 and (B) prepared by addition of 3
(0.5 eq) to an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) after 10 days of stirring. The insets show the height distributions along the directions highlighted by the arrows.
Next, we verified whether the process of effector-induced triggering of self-replication interfered with supramolecular organization of the replicator. Transmission electron microscopy (see Figure 3. 4A and B) and atomic force microscopy (Figure 3. 4C and D) show nanoribbons with dimensions that are very
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similar for the isolated replication subsystem as compared to the case when the subsystems are coupled. Analyses on solutions containing only the host-effector complex 24.3 did not reveal any nanoscale objects
detectable by TEM; and the mixing of 24.3 and pre-formed nanoribbons of 16 revealed nanoribbons
similar to those of 16 by itself (see Figure S3. 7). Thus, subsystem coupling does not appear to affect the
molecular and supramolecular characteristics of the replication process, confirming the modular character of the system and the compatibility between the subsystems.
We further tested subsystem compatibility and whether our approach of triggered replication can be extended to structurally related replication systems. Thus, we prepared DCLs by mixing building blocks
2 + 4 and 2 + 5. Building blocks 4 and 5 contain one ethylene oxide unit less and more than 1,
respectively. Oxidation of compound 4 by itself produces the self-replicating cyclic tetramer 44. In
contrast, compound 5 by itself forms trimers and tetramers, together with a range of large macrocycles and no replicator emerges.[34]
Figure 3. 5. UPLC traces of stirred DCLs composed of equimolar amounts of 4 (A, B) or 5 (C, D) and 2 ([4] = [5] = [2] = 2.0 mM) in aqueous borate buffer (50 mM, pH 8.2) before (A, C) and 1 day after (B, D) the addition of 0.50 equivalents (1.0 mM) of 3.
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In the DCL composed of building blocks 2 and 4, addition of 3 triggered the exclusive formation of replicator 44 (Figure 3. 5B). Analogously, upon mixing building blocks 2 and 5, addition of 3 resulted in
the formation of receptor 24 and the formation of a large diversity of macrocycles (up to 12mers),
composed almost solely of 5 (Figure 3. 5D). These results demonstrate the compatibility of the effector binding subsystem with the subsystems based on 4 and 5 and the transferability of the concept of triggered replication to the tetramer-based replicator made from building block 4.
Figure 3. 6. UPLC traces of DCLs (pH = 8.2, aqueous borate buffer) prepared by mixing a fully oxidized mixture of 1 and 2 ([1] = [2] = 2.0 mM) after partial reduction (10%) by TCEP and 3 days of stirring in the presence of 0.50 equivalents (1.0 mM) of A) spermine (3) B) spermidine C) putrescine D) n-butylamine E) N,N-dimethylbutylamine. Note that efficiency of spermidine and putrescine as an effector is similar to that of spermine (although the relative amount of mixed species is slightly higher), whereas n-butylamine and N,N-dimethylbutylamine do not amplify the receptor 24 and consequently do not trigger self-replication.
As effector specificity is an important characteristic of biological signaling systems, we also investigated whether the ability to trigger self-replication was specific regarding spermine (3) as an effector. We found that among four different, structurally related amines, those that feature the structural moiety required for binding to receptor 24[33] (two secondary amine groups separated by a chain of four
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aliphatic carbon atoms, see Figure 3. 6A-C) are effective triggers, whereas the ones lacking this motif do not induce guest complexation and self-replication (see Figure 3. 6D-E).
Figure 3. 7. Alternating additions of portions of 3 and 2 to an initially equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) in aqueous borate buffer (50 mM, pH = 8.2), resulting in switching between “mixed” (low mol% of 16 ) and self-sorted (high mol% of 16) states of the system.
