Tuning aromatic cage occupancy in prenyltransferases enables selective and efficient production of rare c-prenylated flavonoids

Domain: chemistry
Published: 2026-02-19
Editorial outcome: minor_revision

Abstract
C-prenylated flavonoids possess notable pharmaceutical potential, but their production is hindered by the challenging selective prenylation of their unstable polyphenolic cores. Natural prenyltransferases offer a direct route but suffer from low activity and incomplete mechanistic understanding. Here, we report a directed evolution strategy to reshape the active pocket of the prenyltransferase AtaPT, uncovering an aromatic cage that governs both regioselectivity and donor specificity. By tuning cage occupancy, we engineer three mutants with high chemo- and regioselectivity toward dimethylallyl diphosphate or geranyl pyrophosphate. Structural analysis and molecular simulations validate the role of the cage in guiding flavonoid prenylation. Notably, the aromatic cage mechanism observed in AtaPT is not unique and can be recapitulated in homologous enzymes. Introduction of the aromatic cage consistently enhances both activity and selectivity, confirming its crucial role. AtaPT mutants enable the efficient and scalable synthesis of 27 C-prenylated flavonoids, including 8 previously unreported compounds. With an integrated donor regeneration system, preparative-scale biotransformations achieve product titers up to 400 mg/L. This study establishes a selective and scalable biocatalytic platform for flavonoid prenylation and offers mechanistic insights for enzyme engineering.

Introduction
Prenylated flavonoids represent a distinctive class of natural products, characterized by the attachment of lipophilic prenyl chains to various positions on flavonoid scaffolds (Fig. 1a). This structural diversification imparts a broad spectrum of biological activities, including anti-inflammatory1, anti-bacterial2, antioxidant3, antitumor4, antidiabetic5 and enzyme inhibitory effects6. Owing to these properties, prenylated flavonoids have garnered growing interest as promising scaffolds for drug discovery, attracting attention from both academia and the pharmaceutical industry. Notably, the position, number, and structural diversity of prenyl side substituents play a critical role in modulating bioactivity7. Moreover, the structural diversity of prenyl side chains extends beyond variations in carbon chain length; it also encompasses both normal prenylation (attack at the C-1 primary center of the prenyl carbocation) and reverse prenylation (attack at the C-3 tertiary center of the prenyl carbocation), further expanding the structural repertoire of prenylated flavonoids (Fig. 1b). Therefore, the development of precise and efficient prenylation methodologies to obtain pure products is of paramount importance, particularly in pharmacal research and related fields.

To date, over 1000 naturally occurring prenylated flavonoids have been identified, however, their typically low abundance in nature often limits further investigation of their biological functions and therapeutic potential8. The inherent structural complexity of these secondary metabolites further complicates isolation and purification efforts. To address these challenges and obtain sufficient quantities of diverse, pure prenylated flavonoids, traditional organic synthesis methods—such as Friedel-Crafts alkylation9, cross-coupling reactions10, and rearrangement reactions11, have been employed. Nevertheless, these approaches struggle to achieve precise selectivity and preserve the delicate flavonoid core under the harsh reaction conditions required. Recent advances in molecular biology, including gene sequencing and cost-effective synthesis, have significantly propelled the development of directed evolution technologies broadening the scope of biocatalysis12–17. Biocatalysis offers superior selectivity compared with traditional chemical synthesis, leading to a higher proportion of target products in the reaction mixture. This enhanced selectivity simplifies downstream purification and significantly reduces post-reaction processing costs, thereby improving the overall cost-effectiveness of the biocatalytic approach. Consequently, biocatalysis has emerged as a promising complement to chemical synthesis, offering high selectivity, mild reaction conditions, and environmental sustainability18–20.

Among biocatalysts capable of alkylation, aromatic prenyltransferases (aPTs) have demonstrated utility in producing prenylated flavonoids using dimethylallyl pyrophosphate (DMAPP), geranyl pyrophosphate (GPP), or farnesyl pyrophosphate (FPP) as prenyl donor21–24. Prior studies have established aPTs as valuable tools for constructing flavonoid prenylation platforms, owing to their satisfactory selectivity across diverse flavonoid substrate25–29. However, their practical application is limited by low to moderate catalytic activity and poor tolerance for substrate concentrations exceeding 0.5 mM. Although numerous aPTs have been characterized in metabolic engineering30–33 or evaluated for acceptor substrate scope34–38, few investigations have elucidated the mechanisms governing their activity and selectivity. Fewer than 20 aPT crystal structures have been reported to date, with characterized substrate specificity largely confined to amino acids39–41, dihydroxyl napthine36,42–44 or olivetolic acid19,45, and no studies have detailed structure-based substrate selectivity or regio- and chemoselectivity toward flavonoids. The sole mechanistic insight into regioselectivity engineering of an aPT was reported by Li and co-workers, who successfully switched the product regioselectivity of 5-dimethylallyl tryptophan synthase (5-DMATS) to 6-DMATS via structure-guided mutagenesis; however, this work focused on tryptophan, a structurally simpler substrate than flavonoids46. Thus, structure- and mechanism-guided engineering of aPTs for the selective prenylation of flavonoids remains largely underexplored47. The Dai group reported a prenyltransferase, AtaPT, from Aspergillus terreus, which exhibits remarkable substrate promiscuity toward a broad range of aromatic acceptors (up to 72 types) and prenyl donors (with prenyl residues up to 20 carbons). This enzyme provides a powerful biocatalytic tool for the synthesis of diverse prenylated compounds. Moreover, the structural elucidation of AtaPT has offered valuable insights for further engineering toward enzymes capable of constructing unique bioactive scaffolds48. Building upon this pioneering work, significant progress has been made in tailoring its selectivity toward flavonoids49 and chalcones50. However, given its exceptional catalytic potential, a deeper understanding of the mechanisms governing its activity and selectivity is essential for establishing a versatile platform for efficient and selective prenylation.

In this work, we employed directed evolution to the AtaPT, yielding three mutants with exceptional regioselectivity, chemoselectivity and enhanced activity for the prenylation of kaempferol at the 8-site, 3′-site and for the 3′-site geranylation, respectively. Leveraging structural analysis and molecular dynamics simulations, we identified an aromatic cage structural motif that plays a critical role in modulating regioselectivity and catalytic activity of AtaPT mutants toward flavonoid substrates. By further expanding the substrate scope and enabling the large-scale synthesis of diverse prenylated flavonoid derivatives—facilitated by an in situ DMAPP and GPP regeneration system—we establish a versatile and scalable biocatalytic platform for regioselective flavonoid prenylation.

Results
Testing the Activity of the AtaPT-WT toward Kempferol
8-Prenylkaempferol (8p-KAE) is a direct precursor to the drug Icaridin (Fig. 2a). To date, only seven reported aromatic prenyl-transferases (aPTs) are capable of accepting kaempferol (KAE) as a substrate to produce 8p-KAE (Supplementary Table 6)33,48,51–54, with the highest documented conversion reaching 65% at a low substrate concentration of 0.2 mM. To develop a more efficient biocatalytic route to 8p-KAE, broaden the spectrum of kaempferol prenylation products, and gain mechanistic insight into aPT-mediated regioselectivity, we selected kaempferol (1a) as a model substrate for directed evolution. AtaPT48–50, an aPT known for its promiscuous chemo- and regioselectivity, was chosen as the starting enzyme for directed evolution.

To facilitate high-throughput screening, the use of whole cells as biocatalysts was deemed essential. Accordingly, wild-type AtaPT (AtaPT-WT) was expressed in E. coli, and the resulting cells were employed directly as catalysts. Using dimethylallyl diphosphate (DMAPP) as the prenyl donor, AtaPT-WT catalyzed the formation of four major products from 1a, albeit with low total conversion (Fig. 2b). LC-MS analysis, supported by comparison with a commercial standard (Supplementary Fig. 1, 3 and 32), identified product 2a as 8-prenylkaempferol, with a conversion below 1% and regioselectivity of only 34%. Additional products included C-prenylated derivatives 3a and 4a and an O-prenylated derivative (5a). To improve activity, regio-, and chemoselectivity, a semi-rational directed evolution strategy was subsequently implemented.

Directed evolution of AtaPT
To identify mutation hotspots, 1a and DMAPP were sequentially docked into the crystal structure of wild-type AtaPT (PDB: 5KCG)48. Based on the resulting model (Supplementary Fig. 2), 22 residues within 5 Å of the active site G82, L83, E91, K95, R104, I167, E169, S170, C175, S177, D179, K188, L252, K266, L323, P324, G326, N328, Q342, R396, W397, and N411 were selected for site saturation mutagenesis (SSM)55. A total of 2,024 variants were expressed in E. coli and screened using whole-cell biotransformation in 96-deep-well plates, followed by product quantification via high-performance liquid chromatography (HPLC). Positive mutants were identified in 12 libraries (K95, I167, E169, S170, C175, S177, L323, P324, G326, N328, Q342, and W397). Among them, P324H exhibited the highest selectivity for 2a, achieving 65.5% selectivity at a conversion of 0.5%. Examination of the P324 position in Supplementary Fig. 2 reveals that it is located near the B-ring of substrate 1a. The substitution of the rigid proline residue with the positively charged aromatic residue histidine likely introduced an aromatic cage that facilitates accommodation of the B-ring of 1a, contributing to the observed enhancement in regioselectivity when DMAPP serves as the donor. G326W showed markedly improved activity and selectivity for 3a, reaching 16.6% conversion with 95.8% selectivity. Additionally, S177G preferentially produced 5a with a conversion of 12.39% and selectivity of 46.2% (Supplementary Fig. 3 and Supplementary Table 8). Notably, both S177 and G326 had previously been implicated in modulating DMAPP binding and activity48.

Building on the results from the initial cycle of directed evolution, further optimization was undertaken to enhance activity and selectivity. Iterative saturation mutagenesis (ISM)56–59 was applied to nine promising residues―K95, I167, E169, S170, C175, L323, N328, Q342, and W397―using P324H, G326W, and S177G as templates. In parallel, four double-mutation libraries were constructed: P324H/G326NNK, P324H/S177NNK, G326W/P324NNK, and G326W/S177NNK. From this effort, the double mutant S177G/P324H was identified, exhibiting a conversion of 1.1% and a selectivity of 55.0% for 2a. Iterative combinations led to the triple mutant S177G/P324H/N328M, which achieved a 5.5% conversion with 56.0% selectivity toward 2a. In contrast, iterative mutagenesis based on G326W and S177G yielded less favorable outcomes, with no mutants surpassing the performance of their respective templates.

Given the limited improvement in yield from above efforts, we employed focused rational iterative site-specific mutagenesis (FRISM)13,60–64, integrating insights from SSM and ISM to accelerate the discovery of superior variants. Subsequently, two combinatorial mutations sets were designed based on SSM and ISM results: K95/I167/E169/S170/C175 and E169/S170/S177/P324/N328 (Supplementary Table 7). This strategy generated the quintuple mutant E169A/S170Q/S177A/P324H/N328M (named as M5), which exhibited significantly enhanced activity (14.6% conversion) and selectivity (73.1%) toward 2a. Using M5 as a template, further combinatorial mutagenesis at positions I167, C175, L323 and N328 produced the octuple mutant I167A/E169A/S170Q/C175M/S177A/L323Q/P324H/N328R (named as M8), demonstrating the highest activity and regioselectivity for 2a (44.5% conversion and 77.4% selectivity, Fig. 2d). In contrast, analogous efforts aimed at improving 3a and 5a formation did not yield superior variants.

Given the broad donor promiscuity of AtaPT, we next assessed its activity with GPP using 1a as the acceptor. While the WT enzyme exhibited moderate selectivity for the 3’-geranylated product (6a) at 84.9%, overall conversion remained low (<14.6%). To enhance both catalytic efficiency and selectivity, selected variants from previous mutagenesis rounds were screened. The septuple mutant I167T/E169A/S170Q/S177A/L323Q/P324H/N328M (designated M7) demonstrated a dramatic improvement, achieving >90.7% conversion and 92.6% selectivity for 6a (Fig. 2e). Notably, M7 was identified serendipitously during the second round of combinatorial mutagenesis, highlighting the effectiveness of the iterative strategy.

With the successful evolution of four functional mutants—G326W, M7, M8, and S177G—catalyzing the formation of three distinct C-prenylated KAE derivatives and one O-prenylated product, precise structural characterization was undertaken. This step was essential to confirm the exact prenylation sites and substitution patterns, thereby elucidating the chemo- and regioselectivity profiles of each engineered variant.

Structural Characterization of 2a, 3a, 4a, 5a, and 6a
To characterize the structures of 2a, 3a, 4a, 5a, and 6a, biotransformations were scaled up using whole-cell E. coli catalysts expressing the corresponding AtaPT mutants. Reactions were conducted at a 2 mM concentration of 1a with 4 mM DMAPP (or GPP for 6a) in 50 mL volumes. The isolated yields were 33.6% for 2a, 12.2% for 3a, 6.0% for 4a, 9.1% for 5a, and 78.4% for 6a. The products were purified and structurally validated by NMR and high-resolution mass spectroscopy (HRMS). Spectroscopic analysis revealed that 3a and 6a are C-prenylated and C-geranylated, respectively, at the 3′-position of 1a, 4a is C-prenylated at the 6-position of 1a, while 5a was identified as a 4′-O-prenylated derivative. Interestingly, 2a was determined to be a rare reverse 8-prenylated kaempferol, differing from the anticipated 8p-KAE despite co-elution in HPLC (Supplementary Figs. 3, 55 and 56). This highlights the importance of rigorous structural elucidation in biocatalytic prenylation studies.

Kinetic Characterization of AtaPT Variants
To elucidate the catalytic basis underlying the enhanced performance of variants G326W, M8, and M7, steady-state kinetic assays were conducted using 1a as the acceptor substrate. The wild-type AtaPT (WT) exhibited insufficient activity with dimethylallyl pyrophosphate (DMAPP) to reliably determine kinetic parameters. In contrast, M8 and G326W showed markedly improved catalytic efficiency, with kcat/Km value of 34.79 s⁻¹ M⁻¹ and 15.09 s⁻¹ M⁻¹, respectively, using DMAPP as the donor. When geranyl pyrophosphate (GPP) was employed, M7 demonstrated both an increased turnover number (kcat = 0.64 min-1) and a reduced Michaelis constant (Km = 0.23 mM) relative to WT (kcat = 0.09 min-1; Km = 1.27 mM). As a result, the catalytic efficiency of M7 (kcat/Km = 47.12 s⁻¹ M⁻¹) exceeded that of WT by over 40-fold (1.12 s⁻¹ M⁻¹), indicating substantially improved substrate affinity and catalytic performance (Supplementary Table 9 and Supplementary Fig. 13). We investigated the stability of the purified enzymes M8, G326W, and M7 over extended reaction times. The reaction profiles showed that product formation plateaued after approximately 8h. When additional DMAPP/GPP was supplemented at this point, a further increase in product yield was observed, indicating that enzyme activity was maintained beyond the initial reaction phase. To further confirm enzyme stability, the reaction supernatant was collected after 8h, supplemented with a fresh substrate and DMAPP or GPP, and incubated further. The continued formation of the corresponding products demonstrated that the enzymes retained substantial catalytic activity over prolonged periods (Supplementary Fig. 14). These results suggest that the current limitation lies in donor substrate depletion rather than enzyme instability, highlighting the need for a more efficient donor regeneration system in future applications.

