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. 2020 Jul 6;11(28):7408-7414.
doi: 10.1039/d0sc02234a.

Supramolecular catalysis by recognition-encoded oligomers: discovery of a synthetic imine polymerase

Luca Gabrielli  1   2 , Christopher A Hunter  1
Affiliations

Supramolecular catalysis by recognition-encoded oligomers: discovery of a synthetic imine polymerase

Luca Gabrielli et al. Chem Sci. .

Abstract

All key chemical transformations in biology are catalysed by linear oligomers. Catalytic properties could be programmed into synthetic oligomers in the same way as they are programmed into proteins, and an example of the discovery of emergent catalytic properties in a synthetic oligomer is reported. Dynamic combinatorial chemistry experiments designed to study the templating of a recognition-encoded oligomer by the complementary sequence have uncovered an unexpected imine polymerase activity. Libraries of equilibrating imines were formed by coupling diamine linkers with monomer building blocks composed of dialdehydes functionalised with either a trifluoromethyl phenol (D) or phosphine oxide (A) H-bond recognition unit. However, addition of the AAA trimer to a mixture of the phenol dialdehyde and the diamine linker did not template the formation of the DDD oligo-imine. Instead, AAA was found to be a catalyst, leading to rapid formation of long oligomers of D. AAA catalysed a number of different imine formation reactions, but a complementary phenol recognition group on the aldehyde reaction partner is an essential requirement. Competitive inhibition by an unreactive phenol confirmed the role of H-bonding in substrate recognition. AAA accelerates the rate of imine formation in toluene by a factor of 20. The kinetic parameters for this enzyme-like catalysis are estimated as 1 × 10-3 s-1 for k cat and the dissociation constant for substrate binding is 300 μM. The corresponding DDD trimer was found to catalyse oligomerisation the phosphine oxide dialdehyde with the diamine linker, suggesting an important role for the backbone in catalysis. This unexpected imine polymerase activity in a duplex-forming synthetic oligomer suggests that there are many interesting processes to be discovered in the chemistry of synthetic recognition-encoded oligomers that will parallel those found in natural biopolymers.

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Conflict of interest statement

There are no conflicts to declare.

