ACS Publications. Most Trusted. Most Cited. Most Read
Light-Driven Hybrid Nanoreactor Harnessing the Synergy of Carboxysomes and Organic Frameworks for Efficient Hydrogen Production
My Activity
  • Open Access
Research Article

Light-Driven Hybrid Nanoreactor Harnessing the Synergy of Carboxysomes and Organic Frameworks for Efficient Hydrogen Production
Click to copy article linkArticle link copied!

  • Jing Yang
    Jing Yang
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    More by Jing Yang
  • Qiuyao Jiang
    Qiuyao Jiang
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    More by Qiuyao Jiang
  • Yu Chen
    Yu Chen
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    More by Yu Chen
  • Quan Wen
    Quan Wen
    Hubei Key Laboratory of Agricultural Bioinformatics, College of Informatics, Huazhong Agricultural University, Wuhan 430070, China
    More by Quan Wen
  • Xingwu Ge
    Xingwu Ge
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    More by Xingwu Ge
  • Qiang Zhu
    Qiang Zhu
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Qiang Zhu
  • Wei Zhao
    Wei Zhao
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Wei Zhao
  • Oluwatobi Adegbite
    Oluwatobi Adegbite
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
  • Haofan Yang
    Haofan Yang
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Haofan Yang
  • Liang Luo
    Liang Luo
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Liang Luo
  • Hang Qu
    Hang Qu
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Hang Qu
  • Veronica Del-Angel-Hernandez
    Veronica Del-Angel-Hernandez
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
  • Rob Clowes
    Rob Clowes
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    More by Rob Clowes
  • Jun Gao
    Jun Gao
    Hubei Key Laboratory of Agricultural Bioinformatics, College of Informatics, Huazhong Agricultural University, Wuhan 430070, China
    More by Jun Gao
  • Marc A. Little*
    Marc A. Little
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    Institute of Chemical Sciences, Heriot-Watt University, Edinburgh EH14 4AS, U.K.
    *Email: [email protected]
  • Andrew I. Cooper*
    Andrew I. Cooper
    Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    *Email: [email protected]
  • Lu-Ning Liu*
    Lu-Ning Liu
    Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    College of Marine Life Sciences and Frontiers Science Center for Deep Ocean Multispheres and Earth System, Ocean University of China, Qingdao 266003, China
    *Email: [email protected]
    More by Lu-Ning Liu
Open PDFSupporting Information (1)

ACS Catalysis

Cite this: ACS Catal. 2024, 14, 24, 18603–18614
Click to copy citationCitation copied!
https://doi.org/10.1021/acscatal.4c03672
Published December 6, 2024

Copyright © 2024 The Authors. Published by American Chemical Society. This publication is licensed under

CC-BY 4.0 .

Abstract

Click to copy section linkSection link copied!

Synthetic photobiocatalysts are promising catalysts for valuable chemical transformations by harnessing solar energy inspired by natural photosynthesis. However, the synergistic integration of all of the components for efficient light harvesting, cascade electron transfer, and efficient biocatalytic reactions presents a formidable challenge. In particular, replicating intricate multiscale hierarchical assembly and functional segregation involved in natural photosystems, such as photosystems I and II, remains particularly demanding within artificial structures. Here, we report the bottom-up construction of a visible-light-driven chemical–biological hybrid nanoreactor with augmented photocatalytic efficiency by anchoring an α-carboxysome shell encasing [FeFe]-hydrogenases (H–S) on the surface of a hydrogen-bonded organic molecular crystal, a microporous α-polymorph of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP-α). The self-association of this chemical–biological hybrid system is facilitated by hydrogen bonds, as revealed by molecular dynamics simulations. Within this hybrid photobiocatalyst, TBAP-α functions as an antenna for visible-light absorption and exciton generation, supplying electrons for sacrificial hydrogen production by H–S in aqueous solutions. This coordination allows the hybrid nanoreactor, H–S|TBAP-α, to execute hydrogen evolution exclusively driven by light irradiation with a rate comparable to that of photocatalyst-loaded precious cocatalyst. The established approach to constructing new light-driven biocatalysts combines the synergistic power of biological nanotechnology with the multilength-scale structure and functional control offered by supramolecular organic semiconductors. It opens up innovative opportunities for the fabrication of biomimetic nanoreactors for sustainable fuel production and enzymatic reactions.

This publication is licensed under

CC-BY 4.0 .
  • cc licence
  • by licence
Copyright © 2024 The Authors. Published by American Chemical Society

Introduction

Click to copy section linkSection link copied!

Solar energy harvesting and conversion have the potential to contribute to a sustainable, carbon-neutral environment. (1−3) In nature, multicomponent photosynthetic systems capture solar energy and use enzymatic cascades to orchestrate charge separation and catalytic proton-coupled electron transfer reactions to produce diverse natural products, albeit with certain intrinsic limitations, such as a relatively narrow light-harvesting window and susceptibility to chemical and structural damage by higher energy photons. (4−7) Artificial photocatalysts employ synthetic materials, often inorganic semiconductors, to mimic natural photosynthetic systems, typically by combining various components to achieve the essential functions. (8) For example, semiconductor particles have the capability of absorbing light, and when platinum nanoparticles are placed on the surface of the semiconductor, they could facilitate the reduction of protons to generate hydrogen gas. (9−11) While these synthetic hybrids can achieve high solar-to-hydrogen efficiencies, particularly in the ultraviolet range of the solar spectrum, (12) they lack the complex, hierarchically structured functionality and selectivity that are present in natural photosynthetic systems. Moreover, it is still challenging to develop artificial systems that can use the whole solar spectrum, and the most effective direct photocatalysts for overall water splitting operate in the ultraviolet spectral range, which constitutes only about 7% of total solar energy. (13)
One promising strategy to leverage the advantages of both biological and abiotic systems is to synergistically integrate natural biological machinery (such as enzymes, organelles, or whole cells) with artificial materials to construct new chemical–biological hybrid systems. (14−17) Such hybrid systems can, in principle, combine broadband light absorption and high exciton generation efficiency from synthetic semiconductors and the selectivity and catalytic power of living biocatalysts. Chemical–biological hybrids can be generated through surface engineering (e.g., surface ionization (18,19)), redox mediator addition (e.g., methyl viologen (20)), and enzyme encapsulation in porous materials. (21) However, establishing an efficient physical-chemical connection between biological and nonliving materials, which is crucial for facilitating electron cascades and energy transfer to catalytic centers and exceeding the catalytic potential of each component, poses a significant challenge. (22) A major obstacle lies in the incompatibility between biocatalysts and synthetic materials in their working environment. Thus, there is a pressing need for more designable and effective approaches to constructing chemical–biological hybrid nanoreactors, allowing for precise structural control across various length scales.
Carboxysomes are anabolic bacterial microcompartments (BMCs) responsible for biological carbon fixation in cyanobacteria and some chemoautotrophic bacteria. (23−25) The carboxysome sequesters the central CO2-fixing enzyme, Ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisco), and carbonic anhydrase (CA) within a self-assembling proteinaceous shell. The carboxysome shell is composed of numerous protein paralogs that are in the form of hexamers and pentamers and allows the entry of HCO3 to create a CO2-rich microenvironment for facilitating carboxylation. (26−28) These natural features of carboxysomes have inspired the design and reprogramming of carboxysome structures by using synthetic biology for various biotechnological applications. In addition, hydrogenases are native catalysts for H2 production in bacteria, archaea, and some eukaryotes with exceptional catalytic activity and efficiency and act as valid candidates for biological H2 production. (29,30) In particular, [FeFe]-hydrogenases are highly active for H2 generation but are limited by their high oxygen sensitivity and irreversible inactivation by oxygen. (31) Therefore, the carboxysome shell has demonstrated great potential to serve as novel nanoreactors to boost H2 evolution. (32,33) However, the reported carboxysome-based H2-evolving nanoreactors require a constant supply of electrons from methyl viologen, which is not sustainable.
Here, we report a bottom-up approach to constructing a visible-light-driven chemical–biological hybrid nanoreactor by interfacing a porous crystalline organic semiconductor, microporous α-polymorph of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP-α), with recombinant α-carboxysome shells (with an average diameter of 90 nm) that encase [FeFe]-hydrogenases (H–S). (33) The resulting hybrid photobiocatalyst was characterized by electron microscopy, confocal microscopy, isothermal titration calorimetry, photoluminescence and photoelectrochemical assays, and computational simulations. Our results demonstrate that the intimate bio- and abiotic interface, established through extensive hydrogen bonds, mediates charge transport between the organic semiconductor and the biocatalyst. This allows the light-driven hybrid nanoreactor (H–S|TBAP-α) to exhibit a high sacrificial H2-evolution efficiency compared to the photocatalyst alone, and the efficiency of this photobiocatalyst is comparable with analogous synthetic photocatalysts that use expensive precious metals, such as platinum.

Results and Discussion

Click to copy section linkSection link copied!

Preparation of Porous Crystalline Photocatalyst TBAP-α

Light-driven chemical–biological hybrid nanoreactors should comprise light-harvesting antenna with strong visible-light absorption, high exciton production efficiency in moderate aqueous solution, and favorable hydrolytic stability to integrate with biological machinery with robust stability and high catalytic conversion. Hydrogen-bonding organic framework (HOF) photocatalysts that contain discrete hydrogen-bonding molecular building blocks, in principle, possess great potential for this application due to their inherent structure-tunability, functionality, and solution processability. (34)
1,3,6,8-tetra(4′-carboxyphenyl)pyrene molecule (TBAP) has been investigated as a building block in HOFs, (35) metal–organic frameworks (MOFs), (36) and protein-HOFs assemblies. (37) TBAP has a pyrene core that can π–π stack and carboxylic acid groups that can hydrogen-bond (Figures S1–S3), making it an ideal candidate for assembling HOFs. The α-polymorph of TBAP, referred to hereafter as TBAP-α, was successfully crystallized by slow diffusion of chloroform (CHCl3) into a solution of TBAP dissolved in dimethylformamide (DMF). (35) In the TBAP-α structure, 2-D layers of hydrogen-bonded TBAP molecules are closely packed and interact through π–π stacking of the pyrene units, which are separated by 0.38 nm. This generates a columnar microporous structure with 1.7 nm 1-D pores and unsaturated hydrogen-bonding layers on the surfaces of the needle-shaped crystals (Figures 1b and S4). Here, we characterized TBAP-α using single-crystal X-ray diffraction (sc-XRD) (Table S1), powder X-ray diffraction (PXRD), and gas absorption analysis of the micrometer-sized needle-shaped crystals (Figure S5).

Figure 1

Figure 1. Generation of the organic semiconductor, the hydrogenase-containing carboxysome shell, and the chemical–biological hybrid nanoreactor, H–S|TBAP-α. (a) Chemical structure of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP). (b) Crystal structure of α-polymorph of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP-α). (c) Genetic organization of the α-carboxysome shell operon and hydrogenase-expressing operon for the production of recombinant shells and active [FeFe]-hydrogenases in E. coli, respectively, and a schematic model of the hydrogenase-shell nanoreactor (H–S). EP: CsoS2 C-terminus as the encapsulation peptide. The hydGXEF genes encode the crucial maturase enzymes for hydrogenase formation and activation, including HydE, HydF, and HydG. PDB ID: Ferredoxin (Fd)-[FeFe]-hydrogenase, 2N0S; ferredoxin-NADP reductase (FNR), 2XNJ. (d) 3D model representation of the H–S|TBAP-α hybrid system, in which H–S adheres to the hydrogen-bonding groups on the TBAP-α crystal surface.

TBAP-α exhibited remarkable visible-light absorption (Figure S6) and semiconducting properties with an optical bandgap of 2.30 eV and a conduction band (C.B.) of −1.15 V (Figure S7), determined by using the Mott–Schottky technique, performing a considerable thermodynamic driving force for H2 production from water. In addition, TBAP-α favors neutralized ascorbic acid (AA) aqueous solution as the sacrificial agent, (35,38) a commonly used sacrificial agent in artificial photosynthesis due to its solubility in water and the compatibility to work in a neutral medium. (39)

Self-Association of α-Carboxysome Shells and TBAP-α

The recombinant α-carboxysome shell is composed of CsoS1 and CsoS4 paralogs, as well as the linker protein CsoS2 that is attached to the inner surface of the shell. (32,33) CsoS2 exists as two distinct isoforms in Halothiobacillus neapolitanus: the longer form CsoS2A (∼130 kDa) and the short form CsoS2B (∼85 kDa). (40) The two hexameric shell proteins, CsoS1A and CsoS1C, which show 98% similarity in amino acid sequence, represent the most abundant components on the α-carboxysome shell (23) (Table S2). To verify the physical association between α-carboxysome shells and TBAP-α, we performed molecular dynamics (MD) simulations to study the binding between CsoS1A and TBAP-α, using the crystal structure of CsoS1A (PDB: 2EWH) and TBAP-α structure determined by sc-XRD. The calculated binding free energy is −22.10 kcal mol–1 by the MM/GBSA method in AmberTool22 (41) using the last 30 ns MD trajectory (150 frames) of the TBAP-α and CsoS1A protein binding simulation. MD simulations suggest that the residues mainly in the loop regions of CsoS1A, including Thr5, Arg34, Ser86, Gly89, Asp90, Lys94, Pro96, and Glu97, form strong interactions with the net-carboxylate on the surface of TBAP-α through hydrogen-bonding and salt bridges (Figure 2a–c and Tables S3 and S4). These amino acid residues are conserved among the main hexamer shell proteins, as indicated by multiple sequence alignment (Figure S8) and structural analysis (Figure S9), suggesting that the TBAP-α crystal surface interacts with α-carboxysome shells through cooperative hydrogen-bonding interactions with the shell proteins.