Crystal structure resolution, molecular docking and molecular dynamics simulations
To elucidate the molecular basis for the enhanced activity and regio-/chemoselectivity of engineered AtaPT variants, we determined the crystal structures of M8 (2.50 Å, PDB: 9V5G), G326W (2.69 Å, PDB: 9V5F) and M7 (2.20 Å, PDB: 9V3U) (Supplementary Table 10). Crystals of M8 were obtained in the apo form and subsequently soaked with DMSPP and 1a for varying durations prior to data collection. The overall protein architecture remained largely unchanged relative to the wild-type AtaPT structure, except for a notable movement of DMSPP in M8 away from the substrate binding pocket (Fig. 3a). This rearrangement is attributable to the N328R mutation, whose guanidinium group forms dual hydrogen bonds with the two phosphate moieties, thereby stabilizing the altered geometry. The substrate-binding pocket remained solvent-accessible, with 1a positioned at the base of the cavity. On one side of 1a, the chromenone core was packed against P324H, Y344, W397 and Y413, forming an aromatic cage. Of these residues, the P324H mutation played a pivotal role in anchoring 1a via a hydrogen bond, thereby reducing its conformational flexibility (Fig. 3b). This is consistent with the substantial increase in regioselectivity observed for 2a in this variant (Fig. 2d). To further assess whether this mutation is irreplaceable, we substituted this position with other aromatic residues (F, W, and Y) as well as bulky residues (M and L) in the WT, M7, and M8. Enzymatic assays showed that both activity and selectivity were markedly reduced compared with the original variants (Supplementary Fig. 16), demonstrating that histidine at this position is uniquely optimal for the observed enhancement in catalytic performance. On the other side, the chromenone core and the phenolic group of 1a were stabilized by C175M and L252, respectively. Furthermore, the side chains of E91 and R396 form additional hydrogen bonds with the chromenone core (Supplementary Fig. 17). These interactions together bring the C8 position of 1a into close proximity with the C3 atom of DMSPP (5.2 Å). The hydrogen bond between E91 and the hydroxyl group on C7 may facilitate the attack of C8 on C3 of DMAPP, thereby promoting the formation of the unusual reverse product 2a via M8.

In contrast to M8, no substrate was observed in the structure of M7, even though the crystals were soaked with 1a and GSPP before harvest. The electron density for GSPP was continuous and intact in one protomer of the dimeric structure (Supplementary Fig. 17). While in the other protomer, we could only define the pyrophosphate part unambiguously, implying flexibility of the geranyl chain. To probe the molecular basis of M7’s improved catalytic performance with geranyl pyrophosphate (GPP), we docked both 1a and GPP into M7 and performed molecular dynamics (MD) simulations (Supplementary Fig. 18). The simulations demonstrated that 1a can be stably accommodated within the active site, as supported by the narrow RMSD fluctuations of 1a and GPP during the whole simulations (Supplementary Fig. 18a). The distance between the 3′-positions of 1a and the electrophilic center of the geranyl donor keep steadily around ~5 Å (Supplementary Fig. 18b and 18c). The main anchoring residues for 1a binding through hydrogen bond include E91, N172, S192 and H324 (Supplementary Fig. 18d and 18e). Notably, the extended geranyl chain of GPP projects into the aromatic cage (Fig. 3c). This spatial shift induces a distinct binding orientation, in which the phenolic group of 1a becomes deeply buried at the base of the pocket, forming a stabilizing hydrogen bond with the carboxylate of. This interaction likely activates the phenol ring for nucleophilic attack at the ortho position, thereby favoring C3′-geranylation to yield 6a. The chromenone ring adopts a partially solvent-exposed configuration, directed toward a surface-accessible channel formed by a flexible loop comprising residues 167–170 (Fig. 3d). Within this loop, the S170Q mutation introduces a glutamine side chain that intrudes into the channel, narrowing its aperture and potentially stabilizing the substrate by limiting its mobility. Additionally, the I167T and E169A mutations probably enhance loop flexibility, promoting favorable conformational adjustments for substrate engagement and retention. Collectively, these observations suggest that the 167–170 loop functions as a dynamic gating element that modulates substrate access and positioning, thereby contributing to M7’s improved regioselectivity and activity in GPP-dependent C3′-geranylation of 1a.

Structural and computational analyses collectively reveal two distinct binding conformations of 1a within the AtaPT substrate-binding pocket, which correspond to prenylation at either the chromenone ring (conformation I) or the phenolic moiety (conformation II). Conformation I is stabilized by the combined effects of the C175M mutation and the aromatic cage introduced by P324H mutation. The aromatic cage appears essential for establishing this conformation, as its occupation or disruption triggers a switch to conformation II, favoring prenylation on the phenolic moiety. This mechanistic insight explains the regioselectivity observed for the M7 and G326W mutants. Specifically, in M7, the aromatic cage is occupied by the extended geranyl chain of GPP, whereas in G326W, the cage is disrupted due to the absence of the P324H mutation (Supplementary Fig. 19a). Furthermore, the bulky indole ring introduced by the G326W mutation occupies the cage’s center, accounting for this mutant’s exclusive preference for DMAPP as the prenyl donor, consistent with experimental observations showing no activity with GPP.

The role of C175M is also indispensable for M8, as it may reduce the dynamics of 1a and orient the chromone ring for prenylation. This may be the reason why M7 cannot catalyze the prenylation of 1a effectively as M8 (Fig. 2d and Supplementary Fig. 5), for the absence of C175M in M7 may result in a high degree of flexibility of 1a in the pocket. When overlapping the M7-1a model with the structure of M8, the closest distance between the phenolic ring of 1a and the C175M side chain is only 3 Å (Supplementary Fig. 19b), suggesting a potential steric clash. This observation explains why M8 exhibits very weak activity in modifying the 3′-position of 1a using GPP as the donor, even lower than the WT.

Mechanism-guided design of PTs beyond AtaPT
The mechanisms of both the aromatic cage and the flexible loop have been elucidated; more importantly, it remains essential to determine whether the aromatic cage can be transferred to other PTs to guide the rational mining and design of enzymes with enhanced catalytic efficiency. Using AtaPT as a template for BLAST searches, we selected three homologous enzymes (Supplementary Table 11)—AfaPT (94.81% identity), AaaPT (86.70%), and AmaPT (56.87%)—all of which contain the conserved XXLPXX motif and a highly similar loop region (residues 167–170). We initially tested the WT activities of these enzymes toward kaempferol. With DMAPP, AfaPT-WT exhibited weak activity (1.7%) and produced 2a as the major product (42.0%), whereas AaaPT-WT also showed low activity (1.3%) but favored 3a (37.7%), with only 20.6% 2a. AmaPT-WT showed similarly limited activity (1.4%) and generated only 3a (56.1%) with no detectable 2a. With GPP, AfaPT-WT and AaaPT-WT afforded 6a with low activity (7.5% and 8.9%) but moderate selectivity (82.4% and 80.5%), whereas AmaPT-WT remained largely inactive (<1%) and produced only 7.5% 6a (Fig. 4a).

According to our previous work on rationally tuning the chemoselectivity of glycosyltransferases, the substrate-binding mode governs both chemo- and regioselectivity18. To enhance catalytic performance and target-product formation, we first performed sequence and structural homologous comparisons using AtaPT-M8 and AtaPT-M7 as templates. For AfaPT and AaaPT, the active pockets were highly similar to that of AtaPT (Fig. 4b), suggesting that comparable catalytic properties could be achieved by introducing into these enzymes the key sites identified in AtaPT-M7 and AtaPT-M8 via site-directed mutagenesis. Guided by this strategy, we constructed AfaPT-M7, AfaPT-M8, AaaPT-M7, and AaaPT-M8. As anticipated, when tested with DMAPP, the M8 variants displayed dramatic increases in activity: AfaPT-M8 and AaaPT-M8 showed 14-fold and 19-fold increases in overall activity (22.9% and 25.6%) and 24-fold and 66-fold improvements in 2a formation (16.9% and 18.3%), accompanied by substantially enhanced selectivity (73.8% and 71.6%; Fig. 4c). When tested with GPP, the M7 variants similarly improved catalysis: AfaPT-M7 and AaaPT-M7 increased overall activity to 75.6% and 66.0% and enhanced 6a formation by 11-fold and 9-fold (70.7% and 63.2%), maintaining excellent selectivity (93.6% and 95.8%; Fig. 4c).

In contrast, the active pocket of AmaPT exhibited low homology to AtaPT and contained three bulky residues (Y193, M253, M327) that likely hindered productive binding of 1a in a conformation favorable for generating 2a (Fig. 4d). Therefore, improving the catalytic activity and selectivity of AmaPT required not only introduction of M7/M8 mutations but also remodeling of the pocket through substitution of these obstructive residues with the corresponding smaller residues found in AtaPT. To test this hypothesis, we constructed five variants: AmaPT-SLG (Y193S/M253L/M327G), AmaPT-M7, AmaPT-M8, AmaPT-M7-SLG, and AmaPT-M8-SLG. With DMAPP, AmaPT-M8 showed only modest overall activity improvement (2.5%) but enabled formation of 2a for the first time (9.2%). As expected, AmaPT-SLG exhibited slightly reduced overall activity relative to AmaPT-WT yet produced 2a with 8.7% selectivity. Notably, AmaPT-M8-SLG increased total activity to 7.9% and generated 2a at levels six fold higher than AmaPT-SLG and twofold higher than AmaPT-M8 (Fig. 4e).

When tested with GPP, AmaPT-M7 exhibited limited improvement in total conversion but increased 6a formation nine fold with markedly enhanced selectivity (69.2%). Consistent with our design, AmaPT-SLG and AmaPT-M7-SLG produced greatly enhanced conversions (21.0% and 54.8%) despite moderate selectivity toward 6a (26.0% and 55.4%; Fig. 4f). Ultimately, the optimized mutant AmaPT-M7-SLG increased 6a formation by approximately 862-fold and improved selectivity nearly eightfold relative to AmaPT-WT, demonstrating that the aromatic cage mechanism discovered in AtaPT is transferable and can be effectively reconstituted in homologous PTs.

Substrate Scope Exploration
To evaluate the utility of AtaPT and its engineered C-prenyltransferase mutants for flavonoid prenylation, in vitro assays were conducted with 26 structurally diverse, medicinally relevant flavonoids (Supplementary Table 12). Consistent with prior reports48, wild-type AtaPT (WT) catalyzed the prenylation of 23 out of 26 tested substrates, underscoring its broad substrate scope. However, under high substrate loading (2 mM), whole-cell catalysis by WT generally yielded low conversion and poor selectivity, with the exception of 1g and 1j, which were efficiently converted to single prenylated products (>50% yield). Most other substrates yielded complex product mixtures in low abundance. Notably, neither WT nor the tested mutants (G326W, M5, M7, and M8) exhibited activity toward substrates 1t, 1u, and 1v. Comprehensive analysis of the reactions involving substrates 1t–1z reveals that both the position and number of hydroxyl groups exert a pronounced influence on catalytic activity. Substrates 1t (lacking hydroxyl groups), 1u (bearing one hydroxyl group on ring C), and 1v (bearing one hydroxyl group on ring A, possibly affected by an ortho effect) exhibited no detectable activity. Substrates 1w and 1x, each containing a single hydroxyl group on ring A, showed only weak activity. In contrast, substrates 1y and 1z, each possessing a hydroxyl group on ring B, displayed relatively higher activity with mutants G326W and M7. These results indicate that the hydroxylation pattern critically correlates with enzymatic performance, and substrates with hydroxyl substitution on the B ring generally exhibit broader enzyme compatibility and enhanced activity (Supplementary Fig. 23).

In contrast, M8 displayed markedly improved activity and regioselectivity across seven substrates (1a–1c, 1f, 1h–1i), as confirmed by product isolation and NMR spectroscopic analysis. For 1a, M8 catalyzed formation of the rare reverse 8-prenylkaempferol (2a). Substrates 1b, 1c and 1f yielded 2’-prenylated products (2b, 2c and 2f), attributed to the influence of multiple hydroxyl groups on the B ring guiding substrate orientation. For 1h, bearing a methylated 4′-hydroxyl, prenylation occurred at the 6-position rather than the canonical 8-position. Isoflavones 1i and 1j, characterized by an inverted B-ring topology, were converted to 6-prenylated derivatives. M8 also exhibited moderate reactivity toward additional substrates, including 1g, for which it generated a broader range of products, likely reflecting enhanced conformational adaptability in the active site. In contrast, low activity toward 1e, 1k, and 1s was attributed to the limited number of hydroxyl groups, weakening enzyme–substrate interactions (Fig. 5a and Supplementary Fig. 24).

The G326W mutant also prenylated eight substrates (1a–1b, 1d, 1e, 1g–1i, 1k) with enhanced activity and regioselectivity compared to WT. Products from 1a, 1b, 1e, and 1g were predominantly 3’-prenylated. The two products of 1d are respectively located at the 2’ and 4’-O-positions. Notably, 1h yielded a 2′-substituted product, attributed to the methylation of the 4′-hydroxyl. Isoflavones 1i and 1k were converted into diprenylated derivatives with modifications at both the 3′- and 8-positions, suggesting that G326W reorients the B ring to facilitate proximity to DMAPP. Substrates 1c and 1f showed limited reactivity, likely due to steric hindrance from B-ring hydroxylation, while 1l–1o and 1q–1r exhibited minimal conversion, presumably due to the absence or masking of key hydroxyl groups required for catalysis (Fig. 5b).

