Biological Engineering

In recent years, the calcifying properties of some cyanobacteria have been used in the production of living building materials (LBMs), such as bio-concrete, as a CO₂-friendly alternative for cement. This microbially induced calcium carbonate precipitation (MICP) technique can act as a novel platform technology for carbon capture strategies. Consequently, various research articles have been conducted based on a diverse set of workflows, including several modifications, to manufacture LBMs. However, such articles contain only fragmentary descriptions of the materials and methods used. This protocol is meant to act as a detailed, step-by-step operational manual for the production of LBMs using the cyanobacterial model strain Picosynechococcus sp. PCC 7002. The process is divided into several steps, such as the activation of the cyanobacterial-gel solution with CaCl₂ × 2H₂O and NaHCO₃, casting standardized prisms (160 × 40 × 40 mm), and demolding LBMs. Subsequently, bending tensile and compressive strength tests are performed according to the procedures commonly used in concrete and material testing as proof of concept.

Intravenous hemostats have shown significant promise in prolonging survival for severe noncompressible and internal injuries in preclinical animal models. Existing approaches include the use of liposomes with or without procoagulant enzymes, as well as polymer nanoparticles or soluble biopolymers. While these methods predominantly target or mimic tissue components that are present during coagulation, such as activated platelets and collagen, they may not account for the loss of fibrinogen, which is not only key to clot formation but also the first protein to fall below critical levels in dilutional coagulopathy. This protocol describes the synthesis and in vitro or ex vivo characterization of a crosslinkable nanoparticle system that seeks to address dilutional coagulopathy by leveraging the critical gelation concentration and bioorthogonal click chemistry. The system was shown to only gel at high nanoparticle and crosslinker concentrations, increase the rate of platelet recruitment, and decrease the rate of clot degradation in a low-fibrinogen environment, providing a platform for treating severe hemorrhage in a coagulopathic environment. Ultimately, the contents of this protocol may assist researchers in the in vitro characterization and screening of other crosslinkable nanoparticle systems or hemostats, with potential expansions to other categories of coagulation dysfunction, such as embolism treatment.

Zebrafish offer numerous advantages as a vertebrate model because of their rapid development, high fecundity, transparent embryos, and ease of genetic manipulation. A wide variety of transgenic and mutant fish lines have been generated, and efficiently sharing these resources is crucial for advancing research. Zebrafish lines have typically been exchanged as early embryos, adult fish, or cryopreserved sperm, making transportation costly and logistically challenging. Here, we provide a protocol for preserving functional zebrafish sperm for more than 7 days at room temperature and subsequent in vitro fertilization using the preserved sperm. In this protocol, sperm collected either from the cloaca of an anesthetized male or from dissected testes is stored in L-15-based storage medium. Importantly, the storage medium, originally developed for zebrafish, is also applicable to medaka, another widely used vertebrate model. This sperm storage method allows researchers to ship sperm using low-cost methods and to investigate key factors for motility and fertilizing ability in those sperm.

Bottom-up tissue engineering using cell spheroids offers many advantages in recapitulating native cell–cell and cell–matrix interactions. Many tissues, such as cartilage, bone, cardiac muscle, intestine, and neural tissues, have been tissue-engineered using cell spheroids. However, previous methods for spheroid assembling, such as mold casting, hydrogel-based bioprinting, or needle array, either lack control over final tissue geometry or face challenges in scalability and throughput. In this protocol, we describe a robust and scalable tissue engineering method for assembling cell spheroids into a thin, planar spheroid sheet. The spheroids are sandwiched between two flexible meshes held by a frame, facilitating uniform spheroid fusion while ensuring nutrient exchange and ease of handling. We demonstrate this method by producing thin cartilage tissue from human mesenchymal stem cells undergoing chondrogenic differentiation. This approach offers a practical platform for producing thin membrane-like tissue constructs for many research and therapeutic applications.