Figure 2

Figure 2. Binding between the α-carboxysome shell and TBAP-α. Results of molecular dynamics simulations of CsoS1A (PDB ID: 2EWH) and TBAP-α binding equilibrium states are shown in panels (a–c). (a) Front view. (b) Side view, showing the simulated binding interface between TBAP-α and the CsoS1A hexamer. (c) Zoom-in view of the noncovalent interactions (red dotted lines) formed between amino acid residues of CsoS1A (Pro96, Thr5, Ser86, Asp90, Gly89, Lys94, Arg34, Glut97) and TBAP-α (blue = N atoms, red = O atoms). (d) ITC thermogram resulting from the titration of TBAP (50 μM) into the α-carboxysome shell (3.75 μM) in TN buffer (20 mM Tris-HCl (pH = 8.0), 150 mM NaCl). (e) Thermodynamic profile of TBAP binding to α-carboxysome shell in TN buffer. ΔG, change in Gibbs Free Energy; ΔH, change in enthalpy; T, temperature in Kelvin; ΔS, change in entropy.

Next, the binding potential of the whole recombinant α-carboxysome shells and molecular TBAP-α was detected by using isothermal titration calorimetry (ITC) (Figure 2d). A low dissociation constant (Kd) was measured as 524 (±54.4) × 10–9 M, indicating strong affinity between α-carboxysome shells and TBAP-α. In addition, the negative ΔG value indicates that the association between α-carboxysome shells and TBAP-α is energetically favorable, while the ΔH value reveals that this process is enthalpy-driven, suggesting that hydrogen bonds and van der Waals interactions may be the main driving forces in the association process (Figure 2e). The ITC results were consistent with our computational findings.
To directly visualize the association between α-carboxysome shells and TBAP-α crystals, we generated recombinant α-carboxysome shells that encapsulate mCherry, by fusing mCherry with the encapsulation peptide CsoS2 C-terminus (mCherry-EP) and coexpressing mCherry-EP with shell proteins. This approach led to the generation of mCherry-EP-Shell (C–S) (Figures S10 and S11). The purified C–S and TBAP-α were then incubated in solution for 30 min to enable self-association. Scanning electron microscopy (SEM) confirmed the coating of C–S particles on the surfaces of needle-shaped TBAP-α crystals (Figure 3a–c). Moreover, mCherry exhibits a distinct absorption maximum (λEx = 587 nm) from that of TBAP-α (λEx = 390 nm). Confocal laser scanning microscopy (CLSM) showed that the needle-shaped TBAP-α crystals exhibit bright fluorescence when excited at 488 nm and very weak red fluorescence when excited at 561 nm (Figure 3d, top row). In contrast, strong red fluorescence was recorded along the surface of TBAP-α crystals incubated with C–S when excited at 561 nm, indicating the association of C–S with the TBAP-α crystal surface (Figure 3d, bottom row). Both SEM and CLSM results demonstrate that the surfaces of TBAP-α crystals were densely coated with C–S particles in an aqueous suspension. In comparison, a bulk sample of unfunctionalized pyrene crystals (42) was incubated with C–S using the same procedure to explore the contribution of the organized layer of unsaturated carboxylic acid groups on the TBAP-α surface. The SEM images showed no visible C–S particles on these pyrene crystals (Figure S12), indicating that unsaturated carboxylic acid groups serve as the main binding sites between TBAP-α and α-carboxysome shells.

Figure 3

Figure 3. Attachment of mCherry-encapsulated C–S shells on the surface of TBAP-α crystals. SEM images of (a) uncoated TBAP-α, (b) C–S|TBAP-α (40 × 3.6 μm2 size), and (c) zoom-in view of C–S|TBAP-α. Orange arrows show the C–S particles that were attached to the TBAP-α crystal surface. (d) Fluorescence images of TBAP-α only (top row) and the C–S|TBAP-α hybrid (bottom row). Left, excitation at 488 nm; middle, excitation at 561 nm; right, the merged channel. All fluorescence images were adjusted to have the same brightness and contrast settings.

Construction of a Light-Driven Biomimetic Hybrid Nanoreactor for Hydrogen Production

Given the unique properties of TBAP-α for light absorption and electron–proton transfer (35) as well as the strong binding between TBAP-α crystals and α-carboxysome shells, we hypothesized that the C–S|TBAP-α hybrid may serve as a novel biomimetic photocatalyst enabling electron transfer cascade and enhanced enzymatic performance in biotechnological applications. Intriguingly, sulfate ions can be soaked into the major shell hexamers of both α- and β-carboxysomes, (43,44) indicating that the pores may act as the conduits for negatively charged metabolites. Furthermore, iron–sulfur (Fe–S) clusters have been found in catabolic BMCs, (45−48) which could potentially facilitate the transfer of electrons and protons from external TBAP-α crystals across the shell to retain the internal redox environment. (49,50)
To test the hypothesis, we chose the α-carboxysome shells that encase the oxygen-sensitive redox enzymes, hydrogenases, that bind to the TBAP-α crystals. The hydrogenase-containing shells (H–S) were generated by encapsulating [FeFe]-hydrogenase fused with the CsoS2 C-terminus (HydA-EP), ferredoxin (Fd), and ferredoxin: NADP+-oxidoreductase (FNR) into the α-carboxysome shells, along with coexpression of the crucial maturase enzymes HydE, HydF, and HydG (33) (Figure 1c). H–S has been reported to exhibit enhanced H2 evolution and O2 tolerance, taking advantage of the O2-limiting microenvironment created within the α-carboxysome shell. (32,33) The comparable ζ-potentials of empty α-carboxysome shell (S), mCherry-encapsulated α-carboxysome shell (C–S), and [FeFe]-hydrogenase-encapsulated α-carboxysome shell (H–S), as well as the more negative ζ-potentials of S, C–S, and H–S after binding with TBAP-α individually, indicated that binding of the α-carboxysome shells to the TBAP-α surface was not remarkably affected by cargo encapsulation (Figure S13). Electron microscopy further confirmed that the TBAP-α crystals were coated with H–S particles (Figures 4a and S14), consistent with the observations of C–S shells and TBAP-α crystals (Figure 3).

Figure 4

Figure 4. Characterization of the light-driven H–S|TBAP-α hybrid nanoreactor. (a) SEM image of H–S|TBAP-α. Orange arrows show the H–S particles that were attached to the TBAP-α crystal surface. (b) Photoluminescence (PL) spectrum and (c) Excited-state lifetimes of H–S|TBAP-α and TBAP-α alone in 0.1 M neutralized AA aqueous solution when excited at 390 nm.

The H–S|TBAP-α hybrid biocatalyst (Figure 1d) was generated by dispersing freshly purified H–S and TBAP-α in nitrogen-degassed neutralized AA solution (Figures S15 and S16), followed by incubation for 30 min at room temperature. UV–vis spectroscopy was used to examine the colloidal stability of H–S|TBAP-α. No significant changes were detected, even after 19 h incubation (Figure S17). We also employed photoluminescence (PL) spectroscopy to analyze the electron–hole pair recombination performance of H–S|TBAP-α and TBAP-α. The PL emission intensity of H–S|TBAP-α was quenched greatly compared to TBAP-α alone when excited at 390 nm (Figure 4b), indicating that the recombination rate for the photogenerated electron and hole was drastically reduced. This is beneficial for the transfer of electrons to H–S for the H2 production reaction. In addition, time-correlated single-photon counting (TCSPC) showed that the average weighted lifetime of H–S|TBAP-α (τavg = 4.11 ns) was greater than that of TBAP-α (τavg = 3.51 ns) in an aqueous suspension (Table S4), suggesting that more electrons were transferred to H–S rather than recombining with holes. Moreover, TBAP-α in aqueous suspension exhibited a shorter lifetime (τavg = 2.00 ns), (35) which aligns with its remarkably lower H2 production performance, even when hybridized with H–S (0.79 mmol (g TBAP)−1 h–1, Figure 5a). Therefore, the PL quench and the extended lifetime of H–S|TBAP-α indicate that H–S facilitated the separation of electron–hole pairs.

Figure 5

Figure 5. Visible-light-driven hydrogen production of H–S|TBAP-α. (a) H2-evolution rates of TBAP-α, H–S|TBAP-α, H–Sox|TBAP-α, S|TBAP-α, and H–S|Amorphous TBAP irradiated by a solar simulator for 2 h. (b) H2 evolution of TBAP-α (black dots) and H–S|TBAP-α (blue dots) as a function of time (λ > 420 nm). (c) Schematic diagram of proposed electron transfer pathway in H–S|TBAP-α for hydrogen production. Electrons generated from TBAP-α irradiation are transferred through holes on the α-carboxysome shell to the 4Fe–4S cluster via FNR or directly for proton reduction. Ascorbic acid acts as a sacrificial agent to prevent the photogenerated hole–electron recombination. Error bars represent the SD of the mean of three independent experiments.

