Microbial autotrophy driven by extracellular electrons generated from mechanical forces via flexoelectricity

Abstract

Mechanical force has recently emerged as a microbial energy source through direct mechanical-to-electrical energy conversion. However, the flexoelectric effect, a universal mechanoelectrical coupling phenomenon, remains underexplored as an energy source for microbial life. Here, we report a mechanically-driven microbial growth mechanism whereby mechanical energy is converted into electrical energy via the flexoelectric effect of a mineral. This process supports microbial growth in the absence of sunlight and chemical energy sources. Using flexoelectric manganese oxide and electroactive Rhodopseudomonas palustris as a model biohybrid, we demonstrate that mechanical force bends manganese oxide nanosheets to generate free electrical charge. R. palustris captures and utilizes this electrical energy for growth by coupling carbon fixation and denitrification in the absence of chemical electron donors under dark conditions. This pathway is accessible to various electroactive microorganisms and flexoelectric minerals. These findings reveal a noteworthy energy acquisition pathway for microbial survival in energy-limited settings, providing crucial evidence for the diversity of mechanical-to-metabolic coupling in biological processes.

Mechanical forces are ubiquitous in natural environments, yet the ability of microbes to harness this energy is underexplored. Here, the authors reveal that flexoelectric minerals generate free electrons under mechanical strain, providing electroactive microbes with an energy source for growth independent of sunlight or chemical substrates.

Introduction

Phototrophy and chemotrophy are the dominant microbial energy strategies for most microbial life on Earth. Yet, microbial life also thrives in the most energy-starved environments on the Earth, from subsurface sediments to deep-sea vents, having evolved additional versatile metabolic strategies^1–3. In these remote environments, microbes cannot rely on light or chemical substrates^4,5. Electroactive microorganisms (EAMs) are one such type of diverse group of bacteria, archaea, and eukaryotes that have been shown to directly harvest extracellular electrons to obtain growth and metabolic energy^6,7. Research has shown that non-phototrophic EAMs, such as Moorella thermoacetica, Alcaligenes faecalis, Shewanella oneidensis, and Acidithiobacillus ferrooxidans, support their growth and metabolic activites by utilizing electrons produced from the photocatalytic activity of semiconductive materials^8–11. Additionally, their metabolic flexibility allows them to tap into geo-electrical currents in hydrothermal vents and exploit electron flows generated during water evaporation^7,12. However, the widespread availability and impact of these electron sources in natural settings remain under debate, and further exploration into the viable energy transduction mechanisms is needed.

Mechanical forces, ubiquitous in natural systems ranging from tectonic shifts and fluid flow to biological activities, constitute a vastly underexploited energy reservoir. These forces fundamentally govern geomorphological processes, drive essential ecological functions, and modulate interspecies dynamics^13,14. Pioneering studies have demonstrated that piezoelectric materials (e.g., struvite, barium titanate and zinc oxide) can successfully convert mechanical stress into electrical energy, thereby supporting the growth and metabolism of denitrifying and electrotrophic microorganisms^15–18. These findings suggested a survival mechanism for bacteria and potential applications in biotechnology. The studies are also improving our understanding of the interactions between living organisms and electromechanical materials. However, conventional piezoelectric materials are constrained by their necessity for non-centrosymmetric structures, which limits the variety of materials available and their ecological relevance, as few natural minerals exhibit this property^19,20. In contrast, the flexoelectric effect, driven by strain gradients, overcomes such structural limitations^21,22. The flexoelectric effect, by responding to non-uniform strain (e.g., bending and twisting), achieves practical viability across all dielectric materials and serves as a highly representative mechanism for natural mechanical environments^23,24. Recent studies have successfully demonstrated the flexoelectric effect in various structures and geometries, such as bent-core nematic liquid crystals, polymer films, perovskite oxides membranes, porous biodegradable material, bulk semiconductor materials and free-standing cantilevers^25–30. Notably, even widely available natural materials, such as titanium dioxide (TiO₂) and manganese dioxide (MnO₂), display pronounced flexoelectric responses, particularly at the nanoscale^31,32. Although flexoelectricity generally exhibits lower theoretical energy conversion efficiency than piezoelectricity^30,33, its broad applicability and sensitivity to strain gradients make it a versatile alternative for real-world energy harvesting applications. For instance, flexoelectric effects have wide-ranging applications in sensors, actuators, and energy harvesting systems^34,35. However, despite these advances, the potential of flexoelectricity to power microbial physiological processes, including growth and metabolism, has yet to be explored.

In this study, we present compelling evidence that mechanical-to-electrical energy conversion induced by the flexoelectric effect of dielectric minerals can supply extracellular electrons to sustain microbial growth and metabolism. We demonstrate this mechanism using a model system comprising flexoelectric MnO₂ nanoflowers and the bacterium Rhodopseudomonas palustris. The selection of MnO₂ and R. palustris is grounded in their environmental relevance and complementary functions. MnO₂ is not only a flexoelectric material but also a widespread mineral in natural settings such as soils and sediments³⁶. R. palustris was chosen as the biological counterpart due to its widespread distribution, metabolic versatility, and well-documented competence in extracellular electron uptake (EEU)^37–39. We show that under dark and oligotrophic conditions, the flexoelectric MnO₂ nanosheets convert mechanical strain into bioavailable electrons, which are subsequently utilized by R. palustris to drive CO₂ fixation and nitrate reduction metabolic reactions (Fig. 1). The strain gradient generated by mechanical polarization yields surface-localized free electrons that are readily accessible to the microbes. Crucially, this mechanically-driven metabolic strategy extends to diverse combinations of electroactive microorganisms (EAMs; e.g., Shewanella oneidensis, Geobacter sulfurreducens) and flexoelectric minerals (e.g., SnO₂, WS₂, and birnessite nanoflowers), suggesting its broad ecological significance. By bridging mechanics, materials science, and microbial electrochemistry, our work unveils an energy-harvesting strategy, positioning flexoelectricity as a cornerstone of microbial resilience in energy-scarce environments.

Fig. 1. Schematic illustration of the flexoelectric-driven microbial energy transduction pathway.

Mechanical agitation induces bending of MnO₂ nanosheets, generating strain gradients that produce flexoelectric polarization and free electrons. These electrons are captured by R. palustris via extracellular electron transfer pathways and used to drive CO₂ fixation and denitrification under dark, electron‑donor‑free conditions.

