Engineered Microbial Platform Confers Resistance against Heavy Metals via Phosphomelanin Biosynthesis

doi:10.21203/rs.3.rs-4671167/v1

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Engineered Microbial Platform Confers Resistance against Heavy Metals via Phosphomelanin Biosynthesis

https://doi.org/10.21203/rs.3.rs-4671167/v1

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Environmental concerns are increasingly fueling interest in engineered living materials derived from microbial sources. Melanin biosynthesis in microbes, particularly facilitated by recombinant tyrosinase expression, offers sustainable protection for the habitat of microorganisms against severe environmental stressors. However, there exists a vast urgency to optimize these engineered microbial platforms, which will amplify their protective capabilities, integrate multifaceted functions, and thereby expand their utility and effectiveness. In our work, we have genetically engineered microbial platforms capable of endogenously biosynthesizing phosphomelanin, a unique phosphorus-bearing melanin featured by its distinctive phosphate and phosphonate groups. The ability to heterogeneously biosynthesize phosphomelanin endows the microbes with enhanced resistance to heavy metals, thus safeguarding their survival in adverse conditions. Furthermore, we have upgraded these engineered microbes by integrating PET-degrading enzymes, thereby achieving effective integrated management of metallized plastic waste. This engineered microbial platform, with its phosphomelanin biosynthetic capabilities, presents significant opportunities for microbes to engage in bioengineering manufacturing, potentially serving as the next-generation guardians against global ecological challenges.

Biological sciences/Biological techniques

Physical sciences/Materials science/Biomaterials

Physical sciences/Chemistry/Biosynthesis

Biological sciences/Biochemistry/Biocatalysis

phosphomelanin

synthetic biology

engineered microbial platform

heavy metal resistance

metallized plastic degradation

Heavy metals (HMs) and metallized wastes constitute a persistent and pervasive global environmental challenge^1,2. HMs are a collection of metallic elements characterized by their high density, and toxicity or harmfulness to living organisms³. Being non-biodegradable, they have a tendency to bio-accumulate in the tissues of organisms and undergo bio-magnification across trophic levels. This process can lead to detrimental health effects across the spectrum of life forms, including microbes, plants, animals, and humans. Various strategies have been developed for the remediation of HM pollution. The most commonly used method involves the addition of amendments such as precipitating chelating agents (e.g., limestone, lime)⁴, complexing chelating agents (e.g., ethylenediaminetetraacetic acid, citric acid)⁵, and phosphate compounds⁶. These amendments can help immobilize HMs, reducing their bioavailability and mobility, and thereby decreasing their toxicity. However, conventional methods for HM removal have several drawbacks, including potential secondary pollution risks and significant alterations to the physical and chemical properties of soil and water⁷. Consequently, it is imperative to explore efficient and cost-effective methods to mitigate the ecological damage caused by HMs and metallized wastes, while also minimizing the associated risks to human health during the remediation process.

Microbes play a multitude of vital roles in environmental protection, including the purification of pollutants and the bio-remediation of contaminated sites, thereby contributing to the sustainability and resilience of ecosystems^8–10. Microbial biotechnology has thus emerged as an effective and eco-friendly solution for the treatment of HMs and metallized waste¹¹. It is well known that metals are involved in the growth and metabolism of microbes in the process of their adaptive evolution, either directly or indirectly¹². Precisely for this reason, microbes can exhibit resistance to HMs. However, exposure to high levels of HMs can cause irreversible damage to microbes. This includes the disruption of microbial structures, disturbance of oxidative phosphorylation, conformational alterations of nucleic acids and proteins, and the induction of enzyme inactivation¹³. Enhancing the resistance to HMs is crucial for improving the effectiveness of microbes as a sustainable and efficient approach for HM pollution management.

Melanogenic microbes, leveraging their inherent metal tolerance mechanisms and the endogenous biosynthesis of melanin, demonstrate metallophilic and metal-resistant characteristics superior to those of ordinary microbes¹⁴. This renders them highly effective for the bio-remediation of HM-polluted environments, due to the metal affinity and high adsorption capacity of melanin. A typical example is A. salmonicida subsp. pectinolytica, an extremophile isolated from a heavily polluted river, which utilizes its own oxidase and peroxides to manipulate the cascade biosynthesis of melanin¹⁵. The melanin biosynthesized within this extremophile acts as metal-binding agents, providing protection against high concentrations of HMs¹⁶. Additionally, the Gram-positive bacterium Bacillus megaterium (B. megaterium) is a versatile and stress-resistant melanogenic bacterium capable of resisting HMs^17,18. While melanogenic microbes provide a sustainable and eco-friendly alternative for HM removal, the low concentrations of melanin generated by naturally-occurring strains are currently insufficient to address the escalating issue of HM contamination¹⁰.

Engineering microbial systems aim to customize or optimize microbes with programmable functionalities using genetically engineered and approaches. This nascent field holds the potential to revolutionize diverse sectors, from sustainable manufacturing to environmental management and ecological sustainability¹⁹. Currently, ongoing efforts are focused on the overexpression of genes responsible for melanin-producing enzymes (eg., tyrosinases (TYRs), phenoloxidases) in both melanogenic and non-melanogenic microbes, have not only boosted the inherent melanogenic potential, but also enabled the creation of novel melanin-recombinant microbial strains^20,21. Engineered microbes have successfully facilitated the production of various types of melanin, such as eumelanin, pheomelanin, pyomelanin, neuromelanin, allomelanin. However, despite these advancements, there is considerable scope for further progress in the engineering of strains capable of biosynthesizing novel-type-melanins to combat HM pollution.

The phosphate group, a key component of essential biomolecules such as nucleic acids and adenosine triphosphate (ATP), has a strong tendency to interact with metal ions due to its negatively charged oxygen atoms^22,23. For instance, metachromatic granules are densely packed particles widely found within bacteria, the main component of which is polyphosphate (polyP), and play a significant role in microbial resistance to environmental HM stress. At the same time, most chassis microorganisms are capable of utilizing organophosphorus compounds and converting them into endogenous substances¹⁸. Herein, we have designed and customized an engineered melanogenic microbe that achieves the biosynthesis of a new type of melanin. We leveraged a series of chassis organisms with upregulated tyrosinase gene (TYR gene) and employed a phosphorylated tripeptide as the substrate. Peptides possess flexible tunability^24–26, and by encoding their structure, it is possible to tailor enzymatic reaction substrates, thereby achieving engineered bacteria with customized properties. This engineering microbial system is capable of biosynthetically producing a novel melanin enriched with phosphorus, which we have designated as “phosphomelanin”. This phosphomelanin featured by its distinctive phosphate and phosphonate groups, is a new member of the melanin family. We demonstrate here that the production of phosphomelanin endows the microbes with enhanced resistance to HMs. Moreover, we showcase the potential of combining phosphomelanin biosynthesis with the functionality of poly(ethylene terephthalate) degrading enzymes (PETase) within an engineered melanogenic microbial platform, thereby achieving effective and integrated management of metallized plastic waste.