The electron transfer performance of TBAP-α and H–S|TBAP-α was characterized by transient photocurrent (TPC) and electrochemical impedance spectroscopy (EIS). The transient photocurrent intensity of H–S|TBAP-α reached a value that was 30% higher than that of TBAP-α alone (Figure S18a), indicating that photogenerated electrons and holes could be separated and transferred effectively to H–S in H–S|TBAP-α. The arc radius of EIS plots varied in the order H–S|TBAP-α < TBAP-α (Figure S18b), suggesting that H–S|TBAP-α showed lower electrical resistance than the TBAP-α crystalline, likely through enhanced charge transport. These assays confirmed the effective construction of the biomimetic nanoreactor with accelerated electron transfer and promising photocatalytic potential.
High-throughput sacrificial H2 evolution measurements revealed that the mass-normalized H2 evolution rate of H–S|TBAP-α (approximately 25 wt % of H–S) was 3.97 ± 0.41 mmol (g TBAP-α)−1 h–1 during 2 h irradiation in nitrogen-degassed neutralized AA aqueous solution (0.1 M) using a solar simulator (AM1.5G, 1440 W xenon, see Methods for details). The rate is almost 5 times higher than that obtained from TBAP-α alone under the same conditions (0.79 ± 0.01 mmol (g TBAP-α)−1 h–1; Figure 5a). The H2-evolution activity of TBAP-α alone is associated with the residual palladium present in the sample, which originates from the synthesis of TBAP and acts as a cocatalyst. (35) When empty shell (S) and deactivated H–S with oxygen bubbling (H–Sox) (the deactivation of H–Sox was confirmed by sodium dithionite-methyl viologen assays shown in Figure S19a) were mixed with TBAP-α respectively, the H2-evolution rates of both the resulting S|TBAP-α and H–Sox|TBAP-α declined significantly (0.78 ± 0.15 mmol (g TBAP-α)−1 h–1 for S|TBAP-α, 0.76 ± 0.06 mmol (g TBAP-α)−1 h–1 for H–Sox|TBAP-α), which were comparable to the amount of H2 produced by TBAP-α alone (Figure 5a). These results indicated that H–S in the H–S|TBAP-α hybrid nanoreactor was enzymatically active. Furthermore, no detectable amounts of H2 were produced from H–S|TBAP-α in a dark suspension, an irradiated suspension of H–S in a neutralized AA aqueous solution (0.1 M), or an irradiated suspension of H–S|TBAP-α in pure water over 2 h. Together, these control experiments indicate that TBAP-α donated photogenerated electrons for proton reduction and AA acted effectively as the hole scavenger (Figure S19 and Table S6). When TBAP-α was associated with the α-carboxysome shells that encapsulate only hydrogenases without FNR and Fd (HydA-S), there was a 33% decrease in the H2-evolution rate compared to H–S|TBAP-α (2.68 ± 0.45 mmol (g TBAP-α)−1 h–1 for HydA-S|TBAP-α, 3.97 ± 0.41 mmol (g TBAP-α)−1 h–1 for H–S|TBAP-α) (Figure S19d). These results suggest that FNR and Fd indeed contributed to the transport of photogenerated electrons to the active sites of hydrogenases.
One advantage of crystalline photocatalysts is their ability to accelerate electron transfer through a highly ordered packing pattern. This can be verified by a larger photocurrent density and smaller EIS radius from TBAP-α compared to amorphous TBAP (Figure S20). This observation coincides with the substantially higher H2 produced when H–S was hybridized with TBAP-α compared with an amorphous powdered TBAP sample. Specifically, the crystalline TBAP-α produced 3.97 ± 0.41 mmol–1 (g TBAP-α)−1 h–1, while the amorphous TBAP sample yielded only 0.79 ± 0.15 mmol–1 (g TBAP)−1 h–1 under the same conditions (Figure 5a). Hence, the synergy between H–S and TBAP-α in the hierarchical H–S|TBAP-α assemblies appears to be critical for the attached H–S structures, ensuring electron transfer between the two components and enhanced hydrogen evolution performance. The apparent quantum efficiency (AQY) was measured at different wavelengths to evaluate the photocatalytic H2 production performance of H–S|TBAP-α. The AQY measured at 420 nm was 2.8% over a 1.5 h experiment (Figure S21).
To examine the H2-evolution performance of H–S|TBAP-α in long-term irradiation, the H–S|TBAP-α hybrid nanoreactors were exposed to visible light (300 W Xe light, λ > 420 nm). As depicted in Figure 5b, H–S|TBAP-α exhibited a constant increase in H2 production as a function of time. Under the same conditions, H–S|TBAP-α produced an H2 efficiency that was roughly comparable to the noble metal-loaded system as reference, namely, 1 wt % platinum metal cocatalyst deposited on TBAP-α (Pt|TBAP-α), after 14 h of irradiation (144.3 μmol for H–S|TBAP-α, 170.2 μmol for Pt|TBAP-α) (Figure S22). In contrast, TBAP-α, without the addition of platinum cocatalyst or H–S, produced only 31.9 μmol of H2 after 14 h irradiation (Figure S22). The H2-production ability of H–S|TBAP-α ahowed a relatively steady increase as a function of time for 30 h irradiation, with an H2-evolution rate of 1,137.8 ± 47.2 μmol (g TBAP-α)−1 h–1. Intriguingly, H–S|TBAP-α exhibited remarkable stability during 5 consecutive runs over a total reaction time of 25 h, with no notable decline in the H2-production rate observed after the 5 cycles (Figure S23). From the listed state-of-the-art photocatalyst-hydrogenase hybrids (Table S7), H–S|TBAP-α has the unique advantage of considerable stability with a promising H2 production efficiency. In the long-term H2 evolution experiments, the HydA content was measured using purified HydA as a reference (Figure S24), and the total turnover number (TON) of H–S|TBAP-α was measured as 5,411,174 mol H2 (mol H2ase)−1 on average after 24-h irradiation (Xe lamp, λ > 420 nm), which is comparable to the reported organic materials-hydrogenase hybrids (Table S7). SEM was performed to examine the morphology changes of H–S|TBAP-α after 30 h of continuous irradiation. The attachment of H–S nanoparticles on the TBAP-α surface was not as obvious as before the illumination (Figure S25). Meanwhile, cracks appeared on the crystal surface, which is consistent with the crystallinity decrease found in the PXRD pattern (Figure S26).
Overall, our findings demonstrate the catalytic efficiency and robustness of H–S|TBAP-α under irradiation and the fact that the hybrid H–S|TBAP-α system has an enhanced photocatalytic H2-evolution capacity relative to the organic semiconductor TBAP-α crystal or H–S alone. We speculate that the enhanced photocatalytic H2-evolution performance of H–S|TBAP-α is partially due to accelerated electron flux from TBAP-α, through the ion-containing pores of shell protein complexes at the biotic–abiotic interface, to the catalytically active centers ([4Fe–4S] cluster) of encapsulated [FeFe]-hydrogenases, via FNR/Fd proteins or directly (Figure 5c).

Conclusions

Click to copy section linkSection link copied!

We developed a bottom-up strategy for biomimetic nanoreactor construction based on the inherent structural properties of biological machinery and synthetic organic materials. A crystalline HOF, TBAP-α, acts as a visible-light-capture organic antenna and can funnel energy into a complex, multicomponent biocatalytic nanoreactor, H–S, which mimics the natural photosynthesis process. This hybrid nanoreactor performs as a light-driven photocatalyst for sacrificial H2 evolution in water, and it exhibits improved activity than H–S or TBAP-α alone and comparable activity to the TBAP-α material deposited with 1 wt % of the precious metal cocatalyst, platinum. Our study suggests that H–S|TBAP-α is attributed to the separation of the photogenerated electron and hole pairs at the TBAP-α surface and the transport of electrons into the nanoreactor interior, which enhances the catalytic performance of encapsulated hydrogenases for the production of molecular hydrogen. The construction of H–S|TBAP-α with considerable hydrogen production activity acts as a promising model system to explore the potential of combining catalytic advantages from both artificial materials and natural machinery for solar energy conversion.
The successful assembly of complex, hierarchical structured nanoreactors using multiple noncovalent hydrogen-bonding interactions on organic semiconductor surfaces opens up avenues for designing new families of chemical-biologically active hybrid systems for reactions other than H2 production. Here, we demonstrated the generality and versatility of our approach, which utilizes noncovalent hydrogen-bonding interactions between multiple proteins and the organic semiconductor surface. We coated the TBAP-α crystal surface with three different proteins, C–S, H–S, and H–Sox, each with distinct catalytic properties. We postulate that this approach could be adapted to create biocatalytic nanoreactors for more complex chemical transformations by altering the cargo enzymes within the α-carboxysome shell.

Methods

Click to copy section linkSection link copied!

Generation of Constructs

All assemblies between genes and linearized vectors were achieved by Gibson assembly (Gibson assembly kit, New England BioLabs, U.K.). The mCherry gene was cloned to pCDFDueT-1 linearized by EcoRI and AscI to generate the pCDFDueT-mCherry vector. The mCherry gene with the nucleotide sequence encoding EP fused at the C-terminus was cloned into pCDFDueT-1 and in the frame to create pCDFDueT-mCherry-EP, using primer pairs M-F/M-C-R and M-C-F/C-R (Table S8). The mCherry gene fused to the N-termini of these truncated CsoS2 C-terminal regions was inserted into pCDFDueT linearized by BamHI and AscI. These constructs were verified by PCR and DNA sequencing and transformed into Escherichia coli BL21(DE3) cells. Plasmid maps (Figure S27) and sequences (Table S9) are available in the Supporting Information.

Coexpression and Generation of α-Carboxysome Encapsulated with mCherry (C–S)

E. coli strains containing the pCDTDuet-mCherry-CS2 plasmid and pBAD-cso-2 plasmid were cultivated in LB medium containing 50 μg mL–1 of spectinomycin and 100 μg mL–1 of ampicillin. The mCherry expression was induced by adding 0.5 mM isopropyl β-d-1-thiogalactopyranoside (IPTG) at OD600 = 0.6–0.8. After 4 h induction of the mCherry-EP expression, the shell expression was induced by 1 mM l-Arabinose, and cells were then grown at 25 °C for 16 h with constant shaking. The mCherry induction was performed before the expression of shells to allow mCherry protein expression before shell encapsulation. The cells were then harvested by centrifugation at 5000g for 10 min. The cell pellets were washed with TN buffer (20 mM Tris-HCl pH = 8.0, 150 mM NaCl) and resuspended in TN buffer supplemented with 10% (v/v) CelLytic B cell Lysis reagent (Sigma-Aldrich) and 1% Protein Inhibitor Cocktail (100×) (Sigma-Aldrich). The cell suspensions were lysed by French Press (Stansted Fluid Power, U.K.). Cell debris was removed by centrifugation at 10,000g at 4 °C for 10 min, followed by centrifugation at 50,000g to enrich C–S at 4 °C. The pellets were resuspended in TN buffer and then loaded onto sucrose gradients (10–30% or 10–50%, w/v), followed by ultracentrifugation (Beckman, XL100K Ultracentrifuge) at 105,000g for 30 min. Each sucrose fraction was collected and stored at 4 °C. Electron microscopy, confocal microscopy, SDS-PAGE, and immunoblot analysis were performed, as described in the Supporting Information.

Expression of Mature [FeFe]-Hydrogenase and Generation of α-Carboxysome Shells with Encapsulated Hydrogenases (H–S)

[FeFe]-hydrogenase from the green alga Chlamydomonas reinhardtii was expressed and encapsulated into α-carboxysome shells (H–S) in E. coli, as reported before. (24) In summary, E. coli strains containing the pCDFDuet-hydA-CS2 (GenBank accession code AAL23572.1) plasmid and the pBAD-cso-2 plasmid were cultivated in LB medium containing 0.2 mM ferric ammonium citrate, 50 μg mL–1 spectinomycin, and 100 μg mL–1 ampicillin and degassed with N2 for 30 min at OD600 = 0.6–0.8. The hydA-EP expression was induced by the addition of 0.5 mM IPTG. After 4 h induction of the hydA-EP expression, the shell expression was induced by 1 mM l-Arabinose, and cells were then grown at 25 °C for 16 h. The purification of H–S was carried out following the method for C–S mentioned above. It should be noted that H–S should be expressed and purified under anaerobic conditions, and all of the solutions used should be degassed in advance. As a control, the plasmid for expressing hydrogenases without FNR and Fd (pCDFDuet-hydA-hydGxEF-CS2) was generated by deleting both genes encoding FNR and Fd from the pCDFDuet-hydA-CS2 plasmid. The α-carboxysome shells that encapsulate hydrogenases only without FNR and Fd (HydA-S) were generated following the same procedure as that for H–S.

TBAP-α Construction

Synthesis and characterization of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP) are described in detail in the Supporting Information. As-synthesized TBAP powder (100 mg) was dissolved in DMF (8 mL). The supernatant solutions were filtered into 25 mL vials using a 0.22 μm PTFE syringe filter to remove any particulates and then placed into larger 100 mL vials with antisolvent CHCl3. Vapor diffusion of the antisolvent into the TBAP solution was carried out for several days until the vials were nearly full of solvent. The resulting crystals were activated after the crystallization solvent was exchanged with acetone 10–12 times over a few days. The crystals were then collected by filtration and characterized by PXRD.

Generation of H–S|TBAP-α Hybrid Nanoreactors

A 2 mg portion of TBAP-α and neutralized AA aqueous solution (0.1 M, 4.5 mL) were added into sample vials and purged with N2 to eliminate oxygen in the solution and sample vials. Next, 0.5 mL of H–S solution purified by a sugar gradient strategy (20% sucrose content, thereby diluting the total protein concentration to 1 mg mL–1 as measured via the Nanodrop method) was introduced into the sample vials under an N2 gas flow. The mixture solution was incubated on a rotator at room temperature for 30 min to facilitate the association of H–S on the TBAP-α surface. The H–S|TBAP-α construction was confirmed by SEM and photocatalytic H2 evolution performance.

High-Throughput Photocatalytic H2 Production Experiments

2 mg of TBAP-α or amorphous TBAP powder and neutralized AA aqueous solution (0.1 M, 4.5 mL) were added into separate sample vials (vial size = 12.5 mL) and purged with N2 on a Chemspeed Technologies Sweigher platform for 6 h. Next, the purified H–S solution (0.5 mL in 20% sucrose, 1 mg mL–1 measured using the Nanodrop method) was added under N2 gas flow to make the total volume of 5 mL. For control samples, 0.5 mL of TN buffer was added to the vials to prepare solutions with a total volume of 5 mL.
All sample vials were irradiated using an Oriel Solar Simulator with an output of 1.0 sun (AM1.5G, Class AAA, IEC/JIS/ASTM, 1440 W xenon, 12 × 12 in., MODEL: 94123A) equipped with a rocker/roller device. The vial size was 23 × 46 mm2, and the irradiation area on the samples was 16.61 cm2. Gaseous products were analyzed on a Shimadzu GC-2010 equipped with a Shimadzu HS-20, injecting a sample from the headspace sampler via a transfer line (temperature 150 °C) onto an Rt-Msieve 5 Å column with He as the carrier gas at a flow rate of 30 mL min–1. H2 was detected with a barrier discharge ionization detector and referenced against a standard that contained a known concentration of H2. Data are represented as the mean ± standard deviation (SD) from at least three repeated measurements.