Results

R. palustris harnesses mechanical energy for growth

To evaluate the implications of the flexoelectricity on microbial processes, we first explored the autotrophic growth of R. palustris in the presence of MnO₂ nanoflowers under continuous stirring at 400 rpm (ST-RP-MnO₂ nanoflower) during a 6-day dark culture in an anaerobic reactor (Fig. 2A and Table S1). Scanning electron microscopy (SEM) images confirmed the coexistence of R. palustris cells and MnO₂ nanoflowers after 6 days of culturing (Fig. S1). As shown in Fig. 2B and Fig. S2, mechanical stirring increased R. palustris biomass by 50.0% after 6 days in the ST-RP-MnO₂ nanoflower system, as quantified by total cellular protein. In contrast, cultures without mechanical stirring (RP-MnO₂ nanoflower), those subjected to stirring in the absence of MnO₂ nanoflowers (ST-RP), and cultures stirred with only non-flower-structured MnO₂ particles (ST-RP-MnO₂ particle) all exhibited reduced total protein content (Fig. 2B and Fig. S2). These reductions likely resulted from autolytic processes initiated under nutrient-limited conditions, whereby cells decompose themselves to maintain biomass when external energy sources are insufficient to support the metabolic activity of highly aggregated microbial cells⁴⁰. The growth dynamics of R. palustris were further analyzed using quantitative dry cell weight (DCW) and real-time PCR (qPCR) analyses. Biomass accumulation under flexoelectric culture conditions increased by 0.37 mg DCM/L to reach 0.57 mg DCW/L, which is higher than that of photoautotrophic growth but lower than that of heterotrophic growth (Fig. S2). This intermediate level of productivity indicates that the flexoelectricity can sustain measurable cellular growth, supporting a distinct, non-photosynthetic metabolic mode. After 6 days of autotrophic cultivation, the application of mechanical stirring resulted in a notable 1.74-fold increase in R. palustris copy number of in the ST-RP-MnO₂ nanoflower system (Fig. 2C). Confocal laser scanning microscopy (CLSM) analysis revealed a significantly higher ratio of live/dead cells in the ST-RP-MnO₂ nanoflower system (ST group, ~11:1) compared to the RP-MnO₂ nanoflower system without stirring (w/o ST group, ~2:1) (Fig. S3). To more comprehensively evaluate cell viability, we used flow cytometry (FCM) to assess cell vitality. As shown in Fig. 2D and Fig. S4, the ST group exhibited a significantly higher cell survival rate compared to the w/o ST group (91.0% vs. 62.0%). This indicates that flexoelectric electrons can substitute for traditional energy sources such as light or organic electron donors to sustain the metabolic activity of R. palustris under energy-depleted conditions. Moreover, the ST group also demonstrated higher viability than the regular heterotrophic culture (CK group) (91.0% vs. 82.0%). This difference may be explained by the distinct metabolic fates of the two groups. Following 6 days of cultivation, cells in the CK group reached the late stationary phase and underwent natural cell death due to the exhaustion of organic electron donors. In contrast, the ST group sustained a relatively lower metabolic rate by utilizing flexoelectric electrons, preserving cellular viability over a longer time frame.

Fig. 2. Microbial growth supported by mechanical energy.

A Incubation reactor. B Cell biomass (total protein) accumulation in different systems. Data are presented as the mean ± SD (n = 3 biological replicates). C Variation in gene copy numbers of R. palustris under conditions with and without mechanical stirring. Data are presented as the mean ± SD (n = 6 biological replicates). D Variations of cell survival ratio and integrity monitored by flow cytometry. Data are presented as the mean ± SD (n = 3 biological replicates). E Adenosine triphosphatase (ATP) concentration. Data are presented as the mean ± SD (n = 6 biological replicates). F Relative abundances of transcription genes involved in ATP synthesis and NADH dehydrogenase under various growth conditions. Data are presented as the mean ± SD (n = 3 biological replicates). G NADPH/NADP⁺ and NADH/NAD⁺ ratios. Data are presented as the mean ± SD (n = 3 biological replicates). Statistical analysis was conducted with paired two-tailed t-tests. *P < 0.05, **P < 0.01, ***P < 0.001.

The production of adenosine triphosphate (ATP) is essential for cellular growth⁴¹. As illustrated in Fig. 2E, the intracellular ATP concentration of R. palustris significantly increased from 0.25 to 0.31 μmol/mg protein over 6 days of growth under continuous stirring, revealing the positive impact of mechanical energy on the cellular energy metabolism. In contrast, intracellular ATP levels decreased to approximately 0.15 μmol/mg protein in cells without stirring (Fig. 2E). The analysis of key genes involved in ATP production revealed that the expression of F-type ATP synthase genes (atpA-G) was significantly higher under mechanical stirring than under non-stirring conditions (Fig. 2F and Table S2). Nicotinamide adenine dinucleotide (NAD) and its phosphorylated counterpart NADP, which exist in oxidized (NAD⁺, NADP⁺) and reduced (NADH, NADPH) forms, are essential redox cofactors in cellular energy metabolism⁴². Elevated NADH/NAD⁺ and NADPH/NADP⁺ ratios are widely recognized indicators of enhanced microbial metabolic activity⁴³. In our system, mechanical stirring of MnO₂ nanoflowers led to concurrent increases in both redox ratios (Fig. 2G and Fig. S5), demonstrating that flexoelectric stimulation provides R. palustris with the reducing power necessary to sustain active metabolism. Without mechanical stirring, the NADH/NAD⁺ and NADPH/NADP⁺ ratios both declined (Fig. 2G). Furthermore, we found higher expression levels of genes associated with reducing power substances, including NADH and NADPH, in R. palustris under stirring conditions (Fig. 2F and Table S3). The NADPH/NADP⁺ and NADH/NAD⁺ ratios in the flexoelectric culture were intermediate between those of the photoautotrophic and heterotrophic cultures (Fig. S5), suggesting an effective energy input via the flexoelectric mechanism.

Mechanical energy is converted into flexoelectric energy via MnO₂ nanoflowers

Since we did not supply any light or chemical electron donors to our system, we hypothesized that the growth of R. palustris would be enabled by direct uptake of extracellular electrical energy from the MnO₂ nanoflowers. To validate this hypothesis, we investigated the electrical properties of MnO₂ mineral. We utilized two types of MnO₂ minerals, one with a nanoflower structure and the other with a regular particle structure that lacked the flexostructure (Fig. 3A, B and Fig. S6), to assess the generation of flexoelectricity under mechanical stirring. The elemental composition, chemical states, and crystal structures of two MnO₂-based minerals were systematically investigated using X-ray diffraction (XRD) and X-ray photoelectron spectroscopy (XPS) (Fig. S7). The phase purity of the as-synthesized δ-MnO₂ nanoflowers was confirmed by XRD, with characteristic peaks corresponding to the (001), (002), (111), and (114) planes (Fig. S7). High-resolution transmission electron microscopy (HRTEM) further revealed detailed lattice fringes with spacing of 0.71, 0.36, and 0.24 nm, which were indexed to the (001), (002), and (111) planes of δ-MnO₂, respectively (Fig. S8A). The corresponding fast Fourier transform (FFT) pattern of a single nanosheet shows sharp diffraction spots, confirming its single-crystalline nature and high crystallinity (Fig. S8B). Notably, the FFT pattern exhibits a centrosymmetric 2/m point group symmetry, which implies a crystal structure composed of centrosymmetric MnO₆ octahedra⁴⁴. This structural characteristic precludes a bulk piezoelectric effect, thereby confirming that the observed response is governed by a flexoelectric mechanism. A significant strain gradient was created when a non-homogeneous strain was induced upon mechanical agitation in the δ-MnO2 nanoflowers. This strain led to relative displacement between the cationic and anionic centers in the MnO2 nanosheets, resulting in intrinsic flexoelectric polarization (Fig. S9)⁴⁵.

Fig. 3. Characterization of the MnO₂ nanoflowers.