M5 and M7 exhibited significantly enhanced activity with GPP as the prenyl donor. In particular, M7 catalyzed geranylation of 11 substrates with high efficiency and selectivity, generating more than 15 distinct products. For example, 1c, bearing three hydroxyls on the B ring, yielded two O-geranylated products—3c (4′-O) and 4c (3′-O)—in a 3:1 ratio, although they could not be separated by conventional chromatography. Substrates 1h and 1j gave rise to 4h (2′-substituted) and 3j (6-substituted), respectively, consistent with altered binding geometries in the active site. Product from 1k was not fully characterized due to co-elution, as it exists as a 1:1 mixture, the products are likely geranylation at either the 2’- or 3’-position. The remaining substrates yielded products predominantly prenylated at the 3’-position likely reflects the larger volume and flexible conformation of the active site, which accommodates GPP’s extended isoprenoid chain (Fig. 5c).

Altogether, we successfully prepared and characterized 31 prenylated flavonoids. Notably, eight of these compounds (2f, 3c, 3f, 3h, 3j, 4c, 4f, and 4h) are previously unreported, thereby expanding the chemical diversity of this compound class and providing more molecular scaffolds for future studies.

Coupling the isopentenol utilization pathway with prenyltransferase for prenylflavonoid biosynthesis
Recent strategies to boost intracellular dimethylallyl diphosphate (DMAPP) levels have focused on optimizing the native mevalonate (MVA) and methylerythritol phosphate (MEP) pathways65, as well as engineering synthetic isopentenol utilization pathways (IUPs)66. By fine-tuning precursor availability in concert with prenyltransferase (PT) activity, these approaches facilitate efficient and cost-effective prenylation, holding significant promise for industrial biocatalysis67,68. The IUP offers a streamlined alternative by bypassing complex central carbon metabolism, instead leveraging exogenously supplied isoprenol or prenol, which are converted to isoprenoid diphosphates via sequential phosphorylation. This decoupled system not only simplifies metabolic control but also reduces the metabolic burden on host cells, presenting a competitive advantage over traditional biosynthetic routes. Integration of the IUP with PTs such as ShFPT69, NgFPT70, AnaPT71 and Ad0372 have recently enabled the biosynthesis of diverse prenylated flavonoids. Building on this framework, we envision that coupling a multi-enzyme cascade for enhanced in vivo DMAPP production with engineered PT variants—such as the highly active and selective M8 and G326W—can further increase production titers and reduce costs. Additionally, the biosynthesis of geranyl pyrophosphate (GPP) in vivo, when coupled with the M7 variant, provides a promising route to access geranylated flavonoid scaffolds. Together, these strategies establish a versatile and scalable platform for the sustainable biosynthesis of prenylated natural products with pharmaceutical and nutraceutical relevance.

To evaluate the practical applicability of the biocatalytic cascade system for preparative-scale prenylation, we initiated reactions using whole cells co-expressing the prenyltransferase variant M8 (pET28a-M8) and the donor biosynthesis module PCDF-AtIPK-EcPK73. However, even in small-scale reactions (300 μL), the conversion rate is also limited, at only 5.6%. Cell lysis significantly improved reaction performance, with the lysate exhibiting double (11.3%) the activity of whole cells, though still inferior to that observed with externally supplied DMAPP. To improve donor supply, alternative kinase systems were tested. MjIPK74 and MTHIPK75 were expressed in place of AtIPK, generating the constructs PCDF-MjIPK-EcPK (23.1%) and PCDF-MTHIPK-EcPK (15.3%). Among these, MjIPK showed superior activity, doubling the conversion rate compared to the AtIPK-based system. Consequently, PCDF-MjIPK-EcPK and pET28a-M8 were selected for reaction optimization. Key parameters—including expression temperature, inducer concentration, catalyst ratio, reaction temperature and pH, and concentrations of enzyme, prenol, Mg²⁺, and DMSO—were systematically evaluated. Optimal conditions were determined to be: PCDF-MjIPK-EcPK: pET28a-M8 = 1:1, OD600 = 120, in 50 mM Tris-HCl buffer (pH 7.5), with 2 mM acceptor, 10 mM prenol, 2% DMSO (v/v), and incubation at 35°C. Under these conditions, the reaction plateaued after 10h, indicating efficient in situ DMAPP generation (Supplementary Figs. 25–27).

Following the success of DMAPP production, we extended this strategy to GPP biosynthesis from geraniol. Folk, a farnesol kinase from Arabidopsis thaliana, can also accept geraniol as a substrate76, whereas THAIPK, an isopentenyl phosphate kinase from Thermoplasma acidophilum, naturally acts on DMAPP; however, engineered variant of THAIPKG (THAIPK-Y70A/V130A/I140A) is capable of accommodating geranyl phosphate (GP)77. Initially, the donor module PCDF-Folk-THAIPKG was constructed; however, poor expression and solubility of Folk in E. coli, even after N-terminal truncation (residues 1–65), hindered activity73. In contrast, we discovered by chance an assembled system comprising PCDF-THAIPKG and pET28a-M7 successfully catalyzed the formation of compound 6a, albeit with a conversion of only 15.8%, which is significantly lower than the catalytic efficiency observed with the direct addition of GPP. Nevertheless, this result demonstrates the feasibility of an in situ GPP generation strategy from geraniol coupled with PT catalysis, enabling the production of geranylated products in whole-cell systems. With further optimization, such as enhancing GPP turnover efficiency, this platform could be extended to farnesol for in situ FPP generation and the subsequent biosynthesis of farnesylated products. Furthermore, application to substrates 1b and 1g yielded corresponding geranylated products, confirming the effectiveness of this GPP generation system. Notably, replacing geraniol with its isomer nerol afforded 5b and 4g with high efficiency, underscoring the system’s substrate flexibility (Supplementary Fig. 31).

The versatility of the cascade platform was further validated across a range of flavonoid substrates. Considering both reaction activity and the synthetic value of the products-2a with anti-breast cancer activity78, 2j with anti-diabetic and neuroprotective effects5,79, and analogs of 2f with antibacterial and cytotoxic activities80,81, we selected substrates 1a, 1f, and 1j for scale-up preparation. Preparative-scale reactions were performed for compounds 2a, 2f, and 2j in 1L systems, yielding 218 mg/L, 221 mg/L, and 442 mg/L of isolated products, respectively. An additional 11 products were synthesized in 50 mL reactions, each yielding between ~40–300 mg/L under optimized conditions (Fig. 6). These results highlight the broad applicability and scalability of the engineered prenylation platform, offering a robust and flexible strategy for the biosynthesis of structurally diverse prenylated flavonoids.

Methods
Reagents and instruments
All chemical reagents were purchased from Aladdin, Macklin Biochemical, Meryer (Shanghai) Chemical Technology Co., 9ding Chemistry, Bide PharmTech, Shanghai Yuan Ye Biotechnology Co., or Sangon Biotech (Shanghai, China) unless otherwise stated. The KOD OneTM PCR master Mix polymerase was obtained from Toyobo Ideas & Chemistry. DpnI was purchased from Abclonal Technology. The ClonExpress MultiS One-Step Cloning Kit was obtained from Vazyme Biotech. Primer synthesis and DNA sequencing were conducted at Sangon Biotech Company (Shanghai, China). Microspectrophotometer (Allsheng, Nano-300), ultrasonic homogenizer with noise isolating chamber (SCIENTZ-IID), freeze drier (SCIENTZ-10ND), centrifuge (Thermo Fisher, (Sorvall ST16R) and PICO17), PCR Amplifier (BIO-RAD, T100TM Thermal Cycler), mini shaker (Allsheng, MSC-100), shaker (Minquan, MQD-BIR). Shimadzu Nexis LC 20AT equipped with an electrospray ionization (ESI) source was used for the measurement of substrates and products. High resolution mass spectrometry (HRMS) detection was performed by X500R QTOF from ABSCIEX. The prenylated products were isolated and purified by semi-preparative HPLC on an LC-8A instrument (Shimadzu). Products were characterized by 1H NMR and 13C NMR in DMSO-d6 500 MHz Bruker spectrometer. Chemical shifts (δ) were given in ppm and coupling constants (J) were given in hertz (Hz).

Heterogenous expression and purification of PTs
AtaPT DNA sequence were synthesized by GENEWIZ (China), and then inserted into the NdeI-XhoI site of pET28a. The recombinant plasmid pET28a-AtaPT were transformed into E. coli BL21(DE3) for heterologous expression. E. coli cells were cultured (5 mL) overnight in Luria-Bertani (LB) medium containing 50 μg/mL kanamycin at 37°C with shaking (220 rpm). Subsequently, 1% seeding cultures were transferred into 500 mL TB-medium containing 50 μg/mL of kanamycin and grown at 37°C and 220 rpm in 2 L shake flasks. After the OD600 reached 0.8–1.0, the expression of recombinant AtaPT were induced with 0.2 mM IPTG at 18°C, 220 rpm for 16 h. The cells were harvested by centrifugation at 6,438g for 3 min at 4°C and then resuspended in binding buffer (50 mM Tris-HCl, 5 mM CaCl2, pH 7.5)49,50. After cell rupture by sonication in ice bath, the cell debris was removed by centrifugation at 15,480g and 4°C for 1 h. The soluble fraction was passed through a 0.45 μm syringe filter. The supernatant was applied to an ÄKTA avant 25 system equipped with a 5-mL HisTrapTM HP column following the manufacturer’s instructions, using a linear gradient of 10-500 mM imidazole in 50 mM Tris-HCl, 5 mM CaCl2, pH 7.5 (Supplementary Fig. 12).

Construction and screening of AtaPT mutant libraries
All strains, plasmids, and primers used in this study are listed in Supplementary Tables 2 and 3. PCR assays (Supplementary Table 4) were conducted with KOD OneTM PCR master Mix. Following sequence verification, the recombinant plasmids were transformed into E. coli BL21(DE3) for heterologous protein expression, as described above. Single-site saturation mutagenesis (SSM) and iterative saturation mutagenesis (ISM) was performed using primers containing NNK codons at targeted sites. To ensure adequate library coverage, 92 colonies were screened per mutagenized site. Colonies were cultured in 96-deep-well plates containing 0.3 mL LB medium supplemented with 50 μg/mL kanamycin at 37°C overnight. Subsequently, 0.7 mL TB medium containing 50 μg/mL kanamycin and 0.2 mM IPTG was added, and cultures were incubated at 18°C for 16 h. Cells were harvested and resuspended in 0.2 mL reaction buffer (2 mM substrate 1a, 2 mM DMAPP, 50 mM Tris-HCl, 5 mM CaCl2, pH 7.5) and incubated at 37°C for 24 h with shaking. Reactions were quenched by the addition of 0.6 mL methanol. Relative activity was determined by whole-cell biotransformation in a total volume of 2 mL reaction buffer (2 mM 1a, 2 mM DMAPP, 50 mM Tris-HCl, 5 mM CaCl2, pH 7.5) at 37°C for 24 h, using the wild-type enzyme as a control. Samples were centrifuged at 12,000g for 15 min and analyzed by HPLC/UV using a SHIMADZU Shim-pack GIS C18 column (4.6 × 250 mm, 5 μm) at a flow rate of 1 mL/min and column temperature of 30°C. The gradient elution programs are detailed in Supplementary Table 5.

Determination of kinetic parameters
Kinetic analyses were conducted for AtaPT-M8 and AtaPT-G326W using DMAPP, and for AtaPT-WT and AtaPT-M7 using GPP, with 1a as the substrate. Enzymatic assays were performed in 100 μL reaction volumes containing 50 mM Tris-HCl buffer (pH 7.5), 5 mM CaCl2, 2/5/10 μM enzyme, 2 or 4 mM saturated DMAPP/GPP, and varying concentrations of 1a (40, 60, 80, 100, 150, 200, 250, 300, 500, 1000, 1500, 2000, 2500, and 4000 μM). Reactions were incubated at 37°C for 10 or 20 min, then quenched by the addition of 300 μL methanol and centrifuged at 13,800g for 15 min. Supernatants were analyzed by HPLC as previously described.
All experiments were performed in triplicate. Michaelis–Menten constants (Km) were determined by non-linear regression fitting, and turnover numbers (kcat) were calculated accordingly (Supplementary Table 9 and Supplementary Fig. 13).

Structure determination of AtaPT mutants
The AtaPT variants (M8, G326W, S177G, and M7(11–424)), each bearing an N-terminal 6×His tag, were overexpressed in E. coli BL21(DE3) cells. Protein expression was induced with 0.4 mM IPTG when cultures reached an OD600 of 0.8, followed by incubation at 16°C for 12 h. Cells were harvested and lysed in lysis buffer containing 25 mM Tris-HCl (pH 8.0), 1 M NaCl, 25 mM imidazole, 0.5 mM β-mercaptoethanol (β-ME), and 1 mM phenylmethylsulfonyl fluoride (PMSF). The His-tagged fusion proteins were purified using Ni2+-affinity chromatography. The column was washed with buffer containing 25 mM Tris-HCl (pH 8.0), 25 mM imidazole, and 1 M NaCl, and proteins were eluted with 250 mM imidazole in the same buffer. Further purification was performed by size-exclusion chromatography using a Superdex 75 column (GE Healthcare) pre-equilibrated with buffer containing 25 mM Bis-Tris (pH 6.5), 200 mM NaCl, 5% (v/v) glycerol, and 3 mM dithiothreitol (DTT). The purified proteins were concentrated to 25 mg/mL and stored at –80°C. The small molecule 1a was dissolved in DMSO to a final concentration of 100 mM. Crystallization was performed using the hanging-drop vapor diffusion method at 20°C. Equal volumes (1 μL each) of protein-ligand complex solution and reservoir buffer were mixed and equilibrated against 500 μL of reservoir solution. Crystals were obtained in a buffer containing 0.1 M Bis-Tris (pH 6.5), 0.2 M ammonium sulfate, and 20% (w/v) PEG 3350, typically reaching full size within ~5 days. The crystals were then soaked with 1a (20 mM) and analogues of prenyl donors (DMSPP or GSPP, 10 mM) for 5 min to 12 h. Prior to flash-freezing in liquid nitrogen, crystals were cryo-protected using the reservoir buffer supplemented with 20% (v/v) glycerol. X-ray diffraction data were collected at beamline BL19U1 of the Shanghai Synchrotron Radiation Facility and BL02U1 at Shanghai Synchrotron Radiation Facility. Data were indexed, integrated, and scaled using the HKL3000 software package82. The structures were solved by molecular replacement using PHASER83 with the structure of AtaPT-WT (PDB: 5KCG) as the searching model. Following iterative manual rebuilding and refinement were performed with the use of COOT84 and PHENIX85, respectively.

