生物工程

Bacterial cellulose (BC) is a renewable biopolymer valued for its exceptional purity, biocompatibility, and mechanical strength, with broad applications in biomedicine and sustainable materials. However, achieving reproducible BC production and downstream processing remains a major challenge. Inoculum preparation is particularly difficult to standardize because cellulose-producing strains form pellicles that sequester cells, making optical density measurements unreliable. In addition, recovery and drying procedures can alter fiber accessibility, and enzymatic hydrolysis conditions are often inconsistently defined and lack proper enzyme activity assessment. These issues contribute to substantial variability in BC-derived nanoparticle yields. This protocol describes the production of BC from Komagataeibacter xylinus DSMZ 6513, including culture medium preparation, inoculum generation, and scaling up under static cultivation conditions. It further details BC pellicle purification using NaOH, followed by pulping, freeze-drying, and milling to ensure material stability during storage and use. BC hydrolysis is performed with commercially available cellulase from Trichoderma reesei, with enzyme activity quantified prior to each reaction to ensure reproducibility. This standardized approach enables the reproducible production of bacterial cellulose nanoparticles (BCNPs). The protocol also includes minimal morphological characterization methods. By standardizing culture, recovery, and hydrolysis steps, the workflow reduces experimental variability and improves comparability across laboratories. Overall, it provides an accessible and reproducible method for generating BC and BCNPs of consistent quality without the need for specialized instrumentation.

Genetically encodable reporters that produce signals detectable in deep tissues offer a powerful tool for noninvasive monitoring of molecular events in vivo. Although magnetic resonance imaging (MRI) is a standard technique for noninvasive clinical imaging, its wider application in detecting molecular activities has been constrained by the lack of programmable sensors. This limitation is in stark contrast to the widespread use of fluorescent reporter–derived sensors in cultured cells and in transparent specimens. To overcome this limitation, we recently developed the modular aquaporin-based protease-activatable probe for enhanced reporting (MAPPER) platform. This sensor engineering framework integrates a metal-free MRI reporter derived from human aquaporin-1 (hAqp1) with synthetic protease-based circuits. This integration facilitates the modular and scalable creation of a wide range of sensors by regulating protease activity through precise molecular events, such as protein–protein interactions, pharmacological inhibition, and second messenger signaling. In this paper, we present a detailed protocol for constructing and deploying sensors using the MAPPER paradigm. The protocol encompasses genetic design, lentiviral production, stable cell line generation, biochemical and microscopic validation of sensor function, diffusion-weighted MRI, and MR image analysis to quantify sensor signals in terms of the apparent diffusion coefficient. We describe two distinct MAPPER architectures: DD-MAPPER, which leverages protease-controlled protein degradation, and ER-MAPPER, which utilizes protease-controlled, subcellular trafficking. The MAPPER framework allows adaptation to various molecular targets without the need to redesign the core MRI reporter mechanism, making MAPPER a versatile platform for noninvasive biosensing in living cells and tissues.

Skin models play critical roles in understanding disease mechanisms and advancing therapeutic development. However, conventional systems based on 2D cell cultures, in vivo animal models, and ex vivo tissue explants are limited by insufficient physiological complexity, interspecies differences, and restricted accessibility, respectively. Advances in biofabrication technologies have enabled the engineering of 3D skin equivalents that better balance biological complexity and experimental scalability. Here, we present a biofabrication protocol inspired by the regenerative processes of wound healing to construct vascularized 3D organotypic skin models in a stepwise manner. The approach integrates bioprinting for precise spatial organization of cellular compartments with guided cell self-organization to achieve native-like tissue complexity and heterogeneity. Through a programmable culture strategy, tissue maturation proceeds sequentially through keratinocyte proliferation and collective migration, microchannel endothelialization, basal-to-suprabasal differentiation, and progressive extracellular matrix remodeling within a fibrin-based scaffold. The resulting tissue constructs comprise stratified epidermal layers positioned atop a vascularized, fibroblast-remodeled dermal matrix. Beyond reproducing key structural features of human skin, this protocol recapitulates cellular processes associated with tissue regeneration, providing a dynamic platform for investigating disease pathogenesis, progression, and therapeutic responses.

Cell therapy holds great promise for cancer immunotherapy, but its clinical efficacy is severely hindered by poor post-transplant cell survival, low homing efficiency, and host immune clearance. To address these challenges, this study develops a novel light-controlled immunotherapy strategy that integrates a red/far-red light genetic switch with single-cell encapsulation engineering. The red/far-red light (660/730 nm) reversible regulatory system enables precise spatiotemporal control over the expression of therapeutic proteins in engineered cells (e.g., CAR-T or engineered HEK 293T cells), allowing on-demand activation of anti-tumor immune responses. On this basis, a mild enzyme-mediated single-cell encapsulation technique is further employed to rapidly form a protective hydrogel coating in situ on the cell surface, thereby enhancing the survival of transplanted cells under hostile in vivo microenvironments. This strategy combines precise gene expression regulation with physical protection, improving therapeutic outcomes without the need for genomic modification of the cells. It provides a new paradigm for developing safe, controllable, and efficient cancer immunotherapy.

