Metabolic engineering and late-stage functionalization expand the chemical space of the antimalarial premarineosin A

McBride, Christina M.; McCauley, Morgan; Harris, Natalia R.; Amin, Sahar; Curtis, Brian J.; Verhey-Henke, Linnea; Teklemichael, Awet A.; Oliphant, Erin N.; Dranchak, Patricia; Lev, Katherine L.; Qu, Fengrui; Snodgrass, Harrison M.; Lewis, Jared C.; Inglese, James; Su, Xin-Zhuan; Pereira, Filipa; Sherman, David H.

doi:10.1038/s42004-025-01779-6

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Published: 05 December 2025

Metabolic engineering and late-stage functionalization expand the chemical space of the antimalarial premarineosin A

Communications Chemistry volume 8, Article number: 391 (2025) Cite this article

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Abstract

Diversification of structurally complex natural products remains a key challenge in the discovery of next-generation therapeutics. Premarineosin A, a potent and selective antimalarial natural product, is a promising yet underexplored scaffold due to its limited availability and synthetic complexity. In this work, we overcome both barriers by coupling metabolic engineering with late-stage derivatization, enabling the first systematic exploration of the premarineosin A scaffold. Rational engineering of Streptomyces eitanensis, encoding a premarineosin A biosynthetic gene cluster, increased titers over 170-fold. Sustainable production of (−)-premarineosin A enabled a unique semi-synthetic and biocatalytic derivatization campaign. In this structure-activity relationship study of premarineosin A, we accessed a suite of analogs, including a C12-brominated derivative with nanomolar potency (EC₅₀ < 5 nM). This work establishes (−)-premarineosin A as a tractable and evolvable antimalarial scaffold, demonstrating how chemical biology approaches can unlock structural and pharmacological space from complex microbial metabolites.

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Introduction

Malaria is a life-threatening disease that poses severe health risks for nearly half of the global population. In 2023 alone, malaria was responsible for an estimated 263 million cases and 597,000 deaths¹. As the causative Plasmodium parasites are becoming increasingly drug resistant, developing new antimalarials that match or exceed the efficacy of current therapeutics is paramount^2,3,4. The prodiginine class of microbial natural products, distinguished by a core tripyrrole moiety formed from the condensation of 4-methoxy-2,2′-bipyrrole-5-carboxaldehyde (MBC) with a third, functionalized pyrrole⁵, has been explored for potent antiparasitic activity and selectivity^{6,7,8,9,10,11}. In addition to their antimalarial activity^6,7,8,9,12, these metabolites have exhibited remarkable anticancer^13,14,15, antibacterial^16,17, antifungal¹⁷, and immunosuppressant¹⁴ properties. Cyclic prodiginines are particularly notable, as their constrained conformations encourage the binding of charged ions, leading to enhanced bioactivity compared to their linear counterparts^15,18,19. For instance, marineosin A (1), a cyclic prodiginine isolated from Streptomyces sp. CNQ-617, incorporates a unique spiroaminal structure that confers potent cytotoxicity²⁰. In a key late-stage step in marineosin A (1) biosynthesis, the linear 23-hydroxyundecylprodiginine (23-HUP, 2) undergoes an oxidative bicyclization at the C8-C9 double bond catalyzed by a Rieske oxygenase (MarG), yielding premarineosin A (3)¹². Finally, premarineosin A (3) is converted to marineosin A (1) by a proposed MarA-catalyzed reduction of the C6-C7 double bond¹². Premarineosin A (3) is of particular interest, as it exhibits single-digit nanomolar antimalarial activity in vitro and is notably less cytotoxic than marineosin A^12,20.

Derivatization is an effective strategy for enhancing the antiplasmodial activity and selectivity of prodiginine natural products^{6,7,8,9,10,11}, as functionalization can improve potency, reduce toxicity, optimize metabolism, or increase oral bioavailability in comparison to the native scaffold^{21,22,23,24,25}. Most often, total synthesis has been employed to derivatize linear prodiginines, with modifications occurring early in the route through the functionalization of simple precursors⁸. However, because of the synthetic complexity of the spirocyclic core, premarineosin A (3) is significantly less amenable to total synthesis, as evidenced by all prior efforts resulting in low overall yields^26,27. Synthesis of premarineosin A (3) analogs has been attempted via bioconversion of 23-HUP (2), but was limited by low conversion rates⁶. Late-stage functionalization is a promising alternative strategy but can be challenging for natural product scaffolds due to their complex ring structures and diverse functional groups, which can hinder selectivity and reactive site accessibility^28,29. Hence, identifying late-stage diversification approaches that selectively target sites like the highly reactive pyrrolic carbons would enable rapid functionalization of premarineosin A (3).

Securing access to premarineosin A (3) is also essential to our diversification efforts. While heterologous expression of the marineosin A (1) biosynthetic cluster disrupted in marA enabled the isolation of 0.5 mg of premarineosin A (3)¹², no production was observed in cultures >100 mL, limiting further structural characterization, derivatization, and drug development efforts. Metabolic engineering is a proven approach for enhancing the production of natural products^30,31, including prodiginines^{32,33,34,35,36}. In this work, we report a previously unexplored premarineosin A (pma) biosynthetic gene cluster (BGC) in Streptomyces eitanensis and employ metabolic engineering approaches to improve its production. Access to a sustainable source of (−)-premarineosin A (3) enabled us to define its absolute stereochemistry and develop high-yield strategies for its late-stage derivatization via unique semi-synthetic and biocatalytic approaches. The derivatives were investigated for potency and cytotoxicity in vitro, which identified a brominated analog with high activity against drug-sensitive and drug-resistant Plasmodium falciparum parasites.

Results

Genome and metabolome analysis identified a (−)-premarineosin A biosynthetic gene cluster in S. eitanensis

Recently, we identified the production of 23-HUP (2) in the environmental isolate Streptomyces eitanensis³⁰. Analysis of its genome revealed a cluster with 90% similarity to the canonical marineosin A BGC (MIBiG: BGC0000091)^12,37. This newly identified cluster lacks genes previously shown to encode the putative acyltransferase (MarE) and the reductase (MarA) predicted to generate the 1-pyrroline B-ring in marineosin A (1) biosynthesis (Fig. 1a)¹². However, it does encode the putative Rieske oxygenase PmaG, a homolog (85% similarity, 77% identity) of the MarG enzyme that catalyzes bicyclization of 23-HUP (2) into the marineosin precursor, premarineosin A (3) (Fig. 1b, c). Closer inspection of the S. eitanensis metabolome detected a compound with the same molecular weight and fragmentation pattern as premarineosin A (3) (Supplementary Fig. 1). As marineosin A (1) production was not observed in this strain, we reasoned that the identified BGC (pma BGC) likely encodes premarineosin A (3) as the terminal product, suggesting that S. eitanensis could serve as an ideal strain for optimizing its large-scale production.

Fig. 1: Metabolic engineering of S. eitanensis enhanced (−)-premarineosin A (3) production. — **Fig. 1: Metabolic engineering of *S. eitanensis* enhanced (−)-premarineosin A (3) production.**

Overexpression of the pma cluster-situated regulator (pmaD) and Rieske oxygenase (pmaG) improved (−)-premarineosin A production in S. eitanensis

Trace amounts of (−)-premarineosin A (3) were detected in wild-type S. eitanensis (Fig. 1d, Supplementary Methods 1 and Supplementary Fig. 1). Annotation of the pma BGC (Supplementary Table 1) revealed PmaD, a homolog (40% protein identity, 52% similarity) of the RedD transcriptional activator from the undecylprodigiosin (UDP) pathway in Streptomyces coelicolor A3(2)^38,39,40, suggesting that PmaD may similarly regulate (−)-premarineosin A (3) production in S. eitanensis. Thus, we engineered S. eitanensis for enhanced (−)-premarineosin A (3) production using a regulatory gene expression strategy³⁰. The pmaD gene, amplified from the S. eitanensis genome, was assembled in pSET152k under the strong constitutive promoter kasOp*, yielding pSET152k-pmaD. This plasmid was integrated at the ΦC31 actinophage integrase attB site in the S. eitanensis genome, generating the pmaD overexpressing strain FP10017 (Supplementary Fig. 2). To ensure that the producing phenotype was directly attributed to overexpression of pmaD, we integrated the empty vector into the S. eitanensis genome, generating the strain FP10016. Plasmid integration at the attB site was confirmed by whole genome sequencing for all engineered S. eitanensis strains (Supplementary Fig. 2). Under non-optimized culture conditions (corn gluten meal, GICYE), the FP10017 strain produced significantly more (−)-premarineosin A (3) (14.7 ± 1.4 mg L⁻¹) than the vector control FP10016 strain (1.0 ± 0.4 mg L⁻¹) and the wild-type strain (below the limit of quantification) (Fig. 1d).

