Photocatalyst-mineralized biofilms as living bio-abiotic interfaces for single enzyme to whole-cell photocatalytic applications

Abstract

There is an increasing trend of combining living cells with inorganic semiconductors to construct semi-artificial photosynthesis systems. Creating a robust and benign bio-abiotic interface is key to the success of such solar-to-chemical conversions but often faces a variety of challenges, including biocompatibility and the susceptibility of cell membrane to high-energy damage arising from direct interfacial contact. Here, we report living mineralized biofilms as an ultrastable and biocompatible bio-abiotic interface to implement single enzyme to whole-cell photocatalytic applications. These photocatalyst-mineralized biofilms exhibited efficient photoelectrical responses and were further exploited for diverse photocatalytic reaction systems including a whole-cell photocatalytic CO₂ reduction system enabled by the same biofilm-producing strain. Segregated from the extracellularly mineralized semiconductors, the bacteria remained alive even after 5 cycles of photocatalytic NADH regeneration reactions, and the biofilms could be easily regenerated. Our work thus demonstrates the construction of biocompatible interfaces using biofilm matrices and establishes proof of concept for future sustainable photocatalytic applications.

Fig. 1.. Schematic of living photocatalyst-mineralized biofilms explored for single enzyme to whole-cell photocatalysis.

The A7 peptides within the biofilms are functional peptides specifically displayed on the CsgA_A7 nanofibers that can be mineralized via in situ formation of CdS NPs generating photocatalyst-mineralized biofilms. The mineralized biofilms harbor the CdS NPs that can produce electrons upon light irradiation; the electrons are then harnessed and transferred to the catalytic centers of redox enzymes through electron mediators (MV or NAD) for various photocatalytic applications including reduction of TMP into l-tert-leucine coupling with purified LDH (A) and CO₂ reduction coupling with a single cell coexpressing CsgA_A7 fibers and FDH (B).

Fig. 2.. Characterization of the functional biofilms mineralized with CdS NPs.

(A) Schematic of the formation process for Tc_Receiver/CsgA_A7 biofilms and their TEM and SAED images. (B) Schematic of the in situ mineralization process on biofilm curli nanofibers and TEM and SAED (inset) images of the CdS-mineralized biofilms. The mineralized biofilms exhibited a clear diffraction ring, whereas the Tc_Receiver/CsgA_A7 biofilms did not. The (100) corresponds to the space lattice of 0.35 nm for the CdS NPs in (C). (C and D) HRTEM, HAADF-STEM, and EDS mapping images of the nanofiber/CdS composites in the mineralized biofilms. The green represents Cd atoms, and the orange represents S atoms. (E and F) XPS spectra of CdS-mineralized biofilms, which confirmed the presence of Cd and S atoms, in accordance with the EDS mapping results. The red curves were fitted curves to accurately determine the peak value position. (G) Inductively coupled plasma optical emission spectrometry (ICP-OES) results for measurement of free Cd²⁺ ion concentrations in the supernatant during the process of in situ mineralization of CdS NPs on biofilms in M63 cultivation medium over a 2-day period. (H and I) TEM (H) and SAED (I) images for the Tc_Receiver/ompA_A7 sample mineralized with CdS NPs. (J and K) TEM images for the CdS-mineralized biofilms (J) and CdS-mineralized Tc_Receiver/ompA_A7 cells (K) after irradiation for 24 hours by an artificial blue light. The red arrows pointed to the damaged cell parts.

Fig. 3.. Photoelectric properties of the photocatalyst-mineralized biofilms.