Phosphomelanin Production from Engineered Melanogenic Microbes

TYR functions as a prototypical multifunctional binuclear copper oxidase pivotal in melanogenesis and enzymatic browning processes²⁷. The manipulation of the TYR gene, which is essential for melanin and its analogues production in living organisms, enables the customization of microbes to achieve enhanced TYR expression. Employing a standard genome editing method²⁸, we initially engineered a shuttle plasmid (designated as phis 1525), which incorporates the promoter and repressor genes of the B. megaterium-borne xylose utilization operon. Following this, the TYR gene from B. megaterium (referred to as bmTYR gene) was integrated into phis 1525 under the control of the xylose operon. This step produced the recombinant shuttle plasmid, designated as phis 1525-bmTYR (Fig. 1a). Upon transforming phis 1525-bmTYR into the wild-type (WT) B. megaterium strain WH320, we successfully developed an engineered bacteria named GE-WH320 with high-level expression of the TYR (Fig. 1b-c, Supplementary Fig. 1). By utilizing the regulatory elements of the xylose-utilization operon, we were able to induce the expression of the heterologous bmTYR up to 350-fold in B. megaterium²⁸.

TYR features a spacious active site pocket that is specifically tailored to accommodate Tyr-based substrates, thereby facilitating their conversion into melanin or melanin analogues^29–32. In our study, we designed the tripeptide Ser-Tyr-Gly (SYG) and the phosphorylated tripeptide _pSer-Tyr-Gly (_pSYG) as substrates. The SYG is derived from the reactive sequence found in the green fluorescence proteins of Aequorea victoria³³, wherein Tyr functions as the site for the enzyme-catalyzed reaction, and Ser provides a covalent bonding site for the phosphate group, thereby facilitating the efficient phosphorylation of melanin and its analogues. The molecular structures of these two tripeptides were subsequently elucidated using a combination of nuclear magnetic resonance (NMR) and electrospray ionization mass spectroscopy (ESI-MS) (Supplementary Fig. 2–3). Following a 12-hour co-culturing of the GE-WH320 with _pSYG, a macroscopic colonial browning reaction was evident in the culture medium. This pronounced color change was not detected during the co-incubation of the WH320 with _pSYG (Fig. 1d and Supplementary Fig. 4a). The observed coloration suggests that the engineered bacteria are highly expressing the TYR, which catalyzes the Tyr-containing tripeptide substrate to produce melanin-like substances³⁰. Scanning electron microscopy (SEM) revealed the generation of heterogeneous aggregates within the GE-WH320 following co-culturing with _pSYG. This resulted in a transition from smooth to rough bacterial surfaces, attributable to the deposition of these melanin-like aggregates (Fig. 1e). The co-incubation melanin-like products derived from the GE-WH320 exhibited amplified absorption in the UV-vis region when compared to those from the WT bacteria (Fig. 1f). These observations collectively validate the successful biosynthesis of brown-black pigments within the engineered bacteria, which possess classic characteristics of melanin.

Co-culturing with phosphorylated substrates resulted in a notably amplified phosphorus signal intensity in the GE-WH320 compared to WH320, as evidenced by ³¹P solid-state NMR (ssNMR) spectroscopy (Fig. 1g). Furthermore, quantitative analysis using inductively coupled plasma atomic emission spectrometry (ICP-AES) showcased a substantial 2.9-fold escalation in phosphorus content in the GE-WH320 relative to the WT bacteria (Fig. 1h). These results underscore an augmented accumulation of phosphorus during the biosynthesis process of melanin analogues catalyzed by TYR within the engineered bacteria. These findings signify the successful biosynthesis of a phosphorus-containing melanin analogue, which we have designated as “phosphomelanin”, within the customized engineered bacteria. The intracellular enzymatic cascade oxidation process involving phosphorus, as the critical step in the biosynthesis of phosphomelanin, is found to be an exceedingly mild procedure (Fig. 1i, Supplementary Fig. 4b-c). SEM results unveiled that the cell wall and cytoplasmic membrane of the engineered bacteria retained their structural integrity subsequent to phosphomelanin generation. Additionally, the self-organization phenomenon during the expansion of bacterial colonies was observed. Upon transferring these colonies, which have generated phosphomelanin, to a fresh Luria-Broth (LB) culture medium, they demonstrated exponential growth following an “S-type” growth curve³⁴.

Structural Analysis of Phosphomelanin

The commercially available TYR extracted from the champignon mushroom Agaricus bisporus (referred to as abTYR) exhibits high homology with bmTY (Supplementary Fig. 5a), attributable to their shared dicopper active centers²⁷. This makes abTYR a well-suited model for in vitro studies of melanogenesis. The substrates _pSYG and SYG were each co-incubated with abTYR at 37 ℃ for 48 hours under aerobic conditions. The UV-vis absorption spectra spanning the range of 200 nm to 800 nm confirmed unequivocally the generation and accumulation of melanin analogues (_pSYG_ox and SYG_ox) in vitro (Supplementary Fig. 5b). The ³¹P ssNMR spectroscopic features of the phosphomelanin catalyzed by abTYR closely resembled those of the phosphomelanin generated by bmTYR (Supplementary Fig. 5c). Notably, when compared to the substrate _pSYG, the phosphomelanin (_pSYG_ox) exhibited resonance peaks with comparable ³¹P chemical shifts (δ), signifying a high degree of consistency in their chemical structure³⁰. This suggests that the phosphomelanin is synthesized through the polymerization of the substrate peptide molecules (_pSYG). The generated phosphomelanin was verified as a water-soluble, heterogeneous aggregate with a black, electron-dense structure, as revealed by cryo-transmission electron microscopy (cryo-TEM) (Supplementary Fig. 6). The in vitro synthesis of phosphomelanin via abTYR-catalyzed oxidative polymerization achieved a high yield rate of 57% after 48 hours. Leveraging a combination of ¹³C NMR and ¹³C ssNMR techniques, we unraveled the abTYR-catalyzed oxidative polymerization process responsible for the formation of phosphomelanin.