In the last step of (−)-premarineosin A (3) biosynthesis, the putative Rieske oxygenase PmaG catalyzes the bicyclization of 23-HUP (2)^6,12. As accumulation of 23-HUP (2) was observed in the engineered FP10017 strain (Fig. 1d), we designed pSET152k-pmaDG for constitutive overexpression of pmaG and pmaD in S. eitanensis. This plasmid was integrated into S. eitanensis wild-type, yielding the FP10018 strain. Constitutive overexpression of pmaG and pmaD significantly enhanced (−)-premarineosin A (3) production (23.3 ± 0.5 mg L⁻¹) in FP10018, a 1.6-fold increase compared to the overexpression of pmaD alone. The metabolic shift toward the production of (−)-premarineosin A (3) is also reflected by the difference in color between the pmaG and pmaD overexpressing strain FP10018 (red) and the FP10016 (green) strain (Fig. 1e).

In addition to gene overexpression strategies, optimization of media composition and culture conditions can enhance secondary metabolite production in Streptomyces^30,36,41,42. Cultivation of the engineered FP10018 strain in soybean meal medium (GISYE) significantly (p < 0.0001) enhanced (−)-premarineosin A (3) production by 1.39-fold compared to cultivation in GICYE (Fig. 1d and Supplementary Fig. 3). Target compound production was assessed throughout cultivation, and while maximum titers of (−)-premarineosin A (3) were obtained around day six, the titers of 23-HUP (2) were lowest on day seven (Fig. 1f and Supplementary Fig. 4). Both acetone and methanol were investigated as extraction solvents, with acetone showing, on average, higher (−)-premarineosin A (3) and 23-HUP (2) recovery (Fig. 1d and Supplementary Fig. 5). Under optimal conditions (GISYE at 22 °C for seven days followed by extraction with acetone), (−)-premarineosin A (3) production increased approximately 174-fold in the engineered strains FP10017 and FP10018 (34.8 ± 1.6 mg L⁻¹ vs. 33.7 ± 5.1 mg L⁻¹, respectively) when compared to wild-type (0.2 ± 0.1 mg L⁻¹) (Fig. 1d). In GISYE, the vector control strain FP10016 produced 15-fold more (−)-premarineosin A (3) (3.0 ± 2.1 mg L⁻¹) compared to wild-type. The FP10018 strain produces nearly twice (p < 0.05) as much (−)-premarineosin A (3) per gram of dry cell weight (5.1 mg g⁻¹ DCW) as strain FP10017 (2.6 mg g⁻¹ DCW) (Fig. 1f). Production of 23-HUP (2) was also increased in the engineered strains, with significantly (p < 0.0005) increased production in the FP10018 strain compared to the wild-type when cultured in GISYE (Fig. 1d).

Efforts to maximize (−)-premarineosin A (3) isolation and purification from the engineered strains FP10017 and FP10018 yielded 12.9 ± 1.2 mg L⁻¹ and 11.8 ± 2.9 mg L⁻¹, respectively, with >95% purity (Fig. 1g and Supplementary Fig. 6). Structural characterization was conducted using NMR spectroscopy (Supplementary Data 1 and Supplementary Methods 2)¹², X-ray crystallography (Fig. 1c, Supplementary Note 1 and Supplementary Data 2), and polarimetry. While the relative stereochemistry of (−)-premarineosin A (3) confirms previously described reports^6,20, our X-ray crystal data (Fig. 1c) revealed that its absolute stereochemistry did not agree with the premarineosin A (3) structure obtained by total synthesis^26,27. As the optical rotation value for the compound isolated in this work was [α]_D²⁴ (c 0.4758 in MeOH) = −104.0 ± 0.1°, we refer to the compound as (−)-premarineosin A (3). To address the stereochemical discrepancy, marineosin A (1) was isolated from the original producer strain, Streptomyces sp. CNQ-617²⁰. The structural identity of the isolated marineosin A (1) was confirmed by ¹H and ¹³C NMR (Supplementary Data 1 and Supplementary Methods 2) and by optical rotation. The optical rotation value of [α]_D²⁵ = −105.8 ± 2.1° (c 0.2525, MeOH) indicates that the isolated marineosin A (1), like (−)-premarineosin A (3), is levorotatory.

Isolated compound (−)-premarineosin A (3) was tested against the chloroquine (CQ)-sensitive 3D7 and CQ-resistant Dd2 P. falciparum parasites in an in vitro assay (Fig. 2). The compound was concomitantly evaluated for cytotoxicity against the HEK293 embryonic kidney and MOLT4 T-cell leukemia cell lines. (−)-Premarineosin A (3) exhibits nanomolar potency against the 3D7 (EC₅₀ = 4.0 ± 0.1 nM) and Dd2 (EC₅₀ = 1.0 ± 0.7 nM) P. falciparum parasites with minimal toxicity to mammalian cells (Fig. 2). In comparison, (−)-premarineosin A (3) is over 10- (3D7) and 350-fold (Dd2) more potent than chloroquine and is 6- (3D7) and 8-fold (Dd2) more potent than artesunate (Fig. 2).

Fig. 2: In vitro bioactivities of (−)-premarineosin A (3) and derivatives (4–12) show selectivity against Plasmodium falciparum. — Fig. 2: In vitro bioactivities of (−)-premarineosin A (3) and derivatives (4–12) show selectivity against *Plasmodium falciparum.*

Acid-catalyzed electrophilic aromatic substitution enabled late-stage derivatization of (−)-premarineosin A (3)

With access to sufficient quantities of (−)-premarineosin A (3), we pursued a semi-synthetic approach to conduct the first structure-activity relationship analysis of this scaffold. Dimerization of natural products is a known approach to increase their structural complexity and biological activity^43,44. Previous synthetic studies established that simple 2,5-dimethylpyrroles are susceptible to acid-catalyzed condensation with ketones, such as acetone, to form dimerized substrates at the 3-position in an electrophilic aromatic substitution-like reaction⁴⁵. To our knowledge, this synthetic approach has not been previously applied to any natural product. Thus, we reasoned that exploration of the breadth and flexibility of this chemistry may enable access to previously untapped chemical space. Applying similar methodologies, we found that (−)-premarineosin A (3) dimerizes under acidic conditions with both ketones and aldehydes at the C12 position of its C-ring, forming dimers with a gem-dimethyl-bridge (4), a methylene-bridge (5), and a trifluoromethyl-bridge (6) (Fig. 3a, top). A crystal structure of gem-dimethyl-bridged premarineosin A (4) was obtained to confirm its unique structure, stereochemistry, and absolute configuration (Fig. 3b, Supplementary Note 2 and Supplementary Data 3). Functionalization of (−)-premarineosin A (3) with trifluoroacetone also yielded a monomeric derivative (12-trifluoropropanol premarineosin A, dr 1:0.65 (7)), likely due to early reaction termination before complete substrate consumption (Supplementary Fig. 7). Premature termination using acetone or formaldehyde as the electrophile failed to generate monomeric intermediates, suggesting that isolating these monomeric (−)-premarineosin A (3) derivatives under electrophilic aromatic substitution conditions occurs only when using electron-withdrawing groups⁴⁵. To synthesize other monomeric derivatives of (−)-premarineosin A (3), we used acid chlorides as the electrophile for Friedel-Crafts acylation to form 12-formyl premarineosin A (8) and 12-acetyl premarineosin A (9). Further reduction of 12-formyl premarineosin A (8) with sodium borohydride provided the desired primary alcohol, forming 12-hydroxymethyl-premarineosin A (10) (Fig. 3a, bottom).