Reaction condition optimization
All optimization experiments were performed in triplicate on an analytical scale (300 μL) using whole-cell catalysts, with DMAPP as the prenyl donor and 1a as the acceptor. Reaction conditions—37°C, 50 mM Tris-HCl (pH 7.5), and 5 mM CaCl2—were based on previously reported protocols. specifically at 37°C, 50 mM Tris-HCl (pH 7.5), and 5 mM CaCl2. To evaluate the effect of the co-solvent DMSO, reactions were carried out under optimized conditions with varying DMSO concentrations (2%, 4%, 6%, 8%, and 10% v/v) (Supplementary Fig. 10). The influence of cell density on product formation (2a, 3a, and 6a) was assessed by conducting reactions at different OD600 values (40, 60, 80, 100, and 120) in 50 mM Tris-HCl buffer (pH 7.5). Product titers increased with cell density up to an OD600 of 80, after which a decline was observed at OD600 = 100. To investigate the effect of reaction time, biotransformations were carried out at 37°C in 50 mM Tris-HCl buffer (pH 7.5) containing 6% (v/v) DMSO. Reactions were sampled at various time points (2, 4, 6, 8, 10, 12, and 24 h). Reactions were quenched by the addition of 0.9 mL methanol, and supernatants were analyzed by analytical HPLC as described previously (Supplementary Table 5 and Supplementary Fig. 10).

Preparation of E. coli whole cell catalyst harboring AtaPT mutants
After protein expression, the cells were harvested by centrifugation at 6438g for 2 min at 4°C, then resuspended in binding buffer (50 mM Tris-HCl, 5 mM CaCl2, pH = 7.5) at OD600 = 80. The wet pellets were stored at -80°C until further use.

HPLC-ESI and HRMS-based prenyltransferase activity assay
For the in vitro activity assay, the reaction mixture contained 4 mM DMAPP or GPP, 2 mM acceptor substrate, 6% (v/v) DMSO, and whole cell (OD600 = 80) in a total volume of 300 μL of 50 mM Tris-HCl buffer (pH 7.5) supplemented with 5 mM CaCl2. Reactions were incubated at 37°C for 8 or 12 h and quenched by addition of 900 μL methanol. After centrifugation at 13,800g for 15 min, the supernatants were analyzed by HPLC-ESI and high-resolution mass spectrometry (HRMS). All reactions were performed in triplicate. Gradient programs used for analysis are detailed in Supplementary Table 5 and Supplementary Figs. 32-54. For the substrate scope experiments, conversion rates were determined from the ratio of the product peak area to the substrate peak area in HPLC.

Product identification by NMR
Recombinant E. coli strains harboring pET28a-AtaPT-M8, pET28a-AtaPT-G326W, pET28a-AtaPT-S177G, or pET28a-AtaPT-M7 were prepared and induced as described above. After induction, the whole cells were harvested by centrifugation and washed with 50 mM Tris-HCl buffer (pH 7.5) containing 5 mM CaCl2. The cells were then resuspended in the same buffer, and the optical density was adjusted to OD600 = 80. The reaction mixture (total volume: 50 mL in a 250 mL Erlenmeyer flask) was prepared by adding 2 mM acceptor substrate and 4 mM DMAPP or GPP. Bioconversion was carried out at 37°C with shaking for 8-12 h. After the reaction, the mixture was extracted with an equal volume of ethyl acetate and centrifuged at 6438g for 10 min. The supernatant was collected and transferred to a separating funnel, where the aqueous phase was discarded, and the organic phase was retained. The solvent was removed under reduced pressure, and the resulting residue was dissolved in methanol and purified by reverse-phase semi-preparative HPLC. The purified products were dissolved in DMSO-d6 and characterized by 1H and 13C NMR spectroscopy (Supplementary Figs. 24, 55–119).

Molecular docking and Molecular dynamics (MD) simulations
Molecular docking of compound 1a and GPP into the M7 variant was conducted using the Glide module Schrödinger, LLC, New York, NY, 2024) in Standard Precision (SP) mode. Prior to docking, the M7 structure was prepared by optimizing the hydrogen bond network, removing water molecules, and energy minimization in Protein Preparation Wizard. Compound 1a and GPP were prepared using Ligprep. A receptor grid box centered on the active site was generated using the Receptor Grid Generation module. The docking poses consistent with in vitro enzymatic assay results were selected for molecular dynamics (MD) simulations.
MD simulations were carried out using the Desmond. The protein-ligand complexes were solvated in an SPC water model, and the systems were neutralized with 0.15 M NaCl. All simulations employed the OPLS4 force field, with the default cutoff values used for Coulombic interactions. Following energy minimization, the systems were equilibrated using both NVT and NPT ensembles. Subsequently, production runs were performed for 250 ns at 300 K and 1 atm. And Simulation Interactions Diagram was employed to evaluate the stability and binding interactions of the ligand-protein complexes.

Construction and characterization of other PTs mutants
Using AtaPT as a template for BLAST searches against the NCBI database, three homologous enzymes with distinct degrees of sequence identity were selected: AfaPT (94.81%, NCBI GenBank: KAL5359867.1), AaaPT (86.7%, NCBI GenBank: XP_073546247.1), and AmaPT (56.87%, NCBI GenBank: XP_045942482.1). The DNA sequences of AfaPT, AaaPT, and AmaPT were synthesized by General Biosystems (Anhui) Corp. Ltd and cloned into the NdeI–XhoI sites of the pET28a vector. The resulting recombinant plasmids (pET28a-AfaPT, pET28a-AaaPT, and pET28a-AmaPT) were transformed into Escherichia coli BL21(DE3) for heterologous expression.
E. coli cells were cultured overnight in 5 mL of Luria–Bertani (LB) medium supplemented with 50 μg/mL kanamycin at 37°C with shaking at 220 rpm. The overnight cultures were then inoculated (1%, v/v) into 500 mL of TB medium containing 50 μg/mL kanamycin and grown at 37°C and 220 rpm in 2 L shake flasks. When the OD600 reached 0.8–1.0, recombinant protein expression was induced with 0.2 mM IPTG, followed by incubation at 18°C and 220 rpm for 16 h. Engineered mutants of the PT homologs were constructed by PCR using KOD One™ PCR Master Mix.
Whole-cell biotransformation assays were performed in a final volume of 0.3 mL containing 50 mM Tris–HCl buffer (pH 7.5), 5 mM CaCl2, 4 mM DMAPP or GPP, 2 mM acceptor substrate 1a, 6% (v/v) DMSO, and enzyme preparations adjusted to an OD600 of 80. Reactions were conducted at 37°C for 8 h and terminated by the addition of 0.9 mL methanol. After centrifugation at 13,800×g for 15 min, the supernatants were subjected to HPLC analysis. All experiments were performed in triplicate (n=3).