Finally, we show that the system can be switched between the self-sorted state featuring replicator
16 and the “mixed” state mostly devoid of replicator (Figure 3. 7 and Figure S3. 10). These studies were
motivated by the observation that, upon addition of one equivalent of 2 to a sample containing only 16,
this replicator rapidly disintegrates and a DCL dominated by mixed macrocycles is formed (see Figure S3. 5). We performed a similar experiment, but now starting from an equimolar mixture of 1 and 2 (2.0 mM), which gave rise to a mixed DCL, featuring numerous species (Figure 3. 1A). Addition of 0.25 equivalents of 3 resulted in the formation of a self-sorted DCL, consisting nearly exclusively of 16 and 24
(Figure 3. 1C). When 1 equivalent of 2 was added to a self-sorted DCL (containing 1:2:3 in a 2:2:0.5 molar ratio), the composition of the resulting library was similar to the initial one, featuring only trace amounts of 16. However, ca. 1 equivalent of 24 was still present in the mixture, reflecting the fact that the
binding between 24and 3 is strong. The resulting mixture again showed self-sorting upon addition of 0.5
equivalents of 3 and the resulting self-sorted DCL could again be switched back to the mixed state. Only two switching cycles could be accomplished. Upon the third addition of 3, self-sorting was incomplete and substantial amounts of mixed species were still present. These results demonstrate that the system
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can be alternated between replicator-rich and replicator poor states depending on the ratio between effector and building blocks, but also point to the need of controlling the amount of accumulated waste products in switching systems of this chemical nature.
3.3. Conclusions
In conclusion, we have demonstrated effector-triggered self-replication by coupling a host-guest subsystem with a self-replication subsystem via dynamic combinatorial disulfide chemistry. Mixing the two building blocks that correspond to the two subsystems results in a diverse library, essentially devoid of replicator. Addition of an effector molecule results in binding between an oligomer of the replicating building block and the effector molecule. As a result, the library is depleted of the non-replicating building block, making replication possible. The quantitative characteristics of the self-replication process (length of the lag phase, rate and maximal extent of self-replication) depend on the amount of effector present. Triggering replication by subsystem coupling has an important advantage over previously reported triggered replication systems[11–13] by being modular: the replication system can
in principle evolve independently of the triggering system, since, in contrast to earlier reports, the triggering system does not directly involve the replicating molecules. This method paves the way for better temporal (and possibly also spatial) control over self-replication in dynamic systems. Furthermore, the control over replication through by a single effector provides a handle through which the replication system could potentially be integrated with further subsystems, with the prospect of creating supersystems with more intricate emergent properties, such as negative feedback and oscillatory behavior.
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3.4. Supporting Information
3.4.2. General Procedures
All chemicals, unless otherwise stated, were purchased from Sigma-Aldrich and used as received. Acetonitrile (ULC-MS grade), water (ULC-MS grade) and trifluoroacetic acid (HPLC grade) were purchased from Biosolve BV. The synthesis and characterization of the compounds 1, 4 and 5 was described in the previous chapter. UPLC vials, caps and inserts were purchased from Aireka Cells (Singapore).
Library preparation
For the preparation of the initial DCLs, a solution of 2 (2 mM) in aqueous borate buffer (50 mM, pH = 8.2) was added to the required amount of solid 1 in a 1.5 mL UPLC vial. The solution was stirred at 880 rpm until no further change was observed in the library composition by UPLC (2-3 days).
In a typical triggering experiment, 400 μL of a so-prepared DCL, containing 1 and 2 in equimolar amounts ([1] = [2] = 2.0 mM) was partially reduced with a 20 mM solution of tris(carboxyethyl) phosphine hydrochloride (TCEP) in borate buffer in order to liberate a small amount of thiol to facilitate thiol-disulfide exchange. The extent of reduction was 15 mol% regarding the total amount of disulfides (i.e. 12 μL TCEP solution was required). Subsequently, the required amount of 3 (or an other effector) was added (as 32 mM solution in aqueous borate buffer) and the solution was stirred at 880 rpm.
In the experiments shown in Figure 3. 7, building block 2 was added as a 4.0 mM solution in aqueous borate buffer to the self-sorted DCL samples, whereas effector 3 was added as described in the previous paragraph.
Sample preparation
Generally, samples were diluted to 200 μM with a 7:3 V:V mixture of water (UPLC grade) and isopropanol (HPLC grade), containing 0.1 V/V% TFA (HPLC grade) and 5 μL of this diluted sample was injected for UPLC measurements. For the experiments shown in Figure 5, 5 μL of the original library was transferred into a 250 μL insert contained within a 1.5 mL screw-caped UPLC vial, the sample was diluted by the UPLC instrument with 20 μL of a 1:1 V:V mixture of water (UPLC grade) and isopropanol (HPLC grade) 1 minute prior to measurement and 5 μL of this sample was injected for UPLC measurements UPLC analysis
UPLC measurements were performed on a Waters Acquity H-class system equipped with a PDA detector, at a detection wavelength of 254 nm. UPLC analyses, unless otherwise stated, were performed on an HSS T3 1.8 μm column (100 Å, 150 × 2.1 mm, purchased from Waters) using ULC-MS grade water (eluent A) and ULC-MS grade acetonitrile (eluent B), each containing 0.1 V/V % TFA as a modifier. A flow rate of 0.3 mL/min and a column temperature of 35 °C were applied.