Time-Course H2 Evolution Assays

Time-course H2 evolution measurements were conducted in a 67 mL quartz flask. 10 mg of TBAP-α and 22.5 mL of neutralized AA aqueous solution (0.1 M) were added to the quartz flask and purged with N2 for 30 min, followed by the addition of purified H–S solution (2.5 mL in 20% sucrose, 1 mg mL–1 measured using the Nanodrop method) under N2 gas flow to make the final volume of 25 mL. The reaction mixture was illuminated with a 300 W Newport Xe light source (Model: 6258, Ozone-free) using a λ > 420 nm cutoff filter. The light source was cooled by water circulating through a metal jacket. Gas samples were taken with a gastight syringe and run on a Bruker 450-GC gas chromatograph. H2 was detected with a thermal conductivity detector referencing standard gas with a known concentration of H2. Data are represented as mean ± SD from at least three repeated measurements.

Molecular Dynamics (MD) Simulations

Model Setup

The TBAP-α model (1 row, 5 columns) was obtained from the published data, (35) and the structure of the CsoS1A protein was downloaded from the Protein Data Bank (PDB ID: 2EWH). (44) The CsoS1A was docked to the TBAP-α model using the HDOCK Server (http://hdock.phys.hust.edu.cn/). (51) The structures with the highest scores were manually screened. The best-fit conformation was selected for molecular dynamic simulation using the AMBER20 software package. (52) The ff14SB (53) force field generated the protein parameters. The Generalized Amber Force field (GAFF) force field (54) is used to create the force field of the TBAP molecule. The atomic charges of the TBAP molecule were calculated by the restricted electrostatic potential (RESP) method with B3LYP/6-31G(d) (55,56) level of theory by the Gaussian 09 program. After examining the protonation state of the protein and adding hydrogen atoms, the system was solvated in a truncated octahedral box of the TIP3P water model (57) with a buffer of 35.0 Å. Na+ and Cl (58) were added to neutralize the system and achieve a physiological salt concentration of 150 mM. The resulting system comprised 465,869 atoms.

MD Simulations

The system was minimized by 1000 steps of the fastest descent algorithm and 1000 steps of the conjugation gradient algorithm. Then, the system was heated to 300 K in 1 ns, and 300 ns of simulations were performed with 40 ns of NVT simulations and 260 ns of NPT simulation. The electrostatic interactions were calculated using the particle mesh Ewald (PME) (59) method, and the periodic boundary conditions were used. The Langevin thermostat method (60) is used to control the temperature, and the Berendsen barostat method (61) was used to control the system’s pressure. To mimic the environment of carboxysomes, a restraint of 10 kcal mol–1 Å–2 was applied to the backbone of the protein and TBAP-α during the simulation. The trajectory of the last 30 ns was selected as a production run for analysis.

Analysis of Simulations

The root-mean-square deviation (RMSD) of the trajectory of molecular dynamics simulations is calculated by the RMSD Trajectory Tool in VMD. (62) The hydrogen bond and salt bridge analysis used the PLIP web tool (57) (https://plip-tool.biotec.tu-dresden.de/plip-web/plip/index). The maximum distance of the hydrogen bond was set to 4.1 Å, and the minimum angle of a hydrogen bond was 100°. The cutoff of the salt bridge interaction is 5.5 Å.

Supporting Information

Click to copy section linkSection link copied!

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acscatal.4c03672.

  • Chemical synthesis pathway of TBAP (Scheme S1); NMR spectrum of TBAP (Figures S1 and S2); FI-IR spectrum of TBAP (Figure S3); crystal model of TBAP-α and 3D model of α-carboxysome shell (Figure S4); PXRD patterns of TBAP phases, nitrogen adsorption isotherm and desorption isotherm for activated TBAP-α and amorphous TBAP, and SEM images of amorphous TBAP (Figure S5); solid UV–vis absorption spectrum of amorphous TBAP and TBAP-α (Figure S6); (αhν)1/2 versus hν curve and Mott–Schottky plot, diagram of conduction band and valence band of amorphous TBAP and TBAP-α, and cyclic voltammetry plot of TBAP-α (Figure S7); amino acid sequence alignment of α-carboxysome shell proteins (Figure S8); front and side views of the structures of α-carboxysome shell proteins (Figure S9); confocal microscopy images of E. coli cells expressing mCherry-CsoS2C (mCherry-EP) and coexpressing shells and mCherry-csoS2C (C–S) and SDS-PAGE of purified C–S (Figure S10); TEM image of C–S (Figure S11); SEM and confocal microscopy images of pyrene crystals and pyrene crystals with C–S (Figure S12); ζ-potentials of TBAP-α and α-carboxysome shell-encasing proteins (Figure S13); size distribution of H–S revealed by SEM (Figure S14); SDS-PAGE result of H–S purification (Figure S15); immunoblot analysis of purified H–S (Figure S16); colloidal stability of H–S|TBAP-α (Figure S17); photocurrent responses and EIS analysis for TBAP-α and H–S|TBAP-α (Figure S18); H2 production condition optimization of H–S|TBAP-α (Figure S19); photocurrent response and EIS analysis for amorphous TBAP and TBAP-α (Figure S20); wavelength-dependent AQY value of H–S|TBAP-α (Figure S21); H2 evolution of TBAP-α, H–S|TBAP-α, and 1 wt % Pt|TBAP-α (λ > 420 nm) as a function of time (Figure S22); cycling measurements for the photocatalytic hydrogen evolution of H–S|TBAP-α (Figure S23); immunoblot analysis of purified HydA (Fd-HydA-EP) (Figure S24); SEM images (Figure S25) and PXRD pattern (Figure S26) of H–S|TBAP-α after 30 h irradiation; gene maps of used plasmids (Figure S27); crystal data and structure refinement of TBAP-α (Table S1); protein components in recombinant α-carboxysomes from E. coli (Table S2); statistics of residues for TBAP-α and CsoS1A protein binding calculated by MD simulations (Tables S3 and S4); estimated fluorescence lifetimes of TBAP-α and H–S|TBAP-α (Table S5); a list of hydrogen evolution reaction conditions in this work (Table S6); a comparison of the H–S|TBAP-α assembly performance with the related state-of-the-art photocatalysts (Table S7); primers used for pCDFDuet-mCherry-CS2 plasmid construction (Table S8); and gene sequences of used plasmids (Table S9). (PDF)

Terms & Conditions

Most electronic Supporting Information files are available without a subscription to ACS Web Editions. Such files may be downloaded by article for research use (if there is a public use license linked to the relevant article, that license may permit other uses). Permission may be obtained from ACS for other uses through requests via the RightsLink permission system: http://pubs.acs.org/page/copyright/permissions.html.

Author Information

Click to copy section linkSection link copied!

  • Corresponding Authors
  • Authors
    • Jing Yang - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    • Qiuyao Jiang - Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    • Yu Chen - Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    • Quan Wen - Hubei Key Laboratory of Agricultural Bioinformatics, College of Informatics, Huazhong Agricultural University, Wuhan 430070, China
    • Xingwu Ge - Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    • Qiang Zhu - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Wei Zhao - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.Orcidhttps://orcid.org/0000-0003-0265-2590
    • Oluwatobi Adegbite - Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool L69 7ZB, U.K.
    • Haofan Yang - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Liang Luo - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Hang Qu - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Veronica Del-Angel-Hernandez - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Rob Clowes - Materials Innovation Factory and Department of Chemistry, University of Liverpool, Liverpool L7 3NY, U.K.
    • Jun Gao - Hubei Key Laboratory of Agricultural Bioinformatics, College of Informatics, Huazhong Agricultural University, Wuhan 430070, ChinaOrcidhttps://orcid.org/0000-0002-6692-1828
  • Author Contributions

    L.-N.L. and A.I.C. devised and supervised the project. M.A.L. helped supervise the project and contributed to the design of experiments and data interpretation. J.Y. designed and carried out the experiments and led the experimental work. Q.J. assisted with the construction of protein. X.G. helped with confocal microscopy. Y.C. assisted with plasmid design and preparation. Q.W. and J.G. performed molecular dynamics simulations and analysis. Q.Z. and H.Q. helped with PXRD and gas absorption measurements. W.Z., L.L., H.Y., and V.D.-A.-H. assisted with photoelectrochemical measurements. O.A. helped with the ITC measurement. R.C. assisted with high-throughput photocatalytic H2 production experiments. J.Y., M.A.L., A.I.C., and L.-N.L. wrote the manuscript with all authors’ contributions.

  • Notes
    The authors declare no competing financial interest.

Acknowledgments

Click to copy section linkSection link copied!

The authors thank the Liverpool Biomedical Electron Microscopy Unit and Centre for Cell Imaging for technical assistance and provision for microscopic imaging and the Materials Innovation Factory (MIF) for the provision of analytical equipment. This work was supported by the National Key R&D Program of China (2021YFA0909600), the National Natural Science Foundation of China (32070109), the Biotechnology and Biological Sciences Research Council (BBSRC) (BB/Y008308/1, BB/Y01135X/1), the Royal Society (URF\R\180030, RGF\EA\181061, RGF\EA\180233), and the Leverhulme Trust (RPG-2021-286). The authors acknowledge financial support from the Leverhulme Trust via the Leverhulme Research Centre for Functional Materials Design.

References

Click to copy section linkSection link copied!

This article references 62 other publications.