Coupling of IUP with AtaPT-M8/G326W/M7 for the preparation of prenyl flavonoids using a lysate-based catalyst
The cascade biosynthesis of prenyl flavonoids was optimized using mixed whole-cell lysates of E. coli BL21(DE3) harboring pET28a-M8 and PCDF-MjIPK-EcPK74, with substrate 1a. The influence of various parameters was investigated, including expression temperature (18, 25, 30, or 37°C), IPTG concentration (0.2, 0.5, 0.8, or 1 mM), reaction temperature and pH (18, 25, 30, or 37°C; pH 7.0, 7.5, 8.0, 8.5, or 9.0), enzyme lysate ratio (PCDF-MjIPK-EcPK: pET28a-M8 = 1:3, 1:2, 1:1, 2:1, or 3:1), DMSO content (1-10% v/v), Mg2+ concentration (0, 2, 5, or 10 mM), prenol concentration (2–30 mM), and cell density (OD600 = 40-200). The reaction reached a steady state after approximately 10 h. Reactions were terminated by adding three volumes of methanol, followed by centrifugation at 13,800g for 15 min. Products were analyzed by HPLC, and all concentrations are reported as mean±SD from three biological replicates (Supplementary Figs. 25–27).
For strain cultivation, E. coli harboring the relevant plasmids was grown in TB medium containing 50 μg/mL kanamycin or 100 μg/mL spectinomycin at 37°C until OD600 reached 0.8–1.0. IPTG (0.2 or 0.5 mM) was added to induce expression, followed by incubation at 18 or 25°C for 16 h. To enable large-scale production, a high-density fermentation method was also established using glucose as a carbon source. This process is cost-effective and enables high cell density (Supplementary Table 13). Cultures were grown at 37°C until OD600 reached 25-30, followed by induction with 1 mM IPTG and continued expression overnight at 18 or 25°C. During induction, glucose levels were maintained between 0.5–2 g/L until its consumption ceased. Cells were harvested by centrifugation at 6438g for 2 min at 4°C and resuspended in 50 mM Tris-HCl buffer (pH 7.5) to an OD600 of 120.
The cascade preparation of alkylated flavonoids was conducted with 2 mM of substrates 1a, 1f, or 1j, 10 mM prenol, and 2% DMSO in a total volume of 1 L. The reaction was carried out at 35°C for 10 h. Afterward, samples were extracted with two volumes of ethyl acetate and centrifuged at 6438g for 5 min. The supernatants were collected and the solvents evaporated under reduced pressure. The residues were dissolved in dichloromethane (DCM) and purified via silica gel chromatography (DCM: MeOH = 30: 1). The eluates were concentrated under reduced pressure to obtain the final products, which were subsequently dried.
Additionally, substrates were evaluated using pET28a-G326W and PCDF-MjIPK-EcPK, or pET28a-M7 combined with pET28a-THAIPK-Y70A/V130A/I140A (referred to as THAIPKG)77. The G326W combination exhibited higher reactivity toward 1a, 1e, and 1g, while the M7-THAIPKG system showed enhanced activity for 1a, 1b, 1e, 1g, and 1i, particularly in aromatic geranylation. Interestingly, replacing geraniol with nerol in reactions with substrates 1b and 1g resulted in notable activity, an unexpected yet intriguing outcome. The yield was determined by substituting the peak area into the standard calibration curve.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This work by Professor Wang and co-workers report the directed evolution of AtaPT, structural analysis and MD simulation of its variants and scalable synthesis of 26 C-prenylated flavonoids its variants. Especially, they found an aromatic cage governing both regioselectivity and donor specificity. This work expands the application the reported aromatic prenyltransferase AtaPT, and establishes a selective and scalable biocatalytic platform for flavonoid prenylation. It’s a very comprehensive work and achieved numerous results, thus it’s recommended to be accepted after the author address the following issues.\n1, In Introduction Section, AtaPT should be much more introduced, such as its catalytic properties, and so on. Thus, the reader can understand the advances in this work.\n2, Different from most prenylated products, the prenyl group in 2a was reverse instead of normal, why? Please give an explanation.\n3, Page 8: the authors stated: “Notably, neither WT nor the tested mutants (G326W, M5, M7, and M8) exhibited activity toward substrates 1t and 1u, likely due to the absence of hydroxyl groups on the A and B rings, which are presumed to be critical for substrate recognition and binding.” In view of the reaction mechanism, this deduction isn’t completely corrected.\n4, In Fig. 2: Considering easy-understanding for readers, HPLC chromatograms for the conversions of 1a by M8 and 1a by M7 should be provided here.\n5, The steady-state kinetic constants of AtaPT variants, such as Km = 1.271 mM, it should be written as Km = 1.2 or Km = 1.3. Km = 1.271 mM seems to be strange, and imply an accuracy that will not be reproducible.\n6, In scalable biocatalytic synthesis, why did the authors synthesized the compounds shown in Figure 5? Please give more description on these compounds, such as biological activities, etc.", "reviewer_2": "This is a well-executed study that makes significant contributions to biocatalysis and natural product synthesis. The authors successfully engineered AtaPT variants with dramatically improved activity and selectivity for flavonoid prenylation, identified a novel aromatic cage mechanism governing selectivity, and demonstrated practical applications through preparative-scale synthesis.\nThe work merits publication after addressing the following points.\nMajor Points:\n1. Mechanistic Clarity: While the aromatic cage concept is compelling, the manuscript would benefit from:\na.More detailed explanation of why the \"reverse\" 8-prenylation occurs with M8 versus normal prenylation;\nb. Quantitative analysis of substrate-cage interactions through computational binding energies;\nc. Discussion of how the cage mechanism relates to other prenyltransferases beyond AtaPT.\n2. Structure-Function Analysis:\na. The crystal structures show relatively weak electron density for substrate 1a. Additional validation through anomalous scattering or co-crystallization with substrate analogs would strengthen structural conclusions;\nb. MD simulations are mentioned but insufficiently detailed - include RMSD plots, interaction frequencies, and convergence analysis;\n3. Substrate Scope Limitations:\na. The inability to prenylate substrates 1t and 1u is attributed to missing hydroxyl groups, but this hypothesis needs experimental validation;\nb. Test additional substrates with varied hydroxylation patterns to establish clearer structure-activity relationships.\n4.Scale-up Considerations:\na. While 400 mg/L titers are reported, provide space-time yields and total turnover numbers;\nb. Discuss enzyme stability over extended reaction times;\nc. Address the relatively low conversion (15.8%) with in situ GPP generation compared to direct addition.\nMinor Points:\n1. Methods Section:\na. Include detailed protocols for the high-density fermentation mentioned;\nb. Clarify the criteria for selecting mutation sites in directed evolution;\n2. Data Presentation:\na. Figure 2d would benefit from error bars on all data points;\nb. Include a supplementary table comparing kinetic parameters of all variants with all tested substrates;\nc. Provide NMR spectra for all newly reported compounds in the SI;\n3. Writing and Clarity:\na. Define abbreviations (SSM, ISM, FRISM) at first use in the main text;\nb. The introduction could be condensed to focus more on the specific challenges addressed;\nc. Clarify the distinction between \"reverse\" and normal prenylation earlier in the text.\n4. Comparative Analysis:\na. Include a table comparing your results with other reported prenyltransferases for the same substrates;\nb. Discuss how your aromatic cage mechanism relates to selectivity mechanisms in other enzyme families.\n5. Practical Applications:\na. Discuss potential pharmaceutical applications of the newly synthesized compounds;\nb. Address the cost-effectiveness of the biocatalytic approach versus chemical synthesis;\nc. Comment on the environmental benefits of your approach.\n6. Statistical Analysis:\na. Specify the number of independent experiments for all data;\nb. Include statistical significance tests when comparing variants;\nTechnical Corrections:\na. Page 5: \"Folk\" should be defined (farnesol kinase?);\nb. Figure 3 caption: Specify the resolution of the crystal structures;\nc. Supplementary Table 6 reference appears before proper introduction;\nd. Ensure consistent use of compound numbering throughout.\nStrengths of the manuscript:\n1. Novel mechanistic insights into prenyltransferase selectivity.\n2. Impressive 45-fold improvement in catalytic efficiency.\n3. Comprehensive substrate scope analysis.\n4. Strong structural biology support for mechanistic conclusions.\n5. Practical demonstration through preparative-scale synthesis.\nThis manuscript represents a significant advance in biocatalytic prenylation and provides valuable mechanistic insights. With the suggested revisions, particularly strengthening the mechanistic discussion and providing additional validation of the aromatic cage concept, this work would make an excellent contribution to Nature Communications. The combination of directed evolution, structural biology, and practical synthesis makes this a comprehensive study that will be of broad interest to the biocatalysis and natural products communities.", "reviewer_3": "This study focuses on the discovery and yield optimization of C-prenylated flavonoids by employing directed evolution strategies to engineer the active pocket of the prenyltransferase AtaPT. The proposed aromatic cage, which governs regioselectivity and donor specificity, holds significant scientific value. Furthermore, the research achieved high-yield bioproduction of specific prenylated flavonoids through an integrated donor regeneration system. Although progress has been made in scaling up the production of C-prenylated flavonoids, several unresolved issues remain perplexing.\n1. Is the aromatic cage governing regioselectivity and donor specificity applicable to similar prenyltransferases? Alternatively, is there more robust experimental evidence supporting the crucial role of the aromatic cage? Additionally, the ability to produce specific regiospecific substitution patterns is not uncommon for prenyltransferases, as numerous enzymes have already demonstrated remarkable plasticity in this regard.\n2. The current data appear insufficient to robustly support the proposed 'aromatic cage' concept. This is primarily because the experimental evidence involves too few amino acid residues. Crucially, the manuscript lacks experimental data for mutations of P324 to other aromatic amino acids, such as phenylalanine (P324F). The observed changes in activity might be primarily attributed to enhanced interactions between the introduced histidine residue and the substrate, rather than definitively demonstrating the formation or function of an aromatic cage.\n3. In the engineering of donor plasticity across multiple prenyltransferases, bulky residues are commonly substituted with smaller ones to expand the donor-binding cavity, thereby accommodating larger donor molecules. Consequently, the authors' explanation for the G326W mutant's inactivity toward GPP—compared to previous studies—does not present particularly novel insights.\n4. The use of DMSPP as an alternative donor to DMAPP for crystallographic studies of prenyltransferases has become an established methodology. Therefore, we recommend that the authors reattempt cocrystal acquisition using DMSPP to better elucidate the conformational arrangements of both donor- and acceptor-proximal residues.\n5. The manuscript predominantly employs percentage yield for quantitative description of the data, yet fails to specify the calculation methodology in the Methods section.\n6. The text contains numerous discrepancies between the descriptions and referenced figures/tables, or lacks adequate graphical support. For example – When overlapping the M7-1a model with the structure of M8, the closest distance between the phenolic ring of 1a and the C175M side chain is only 3 Å, suggesting a potential steric clash. A thorough revision of the entire manuscript is strongly recommended to address these issues.\n7. I have a few other minor considerations for revision.\nTo date, exactly how many aPTs have been experimentally validated to accept KAE as a substrate—six or seven?\nThe manuscript lacks structural characterization of compound 4a, and critically, the supporting materials fail to provide standardized NMR data listings, particularly for the new compounds.\nPlease provide the correct PDB accession codes for the structural data.\nHow was the reaction time determined for measuring the Michaelis-Menten kinetic parameters?"}, "author_response": "Point-by-point Responses\nReviewer #1 (Remarks to the Author):\nThis work by Professor Wang and co-workers report the directed evolution of AtaPT,\nstructural analysis and MD simulation of its variants and scalable synthesis of 26 C-\nprenylated flavonoids its variants. Especially, they found an aromatic cage governing\nboth regioselectivity and donor specificity. This work expands the application the\nreported aromatic prenyltransferase AtaPT, and establishes a selective and scalable\nbiocatalytic platform for flavonoid prenylation. It’s a very comprehensive work and\nachieved numerous results, thus it’s recommended to be accepted after the author\naddress the following issues.\nThank you for your positive feedback. We have revised the manuscript and supporting\ninformation based on your suggestions. The point-by-point responses are detailed\nbelow:\n1, In Introduction Section, AtaPT should be much more introduced, such as its catalytic\nproperties, and so on. Thus, the reader can understand the advances in this work.\nThank you for this valuable suggestion. To provide readers with a clearer understanding\nof AtaPT and its significance, we have added additional information in the revised\nIntroduction, describing its catalytic properties and related characteristics. The\nsupplemented text reads as follows: “The Dai group reported a prenyltransferase, AtaPT,\nfrom Aspergillus terreus, which exhibits remarkable substrate promiscuity toward a broad\nrange of aromatic acceptors (up to 72 types) and prenyl donors (with prenyl residues up\nto 20 carbons). This enzyme provides a powerful biocatalytic tool for the synthesis of\ndiverse prenylated compounds. Moreover, the structural elucidation of AtaPT has offered\nvaluable insights for further engineering toward enzymes capable of constructing unique\nbioactive scaffolds. Building upon this pioneering work, significant progress has been\nmade in tailoring its selectivity toward flavonoids and chalcones. However, given its\nexceptional catalytic potential, a deeper understanding of the mechanisms governing its\nactivity and selectivity is essential for establishing a versatile platform for efficient and\nselective prenylation.” Relevant information has been added to the fourth paragraph of\nthe Introduction section (Page 3).\n2, Different from most prenylated products, the prenyl group in 2a was reverse instead\nof normal, why? Please give an explanation.\nThank you very much for pointing out this unclear description. In the revised\nmanuscript, we provided the crystal structure of M8 in complex with 1a and DMSPP.\nThe structure shows that 1a is bound in the active site by forming an aromatic cage on\none side and interacting with C175M and L252 on the other side. Hydrogen bonds\nformed between 1a and P324H, E91 and R396 further stabilize 1a in a configuration\nthat bring its C8 position into close proximity with the C3 atom of DMSPP (5.2 Å). The\nhydrogen bond between E91 and the hydroxyl group on C7 may facilitate the attack of\nC8 on C3 of DMAPP, thereby promoting the formation of the unusual reverse product\n2a via M8 (Figure 1R1, Figure 3, Page 25).\nFigure 1R1. The crystal structure of M8 in complex with 1a and DMSPP\n3, Page 8: the authors stated: “Notably, neither WT nor the tested mutants (G326W, M5,\nM7, and M8) exhibited activity toward substrates 1t and 1u, likely due to the absence\nof hydroxyl groups on the A and B rings, which are presumed to be critical for substrate\nrecognition and binding.” In view of the reaction mechanism, this deduction isn’t\ncompletely corrected.\nThank you for this insightful comment. To further clarify this point, we tested five\nadditional substrates (1v–1z) (Figure. 1R2 and Figure. 5). The results (Figures 1R3-1R6,\nSupplementary Figures 51-54) show that 1v, although containing a hydroxyl group on\nring A, remained inactive—likely due to an ortho effect. Substrates 1w and 1x displayed\nonly weak activity, suggesting that both the number and position of hydroxyl groups on\nring A strongly influence catalytic performance. In contrast, substrates 1y and 1z, which\nbear hydroxyl groups on ring B, exhibited good reactivity with mutants G326W and M7,\nindicating that hydroxyl substitution on ring B is more\nfavorable for substrate recognition and catalysis. Overall, these results demonstrate that\nthe distribution and position of hydroxyl groups significantly affect enzyme activity.