A, B SEM images of the synthesized MnO₂ nanoflowers and MnO₂ particles. Scale bar: 2 μm. C Flexoelectric properties of MnO₂ nanoflowers, including topography, amplitude, phase images, butterfly loops and piezoelectric hysteresis loops. D Flexocurrent intensities of R. palustris-MnO₂ nanoflower and R. palustris-MnO₂ particle with increasing agitation rates (0–1000 rpm). E EPR spectra using TEMPO as a probe. F hν plot spectrum for MnO₂ nanoflowers and MnO₂ particles. G XPS valence band spectrum of MnO₂ nanoflowers and particles. H The full energy level diagram with R. palustris-MnO₂ nanoflowers mixture. Cyts represent C-type cytochromes, NFs represent nanofilaments of R. palustris such as protein nanowires and flagella, H₂ase represents hydrogenases and UQ represents ubiquinone. Experiments were repeated independently three times with similar results in A and B.

To experimentally verify the flexoelectric characteristics of MnO₂ nanoflowers, we employed piezoresponse force microscopy (PFM) to quantify the spontaneous flexoelectric polarization of the MnO₂ nanoflowers. Notably, the observed piezoresponse signals in the centrosymmetric MnO₂, a mineral material classically regarded as non-piezoelectric, can be unambiguously attributed to the flexoelectric effect³². The topography, PFM amplitude, and phase images of the MnO₂ nanoflowers are shown in Fig. 3C. The powder exhibited an average particle size <2 μm, with PFM amplitude consistent with both the topography and corresponding SEM image (Fig. 3A). The presence of varying polarization orientations was demonstrated by the distinct contrast in the phase image. The flexoresponse hysteresis loop revealed significant phase angle changes (180°) upon application of a 20 V bias voltage (Fig. 3C), confirming polarization switching. The displacement-voltage butterfly loop further demonstrated an exceptional local flexoelectric response. Finite element simulations using COMSOL Multiphysics were further employed to simulate the distribution of the resultant strain gradient and the corresponding electric response in MnO₂ nanosheets of varying dimensions. The application of shear force to the MnO₂ nanoflowers induces a significant and highly non-uniform strain field within the nanosheets (Fig. S10A). The simulation results indicate that the deformation intensity increases with decreasing nanosheet size, suggesting that smaller nanosheets are more effective in generating localized mechanical strain under the same stimulus. The strain gradients (∂ε/∂x, ∂ε/∂y, and ∂ε/∂z) exhibit a distinctly heterogeneous spatial distribution, with maximum values concentrated at the edges and tips, which are the regions of the highest curvature induced by bending deformation (Fig. S10B). This deformation resulted in a gradient-distributed electric potential field across the nanosheets, accompanied by a strong electric field at the nanosheet apex (Fig. S10C). These results demonstrated that the MnO₂ nanoflowers intrinsically exhibited excellent flexoelectric properties.

We further verified the flexoelectric properties of MnO₂ nanoflowers under mechanical stirring through electrochemical analysis and electron paramagnetic resonance (EPR) tests. As shown in Fig. 3D, we observed a strong correlation between the magnitude of the flexoelectric current and the stirring rate. Furthermore, the EPR signal of 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO), a probe for free electrons, decreased significantly under mechanical stirring (Fig. 3E), confirming efficient electron separation and transfer induced by flexoelectric polarization. Due to the absence of a flexostructure and its inherent non-piezoelectric properties, MnO₂ nanoparticles did not generate an obvious flexoelectric field and free electrons under mechanical stirring. Although flexoelectricity has been identified in many materials, the mechanism of free-carrier (electron and hole) generation is still insufficiently understood. The flexoelectric effect could disrupt local centrosymmetry of MnO₂ nanosheets, leading to the induction of spontaneous electric polarization^46,47. Meanwhile, there is generation of inner crystal flexoelectric polarization in the band-structure engineering of MnO₂ nanosheets which has been observed to facilitate the separation and transportation of electrons and holes^32,48. The nanoscale thickness and high surface area of these nanosheets enable efficient transport to MnO₂ interface and direct cellular uptake by microorganisms. The band structure properties of MnO₂ nanoflowers were determined using X-ray photoelectron spectroscopy (XPS) valence band spectra and the hν curve derived from ultraviolet diffuse reflectance spectra (Figs. 3F and 3G). These analyses revealed a band gap (E_g) of 2.75 eV for MnO₂ nanoflowers, with the conduction band (CB) and valence band (VB) edges positioned at approximately −0.63 and +2.12 eV, respectively. Mechanical stirring is expected to enhance the separation of electrons and holes in MnO₂ nanoflowers. To further elucidate the generation dynamics of free electrons and holes, additional TEMPO-based EPR experiments were conducted. We discriminated between the holes and free electrons generated via flexoelectric polarization by probing electrons and holes in acetonitrile and pure aqueous solution, respectively. Pronounced attenuation of the TEMPO signal was observed for the ST-MnO₂ nanoflower system in both solvents, confirming the simultaneous generation of electron–hole pairs (Fig. S11). Notably, the addition of glycerol (a hole scavenger) to the aqueous solution resulted in a significantly enhanced signal attenuation (Fig. S11). This enhancement demonstrates that effective suppression of electron–hole recombination extends the free-electron lifetime, further validating the flexoelectric-driven charge separation process. Moreover, using 5,5-dimethyl-1-pyrroline N-oxide (DMPO) as a spin-trapping agent, characteristic signals corresponding to ·OH were detected (Fig. S12), verifying the consumption of holes via water oxidation and the consequent generation of ·OH within the system. After applying sustained mechanical strain to the MnO₂ nanoflowers, no discernible surface potential difference was detected by Kelvin probe force microscopy (KPFM) (Fig. S13), indicating that the surface electronic states remained essentially unaltered following mechanical bending. XPS analysis further confirmed the absence of detectable changes in the surface chemical composition or the oxidation state of manganese after mechanical deformation (Fig. S14). Therefore, strain-induced surface redox reactions or defect formation are ruled out as the dominant mechanism for electron generation. All these findings collectively demonstrate that flexoelectric polarization induces the production of free electrons, aligning with the reaction mechanism illustrated in Fig. 1. Figure S15 shows the Nyquist plots for the ST and w/o ST groups, where the diameter of the semicircular segment is inversely related to the electrochemical impedance of the material. A smaller diameter indicates lower impedance and greater charge transfer capacity. Therefore, the decreased electrochemical impedance under mechanical stirring conditions suggests that mechanical stirring significantly promotes charge transfer. Based on the integrated analyses above, we propose an electron transfer pathway that enables the sustained growth and metabolism of R. palustris (Fig. 3H). Under mechanical stirring, electrons first move from the VB to the CB of MnO₂ nanoflowers. Since the reduction potential of MnO₂ nanoflowers is higher than that of hydrogenase (H₂ase, −0.3 V) and cytochrome b (0.2 V)^15,49, the electrons are then efficiently transferred from MnO₂ nanoflowers to the receptor sites on the surface of R. palustris (Fig. 3H).