**Fig. 3: Semi-synthetic electrophilic aromatic substitution at the C12 position facilitated robust derivatization of the (−)-premarineosin A (3) scaffold.**

The antiparasitic activity and cytotoxicity of each semi-synthetic derivative was evaluated in comparison to (−)-premarineosin A (3) (Fig. 2). The monomeric derivatives 7, 8, and 9 display sub-micromolar activity against the 3D7 and Dd2 P. falciparum parasites. Despite the reduced activity compared to (−)-premarineosin A (3), derivatives 7, 8, and 9 perform better than chloroquine (EC₅₀ = 382 ± 22 nM) against Dd2. 12-Trifluoropropanol premarineosin A (7) is the most promising semi-synthetic derivative in terms of selectivity, displaying moderate potency against Dd2 (EC₅₀ = 87 ± 15 nM) with limited apparent toxicity to HEK293 cells (EC₅₀ = 4210 ± 481 nM). Compound 5 (EC₅₀ = 548 ± 76 nM) and 10 (EC₅₀ = 704 ± 304 nM) also exhibited sub-micromolar potency against the multidrug-resistant P. falciparum strain yet had limited potency against the drug-sensitive strain. While these synthetic modifications at the C-ring reduced the antimalarial potency of the (−)-premarineosin A (3) scaffold, the analogs broadly exhibit decreased cytotoxicity (Fig. 2).

Biocatalytic and semi-synthetic bromination enabled late-stage C-H functionalization of (−)-premarineosin A (3)

Bromine holds significant value in medicinal chemistry due to its capacity to improve key pharmacokinetic properties, such as membrane permeability, target binding affinity, and metabolic stability—factors crucial for combating Plasmodium species effectively^46,47. Hence, we sought to site-selectively brominate (−)-premarineosin A (3) using biocatalysis. Molecular docking of (−)-premarineosin A (3) with D3, an orphan flavin-dependent halogenase (FDH) from Saccharophagus degradans⁴⁸, revealed strong substrate binding (−8.6 kcal mol⁻¹), with (−)-premarineosin A (3) oriented within the active site in a conformation that suggests the A-ring pyrrole would be most accessible for bromination (Fig. 4a). Indeed, reactions of D3 with (−)-premarineosin A (3) in vitro generated a single brominated derivative, 1-bromo premarineosin A (11), with ~50% and 10% substrate conversion efficiency at small- and large-scale, respectively (Fig. 4b, c).

**Fig. 4: Site-selective biocatalytic and semi-synthetic strategies enabled the late-stage bromination of (−)-premarineosin A (3).**

To evaluate product formation and regioselectivity, we compared this enzyme-catalyzed bromination with a synthetic approach. N-bromosuccinimide (NBS) was selected as the brominating agent because of its proven effectiveness in regioselective electrophilic aromatic bromination⁴⁹. Reaction with NBS at 4 °C in DCM for 2 h nearly consumed (−)-premarineosin A (3), primarily producing a C-ring brominated product (55% isolated yield, 10 mg reaction), 12-bromo premarineosin A (12, Fig. 4b, d).

The antiparasitic and cytotoxic activities of the brominated premarineosin A derivatives were also assessed in vitro. 1-Bromo premarineosin A (11) exhibited sub-micromolar antiparasitic activity, with EC₅₀ values of 473 ± 113 nM (3D7) and 259 ± 24 nM (Dd2). On the other hand, 12-bromo premarineosin A (12) demonstrated high potency against 3D7 (EC₅₀ = 2 nM) and Dd2 (EC₅₀ = 22 ± 11 nM) with low cytotoxicity (Fig. 2). These findings highlight 12-bromo premarineosin A (12) as a lead compound, combining exceptional antimalarial potency and lower cytotoxicity with remarkable selectivity.

Discussion

This work presents expanded efforts that highlight (−)-premarineosin A (3) as a promising scaffold for the development of antimalarial therapeutics. Under optimal conditions, the titer of (−)-premarineosin A (3) in the engineered strain FP10017 was 34.8 ± 1.6 mg L⁻¹, a 174-fold increase compared to S. eitanensis wild-type (0.2 ± 0.1 mg L⁻¹). While the difference in (−)-premarineosin A (3) levels between the engineered FP10017 and FP10018 strains under optimized conditions was not statistically significant, we did observe that FP10018 produces nearly twice as much (−)-premarineosin A (3) per gram of dry cell weight (Fig. 1F). As the accumulation of (−)-premarineosin A (3) and 23-HUP (2) might inhibit cell growth, we reason that further culture optimization can be employed to increase cell viability in the engineered FP10018 strain, enhancing compound titers. Nevertheless, the engineered S. eitanensis FP10017 and FP10018 strains boast the highest production and isolation titers of (−)-premarineosin A (3) reported to date. Interestingly, the strain integrating the empty vector (FP10016) also produced 15-fold more (−)-premarineosin A (3) than the wild-type in soybean media. Increased production of the target compound in the engineered FP10016 strain could be a result of plasmid integration at the ΦC31 integrase (attB) site, which has been shown to alter the availability of malonyl-CoA and acetyl-CoA⁵⁰ and induce a stress response⁵¹, stimulating secondary metabolite production.

Contradictory data regarding the stereochemistry of premarineosin-associated molecules have been reported; thus, we sought to definitively establish its absolute configuration. The stereocenters of the (−)-premarineosin A (3) isolated in this work have the same relative configuration as previously reported structures of premarineosin A (3) from both natural product isolation^12,20 and total synthesis^26,27 studies. However, the crystal structures of (−)-premarineosin A (3) and the gem-dimethyl-bridged premarineosin A (4) support an absolute configuration of 8S, 9R, 21R, 23R, which is enantiomeric compared to the previously reported stereochemistry^6,12,26,27 (Figs. 1c and 3b). Efforts toward the total synthesis of marineosin A (1) by Shi et al.²⁷ and Harran et al.²⁶ followed the absolute stereochemistry proposed by Reynolds et al.^6,12 based on the conclusion that marineosin A (1) is biosynthesized from (23S)-HUP rather than (23R)-HUP. The optical rotation values for marineosin A (1) obtained by total synthesis ([α]_D²⁵ = +62.6° (c 0.16, MeOH) by Shi et al.²⁷, and [α]_D²⁵ = +138.7° (c 0.02, MeOH) by Harran et al.²⁶) oppose the value originally reported by Fenical et al.²⁰ for the natural product isolated from Streptomyces sp. CNQ-617 ([α]_D²⁴ = − 101.7° (c 0.06, MeOH)). Shi et al. noted additional differences in compound properties and suggested that their synthetic compound was not identical to the isolated marineosin A (1) but could instead be an isomer²⁷. Harran et al. instead postulated that the difference could be explained by an accidental interchange of the marineosin A and B optical rotation values reported by Fenical et al., claiming that, like their synthetic product, the original isolated marineosin A (1) could be dextrorotatory²⁶. As the absolute configuration of the (−)-premarineosin A (3) isolated in this study is enantiomeric respective to that reported by Reynolds, Shi, and Harran, we sought to confirm the absolute configuration of isolated marineosin A (1) from Streptomyces sp. CNQ-617. Optical rotation analysis for marineosin A (1) isolated in this work supported the polarity value (−) and was of similar magnitude to the value initially reported by Fenical et al.²⁰. Together, these results suggest that the structure-confirmed dextrorotatory product obtained by total synthesis, (+)-marineosin A, is likely the enantiomer of the confirmed levorotatory marineosin A (1) isomer isolated from Streptomyces sp. CNQ-617. Hence, the natural product marineosin A (1) likely shares the same absolute stereochemistry at C8 (S), C9 (R), C21 (R), and C23 (R) as the (−)-premarineosin A (3) isolated from S. eitanensis.