Satellite cells are adult skeletal muscle stem cells that play essential roles in muscle regeneration. Understanding their behavior is critical for elucidating the mechanisms of muscle repair and advancing muscle regenerative therapies. This requires efficient methods for genetic manipulation in these cells. Retroviral-mediated gene delivery is commonly used for stable transgene expression in immortalized cell lines. However, existing approaches are not optimized for primary satellite cells, often resulting in variable efficiency and inconsistent outcomes. Here, we describe an optimized protocol for satellite cell isolation and culture, as well as retroviral production and infection of primary satellite cells that achieves high transduction efficiency. The satellite cell isolation procedure enriches for myofiber fragments prior to satellite cell release, thereby reducing contamination by non-myogenic cells and improving cell purity. Another key feature of this protocol is the concentration of retroviral particles and their resuspension in satellite cell growth medium prior to infection, which minimizes satellite cell exposure to packaging cell-conditioned medium. Compared to standard approaches, this protocol improves both infection efficiency and reproducibility. It is readily adaptable to a wide range of downstream applications, including microscopies, biochemical assays, and molecular biology analyses.

Mitochondrial transplantation is an emerging strategy for cellular repair, yet its efficiency is often limited by poor targeting and environmental instability. This protocol details the fabrication and comprehensive characterization of neutrophil membrane-fused mitochondria (nMITO), a hybrid organelle platform designed to combine the metabolic vigor of natural mitochondria with the targeting and anti-inflammatory properties of neutrophil membranes. We describe an optimized workflow for mouse heart mitochondrial isolation, lipopolysaccharide (LPS)-activated neutrophil membrane (NEM) extraction, and the subsequent sonication-mediated fusion process. Characterization techniques include dynamic light scattering (DLS) for size and zeta potential, transmission electron microscopy (TEM) for ultrastructural integrity, and bioenergetic assays [ATP synthesis and tetramethylrhodamine methyl ester (TMRM)-based membrane potential] to ensure functional preservation.

Real-time measurement of blood flow and nanocarrier transport in the cerebral microvasculature is crucial for understanding neurovascular physiology and nanocarrier-based drug delivery. Existing techniques lack the ability to measure blood flow rates in individual vessels with high spatial and temporal resolution in real time. Two-photon fluorescence correlation spectroscopy (2P-FCS) provides a powerful approach for monitoring tracer molecules within a small confocal observation volume. This enables the simultaneous determination of particle number and flow dynamics in vivo. Here, we present a detailed protocol for in vivo 2P-FCS measurements in the mouse cerebral microvasculature. The protocol includes preparation of the cranial window, delivery of fluorescent dextran tracers for vascular visualization, and FCS measurements. It also includes two-photon imaging of the cerebrovascular network and acquisition and analysis of fluorescence correlation data. The protocol describes calibration of the confocal volume diameter and optimization of two-photon excitation parameters. This workflow enables real-time measurement of tracer concentration and flow velocity in individual cerebral microvessels with high spatial and temporal resolution. The method can be adapted to study blood flow dynamics, nanoparticle transport, and microvascular physiology in a variety of in vivo imaging systems equipped with multiphoton microscopy and FCS capabilities.

Understanding epithelial barrier function is essential for studying both its normal physiology and its role in disease, yet choosing an appropriate experimental model remains challenging. Animal models are commonly used but often suffer from interspecies differences that limit translational relevance. Human-derived cell lines offer a more suitable alternative, although establishing them often requires immortalisation strategies that involve overexpression of oncogenes, which can introduce phenotypic and functional changes. In contrast, primary cells, such as human small airway epithelial cells (HSAECs), provide a more physiologically accurate model. A critical aspect of replicating the native respiratory environment is maintaining continuous air exposure, which can be achieved through air–liquid interface (ALI) culture. This protocol provides a unified, step-by-step workflow for cultivating primary HSAECs under ALI conditions, covering the entire process from initial recovery after cryopreservation to the formation of a barrier-like layer. The protocol incorporates non-invasive methods such as transepithelial electrical resistance (TEER) measurements to monitor its integrity. While individual elements of this workflow have been described separately in different studies, a consolidated version encompassing the full workflow has not been widely available. This resource is intended for researchers with limited experience in airway epithelial culture and offers practical, clear guidance through each step of the process.

3-nitro-tyrosine (nitroTyr) is one of numerous oxidative protein modifications implicated in diseases such as cardiovascular disease, cancer, and amyotrophic lateral sclerosis (ALS). Because of this, the ability to site-specifically encode nitroTyr into recombinant proteins is a powerful approach for studying these disease pathways. However, producing proteins with defined nitration sites is technically challenging due to the limitations of traditional chemical nitration via peroxynitrite, which lacks residue and site-specificity. Genetic code expansion (GCE) offers a solution by enabling precise incorporation of nitroTyr at designated TAG codons using engineered aminoacyl-tRNA synthetase/tRNA pairs from Methanocaldococcus jannaschii and Methanomethylophilus alvus. This protocol provides a reliable, optimized workflow for incorporating nitroTyr into proteins in E. coli using GCE. It guides users through key considerations in selecting cell lines, media conditions, and GCE systems to minimize off-target effects such as release factor 1 competition, near-cognate suppression, and chemical reduction of nitroTyr. The method is demonstrated using wild-type and TAG-containing superfolder GFP but is broadly applicable to other proteins of interest.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.