In the ¹³C NMR spectra, signals at positions 1/1' (δ = 130 ppm) and 2/2' (δ = 115 ppm), corresponding to the meta- and ortho- carbons of the hydroxyl carbon within the aromatic region, demonstrated a strong consistency between the structure of substrates (_pSYG) and its oxidized products (_pSYG_ox). This consistency served as evidence of the generation of polymers with repeating units (Fig. 2a). Moreover, the intensities of these two signals assigned to meta-carbon and ortho-carbon were notably augmented in the cross-polarization/magic angle spinning (CP/MAS) ¹³C ssNMR spectrum compared to those detected by ¹³C NMR (Fig. 2b). This observation indicated that the abTYR-catalyzed oxidative polymerization on the phenolic hydroxyl group in the substrate monomer _pSYG resulted in the formation of densely packed solid-state structures. Furthermore, matrix-assisted laser desorption/ionization (MALDI) mass spectrometry (MS) provided additional evidence for the polymeric structure of _pSYG_ox (Supplementary Fig. 7), revealing characteristic structurally heterogeneous and multi-dispersed oligomeric fragments. The Fourier transform infrared (FTIR) spectrum of the substrate _pSYG revealed the presence of the phenolic groups of Tyr, evidenced by the distinct band at 1518 cm^− 1. Subsequent to the abTYR-catalyzed oxidative polymerization, the FTIR spectrum of the _pSYG_ox displayed characteristic bands at 1601 cm^− 1 and 1680 cm^− 1, which are attributed to the catechol and quinone moieties, respectively (Fig. 2c). These findings conclusively establish that the enzymatic polymerization process of phosphomelanin is a phenol-to-quinone oxidative reaction³⁵.

The existing form and distribution of phosphorus within phosphomelanin _pSYG_ox were further elucidated using ³¹P NMR spectroscopy by the chemical shifts. For substrate _pSYG, its ³¹P NMR spectrum featured a distinct and narrow peak signal at 3 (δ = − 0.32 ppm), which is indicative of a phosphomonoester group, indicating the presence of a phosphate group connected to the hydroxyl group of the Ser side chain. This characteristic group was also detected in _pSYG_ox (3', δ = 0.76 ppm). Additionally, an additional peak signal at 4 (δ = 2.6 ppm) emerged for _pSYG_ox, indicating the formation of a newly formed phosphorus-bearing group (Fig. 2d). These above two types of phosphorus-bearing groups were also detectable through ³¹P ssNMR, with corresponding peak signals at δ = − 0.98 ppm and δ = 2.04 ppm, respectively (Supplementary Fig. 8). A comparison analysis of the ¹³C NMR spectra for SYG_ox and _pSYG_ox has facilitated further structural elucidation of the newly formed phosphorus-bearing group. In SYG_ox, a sharp single peak signal was observed at δ = 119 ppm (Supplementary Fig. 9), whereas in _pSYG_ox, a set of split peak signal was detected (Supplementary Fig. 10a-b), suggesting that the newly formed phosphorus-bearing group is linked to the ortho-carbon of the phenol ring within Tyr residue. Within _pSYG, the signals were additionally split with a coupling constant of ₂J_C−P = 7.04 Hz, indicative of the existence of P-O bonds. In the case of _pSYG_ox, in addition to the P-O bonds, new signals were observed with additional splitting and coupling constants of ₁J_C−P = 103.84 Hz (Fig. 2e, Supplementary Fig. 10c), which implies the presence of P-C bonds³⁶. X-ray photoelectron spectroscopy (XPS) peak fitting of _pSYG_ox revealed two distinct absorption peaks: one at 133.7 eV, also detectable in the substrate _pSYG, corresponding to the phosphorus group connected to Ser via a P-O bond; and a newly emerged peak at a binding energy of 132.8 eV (Fig. 2f), attributable to the P-C bonds within the newly formed phosphorus group^37,38. The above results confirm the coexistence of two types of phosphorus-bearing species in the _pSYG_ox: one being the phosphate groups existing as P-O bonds, inherited from substrate molecules _pSYG; the other being the phosphonate groups existing as P-C bonds, presumably formed through a dehydration condensation reaction between the phosphate group and the dopa-quinone group, where the quinone radical anions binds to the phosphonate groups (Supplementary Fig. 10d).

The process of phosphomelanin generation through abTYR-catalyzed oxidative polymerization can be speculated as follows. In Stage I, a sequential cascade of oxidation catalysis was initiated, commencing from mono-phenol and proceeding to di-phenol, and ultimately leading to the formation of the quinone intermediate. Advancing to Stage II, the essential repetitive building blocks of phosphomelanin were formed, involving the P-C bond-forming reaction. Subsequently, in Stage III, the oxidative polymerization process continued, leading to the elongation and diversification of the polymer chains, ultimately yielding the heterogeneous phosphomelanin (Fig. 2g, Supplementary Fig. 11). In natural systems, there are several classical types of protein phosphorylation that play essential roles in cellular signaling and regulation³⁹. These types, apart from Ser phosphorylation, typically include Thr phosphorylation and Tyr phosphorylation. In our study, we further purposefully designed and synthesized phosphate-bearing tripeptides including _pThr-Tyr-Gly (_pTYG) and _pTyr-Tyr-Gly (_pYYG) (Supplementary Fig. 12–13), to simplify mimic the phosphorylated proteins found in nature. Co-incubation of these two substrates with abTYR can also resulted in generation of phosphomelanin (Supplementary Fig. 14–15). The experimental results obtained from the UV-vis absorption spectra, ³¹P NMR spectra, and ¹³C NMR spectra provided evidence that, in the presence of abTYR, the substrate molecules underwent oxidative polymerization, resulting in the formation of phosphomelanin (_pTYG_ox and _pYYG_ox). Given the important role of phosphorylation in organisms, elucidating the biosynthesis of phosphomelanin will offer new insights into the evolution of melanin function and the genetic foundation of melanization in microorganisms⁴⁰.

Kinetics of Enzymatic Production of Phosphomelanin

TYR is composed of a four-helix bundle core bearing a magnetically coupled dinuclear copper center in a deep active site pocket, capable of oxidizing phenols attached to cargo molecules to yield o-quinones²⁷. These copper ions facilitate the transfer of an atom from molecular oxygen (O₂) to the orthoposition of a bound phenol molecule. With the aim to elucidate the nature of the copper binding sites and the presence of radical intermediates of the copper ions in abTYR, spectroscopic studies by EPR were carried out. Results revealed the existence of an “EPR-silent” state around g = 2.5 ~ 2.05 (Extended Data Fig. 1a). The absence of the EPR signal has been shown to derive from antiferromagnetic coupling between the copper (II)s, which requires a superexchange pathway associated with a bridging ligand²⁷.