Engineering in vivo production for sustainable and scalable access to (−)-premarineosin A (3) not only allowed us to resolve this stereochemical ambiguity but also enabled our expansion of the chemical space around this scaffold, culminating in the synthesis of nine analogs. We first focused on the surprising discovery of pyrrole dimerization chemistry to diversify the C12 position of the (−)-premarineosin A (3) C-ring. While electrophilic aromatic substitution has been utilized previously to functionalize the β-position of pyrroles^52,53, to our knowledge this methodology has not been reported with any complex natural product scaffold. This reaction platform revealed an underexplored acid-catalyzed electrophilic aromatic substitution diversification strategy for pyrrolic alkaloids, highlighting (−)-premarineosin A (3) as a synthetically pliable and chemically evolvable scaffold. Of these seven semi-synthetic premarineosin A derivatives (4-10), five show sub-micromolar potency against the chloroquine-resistant P. falciparum Dd2, with three having a lower EC₅₀ than chloroquine (Fig. 2). However, they all have reduced bioactivity compared to (−)-premarineosin A (3), suggesting that the C12 position plays a crucial role in the antimalarial activity of the premarineosin A scaffold (Figs. 2 and 5). Remarkably, 4–10 are overall less cytotoxic than (−)-premarineosin A (3), in agreement with the bioactivity of other C-ring functionalized prodiginines^54,55,56. Hence, functionalizing the C12 position of the C-ring is an encouraging avenue to increase the selectivity of (−)-premarineosin A (3) and improve its therapeutic potential. To explore this trend further, we carried out NBS-mediated bromination of (−)-premarineosin A (3). The resulting 12-bromo premarineosin A (12) derivative was substantially more potent (17-fold) than chloroquine against the chloroquine-resistant P. falciparum Dd2 strain. While 12-bromo premarineosin A (12) did not outperform the outstanding potency and selectivity of (−)-premarineosin A (3) against the Dd2 strain, it was twice as potent as (−)-premarineosin A (3) against the drug-sensitive P. falciparum 3D7 with improved selectivity relative to HEK293 mammalian cells (Figs. 2 and 5). Bromination of the C12-position of (−)-premarineosin A (3) decreased cytotoxicity while maintaining single-digit nanomolar potency against P. falciparum (EC₅₀ < 5 nM), offering a unique opportunity to enhance drug efficacy, stability, and bioavailability^57,58. Given that its potency exceeds both artesunate and chloroquine in the 3D7 strain, these results underscore 12-bromo premarineosin A (12) as a compelling lead compound for antimalarial drug development (Fig. 5).

Fig. 5: Diversifying the (−)-premarineosin A (3) scaffold improved selectivity against Plasmodium falciparum. — Fig. 5: Diversifying the (−)-premarineosin A (3) scaffold improved selectivity against *Plasmodium falciparum.*

As a complement to synthetic chemistry methods, we turned to biocatalysis to probe unexplored regions of the premarineosin A (3) chemical space. To modify (−)-premarineosin A (3) beyond the C12 position, we explored FDHs for late-stage C-H bromination of the A-ring pyrrole, which is beyond the reach of our semi-synthetic approach (Fig. 4). While previous studies have used FDHs to halogenate simple pyrrole-containing precursors in early biosynthetic steps⁵⁹, their application to fully assembled, highly complex natural products has remained largely unexplored. As such, our strategy of using the orphan FDH D3 from S. degradans to selectively brominate (−)-premarineosin A (3) on the minimally substituted (A-ring) pyrrole holds significant promise for expanding the scope of enzymatic halogenation (Fig. 4). This result not only highlights the catalytic flexibility of D3 but also demonstrates the feasibility of using non-native halogenases to derivatize densely functionalized, late-stage scaffolds. As depicted in the AlphaFold-generated structure of D3 docked with (−)-premarineosin A (3) (Fig. 4a), the enzyme’s unusually large and flexible active site pocket appears well-suited to readily accommodate the bulky (−)-premarineosin A (3) scaffold in a conformation conducive for strong substrate binding and regio-selective bromination at the A-ring C1-position. The contrasting regioselectivity of the semi-synthetic and biosynthetic methods motivates further enzyme engineering studies to assess the efficiency and selectivity of C-H halogenation of (−)-premarineosin A (3). Although the resulting 1-bromo premarineosin A (11) derivative showed enhanced potency relative to chloroquine against the chloroquine-resistant P. falciparum Dd2 strain and reduced cytotoxicity compared to (−)-premarineosin A (3) and 12-bromo premarineosin A (12) against both cell lines, it exhibited decreased selectivity and antimalarial activity overall, suggesting the C1-position may play a critical role in the therapeutic efficacy of (−)-premarineosin A (3) (Figs. 2 and 5). Nevertheless, this work shows that even without prior engineering, naturally occurring and previously uncharacterized enzymes can be identified through computational modeling and repurposed to modify complex cyclic natural products. The ability of D3 to accept a bulky, non-native scaffold like (−)-premarineosin A (3) and catalyze regioselective halogenation, despite being predicted to be a tryptophan halogenase, underscores the untapped potential of orphan biocatalysts.

Together, our results establish a tractable and modifiable framework for late-stage functionalization of (−)-premarineosin A (3) and demonstrate an approach for strategic expansion of scaffold-level chemical space using both synthetic and biocatalytic approaches. In addition, the remarkable selectivity of 12-bromo premarineosin A (12) against P. falciparum warrants further investigation into its mechanism of action and in vivo efficacy studies. By combining metabolic engineering, biocatalysis, computational modeling, and semi-synthesis, we define a versatile platform for late-stage functionalization of complex pyrrolic alkaloids and create a roadmap for the systematic exploration of (−)-premarineosin A (3) chemical space in the search for next-generation antimalarial agents.

Methods

Strains and culture conditions

All strains that were designed and/or cultured in this work are listed in Supplementary Table 2. The Streptomyces eitanensis strain is an environmental isolate previously characterized by our groups³⁰. Plasmid assembly, replication, and preservation were performed using high-efficiency Escherichia coli DH5α (NEB). E. coli S17-1 was used as the mobilization host for conjugation with S. eitanensis. All E. coli strains were cultivated in LB medium (10 g L⁻¹ tryptone, 10 g L⁻¹ NaCl, and 5 g L⁻¹ yeast extract in ultrapure water) at 37 °C. The LB media was supplemented with 50 µg mL⁻¹ kanamycin for plasmid maintenance and selection. All Streptomyces strains were cultivated in 2xYT (16 g L⁻¹ tryptone, 10 g L⁻¹ yeast extract, and 5 g L⁻¹ NaCl in ultrapure water) or peanut meal & starch media (10 g L⁻¹ glucose, 30 g L⁻¹ starch, 5 g L⁻¹ bacto peptone, 10 g L⁻¹ peanut meal, 5 g L⁻¹ yeast extract, 2 g L⁻¹ CaCO₃ in ultrapure water) for 3 d at 28 °C for seed culture. Strains were sporulated in OPAH (1 g L⁻¹ oatmeal, 1 g L⁻¹ pharmamedia, 1 g L⁻¹ arabinose, 0.5 g L⁻¹ humic acid, 0.5 mM KH₂PO₄, 0.5 mM CaCl₂, 0.5 mM MgSO₄, 1.9 mg L⁻¹ Na₂-EDTA·2H₂O, 1.4 mg L⁻¹ FeSO₄·7H₂O, 0.2 mg L⁻¹ H₃BO₃, 0.05 mg L⁻¹ MnSO₄·H₂O, 0.01 mg L⁻¹ ZnSO₄·7H₂O, 0.01 mg L⁻¹ Na₂MoO₄·2H₂O, 0.01 mg L⁻¹ CuSO₄, and 0.01 mg L⁻¹ CoCl₂ in ultrapure water). Media was supplemented with 50 µg mL⁻¹ kanamycin (Goldbio, K-120-25) and/or 25 µg mL⁻¹ nalidixic acid (Cayman, 19807) as needed.

Plasmid design and genome editing

All plasmids generated in this study are described in Supplementary Table 3. Primers were designed with guidance from pyDNA⁶⁰ (Supplementary Table 4). The pmaD and pmaG genes were amplified from wild-type S. eitanensis gDNA (Supplementary Table 5), then assembled via Gibson Assembly in the plasmid pSET152k-kasOp* between the BamHI and EcoRI restriction sites (New England Biolabs, R0136S and R0101S), as previously described³⁰. Synthetic ribosomal binding sites⁶¹ were used as overlapping regions for multi-gene vector design. An integrative system, mediated by the attP site of the Streptomyces phage ΦC31⁶², was used for constitutive overexpression in S. eitanensis. Interspecies conjugation was performed to transfer DNA from E. coli S171 to S. eitanensis. Following incubation (12 h, 28 °C), the plates were overlaid with 1 mL sterilized water with 1.25 mg kanamycin and 0.5 mg nalidixic acid. After 3–5 d, recombinants were transferred to OPAH plates with 25 µg mL⁻¹ nalidixic acid and 50 µg mL⁻¹ kanamycin.