\nThe corresponding discussion has been revised and incorporated into the substrate scope\nsection of the manuscript (Page 9).\nFigure 1R2. The structural formula of the addition tested substrates.\nFigure 1R3. HPLC-DAD/HRMS analysis of the prenylation of 1w catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1w and its products 2w. b, HPLC\nchromatogram of 1w and its product 3w and 4w. c-e, Typical negative ion HRMS\nspectra for product 2w (c), 3w (d), and 4w (e). Molecular weights: 1w = 238; 2w = 306;\n3w, and 4w = 374. The detection wavelength is 308 nm.\nFigure 1R4. HPLC-DAD/HRMS analysis of the prenylation of 1x catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1x and its products 2x and 3x. b, HPLC\nchromatogram of 1x and its product 4x and 5x. c-f, Typical negative ion HRMS spectra\nfor product 2x (c), 3x (d), 4x (e) and 5x (f). Molecular weights: 1x = 238; 2x, and 3x =\n306; 4x, and 5x = 374. The detection wavelength is 308 nm.\nFigure 1R5. HPLC-DAD/HRMS analysis of the prenylation of 1y catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1y and its products 2y. b, HPLC\nchromatogram of 1y and its product 3y. c-d, Typical negative ion HRMS spectra for\nproduct 2y (c), and 3y (d). Molecular weights: 1y = 238; 2y = 306; 3y = 374. The\ndetection wavelength is 308 nm.\nFigure 1R6. HPLC-DAD/HRMS analysis of the prenylation of 1z catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1z and its products 2z. b, HPLC\nchromatogram of 1z and its product 3z. c-d, Typical negative ion HRMS spectra for\nproduct 2z (c), and 3z (d). Molecular weights: 1z = 238; 2z = 306; 3z= 374. The\ndetection wavelength is 320 nm.\n4, In Fig. 2: Considering easy-understanding for readers, HPLC chromatograms for the\nconversions of 1a by M8 and 1a by M7 should be provided here.\nThank you for your constructive suggestion. We agree that including these\nchromatograms will help readers better understand the experimental results.\nAccordingly, we have added the HPLC chromatograms for the conversions of 1a by M8\nand 1a by M7 to Figure 2 in the revised manuscript (Page 24).\nFigure 1R7. Evolution of AtaPT for efficient synthesis of prenylated kempferol\nderivatives. a, Reactions starting with KAE to form 8P-KAE and Icaritin. b, HPLC\nchromatogram showing the conversion of 1a with DMAPP to products 2a, 3a, 4a, and 5a\ncatalyzed by AtaPT-WT. c, HPLC chromatogram of 1a with GPP resulting in the formation\nof product 6a catalyzed by AtaPT-WT. d, Improvement in catalytic activity across five\nrounds of directed evolution using site-saturation mutagenesis (SSM), iterative saturation\nmutagenesis (ISM), and focused rational iterative site mutagenesis (FRISM). M2:\nS177G/P324H, M3: S177G/P324H/N328M, M5: E169A/S170Q/ S177A/P324H/N328M,\nM7: I167T/E169A/S170Q/S177A/L323Q/P324H/N328M and M8:\nI167A/E169A/S170Q/C175M/S177A/L323Q/P324H/N328R. e, Rescreening of mutants\nusing GPP to enhance the conversion efficiency of 6a. f, HPLC chromatogram showing the\nconversion of 1a with DMAPP to products 2a, catalyzed by AtaPT-M8. g, HPLC\nchromatogram of 1a with GPP resulting in the formation of product 6a catalyzed by AtaPT-\nM7.\n5, The steady-state kinetic constants of AtaPT variants, such as K = 1.271 mM, it\nm\nshould be written as K = 1.2 or K = 1.3. K = 1.271 mM seems to be strange, and\nm m m\nimply an accuracy that will not be reproducible.\nThank you very much for your valuable suggestion. We have revised all numerical\nvalues to maintain consistent precision by retaining two decimal places. Accordingly,\nthe K value has been updated to 1.27 mM in the revised manuscript (Page 5).\nm\n6, In scalable biocatalytic synthesis, why did the authors synthesized the compounds\nshown in Figure 5? Please give more description on these compounds, such as\nbiological activities, etc.\nThank you for pointing this out. We agree that highlighting the practical utility of the\nsynthesized compounds is important. The preparation-scale synthesis of 2a, 2f, and 2j\nserves two purposes: (i) to validate the feasibility of the engineered mutants and the\nestablished cascade reaction, and (ii) to demonstrate the biological significance of the\nproducts. Specifically, 2a exhibits anti-breast cancer activity (Phytochemistry 130, 170-\n18 2016), 2j shows anti-diabetic and neuroprotective effects (Phytochemistry 111, 141-\n148, 2015; Bioorg. Chem. 114, 105098, 2021), and analogs of 2f possess antibacterial\nand cytotoxic properties (Nat. Prod. Res. 34, 1256-1263, 2019; Phytochemistry 164,\n33-40, 2019). These results underscore the synthetic value of our approach and provide\na solid foundation for future biological investigations. This information has been\nincorporated into the revised manuscript. Relevant information has been supplemented\nin the final paragraph of section 'Coupling the isopentenol utilization pathway with\nprenyltransferase for prenylflavonoid biosynthesis' (Page 11).\nReviewer #2 (Remarks to the Author):\nThis is a well-executed study that makes significant contributions to biocatalysis and\nnatural product synthesis. The authors successfully engineered AtaPT variants with\ndramatically improved activity and selectivity for flavonoid prenylation, identified a\nnovel aromatic cage mechanism governing selectivity, and demonstrated practical\napplications through preparative-scale synthesis.\nThe work merits publication after addressing the following points.\nThank you for your positive feedback. We have revised the manuscript and supporting\ninformation based on your suggestions. The point-by-point responses are detailed\nbelow:\nMajor Points:\n1. Mechanistic Clarity: While the aromatic cage concept is compelling, the manuscript\nwould benefit from:\na. More detailed explanation of why the \"reverse\" 8-prenylation occurs with M8 versus\nnormal prenylation;\nThank you very much for your helpful suggestion. We provided the crystal structure of M8\nin complex with 1a and DMSPP. The structure shows that 1a is bound in the active site by\nforming an aromatic cage on one side and interacting with C175M and L252 on the other\nside. Hydrogen bonds formed between 1a and P324H, E91 and R396 further stabilize 1a\nin a configuration that bring its C8 position into close proximity with the C3 atom of\nDMSPP (5.2Å). The hydrogen bond between E91 and the hydroxyl group on C7 may\nfacilitate the attack of C8 on C3 of DMAPP, thereby promoting the formation of the\nunusual reverse product 2a via M8 (Figure 2R1, Figure 3, Page 25).\nFigure 2R1. The crystal structure of M8 in complex with 1a and DMSPP\nb. Quantitative analysis of substrate-cage interactions through computational binding\nenergies;\nThank you for your insightful suggestion. We have provided crystal structure of M8 in\ncomplex with 1a and DMSPP in the revised manuscript, which shows clearly how 1a\ninteracts with the aromatic cage. Therefore, we have removed the related results of MD\nsimulations on M8.\nc. Discussion of how the cage mechanism relates to other prenyltransferases beyond\nAtaPT.\nThank you for your insightful suggestion. To explore whether the aromatic cage\nmechanism is applicable to other prenyltransferases, we performed mechanism-guided\nengineering on three additional prenyltransferases: AfaPT (NCBI GenBank:\nKAL5359867.1), AaaPT (NCBI GenBank: XP_073546247.1), and AmaPT (NCBI\nGenBank: XP_045942482.1), which share 94.81%, 86.70%, and 56.87% sequence identity\nwith AtaPT, respectively. Sequence alignment revealed that all four enzymes conserve the\nXXLPXX motif, suggesting that the aromatic cage mechanism may be generalizable.\nAdditionally, the loop region (residues 167–170) is highly similar among these enzymes\n(Figure 2R2, Supplementary Figure 20). We introduced the corresponding M8 and M7\nmutations into the homologs. Variants derived from AfaPT and AaaPT showed enhanced\nactivity and selectivity toward their respective products (Figure 2R3, Figure 4c and\nSupplementary Figure 22). In contrast, AmaPT, with lower sequence homology, exhibited\nminimal activity in both the wild type and the engineered variants. Structure alignment\nanalysis revealed that AmaPT’s active pocket contains bulky residues (Y193, M253,\nM327) that hinder substrate accommodation. Subsequent mutagenesis to Y193S, M253L,\nand M327G enabled AmaPT variants to accept 1a as a substrate. Notably, unlike the other\ntested PTs, AmaPT preferentially produces 3a in both the wild-type and AmaPT-M8\nvariants. No detectable 2a formation was observed when using the wild-type enzyme.\nHowever, upon introducing mutations around the active pocket, two variants—\nY193S/M253L /M327G and AmaPT-M8-SLG—exhibited significantly enhanced activity\ntoward 2a, indicating that aromatic cage engineering plays a decisive role in modulating\nregioselectivity (Figure 2R3, Figure 4e and Supplementary Figure 22). These results\nindicate that the aromatic cage mechanism\nidentified in AtaPT is not unique but can be successfully recapitulated in its homologous\nenzymes, providing a generalizable strategy for engineering prenyltransferases. Relevant\ninformation has been added to the section of ‘Mechanism-guided design of PTs beyond\nAtaPT (Page 8).\nFigure 2R2. Sequence alignments of AtaPT with AfaPT, AaaPT, and AmaPT.\nFigure 2R3. Activity tests of other PT variants. a, Activity tests of AfaPT-WT and\nvariants. b, Activity tests of AaaPT-WT and variants. c and d, Activity tests of AmaPT-\nWT and variants. Whole cells harboring AtaPT mutants were resuspended in 50 mM\nTris-HCl buffer (pH 7.5) containing 5 mM CaCl2 and 6% (v/v) DMSO, with the cell\ndensity adjusted to OD = 80. Substrate 1a and DMAPP/GPP were added at final\n600\nconcentrations of 2 mM and 4 mM, respectively, in a total reaction volume of 300 μL.\nBioconversion reactions were performed at 37 °C with shaking at 1200 rpm for 24 h.\nRelative activities are expressed as mean ± SD from three independent biological\nreplicates (n = 3).\n2. Structure-Function Analysis:\na. The crystal structures show relatively weak electron density for substrate 1a.\nAdditional validation through anomalous scattering or co-crystallization with substrate\nanalogs would strengthen structural conclusions;\nThank you very much for pointing this out. We have provided the structure of M8 in\ncomplex with 1a and DMSPP in the revised manuscript. We also tried to obtain similar\nstructure of M7 by soaking GSPP and 1a or different substrate analogs, but only\nobserved GSPP in the final structure. GSPP is actually flexible in the active site, as we\nonly observed one whole GSPP molecule in the dimeric structure of M7.\nb. MD simulations are mentioned but insufficiently detailed - include RMSD plots,\ninteraction frequencies, and convergence analysis;\nThank you very much for your valuable suggestions. As we have now obtained the ternary\ncomplex structure of M8 with 1a and DMSPP, the MD simulation data for M8 have been\nremoved from the revised manuscript. Instead, we have provided more detailed molecular\ndynamics analyses for the M7 complex, including RMSD analyses of 1a and GPP during\nthe simulations, interaction frequency analyses, and measurements of the distance\n′\nbetween the 3-position of 1a and the electrophilic center of the geranyl donor. The RMSD\nfluctuations of both 1a and GPP throughout the simulations indicate that the predicted\nbinding pose of 1a remains conformationally stable within the active site of M7 (Figure\n2R4 and Supplementary Figure 18).\nFigure 2R4. Docking model and molecular dynamics simulations of the M7-1a-GPP\ncomplex\n3. Substrate Scope Limitations:\na. The inability to prenylate substrates 1t and 1u is attributed to missing hydroxyl\ngroups, but this hypothesis needs experimental validation;\nb. Test additional substrates with varied hydroxylation patterns to establish clearer\nstructure-activity relationships.\nThank you for these thoughtful suggestions. Since comments (a) and (b) address the\nsame issue, we respond to them together. Following your advice, we tested five\nadditional substrates (1v–1z, Figure 2R5, Figure 5) bearing a single hydroxyl group at\ndifferent positions on the A or B ring. The results showed that 1v was inactive, while\nthe other four substrates were accepted by AtaPT variants. Notably, in 1v, the hydroxyl\ngroup is located at the meta position, which can readily form a hydrogen bond with the\ncarbonyl group of the C-ring. This interaction likely hinders the formation of the reactive\ncarbon anion, explaining its inactivity. These findings experimentally confirm that both\nthe position and number of hydroxyl groups play critical roles in substrate recognition\nand catalytic efficiency. Based on these results, we have revised the corresponding\ndiscussion in the main text and Supporting Information (Figures 2R6-2R9,\nSupplementary Figures 51-54).\nFigure 2R5. The structural formula of the addition tested substrates.\nFigure 2R6. HPLC-DAD/HRMS analysis of the prenylation of 1w catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1w and its products 2w. b, HPLC\nchromatogram of 1w and its product 3w and 4w. c-e, Typical negative ion HRMS\nspectra for product 2w (c), 3w (d), and 4w (e). Molecular weights: 1w = 238; 2w = 306;\n3w, and 4w = 374. The detection wavelength is 308 nm.\nFigure 2R7. HPLC-DAD/HRMS analysis of the prenylation of 1x catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1x and its products 2x and 3x. b, HPLC\nchromatogram of 1x and its product 4x and 5x. c-f, Typical negative ion HRMS spectra\nfor product 2x (c), 3x (d), 4x (e) and 5x (f). Molecular weights: 1x = 238; 2x, and 3x =\n306; 4x, and 5x = 374. The detection wavelength is 308 nm.\nFigure 2R8. HPLC-DAD/HRMS analysis of the prenylation of 1y catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1y and its products 2y. b, HPLC\nchromatogram of 1y and its product 3y. c-d, Typical negative ion HRMS spectra for\nproduct 2y (c), and 3y (d). Molecular weights: 1y = 238; 2y = 306; 3y = 374. The\ndetection wavelength is 308 nm.\nFigure 2R9. HPLC-DAD/HRMS analysis of the prenylation of 1z catalyzed by\nAtaPT mutants. a, HPLC chromatogram of 1z and its products 2z. b, HPLC\nchromatogram of 1z and its product 3z. c-d, Typical negative ion HRMS spectra for\nproduct 2z (c), and 3z (d). Molecular weights: 1z = 238; 2z = 306; 3z= 374. The\ndetection wavelength is 320 nm.\n4.Scale-up Considerations:\na. While 400 mg/L titers are reported, provide space-time yields and total turnover\nnumbers;\nThank you for your helpful comment. We have now included the space–time yields and\ntotal turnover numbers (TONs) in the revised manuscript. Based on the 10-hour reaction\ntime of the scale-up cascade system, the space–time yields for 2a, 2f, and 2j were\ncalculated to be 21.8, 22.1, and 44.2 mg L-1 h-1, respectively. For TON determination, the\ncascade reactions were performed using prenol as the donor substrate, with the lysate of\nPCDF-MjIPK-EcPK and the purified M8 enzyme catalyzing 1a, 1f, and 1j, respectively.\nThe calculated TONs were 12, 21, and 27, as summarized in Table 2R1.\nTable 2R1. TON for 1a/1f/1j Catalyzed by AtaPT-M8.\nspace-time yields (mg L-1\nEntry Reaction Enzyme TON\nh-1)\n1 1a→2a AtaPT-M8 21.8 12\n2 1f→2f AtaPT-M8 22.1 21\n3 1j→2j AtaPT-M8 44.2 27\nEnzymatic assays containing 50mM Tris-HCl, pH 7.5, 10 mM prenol, 100 μL lysate of\nPCDF-MjIPK-EcPK, 20 μM purified enzyme and varying concentrations (500, 1000,\n1500, 2000 µM) of 1a/1f/1j was performed at 35 °C for 10 h in a final volume of 200\nμL. All the reactions were terminated by adding 600 μL methanol and centrifuged at\n12,000 rpm for 15 min.\nb. Discuss enzyme stability over extended reaction times;\nThank you for your insightful comment. We investigated the stability of the purified\nenzymes M8, G326W, and M7 over extended reaction times. The reaction profiles showed\nthat product formation plateaued after approximately 8 hours. When additional\nDMAPP/GPP was supplemented at this point, a further increase in product yield was\nobserved, indicating that enzyme activity was maintained beyond the initial reaction phase.\nTo further validate this, we collected the supernatant after 8 hours, added fresh substrate\nalong with DMAPP/GPP, and continued the reaction. The successful formation of the\ncorresponding products confirmed that the enzymes retained catalytic activity over an\nextended period (Figure 2R10, Supplementary Figure 14). These results suggest that the\ncurrent limitation lies in donor substrate depletion rather than enzyme instability,\nhighlighting the need for a more efficient donor regeneration system in future applications.\nAll these contents have been included in the revised manuscript.