Flexoelectricity drives denitrification and CO₂ fixation in R. palustris

To understand the metabolism behind flexoelectricity-driven growth in R. palustris, we examined two key pathways for energy conservation and biomass synthesis of this strain—denitrification (dissimilatory nitrate reduction) and CO₂ fixation (carbon assimilation) (Fig. 2). We directly measured the activity of these pathways by tracking the consumption of NO₃^- and the incorporation of CO₂ into biomass in the R. palustris-MnO₂ nanoflower system under mechanical agitation. As shown in Fig. 4A, the ST-RP-MnO₂ nanoflower system achieved complete degradation of NO₃⁻ within 6 days, whereas other conditions showed minimal NO₃⁻ reduction. The ST-RP-MnO₂ particle system exhibited weak NO₃⁻ reduction, demonstrating that the electron transfer to R. palustris relied on the nanosheet structure of MnO₂ nanoflowers. The release of Mn²⁺ into the culture system was found to be negligible (<0.5 mg/L) (Fig. S16), a concentration well below known inhibitory thresholds^50,51. Furthermore, control experiments confirmed that trace levels of Mn²⁺ alone did not promote microbial growth or nitrate reduction (Fig. S17). Therefore, the observed biological activity can be attributed to the flexoelectric effect, rather than to dissolved manganese species.

Fig. 4. Metabolic performances of R.

palustris driven by flexoelectric energy. A NO₃⁻ reduction by R. palustris-MnO₂ nanoflowers and various controls (ST-RP: R. palustris with stirring; ST-RP-MnO₂: R. palustris-MnO₂ particles with stirring; RP-MnO₂ nanoflower: R. palustris-MnO₂ nanoflowers without stirring; ST-MnO₂ nanoflower: MnO₂ nanoflowers with stirring; ST-RP-MnO₂ nanoflower: R. palustris-MnO₂ nanoflowers with stirring). B Typical time course of NO₃⁻ reduction with R. palustris-MnO₂ nanoflowers over five successive cycles. C The changes of nitrogenous products during the NO₃⁻ reduction with R. palustris-MnO₂ nanoflowers under mechanical agitation. D ¹⁵N-KNO₃⁻ labeled NO₃⁻ reduction with R. palustris-MnO₂ nanoflowers with (ST) or without mechanical stirring (w/o ST). E Labeled ¹³C-NaHCO₃ fixation by R. palustris-MnO₂ nanoflowers system. F Raman spectra of R. palustris grown in R. palustris-MnO₂ nanoflowers system containing ¹³C-NaHCO₃. Data are presented as the mean ± SD (n = 3 biological replicates) in (A–C).

We further determined 400 rpm and 100 mg/L to be the optimal conditions for efficient nitrate reduction over 6 days through gradient experiments of stirring speed and nanoflower dosage, and established them as the optimal parameters (Fig. S18). These optimal parameters represent a balance between maximizing the flexoelectric driving force and maintaining an effective biophysical interface. A stirring speed of 400 rpm generates sufficient shear force to induce substantial dynamic bending and generate a strong strain gradient. Lower speeds may lead to a weak strain gradient and poor polarization, whereas excessively high speeds can cause damage to cells or disrupt the nanoflower structure due to excessive shear⁵². Besides, our results showed that 100 mg/L achieved a stoichiometric balance between electron supply and microbial uptake, thereby maximizing metabolic activity. Suboptimal performance at lower concentrations may stem from insufficient electron generation and limited interfacial contact. In contrast, higher concentrations may lead to nanoflower aggregation, which diminishes the effective surface area and hinders interface contact. To confirm the sustainability of this performance, we monitored the NO₃⁻ reduction across five cycles, with periodic replenishment of NO₃⁻. Despite these repeated cycles, R. palustris maintained consistent denitrification activity (Fig. 4B). Material stability tests revealed no significant phase transformation of MnO₂ nanoflowers after 30 days of incubation with R. palustris (Figs. S13 and S14). While MnO₂ nanoflowers have been shown to be antibacterial under aerobic conditions with high-intensity ultrasonication⁵³, we observed no inhibitory effect in our system. This discrepancy is likely attributed to the combined effect of the anaerobic environment and the applied gentle mechanical stirring. Analysis of nitrogenous products revealed the generation of low-concentration NH₄⁺-N and trace amounts of NO₂^--N following 6 days of activity (Fig. 4C). Isotopic labeling with ¹⁵N-NO₃⁻ further confirmed the reduction of NO₃⁻ to gaseous N₂ and N₂O, detected at m/z = 30 (¹⁵N₂) and m/z = 46 (¹⁵N₂O), respectively (Fig. 4D).

We performed additional analyses to confirm the CO₂ fixation property of R. palustris under dark and anoxic conditions by direct uptake of extracellular electrons induced by flexoelectric polarization. R. palustris fixes CO₂ via the CBB cycle and subsequently synthesizes high-value lycopene through the methylerythritol phosphate (MEP) pathway⁵⁴. In this regard, we investigated lycopene biosynthesis in microbial metabolic processes utilizing free electrons from flexoelectric polarization. As shown in Fig. S19, the lycopene production achieved by the ST-RP-MnO₂ nanoflower system was only marginally lower than that of the photoheterotrophic cultivation, yet it significantly outperformed both the static control (RP-MnO₂ nanoflower) and the non-flexoelectric system (ST-RP). These results demonstrated that the R. palustris-MnO₂ nanoflower hybrid system enabled a robust biomanufacturing platform. In this system, mechanical energy was efficiently transduced into bioavailable electrical energy via the flexoelectric effect, offering a sustainable energy solution for bioprocesses. Isotope tracing with labeled ¹³C-NaHCO₃ showed significant incorporation of ¹³C into the biomass of the R. palustris-MnO₂ nanoflowers system under mechanical stirring, far exceeding that of the control without stirring (Fig. 4E and Fig. S20). The shift in Raman spectra from cells grown in medium with ¹³C-NaHCO₃ highlighted a successful change in biomass accumulation. This was evidenced by the red-shift from 1507.27, 1148.62, and 1005.7 cm⁻¹ for ¹²C cells to 1505.71, 1146.99, and 1002.39 cm⁻¹ for ¹³C-labeled cells (Fig. 4F).