Method for the analysis of DCLs shown in the main text: t / min % A 0 90 0.5 90 3 70 9 38 12 34 18 5 19 5 20 90 23 90
For the control experiments shown in Figure S3. 1, an Aeris WIDEPORE 3.6 μm XB-C18 column (150 × 2.1 mm, purchased from Phenomenex) was used, with the following method:
t / min % A 0 60 7 38 10 34 17 5 18 5 19 60 20 60
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For the experiments shown in Figure S3. 5 and Figure S3. 10, an HSS T3 1.8 μm column (100 Å, 150 × 2.1 mm, purchased from Waters), was used with the UPLC method identical to those used for the experiments shown in Figure S3. 1. UPLC-MS analysis
UPLC-MS measurements were performed using a Waters Acquity UPLC H-class system coupled to a Waters Xevo-G2 TOF. The mass spectrometer was operated in the positive electrospray ionization mode with the following ionization parameters: capillary voltage: 2.5 kV, sampling cone voltage: 30 V, extraction cone voltage : 4 V, source gas temperature: 150°C, desolvation gas temperature: 500°C, cone gas flow (nitrogen): 5 L/h, desolvation gas flow (nitrogen): 800 L/h.
Negative-staining Transmission Electron Microscopy
Samples were diluted to 60-fold using UPLC grade water. A small drop (5 µL) of sample was then deposited on a 400 mesh copper grid covered with a thin carbon film (supplied by Agar Scientific). After 30 seconds, the droplet was blotted on filter paper. The sample was then stained with a solution of 2% uranyl acetate (4 µL) deposited on the grid, subsequently washed and blotted on filter paper after 30 seconds. The staining procedure was repeated a second time, this time without the washing and blotting step. The grids were observed in a Philips CM12 electron microscope operating at 120 kV. Images were recorded on a slow scan CCD camera.
Atomic Force Microscopy
AFM samples were prepared by depositing 100 µL of the sample (diluted to 10 µM with respect to building block with UPLC grade water) onto a clean mica surface (Grade V1, Van Loenen Instruments). Subsequently, the solvent was evaporated in a gentle stream of air in ca. 20 minutes. The surface was then washed with 100 µL of UPLC grade water and blotted into a piece of paper twice and finally air-dried. The AFM measurements have been performed using a Bruker Multimode 8 instrument in Scan Asyst-Air imaging mode. Measurements were performed in air at room temperature. As a probe, a ScanAsyst Air (Bruker) silicon tip on a nitride cantilever was used with the following parameters: length: 115 µm, width: 25 µm, resonance frequency: 70 kHz, force constant: 0.4 N/m. The images were recorded with frequencies between 0.5 and 1.5 Hz and analyzed with NanoScope Analysis 1.50 software (Bruker Corporation, 2015).
3.4.3. LC-MS analyses of DCLs
From environmental reasons, the large number of LC-MS spectra are not published in the printed version of this thesis. However, they are accessible free of charge in the online Supporting Information of the original publication.
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3.4.4. Control Experiments
Figure S3. 1. UPLC traces of non-agitated (A, B) and stirred (C, D) DCLs prepared from 1 (2.0 mM) in aqueous borate buffer (50 mM, pH = 8.2) in the absence (A, C) and in the presence (B, D) of 3 (1.0 mM).
Figure S3. 2. UPLC traces of stirred DCLs (pH = 8.2, aqueous borate buffer) prepared from 2.0 mM 2 and A) 2.0 mM B) 3.0 mM, C) 4.0 mM D) 5.0 mM, E) 6.0 mM 1.
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Figure S3. 3. UPLC traces of stirred DCLs (pH = 8.2, aqueous borate buffer) prepared from A) an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) followed by the addition of 3 (1 mM) and additional stirring and B) an equimolar mixture of 2 and 3 ([2] = [3] = 2.0 mM).
Figure S3. 4. UPLC trace of a 4 months old stirred DCL (pH = 8.2, aqueous borate buffer) prepared from an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM).