Figures

Fig. 1
Fig. 1. Chemical structure of the AAA·DDD duplex. The complementary recognition modules are phosphine oxide H-bond acceptors (red) and 2-trifluoromethyl phenol H-bond donors. Synthesis is based on reductive amination of bisaniline linkers (green) with dialdehydes monomer building blocks. The backbone modules (black) provide a geometry compatible with duplex formation. The 2-ethylhexyl groups ensure solubility in non-polar solvents.
Fig. 2
Fig. 2. Non-covalent template-directed synthesis of recognition-encoded oligomers. The template is an amine (green) oligomer bearing recognition units (blue and red). Dynamic imine libraries can be built from dialdehydes and diamines equipped with recognition units. The template shifts the equilibrium, favouring the formation of the complementary sequence, which can be trapped as an amine oligomer.
Fig. 3
Fig. 3. Structures of the acceptor and donor dialdehyde (D, A) and dianiline (N) monomers used for building libraries of imine oligomers. Structures of monoaldehyde (D′, A′) and monoaniline (N′) monomers used as chain end capping units (R = 2-ethylhexyl, R′ = pentyl).
Fig. 4
Fig. 4. Equilibria involved in the formation of a library of imine oligomers. If no cyclic oligomers are formed, the degree of oligomerization can be controlled with the amount of dianiline. X is a recognition module, and R is a solubilizing group.
Fig. 5
Fig. 5. HPLC traces of a toluene-d8 solution of donor D (10 mM) 5 minutes after the addition of dianiline N (16 mM) in the presence of phosphine oxides. (a) 10 mM nBu3PO. The peak corresponding to the monomeric mono-imine mono-aldehyde is labelled D′. (b) 3.3 mM AAA. The imine oligomers are labelled D, DD, DDD, D4, D5etc. For each oligomer, three peaks were observed, corresponding to dialdehyde (traces), mono-imine mono-aldehyde and diimine terminal groups. HPLC method: coagent phenyl hydride 2.o 3 cm × 3 mm column, flow rate 0.4 ml min−1, injection volume 0.1 μL, 45 °C; eluent A 60% NH4OAc 10 mM, pH 5.70, 40% THF; eluent B 85% THF, 10% IPA, 5% A; 45% B for 1 min, then increased to 75% over 3.5 min, and then increased to 80% over 2 min. The UV/vis absorbance was recorded at 292–308 nm.
Fig. 6
Fig. 6. Effects of phosphine oxide oligomers on the rate of imine formation for substrates with no recognition sites. (a) Reaction of 4-pentylbenzaldehyde with 4-pentylaniline (R′ = n-pentyl). (b) Half-life of the aldehyde starting material measured by integration of 500 MHz 1H NMR signals as a function of time. Reaction conditions: 20 mM aldehyde, 40 mM aniline and 10 mM phosphine oxide in toluene-d8 (i.e. 10 mM nBu3PO, 5 mM AA, or 3.3 mM AAA). (c) Chemical structure of AA.
Fig. 7
Fig. 7. Kinetics of imine formation measured by integration of all imine signals in the 500 MHz 1H NMR spectra of a mixture of N (20 mM) and D (10 mM) in toluene-d8 in the presence of 10 mM nBu3PO (red data), 5 mM AA (yellow data) or 3.3 mM AAA (green data).
Fig. 8
Fig. 8. (a) Kinetics of imine formation measured by integration of all imine signals in the 500 MHz 1H NMR spectra of a mixture of N (20 mM) and D (10 mM) in toluene-d8 in the presence of different amounts of AAA: 0 (red), 0.83 (yellow), 1.65 (blue) or 3.3 (green) mM. The dotted lines show the linear region where the initial rate (Vinit) was measured. (b) Relationship between the initial rate (Vinit) and the concentration of AAA. The straight line of best fit is shown.
Fig. 9
Fig. 9. Kinetics of imine formation measured by integration of all imine signals in the 500 MHz 1H NMR spectra of a mixture of N (20 mM) and D (10 mM) in toluene-d8 in the presence of 3.3 mM AAA in the absence (filled circles) or presence of 10 mM 2-trifluoromethyl phenol (open circles).
Fig. 10
Fig. 10. Half-life for reactions in the presence of AAA (3.3 mM) relative to the corresponding half-life in the presence of nBu3PO (10 mM) (t1/2 (rel)). Half-lives were measured by integration of the 500 MHz 1H NMR signal due to the aldehyde as a function of time for mixtures of 20 mM total aldehyde (20 mM D′, 10 mM D, or 20 mM P) and 40 mM total aniline (40 mM N′, or 20 mM N) in toluene-d8.
Fig. 11
Fig. 11. a) Half-life for reactions in the presence of DD (5 mM, shaded data) or DDD (3.3 mM, black data) relative to the corresponding half-life in the presence of 2-trifluoromethyl phenol (10 mM) (t1/2 (rel)). Half-lives were measured by integration of the 500 MHz 1H NMR signal due to the aldehyde as a function of time for mixtures of 20 mM aldehyde (P or A′) and 40 mM total aniline (40 mM N′, or 20 mM N) in toluene-d8. (b) Chemical structure of DD.
Fig. 12
Fig. 12. Partial 500 MHz 1H-NMR spectra of a mixture of D (10 mM) and B (10 mM) in toluene-d8 solution 5 minutes after the addition of N (20 mM) in the presence of (a) AAA (3.3 mM) or (b) nBu3PO (10 mM). The signals at 10.4–10.5 ppm are due to the aldehyde protons at the end of D oligomers; the signal at 10.1–10.2 ppm is due to the aldehyde proton of the mono-imine of B.
Fig. 13
Fig. 13. Elongation of an imine oligomer of D catalysed by AAA. H-bonding interactions between phenol and phosphine oxide recognition units are required for catalysis.
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