Transcriptional response of R. palustris to flexoelectricity

To elucidate the mechanisms underlying flexoelectricity-driven growth and metabolic activity in R. palustris, we performed transcriptomic profiling within the biohybrid system. Principal component analysis (PCA) revealed a clear separation between transcriptional profiles of cells subjected to agitated and static conditions (Fig. 5A). Transcriptional analysis identified 211 upregulated and 150 downregulated genes, indicating a substantial reprogramming of gene expression in response to flexoelectric cues (Fig. 5B). The broad upregulation of genes encoding cytochromec (Cytc) proteins was among the most notable changes under mechanical agitation (Fig. 5C, Fig. S21), and Table S4). For instance, the gene encoding cytochromec oxidase subunit IV showed a log₂(fold change) of 2.57. This coordinated induction suggests that Cytc proteins act to uptake electrons generated by flexoelectric MnO₂ nanoflowers, serving as a critical interface that transduces extracellular physical stimuli into intracellular electron transport pathways⁵⁵. Concurrently, hydrogenase (H₂ase)-encoding genes were significantly upregulated under agitation (Fig. 5C). Notably, no H₂ was detected in the biohybrid culture system (Fig. S22), and it has been reported that H₂ cannot support R. palustris growth under our dark, anoxic conditions³⁹. Therefore, the upregulated H₂ase genes should not due to the presence of H₂ metabolism in this system. Instead, given that H₂ase is known to mediate direct electron transfer between intracellular enzymes and extracellular acceptors⁵⁶, the observed gene upregulation can be attributed to its role in shuttling electrons from the flexoelectric materials to intracellular electron transfer components. Beyond redox proteins, genes encoding nanofilaments (nanowires and flagella) were significantly upregulated (Fig. 5C, Fig. S21), aligning with their reported function in electron exchange with extracellular acceptors/donors⁵⁷. As shown in Fig. 5C, the upregulation of genes encoding the proton-pumping cytochrome bc₁ complex (Complex III) and cbb₃ oxidase was also observed (Fig. 5C), indicating the enhanced respiratory activity. This suggests that the harvested electrons are channeled into the quinone pool, driving these complexes to promote proton translocation across the membrane, thereby establishing the proton motive force (PMF)⁵⁸. This PMF powered the concurrently upregulated F-type ATP synthase to generate ATP (Fig. 5C, Table S4), transducing mechanical energy into biochemical energy to drive active anabolism. The functional indispensability of this electron transfer chain and the resultant PMF for flexoelectric-driven metabolism was further verified by inhibitor assays (Fig. S23A). Inhibition of the cytochrome bc₁ complex by antimycin A, dissipation of the PMF by carbonyl cyanide m-chlorophenyl hydrazine (CCCP), or blockage of electron entry into the quinone pool by rotenone reduced nitrate reduction by 56.41%, 66.42%, and 86.21%, respectively (Fig. S23B). The Calvin–Benson–Bassham (CBB) cycle is established as a central pathway for bacterial carbon fixation in R. palustris^38,39. Transcriptional activity of the CBB cycle was markedly enhanced under flexoelectric stimulation. RubisCO genes were upregulated, reaching a log₂(fold change) of 1.0 (Fig. 5C, Table S4), which indicates enhanced carbon fixation. Moreover, denitrification operons (nar, nir, nor, and nos) exhibited significant transcriptional enhancement, with peak log₂(Fold Change) values of 0.63, 4.05, 0.68, and 0.39, respectively (Table S4). These operons encode nitrate reductase, nitrite reductase, nitric oxide reductase, and nitrous oxide reductase, respectively⁵⁹. Collectively, these results demonstrate that the flexoelectric effect effectively drives R. palustris metabolism by upregulating key genes for extracellular electron uptake, carbon fixation, and denitrification.

Fig. 5. Transcriptomic profiling.

A Principal component analysis (PCA, 95% confidence interval). B Volcano plot of differentially expressed genes in R. palustris in the R. palustris-MnO₂ nanoflowers system under stirred versus static conditions. (C) Differential expression of key genes associated with electron transfer and energy metabolism was compared among mechanically agitated and static nanoflower systems. Analyzed components included: (i) c-type cytochromes (Cyts), (ii) protein nanowires/flagella (NFs), (iii) hydrogenase (H₂ase), (iv) ATP synthases, (v) NADH dehydrogenase complex (Complex I), and (vi) cytochrome bc₁ complex (Complex III), with Q (quinone) and QH₂ (quinol) indicated. Full form of abbreviations: 3-phosphoglycerate (3-PGA), 1,3-bisphosphoglycerate (1,3-BPG), fructose 1,6-bisphosphate (FBP), fructose 6-phosphate (F6P), xylulose 5-phosphate (X5P), ribulose 5-phosphate (Ru5P), ribose 5-phosphate (R5P), ribulose 1,5-bisphosphate (RuBP), dihydroxy-acetone-phosphate (SDP), sedoheptulose-7-phosphate (S7P), glyceraldehyde 3-phosphate (G3P), dihydroxyacetone phosphate (DHAP), and sedoheptulose-1,7-bisphosphate (SBP), nar (nitrate reductase), nir (nitrite reductase), nor (nitric oxide reductase), and nos (nitrous oxide reductase). Corresponding transcriptomic data are presented in Table S4. All P values are provided in the Source Data.

Harnessing of flexoelectricity in diverse microbe-mineral biohybrids

In exploring microbial metabolism driven by flexoelectric effects, we expanded our investigation to include other flexoelectric mineral materials such as SnO₂, WS₂ and birnessite nanoflowers. The morphology and flexoelectric responses of these minerals are characterized by SEM and electrochemical measurements, respectively. All materials exhibited well-defined nanoflower structures and generated significant remarkable flexocurrents under mechanical stirring (Figs. S24 and S25). After 6 days of mechanical stirring, the NO₃⁻-N reduction efficiencies were 40.9% for the R. palustris-SnO₂, 66.1% for the R. palustris-WS₂, and 87.3% for the R. palustris-birnessite, respectively (Fig. 6A). These metabolic enhancements were consistently observed across all these materials under identical experimental conditions. Additionally, our results demonstrated that alternative forms of mechanical stimulation, including gentle ultrasound and vibration, could also be utilized to support nitrate reduction metabolism in the R. palustris-MnO₂ nanoflower system (Fig. 6A). Further experimentation included testing different microbial species to determine their ability to harness flexoelectric polarization-induced free electrons for metabolic activities. Under mechanical stimulation, G. sulfurreducens and S. oneidensis effectively harnessed flexoelectric polarization-induced electrons from MnO₂ nanoflowers, achieving 22.3% fumarate reduction and 43.9% NO₃⁻ removal, respectively (Fig. 6A). To validate EEU in this biohybrid system, we employed an S. oneidensis ΔomcA-ΔmtrC double mutant deficient in key outer-membrane cytochromes essential for direct electron transfer. This mutant exhibited a 65.63% decrease in NO₃^- reduction compared to the wild type (Fig. S26), confirming the essential role of EEU in harvesting flexoelectric-generated electrons. In contrast, non-electroactive strains lacking EEU capabilities, such as Escherichia coli and Bacillus subtilis, were unable to exploit mechanical energy for growth (Fig. 6A). These findings collectively suggest that the electronic energy derived from mechanical input by flexoelectric minerals can be universally harnessed by electroactive microbes (Fig. 6B), potentially playing a pivotal role in driving the biogeochemical cycles of Earth regardless of their extracellular electron transfer proficiency.

Fig. 6. Microbial growth and metabolism driven by the flexoelectric effect based on diverse systems.

A R. palustris-different flexoelectric minerals (SnO₂, WS₂, birnessite and MnO₂ nanoflower) and mechanical stirring, different mechanical forces (mechanical stir, vibration, and weak ultrasound) in the R. palustris-MnO₂ nanoflowers system, different electroactive microorganisms (G. sulfurreducens, S. oneidensis and R. palustris)-MnO₂ nanoflowers and mechanical stirring and cell growth in different microorganisms (B. subtilis, E. coli and R. palustris)-MnO₂ nanoflowers system. B Schematic diagram of mechanism of flexoelectric effect acting on non-electroactive microorganisms and electroactive microorganisms. Data are presented as the mean ± SD (n = 3 biological replicates). Statistical analysis was conducted with paired two-tailed t-tests. *P < 0.05, **P < 0.01, ***P < 0.001.