  1. 1
    Tachibana, Y.; Vayssieres, L.; Durrant, J. R. Artificial photosynthesis for solar water-splitting. Nat. Photonics 2012, 6, 511518,  DOI: 10.1038/nphoton.2012.175
  2. 2
    Blankenship, R. E.; Tiede, D. M.; Barber, J.; Brudvig, G. W.; Fleming, G.; Ghirardi, M.; Gunner, M. R.; Junge, W.; Kramer, D. M.; Melis, A.; Moore, T. A.; Moser, C. C.; Nocera, D. G.; Nozik, A. J.; Ort, D. R.; Parson, W. W.; Prince, R. C.; Sayre, R. T. Comparing Photosynthetic and Photovoltaic Efficiencies and Recognizing the Potential for Improvement. Science 2011, 332, 805809,  DOI: 10.1126/science.1200165
  3. 3
    Wang, Y.; Vogel, A.; Sachs, M.; Sprick, R. S.; Wilbraham, L.; Moniz, S. J. A.; Godin, R.; Zwijnenburg, M. A.; Durrant, J. R.; Cooper, A. I.; Tang, J. Current understanding and challenges of solar-driven hydrogen generation using polymeric photocatalysts. Nat. Energy 2019, 4, 746760,  DOI: 10.1038/s41560-019-0456-5
  4. 4
    Barber, J. Photosynthetic energy conversion: Natural and artificial. Chem. Soc. Rev. 2009, 38, 185196,  DOI: 10.1039/B802262N
  5. 5
    Zhang, J. Z.; Reisner, E. Advancing photosystem II photoelectrochemistry for semi-artificial photosynthesis. Nat. Rev. Chem. 2020, 4, 621,  DOI: 10.1038/s41570-019-0149-4
  6. 6
    Pi, X.; Zhao, S.; Wang, W.; Liu, D.; Xu, C.; Han, G.; Kuang, T.; Sui, S. F.; Shen, J. R. The pigment-protein network of a diatom photosystem II–light-harvesting antenna supercomplex. Science 2019, 365, 447457,  DOI: 10.1126/science.aax4406
  7. 7
    Vayghan, H. S.; Nawrocki, W. J.; Schiphorst, C.; Tolleter, D.; Hu, C.; Douet, V.; Glauser, G.; Finazzi, G.; Croce, R.; Wientjes, E.; Longoni, F. Photosynthetic Light Harvesting and Thylakoid Organization in a CRISPR/Cas9 Arabidopsis Thaliana LHCB1 Knockout Mutant. Front. Plant Sci. 2022, 13, 833032833050,  DOI: 10.3389/fpls.2022.833032
  8. 8
    Koepf, M.; Teillout, A.-L.; Llansola-Portoles, M. J. Artificial Photosynthesis: An Approach for a Sustainable Future. In Handbook of Ecomaterials; Martínez, L. M. T.; Kharissova, O. V.; Kharisov, B. I., Eds.; Springer International Publishing: Cham, 2017; pp 125.
  9. 9
    Guo, Y.; Zhou, Q.; Nan, J.; Shi, W.; Cui, F.; Zhu, Y. Perylenetetracarboxylic acid nanosheets with internal electric fields and anisotropic charge migration for photocatalytic hydrogen evolution. Nat. Commun. 2022, 13, 2067  DOI: 10.1038/s41467-022-29826-z
  10. 10
    Kosco, J.; Bidwell, M.; Cha, H.; Martin, T.; Howells, C. T.; Sachs, M.; Anjum, D. H.; Gonzalez Lopez, S.; Zou, L.; Wadsworth, A.; Zhang, W.; Zhang, L.; Tellam, J.; Sougrat, R.; Laquai, F.; DeLongchamp, D. M.; Durrant, J. R.; McCulloch, I. Enhanced photocatalytic hydrogen evolution from organic semiconductor heterojunction nanoparticles. Nat. Mater. 2020, 19, 559565,  DOI: 10.1038/s41563-019-0591-1
  11. 11
    Wang, Q.; Domen, K. Particulate Photocatalysts for Light-Driven Water Splitting: Mechanisms, Challenges, and Design Strategies. Chem. Rev. 2020, 120, 919985,  DOI: 10.1021/acs.chemrev.9b00201
  12. 12
    Takata, T.; Jiang, J.; Sakata, Y.; Nakabayashi, M.; Shibata, N.; Nandal, V.; Seki, K.; Hisatomi, T.; Domen, K. Photocatalytic water splitting with a quantum efficiency of almost unity. Nature 2020, 581, 411414,  DOI: 10.1038/s41586-020-2278-9
  13. 13
    Searle, N. Z.; Hirt, R. C. Ultraviolet Spectral Energy Distribution of Sunlight. J. Opt. Soc. Am. 1965, 55, 14131421,  DOI: 10.1364/JOSA.55.001413
  14. 14
    Cestellos-Blanco, S.; Zhang, H.; Kim, J. M.; Shen, Yx.; Yang, P. Photosynthetic semiconductor biohybrids for solar-driven biocatalysis. Nat. Catal. 2020, 3, 245255,  DOI: 10.1038/s41929-020-0428-y
  15. 15
    Kornienko, N.; Zhang, J. Z.; Sakimoto, K. K.; Yang, P.; Reisner, E. Interfacing nature’s catalytic machinery with synthetic materials for semi-artificial photosynthesis. Nat. Nanotechnol. 2018, 13, 890899,  DOI: 10.1038/s41565-018-0251-7
  16. 16
    Özgen, F. F.; Runda, M. E.; Schmidt, S. Photo-biocatalytic Cascades: Combining Chemical and Enzymatic Transformations Fueled by Light. ChemBioChem 2021, 22, 790806,  DOI: 10.1002/cbic.202000587
  17. 17
    Schmermund, L.; Jurkaš, V.; Özgen, F. F.; Barone, G. D.; Büchsenschütz, H. C.; Winkler, C. K.; Schmidt, S.; Kourist, R.; Kroutil, W. Photo-Biocatalysis: Biotransformations in the Presence of Light. ACS Catal. 2019, 9, 41154144,  DOI: 10.1021/acscatal.9b00656
  18. 18
    Holá, K.; Pavliuk, M. V.; Németh, B.; Huang, P.; Zdražil, L.; Land, H.; Berggren, G.; Tian, H. Carbon Dots and [FeFe] Hydrogenase Biohybrid Assemblies for Efficient Light-Driven Hydrogen Evolution. ACS Catal. 2020, 10, 99439952,  DOI: 10.1021/acscatal.0c02474
  19. 19
    Gai, P.; Yu, W.; Zhao, H.; Qi, R.; Li, F.; Liu, L.; Lv, F.; Wang, S. Solar-Powered Organic Semiconductor-Bacteria Biohybrids for CO2 Reduction into Acetic Acid. Angew. Chem., Int. Ed. 2020, 59, 72247229,  DOI: 10.1002/anie.202001047
  20. 20
    Wang, X.; Saba, T.; Yiu, H. H. P.; Howe, R. F.; Anderson, J. A.; Shi, J. Cofactor NAD(P)H Regeneration Inspired by Heterogeneous Pathways. Chem 2017, 2, 621654,  DOI: 10.1016/j.chempr.2017.04.009
  21. 21
    Gentil, S.; Che Mansor, S. M.; Jamet, H.; Cosnier, S.; Cavazza, C.; Le Goff, A. Oriented Immobilization of [NiFeSe] Hydrogenases on Covalently and Noncovalently Functionalized Carbon Nanotubes for H2/Air Enzymatic Fuel Cells. ACS Catal. 2018, 8, 39573964,  DOI: 10.1021/acscatal.8b00708
  22. 22
    Zhang, S.; Liu, S.; Sun, Y.; Li, S.; Shi, J.; Jiang, Z. Enzyme-photo-coupled catalytic systems. Chem. Soc. Rev. 2021, 50, 1344913466,  DOI: 10.1039/D1CS00392E
  23. 23
    Sun, Y.; Lin, Y.; Harman, V. M.; Beynon, R. J.; Johnson, J. R.; Liu, L.-N. Decoding the Absolute Stoichiometric Composition and Structural Plasticity of a-Carboxysomes. mBio 2022, 13, e03629-21  DOI: 10.1128/mbio.03629-21
  24. 24
    Gonzalez-Esquer, C. R.; Newnham, S. E.; Kerfeld, C. A. Bacterial microcompartments as metabolic modules for plant synthetic biology. Plant J. 2016, 87, 6675,  DOI: 10.1111/tpj.13166
  25. 25
    Liu, L. N. Advances in the bacterial organelles for CO2 fixation. Trends Microbiol. 2022, 30, 567580,  DOI: 10.1016/j.tim.2021.10.004
  26. 26
    Huang, J.; Jiang, Q.; Yang, M.; Dykes, G. F.; Weetman, S. L.; Xin, W.; He, H. L.; Liu, L. N. Probing the internal pH and permeability of a carboxysome shell. Biomacromolecules 2022, 23, 43394348,  DOI: 10.1021/acs.biomac.2c00781
  27. 27
    Faulkner, M.; Szabó, I.; Weetman, S. L.; Sicard, F.; Huber, R. G.; Bond, P. J.; Rosta, E.; Liu, L.-N. Molecular simulations unravel the molecular principles that mediate selective permeability of carboxysome shell protein. Sci. Rep. 2020, 10, 17501  DOI: 10.1038/s41598-020-74536-5
  28. 28
    Mahinthichaichan, P.; Morris, D. M.; Wang, Y.; Jensen, G. J.; Tajkhorshid, E. Selective permeability of carboxysome shell pores to anionic molecules. J. Phys. Chem. B 2018, 122, 91109118,  DOI: 10.1021/acs.jpcb.8b06822
  29. 29
    Cammack, R.; Frey, M.; Robson, R. Hydrogen as a Fuel, Learning from Nature; CRC Press, 2001.
  30. 30
    Vignais, P. M.; Billoud, B. Occurrence, Classification, and Biological Function of Hydrogenases: An Overview. Chem. Rev. 2007, 107, 42064272,  DOI: 10.1021/cr050196r
  31. 31
    Lubitz, W.; Ogata, H.; Rudiger, O.; Reijerse, E. Hydrogenases. Chem. Rev. 2014, 114, 40814148,  DOI: 10.1021/cr4005814
  32. 32
    Jiang, Q.; Li, T.; Yang, J.; Aitchison, C. M.; Huang, J.; Chen, Y.; Huang, F.; Wang, Q.; Cooper, A. I.; Liu, L.-N. Synthetic engineering of a new biocatalyst encapsulating [NiFe]-hydrogenases for enhanced hydrogen production. J. Mater. Chem. B 2023, 11, 26842692,  DOI: 10.1039/D2TB02781J
  33. 33
    Li, T.; Jiang, Q.; Huang, J.; Aitchison, C. M.; Huang, F.; Yang, M.; Dykes, G. F.; He, H. L.; Wang, Q.; Sprick, R. S.; Cooper, A. I.; Liu, L. N. Reprogramming bacterial protein organelles as a nanoreactor for hydrogen production. Nat. Commun. 2020, 11, 5448  DOI: 10.1038/s41467-020-19280-0
  34. 34
    Qin, W. K.; Tung, C. H.; Wu, L. Z. Covalent organic framework and hydrogen-bonded organic framework for solar-driven photocatalysis. J. Mater. Chem. A 2023, 11, 1252112538,  DOI: 10.1039/D2TA09375H
  35. 35
    Aitchison, C. M.; Kane, C. M.; McMahon, D. P.; Spackman, P. R.; Pulido, A.; Wang, X.; Wilbraham, L.; Chen, L.; Clowes, R.; Zwijnenburg, M. A.; Sprick, R. S.; Little, M. A.; Day, G. M.; Cooper, A. I. Photocatalytic proton reduction by a computationally identified, molecular hydrogen-bonded framework. J. Mater. Chem. A 2020, 8, 71587170,  DOI: 10.1039/D0TA00219D
  36. 36
    Stylianou, K. C.; Heck, R.; Chong, S. Y.; Bacsa, J.; Jones, J. T. A.; Khimyak, Y. Z.; Bradshaw, D.; Rosseinsky, M. J. A guest-responsive fluorescent 3D microporous metal-organic framework derived from a long-lifetime pyrene core. J. Am. Chem. Soc. 2010, 132, 41194130,  DOI: 10.1021/ja906041f
  37. 37
    Chen, G.; Huang, S.; Shen, Y.; Kou, X.; Ma, X.; Huang, S.; Tong, Q.; Ma, K.; Chen, W.; Wang, P.; Shen, J.; Zhu, F.; Ouyang, G. Protein-directed, hydrogen-bonded biohybrid framework. Chem 2021, 7, 27222742,  DOI: 10.1016/j.chempr.2021.07.003
  38. 38
    Zhou, Q.; Guo, Y.; Zhu, Y. Photocatalytic sacrificial H2 evolution dominated by micropore-confined exciton transfer in hydrogen-bonded organic frameworks. Nat. Catal. 2023, 6, 574584,  DOI: 10.1038/s41929-023-00972-x
  39. 39
    Pellegrin, Y.; Odobel, F. Sacrificial electron donor reagents for solar fuel production. C. R. Chim. 2017, 20, 283295,  DOI: 10.1016/j.crci.2015.11.026
  40. 40
    Baker, S. H.; Lorbach, S. C.; Rodriguez-Buey, M.; Williams, D. S.; Aldrich, H. C.; Shively, J. M. The correlation of the gene csoS2 of the carboxysome operon with two polypeptides of the carboxysome in Thiobacillus neapolitanus. Arch. Microbiol. 1999, 172, 233239,  DOI: 10.1007/s002030050765
  41. 41
    Case, D. A.; Aktulga, H. M.; Belfon, K.; Cerutti, D. S.; Cisneros, G. A.; Cruzeiro, V. W. D.; Forouzesh, N.; Giese, T. J.; Götz, A. W.; Gohlke, H.; Izadi, S.; Kasavajhala, K.; Kaymak, M. C.; King, E.; Kurtzman, T.; Lee, T.-S.; Li, P.; Liu, J.; Luchko, T.; Luo, R.; Manathunga, M.; Machado, M. R.; Nguyen, H. M.; O’Hearn, K. A.; Onufriev, A. V.; Pan, F.; Pantano, S.; Qi, R.; Rahnamoun, A.; Risheh, A.; Schott-Verdugo, S.; Shajan, A.; Swails, J.; Wang, J.; Wei, H.; Wu, X.; Wu, Y.; Zhang, S.; Zhao, S.; Zhu, Q.; Cheatham, T. E., III; Roe, D. R.; Roitberg, A.; Simmerling, C.; York, D. M.; Nagan, M. C.; Merz, K. M., Jr. AmberTools. J. Chem. Inf. Model. 2023, 63, 61836191,  DOI: 10.1021/acs.jcim.3c01153
  42. 42
    Karthi, N.; Venkatachalam, M. Growth and Characterization Novel Organic Nonlinear Optical Crystal of Pyrene. Int. J. Sci. 2013, 1, 812
  43. 43
    Tanaka, S.; Kerfeld, C. A.; Sawaya, M. R.; Cai, F.; Heinhorst, S.; Cannon, G. C.; Yeates, T. O. Atomic-level models of the bacterial carboxysome shell. Science 2008, 319, 10831086,  DOI: 10.1126/science.1151458
  44. 44
    Tsai, Y.; Sawaya, M. R.; Cannon, G. C.; Cai, F.; Williams, E. B.; Heinhorst, S.; Kerfeld, C. A.; Yeates, T. O. Structural analysis of CsoS1A and the protein shell of the Halothiobacillus neapolitanus carboxysome. PLoS Biol. 2007, 5, e144,  DOI: 10.1371/journal.pbio.0050144
  45. 45
    Parsons, J. B.; Dinesh, S. D.; Deery, E.; Leech, H. K.; Brindley, A. A.; Heldt, D.; Frank, S.; Smales, C. M.; Lünsdorf, H.; Rambach, A.; Gass, M. H.; Bleloch, A.; McClean, K. J.; Munro, A. W.; Rigby, S. E. J.; Warren, M. J.; Prentice, M. B. Biochemical and Structural Insights into Bacterial Organelle Form and Biogenesis. J. Biol. Chem. 2008, 283, 1436614375,  DOI: 10.1074/jbc.M709214200
  46. 46
    Parsons, J. B.; Lawrence, A. D.; McLean, K. J.; Munro, A. W.; Rigby, S. E. J.; Warren, M. J. Characterisation of PduS, the pdu Metabolosome Corrin Reductase, and Evidence of Substructural Organisation within the Bacterial Microcompartment. PLoS One 2010, 5, e14009,  DOI: 10.1371/journal.pone.0014009
  47. 47
    Silva, D. A.; Yu, S.; Ulge, U. Y.; Spangler, J. B.; Jude, K. M.; Labão-Almeida, C.; Ali, L. R.; Quijano-Rubio, A.; Ruterbusch, M.; Leung, I.; Biary, T.; Crowley, S. J.; Marcos, E.; Walkey, C. D.; Weitzner, B. D.; Pardo-Avila, F.; Castellanos, J.; Carter, L.; Stewart, L.; Riddell, S. R.; Pepper, M.; Bernardes, G. J. L.; Dougan, M.; Garcia, K. C.; Baker, D. De novo design of potent and selective mimics of IL-2 and IL-15. Nature 2019, 565, 186191,  DOI: 10.1038/s41586-018-0830-7
  48. 48
    Thompson, M. C.; Wheatley, N. M.; Jorda, J.; Sawaya, M. R.; Gidaniyan, S. D.; Ahmed, H.; Yang, Z.; McCarty, K. N.; Whitelegge, J. P.; Yeates, T. O. Identification of a Unique Fe-S Cluster Binding Site in a Glycyl-Radical Type Microcompartment Shell Protein. J. Mol. Biol. 2014, 426, 32873304,  DOI: 10.1016/j.jmb.2014.07.018
  49. 49
    Zeng, Z.; Boeren, S.; Bhandula, V.; Light, S. H.; Smid, E. J.; Notebaart, R. A.; Abee, T. Bacterial Microcompartments Coupled with Extracellular Electron Transfer Drive the Anaerobic Utilization of Ethanolamine in Listeria monocytogenes. mSystems 2021, 6, e01349-20  DOI: 10.1128/msystems.01349-20
  50. 50
    Ferlez, B.; Markus, S.; Kerfeld, C. A. Glycyl Radical Enzyme-Associated Microcompartments: Redox-Replete Bacterial Organelles. mBio 2019, 10, e02327-18  DOI: 10.1128/mBio.02327-18
  51. 51
    Yan, Y. M.; Tao, H. Y.; He, J. H.; Huang, S.-Y. The HDOCK server for integrated protein–protein docking. Nat. Protoc. 2020, 15, 18291852,  DOI: 10.1038/s41596-020-0312-x
  52. 52
    Case, A. D.; Belfon, K.; Ido, B.-S.; Scott, R. B.; Cerutti, D. S.; Cheatham, T. E., III; Cruzeiro, V. W. D.; Darden, T. A.; Duke, R. E.; Giambasu, G.; Gilson, M. K.; Gohlke, H.; Goetz, A. W.; Harris, R.; Izadi, S.; Izmailov, S. A.; Kasavajhala, K.; Kovalenko, A.; Krasny, R.; Kurtzman, T.; Lee, T. S.; LeGrand, S.; Li, P.; Lin, C.; L, J.; Luchko, T.; Luo, R.; Man, V.; Merz, K. M.; Miao, Y.; Mikhailovskii, O.; Monard, G.; Nguyen, H.; Onufriev, A.; Pan, F.; Pantano, S.; Qi, R.; Roe, D. R.; Roitberg, A.; Sagui, C.; Schott-Verdugo, S.; Shen, J.; Simmerling, C. L.; Skrynnikov, N. R.; Smith, J.; Swails, J.; Walker, R. C.; Wang, J.; Wilson, L.; Wolf, R. M.; Wu, X.; Xiong, Y.; Xue, Y.; D M AMBER 2020; University of California: New York, 2020.
  53. 53
    Maier, J. A.; Martinez, C.; Kasavajhala, K.; Wickstrom, L.; Hauser, K. E.; Simmerling, C. ff14SB: Improving the Accuracy of Protein Side Chain and Backbone Parameters from ff99SB. J. Chem. Theory Comput. 2015, 11, 36963713,  DOI: 10.1021/acs.jctc.5b00255
  54. 54
    Wang, J.; Wolf, R. M.; Caldwell, J. W.; Kollman, P. A.; Case, D. A. Development and testing of a general amber force field. J. Comput. Chem. 2004, 25, 11571174,  DOI: 10.1002/jcc.20035
  55. 55
    Wang, Z.; Li, J.; Liu, J.; Wang, L.; Lu, Y.; Liu, J. P. Molecular insight into the selective binding between human telomere G-quadruplex and a negatively charged stabilizer. Clin. Exp. Pharmacol. Physiol. 2020, 47, 892902,  DOI: 10.1111/1440-1681.13249
  56. 56
    Kim, M.; Kreig, A.; Lee, C. Y.; Rube, H. T.; Calvert, J.; Song, J. S.; Myong, S. Quantitative analysis and prediction of G-quadruplex forming sequences in double-stranded DNA. Nucleic Acids Res. 2016, 44, 48074817,  DOI: 10.1093/nar/gkw272
  57. 57
    Price, D. J.; Charles, L. B. A modified TIP3P water potential for simulation with Ewald summation. J. Chem. Phys. 2004, 121, 1009610103,  DOI: 10.1063/1.1808117
  58. 58
    Li, P. F.; Merz, K. Metal ion modeling using classical mechanics. Chem. Rev. 2017, 117, 15641686,  DOI: 10.1021/acs.chemrev.6b00440
  59. 59
    Cheatham, T. E. I.; Miller, J. L.; Fox, T.; Darden, T. A.; Kollman, P. A. Molecular Dynamics Simulations on Solvated Biomolecular Systems: The Particle Mesh Ewald Method Leads to Stable Trajectories of DNA, RNA, and Proteins. J. Am. Chem. Soc. 1995, 117, 41934194,  DOI: 10.1021/ja00119a045
  60. 60
    Loncharich, R. J.; Brooks, R. B.; Pastor, W. R. Langevin dynamics of peptides: The frictional dependence of isomerization rates of N-acetylalanyl-N′-methylamide. Biopolymers 1992, 32, 523535,  DOI: 10.1002/bip.360320508
  61. 61
    Berendsen, H. J. C.; Postma, J. P. M.; Gunsteren, W. F.; DiNola, A.; Haak, J. R. Molecular dynamics with coupling to an external bath. J. Chem. Phys. 1984, 81, 36843690,  DOI: 10.1063/1.448118
  62. 62
    Humphrey, W.; Dalke, A.; Schulten, K. VMD: Visual Molecular Dynamics. J. Mol. Graphics 1996, 14, 3338,  DOI: 10.1016/0263-7855(96)00018-5