The kinetics of abTYR-catalyzed oxidative polymerization of phosphomelanin can be elucidated based on three historically recognized states of the active center copper atom: deoxy [Cu(I)-Cu(I)]; oxy [Cu(II)-O₂²⁻-Cu(II), µ-η²:η²-peroxido dicopper (II)]; and met [Cu(II)-O-Cu(II), µ-hydroxido dicopper (II)]^31,41,42. In the initial stage of phosphomelanin biosynthesis, the abTYR in the deoxy-form state activates O₂ to generate a µ-η²:η²-peroxido dicopper (II) core, representing the oxy-form state. This leads to the regioselective hydroxylation of phenols, resulting in the formation of catechols, while concurrently forming a µ-hydroxido dicopper (II) core (met-form state). Subsequently, abTYR facilitates the further oxidation of catechols to quinones, while the copper core in the met-form state reverts back to the deoxy-form state (Supplementary Fig. 16a). Kinetic experiments using _pSYG and SYG as substrates respectively, were performed by monitoring spectrophotometrically the corresponding melanin analogues in the region of 300 ~ 800 nm. Results showed that under the commensurate enzyme-catalyzed reaction conditions, phosphorus-bearing substrates (_pSYG) exhibit higher enzymatic reaction efficiency compared to substrates without phosphorus (SYG) (Extended Data Fig. 1b and Supplementary Fig. 16b-c). In addition to the pathway in which TYR catalyzes the oxidative polymerization of Tyr residues resulting in the production of melanin, a parallel competitive pathway exists. This pathway involves TYR catalyzing the oxidative cross-linking of Tyr residues to form stable and covalent o, o’-dityrosine products with characteristic fluorescence. Excitation of those fluorescent intermediates at a wavelength of 315 nm results in a characteristic emission centered on 400 nm³². It was observed that the fluorescence generated in the abTYR-catalyzed process using _pSYG as the substrate was significantly lower than that using SYG as the substrate (Supplementary Fig. 16d-e and 17). These results indicated that the introduction of phosphorus in the substrates can significantly enhance the efficiency of abTYR in catalyzing the oxidative polymerization, thereby leading to improved melanin production.

Further, energy analysis and molecular docking studies were conducted to reveal the influence of phosphorus-containing groups on the abTYR-catalyzed oxidative reactions. In the simulation, the subunit of recombinant mushroom TYR (abPPO4, PDB ID: 5M6B, chain A) was taken as the receptor, and substrates including _pSYG and SYG were used as the ligands⁴³. The computational docking results were visualized to illustrate the strength of the bonding forces and the best geometrical arrangements of the substrate molecule (_pSYG and SYG) within the active pocket of the abPPO4 enzyme (Extended Data Fig. 1c-d). Results showed that phosphorus-containing groups in substrates _pSYG provided abundant hydrogen bonding sites inside TYR (abPPO4) enzyme. Amino acid residues Cys80, Ser83, Lys421, and Thr457 formed six hydrogen bonds with _pSYG. Comparably, only two hydrogen bonds located on the Cys80 and Thr457 were found between this enzyme and SYG. The predicted binding energy for _pSYG is -2.7 kcal mol^− 1, which is stronger than that for SYG at -2.4 kcal mol^− 1. Consequently, it can be concluded that _pSYG demonstrates a higher binding affinity to abTYR than SYG, implying that phosphorus-containing substrates may enhance the activity of abTYR-catalyzed oxidative reactions.

Metal Binding Ability of Phosphomelanin

Given the bioaccumulation, persistence, and toxicity of HMs, there is an urgent need for sustainable remediation approaches to address their environmental hazards³. Melanin have the ability to form stable complexes with HMs through various functional groups, such as hydroxyl, carboxyl, and phenolic hydroxyl⁴⁴. These complexes are typically insoluble, facilitating their precipitation from solutions. This property can be effectively utilized in environmental remediation processes, especially for the adsorption and elimination of HMs from aquatic systems. Building on this eco-friendly strategy, we have investigated the metal adsorption and removal capabilities of phosphomelanin with a focus on its interactions with a selection of representative HMs, including Cr^II, Pb^II, Yb^III, Dy^III, and Tb^III (Fig. 3a). Upon mixing the nitrate salts of the aforementioned HMs with _pSYG_ox in aqueous solutions, the initially tawny-colored _pSYG_ox solution lightened to a pale yellow (for Cr^II) and became colorless (for Pb^II, Yb^III, Dy^III, and Tb^III). Concurrently, insoluble dark-colored complexes rapidly formed and precipitated (Fig. 3b). The findings indicate that phosphomelanin forms effective complexes with heavy metals, transforming them into easily manageable precipitates, and surpasses traditional melanin in sequestration capabilities. The strong binding capabilities of phosphomelanin with HMs was also supported by the results from UV-vis spectroscopy (Fig. 3c) and ICP-AES (Fig. 3d, Supplementary Fig. 18). Under identical conditions, the incorporation of phosphorus significantly enhanced the melanin’s ability to remove Cr^II, Pb^II, Yb^III, Dy^III, and Tb^III, with the concentrations of HMs in the precipitates increasing by 41.8%, 4.01%, 32.18%, 51.70%, and 8.67%, respectively.

Subsequently, we employed a suite of ssNMR techniques, including 1D ¹H NMR spectra, 1D ³¹P NMR spectra, as well as 2D ¹H-³¹P heteronuclear chemical shift correlation experiments (HETCOR) under magic angle spinning (MAS), to elucidate the intermolecular interactions within HM-phosphomelanin complexes. These analyses were instrumental in revealing the mechanisms by which phosphorus-bearing melanin augment the binding capacity for HMs. Specifically, we selected the Pb^II-_pSYG_ox complex as an example to provide insights into the role of phosphorus-bearing moieties in the sequestration of HMs. As shown in Fig. 3e-f, the 1D ³¹P and ¹H NMR spectra were utilized to investigate the alterations in electron density of the Pb^II-bound and Pb^II-free states of _pSYG_ox. After the formation of the Pb^II-_pSYG_ox complex, the ³¹P resonance shifted to a higher frequency region. This indicated that Pb^II caused a decrease in the electron density around the phosphorus atom. This can be attributed to the electrostatic interaction between the negatively charged phosphorus groups and the positively charged Pb^II. And, the ¹H resonances indicated that the aromatic hydrogen (H_aromatic) and the active hydrogen (H_active) in Pb^II-_pSYGox complex all shifted to a higher frequency region, while the resonance of aliphatic hydrogen (H_aliphatic) stayed the same. These results above indicate that Pb^II interacts with the carboxyl, dopaquinone, and phosphoryl groups of _pSYGox, facilitating the adsorption and removal of HMs by the phosphomelanin. Furthermore, 2D ¹H-³¹P HETCOR spectra were measured to provide more detailed structural information of Pb^II-_pSYG_ox complex^45,46. Two groups of ³¹P resonances were detected from _pSYGox, which correspond to the phosphonate group (around − 0.94ppm) and phosphate group (around 0.23 ppm), respectively (Fig. 3g and Supplementary Table 1). Upon complexation with Pb^II, the ³¹P resonances observed in _pSYG_ox coalesce and relocate to chemical shifts of -0.43 ppm, togetherly. These chemical shifts indicate the column force between Pb^II ions and the phosphate and phosphonate groups within _pSYG_ox, resulting in the formation of the Pb^II-_pSYG_ox complex.