\nFigure 2R10. Temporal Stability of purified enzyme. a, Time-course analysis of the\nreaction by AtaPT-M8. b, Time-course analysis of the reaction by AtaPT-G326W. c,\nTime-course analysis of the reaction by AtaPT-M7. d, Test of purified enzyme. After 8\nhours, the reaction was continued by adding identical substrate for another 8 hours. In a\nsecond approach, after 8 hours, centrifuge and collect the supernatant, a different\nsubstrate was added and the reaction was allowed to proceed for 8 hours. Enzymatic\nassays containing 1mM prenyl acceptor, 2mM DMAPP (2mM prenyl acceptor, 4mM\nGPP for M7) and 20μM purified enzyme, 37°C for 8/16 h in a final volume of 200 μL\n(50mM Tris-HCl, 5mM CaCl , pH 7.5). All the reactions were terminated by adding 600\n2\nμL methanol and centrifuged at 12,000 rpm for 15 min.\nc. Address the relatively low conversion (15.8%) with in situ GPP generation compared\nto direct addition.\nThank you for your valuable comment. The relatively low conversion observed with the\nin situ GPP generation system is primarily due to insufficient GPP supply. In theory,\ngeraniol is first phosphorylated to geranyl phosphate (GP) by geraniol kinase and\nsubsequently converted to GPP via isopentenyl phosphate kinase (IPK). Initially, we\nconstructed the PCDF-Folk-THAIPKG system following literature reports; however,\nFolk exhibited poor expression in E. coli and interfered with THAIPKG expression,\nresulting in minimal product formation in the cascade with pET28a-M7. Interestingly,\nthe PCDF-THAIPKG and pET28a-M7 cascade alone performed well, leading us to\nadopt this system for geranyl-derived product synthesis.\nTo further investigate in situ GPP biosynthesis, we purified THAIPKG and M7 and\nconducted an in vitro cascade reaction, but no product was detected, suggesting that\nTHAIPKG cannot directly diphosphorylate geraniol to GPP. This implies that an\nendogenous E. coli enzyme likely catalyzes the conversion of geraniol to GP. We\ntherefore examined EcPK and EcThiK (NCBI GenBank: CAD6016682.1), but neither\nenzyme produced GP. We are currently exploring potential endogenous enzymes\nresponsible for this step. Identification of such an enzyme and elucidation of the\ncomplete GPP biosynthetic pathway may substantially enhance the overall conversion\nyield.\nMinor Points:\n1. Methods Section:\na. Include detailed protocols for the high-density fermentation mentioned;\nThanks very much for your reminder. Firstly, the high-density fermentation expression\nformula is presented in tabular form within the supplementary materials (Supplementary\nTable 13), as detailed below. In accordance with the protocol, the medium was prepared\nin a fermentation tank and sterilized (the medium is acidic at this stage, with a pH around\n4.5–5). After cooling, ensure adequate aeration to prevent contamination, and\nsimultaneously prepare 200 mL of TB inoculated and cultured overnight to serve as the\nseed culture.\nEarly the next morning, adjust the pH to approximately 6.9–7.0 using 50% ammonium\nhydroxide, then add 25 g of glucose (pre-dissolved in water and sterilized). Inoculate\nwith the seed culture and supplement with kanamycin (Km, 50 μg/mL). Incubate at 37 °\nC and 600 rpm for approximately 5–6 h, until the OD600 reaches 25–30. Subsequently,\ncool the culture to 18°C, add 200 μL of antifoam agent, and induce\nprotein expression by adding 1 mM IPTG.\nThroughout this process, the 50% ammonium hydroxide is added automatically to\nmaintain the pH around 7.0 (not exceeding 7.4). If a sugar electrode is not available to\nmonitor glucose, periodically withdraw a small volume of culture for sugar testing with\ntest strips, and measure OD as well. The objective is to maintain glucose\n600\nconcentration in the range of 0.5–2 g/L (this is critical for enzyme activity). Continue\nuntil consumption ceases (i.e., OD stops increasing). Expression typically lasts for\n600\nmore than 16 h. After completion, harvest the cells for subsequent reactions.\nTable 2R2. Fermenter high density fermentation expression formula.\nMass\nComponents\nConcentration (take 2 L as an example)\nMgSO ·7H O 1 g/L 2 g\n4 2\n(NH4) SO 5 g/L 10 g\n2 4\nKH PO 3 g/L 6 g\n2 4\nCitric acid 2 g/L 4 g\nMethionine 1 g/L 2 g\nDefoamer 1 mL/L 2 mL\nTap water - to 2 L\nb. Clarify the criteria for selecting mutation sites in directed evolution;\nThank you for pointing this out. In our directed evolution workflow, the first-round site-\ndirected saturation mutagenesis targeted 22 residues located within 5 Å of the active\npocket, as identified through substrate docking analysis. Based on the screening results,\nthe mutation P324H, which showed the most pronounced improvement in selectivity,\nwas selected as the template for the second round. Additional beneficial mutations were\nsubsequently introduced to further enhance activity (Supplementary Figure 4),\ngenerating variants M2 and M3.\nDuring iterative mutagenesis, we noted that activity improvements became incremental.\nTherefore, leveraging the amino acid information obtained from site-directed saturation\nmutagenesis, we performed two additional rounds of FRISM to explore combinatorial\neffects. This strategy focused on integrating mutations that positively influenced activity\nand/or selectivity, ultimately yielding variants M5, M8, and the unexpectedly high-\nperforming M7 (Supplementary Table 7).\nOverall, the selection of mutation sites was guided by the goal of maximizing catalytic\nactivity and product selectivity. Combinations of beneficial residues identified from\nprevious rounds were prioritized for recombination and screening, which effectively\nproduced the most favorable mutants.\nFigure 2R11. The process of directed evolution.\n2. Data Presentation:\na. Figure 2d would benefit from error bars on all data points;\nThank you very much for your reminder. The error bars for all data points were indeed\nincluded; however, because they were displayed in gray and the variations were small,\nthey were not clearly visible in the original figure. We have now replotted and updated\nFigure 2d in the revised manuscript to improve visual clarity. We apologize for any\nconfusion caused and appreciate your careful attention to this detail (Page 24).\nFigure 2R12. Evolution of AtaPT for efficient synthesis of prenylated kempferol\nderivatives. a, Reactions starting with KAE to form 8P-KAE and Icaritin. b, HPLC\nchromatogram showing the conversion of 1a with DMAPP to products 2a, 3a, 4a, and\n5a catalyzed by AtaPT-WT. c, HPLC chromatogram of 1a with GPP resulting in the\nformation of product 6a catalyzed by AtaPT-WT. d, Improvement in catalytic activity\nacross five rounds of directed evolution using site-saturation mutagenesis (SSM), iterative\nsaturation mutagenesis (ISM), and focused rational iterative site mutagenesis (FRISM). M2:\nS177G/P324H, M3: S177G/P324H/N328M, M5: E169A/S170Q/ S177A/P324H/N328M,\nM7: I167T/E169A/S170Q/S177A/L323Q/P324H/N328M and M8:\nI167A/E169A/S170Q/C175M/S177A/L323Q/P324H/N328R. e, Rescreening of mutants\nusing GPP to enhance the conversion efficiency of 6a. f, HPLC chromatogram showing the\nconversion of 1a with DMAPP to products 2a, catalyzed by AtaPT-M8. g, HPLC\nchromatogram of 1a with GPP resulting in the formation of product 6a catalyzed by AtaPT-\nM7.\nb. Include a supplementary table comparing kinetic parameters of all variants with all\ntested substrates;\nThank you very much for your valuable suggestion. We have compiled the kinetic\nparameters of all variants with all tested substrates and included the data in the revised\nsupplementary materials (Supplementary Table 9), as shown in the table below.\nTable 2R3. Kinetic test of AtaPT WT and mutants in Reactions of 1a.\nDonor Enzyme k cat (h-1) K m (μM) k cat / K m (M-1 S-1)\nWT - - -\nM8 6.23 ± 0.13 49.74 ± 5.37 34.79\nDMAPP\nG326W 1.17 ± 0.18 21.54 ± 2.11 15.09\nS177G - - -\nWT 5.13 ± 0.11 1270.52 ± 49.78 1.12\nGPP\nM7 38.22 ± 0.15 225.29 ± 6.96 47.12\nEnzymatic assays were performed in a total volume of 100 μL containing 50 mM Tris-\nHCl (pH 7.5), 5 mM CaCl2, 2/5/10 μM enzyme, 2 or 4 mM DMAPP or GPP (saturated),\nand varying concentrations of substrate 1a (40, 60, 80, 100, 150, 200, 250, 300, 500,\n1000, 1500, 2000, 2500, and 4000 μM). Reactions were incubated at 37 °C for 10 or 20\nmin and terminated by the addition of 300 μL methanol, followed by centrifugation at\n12,000 rpm for 15 min.\nc. Provide NMR spectra for all newly reported compounds in the SI;\nThank you for your suggestion. We have compiled the NMR spectra for all newly\nreported compounds described in this study. These spectra are now included in the\nrevised supplementary materials under the section “Characterization of prenylated\nproducts” (Page S136).\n3. Writing and Clarity:\na. Define abbreviations (SSM, ISM, FRISM) at first use in the main text;\nThank you for your reminder. We have revised the manuscript to define all\nabbreviations—SSM (site-saturation mutagenesis), ISM (iterative saturation\nmutagenesis), and FRISM (focused rational iterative site mutagenesis)—at their first\noccurrence in the main text, and have included the corresponding references (Page 4).\nWe appreciate your careful attention to detail.\nb. The introduction could be condensed to focus more on the specific challenges\naddressed;\nThank you very much for your suggestion. To make the introduction more concise and\nfocused on the specific challenges addressed in this study, we have removed redundant\nbackground information and general context (e.g., the original first paragraph). These\nrevisions streamline the narrative while retaining all essential content relevant to the\nstudy’s objectives. The updated introduction is included in the revised manuscript.\nc. Clarify the distinction between \"reverse\" and normal prenylation earlier in the text.\nThank you very much for your suggestion. We have added a clear explanation\ndistinguishing “reverse prenylation” from normal prenylation in the Introduction (first\nparagraph) to aid reader understanding (page 2). In addition, we have revised the legend of\nFigure 1 to include this clarification in the revised manuscript (Page 23).\n4. Comparative Analysis:\na. Include a table comparing your results with other reported prenyltransferases for the\nsame substrates;\nThank you for your valuable suggestion. We agree that comparing our results with\npreviously reported prenyltransferases acting on the same substrate (1a) provides\nimportant context. Accordingly, we have compiled the relevant data and included them\nin the revised manuscript as Supplementary Table 6 (Table 2R4 below), which\nsummarizes the reported activities of prenyltransferases toward kaempferol.\nTable 2R4. Previous work on the activity of PTs on kaempferol.\nConcentration Conversion Concentration Expressing\nEnzyme Ref.\nRate strain\nof substrate of product\n0.1 mM 98.25% 0.098 mM A. thaliana 8\nLaPT2\n0.7 mM 4.6% 0.032 mM S. cerevisiae 9\nEsPT2 0.2 mM 55% 0.11 mM S. cerevisiae 10\ntEsPT - - >200 mg/L Y. lipolytica 11\nEpPT8 0.2 mM 65% 0.13 mM S. cerevisiae 12\nSfFPT 0.4 mM 13% 0.052 mM S. cerevisiae 13\nEkF8DT3 - - >33.1 mg/L S. cerevisiae 14\nComplex\nAtaPT 0.4 mM <65% E. coli 1\nproduct\nb. Discuss how your aromatic cage mechanism relates to selectivity mechanisms in\nother enzyme families.\nThank you for your insightful suggestion. To our knowledge, this study is the first to\ndefine an aromatic cage in prenyltransferases (PTs) as a structural feature regulating\nselectivity. To assess whether this mechanism could be generalized, we performed\nmechanism-guided modifications of the aromatic cage in three additional PTs. The\nresults, provided in the response to Major Point 1c, demonstrate that the aromatic cage\ncan similarly influence selectivity in these homologous enzymes. These findings\nindicate that the aromatic cage mechanism is not unique to AtaPT and suggest potential\nparallels with selectivity-determining structural motifs in other enzyme families.\nRelevant information has been added to the section of ‘Mechanism-guided design of\nPTs beyond AtaPT’ (Page 8).\n5. Practical Applications:\na. Discuss potential pharmaceutical applications of the newly synthesized compounds;\nThank you for your comments. The newly synthesized compounds exhibit notable\nbiological activities: 2a shows anti-breast cancer activity (Phytochemistry 130, 170-18\n2016), 2j displays anti-diabetic and neuroprotective effects (Phytochemistry 111, 141-\n148, 2015; Bioorg. Chem. 114, 105098, 2021), and analogs of 2f possess antibacterial\nand cytotoxic properties (Nat. Prod. Res. 34, 1256-1263, 2019; Phytochemistry 164, 33-\n40, 2019). These results highlight both their synthetic value and potential pharmaceutical\napplications, providing a strong foundation for further investigation. This information\nhas been incorporated into the revised manuscript. Relevant information has been\nsupplemented in the final paragraph of section 'Coupling the isopentenol utilization\npathway with prenyltransferase for prenylflavonoid biosynthesis' (Page 11). In addition,\nwe have initiated collaborative studies with other research teams to explore the specific\ntherapeutic potential of these compounds.\nb. Address the cost-effectiveness of the biocatalytic approach versus chemical synthesis;\nThank you for your comments. We have added the following statement to the revised\nmanuscript: Biocatalysis offers superior selectivity compared with traditional chemical\nsynthesis, leading to a higher proportion of target products in the reaction mixture. This\nenhanced selectivity simplifies downstream purification and significantly reduces post-\nreaction processing costs, thereby improving the overall cost-effectiveness of the\nbiocatalytic approach (Page 2).\nc. Comment on the environmental benefits of your approach.\nThank you for your comments. We have added the following statement to the revised\nmanuscript to address this point: The developed approach enables the efficient\nbioproduction of prenylated flavonoids, thereby reducing reliance on large-scale plant\ncultivation and minimizing waste associated with traditional extraction methods that\ntypically yield low product content. Moreover, the reactions are performed in aqueous\nmedia under mild temperature conditions, which decreases energy consumption and\nchemical waste, contributing to a more sustainable and environmentally friendly\nproduction process (Page 12).\n6. Statistical Analysis:\na. Specify the number of independent experiments for all data;\nThank you very much for your reminder. We have indicated the number of experiments\nin each figure caption (n = 3). We apologize for any confusion caused and appreciate\nyour attention to detail.\nb. Include statistical significance tests when comparing variants;\nThank you for pointing this out. In this study, each reaction test was performed in\ntriplicate, and the results were presented in a bar chart or broken line chart.\nTechnical Corrections:\na. Page 5: \"Folk\" should be defined (farnesol kinase?);\nThank you for identifying the unclear description. To make it clear, we add the\nfollowing supplement in the revised manuscript: “Folk is a farnesol kinase derived from\nArabidopsis thaliana and can accept geraniol as a substrate, whereas THAIPK is an\nisopentenyl phosphate kinase derived from Thermoplasma acidophilum, whose\nnatural substrate is DMAP; however, engineered variants of THAIPKG (THAIPK-\nY70A!V130A!I140A) can accommodate GP.” (Page 11).\nb. Figure 3 caption: Specify the resolution of the crystal structures;\nThanks very much for your reminder. We have specified the resolution of the crystals\nin the revised manuscript as: we determined the crystal structures of M8 (2.50Å, PDB:\n9V5G), G326W (2.69 Å, PDB: 9V5F) and M7 (2.20 Å, PDB: 9V3U) (Supplementary\nTable 10, Page 6).\nc. Supplementary Table 6 reference appears before proper introduction;\nThank you for pointing this out. We have adjusted it in the revised version.\nd. Ensure consistent use of compound numbering throughout.\nThanks very much for your reminder. We have carefully verified and ensured the\nuniformity of the compound numbering. We apologize for any confusion caused and\nappreciate your attention to detail.\nStrengths of the manuscript:\n1. Novel mechanistic insights into prenyltransferase selectivity.\n2. Impressive 45-fold improvement in catalytic efficiency.\n3. Comprehensive substrate scope analysis.\n4. Strong structural biology support for mechanistic conclusions.\n5. Practical demonstration through preparative-scale synthesis.\nThis manuscript represents a significant advance in biocatalytic prenylation and\nprovides valuable mechanistic insights. With the suggested revisions, particularly\nstrengthening the mechanistic discussion and providing additional validation of the\naromatic cage concept, this work would make an excellent contribution to Nature\nCommunications. The combination of directed evolution, structural biology, and\npractical synthesis makes this a comprehensive study that will be of broad interest to the\nbiocatalysis and natural products communities.\nThank you very much for your work and positive comments.\nReviewer #3 (Remarks to the Author):\nThis study focuses on the discovery and yield optimization of C-prenylated flavonoids by\nemploying directed evolution strategies to engineer the active pocket of the\nprenyltransferase AtaPT. The proposed aromatic cage, which governs regioselectivity and\ndonor specificity, holds significant scientific value. Furthermore, the research achieved\nhigh-yield bioproduction of specific prenylated flavonoids through an integrated donor\nregeneration system. Although progress has been made in scaling up the production of C-\nprenylated flavonoids, several unresolved issues remain perplexing.