Cited By

Click to copy section linkSection link copied!

This article has not yet been cited by other publications.

ACS Catalysis

Cite this: ACS Catal. 2024, 14, 24, 18603–18614
Click to copy citationCitation copied!
https://doi.org/10.1021/acscatal.4c03672
Published December 6, 2024

Copyright © 2024 The Authors. Published by American Chemical Society. This publication is licensed under

CC-BY 4.0 .

Article Views

2003

Altmetric

-

Citations

-
Learn about these metrics

Article Views are the COUNTER-compliant sum of full text article downloads since November 2008 (both PDF and HTML) across all institutions and individuals. These metrics are regularly updated to reflect usage leading up to the last few days.

Citations are the number of other articles citing this article, calculated by Crossref and updated daily. Find more information about Crossref citation counts.

The Altmetric Attention Score is a quantitative measure of the attention that a research article has received online. Clicking on the donut icon will load a page at altmetric.com with additional details about the score and the social media presence for the given article. Find more information on the Altmetric Attention Score and how the score is calculated.

  • Abstract

    Figure 1

    Figure 1. Generation of the organic semiconductor, the hydrogenase-containing carboxysome shell, and the chemical–biological hybrid nanoreactor, H–S|TBAP-α. (a) Chemical structure of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP). (b) Crystal structure of α-polymorph of 1,3,6,8-tetra(4′-carboxyphenyl)pyrene (TBAP-α). (c) Genetic organization of the α-carboxysome shell operon and hydrogenase-expressing operon for the production of recombinant shells and active [FeFe]-hydrogenases in E. coli, respectively, and a schematic model of the hydrogenase-shell nanoreactor (H–S). EP: CsoS2 C-terminus as the encapsulation peptide. The hydGXEF genes encode the crucial maturase enzymes for hydrogenase formation and activation, including HydE, HydF, and HydG. PDB ID: Ferredoxin (Fd)-[FeFe]-hydrogenase, 2N0S; ferredoxin-NADP reductase (FNR), 2XNJ. (d) 3D model representation of the H–S|TBAP-α hybrid system, in which H–S adheres to the hydrogen-bonding groups on the TBAP-α crystal surface.

    Figure 2

    Figure 2. Binding between the α-carboxysome shell and TBAP-α. Results of molecular dynamics simulations of CsoS1A (PDB ID: 2EWH) and TBAP-α binding equilibrium states are shown in panels (a–c). (a) Front view. (b) Side view, showing the simulated binding interface between TBAP-α and the CsoS1A hexamer. (c) Zoom-in view of the noncovalent interactions (red dotted lines) formed between amino acid residues of CsoS1A (Pro96, Thr5, Ser86, Asp90, Gly89, Lys94, Arg34, Glut97) and TBAP-α (blue = N atoms, red = O atoms). (d) ITC thermogram resulting from the titration of TBAP (50 μM) into the α-carboxysome shell (3.75 μM) in TN buffer (20 mM Tris-HCl (pH = 8.0), 150 mM NaCl). (e) Thermodynamic profile of TBAP binding to α-carboxysome shell in TN buffer. ΔG, change in Gibbs Free Energy; ΔH, change in enthalpy; T, temperature in Kelvin; ΔS, change in entropy.

    Figure 3

    Figure 3. Attachment of mCherry-encapsulated C–S shells on the surface of TBAP-α crystals. SEM images of (a) uncoated TBAP-α, (b) C–S|TBAP-α (40 × 3.6 μm2 size), and (c) zoom-in view of C–S|TBAP-α. Orange arrows show the C–S particles that were attached to the TBAP-α crystal surface. (d) Fluorescence images of TBAP-α only (top row) and the C–S|TBAP-α hybrid (bottom row). Left, excitation at 488 nm; middle, excitation at 561 nm; right, the merged channel. All fluorescence images were adjusted to have the same brightness and contrast settings.

    Figure 4

    Figure 4. Characterization of the light-driven H–S|TBAP-α hybrid nanoreactor. (a) SEM image of H–S|TBAP-α. Orange arrows show the H–S particles that were attached to the TBAP-α crystal surface. (b) Photoluminescence (PL) spectrum and (c) Excited-state lifetimes of H–S|TBAP-α and TBAP-α alone in 0.1 M neutralized AA aqueous solution when excited at 390 nm.

    Figure 5

    Figure 5. Visible-light-driven hydrogen production of H–S|TBAP-α. (a) H2-evolution rates of TBAP-α, H–S|TBAP-α, H–Sox|TBAP-α, S|TBAP-α, and H–S|Amorphous TBAP irradiated by a solar simulator for 2 h. (b) H2 evolution of TBAP-α (black dots) and H–S|TBAP-α (blue dots) as a function of time (λ > 420 nm). (c) Schematic diagram of proposed electron transfer pathway in H–S|TBAP-α for hydrogen production. Electrons generated from TBAP-α irradiation are transferred through holes on the α-carboxysome shell to the 4Fe–4S cluster via FNR or directly for proton reduction. Ascorbic acid acts as a sacrificial agent to prevent the photogenerated hole–electron recombination. Error bars represent the SD of the mean of three independent experiments.

  • References


    This article references 62 other publications.