Discussion

R. palustris typically grows in an autotrophic manner, utilizing CO₂ as a carbon source and drawing energy from light and inorganic electron donors⁶⁰. This study introduces an innovative mechanical-to-metabolic coupling pathway through energy transduction via the flexoelectric effect. By converting mechanical energy into bioavailable electrons, MnO₂ nanoflowers effectively support R. palustris growth, enabling the continued growth of these bacteria through the coupling of CO₂ fixation and nitrate reduction. This unique mechanism applies broadly to other EAMs and a variety of flexoelectric minerals. Preliminary calculations reveal total mechanical energy conversion efficiencies of approximately 1.12‰, 0.36‰, and 0.74‰ for the R. palustris-MnO₂ nanoflowers, S. oneidensis-MnO₂ nanoflowers, and G. sulfurreducens-MnO₂ nanoflowers systems, respectively (Supplementary Text and Tables S5 and S6). Although these efficiency values remain relatively low, this energy can be harvested from ambient mechanical sources. Therefore, this flexoelectric-driven microbial energy capture offers a promising route for powering life in energy-scarce environments like deep-sea vents and subsurface sediments. Notably, under comparable culture conditions, R. palustris exhibited slightly higher growth and metabolic efficiency with flexoelectric stimulation than with piezoelectric stimulation (Table S7). Beyond efficiency, flexoelectricity offers broader material applicability, including centrosymmetric and high-temperature stable materials³¹. This fundamental distinction likely positions flexoelectricity as a complementary and even versatile pathway for harnessing ubiquitous mechanical energy to power microbial processes in complex natural environments. This mechanism also reveals a distinct microbe-mineral interaction mode driven by the energy transduction process, facilitating direct energy exchange between minerals and microbes.

The transduction of mechanical energy by microorganisms, with factors such as shear stress, hydrostatic pressure, and vibrational forces having direct implications on metabolic processes, represents a burgeoning area of interest within microbial ecology^61,62. This emerging mechanobiological paradigm is revolutionizing our understanding of microbial adaptation across diverse Earth ecosystems. In marine biofilms, fluid shear stress critically influences community assembly by activating metabolic pathways in both prokaryotic and eukaryotic organisms, thereby facilitating structural reorganization that optimizes nutrient transfer⁶². This hydrodynamic-microbial interaction, driven by disturbance shear, can significantly influence marine biofilm community success, acting as a selective filter that is influenced by the frequency of shear exposure⁶³. Leveraging foundational insights into microbial mechano-sensitivity has led to various industrial applications. For instance, intermittent weak ultrasound could induce microbial adaptation and physiological regulation to enhance pollutant removal efficiency and reduce membrane fouling in ultrasound-coupled membrane bioreactor systems⁶⁴. In industry, further refinement through targeted hydrodynamic management has been employed to improve bioprocessing efficiency⁶⁵. Collectively, these findings underscore hydrodynamic parameters as pivotal variables governing microbial ecology across scales, from biofilm microenvironments to nutrient fluxes of ocean basin. However, the mechanisms underlying the mechanical-to-metabolic coupling in these systems remains unknown. Specifically, from an energy perspective, it is unclear whether these interactions involve energy transduction via direct electron transfer, which is a crucial aspect of microbial metabolism. Herein, we have provided evidence that a mechanical force, through the flexoelectric effect, can drive the growth and metabolism of certain microorganisms, thereby advancing the understanding of interactions between microorganisms and their surrounding environment.

Our current study emphasizes the potential of flexoelectric minerals in supporting microbial processes in environments characterized by mechanical forces, such as coastal areas and industrial zones, opening avenues for bioremediation and environmental conservation. However, as described in Fig. 6, only electroactive microorganisms possessing extracellular electron uptake capabilities can effectively harness this unique extracellular flexoelectric-derived energy for growth and metabolic activities. Despite their pivotal roles in biogeochemical cycles, electroactive microorganisms constitute merely a niche subset within the vast microbial biosphere. The vast majority of microorganisms are unable to access extracellular electron sources, which fundamentally limits the widespread adoption and application of extracellular flexoelectric energy conversion to non-electroactive microbial populations. From an engineering standpoint, the direct integration of flexoelectric minerals into microbial cells holds great promise for enabling intracellular mechanical-to-metabolic coupling in non-electroactive microbes, thus eliminating the need for extracellular electron transfer. This strategy could significantly expand the scope of bioenergy applications of flexoelectric effects by tapping into a wide variety of microorganisms that do not naturally engage in extracellular electron exchange. From a nature perspective, microorganisms have an innate ability to spontaneously internalize nanomaterials⁹, which provides a biologically relevant and practical framework for flexoelectric energy harvesting among non-electroactive microbes. In this regard, microbial cells are capable of harnessing intracellular flexoelectric energy for metabolism by means that have yet to be fully recognized in natural environments. Given that the efficiency of flexoelectric effects is known to increase at the nanoscale, it is plausible that this mechanism is not only viable but also potentially widespread in various microbial life forms. Therefore, future in-depth investigations into this phenomenon should be able to fully uncover its potential and prevalence within the microbial world.

Methods

Synthesis of flexoelectric minerals

The hierarchical MnO₂ nanostructures were fabricated through a modified hydrothermal approach employing stoichiometrically controlled precursors (KMnO₄:KCl = 3:1 molar ratio)³². In a standard synthesis protocol, potassium permanganate (KMnO₄, 0.4741 g, 3.00 mmol, ≥99.0%, Sigma-Aldrich) and potassium chloride (KCl, 0.0746 g, 1.00 mmol, ≥99.5%, Sigma-Aldrich) were co-dissolved in 40 mL Milli-Q water (18.2 MΩ·cm) under ambient conditions. The homogeneous solution was magnetically stirred (500 rpm) for 10 min. The precursor solution was then transferred to a 100 mL Teflon-lined stainless steel autoclave and heated at 180 °C for 13 h in a convection oven. After cooling to room temperature, the product was collected by centrifugation (6000 × g, 10 min), washed three times with ethanol, and dried under vacuum at 60 °C for 12 h. The isolated mass was 0.215 g, calculated based on the manganese content assuming complete conversion of KMnO₄ to δ-MnO₂, with a theoretical yield of 0.261 g and an isolated yield of 82.4%. The detailed synthesis procedures for the other several nanoflower materials are provided in the SI Text.

Bacterial culture

R. palustris strain (GDMCC 1.485) was obtained from the Guangdong Microbial Culture Collection Center and initially cultured in 50 mL of mineral medium (10% v/v inoculum ratio, see Table S1). Primary cultivation was conducted in borosilicate glass reactors maintained at 30 ± 0.5 °C using a constant temperature incubator (Model LRH-1500F, Bluepard Instruments Co., Ltd., Shanghai). Four other bacterial strains, including G. sulfurreducens ATCC 51573, S. oneidensis ATCC 700550, E. coli BAA-3219, and B. subtilis ATCC 6051, were obtained from the American Type Culture Collection (ATCC). A pre-culture of R. palustris was prepared by inoculating 50 mL of heterotrophic medium (Table S1; pH 7.0 ± 0.1) at a 1:20 ratio, followed by incubation at 30 °C with LED illumination (20 W/m²)⁶⁶. G. sulfurreducens was cultured under strict anaerobic conditions (80% N₂, 20% CO₂) at 30 °C in NBAF medium supplemented with 20 mM sodium acetate and 40 mM sodium fumarate⁶⁷. S. oneidensis, E. coli, and B. subtilis were cultured aerobically in Luria-Bertani (LB) broth at 30 °C or 37 °C, respectively, until mid-exponential growth phase. Cells were harvested by centrifugation (6000 × g, 10 min, 4 °C) and washed three times with sterile 0.9% NaCl solution (anaerobic for Geobacter; aerobic for others) to remove residual organic substrates. The final cell pellets were resuspended in a minimal volume of saline solution.