Genome extraction, sequencing, and assembly

High-quality genomic DNA was prepared using reagents from the MasterPure Complete DNA and RNA Purification Kit (Lucigen, MC85200). The wild-type and engineered Streptomyces strains were cultivated in 2xYT for 3 d at 28 °C. Cells were pelleted via centrifugation, resuspended in 480 µL EDTA (50 mM, pH 8, sterile filtered) and 120 µL lysozyme (10 mg mL⁻¹, DotScientific DSL38100-10) then incubated for 35 min at 37 °C. Post-incubation, the cells were centrifuged (1 min, 17,000 × g, RT) and resuspended in 200 µL MasterPure Tissue and Cell Lysis Solution treated with 1 µL Proteinase K (Qiagen RP107B-5). Cells were incubated for 15 min at 65 °C, incubated for 2 min at 95 °C, then cooled to RT. 30 µL of RNase A (10 mg mL⁻¹, Sigma Aldrich R5503-1G) was added prior to incubation (1.5 h, 37 °C). The genomic DNA was precipitated following the MasterPure Kit Precipitation of Total Nucleic Acids protocol. Genomic DNA quality was assessed using gel electrophoresis. Bacterial genome sequencing was performed by Plasmidsaurus using Oxford Nanopore Technology with custom analysis and annotation.

Cluster comparison to confirm genome integration

To confirm the correct integration of the designed plasmids into the wild-type S. eitanensis genome, the whole genome of all engineered strains (100% completeness, Supplementary Table 2) was sequenced using long-read Oxford Nanopore Technology (Plasmidsaurus). Correct genome integration of target plasmids was confirmed by extracting the sequence information surrounding the integration site attB using the Integrative Genomics Viewer⁶³ and aligned using clinker⁶⁴ with default settings (Supplementary Fig. 2).

(−)-Premarineosin A (3) production conditions

Spores from OPAH plates were inoculated into 50 mL of peanut meal & starch media (10 g L⁻¹ glucose, 30 g L⁻¹ starch, 5 g L⁻¹ bacto peptone, 10 g L⁻¹ peanut meal, 5 g L⁻¹ yeast extract, 2 g L⁻¹ CaCO₃ in ultrapure water) and incubated (3 d, 28 °C, 200 rpm). 10 mL of this pre-inoculum culture was used to inoculate 1 L of producing media: GICYE (10 g L⁻¹ glucose, 30 g L⁻¹ inulin, 5 g L⁻¹ bacto peptone, 10 g L⁻¹ corn gluten meal, 5 g L⁻¹ yeast extract, 2 g L⁻¹ CaCO₃ in ultrapure water, adjusted to pH 7.0) or GISYE (10 g L⁻¹ glucose, 30 g L⁻¹ inulin, 5 g L⁻¹ bacto peptone, 10 g L⁻¹ soybean meal, 5 g L⁻¹ yeast extract, 2 g L⁻¹ CaCO₃ in ultrapure water, adjusted to pH 7.0) in 2.8 L Fernbach flasks. Cultures were incubated for 7 d at 22 °C and 170 rpm.

Sample preparation and quantification of (−)-premarineosin A (3)

Quantification of (−)-premarineosin A (3) production in 1 L cultures was performed by extracting 1 mL biomass samples from three independent replicates. Target compounds were quantified from cell extracts collected on day seven unless stated otherwise (Fig. 1d and Supplementary Figs. 3, 5). Production over time was assessed from extracts collected every day for 7 d (Fig. 1f and Supplementary Fig. 4). On each collection day, 1 mL was collected from each 1 L culture and the biomass was pelleted via centrifugation (4 min, 20,800 rpm, RT). The cell pellet was resuspended in 1 mL of solvent (methanol or acetone) and 100 μL of glass beads. Supernatant was combined 50:50 with solvent and stored at −20 °C until quantification. Tubes were vortexed (2 h, 4 °C), followed by centrifugation (4 min, 20,800 rpm, RT). The supernatant was retained as the extract and was stored at −20 °C until quantification. Quantification of (−)-premarineosin A (3) production was performed using analytical HPLC (Shimadzu) equipped with a PDA detector and analyzed with a Phenyl-Hexyl column (Luna 5 μM Phenyl-Hexyl 100 Å, LC Column 250 × 4.6 mm, heated to 40 °C). Water + 0.1% formic acid (A)/acetonitrile + 0.1% formic acid (B) (10% to 100% B) was used for the mobile phase at 2 ml min⁻¹. Calibration curves were prepared with known concentrations of pure (−)-premarineosin A (3) and 23-HUP (2). Production titers were determined by comparing the calibration curve and sample peak areas (AUC) at 346 nm (3) and 520 nm (2). These compounds were only detected in culture supernatants at trace levels for all tested strains. Therefore, quantifications reported in this document correspond to their intracellular concentrations. Statistical analysis and visualization were performed with GraphPad Prism v. 10.4.2., with p-values calculated as the result of a two-tailed T-test.

Biomass quantification

Daily 1 mL samples were collected from each of the three independent replicate cultures and vacuum filtered on a pre-dried membrane filter. The retained cells were dried in a microwave oven for 1.5 min⁶⁵. As S. eitanensis is known to form small, spherical pellets in liquid culture, sampling was performed using sterile wide-bore pipette tips and serological pipettes.

Isolation and purification of (−)-premarineosin A (3) and 23-HUP (2)

The total biomass (from 1 L culture) was separated via vacuum filtration. Cell pellets were broken by coating the cells with 750 mL of 100% acetone and then shaking overnight. The acetone was obtained by vacuum filtration, concentrated, and liquid-liquid extracted with ethyl acetate. The dried organic layer was concentrated with silica to be dry loaded for normal phase purification on a Biotage Isolera flash column system. Initial rounds of purification utilized a 40 g silica column with an ethyl acetate/hexane gradient (15–100%) with 1% acetic acid; (−)-premarineosin A (3) eluted at 34–50% and 23-HUP (2) eluted at 20–34%. (−)-Premarineosin A (3) was further purified on a 40 g silica column with a 3% ammonia (7 N in methanol) in DCM solution / hexane gradient (15–100%); (−)-premarineosin A (3) eluted at 20–30%. 23-HUP (2) fraction(s) were further purified using preparative thin-layer chromatography with a 3% ammonia (7 N in methanol) in DCM mobile phase. Purity assessment of isolated compounds was performed using HPLC (Supplementary Fig. 6), NMR (Supplementary Data 1), and LC-MS/MS (Supplementary Fig. 6). All samples for bioactivity testing were >95% pure. Optical rotations were obtained using a Jasco P2000 polarimeter with a 100 mm cell.

Marineosin A (1) production, isolation, and purification conditions

Streptomyces sp. CNQ-617 spores from modified A1 plates²⁰ (10 g L⁻¹ starch, 4 g L⁻¹ yeast extract, 2 g L⁻¹ peptone, 36 g L⁻¹ Instant Ocean) were inoculated into 50 mL of peanut meal & starch media and incubated (3 d, 28 °C, 200 rpm). 10 mL of this pre-inoculum culture was used to inoculate 1 L of GICYE producing media in 2.8 L Fernbach flasks. Cultures were incubated for 7 d at 22 °C and 170 rpm. The total biomass (from 1 L culture) was separated via centrifugation. Cell pellets were broken by coating the cells with 750 mL of 100% acetone and then shaking overnight. The acetone extract was obtained by vacuum filtration, concentrated, and liquid-liquid extracted with ethyl acetate. The dried organic layer was concentrated with silica to be dry loaded for normal phase purification on a Biotage Isolera flash column system, utilizing a 40 g silica column with an ethyl acetate/hexane gradient (15–100%) with 1% acetic acid; marineosin A (1) eluted at 34–50%. Marineosin A (1) fraction(s) were further purified using preparative thin-layer chromatography with a 3% ammonia (7 N in methanol) in DCM mobile phase; marineosin A (1) eluted as a strong UV-active band near the top of the plate. Purity assessment of isolated compounds was performed using HPLC, NMR (Supplementary Data 1), and LC-MS/MS. Optical rotations were obtained using a Jasco P2000 polarimeter with a 100 mm cell.