\nThank you very much for your comments. We have incorporated new experimental data\nand revised both the manuscript and Supporting Information in accordance with your\nsuggestions to address the highlighted points. The point-by-point responses are detailed\nbelow:\n1. Is the aromatic cage governing regioselectivity and donor specificity applicable to\nsimilar prenyltransferases? Alternatively, is there more robust experimental evidence\nsupporting the crucial role of the aromatic cage? Additionally, the ability to produce\nspecific regiospecific substitution patterns is not uncommon for prenyltransferases, as\nnumerous enzymes have already demonstrated remarkable plasticity in this regard.\nThank you for your insightful suggestions. To evaluate the generality of the aromatic\ncage mechanism in prenyltransferases (PTs), we performed mechanism-guided\nengineering on three additional PTs: AfaPT (NCBI GenBank: KAL5359867.1), AaaPT\n(NCBI GenBank: XP_073546247.1), and AmaPT (NCBI GenBank: XP_045942482.1),\nwhich share 94.81%, 86.70%, and 56.87% sequence identity with AtaPT, respectively.\nSequence alignment revealed that all four enzymes conserve the XXLPXX motif, and\nthe loop region (residues 167–170) shows high similarity (Figure 3R1, Supplementary\nFigure 20), suggesting that the aromatic cage mechanism may be generalizable across\nthese homologs.\nWe introduced the corresponding M8 and M7 mutations into these enzymes. Variants\nderived from AfaPT and AaaPT exhibited enhanced activity and selectivity toward their\nrespective products. In contrast, AmaPT, due to lower homology, showed minimal\nresponse in both the wild type and M8/M7-like variants (Figure 3R2, Supplementary\nFigure 22). Homologous alignment analysis revealed that AmaPT possesses bulky residues\n(Y193, M253, M327) in its active pocket; mutagenesis to Y193S, M253L, M327G enabled\nAmaPT variants to accommodate 1a as a substrate. Notably, unlike the other tested PTs,\nAmaPT preferentially produces 3a in both the wild-type and AmaPT-M8 variants. No\ndetectable 2a formation was observed when using the wild-type enzyme. However, upon\nintroducing mutations around the active pocket, two variants — AmaPT-\nY193S/M253L/M327G and AmaPT-M8-SLG — exhibited significantly enhanced activity\ntoward 2a, indicating that aromatic cage engineering plays a decisive\nrole in modulating regioselectivity.\nThese results demonstrate that the aromatic cage mechanism observed in AtaPT is not\nunique and can be recapitulated in homologous enzymes. Introduction of the aromatic\ncage consistently enhanced both activity and selectivity, confirming its crucial role.\nRelevant information has been added to the section of ‘Mechanism-guided design of\nPTs beyond AtaPT’ (Page 8).\nWe agree that prenyltransferases (PTs) with intrinsic regioselectivity are common,\nparticularly for flavonoid substrates. Most characterized PTs exhibit strong specificity\ntoward flavanones (e.g., naringenin), isoflavones (e.g., genistein), and chalcones (e.g.,\nisoliquiritigenin) (Adv. Synth. Catal. 355, 1817-1828, 2013; ChemBioChem 15, 1673-\n1681, 2014; ChemBioChem 12, 2280-2283, 2011), whereas activity toward flavonols\n(e.g., kaempferol) is often low or undetectable, with limited potential to expand\nsubstrate scope. Moreover, PTs capable of recognizing kaempferol are primarily plant-\nderived, and their heterologous expression and catalytic performance present significant\nchallenges (Front. Plant Sci. 12, 673337, 2021; Front. Plant Sci. 13, 1034943, 2022;\nSci. Bull. 66, 1906-1916, 2021). Investigations into regioselectivity for the same\nsubstrate are also relatively scarce. To date, only one example has been reported in\nwhich directed evolution of AnaPT expanded its active pocket, producing multiple\ndistinct di- or tri-substituted products from dihydrochalcone (phloretin), albeit with\nmodest activity and selectivity (J. Agric. Food Chem. 72, 8018-8026, 2024). Systematic\nmechanistic studies and rational engineering strategies that can simultaneously adjust\nselectivity and enhance activity—particularly for flavonol substrates—remain largely\nunexplored. Our work addresses this gap by providing a practical approach to elucidate\nthe underlying mechanism and enabling high-yield bioproduction of prenylated\nflavonoids.\n20 39 • 9 50\nAl I SQ Y\nA A A A t f a m a a a a P P P P T T T T L . . P M M M L L L P • A • P P A S A A D D T N I I S A S Q E Q G D Y D D D ELW E L L W W NN T N E L TT W T i AA N A TA EE I E N 1 L LN N L N 4 4 4 4C C C CD D D DY Y\n60 7? 60 99 100 110\nAtaPT KY A\nAfaPT YILPSLG FRR G EP I EISVK D T CR GLQ IGP AGT\nAaaPT I PSLG LPSLFGIFRRRPIGI.EP G EPEFI IIGGLGLSLHSSGHGHGHGPGHIPGE IHSVPKI D EISVK D T TCRI CRIGLQ GLQ IGP IGPAIGAGTT S\nAmaPT HR LP G FRR G EP I EISVK ONT T CR G SPIGP AGT K\nGLSHGGHP\n129 130 140 150 160 17\nAtaPT PLNS ?\nA A Am f a a a a P P P T T T P P P L L L N N S N S H S S F G D G G D D\nI\nE L E LE\nL\nC\nE\nS\nS\nVC\nG\nT V T S C S\nS\nS D\nD\nD D\n180 190 200 210 220\nAtaPT L KDGE P MIE IEVYHKtAPL\nA£aPT L KDGEV YFSP\"T IPK MQI E IEVYHRKT' APLR\nA A a m a a P i T n L F R K N D E G D D V V P I \"\" P Y K F 1 YFSGTILPVKG L V YFSTIP P KG IG L PS V n M i QQ I E , ATR E. IEVYH RTK S• • A PL G K S•\n240 250 269 270 289 290\nAtaPT PTDAS PPL G DC P SRL YL FRMDL S\nAfaPT PTDAS PPL G DC DPM SRL YL FRMDL I YWT GGL TDAST KGLE\nAaaPT P DAS PPL G DC DP SRL YL FRMDL I IiillYWTIG TDAGTIKGLE\nAmaPT PA GTMNGSL I GMVG DC PS SRL YL FRMDL I G GGV GG S S D L D LS T K K G G L L E E\nY\nY\nW\nW\nT\nT\nA A A A t f a m a a a a P P P P T T T T M M ET L L L A A A A 2P 1 2 I K K N L L L G G G D D O E E A A K C C A E E C GDE E A C L L I i AER A A P E D E R R R L L L P P P F G P I G G M I I L A AM M A E M V. K P PP P G GG G TT • K P P P i • • Q Q Q Q I I I I Y Y Y Y F F F F P P P P L L L L L L L L G G G G P N N N N D D D D F F F F I I I I A A A A D D D D A A A A L L L L\nD I\n360 37? 400 410\nAtaPT VE N RWL AYS KGPSMNIY VA\nAfaPT VQ N RWLGVAYS KGPS YD NIYYD VA\nAaaPT VE AWL AYS VA\nAmaPT MD N AWL SYS 4KGP KGPSMNIYNIYYD,VA\nN *Il YD\nAfaPT AGRV V\nAaaPT LIGSAKA\nAmaPT\nFigure 3R1. Sequence alignments of AtaPT with AfaPT, AaaPT, and AmaPT.\nFigure 3R2. Activity tests of other PTs variants. a, Activity tests of AfaPT-WT and\nvariants. b, Activity tests of AaaPT-WT and variants. c, Activity tests of AmaPT-WT\nand variants. Whole cells harboring AtaPT mutants were resuspended in 50 mM Tris-\nHCl buffer (pH 7.5) containing 5 mM CaCl2 and 6% (v/v) DMSO, with the cell density\nadjusted to OD = 80. Substrate 1a and DMAPP/GPP were added at final\n600\nconcentrations of 2 mM and 4 mM, respectively, in a total reaction volume of 300 μL.\nBioconversion reactions were performed at 37 °C with shaking at 1200 rpm for 24 h.\nRelative activities are expressed as mean ± SD from three independent biological\nreplicates (n = 3).\n2. The current data appear insufficient to robustly support the proposed 'aromatic cage'\nconcept. This is primarily because the experimental evidence involves too few amino\nacid residues. Crucially, the manuscript lacks experimental data for mutations of P324\nto other aromatic amino acids, such as phenylalanine (P324F). The observed changes in\nactivity might be primarily attributed to enhanced interactions between the introduced\nhistidine residue and the substrate, rather than definitively demonstrating the formation\nor function of an aromatic cage.\nThank you for your insightful comments. We constructed and evaluated additional\nvariants at position 324, including P324F, P324Y, and P324W, all of which exhibited\nlower activity and selectivity than P324H. Introducing these substitutions into the M7\nand M8 further diminished catalytic performance. To assess the influence of side-chain\nvolume, we also generated the P324L and P324M variants, both of which similarly\nresulted in reduced activity. Moreover, reverting H324 back to proline again weakened\nthe reaction (Figure 3R3 and Supplementary Figure 16). Collectively, these results\ndemonstrate that histidine provides the optimal balance of polarity and steric properties\nat this position, thereby supporting efficient catalysis (Page 6).\nThe aromatic cage concept itself is well established in enzymology (Biochemistry 60,\n259-273, 2021; PLoS Comput. Biol. 18, e1009871, 2022; Chem. Biol. Drug Des. 89,\n141-151, 2016), where it typically serves to anchor and orient the substrate in a reactive\nconformation. Our findings extend this concept by demonstrating that, beyond aromatic\nstacking, residue volume and polarity can also critically modulate catalytic\nperformance, broadening the mechanistic understanding of aromatic cage-mediated\nselectivity.\nFigure 3R3. Relevant exploration of residue 324. Whole cells harboring AtaPT mutants\nwere resuspended in 50 mM Tris-HCl buffer (pH 7.5) containing 5 mM CaCl2 and 6%\n(v/v) DMSO, and the cell density was adjusted to OD = 80. Substrate 1a and DMAPP\n600\nor GPP were added to the reaction mixture at final concentrations of 2 mM\nand 4 mM, respectively, in a total volume of 300 μL. Bioconversion reactions were\nincubated at 37 °C with shaking at 1200 rpm for 24 h. Relative activities are reported as\nmean ± SD of three independent biological replicates (n = 3).\n3. In the engineering of donor plasticity across multiple prenyltransferases, bulky\nresidues are commonly substituted with smaller ones to expand the donor-binding\ncavity, thereby accommodating larger donor molecules. Consequently, the authors'\nexplanation for the G326W mutant's inactivity toward GPP—compared to previous\nstudies—does not present particularly novel insights.\nThank you for your comments. We agree that, in general, substituting bulkier residues with\nsmaller ones can expand the donor-binding pocket and enhance activity toward larger\ndonor molecules. However, identifying and precisely targeting the key “hot spots” that\nregulate donor accommodation remains challenging. In our study, both regioselectivity and\ndonor specificity were considered simultaneously. While the G326W mutation indeed\nblocked GPP acceptance, substituting this site with other bulky residues led to a drastic\nloss in both activity and regioselectivity. When analyzing M7, M8, and G326W together,\nour results indicate that the binding mode of the acceptor substrate also critically influences\ndonor recognition. These findings underscore the importance of coordinated donor–\nacceptor interactions in modulating enzyme plasticity rather than relying solely on cavity\nexpansion.\n4. The use of DMSPP as an alternative donor to DMAPP for crystallographic studies of\nprenyltransferases has become an established methodology. Therefore, we recommend\nthat the authors reattempt cocrystal acquisition using DMSPP to better elucidate the\nconformational arrangements of both donor- and acceptor-proximal residues.\nThank you very much for your suggestions. In the revised manuscript, we have included\nthe crystal structure of M8 in complex with DMSPP and 1a (Figure 3R4 and Figure 3).\nFigure 3R4. The crystal structure of M8 in complex with 1a and DMSPP\n5. The manuscript predominantly employs percentage yield for quantitative description of\nthe data, yet fails to specify the calculation methodology in the Methods section.\nThank you for pointing this out. To clarify, we have added detailed descriptions in the\nMethods section and updated the relevant figure captions in the revised manuscript.\nSpecifically, both the model and cascade reactions were quantified by integrating the\nHPLC peak areas of the products and applying the corresponding calibration curves. For\nsubstrate scope experiments, conversion rates were determined from the ratio of product\npeak area to substrate peak area in HPLC. These additions ensure that the quantitative\nmethodology is now clearly defined and fully reproducible.\n6. The text contains numerous discrepancies between the descriptions and referenced\nfigures/tables, or lacks adequate graphical support. For example – When overlapping\nthe M7-1a model with the structure of M8, the closest distance between the phenolic\nring of 1a and the C175M side chain is only 3 Å, suggesting a potential steric clash. A\nthorough revision of the entire manuscript is strongly recommended to address these\nissues.\nThank you very much for your reminder. We have added a new figure in the\nSupplementary Information showing the distance between the phenolic ring of 1a and\nthe C175M side chain (Figure R5; Supplementary Fig. 19). In addition, following your\nsuggestion, we carefully reviewed the entire manuscript to identify and address any\nsimilar issues.\nFigure 3R5. The close distance distance between the phenolic ring of 1a and the C175M\nside chain in M8 when in the M7 docking model is overlapped with M8.\n7. I have a few other minor considerations for revision.\nTo date, exactly how many aPTs have been experimentally validated to accept KAE as\na substrate—six or seven?\nThank you for pointing out the error. We have carefully verified and fixed the “six” in\nthe text to \"seven\". We apologize for any confusion caused and appreciate your attention\nto detail.\nThe manuscript lacks structural characterization of compound 4a, and critically, the\nsupporting materials fail to provide standardized NMR data listings, particularly for the\nnew compounds.\nThank you very much for your valuable suggestions. Previously, we did not synthesize or\ncharacterize 4a because no suitable mutant exhibited both high activity and selectivity\ntoward this product. However, we agree that its structural characterization is essential.\nAccordingly, we prepared 4a via enlarged-scale bioconversion and performed NMR\nanalysis, which confirms that 4a is a C-6 prenylated derivative of 1a (Figures R6 and R7;\nSupplementary Figures 60 and 61). In addition, we have compiled and standardized the\nNMR data for all synthesized products reported in this study. These data have been\nincorporated into the revised Supplementary Information under the section\n“Characterization of Prenylated Products” (page S136).\nFigure 3R6. 1H NMR spectrum of 4a (400 MHz, DMSO-d ).\nThank you very much for your reminder. We have added supplementation in the\nrelevant sections to include the required information in the revised manuscript (Page 6,\nSupplementary Table 10). We apologize for any confusion caused and appreciate your\nattention to detail.\nHow was the reaction time determined for measuring the Michaelis-Menten kinetic\nparameters?\nThank you for your comments. Initially, we referred to previously reported procedures for\nsimilar prenyltransferases (J. Agric. Food Chem. 72, 8018-8026, 2024; J. Agric. Food\nChem. 71, 4675-4682, 2023; Phytochemistry 224, 114149, 2024). Based on these\nreferences, we conducted preliminary time-course experiments to evaluate product\nformation rates under our assay conditions. To ensure the accuracy of the kinetic\nmeasurements, we further optimized enzyme concentration and reaction time according to\nthe catalytic activity of each variant, thereby determining the optimal time point at which\nthe reaction remained within the initial velocity range for Michaelis–Menten kinetic\nanalysis.\nPoint-by-point Responses\nReviewer #1 (Remarks to the Author):\nThe authors have well addressed the concerns raised by this reviewer, and the present\nmanuscript can be accepted. Congratulations!\nWe sincerely thank the reviewer for the positive evaluation of our work and for\nacknowledging that all concerns have been satisfactorily addressed. We appreciate your\nconstructive input and encouragement.\nReviewer #2 (Remarks to the Author):\nThe authors have sufficiently addressed my concerns in their current revision.\nWe thank the reviewer for the positive feedback and careful reassessment of the revised\nmanuscript. We are grateful for your professional review, constructive comments, and\nvaluable suggestions, which have significantly improved the quality of our work.\nReviewer #3 (Remarks to the Author):\nThe revised manuscript has effectively addressed our concerns with modifications and\nrefinements, making the current version suitable for publication.\nWe sincerely thank the reviewer for the thorough evaluation and positive assessment of\nthe revised manuscript. Your insightful comments and suggestions were instrumental in\nrefining the manuscript and improving its overall quality."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have well addressed the concerns raised by this reviewer, and the present manuscript can be accepted. Congratulations!", "reviewer_2": "The authors have sufficiently addressed my concerns in their current revision.", "reviewer_3": "The revised manuscript has effectively addressed our concerns with modifications and refinements, making the current version suitable for publication."}, "author_response": ""}]

Decision letter