Alternative design strategies to help build the enzymatic retrosynthesis toolbox

Abstract

Most of the complex molecules found in nature still cannot be synthesized by current organic chemistry methods. Given the number of enzymes that exist in nature and the incredible potential of directed evolution, the field of synthetic biology contains perhaps all the necessary building blocks to bring about the realization of applied enzymatic retrosynthesis. Current thinking anticipates that enzymatic retrosynthesis will be implemented using conventional cell-based synthetic biology approaches where requisite native, heterologous, designer, and evolved enzymes making up a given multi-enzyme pathway are hosted by chassis organisms to carry out designer synthesis. In this perspective, we suggest that such an effort should not be limited by solely exploiting living cells and enzyme evolution and describe some useful yet less intensive complementary approaches that may prove especially productive in this grand scheme. By decoupling reactions from the environment of a living cell, a significantly larger portion of potential synthetic chemical space becomes available for exploration; most of this area is currently unavailable to cell-based approaches due to toxicity issues. In contrast, in a cell-free reaction a variety of classical enzymatic approaches can be exploited to improve performance and explore and understand a given enzyme's substrate specificity and catalytic profile towards non-natural substrates. We expect these studies will reveal unique enzymatic capabilities that are not accessible in living cells.

Fig. 1. Multienzymatic pathway converting glucose into monoterpenes. Schematic of the synthetic enzyme system converting glucose to monoterpenes. Glycolytic and mevalonate enzymes are highlighted in blue and orange respectively. Auxiliary enzymes constituting the purge valve, (mGap and NoxE) and phosphate recycling by pyrophosphatase are highlighted in red, purple, and cyan. Different monoterpenes can be made by using alternative terpene synthases (green). Abbreviations: Hex – hexokinase, Pgi – glucose-6-phosphate isomerase, Pfk – phosphofructokinase, Fba – fructose-1,6-bisphosphate aldolase, Tpi – triose phosphate isomerase, Gap – gald-3-P dehydrogenase, mGap – gald-3-P dehydrogenase, Pgk – phosphoglycerate kinase, dPgm – 2,3-bisphosphoglycerate-dependent phosphoglycerate mutase, Eno – phosphoenolpyruvate hydratase, PykF – pyruvate kinase, PDH – pyruvate dehydrogenase, PhaA – acetyl-CoA acetyltransferase, Hmgs – β-hydroxy β-methylglutaryl (HMG)-CoA synthase, Hmgr – HMG-CoA reductase, Mvk – mevalonate kinase, Pmvk – phosphomevalonate kinase, Mdc – mevalonate-PP decarboxylase, Idi – isopentenyl-PP isomerase, Fpps – farnesyl-PP synthase, NoxE – NADH oxidase, PPase – pyrophosphatase, Gald-3-P – glyceraldehyde 3-phosphate, DHAP – dihydroxyacetone phosphate, P – phosphate, PEP – 2-phosphophenolpyruvate, HMG – 3-hydroxy-3-methylglutaryl, PP – diphosphate. Schematic drawn from ref. .

Fig. 2. Nanoparticle enhancement of enzyme activity. (A) Schematic of a CdSe/ZnS core/shell semiconductor quantum dot (QD) surface-functionalized with the DHLA-CL4 ligand to make it colloidally stable in buffer. Phosphotriesterase (PTE) is ratiometrically self-assembled to the QD surface by its terminal hexahistidine (His)₆ sequence. The average number of PTE per QD is controlled through the molar stoichiometry added during assembly. PTE hydrolysis of paraoxon substrate to p-nitrophenol product, which absorbs at 405 nm, is also shown schematically. Structure of the PTE competitive inhibitor triethyl phosphate. QD phosphotriesterase bioconjugate triethyl phosphate inhibition assays. Three-dimensional plots of PTE initial rates versus increasing paraoxon concentration in the presence of increasing triethyl phosphate inhibitor for (B) free enzyme and (C) 625 QD nm emitting (diameter 9.3 nm)–(PTE)₉ bioconjugates. Estimated K_i, k_cat, and K_M values shown for each. Figure reproduced with permission from ref. Copyright 2015 American Chemical Society.

Fig. 3. Coupled enzymatic and channeling activity when assembled on a QD scaffold. (A) Reaction scheme for coupled PykA and LDH activity. Individual and coupled enzyme activity monitored by NADH consumption with loss of its absorption at 340 nm. (B) Schematic of the coupled PykA–LDH enzyme system colocalized on a QD surface. The propensity of the enzymes to form cross-linked QD dimers and, to a lesser extent, trimers via the enzyme's tetrameric polyhistidine tags located at each monomers distal end is also schematically indicated. Note, not to scale. (C) LDH tetramer stability monitored as its concentration is reduced when free in solution and as assembled to QDs. Specific activity of LDH (μM NADH consumed s⁻¹ μM LDH⁻¹) determined at various enzyme concentrations in the absence (black) or presence of 605 nm emitting QD (blue, diameter ∼10.1 nm) or 525 nm emitting QDs (red, diameter 4.3 nm). Data fit to a dissociation equation. Note how LDH activity when assembled to the QDs significantly extends its activity profile below the 10 nM dissociation constant of the wildtype enzyme. (D) Coupled PykA–LDH enzymatic kinetics. NADH consumption in a combined PykA–LDH reaction monitored with enzymes assembled to 605 (red) or 525 QDs (green) and QD-free (black). The reactions were at a concentration of 8 LDH (2.4 nM)/4 PykA (1.2 nM)/1 QD (0.3 nM). Enzyme-only contained the equivalent amount of free enzyme. Figure reproduced with permission from ref. Copyright 2018 American Chemical Society.

Fig. 4. Schematic highlighting potential contributions from the studies suggested here towards augmenting retrosynthetic enzyme synthesis. The concepts behind each are outlined in the text along with how they can contribute to improving the data that will give rise to retrosynthetic enzymatic capabilities. E1 and E2 represent enzymes in an assembled pathway while the subscripts a–c or a–e represent homologs with different substrate and reactivity profiles.