(−)-Premarineosin A (3)

¹H NMR (600 MHz, acetone-D₆) δ 13.67 (br s, 1H), 12.06 (br s, 1H), 9.82 (br s, 1H), 7.67–7.65 (m, 1H), 7.49–7.44 (m, 1H), 6.54–6.50 (m, 1H), 6.29–6.26 (m, 1H), 5.72 (t, J = 2.9 Hz, 3H), 5.52 (t, J = 2.9 Hz, 1H), 4.46–4.38 (m, 1H), 4.15 (s, 3H), 3.03 (d, J = 12.5 Hz, 1H), 2.69 (dp, J = 13.4, 4.9 Hz, 1H), 2.39 (dt, J = 14.8, 4.4 Hz, 1H), 2.28 (ddd, J = 14.7, 11.7, 3.3 Hz, 1H), 1.95 (dt, J = 13.8, 4.2 Hz, 1H), 1.92–1.83 (m, 2H), 1.72 (ddd, J = 14.5, 10.1, 5.1 Hz, 1H), 1.45 (d, J = 6.8 Hz, 3H), 1.43–1.36 (m, 4H), 1.36–1.30 (m, 1H), 1.25–1.17 (m, 1H), 1.11–1.01 (m, 2H), 0.89–0.77 (m, 2H), 0.60–0.51 (m, 1H). ¹³C NMR (151 MHz, acetone-D₆) δ 181.77, 165.95, 163.25 (q, J = 35.36 Hz), 135.05, 134.18, 127.55, 125.09, 121.64, 117.74 (q, J = 292.0 Hz), 114.62, 110.64, 105.99, 97.24, 93.84, 71.23, 60.92, 46.32, 38.27, 33.56, 28.70, 28.19, 27.26, 25.87, 25.23, 25.18, 24.82, 20.92. LC-MS: calculated for C₂₅H₃₃N₃O₂ [M+H⁺] 408.2651 m/z; found 408.2673 m/z.

Acid-catalyzed electrophilic aromatic substitution-based derivatization general procedure

(−)-Premarineosin A (3) was dissolved in acetonitrile (MeCN) before trifluoroacetic acid (TFA) and a carbonyl-containing substrate was added at RT. The reaction was run until TLC (50% ethyl acetate in hexanes with 1% acetic acid) showed consumption of starting material (15 min–6 h). After completion, the reaction was diluted with ethyl acetate and washed with 1 M NaOH to quench the TFA. The aqueous layer was further extracted with ethyl acetate (×3) and organics were combined, dried over sodium sulfate, filtered, and concentrated. The material was purified through a combination of acidic and/or basic conditions. Acidic conditions: 50% ethyl acetate (with 1% acetic acid) and hexanes (with 1% acetic acid). Basic conditions: 20–50% of a 3% ammonia (7 N in methanol) in DCM solution in hexane gradient. See Supplementary Method 3 for detailed synthetic methods; see Supplementary Data 1 for all NMR spectra.

D3 docking study

The amino acid sequence of the flavin-dependent halogenase D3 (UniProt Accession Number: Q21N77) was used as input for AlphaFold2 (version 2.2.0) to generate a predicted three-dimensional structure of D3. The default parameters and model configuration provided by the AlphaFold2 pipeline were utilized, with a focus on the most confident prediction based on the pLDDT score. Five models were generated, and the one with the highest overall pLDDT score was selected for subsequent studies. The substrate, (−)-premarineosin A (3), was drawn and minimized using ChemDraw (version 21.0) and Chem3D (version 21.0). The minimized structure was exported in PDB format and converted to PDBQT format using AutoDockTools. Protonation states of (−)-premarineosin A (3) were assigned at pH 7.4, and torsional bonds were defined to enable flexible docking. Docking simulations were performed using AutoDock Vina (version 1.1.2). The AlphaFold-predicted D3 structure was prepared for docking by adding polar hydrogens, assigning Kollman charges, and converting the files to PDBQT format using AutoDockTools. The putative binding site was identified using structural alignment with the crystal structure of the D3 homolog, RebH (PDB ID: 2OA1)⁶⁶ and manual inspection of conserved active site residues. A grid box was defined to encompass the putative binding pocket with dimensions of 20 × 20 × 20 Å, centered on the active site. Docking simulations were run with an exhaustiveness parameter of 8 to ensure thorough sampling of conformational space. The docking scores were recorded in kcal mol⁻¹, and the best-ranked poses were selected based on binding energy and alignment with the expected binding mode. The top docking poses were visualized using PyMOL to assess binding orientation and interactions with active site residues. Comparative analyzes of the docking poses were conducted to infer similarities and differences in substrate binding.

Expression and purification of D3 halogenase

For large-scale D3 halogenase production, 10 mL culture tubes containing 5 mL LB with kanamycin were inoculated with BL-21(λDE3) E. coli cells harboring pET28b containing the D3 halogenase gene⁶⁷. The cultures were incubated overnight at 37 °C, 250 rpm. The next day, 750 mL TB supplemented with kanamycin was inoculated with the entire overnight culture. The inoculated expression cultures were incubated at 37 °C, 225 rpm until OD600 was between 0.6 and 0.8. The incubator was allowed to cool to 30 °C, and gene expression was induced with 100 µM IPTG, and the expression culture was incubated for 20 h. Cells were harvested by centrifugation at (6000 × g, 4 °C, 15 min), and the cell pellet was stored at −80 °C until purification of the protein. Cells were lysed via sonication with a total processing time of 5 min with 1 min on/off cycles. The cell lysate was centrifugated (60,000 × g, 25 min), and the resulting clarified lysate was transferred to a fresh 50 mL centrifuge tube and added to pre-equilibrated Ni-NTA (equilibration buffer: 20 mM phosphate, 300 mM NaCl, 10 mM imidazole pH 7.4). Clarified lysate was incubated with resin for ~30 min, at which point it was transferred to uncapped spin columns, and the lysate was allowed to flow through. The resin was washed with at least 5 CV wash buffer (20 mM phosphate, 300 mM NaCl, 25 mM imidazole pH 7.4), at which point the spin column was transferred to a new centrifuge tube, and the resin was washed with elution buffer (20 mM phosphate, 300 mM NaCl, 250 mM imidazole, pH 7.4). Eluted protein was concentrated via diafiltration using Amicon spin filters Ultra 30 K MWCO spin filters and the buffer was exchanged for storage buffer (25 mM HEPES and 10% glycerol, pH 7.4). For long-term storage, proteins were immediately frozen in liquid nitrogen and stored at −80 °C until use.

Expression and purification of HpaC flavin reductase

For large-scale HpaC production, 10 mL culture tubes containing 5 mL LB with ampicillin were inoculated with BL-21(λDE3) pRare E. coli cells harboring pET21a containing the HpaC flavin reductase gene. The HpaC flavin reductase (phaC) plasmid was obtained from Prof. David Ballou (University of Michigan). The cultures were incubated overnight at 37 °C, 250 rpm. The next day, 750 mL TB with ampicillin was inoculated with the entire overnight culture. The inoculated expression cultures were incubated at 37 °C, 225 rpm until OD600 was between 0.6 and 0.8. The incubator was allowed to cool to 20 °C, and gene expression was induced with 100 µM IPTG, and the expression culture was incubated for 20 h. Cells were harvested by centrifugation (6000 × g, 4 °C, 15 min) and the cell pellet was stored at −80 °C until purification of the protein. Cells were lysed via sonication with a total processing time of 5 min with 1 min on/off cycles. The cell lysate was centrifugated (60,000 × g, 25 min), and the resulting clarified lysate was transferred to a fresh 50 mL centrifuge tube and added to pre-equilibrated Ni-NTA (equilibration buffer: 20 mM HEPES, 300 mM NaCl, 50 μM FAD, 10% glycerol, pH 7.4). Clarified lysate was incubated with resin for ~30 min, at which point it was transferred to uncapped spin columns, and the lysate was allowed to flow through. The resin was washed with at least 5 CV wash buffer (20 mM HEPES, 300 mM NaCl, 50 mM imidazole, 10% glycerol, pH 7.4), at which point the spin column was transferred to a new centrifuge tube, and the resin was washed with elution buffer (20 mM HEPES, 300 mM NaCl, 500 mM imidazole, 10% glycerol, pH 7.4). Eluted protein was concentrated via diafiltration using Amicon spin filters Ultra 30 K MWCO spin filters and the buffer was exchanged for storage buffer (20 mM HEPES, 100 mM NaCl, 10% glycerol, pH 7.4). For long-term storage, proteins were immediately frozen in liquid nitrogen and stored at – 80 °C until use.