(A) Ultraviolet-visible (UV-vis) spectra of the Tc_Receiver/CsgA_A7 biofilms and the photocatalyst-mineralized biofilms. (B) The bandgap of the CdS NPs formed within the photocatalyst-mineralized biofilms. A is the normalized absorption value, h is the Planck’s constant (6.626 × 10⁻³⁴ J·s), and ν is the frequency. (C) Transient photocurrent curves measured through a three-electrode system, where FTO-conductive glass deposited with biofilm samples served as the working electrode and a platinum wire served as the counter electrode; there was also a Ag/AgCl reference electrode. The blue curve represents the photocurrent of the FTO glass (1 cm by 1 cm) deposited with photocatalyst-mineralized biofilms, upon illumination (light on) or shielding (light off) from a Xe lamp. The red and black curves represent the photocurrent of the FTO glass (1 cm by 1 cm) deposited with Tc_Receiver/CsgA_A7 biofilms or bare FTO glass, respectively. (D) Schematic of photocatalytic electron transfer to MV via the photocatalyst-mineralized biofilms. The 3-ml tris-HCl reaction solution was put into a cuvette containing 1 mM MV²⁺ (as the electron mediator), 1.5% TEOA (as the sacrificial agent), and the photocatalyst-mineralized biofilms. The reaction solution was irradiated for 12 hours, during which colorless MV²⁺ was converted to violet MV^+•. (E) UV-vis spectra for MV²⁺ and its reduced form MV^+•. In (A and C), “CsgA_A7” refers to Tc_Receiver/CsgA_A7 biofilms. “CsgA_A7/CdS” refers to photocatalyst-mineralized biofilms.

Fig. 4.. Photocatalytic reduction of TMP into l-tert-leucine coupling with purified LDH enzyme.

(A) Schematic of the TMP reduction process, where CdS NPs within the photocatalyst-mineralized biofilms absorb light and produce electrons. (B) NAD⁺ reduction under different reaction systems using artificial blue light-emitting diode (LED) arrays as the light source. “CsgA_A7 biofilm (light)” refers to the reaction system using Tc_Receiver/CsgA_A7 biofilms; “CsgA_A7/CdS (dark)” refers to the reaction system without illumination. (C) Relative activity comparison for five consecutive reactions, for which the photocatalyst-mineralized biofilms were recycled for use each time after 3 hours of illumination. These 1-ml reaction solutions (in tris-HCl solution, pH 8.0) contained 1 mM NAD⁺, 1.5% TEOA, 0.25 mM [Cp*Rh(bpy)H₂O]²⁺, and the recycled photocatalyst-mineralized biofilms (with pellets were originally collected from 20 ml of M63 culture medium and recovered via centrifugation and resuspension after each of the reactions). (D) Reduction of TMP to l-tert-leucine under different reaction systems using artificial blue LED arrays as the light source. The 1-ml tris-HCl reaction solution (pH 8.0) contained LDH (1, 2, or 3 mg/ml), 1.5% TEOA, NAD⁺ (1, 5, or 10 mM), 0.25 mM [Cp*Rh(bpy)H₂O]²⁺, 10 mM TMP, 10 mM NH₄Cl, and photocatalyst-mineralized biofilms. Each experiment was repeated three times. Student’s t test, **P < 0.01, and ***P < 0.001.

Fig. 5.. Whole cell–enabled photocatalytic CO₂ reduction.

(A) Schematic of a photocatalytic CO₂ reduction system based on a single strain coexpressing CsgA_A7 nanofibers for CdS mineralization and FDH enzymes for CO₂ reduction. (B) TEM image of the photocatalyst-mineralized biofilms through in situ mineralization on Tc_Receiver/CsgA_A7-FDH biofilms. (C) SDS-PAGE image, Western blot image, and LC-MS spectrum for the purified FDH enzymes from Tc_Receiver/CsgA_A7-FDH biofilms. (D) Photocatalytic CO₂ reduction under different reaction conditions. “Light” refers to a reaction system comprising 5 mM MV, 10 mM ascorbic acid, 100 mM NaHCO₃, and the photocatalyst-mineralized biofilms in PBS solution (pH 7.4) under normal illumination condition. “Dark” refers to the same reaction system but lacking illumination. “−FDH” refers to an illuminated reaction system without expression of FDH enzymes. “−CdS” refers to an illuminated reaction system with Tc_Receiver/CsgA_A7-FDH biofilms that were not mineralized with CdS NPs. “−MV” refers to an illuminated reaction system without MV.