    1. 1
      Tachibana, Y.; Vayssieres, L.; Durrant, J. R. Artificial photosynthesis for solar water-splitting. Nat. Photonics 2012, 6, 511518,  DOI: 10.1038/nphoton.2012.175
    2. 2
      Blankenship, R. E.; Tiede, D. M.; Barber, J.; Brudvig, G. W.; Fleming, G.; Ghirardi, M.; Gunner, M. R.; Junge, W.; Kramer, D. M.; Melis, A.; Moore, T. A.; Moser, C. C.; Nocera, D. G.; Nozik, A. J.; Ort, D. R.; Parson, W. W.; Prince, R. C.; Sayre, R. T. Comparing Photosynthetic and Photovoltaic Efficiencies and Recognizing the Potential for Improvement. Science 2011, 332, 805809,  DOI: 10.1126/science.1200165
    3. 3
      Wang, Y.; Vogel, A.; Sachs, M.; Sprick, R. S.; Wilbraham, L.; Moniz, S. J. A.; Godin, R.; Zwijnenburg, M. A.; Durrant, J. R.; Cooper, A. I.; Tang, J. Current understanding and challenges of solar-driven hydrogen generation using polymeric photocatalysts. Nat. Energy 2019, 4, 746760,  DOI: 10.1038/s41560-019-0456-5
    4. 4
      Barber, J. Photosynthetic energy conversion: Natural and artificial. Chem. Soc. Rev. 2009, 38, 185196,  DOI: 10.1039/B802262N
    5. 5
      Zhang, J. Z.; Reisner, E. Advancing photosystem II photoelectrochemistry for semi-artificial photosynthesis. Nat. Rev. Chem. 2020, 4, 621,  DOI: 10.1038/s41570-019-0149-4
    6. 6
      Pi, X.; Zhao, S.; Wang, W.; Liu, D.; Xu, C.; Han, G.; Kuang, T.; Sui, S. F.; Shen, J. R. The pigment-protein network of a diatom photosystem II–light-harvesting antenna supercomplex. Science 2019, 365, 447457,  DOI: 10.1126/science.aax4406
    7. 7
      Vayghan, H. S.; Nawrocki, W. J.; Schiphorst, C.; Tolleter, D.; Hu, C.; Douet, V.; Glauser, G.; Finazzi, G.; Croce, R.; Wientjes, E.; Longoni, F. Photosynthetic Light Harvesting and Thylakoid Organization in a CRISPR/Cas9 Arabidopsis Thaliana LHCB1 Knockout Mutant. Front. Plant Sci. 2022, 13, 833032833050,  DOI: 10.3389/fpls.2022.833032
    8. 8
      Koepf, M.; Teillout, A.-L.; Llansola-Portoles, M. J. Artificial Photosynthesis: An Approach for a Sustainable Future. In Handbook of Ecomaterials; Martínez, L. M. T.; Kharissova, O. V.; Kharisov, B. I., Eds.; Springer International Publishing: Cham, 2017; pp 125.
    9. 9
      Guo, Y.; Zhou, Q.; Nan, J.; Shi, W.; Cui, F.; Zhu, Y. Perylenetetracarboxylic acid nanosheets with internal electric fields and anisotropic charge migration for photocatalytic hydrogen evolution. Nat. Commun. 2022, 13, 2067  DOI: 10.1038/s41467-022-29826-z
    10. 10
      Kosco, J.; Bidwell, M.; Cha, H.; Martin, T.; Howells, C. T.; Sachs, M.; Anjum, D. H.; Gonzalez Lopez, S.; Zou, L.; Wadsworth, A.; Zhang, W.; Zhang, L.; Tellam, J.; Sougrat, R.; Laquai, F.; DeLongchamp, D. M.; Durrant, J. R.; McCulloch, I. Enhanced photocatalytic hydrogen evolution from organic semiconductor heterojunction nanoparticles. Nat. Mater. 2020, 19, 559565,  DOI: 10.1038/s41563-019-0591-1
    11. 11
      Wang, Q.; Domen, K. Particulate Photocatalysts for Light-Driven Water Splitting: Mechanisms, Challenges, and Design Strategies. Chem. Rev. 2020, 120, 919985,  DOI: 10.1021/acs.chemrev.9b00201
    12. 12
      Takata, T.; Jiang, J.; Sakata, Y.; Nakabayashi, M.; Shibata, N.; Nandal, V.; Seki, K.; Hisatomi, T.; Domen, K. Photocatalytic water splitting with a quantum efficiency of almost unity. Nature 2020, 581, 411414,  DOI: 10.1038/s41586-020-2278-9
    13. 13
      Searle, N. Z.; Hirt, R. C. Ultraviolet Spectral Energy Distribution of Sunlight. J. Opt. Soc. Am. 1965, 55, 14131421,  DOI: 10.1364/JOSA.55.001413
    14. 14
      Cestellos-Blanco, S.; Zhang, H.; Kim, J. M.; Shen, Yx.; Yang, P. Photosynthetic semiconductor biohybrids for solar-driven biocatalysis. Nat. Catal. 2020, 3, 245255,  DOI: 10.1038/s41929-020-0428-y
    15. 15
      Kornienko, N.; Zhang, J. Z.; Sakimoto, K. K.; Yang, P.; Reisner, E. Interfacing nature’s catalytic machinery with synthetic materials for semi-artificial photosynthesis. Nat. Nanotechnol. 2018, 13, 890899,  DOI: 10.1038/s41565-018-0251-7
    16. 16
      Özgen, F. F.; Runda, M. E.; Schmidt, S. Photo-biocatalytic Cascades: Combining Chemical and Enzymatic Transformations Fueled by Light. ChemBioChem 2021, 22, 790806,  DOI: 10.1002/cbic.202000587
    17. 17
      Schmermund, L.; Jurkaš, V.; Özgen, F. F.; Barone, G. D.; Büchsenschütz, H. C.; Winkler, C. K.; Schmidt, S.; Kourist, R.; Kroutil, W. Photo-Biocatalysis: Biotransformations in the Presence of Light. ACS Catal. 2019, 9, 41154144,  DOI: 10.1021/acscatal.9b00656
    18. 18
      Holá, K.; Pavliuk, M. V.; Németh, B.; Huang, P.; Zdražil, L.; Land, H.; Berggren, G.; Tian, H. Carbon Dots and [FeFe] Hydrogenase Biohybrid Assemblies for Efficient Light-Driven Hydrogen Evolution. ACS Catal. 2020, 10, 99439952,  DOI: 10.1021/acscatal.0c02474
    19. 19
      Gai, P.; Yu, W.; Zhao, H.; Qi, R.; Li, F.; Liu, L.; Lv, F.; Wang, S. Solar-Powered Organic Semiconductor-Bacteria Biohybrids for CO2 Reduction into Acetic Acid. Angew. Chem., Int. Ed. 2020, 59, 72247229,  DOI: 10.1002/anie.202001047
    20. 20
      Wang, X.; Saba, T.; Yiu, H. H. P.; Howe, R. F.; Anderson, J. A.; Shi, J. Cofactor NAD(P)H Regeneration Inspired by Heterogeneous Pathways. Chem 2017, 2, 621654,  DOI: 10.1016/j.chempr.2017.04.009
    21. 21
      Gentil, S.; Che Mansor, S. M.; Jamet, H.; Cosnier, S.; Cavazza, C.; Le Goff, A. Oriented Immobilization of [NiFeSe] Hydrogenases on Covalently and Noncovalently Functionalized Carbon Nanotubes for H2/Air Enzymatic Fuel Cells. ACS Catal. 2018, 8, 39573964,  DOI: 10.1021/acscatal.8b00708
    22. 22
      Zhang, S.; Liu, S.; Sun, Y.; Li, S.; Shi, J.; Jiang, Z. Enzyme-photo-coupled catalytic systems. Chem. Soc. Rev. 2021, 50, 1344913466,  DOI: 10.1039/D1CS00392E
    23. 23
      Sun, Y.; Lin, Y.; Harman, V. M.; Beynon, R. J.; Johnson, J. R.; Liu, L.-N. Decoding the Absolute Stoichiometric Composition and Structural Plasticity of a-Carboxysomes. mBio 2022, 13, e03629-21  DOI: 10.1128/mbio.03629-21
    24. 24
      Gonzalez-Esquer, C. R.; Newnham, S. E.; Kerfeld, C. A. Bacterial microcompartments as metabolic modules for plant synthetic biology. Plant J. 2016, 87, 6675,  DOI: 10.1111/tpj.13166
    25. 25
      Liu, L. N. Advances in the bacterial organelles for CO2 fixation. Trends Microbiol. 2022, 30, 567580,  DOI: 10.1016/j.tim.2021.10.004
    26. 26
      Huang, J.; Jiang, Q.; Yang, M.; Dykes, G. F.; Weetman, S. L.; Xin, W.; He, H. L.; Liu, L. N. Probing the internal pH and permeability of a carboxysome shell. Biomacromolecules 2022, 23, 43394348,  DOI: 10.1021/acs.biomac.2c00781
    27. 27
      Faulkner, M.; Szabó, I.; Weetman, S. L.; Sicard, F.; Huber, R. G.; Bond, P. J.; Rosta, E.; Liu, L.-N. Molecular simulations unravel the molecular principles that mediate selective permeability of carboxysome shell protein. Sci. Rep. 2020, 10, 17501  DOI: 10.1038/s41598-020-74536-5
    28. 28
      Mahinthichaichan, P.; Morris, D. M.; Wang, Y.; Jensen, G. J.; Tajkhorshid, E. Selective permeability of carboxysome shell pores to anionic molecules. J. Phys. Chem. B 2018, 122, 91109118,  DOI: 10.1021/acs.jpcb.8b06822
    29. 29
      Cammack, R.; Frey, M.; Robson, R. Hydrogen as a Fuel, Learning from Nature; CRC Press, 2001.
    30. 30
      Vignais, P. M.; Billoud, B. Occurrence, Classification, and Biological Function of Hydrogenases: An Overview. Chem. Rev. 2007, 107, 42064272,  DOI: 10.1021/cr050196r
    31. 31
      Lubitz, W.; Ogata, H.; Rudiger, O.; Reijerse, E. Hydrogenases. Chem. Rev. 2014, 114, 40814148,  DOI: 10.1021/cr4005814
    32. 32
      Jiang, Q.; Li, T.; Yang, J.; Aitchison, C. M.; Huang, J.; Chen, Y.; Huang, F.; Wang, Q.; Cooper, A. I.; Liu, L.-N. Synthetic engineering of a new biocatalyst encapsulating [NiFe]-hydrogenases for enhanced hydrogen production. J. Mater. Chem. B 2023, 11, 26842692,  DOI: 10.1039/D2TB02781J
    33. 33
      Li, T.; Jiang, Q.; Huang, J.; Aitchison, C. M.; Huang, F.; Yang, M.; Dykes, G. F.; He, H. L.; Wang, Q.; Sprick, R. S.; Cooper, A. I.; Liu, L. N. Reprogramming bacterial protein organelles as a nanoreactor for hydrogen production. Nat. Commun. 2020, 11, 5448  DOI: 10.1038/s41467-020-19280-0
    34. 34
      Qin, W. K.; Tung, C. H.; Wu, L. Z. Covalent organic framework and hydrogen-bonded organic framework for solar-driven photocatalysis. J. Mater. Chem. A 2023, 11, 1252112538,  DOI: 10.1039/D2TA09375H
    35. 35
      Aitchison, C. M.; Kane, C. M.; McMahon, D. P.; Spackman, P. R.; Pulido, A.; Wang, X.; Wilbraham, L.; Chen, L.; Clowes, R.; Zwijnenburg, M. A.; Sprick, R. S.; Little, M. A.; Day, G. M.; Cooper, A. I. Photocatalytic proton reduction by a computationally identified, molecular hydrogen-bonded framework. J. Mater. Chem. A 2020, 8, 71587170,  DOI: 10.1039/D0TA00219D
    36. 36
      Stylianou, K. C.; Heck, R.; Chong, S. Y.; Bacsa, J.; Jones, J. T. A.; Khimyak, Y. Z.; Bradshaw, D.; Rosseinsky, M. J. A guest-responsive fluorescent 3D microporous metal-organic framework derived from a long-lifetime pyrene core. J. Am. Chem. Soc. 2010, 132, 41194130,  DOI: 10.1021/ja906041f
    37. 37
      Chen, G.; Huang, S.; Shen, Y.; Kou, X.; Ma, X.; Huang, S.; Tong, Q.; Ma, K.; Chen, W.; Wang, P.; Shen, J.; Zhu, F.; Ouyang, G. Protein-directed, hydrogen-bonded biohybrid framework. Chem 2021, 7, 27222742,  DOI: 10.1016/j.chempr.2021.07.003
    38. 38
      Zhou, Q.; Guo, Y.; Zhu, Y. Photocatalytic sacrificial H2 evolution dominated by micropore-confined exciton transfer in hydrogen-bonded organic frameworks. Nat. Catal. 2023, 6, 574584,  DOI: 10.1038/s41929-023-00972-x
    39. 39
      Pellegrin, Y.; Odobel, F. Sacrificial electron donor reagents for solar fuel production. C. R. Chim. 2017, 20, 283295,  DOI: 10.1016/j.crci.2015.11.026
    40. 40
      Baker, S. H.; Lorbach, S. C.; Rodriguez-Buey, M.; Williams, D. S.; Aldrich, H. C.; Shively, J. M. The correlation of the gene csoS2 of the carboxysome operon with two polypeptides of the carboxysome in Thiobacillus neapolitanus. Arch. Microbiol. 1999, 172, 233239,  DOI: 10.1007/s002030050765
    41. 41
      Case, D. A.; Aktulga, H. M.; Belfon, K.; Cerutti, D. S.; Cisneros, G. A.; Cruzeiro, V. W. D.; Forouzesh, N.; Giese, T. J.; Götz, A. W.; Gohlke, H.; Izadi, S.; Kasavajhala, K.; Kaymak, M. C.; King, E.; Kurtzman, T.; Lee, T.-S.; Li, P.; Liu, J.; Luchko, T.; Luo, R.; Manathunga, M.; Machado, M. R.; Nguyen, H. M.; O’Hearn, K. A.; Onufriev, A. V.; Pan, F.; Pantano, S.; Qi, R.; Rahnamoun, A.; Risheh, A.; Schott-Verdugo, S.; Shajan, A.; Swails, J.; Wang, J.; Wei, H.; Wu, X.; Wu, Y.; Zhang, S.; Zhao, S.; Zhu, Q.; Cheatham, T. E., III; Roe, D. R.; Roitberg, A.; Simmerling, C.; York, D. M.; Nagan, M. C.; Merz, K. M., Jr. AmberTools. J. Chem. Inf. Model. 2023, 63, 61836191,  DOI: 10.1021/acs.jcim.3c01153
    42. 42
      Karthi, N.; Venkatachalam, M. Growth and Characterization Novel Organic Nonlinear Optical Crystal of Pyrene. Int. J. Sci. 2013, 1, 812
    43. 43
      Tanaka, S.; Kerfeld, C. A.; Sawaya, M. R.; Cai, F.; Heinhorst, S.; Cannon, G. C.; Yeates, T. O. Atomic-level models of the bacterial carboxysome shell. Science 2008, 319, 10831086,  DOI: 10.1126/science.1151458
    44. 44
      Tsai, Y.; Sawaya, M. R.; Cannon, G. C.; Cai, F.; Williams, E. B.; Heinhorst, S.; Kerfeld, C. A.; Yeates, T. O. Structural analysis of CsoS1A and the protein shell of the Halothiobacillus neapolitanus carboxysome. PLoS Biol. 2007, 5, e144,  DOI: 10.1371/journal.pbio.0050144
    45. 45
      Parsons, J. B.; Dinesh, S. D.; Deery, E.; Leech, H. K.; Brindley, A. A.; Heldt, D.; Frank, S.; Smales, C. M.; Lünsdorf, H.; Rambach, A.; Gass, M. H.; Bleloch, A.; McClean, K. J.; Munro, A. W.; Rigby, S. E. J.; Warren, M. J.; Prentice, M. B. Biochemical and Structural Insights into Bacterial Organelle Form and Biogenesis. J. Biol. Chem. 2008, 283, 1436614375,  DOI: 10.1074/jbc.M709214200
    46. 46
      Parsons, J. B.; Lawrence, A. D.; McLean, K. J.; Munro, A. W.; Rigby, S. E. J.; Warren, M. J. Characterisation of PduS, the pdu Metabolosome Corrin Reductase, and Evidence of Substructural Organisation within the Bacterial Microcompartment. PLoS One 2010, 5, e14009,  DOI: 10.1371/journal.pone.0014009
    47. 47
      Silva, D. A.; Yu, S.; Ulge, U. Y.; Spangler, J. B.; Jude, K. M.; Labão-Almeida, C.; Ali, L. R.; Quijano-Rubio, A.; Ruterbusch, M.; Leung, I.; Biary, T.; Crowley, S. J.; Marcos, E.; Walkey, C. D.; Weitzner, B. D.; Pardo-Avila, F.; Castellanos, J.; Carter, L.; Stewart, L.; Riddell, S. R.; Pepper, M.; Bernardes, G. J. L.; Dougan, M.; Garcia, K. C.; Baker, D. De novo design of potent and selective mimics of IL-2 and IL-15. Nature 2019, 565, 186191,  DOI: 10.1038/s41586-018-0830-7
    48. 48
      Thompson, M. C.; Wheatley, N. M.; Jorda, J.; Sawaya, M. R.; Gidaniyan, S. D.; Ahmed, H.; Yang, Z.; McCarty, K. N.; Whitelegge, J. P.; Yeates, T. O. Identification of a Unique Fe-S Cluster Binding Site in a Glycyl-Radical Type Microcompartment Shell Protein. J. Mol. Biol. 2014, 426, 32873304,  DOI: 10.1016/j.jmb.2014.07.018
    49. 49
      Zeng, Z.; Boeren, S.; Bhandula, V.; Light, S. H.; Smid, E. J.; Notebaart, R. A.; Abee, T. Bacterial Microcompartments Coupled with Extracellular Electron Transfer Drive the Anaerobic Utilization of Ethanolamine in Listeria monocytogenes. mSystems 2021, 6, e01349-20  DOI: 10.1128/msystems.01349-20
    50. 50
      Ferlez, B.; Markus, S.; Kerfeld, C. A. Glycyl Radical Enzyme-Associated Microcompartments: Redox-Replete Bacterial Organelles. mBio 2019, 10, e02327-18  DOI: 10.1128/mBio.02327-18
    51. 51
      Yan, Y. M.; Tao, H. Y.; He, J. H.; Huang, S.-Y. The HDOCK server for integrated protein–protein docking. Nat. Protoc. 2020, 15, 18291852,  DOI: 10.1038/s41596-020-0312-x
    52. 52
      Case, A. D.; Belfon, K.; Ido, B.-S.; Scott, R. B.; Cerutti, D. S.; Cheatham, T. E., III; Cruzeiro, V. W. D.; Darden, T. A.; Duke, R. E.; Giambasu, G.; Gilson, M. K.; Gohlke, H.; Goetz, A. W.; Harris, R.; Izadi, S.; Izmailov, S. A.; Kasavajhala, K.; Kovalenko, A.; Krasny, R.; Kurtzman, T.; Lee, T. S.; LeGrand, S.; Li, P.; Lin, C.; L, J.; Luchko, T.; Luo, R.; Man, V.; Merz, K. M.; Miao, Y.; Mikhailovskii, O.; Monard, G.; Nguyen, H.; Onufriev, A.; Pan, F.; Pantano, S.; Qi, R.; Roe, D. R.; Roitberg, A.; Sagui, C.; Schott-Verdugo, S.; Shen, J.; Simmerling, C. L.; Skrynnikov, N. R.; Smith, J.; Swails, J.; Walker, R. C.; Wang, J.; Wilson, L.; Wolf, R. M.; Wu, X.; Xiong, Y.; Xue, Y.; D M AMBER 2020; University of California: New York, 2020.
    53. 53
      Maier, J. A.; Martinez, C.; Kasavajhala, K.; Wickstrom, L.; Hauser, K. E.; Simmerling, C. ff14SB: Improving the Accuracy of Protein Side Chain and Backbone Parameters from ff99SB. J. Chem. Theory Comput. 2015, 11, 36963713,  DOI: 10.1021/acs.jctc.5b00255
    54. 54
      Wang, J.; Wolf, R. M.; Caldwell, J. W.; Kollman, P. A.; Case, D. A. Development and testing of a general amber force field. J. Comput. Chem. 2004, 25, 11571174,  DOI: 10.1002/jcc.20035
    55. 55
      Wang, Z.; Li, J.; Liu, J.; Wang, L.; Lu, Y.; Liu, J. P. Molecular insight into the selective binding between human telomere G-quadruplex and a negatively charged stabilizer. Clin. Exp. Pharmacol. Physiol. 2020, 47, 892902,  DOI: 10.1111/1440-1681.13249
    56. 56
      Kim, M.; Kreig, A.; Lee, C. Y.; Rube, H. T.; Calvert, J.; Song, J. S.; Myong, S. Quantitative analysis and prediction of G-quadruplex forming sequences in double-stranded DNA. Nucleic Acids Res. 2016, 44, 48074817,  DOI: 10.1093/nar/gkw272
    57. 57
      Price, D. J.; Charles, L. B. A modified TIP3P water potential for simulation with Ewald summation. J. Chem. Phys. 2004, 121, 1009610103,  DOI: 10.1063/1.1808117
    58. 58
      Li, P. F.; Merz, K. Metal ion modeling using classical mechanics. Chem. Rev. 2017, 117, 15641686,  DOI: 10.1021/acs.chemrev.6b00440
    59. 59
      Cheatham, T. E. I.; Miller, J. L.; Fox, T.; Darden, T. A.; Kollman, P. A. Molecular Dynamics Simulations on Solvated Biomolecular Systems: The Particle Mesh Ewald Method Leads to Stable Trajectories of DNA, RNA, and Proteins. J. Am. Chem. Soc. 1995, 117, 41934194,  DOI: 10.1021/ja00119a045
    60. 60
      Loncharich, R. J.; Brooks, R. B.; Pastor, W. R. Langevin dynamics of peptides: The frictional dependence of isomerization rates of N-acetylalanyl-N′-methylamide. Biopolymers 1992, 32, 523535,  DOI: 10.1002/bip.360320508
    61. 61
      Berendsen, H. J. C.; Postma, J. P. M.; Gunsteren, W. F.; DiNola, A.; Haak, J. R. Molecular dynamics with coupling to an external bath. J. Chem. Phys. 1984, 81, 36843690,  DOI: 10.1063/1.448118
    62. 62
      Humphrey, W.; Dalke, A.; Schulten, K. VMD: Visual Molecular Dynamics. J. Mol. Graphics 1996, 14, 3338,  DOI: 10.1016/0263-7855(96)00018-5
  • Supporting Information