D3 Halogenase reaction conditions

Analytical reactions were performed using 20 μM D3, 5 μM HPAC flavin reductase, 100 μM FAD, 50 mM sodium bromide, and 500 μM of (−)-premarineosin A (3) dissolved in DMSO was added and then diluted to a final volume of 250 μL with reaction buffer (10 mM HEPES, 10% glycerol, pH 7.4) and initiated by adding NADH (5 mM). Two control reactions were performed which included all contents except D3. Reactions were incubated at 30 °C for 4 h agitating at 600 rpm in a thermoshaker (Multithermoshaker, Benchmark) and quenched via addition of 750 μL of methanol, followed by vortexing at full speed S3 for 30 s. Quenched reactions were centrifuged at 17,000 × g to remove insoluble material, and the supernatant was analyzed with chromatographic conditions identical to analytical reactions and were performed using an Agilent G6230B time-of-flight (TOF) mass spectrometer system operating in positive mode, monitoring a mass range of 200 to 1200 amu with ESI-MS, and UV (195−400 nm) detection. ESI conditions were set with the capillary temperature at 320 °C, source voltage at 3.5 kV, and a sheath gas flow rate of 11 L min⁻¹, and the first 1 min of flow was diverted to waste. Reaction supernatants were also analyzed in comparison to standards using an analytical HPLC (Shimadzu) equipped with a PDA detector and analyzed with a Phenyl-Hexyl column (Luna 5 μM Phenyl-Hexyl 100 Å, LC Column 250 × 4.6 mm, RT). Water + 0.1% formic acid (A)/acetonitrile + 0.1% formic acid (B) (20% to 80% B) was used for the mobile phase at 1 ml min⁻¹. Preparatory reactions were performed using 40 μM D3, 5 μM HPAC flavin reductase, 100 μM FAD, 50 mM sodium bromide, and 500 μM of (-)-premarineosin A dissolved in DMSO at a final volume of 5 mL in reaction buffer (10 mM HEPES, pH 7.4 containing 10% glycerol). The reaction was initiated upon the addition of 5 mM NADH. The reaction was quenched via the addition of 20 mL of HPLC-grade methanol, vortexed on the highest setting for 20 s, then water bath sonicated for 30 s. The resulting mixture from all combined 5 mL reactions was then passed through a pad of Celite© and the filter cake was washed with an additional 20 mL of methanol. To purify the biocatalytic byproduct, the aqueous solution was extracted with DCM. The organics were combined and concentrated. The resulting material was further purified by preparative HPLC with a Phenyl-Hexyl column (5 μm, 100 Å, 250 × 10 mm) using a gradient of 10–55% of acetonitrile and water, both modified with 0.1% formic acid, over 50 min at a flow rate of 5 mL min⁻¹. Products were analyzed using LC-MS/MS (Supplementary Fig. 8), HPLC, and NMR (Supplementary Data 1).

NBS chemical bromination

To a solution of (−)-premarineosin A (3) (13.8 mg, 5 mmol) in dichloromethane at 4 °C was added N-Bromo succinimide (10 mg, 10 mmol, DCM) drop wise. The mixture was stirred at 4 °C. After 2 h, the reaction was quenched with sodium bicarbonate. Following a liquid-liquid extraction with DCM, the organic layer was extracted and dried. Reaction supernatants were analyzed in comparison to standards using an analytical HPLC (Shimadzu) equipped with a PDA detector and analyzed with a Phenyl-Hexyl column (Luna 5 μM Phenyl-Hexyl 100 Å, LC Column 250 × 4.6 mm, RT). Water + 0.1% formic acid (A)/acetonitrile + 0.1% formic acid (B) (20% to 80% B) was used for the mobile phase at 1 ml min⁻¹. The resulting material was further purified by preparative HPLC with a phenyl hexyl column (5 μm, 100 Å, 250 × 10 mm) using a gradient of 10–55% of acetonitrile and water, both modified with 0.1% formic acid, over 50 min at a flow rate of 5 mL min⁻¹. Products were analyzed using LC-MS/MS (Supplementary Fig. 8), HPLC, and NMR (Supplementary Data 1).

Mammalian cell lines and cell culture

Cells were obtained from the following sources: HEK293 (ATCC, cat # CRL-1573), MOLT-4 (ATCC, cat # CRL-1582). Cell lines were routinely tested for Mycoplasma contamination with the MycoAlert PLUS Mycoplasma Detection Kit (Lonza Bioscience, cat # LT07) according to manufacturer protocol. Cell number and cell viability were measured using the Countess Automated Cell Counter (Invitrogen) using Trypan blue stain (Invitrogen, cat # T10282). Human embryonic kidney HEK293 cells were maintained in DMEM with high glucose and GlutaMAX (Gibco, cat # 10566016) supplemented with 10% HyClone Characterized Fetal Bovine Serum (Cytiva, cat # SH30071.03), and 100 µ mL⁻¹ Penicillin-Streptomycin (Gibco, cat # 15140122). Human T lymphoblast MOLT-4 cells were maintained in RPMI 1640 Medium (Gibco, cat # 11875093) supplemented with 10% HyClone Characterized Fetal Bovine Serum (Cytiva, cat # SH30071.03), 100 µ mL⁻¹ Penicillin-Streptomycin (Gibco, cat # 15140122), and 10 mM HEPES (Gibco, 15630080). Cells were cultured in incubators maintained at 37 °C with 5% CO₂ and 95% relative humidity.

CellTiter-Glo cell viability assay (CTG) and data analysis

The CTG assay (Promega, cat # G7572) was used to quantify cellular ATP according to manufacturer protocols. The Multidrop Combi liquid dispenser (ThermoFisher) was used to dispense 4 µL of cells into white 1536-well microplates (Corning, cat # 7464) at a density of 1500 cells/well. Plated cells were incubated overnight. (−)-Premarineosin A (3) and derivatives were transferred into respective wells of each plate at 25 nL/well with the mosquito (SPT Labtech) in 16-pt, 1:3 titrations for a final concentration of 125 µM–8.71 pM for compounds with a high concentration of 20 mM, or 62.5 µM–4.36 pM for compounds with a high concentration of 10 mM. Controls were transferred into respective wells of each plate at 25 nL/well with the mosquito with a 16-pt, 1:2 titration of Digitonin for a final concentration of 125 µM–3.81 nM. The final concentration of DMSO was 0.58%. Cells were incubated for either 24, 48, and 72 h, after which point 3 µL/well of CTG was transferred to each plate with the BioRaptr 2.0 FRD (LetsGoRobotics). Plates were incubated for 10 min in the dark at ambient temperature for 10 min. Luminescence was measured using a ViewLux 1430Ultra HTS (PerkinElmer) with the following optical settings: exposure = 1 s, gain = high, speed = slow, binning = 2X. Dimethyl sulfoxide (“DMSO”; AMRESCO, cat # RGE-3070) was used as a vehicle control. Digitonin (Sigma-Aldrich, cat # D141) was prepared as a 20 mM stock solution in DMSO and stored at −30 °C for use as a cytotoxicity control. (−)-Premarineosin A (3) and derivatives were prepared at either 10 mM or 20 mM stock solutions in DMSO. Data were normalized to 125 µM Digitonin as −100% inhibition in a row-wise manner across the plate. Normalization was performed in Excel (Microsoft) and Concentration Response Curves (CRCs) were plotted in GraphPad Prism (GraphPad Software, Inc.) with error bars representing the standard deviation of four replicate wells (Supplementary Fig. 9).