In situ Biosynthesis of Phosphomelanin in Microbes Enabling Resistance to HMs

Encouraged by the exceptional performance of phosphomelanin on the removal of HMs in vitro, we were motivated to develop novel microbes capable of in situ biosynthesis of phosphomelanin to combat HM pollution. In our work, we successfully constructed an engineered bacteria based on the typical Gram-negative platform strain (E. coli TOP 10), capable of heterologous expression of bmTYR (Fig. 4a and Supplementary Fig. 19). Initially, we synthesized the bmTYR gene and placed it under the control of the strong constitutive promoter J23100 (ref.47). Subsequently, we transformed the recombinant plasmid pUC57-bmTYR into the E. coli TOP 10, resulting in high levels of TYR expression. Following a 12-hour incubation with the substrate _pSYG, the color of the engineered bacteria solution progressively intensified. Imaging from cryo-transmission electron microscopy (cryo-TEM) showed that the _pSYG-incubated engineered bacteria were coated with heterogeneous aggregates, signifying the in situ production of phosphomelanin within bacteria (Fig. 4a). The engineered microbes capable of biosynthesizing phosphomelanin can effectively bind to HMs. This was confirmed through analysis using Energy-dispersive X-ray spectroscopy (EDS) and XPS (Fig. 4b and Supplementary Fig. 20).

A four-sample parallel experiment was designed to investigate the resistance of phosphomelanin-producing engineered E.coli to HMs. The WT E. coli TOP 10 served as the negative control, while the WT B. megaterium, which naturally expresses TYR, served as the positive control. Additionally, the engineered E. coli TOP 10, which has produced melanin after incubation of 24 h with SYG and _pSYG separately, were included as the experimental groups. Engineered microbes synthesize melanin analogues (_pSYG_ox and SYG_ox) in situ without significantly altering the surface charge of the biological cell membrane (Fig. 4c), which reflects the mildness of the melanin biosynthesis process. Compared to the “γ” group, engineered bacteria with biosynthesized phosphomelanin display a more negative surface potential, which can be attributed to the auto-ionization of phosphorus-bearing groups around the membrane environments. These four microbial strains were incubated with five typical toxic-level HMs, including Cr^II, Pb^II, Yb^III, Dy^III, and Tb^III, to evaluate their resistance to these HMs (Fig. 4d). The resistance ability was assessed using optical density measurements at a wavelength of 600 nm (OD₆₀₀), a method widely utilized to estimate the cell density in liquid cultures⁶. The engineered E. coli TOP 10 containing phosphomelanin (_pSYG_ox) demonstrated high resistance to all toxic-level of HMs. In comparison, the engineered E. coli TOP 10 containing phosphorus-free melanin (SYG_ox) exhibited decreased resistance to HMs, with nearly no survival at the equivalent concentration level of Cr^II and Yb^III, and showed significant growth suppression in Dy^III and Tb^III. The WT B. megaterium and the non-genome-edited WT E. coli TOP 10 were unable to survive under any of the toxic-level HMs conditions. It is worth noting that when we re-inoculated these phosphomelanin-producing microbes, which had bound HMs, into a fresh culture medium and subjected them to shaking incubation, we observed that all these microbes grew vigorously and were capable of continuous division and proliferation (Fig. 4e). The SEM results show that the microbes maintained intact morphology, uniform size, and clear surface features. Microbes that generate phosphomelanin possess more advanced protective functions against HMs than their natural counterparts⁴⁸. Phosphomelanin, within these microbes, exhibits high metal-binding affinity and serves as an impermeable barrier to toxic HMs. This barrier effect prevents the HMs from entering the cytoplasm and stops them from complexing with cellular components such as DNA and proteins, thereby enhancing the microbes’ resistance and tolerance to environments rich in HMs⁴⁹.

By constructing another recombinant shuttle plasmid (pP43-bmTYR), the bmTYR gene can also be successfully expressed in B. subtilis 168 strain. This indicates that the engineering of microbes to produce phosphomelanin is not only achievable in the Gram-negative bacterium but is also feasible across the Gram-positive bacteria (Fig. 4f). Following co-incubation with the substrate _pSYG, the engineered bacteria exhibited significant production of deep-black phosphomelanin. This was further validated by cryo-TEM imaging equipped with EDS, which depicted the deposition of aggregates on the bacterial surface with a distinct contrast to the bacteria themselves (Fig. 4g), and by UV-vis absorption spectra that showcased characteristic peaks for melanin or its analogues (Supplementary Fig. 21). ICP-AES measurement and elemental mapping confirmed the presence of phosphorus within these melanin-based aggregates (Fig. 4g,h). Collectively, these findings confirm the biosynthesis of phosphomelanin by the engineered B. subtilis 168. These engineered B. subtilis 168 also demonstrated robust growth under conditions with toxic-level concentrations of HMs, including Cr^II, Pb^II, Yb^III, Dy^III, Tb^III, Co^II, and Mn^II (Fig. 4i). Notably, the engineered strain can achieve resistance to high concentrations of HMs through the refactoring of the phosphomelanin biosynthesis pathway. This straightforward enzymatic biosynthesis route is distinct from the complex gene regulatory mechanisms³². It serves as a simple protective mechanism that not only enables microbes to thrive in extremely harsh HM environments but also facilitates the management of HMs. Synthetic biology, with a sophisticated arsenal of tools, presents an exciting avenue for the creation of a comprehensive microbial platform that produces phosphomelanin in situ a compound with significant potential in neutralizing HM hazards¹⁹.

Phosphomelanin-Producing Microbes in situ Secreted Hydrolase for PET Depolymerization in HM Surroundings

Plastics, particularly microplastics (MPs), when associated with HMs, have emerged as a new class of pollutants. This association can modify the environmental behaviors, bioavailability, and toxicity of the HMs, posing significant ecological risks. However, to date, attention has primarily been focused on the enzymatic degradation of plastic pollutants, with the impact of HMs being largely overlooked⁵⁰. Based on the aforementioned research on the genetic engineering of microorganisms, we have further designed a gene cluster with the aim of simultaneously achieving the biodegradation of PET plastic and the adsorption of HMs. This integrated approach could potentially offer a more efficient and targeted solution for environmental remediation, particularly for the treatment of plastic-HMs composite pollutants (Fig. 5a). Here, B. subtilis 168 was chosen as a platform to exogenous express recombinant PET hydrolase (PETase). A biosynthetic gene cluster was constructed within a shuttle plasmid pP43-NMK for E.coli and B.subtilis. And then, a constitutive promoter for B. sublitis, P43 promoter, was contained within this shuttle plasmid to achieve a non-induced expression. Furthermore, the Bacillus signal peptide fragment (SP_amy) was fused to the N-terminus of PETase gene from Indeonella sakainesis to maximize the excellualr secretion of PETase⁵¹. Simultaneously, the bmTYR gene was also integrated to facilitate the intracellular biosynthesis of phosphomelanin. WB measurements confirmed that PETase was secreted into the extracellular environment, while bmTYR remained within the engineered bacteria (Supplementary Fig. 22). The PETase and bmTYR encoding genes are each preceded by the B. subtilis mntA ribosomal binding site (RBS) (AAGAGGAGG), which is a considered as a strong Shine-Dalgarno sequence (ΔG > 50.4 kJ mol^− 1), to ensure efficient translation⁵².