Cell incubation under mechanical force

The washed R. palustris cells were then transferred into autotrophic medium (Table S1), which had been pre-supplemented with flexoelectric mineral and nitrate, excluding organic electron donors and adjusted pH to 7.0 ± 0.1. The Final concentrations were 5 mg of flexoelectric material per 50 mL and 20 mg/L nitrate. This cell-mineral suspension, designated R. palustris-MnO₂ nanoflowers with stirring (ST-RP-MnO₂ nanoflower), was subjected to dark starvation for 24 h at ambient temperature. To establish anaerobic conditions, the headspace of serum bottles (125 mL) was purged with filter-sterilized N₂/CO₂ gas mixture (80:20, v/v) for 20 min, followed by sealing with butyl rubber stoppers. The bottles were then placed on a magnetic stirrer (SP200-2T, MIULAB) operating at 400 rpm and incubated under continuous darkness at 30 °C. Control systems, including R. palustris with stirring (ST-RP), MnO₂ nanoflowers with stirring (ST-MnO₂ nanoflower), and R. palustris-MnO₂ nanoflowers without stirring (RP-MnO₂ nanoflower), were prepared identically. An additional control, ST-RP-MnO₂ particle, was established by replacing MnO₂ nanoflowers with commercial MnO₂ particles (AR grade, Tianjin Damao Chemical Reagent Co. Ltd., China). These particles lack the hierarchical nanoflower architecture and intrinsic flexoelectric response. The biohybrid systems with other strains (G. sulfurreducens, S. oneidensis, E. coli, and B. subtilis) or flexoelectric materials (SnO₂, WS₂, or birnessite) were setup in the same autotrophic basal medium used for R. palustris (Table S1). For strain-specific cultivation, G. sulfurreducens was supplied with 40 mM fumarate as the electron acceptor, whereas the other strains received 20 mg/L nitrate. Other flexoelectric minerals were also supplied at a final concentration of 100 mg/L. The methods for synthesizing these materials are described in Supplementary Information Text (SI Text). The bottles were purged with N₂/CO₂ (80:20, v/v) for 20 min, sealed with butyl rubber stoppers, and incubated on a magnetic stirrer (400 rpm) at 30 °C in the dark.

To investigate the influence of key parameters, the effects of MnO₂ nanoflower concentration and stirring speed on nitrate reduction performance were systematically examined within the ST-RP-MnO₂ nanoflower system. For cyclic nitrate reduction experiments, R. palustris-MnO₂ nanoflowers were cultivated over five consecutive 6-day cycles (total 30 days) under continuous stirring (400 rpm). At the end of each cycle, the spent medium was replaced under strict anaerobic conditions inside an anaerobic chamber (MITR, MT002-C). The cell-mineral suspension was gently centrifuged (6000 × g, 5 min), the supernatant was discarded, and the pellet was resuspended in 50 mL of fresh, pre-reduced autotrophic basal medium containing 20 mg/L NO₃⁻. The resuspended culture was transferred to a new sterile serum bottle, purged with N₂/CO₂ (80:20, v/v) for 20 min, sealed, and returned to incubation conditions to initiate the next cycle.

Characterization

Microstructural characterization of the R. palustris-MnO₂ nanoflowers mixture interface was performed using scanning electron microscopy (SEM, MIRA LMS, TESCAN, 5 kV accelerating voltage). Specimen preparation involved chemical fixation with 2.5% glutaraldehyde (12 h, 4 °C) followed by ethanol gradient dehydration (25–100% v/v, 15 min/step) and freeze drying. Elemental mapping was conducted via energy-dispersive X-ray spectroscopy (EDS, Quantax 200/XFlash^®6-60, Bruker) coupled with SEM at 15 kV. For crystallographic analysis, samples were subjected to X-ray diffraction (XRD, D8 ADVANCE, Bruker) with Cu Kα radiation (λ = 1.5406 Å, 40 mA, 40 kV) over 10–80° 2θ range. Chemical speciation was investigated through complementary spectroscopic techniques, including X-ray photoelectron spectroscopy (XPS, Escalab 250Xi, Thermo Scientific) with a monochromatic Al Kα source (1486.6 eV), UV–vis–NIR spectroscopy (UV-2600, Shimadzu) for Tauc plot derivation (200–1200 nm), and Kelvin probe force microscope (KPFM, Bruker Dimension Icon, USA). Flexoelectric properties were assessed via switching spectroscopy piezoelectric force microscopy (SS-PFM, MFP-3D Origin+, Oxford Instruments) using Pt/Ir-coated tips (AC240TM, 70 kHz resonance). For the flexoelectric simulation in this study, a coupled analysis of solid mechanics and electrostatics was performed using COMSOL Multiphysics, which was detailed in SI Text. Electrochemical performance evaluation was performed using a three-electrode system (CHI660E, Shanghai Chenhua Instrument Co., LTD., China). The working electrode was a polished boron-doped diamond substrate (1 × 2 cm²). The electrochemical cell configuration included a platinum sheet (1 × 1 cm²) and saturated calomel reference electrode (SCE, 0.241 V vs. SHE at 25 °C). Chronoamperometry was executed under programmed rotational control (0-1200 rpm cyclic modulation, 400 s/cycle) with simultaneous potential tracking. Electrochemical impedance spectroscopy (EIS) measurements spanned 10⁶–10⁻² Hz with a sinusoidal perturbation amplitude of 5 mV under an open-circuit potential.

Cell viability tests

To confirm microbial proliferation under mechanically stimulated denitrification conditions, R. palustris-MnO₂ nanoflower mixtures cultivated for 30 days under continuous agitation were harvested. These mixtures were subsequently processed through sequential centrifugation, washed with 0.9% saline solution (NaCl), and finally resuspended in 5 mL of identical saline. Total protein was then isolated from the collected mixture utilizing a One-Step Bacterial Active Protein Extraction Kit (Sangon Biotech). Protein concentration was quantified using a multimode microplate reader (SpectraMax iD3, Molecular Devices) in conjunction with the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). The cell viability was evaluated via staining with the Live/Dead BacLight Bacterial Viability Kit (Invitrogen). Stained samples were visualized using a confocal laser scanning microscope (CLSM, LSM880, ZEISS) and imaged data were processed with ZEN software. Intracellular ATP levels were quantified using a commercial assay kit (micro-method, Mlbio, ml076315). The redox states were assessed by determining the ratios of NADH/NAD⁺ and NADPH/NADP⁺ using specific WST-8-based assay kits (Beyotime, S0175 and S0179, respectively).