Figure S3. 5. UPLC traces of stirred DCLs (pH = 8.2, aqueous borate buffer) prepared by A) stirring 1 ([1] = 6.0 mM) for several days to give mainly 16 B) mixing the previous solution with an equimolar amount of 2 and stirring for 10 minutes and C) mixing
100
Figure S3. 6. Time evolution of a fully oxidized sample of 2 (2.0 mM) after addition of 1 equivalent of 3.
Figure S3. 7. Negative-stain TEM micrographs of a sample prepared A) by addition of 3 (0.50 eq) to an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) after 10 days of stirring, B) by mixing a 2.0 mM solution of 24.3 and a DCL containing solely 16 ([1] = 2.0 mM), C) from an equimolar amount of 1 and 2 ([1] = [2] = 2.0 mM) after 10 days of stirring (the DCL composition was similar to that shown in Figure 1A) D) from an equimolar amount of 2 and 3 ([2] = [3] = 2.0 mM; according to UPLC analysis, the only species observed was 24). Note the similarity of assemblies in panels A) and B), as well as the absence of any type of nanoscale assemblies in panels C) and D).
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3.4.5. Additional Experiments
Figure S3. 8. Time evolution of DCLs prepared from an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) after 2 days of stirring and by subsequent seeding (addition) with A) 20 mol % B) 40 mol % C) 60 mol % of 16 at t = 0 minutes (black and red traces correspond to the relative peak areas 16 of and 24, respectively). Note that the amount of 16 does not grow after seeding.
Figure S3. 9. Time evolution of DCLs prepared from an equimolar mixture of 1 and 2 ([1] = [2] = 2.0 mM) after adding 0.3, 0.6 and 0.9 mM of 3. A), B) and C) show the results from three independent repeats.
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Note that while the qualitative properties of the time evolution process are similar in all cases, their quantitative properties differ significantly. This low reproducibility might arise from several factors. First, the system is heterogeneous and the size distribution of the nanoribbons formed by 16 are polydisperse, as clearly shown in our previous publication on the 1-only
system[32] (see previous chapter, Figures 5 and 6). Thus, the amount of catalytically active nanoribbon edges is quite variable
from sample to sample; hence the high variability in the length of the growth regime. Second, nucleation events which determine the onset of replication are stochastic, resulting in low reproducibility of the length of the lag phase. Third, oxidation of free thiols proceed independently from nucleation and assembly. We already saw that the length of the lag phase can significantly vary, it is highly possible that at the point where replication sets on, the system is essentially devoid of free thiols. However, disulfides undergo bond exchange much more slowly in the absence of thiols as in the presence thereof. Thus, in this case, bond exchange and replicator formation might in practice be halted at lower replicator content, which might explain the variability in the extent of replication.
Figure S3. 10. UPLC chromatograms of DCLs prepared by mixing building blocks A) 1 and 2 in equimolar ratios ([1] = [2] = 2.0 mM), B) addition of 0.25 equivalents (0.5 mM) of 3 to the previous mixture, C) addition of 1 equivalent (2.0 mM) of 2 to the previous mixture, D) addition of 0.25 equivalents (0.5 mM) of 3 to the previous mixture, E) addition of 1 equivalent (2.0 mM) of 2 to the previous mixture, F) addition of 0.5 equivalents (1.0 mM) of 3 to the previous mixture. All chemicals were added in borate buffer 950 mM, pH = 8.2).
3.5. Author Contributions
D. K. and G. S. conceived the project, D. K, G. S and S. O. designed the experiments. D. K. prepared the libraries, performed UPLC and LC-MS analysis, M. T. performed TEM analyses, I. M. and D. K. performed AFM analyses, D. K. analyzed the data and D. K. and S. O. wrote the manuscript.
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3.6. References
[1] T. Gánti, The Principles of Life, Oxford University Press, 2003.
[2] K. Ruiz-Mirazo, C. Briones, A. de la Escosura, Chem. Rev. 2014, 114, 285–366. [3] V. Patzke, G. von Kiedrowski, Arkivoc 2007, (5), 293–310.
[4] A. Vidonne, D. Philp, Eur. J. Org. Chem. 2009, 2009, 593–610.
[5] A. J. Bissette, S. P. Fletcher, Angew. Chem. Int. Ed. 2013, 52, 12800–12826. [6] H. Duim, S. Otto, Beilstein J. Org. Chem. 2017, 13, 1189–1203.