Additionally, the formation of phosphomelanin in engineered bacteria and their HM resistance to Co^II and Mn^II were demonstrated by cyro-TEM, EDS and selected area electron diffraction (SAED) analyses (Fig. 5b and Supplementary Fig. 23). Selecting the monomer of PET, bis (2-hydroxyethyl) terephthalate (BHET) as a model, the biodegradation capability of engineered bacteria in the presence of HMs was further evaluated. As shown in Fig. 5c, these PETase secreted by the engineered bacteria catalyzed in vitro hydrolysis reactions of ester bonds within BHET. The high-resolution electrospray ionization (HR-ESI) measurements confirmed that these ester bonds of BHET were hydrolyzed into terephthalic acid (TPA) and ethylene glycol (EG) monomers (Supplementary Fig. 24). The BHET block degraded into fragments within 1 h and was further digested into smaller pieces after 6 h. Over a period of 36 h, the BHET block was completely decomposed, resulting in a visible clear solution. While there were no morphological changes observed, when the BHET block was inciubated with 50 mM Gly-NaOH buffer (Fig. 5d). The PET film can also be degraded by the engineered bacteria. We immersed BHET blocks and PET films in a solution of engineered bacteria and assessed their weight loss at various time points to confirm the biodegradation capability of the engineered bacteria on plastics (Supplementary Fig. 25). We can observe that by the 28th day, the intact PET film had been broken down into smaller fragments (Fig. 5c insert). After that, SEM was employed to visually confirm the changes in surface morphology during the PET film degradation process (Fig. 5e). The surface of PET film remained smooth and even when incubated in a culture medium or co-incubated with non-engineered bacteria. However, when incubated with the bmTYR-expressing B. subtilis 168 strain, the PET film’s surface lost its original luster, attributing to the generation of phosphomelanin. Notably, incubation of the PET film with the engineered PETase-bmTYR-expressing B. subtilis 168, resulted in a visibly roughened and wrinkled film surface, indicating the biodegradation of the PET film by the engineered bacteria. This rough surface exposed more reactive sites for PET hydrolysis reactions and sped the whole degeneration course eventually. Moreover, 2D XRD measurements confirmed changes in the crystallographic orientation of the PET film as the degeneration progressed (Extended Data Fig. 2). The PET film exhibited three distinct intensity peaks, where the in-plane crystallographic orientations were represented by peaks with d-spacings of 6.8 Å and 3.9 Å, and the out-of-plane orientation was indicated by a peak with a d-spacing of 3.4 Å. The results showed a decrease in all three crystallographic orientations during the engineered bacteria-induced degeneration, with the out-of-plane orientation (d = 3.4 Å) demonstrating a more pronounced reduction compared to the in-plane orientations (d = 6.8 Å and d = 3.9 Å). This suggests that the PET polymer films, which are packed perpendicularly to the film plane, are more readily catalyzed by PETase secreted by engineered bacteria.

The recombinant PETase enzyme exhibits its highest catalytic activity at approximately 50 ℃. However, the chassis microbe B. subtilis 168 does not thrive at this high temperature⁵³. To address this, we have further developed a novel thermophilic engineered bacteria based on the WT B. subtilis HT50 (Extended Data Fig. 3). As anticipated, this engineered thermophilic bacteria is not only capable of in situ biosynthesis of phosphomelanin within the bacteria, which effectively binds to HMs such as Co^II and Mn^II, but also demonstrate excellent biodegradation efficiency for PET at approximately 50°C. This engineered thermophilic bacteria shows potential as a suitable candidate for industrial strains, where a moderate increase in temperature can enhance the efficiency of industrial processes for treating plastic-HM composites. In summary, applying principles of synthetic biology (Supplementary Table 2), we have constructed an engineered bacterial platform that offers functionality: it utilizes in situ biosynthesized phosphomelanin for resistance against HMs and also employs the release of PETase enzyme for plastic degradation. The establishment of this engineered bacterial platform provides an efficient and sustainable solution for the treatment of plastic-HM composite pollutants.

Here, we have customized and optimized engineered microbial systems that confer resistance against heavy metals and metalized plastic wastes via phosphomelanin biosynthesis, employing biological synthetic methodologies. We have demonstrated that tyrosinase expression in engineered melanogenic microbe is proficient to produce a novel melanin enriched with phosphorus, which we have designated as “phosphomelanin”, using a phosphorylated tripeptide as the substrate. We have further delineated the chemical architecture of phosphomelanin, identifying the critical presence of two distinct phosphorus-containing moieties: phosphate groups existing as P-O bonds and phosphonate groups existing as P-C bonds. These endogenously biosynthesized phosphomelanins within engineered microbe serve a pivotal role; they exhibit excellent metal-binding capabilities, thereby enhancing the engineered microbes’ resistance and tolerance to heavy metals. Furthermore, we have upgraded these engineered microbes by integrating PET-degrading enzymes, thereby conferring upon them with the dual capabilities of endogenous phosphomelanin biosynthesis and extracellular secretion of PET hydrolases, successfully achieving the biodegradation of metallized plastic waste. These findings underscore the vast potential of engineered microbes capable of producing phosphomelanin, extending their applications far beyond traditional bioremediation. The phosphomelanin, as a new member of the melanin family, endows these microbes with an unprecedented opportunity to function effectively in extreme situations, which is poised to foster significant advancements in sustainable manufacturing processes. Moreover, it enables a more holistic and comprehensive approach to the management and treatment of the environmental waste.

Figures

Plasmids and chassis microbes

The plasmid (pUC57-bmTYR) and the E. coli TOP 10 were purchased from Tsingke Biotechnology. The WT B. megaterium was sourced from the China General Microbiological Culture Collection Center. The B. megaterium WH320, the B. subtilis 168, and the shuttle plasmids (phis 1525-bmTYR, pP43 NMK-bmTYR, pP43 NMK-amyPETase-bmTYR, pHT304) were purchased from Forhigh Biotechnology. The B. subtilis HT 50 was isolated from the sewer pipe of the lab.

