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Living materials fabricated via gradient mineralization of light-inducible biofilms


Living organisms have evolved sophisticated cell-mediated biomineralization mechanisms to build structurally ordered, environmentally adaptive composite materials. Despite advances in biomimetic mineralization research, it remains difficult to produce mineralized composites that integrate the structural features and ‘living’ attributes of their natural counterparts. Here, inspired by natural graded materials, we developed living patterned and gradient composites by coupling light-inducible bacterial biofilm formation with biomimetic hydroxyapatite (HA) mineralization. We showed that both the location and the degree of mineralization could be regulated by tailoring functional biofilm growth with spatial and biomass density control. The cells in the composites remained viable and could sense and respond to environmental signals. Additionally, the composites exhibited a maximum 15-fold increase in Young’s modulus after mineralization and could be applied to repair damage in a spatially controlled manner. Beyond insights into the mechanism of formation of natural graded composites, our study provides a viable means of fabricating living composites with dynamic responsiveness and environmental adaptability.

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Fig. 1: Engineering light-responsive E. coli functional biofilms as scaffolds for HA mineralization.
Fig. 2: Spatially controllable mineralization of light-inducible biofilms for living patterned composites.
Fig. 3: Density-controllable mineralization in light intensity-regulated gradient biofilms used to fabricate living graded composites.
Fig. 4: Light intensity-regulated biofilm biomass density dictates the mineral density and the eventual mechanical properties of the resultant living graded composites.
Fig. 5: Coupling engineered biofilms with mineralization for robust capturing and immobilization of microspheres on substrates.
Fig. 6: Application of controllable living mineralization for site-specific damage repairs.

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Data availability

The main data supporting the findings of this study are available within the article and its Supplementary Information. Additional data are available from the corresponding author upon reasonable request.


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We thank X. Wang for AFM training and W. Liu and Y. Jiang for TEM and SEM training, respectively. AFM characterization was executed at the Analytical Instrumentation Center (AIC); SEM and TEM experiments were supported by the Center for High-resolution Electron Microscopy (CћEM) at ShanghaiTech University. We also thank K. Kang and W. Xing from the animal core facility at the Shanghai Institute of Biochemistry and Cell Biology for helping with µCT experiments and image analysis. This work was partially sponsored by the National Science and Technology Major Project of the Ministry of Science and Technology of China (grant no. 2020YFA0908100 and grant no. 2018YFA0902804), the Joint Funds of the National Natural Science Foundation of China (key program no. U1932204), the Commission for Science and Technology of Shanghai Municipality (grant no. 17JC1403900) and the China Postdoctoral Science Foundation (grant no. 2019M661676). C.Z. also acknowledges start-up funding support from the 1000 Youth Talents Program, granted by the Chinese Central Government.

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Authors and Affiliations



C.Z. conceived the concept and directed the research. C.Z., Y.W. and B.A. designed and conducted the experiments and data analysis. Y.C. and B.X. participated in AFM nano-indentation and SMFS experiments and relevant data analysis. J.P. conducted the QCM experiments and analyzed the resulting data. Y.Y. and X.Z. contributed to acquiring selected-area electron diffraction patterns and relevant data analysis. Y.H. assisted with western blot experiments and the cell viability assay. C.Z., Y.W. and B.A. wrote the manuscript with help from all authors.

Corresponding author

Correspondence to Chao Zhong.

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Extended data

Extended Data Fig. 1 Comparison of biofilm formation for genetically engineered E.coli strains that express different CsgA-Mfp fusion proteins.

a, TEM characterization of biofilms formed by E.coli strains consisting of different CsgA-Mfp fusion proteins. Scale bars, 500 nm. b, Quantitative Congo Red (CR) binding assay and Crystal Violet (CV) staining revealed that few amounts of biofilm biomass were produced by aTcReceiver/CsgA-Mefp5, while aTcReceiver/CsgA-Mfp3S-pep yielded almost the same amount of biofilm biomass as that of the aTcReceiver/CsgA biofilms. c, Biofilm growth monitored by quantitative CR binding assay revealed that aTcReceiver/CsgA-Mfp3S exhibited a much slower growth rate and produced lower amounts of biofilm biomass compared with both the CsgA and CsgA-Mfp3S-pep biofilms. Results in b,c are presented as Mean ± s.d. Data are representative of n = 4 independent experiments.