The flow cytometry protocol commenced with homogenization of bacterial suspensions. Aliquots (1 mL) were transferred to 2 mL centrifuge tubes and centrifuged at 8000 × g for 3 min; the supernatant was discarded. The resulting pellets were resuspended in 1 mL phosphate-buffered saline (PBS), subjected to a tenfold dilution, and 200 μL of this diluted suspension was aliquoted into fresh 2 mL tubes followed by vortex mixing. Each suspension was then divided equally. One portion served as the native control without treatment, while the other was heat-treated at 100 °C for 5 min. After treatment, the two portions were recombined and subdivided into three equal aliquots designated as follows: 1) an unstained Blank control; 2) a sample stained with 2 μL of 10× diluted SYTO 9; and 3) a sample stained with 2 μL of 10× diluted SYTO 9 plus 2 μL of 10× diluted PI. All other tubes received both SYTO 9 and PI stains (2 μL each of 10× diluted solutions). Flow cytometry (Guava^®EasyCyte^TM, Luminex) analysis utilized the Green-B (Ex/Em: 485/498 nm) and Red-B (Ex/Em: 535/617 nm) channels to quantify apoptosis dynamics.

Mechanical energy-driven denitrification experiments

Denitrification kinetics were conducted in triplicate batch reactors (125 mL serum bottles) that were sealed with Teflon®-coated rubber and aluminum crimp caps. Each reactor received 50 mL of sterile basal medium (composition in Table S1) supplemented with 5 mg of flexoelectric materials and continuously purged with N₂/CO₂ (80:20 v/v, 0.22 μm filtered) at 5 mL/min for 20 min. These serum bottles were then sealed and placed on the magnetic stirrer (SP200-2T, MIULAB) with an agitation speed of 400 rpm. Control experiments were conducted under the same conditions, with systematic removal of R. palustris, flexoelectric materials or mechanical agitation. The stirring speed and flexoelectric materials dosage in the biotic-abiotic mixture system were also sequentially optimized.

Nitrogen speciation dynamics during nitrate reduction were quantitatively tracked. Aqueous phase nitrogen species ((NO₃⁻-N, NO₂⁻-N, NH₄⁺-N) were analyzed via UV–Vis spectrophotometry (UV-2600, Shimadzu) following standard methods: cadmium reduction (450 nm) for NO₃⁻, diazotization (540 nm) for NO₂⁻, and salicylate-hypochlorite (655 nm) for NH₄⁺. Gaseous N₂O was quantified using gas chromatography (7890B, Agilent) equipped with electron capture detector (ECD) and Porapak Q column (80–100 mesh), operated isothermally at 80 °C with Ar/CH₄ (95:5) carrier gas (1.2 mL/min). Molecular nitrogen was determined via thermal conductivity detector (TCD) with pre-column CO₂ trap (Ascartic II, Sigma).

Extraction and determination of lycopene

R. palustris cells were collected via centrifugation at 6000 × g for 6 min. Following supernatant removal, the pellet underwent two washes with 0.9% NaCl solution. The washed cells were subsequently dried at 70 °C for 24 h. Lycopene extraction employed 3 mL of n-hexane/methanol (1:1, v/v). Cellular debris was separated by centrifugation at 8000 × g for 10 min at 4 °C, and the resulting supernatant was subjected to high-performance liquid chromatography (HPLC) for lycopene quantification. HPLC separation utilized a WondaSil^TM C18 column (4.6 × 150 mm, 5 µm) under isocratic conditions with a mobile phase consisting of methanol, acetonitrile, and dichloromethane (40:40:20, v/v/v). The flow rate was maintained at 1 mL/min, the column temperature at 30 °C, and detection occurred at 472 nm. The injection volume was 5 μL.

Isotope analysis

Dual-isotope labeling was performed using NaH¹³CO₃ (99 atom% ¹³C, Cambridge Isotope Laboratories) and K¹⁵NO₃ (99 atom% ¹⁵N, Sigma-Aldrich). Following cryo-fixation (liquid N₂ slush), samples were freeze-fractured and coated with 5-nm gold (Quorum Q150T ES). Isotopic composition was assessed by GC-MS (Agilent 7890–5975c) and ToF-SIMS (TOF-SIMS IONTOF M6). SurfaceLab 6.7 software enabled multivariate analysis of ToF-SIMS hyperspectral images (256 × 256 pixels). Post-harvest cell samples were washed in deionized water for Raman microscopy. Suspensions were dried on CaF₂ slides prior to analysis. Raman spectra were obtained on a LabRAM HR Evolution confocal system (HORIBA Jobin Yvon) fitted with a 100× objective (NA 0.90). Precise single-cell laser positioning (532 nm Nd:YAG laser, Torus; ~2.5 mW) was achieved using a motorized stage (0.1 mm step precision) synchronized with an onboard camera. The spectral range 100–2000 cm⁻¹ was examined.

Transcriptomic analysis

Transcriptomic analysis of the R. palustris-MnO₂ nanoflowers system employed Illumina sequencing (HiSeq/MiSeq platforms). Biomass was harvested by differential centrifugation (10,000 × g, 30 min, 4 °C) during mid-exponential growth. Total RNA extraction utilized Invitrogen TRIzol reagent (USA), followed by rRNA depletion via subtractive hybridization. RNA libraries were constructed using the NEBNext Ultra II Directional RNA Library Prep Kit (New England Biolabs, China). After quality control and filtering of raw sequencing reads, sequences aligning with 16S and 23S rRNA genes were excluded. The remaining reads were aligned to the R. palustris CGMCC 1.2180 reference genome (NZ_CP058907.1). Differential gene expression analysis was conducted using DESeq2. Significance was tested using the Wald test, and p-values were adjusted for multiple comparisons using the Benjamini-Hochberg method to control the false discovery rate (FDR). Expression levels were quantified as FPKM (Fragments Per Kilobase per Million mapped reads) for visualization.

qPCR analysis

Quantitative real-time PCR (qPCR), with primers RpFor (5′-CTGGAAGTCTTGAGTATGGC-3′) and RpRev (5′-AGTAAACCCACTAACGGCTG-3′), was employed to determine 16S rRNA gene copy numbers in planktonic R. palustris. The 20 μL reaction mixture comprised 1 μL DNA template, 0.8 μL each of 5 μM forward and reverse primers, 10 μL 2× Taq Plus Master Mix, and 7.4 μL ultrapure water. Amplification conditions consisted of initial denaturation at 95 °C for 5 min, followed by 35 cycles of denaturation (95 °C, 30 s), annealing (58 °C, 30 s), and extension (72 °C, 40 s).

Reporting summary

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

Author contributions

Y.L., Y.Y. and S.Z. conceived the idea for this work. Y.L. and J.W. cultured the microbial strains. Y.L., J.W. and G.R. performed the characterizations, catalytic measurements and transcriptomic experiments. Y.L. and Y.Y. analyzed the data. Y.L., Y.Y. and S.Z. wrote the manuscript. All the authors contributed to the interpretation of the data and preparation of the manuscript.

Peer review

Peer review information

Nature Communications thanks Guibing Zhu, Junyi Zhai, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Funding

S.Z. discloses support for the research of this work from the National Natural Science Foundation of China [grant number 42525702]. Y.Y. discloses support for publication of this work from the National Natural Science Foundation of China [grant number U24A20624], the Basic and Applied Basic Research Foundation of Guangdong Province [grant number 2023B1515040022]. Y.L., J.W., and G.R. declare no relevant funding.

Data availability

All data supporting the findings of this study are available within the paper and its Supplementary Information files. The RNA-seq data generated in this study have been deposited in the NCBI Trace Archive database under accession code PRJNA1438059. Source data are provided with this paper. All other data underlying this study are available from the corresponding author upon request.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-74030-y.