Media and reagents

Peptides were sourced from GL Biochem Ltd (Shanghai, China), while abTYR was purchased from Sigma-Aldrich Co., Ltd. (Darmstadt, Germany). LB, TB, and LA cultural media were purchased from Solarbio Science & Technology Co., Ltd. (Beijing, China). Additional reagents, including D(+)-xylose, tetracycline, and kanamycin sulfate, were purchased from Solarbio Science & Technology Co., Ltd. (Beijing, China). BHET and dialysis tubes (D-Tube Dialyzer Maxi) were purchased from Sigma-Aldrich Co., Ltd. (Darmstadt, Germany). PET filmswith a thickness of 0.0125 mm were purchased from DuPont (France). All other chemicals were purchased from Beijing Chemical Co. Ltd. (Beijing, China) and were used without further purification or modification.

Preparation and purification of melanin analogues

Enzymatic oxidation to produce melanin analogues was performed using abTYR under aerobic conditions. Briefly, the TYR-containing precursors (_pYYG, _pTYG, _pSYG, and SYG) were dissolved in PBS (0.1 M, pH 8.5) to a concentration of 5.0 mg mL^-1 in 10 mL quartz tubes. These tubes were then equipped with polytetrafluoroethylene stoppers and stoppers were sealed with parafilm to ensure airtight conditions. The solutions were saturated with pure oxygen for 30 minutes to create an oxygen-rich environment. Following saturation, abTYR at a concentration of 1.0 KU mL^-1 was added to the above solutions. The tubes were incubated at room temperature for 24 hours with continuous oxygen saturation to facilitate the enzymatic reaction and promote the formation of melanin analogues.

Following the enzymatic reactions, the melanin analogues were purified using a chromatography column packed with Bio-Gel P gels (G2, Extra Fine). The column was operated at a flow rate of 0.05 mL min-1 with MilliQ water as the mobile phase. The purified melanin analogues (_pYYG_ox, _pTYG_ox, _pSYG_ox, and SYG_ox) were then sealed individually into dialysis tubes with a molecular weight cut-off of 3.5 kDa. These tubes were immersed in 5 liters of MilliQ water for 48 hours to eliminate inorganic phosphates from the PBS. The water was refreshed six times during this period.

Metal-binding study

Melanin analogues were lyophilized over 48 hours. Subsequently, all melanin analogues were redissolved in MilliQ water to a final concentration of 10.0 mg mL^-1 for both SYG_ox, and _pSYG_ox. Solutions of these melanin analogues (100 μL at 10.0 mg mL^-1) were mixed with 900 μL of various metal nitrate solutions (Cr, Pb, Yb, Dy, and Tb at a concentration of 0.5 mmol mL^-1). The resulting insoluble metal-binding complexes were obtained by high-speed centrifugation at 12,000 rpm for 30 minutes. The complexes were then dialyzed against MilliQ water to remove any free metal ions. For the dialysis process, the insoluble complexes were dispersed in 1.0 mL of MilliQ water, and the samples were sealed in dialysis tubes with a molecular weight cut-off of 3.5 kDa. The tubes were immersed in 1.0 L of MilliQ water for 48 hours, with the water being replaced six times over this period.

Morphology characterization

SEM images were captured using a Hitachi S-4800 SEM, with samples placed on the silicon slices. To enhance sample conductivity, a layer of platinum (Pt) was sputtered onto them. Cryo-TEM images were obtained using a JEOL JEM-2310 and F200 instrument. EDS images were obtained using a JEOL JEM-2100F and F200 instrument at a temperature of 100 K, with samples protected by a thin layer of amorphous ice. Zeta potential measurements were conducted using a Malvern Zetasizer Nano ZS ZEN3600. Micrographs were captured using a charge-coupled device (CCD) camera from Gantega, integrated with Olympus Soft Imaging Solutions.

Spectroscopy characterization

FTIR spectra were recorded using a TENSOR-27 infrared spectrometer. 1D ¹H, ¹³C, ³¹P, and 2D in-situ ¹H-¹H COSY NMR spectra were obtained on a 700-MHz Bruker Avance spectrometer equipped with a triple resonance cryogenic probe and utilizing built-in 1D and 2D pulse sequences. 1D ¹³C, 1D ³¹P, and 2D in-situ ¹H-³¹P HECTOR ssNMR spectra were obtained using a Bruker AVANCE III 600. The spectrometer is equipped with a magic angle spinning (MAS) 4.0 mm probe. All chemical shifts were referenced to adamantane (d_H=1.91 ppm, d_C=38.48 ppm). Compressed air was used to regulate the sample’s temperature at room temperature. ESI-MS were obtained using a Bruker APEX II-FT-ICR-MS. MALDI-MS were acquired on a Bruker Autoflex III-MALDI-TOF-MS. UV-vis absorption spectra were collected using a Shimadzu UV-2600 spectrophotometer with a 1.0 mm path-length quartz cuvette. Fluorescence measurements were performed using a Hitachi Model FL-4500 spectrofluorometer with a 1.0 cm quartz cuvette. Fluorescence decay measurements were conducted on an Edinburgh FLS1000 steady-state and transient fluorescence spectrometer. EPR measurements at 9.37 GHz were conducted using a Bruker EMXplus-9.5/12 with a microwave power of 20 mW, and all samples were analyzed at 100 K under aerobic conditions. XPS measurements were measured by ESCALab250Xi. ICP-MS values were obtained using a Thermo Scientific iCAP RQ ICP-MS, with all samples requiring lyophilization for 48 hours prior to analysis.

Construction of heterogeneous TYR-expressing E. coli

The prokaryotic TYR gene from B. megaterium (bmTYR) was inserted into the relaxed plasmid pUC57-kan. These plasmids were then introduced into the E. coli TOP 10 to express bmTYR. A constitutive promoter, J23100, was utilized to ensure overexpression of bmTYR within Gram-negative bacteria. Subsequently, AGE was performed to verify the correctness of the target sequence in the plasmids. Upon digestion with XbaI and PstI, two electrophoretic bands were observed, located between 800 to 1200 bp and 2000 to 3000 bp, respectively. These results were consistent with the predicted sizes of 1104 bp and 2547 bp.

Construction of endogenous TYR-expressing B. megaterium

The endogenous bmTYR was inserted into the shuttle plasmid phis1525, which is capable of transferring between the E. coli TOP 10 and the B. megaterium WH320. The transformation method for the B. megaterium WH320 has been previously reported^54,55. The plasmid phis1525 includes the promoter and repressor gene from the B. megaterium operon responsible for xylose utilization. Expression of bmTYR is inducible by xylose, with 0.5% xylose added to the culture medium to induce expression; non-inducing broth contains no xylose. All media for culturing the B. megaterium WH320 are supplemented with tetracycline at a concentration of 20 μg mL^-1.