[7] P. Stano, P. L. Luisi, Curr. Opin. Biotechnol. 2013, 24, 633–638. [8] J. C. Blain, J. W. Szostak, Annu. Rev. Biochem. 2014, 83, 615–640. [9] P.-A. Monnard, P. Walde, Life 2015, 5, 1239–1263.
[10] K. Kurihara, M. Tamura, K. Shohda, T. Toyota, K. Suzuki, T. Sugawara, Nat. Chem. 2011, 3, 775–781. [11] Z. Dadon, M. Samiappan, E. Y. Safranchik, G. Ashkenasy, Chem. Eur. J. 2010, 16, 12096–12099. [12] Z. Dadon, M. Samiappan, N. Wagner, G. Ashkenasy, Chem. Commun. 2012, 48, 1419–1421. [13] P. Nowak, M. Colomb-Delsuc, S. Otto, J. Li, J. Am. Chem. Soc. 2015, 137, 10965–10969. [14] S. Otto, Acc. Chem. Res. 2012, 45, 2200–2210.
[15] V. E. Campbell, X. de Hatten, N. Delsuc, B. Kauffmann, I. Huc, J. R. Nitschke, Nat. Chem. 2010, 2, 684–687. [16] P. T. Corbett, J. K. M. Sanders, S. Otto, Angew. Chem. Int. Ed. 2007, 46, 8858–8861.
[17] F. J. Rizzuto, J. R. Nitschke, Nat. Chem. 2017, 9, 903–908.
[18] O. Shyshov, R.-C. Brachvogel, T. Bachmann, R. Srikantharajah, D. Segets, F. Hampel, R. Puchta, M. von Delius, Angew.
Chem. Int. Ed. 2017, 56, 776–781.
[19] A. M. Valdivielso, F. Puig-Castellvi, J. Atcher, J. Sola, R. Tauler, I. Alfonso, Chem. Eur. J. 2017, 23, 10789–10799. [20] J. W. Sadownik, D. Philp, Angew. Chem. Int. Ed. 2008, 47, 9965–9970.
[21] R. Nguyen, L. Allouche, E. Buhler, N. Giuseppone, Angew. Chem. Int. Ed. 2009, 48, 1093–1096.
[22] C. Chen, J. Tan, M.-C. Hsieh, T. Pan, J. T. Goodwin, A. K. Mehta, M. A. Grover, D. G. Lynn, Nat. Chem. 2017, DOI 10.1038/nchem.2737.
[23] V. Del Amo, D. Philp, Chem. - A Eur. J. 2010, 16, 13304–13318.
[24] V. C. Allen, C. C. Robertson, S. M. Turega, D. Philp, Org. Lett. 2010, 12, 1920–1923.
[25] J. M. A. Carnall, C. A. Waudby, A. M. Belenguer, M. C. A. Stuart, J. J.-P. Peyralans, S. Otto, Science 2010, 327, 1502– 1506.
[26] M. Malakoutikhah, J. J.-P. Peyralans, M. Colomb-Delsuc, H. Fanlo-Virgós, M. C. A. Stuart, S. Otto, J. Am. Chem. Soc. 2013, 135, 18406–18417.
[27] G. Leonetti, S. Otto, J. Am. Chem. Soc. 2015, 137, 2067–2072.
[28] M. Colomb-Delsuc, E. Mattia, J. W. Sadownik, S. Otto, Nat. Commun. 2015, 6, 7427.
[29] T. Kosikova, N. I. Hassan, D. B. Cordes, A. M. Z. Slawin, D. Philp, J. Am. Chem. Soc. 2015, 137, 16074–16083. [30] J. W. Sadownik, E. Mattia, P. Nowak, S. Otto, Nat. Chem. 2016, 8, 264–269.
[31] S. P. Black, J. K. M. Sanders, A. R. Stefankiewicz, Chem. Soc. Rev. 2014, 43, 1861–1872.
[32] D. Komáromy, M. C. A. Stuart, G. Monreal Santiago, M. Tezcan, V. V. Krasnikov, S. Otto, J. Am. Chem. Soc. 2017, 139, 6234–6241.
[33] L. Vial, R. F. Ludlow, J. Leclaire, R. Pérez-Fernandez, S. Otto, J. Am. Chem. Soc. 2006, 128, 10253–10257.
[34] D. Komáromy, M. C. A. Stuart, G. Monreal Santiago, M. Tezcan, V. V. Krasnikov, S. Otto, J. Am. Chem. Soc. 2017, 139, 6234–6241.