Extended Data Fig. 2 Comparison of HA precipitation on CsgA and CsgA-Mfp3S-pep nanofibers.

TEM images of the precipitated minerals on different protein nanofibers after mineralization for 1, 3, 5, and 7 days. Scale bars, 500 nm.

Extended Data Fig. 3 Comparison of HA crystallization on CsgA and CsgA-Mfp3S-pep biofilms.

TEM images and corresponding SAED patterns of the precipitated minerals on different biofilms after 5-day mineralization.

Extended Data Fig. 4 Adsorption behavior comparison of CsgA and CsgA-Mfp3S-pep on HA surfaces measured by QCM.

a, Adsorption behavior of the CsgA and CsgA-Mfp3S-pep monomers (0.5 mg/mL) on HA-coated QCM chip. b, Adsorption behavior of the CsgA and CsgA-Mfp3S-pep nanofibers (initial monomer concentration at 0.5 mg/mL) on HA-coated QCM chip.

Extended Data Fig. 5 SMFS experiments for quantifying the interactions between the Mfp3S-pep proteins and hydroxyapatite.

a, Schematic of the AFM single-molecule spectroscopy experiments. The SNAP-Mfp3S-pep was connected to the cantilever tip via an O6−benzylguanine (BG)-terminated PEG linker. The substrate was hydroxyapatite-coated. b, Typical force-extension curves of Mfp3S-pep on HA-coated surfaces. Force-extension curves were fitted with WLC (green). All data are collected at the pulling speed of 1000 nm·s-1. c, Histograms (bars, N = 107) and Gaussian function (blue) fitting of the interaction strength between the Mfp3S-pep and HA. d, Histograms (bars, N = 107) and Gaussian function (black) fitting of the fracture length for the rupture of the Mfp3S-pep and HA interactions. e, Schematic of the AFM single-molecule spectroscopy experiments for the interaction between SNAP and HA. f, Typical force-extension curves for the interaction between the SNAP and HA. All data are collected at the pulling speed of 1000 nm·s-1.

Extended Data Fig. 6 TEM observation of morphology and phase evolution.

TEM images (Top) and corresponding SAED patterns (bottom) of the extracellularly self-assembled CsgA-Mfp3S-pep nanofibers (0 day) and the precipitated minerals on nanofibers after mineralization of the CsgA-Mfp3S-pep biofilm samples for 1, 3, 5, and 7 days. Scale bars, 500 nm.

Extended Data Fig. 7 Various illumination images (concentric circles, grids) are recapitulated as patterned biofilms (top) and patterned composites.

The grid pattern of the mineralized composite has a 1 mm spatial resolution. Scale bars, 1 cm.

Extended Data Fig. 8 Mineral contents of mineralized composites with different incubation time determined by TGA analyses.

a, Representative TGA measurements of the dried biofilm samples and living composite samples after mineralization for 1, 3, 5, and 7 days. b, Inorganic contents in the living composites with different mineralization time calculated based on TGA analyses. Results are presented as Mean ± s.d. Data are representative of n = 3 independent experiments.

Extended Data Fig. 9 Mechanical properties of biofilms and mineralized composites.

a, Young’s modulus of biofilms and mineralized living composites, measured using micro-indentation technique with a spherical probe (diameter = ~ 93 microns). The experiments were performed in an aqueous environment. Results are presented as Mean ± s.d. P = 0.00000027. *P < 0.05, **P < 0.01, ***P < 0.001. Statistics are derived using a two-sided t-test. Data are representative of n = 5 independent experiments. b, Representative indentation curves of the biofilms and living composites.

Extended Data Fig. 10 Local nanoscale mechanical properties of the dried gradient living composites measured by nano-indentation through atomic force microscopy (AFM).

a, Schematic model showing how AFM nano-indentation is used to measure the mechanical properties of composite microstructures. AFM nano-indentation is used to estimate Young’s modulus b, and stiffness c, of the different regions in the dried gradient composites. Results are presented as Mean ± s.d., at least 1200 counts were used for Young’s modulus, stiffness statistics.

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Wang, Y., An, B., Xue, B. et al. Living materials fabricated via gradient mineralization of light-inducible biofilms. Nat Chem Biol 17, 351–359 (2021).

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