Construction of heterogeneous TYR-expressing B. subtilis

The bmTYR was inserted in the shuttle plasmid pP43 NMK^56,57. This recombinant plasmid was then introduced into the B. subtilis 168 to express bmTYR. The constitutive promoter P43 was utilized to ensure the overexpression of bmTYR within the gram-positive bacteria⁵⁸.

Construction of heterogeneous bi-enzyme-expressing B. subtilis

The bmTYR, the PETase gene from Ideonella sakaiensis, and the signal peptide (amy) gene from Bacillus amyloliquefaciens were inserted into the shuttle plasmid pP43-NMK-kan. The recombinant plasmids were then introduced into the B. subtilis to enable the co-expression of bmTYR and the secretion of PETase. For the B. subtilis HT 50, the three genes were inserted into the shuttle plasmid pHT304. Electroporation was employed to transform all these plasmids into the bacteria.

WB analysis

Total bacterial protein from B. megaterium and B. subtilis was extracted using the BestBio bacterial total protein extraction kit. Protein quantification was performed using the BestBio BCA protein quantification kit. A cell lysate containing 30 mg of WH320 was mixed with Laemmli buffer (Bio-Rad) containing 5% 2-mercaptoethanol and boiled at 95°C for sample preparation. SDS-PAGE was conducted using 15 mL, 15-well precast mini-PROTEAN TGX gels (Bio-Rad) in a Mini-PROTEAN tetra cell at 200 V with 1× Tris/glycine/SDS running buffer (Bio-Rad). A PageRuler Plus pre-stained protein ladder (Thermo Fisher) was used as a molecular weight marker. Proteins were then transferred onto Immobilon-FL PVDF membranes (Millipore Sigma) at 100 V for 1 hour at 4°C using 1× Tris/glycine transfer buffer (Bio-Rad) with 0.2% methanol. Membranes were blocked at room temperature for 1 hour with Odyssey blocking buffer (TBS, LI-COR).

Production of phosphomelanin within the engineered bacteria

The engineered bacteria (E. coli TOP 10, B. megaterium WH320, or B. subtilis 168) were cultured in LB medium. Specifically, a 10 mL overnight culture of engineered bacteria was inoculated into a 200 mL flask with 50 mL of sterile LB. Here, 100 µg mL^-1 kanamycin was added to the engineered E. coli TOP 10 and engineered B. subtilis 168, while 20 µg mL^-1 tetracycline was added to the engineered B. megaterium WH320. The engineered bacteria were incubated at 37 °C with shaking at 220 rpm for 24 hours. Subsequently, the substrate (_pSYG) was added to the culture media. The engineered bacteria were then broken using ultrasound, and the resulting suspension was centrifuged at 4°C to separate the components. The dark, clear supernatant was collected and lyophilized eventually. The crude phosphomelanin produced by the engineered bacteria was further purified using a chromatography column with MilliQ water as the mobile phase. Finally, ³¹P ssNMR measurements were conducted to reveal the structural information of the purified products.

Identification and quantitation of products in BHET or PET degradation in vivo

The optimal reaction buffer for the recombinant PETase is 50 mM glycine-NaOH buffer at pH 9.0, which contains 10% (v/v) crude enzyme solution. The enzyme was used to hydrolyze a block of BHET measuring 1.0 x 1.5 x 0.3 cm at 50°C for 36 hours. The hydrolytic products were subsequently analyzed directly using MS.

The engineered B. subtilis 168 was cultured in TB medium. Specifically, a 10 mL overnight culture of engineered bacteria was inoculated into a 200 mL flask with 100 mL of sterile LB, 10 µg mL^-1 kanamycin, 10 µg mL^-1 _pSYG, 100 µg mL^-1 Co^II, and 100 µg mL^-1 Mn^II ion. The engineered bacteria were incubated at 37 °C with shaking at 220 rpm for 24 hours with the BHET block (100 mg) or PET film (5.0*5.0 cm). As for the engineered HT 50, the culture temperature remains at 50 °C.

Molecular docking

A Lamarckian genetic algorithm in AutoDock (Molecular Graphics Laboratory, La Jolla, CA, USA) was applied for the molecular docking study^59,60. The structural information of abTYR (PDB ID: 5m6b) was retrieved from the Protein Data Bank. The initial conformations of _pSYG and SYG within the active pocket of abTYR for the docking simulation were determined using density functional theory (DFT) calculations at the B3LYP/6-31G* level, which were carried out using CP2K (version 2023.1) package.

References

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Online content

Any methods, additional references, Nature Portfolio reporting summaries, source data, supplementary information, acknowledgments, peer review information; details of author contributions and competing interests; and statements of data and code availability are available at https://doi.org/xxxxxx.

Acknowledgments

We are grateful to Dr. Zonglin Liang from the Institute of Microbiology, Chinese Academy of Sciences and Forhigh Biotechnology for their insightful discussions, which greatly assisted in the design of the TYR-expressing system. We also extend our thanks to Dr. Junfeng Xiang at the Center for Analysis and Measurement, Institute of Chemistry, Chinese Academy of Sciences for his valuable input on the design of the NMR measurements. This work was financially supported by the National Science Fund for Distinguished Young Scholars of China (No. 22025207), the Innovation Research Community Science Fund of China (No. 21821005), National Natural Science Foundation of China (Nos. 22232006, 22072154 and 22377127), the Youth Innovation Promotion Association of Chinese Academy of Sciences (No. 2021048), and the Beijing Nova Program (No. 20230484352).

Author contributions

X. Yan, R. Xing and X. Ren conceived the idea and designed the experiments. X. Ren carried out the sample preparation, purification and characterization. L. Zhao, C. Yuan assisted with the chemical analysis of MS and NMR measurements. L. Zhao and P. Zhou contributed to the MD simulations and kinetic analysis. K. Zhao contributed to PET degeneration experiment, film structural characterization and data analysis. X. Ren, R. Xing and X. Yan wrote the paper with inputs from all authors.

Competing interests

The authors declare no competing interests.

Additional information

Extended data is available for this paper at https://doi.org/xxxxxx.

Supplementary information The online version contains supplementary material available at https://doi.org/xxxxxx.

Correspondence and requests for materials should be addressed to Xuehai Yan.

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Engineered Microbial Platform Confers Resistance against Heavy Metals via Phosphomelanin Biosynthesis

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Abstract

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Main

Results

Phosphomelanin Production from Engineered Melanogenic Microbes

Structural Analysis of Phosphomelanin

Kinetics of Enzymatic Production of Phosphomelanin

Metal Binding Ability of Phosphomelanin

Conclusion

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Methods

Declarations

References

Additional Declarations

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