    Supporting Information


    The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acscatal.4c03672.

    • Chemical synthesis pathway of TBAP (Scheme S1); NMR spectrum of TBAP (Figures S1 and S2); FI-IR spectrum of TBAP (Figure S3); crystal model of TBAP-α and 3D model of α-carboxysome shell (Figure S4); PXRD patterns of TBAP phases, nitrogen adsorption isotherm and desorption isotherm for activated TBAP-α and amorphous TBAP, and SEM images of amorphous TBAP (Figure S5); solid UV–vis absorption spectrum of amorphous TBAP and TBAP-α (Figure S6); (αhν)1/2 versus hν curve and Mott–Schottky plot, diagram of conduction band and valence band of amorphous TBAP and TBAP-α, and cyclic voltammetry plot of TBAP-α (Figure S7); amino acid sequence alignment of α-carboxysome shell proteins (Figure S8); front and side views of the structures of α-carboxysome shell proteins (Figure S9); confocal microscopy images of E. coli cells expressing mCherry-CsoS2C (mCherry-EP) and coexpressing shells and mCherry-csoS2C (C–S) and SDS-PAGE of purified C–S (Figure S10); TEM image of C–S (Figure S11); SEM and confocal microscopy images of pyrene crystals and pyrene crystals with C–S (Figure S12); ζ-potentials of TBAP-α and α-carboxysome shell-encasing proteins (Figure S13); size distribution of H–S revealed by SEM (Figure S14); SDS-PAGE result of H–S purification (Figure S15); immunoblot analysis of purified H–S (Figure S16); colloidal stability of H–S|TBAP-α (Figure S17); photocurrent responses and EIS analysis for TBAP-α and H–S|TBAP-α (Figure S18); H2 production condition optimization of H–S|TBAP-α (Figure S19); photocurrent response and EIS analysis for amorphous TBAP and TBAP-α (Figure S20); wavelength-dependent AQY value of H–S|TBAP-α (Figure S21); H2 evolution of TBAP-α, H–S|TBAP-α, and 1 wt % Pt|TBAP-α (λ > 420 nm) as a function of time (Figure S22); cycling measurements for the photocatalytic hydrogen evolution of H–S|TBAP-α (Figure S23); immunoblot analysis of purified HydA (Fd-HydA-EP) (Figure S24); SEM images (Figure S25) and PXRD pattern (Figure S26) of H–S|TBAP-α after 30 h irradiation; gene maps of used plasmids (Figure S27); crystal data and structure refinement of TBAP-α (Table S1); protein components in recombinant α-carboxysomes from E. coli (Table S2); statistics of residues for TBAP-α and CsoS1A protein binding calculated by MD simulations (Tables S3 and S4); estimated fluorescence lifetimes of TBAP-α and H–S|TBAP-α (Table S5); a list of hydrogen evolution reaction conditions in this work (Table S6); a comparison of the H–S|TBAP-α assembly performance with the related state-of-the-art photocatalysts (Table S7); primers used for pCDFDuet-mCherry-CS2 plasmid construction (Table S8); and gene sequences of used plasmids (Table S9). (PDF)


    Terms & Conditions

    Most electronic Supporting Information files are available without a subscription to ACS Web Editions. Such files may be downloaded by article for research use (if there is a public use license linked to the relevant article, that license may permit other uses). Permission may be obtained from ACS for other uses through requests via the RightsLink permission system: http://pubs.acs.org/page/copyright/permissions.html.