In vitro antimalarial assay

The antimalarial growth inhibition assay was performed as described previously⁶⁸. Briefly, the Plasmodium falciparum Dd2 and 3D7 parasites were diluted to 0.75% parasitemia with 2% hematocrit, and 50 μL diluted parasites were added to each well in a 96-well plate containing 50 μL of properly diluted drugs. Tested compounds were threefold diluted in triplicate with concentrations ranging from 20 – 0.001 µM/10 – 0.0005 µM. The parasites were incubated with the drug at 37 °C under mixed gas (5% O₂, 5% CO₂, and 90% N₂) conditions for 48 h. DNA was released from the cultured parasite and stained with SYBR green dye. The plate was placed in the dark with gentle agitation for 1 h, and signals were read in a FLUOstar OPTIMA reader (BMG Labtech, Germany). Data from the microplate reader were analyzed as described previously⁶⁸ and plotted using Prism 9.0 software (GraphPad Software, Inc., San Diego, CA). Each in vitro experiment was performed in triplicate wells and repeated twice. Response Curves (CRCs) were plotted in GraphPad Prism (GraphPad Software, Inc.) with error bars representing the standard deviation of three replicate wells (Supplementary Fig. 10).

General information

Reagents and solvents were purchased from EMD Millipore, Sigma-Aldrich, Oakwood Chemical, Chem Impex, Thermo-Fisher Scientific, AABlocks, Advanced Chem Blocks, TCI, or Arctom unless indicated otherwise. NMR spectra were recorded on a Bruker 600 NMR system (600 MHz). High-resolution mass spectra as well as analytical reaction analysis were recorded on an Agilent Technologies 6250 TOF LC/MS equipped with an Agilent 1290 Infinity II HPLC. Samples were also analyzed via ultra-high-performance liquid chromatography-quadrupole time-of-flight mass spectrometry (UHPLC-LCMS) on an Agilent 1290 Infinity II UHPLC coupled to an Agilent 6545 ESI-Q-TOF-MS. Samples were injected and data was acquired using auto-MS/MS in positive mode with a Phenomenex Kinetex Phenyl-Hexyl (1.7 μm, 2.1 × 50 mm) column. Compounds were eluted with an isocratic elution of 90% solvent A (100% water + 0.1% formic acid) for 1 min followed by a 9 min linear gradient elution to 100% solvent B (95% acetonitrile + 5% water + 0.1% formic acid). The capillary temperature was 320 °C with a source voltage of 3.5 kV and a sheath gas flow rate of 11 L min⁻¹ for electrospray ionization (ESI).

Statistical information

Statistical analysis was performed with GraphPad Prism version 10.4.2 unless otherwise described. Specific details on statistical tests are provided in figure legends and relevant methods.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Crystal structure data has been deposited in the Cambridge Crystallographic Data Centre (CCDC) under deposition numbers CCDC 2455429 for (−)-premarineosin A (3) and CCDC 2455430 for gem-dimethyl-bridged premarineosin A (4). Data generated during this study are included in the published article and in the supplemental information files supplied (Supplementary Data 1–4). All other datasets generated during this study are available from the corresponding authors upon reasonable request.

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Acknowledgements

We are grateful for support from NIH grant R35GM118101 and the Hans W. Vahlteich Professorship (to D.H.S.), NIH grant R35GM159915 (to F.P.), a National Science Foundation Graduate Research Fellowship #DGE2241144 (to C.M.M.), an NIH F-31 #5F31DA055451-03 (to N.R.H.), the University of Michigan College of Pharmacy Duellman Graduate Student Research Fund #PGG030232 (to S.A.), a Michigan Pioneer Fellowship (to B.J.C.), and an NIH F-31 #1F31AI186432-01 (to K.L.L.). This research was also supported in part by the Intramural Research Programs of the National Center for Advancing Translational Sciences, NIH under project 1ZIA TR000495-01 (to J.I.) and the Division of Intramural Research of the National Institute of Allergy and Infectious Diseases, NIH under project # ZIA AI000892-23 (to Xz.S). This work was also supported by the University of Michigan BioNMR Core Facility (U-M BioNMR). The U-M BioNMR Core is grateful for support from U-M including the College of Literature, Sciences and Arts, the Life Sciences Institute, the College of Pharmacy, and the U-M Biosciences Initiative. We thank Prof. William Fenical (Scripps Institute of Oceanography, San Diego, CA) for helpful conversations and the gift of Streptomyces sp. CNQ-617. We are grateful to Rajani Arora for her contributions to the graphical abstract.

Author information

These authors contributed equally: Christina M. McBride, Morgan McCauley.
These authors jointly supervised this work: Filipa Pereira, David H. Sherman.

Authors and Affiliations

Life Sciences Institute, University of Michigan, Ann Arbor, MI, USA
Christina M. McBride, Morgan McCauley, Natalia R. Harris, Sahar Amin, Brian J. Curtis, Linnea Verhey-Henke, Katherine L. Lev, Filipa Pereira & David H. Sherman
Program in Chemical Biology, University of Michigan, Ann Arbor, MI, USA
Christina M. McBride & Katherine L. Lev
Department of Biological Chemistry, University of Michigan Medical School, Ann Arbor, MI, USA
Natalia R. Harris & Filipa Pereira
Department of Medicinal Chemistry, University of Michigan, Ann Arbor, MI, USA
Sahar Amin & David H. Sherman
Laboratory of Malaria and Vector Research, National Institute of Allergy and Infectious Diseases, NIH, Rockville, MD, USA
Awet A. Teklemichael & Xin-Zhuan Su
National Center for Advancing Translational Sciences (NCATS), NIH, Rockville, MD, USA
Erin N. Oliphant, Patricia Dranchak & James Inglese
Department of Chemistry, University of Michigan, Ann Arbor, MI, USA
Fengrui Qu & David H. Sherman
Department of Chemistry, Indiana University, Bloomington, IN, USA
Harrison M. Snodgrass & Jared C. Lewis
Metabolic Medicine Branch, National Human Genome Research Institute, NIH, Bethesda, MD, USA
James Inglese
Department of Microbiology and Immunology, University of Michigan, Ann Arbor, MI, USA
David H. Sherman

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Christina M. McBride
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Morgan McCauley
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Contributions

C.M.M. performed experiments, analyzed data, and wrote the manuscript. M.M. designed experiments, performed experiments, analyzed data, and wrote the manuscript. N.R.H. performed experiments, analyzed data, and wrote the manuscript. S.A. performed experiments, analyzed data. B.J.C. analyzed data. L.V.H. performed experiments. A.A.T., E.N.O., K.L., P.D., F.Q., performed experiments, analyzed data. H.M.S. performed experiments. J.C.L. supervised the project. J.I. analyzed data, supervised the project, designed experiments, provided feedback on the manuscript. X.S. supervised the project, designed experiments, provided feedback on the manuscript. F.P. conceived the project, designed experiments, performed experiments, analyzed data, supervised the project, and wrote the manuscript. D.H.S. conceived the project, designed experiments, supervised the project, and wrote the manuscript. All authors approved the final version of the manuscript.

Corresponding authors

Correspondence to Jared C. Lewis, Filipa Pereira or David H. Sherman.

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Competing interests

The authors declare the following competing interests: C.M.M., M.M., N.R.H., S.A., F.P., and D.H.S. are inventors on a filed provisional patent application related to findings described in this manuscript (U.S.A. Provisional Patent Application Number: 63/811,085, “Engineering Premarineosin Production in Streptomyces and Derivatization of the Premarineosin Scaffold for Drug Development”). All other authors declare no competing interests.

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McBride, C.M., McCauley, M., Harris, N.R. et al. Metabolic engineering and late-stage functionalization expand the chemical space of the antimalarial premarineosin A. Commun Chem 8, 391 (2025). https://doi.org/10.1038/s42004-025-01779-6

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Received: 03 July 2025
Accepted: 29 October 2025
Published: 05 December 2025
Version of record: 05 December 2025
DOI: https://doi.org/10.1038/s42004-025-01779-6