Oxygen tension is a key regulator of early human neurogenesis; however, quantifying intra-tissue O₂ in 3D models for an extended period remains difficult. Existing approaches, such as needle-type fiber microsensors and intensity-based oxygen probes or time-domain lifetime imaging, either perturb the organoids or require high excitation doses that limit the measurement period. Here, we present a step-by-step protocol to measure intra-organoid oxygen in human cerebral organoids (hCOs) using embedded ruthenium-based CPOx microbeads and widefield frequency-domain fluorescence lifetime imaging microscopy (FD-FLIM). The workflow covers dorsal/ventral cerebral organoid patterning, organoid fusion at day 12 with co-embedded CPOx beads, standardized FD-FLIM acquisition (470-nm external modulation, 16 phases at 50 kHz, dual-tap camera), automated bead detection and lifetime extraction in MATLAB, and session-matched Stern–Volmer calibration with Ru(dpp)₃(ClO₄)₂ to convert lifetimes to oxygen concentration. The protocol outputs per-bead oxygen maps and longitudinal patterns stratified by bead location (intra-organoid vs. gel) and sample state (healthy vs. abnormal), enabling direct linkage between developmental growth and oxygen dynamics.

Extracellular vesicles (EVs) have emerged as promising carriers for the targeted delivery of therapeutic proteins to specific cells. Previously, we demonstrated that genetically engineered EVs can be used for targeted protein delivery. This protocol details the generation of mannose receptor (CD206)-targeted EVs using a modular plasmid system optimized for production in HEK293T cells. Three plasmids enable customizable EV budding, cargo loading, and surface modification for targeting to antigen-presenting cells (APCs). EVs are isolated via differential centrifugation and chromatography, characterized using transmission electron microscopy (TEM) and nanoparticle tracking analysis (NTA), and validated through functional uptake assays in primary human activated dendritic cells. Our approach combines flexibility in engineering required EVs with robust, reproducible isolation and characterization workflows. Its modularity allows easy adaptation to alternative targets or cargoes, which can be validated immediately through in vitro testing.

Diabetes lacks concrete curative strategies due to diverse aetiologies and, therefore, represents the perfect candidate for cell replacement therapy, since it is caused by either an absolute (type 1 diabetes) or relative (type 2 diabetes) defect in the insulin-producing beta cells of the pancreas. Pancreatic alpha cells are a promising source for transdifferentiation into insulin-producing cells as they share a common developmental origin with beta cells and exhibit a certain degree of cellular plasticity. Furthermore, impairment of glucagon signaling in diabetes leads to a marked increase in alpha cell mass, raising the possibility that such alpha cell hyperplasia provides an increased supply of alpha cells for their transdifferentiation into new beta cells.

In this protocol, we used the modular epigenetic CRISPR/dCas9 toolbox for targeted DNA methylation (EpiCRISPR) and silencing of the Arx gene (Aristaless Related Homeobox, Arx), which is essential for the maintenance of alpha cell identity. Methylation-based silencing of Arx initiates the reprogramming of pancreatic alpha cells into insulin-producing cells. As a key novelty, this protocol provides a direct route for epigenetically induced transdifferentiation of mouse pancreatic alpha TC1-6 cells into insulin-producing cells and thereby confirms a proof of concept of reversible cellular epigenetic reprogramming in vitro. In addition, this streamlined workflow addresses the inherent challenges of transfecting clustered alpha TC1-6 cells by optimizing their dissociation into single-cell suspensions, thereby improving uptake and reproducibility.

In summary, this approach for cell transdifferentiation involves precise epigenetic editing of a lineage-specific marker gene, thereby enabling direct lineage conversion in a safe and versatile strategy to generate insulin-producing cells by epigenetic reprogramming. In contrast to approaches that rely on viral vectors or permanent genome editing, this method reduces the risk of off-target effects and immunogenic responses while ensuring reproducibility. The combination of efficiency and precision makes it a valuable tool to advance regenerative approaches for diabetes therapy and to explore the epigenetic regulation of cell identity.

Micro milling is a subtractive manufacturing method for fabricating micro-scale three-dimensional features from hard substrates like acrylic, wood, or metal. It enables rapid prototyping of biomicrofluidic devices and master molds, offering advantages over traditional fabrication methods like photolithography. Micro milling is seldom applied in the fabrication of organs-on-a-chip, in part due to its requirement for knowledge of computer numerical machining techniques that are required to program and operate micro mills. This protocol provides practical guidelines for micro milling–based fabrication of organs-on-a-chip, including toolpath optimization, SolidWorks and Fusion workflows, and troubleshooting tips. A case study demonstrates the design and fabrication of master molds for a human airway-on-a-chip, validated in a recent publication. This resource supports the expansion of micro milling techniques into organs-on-a-chip, which will enhance capacity for rapid device prototyping and design of more complex 3D features that better recapitulate human physiology.

Artificial metalloenzymes (ArMs) hold great promise for expanding the toolbox of non-natural transformations usable in living systems, such as cells, plants, and animals. However, their practical application remains challenging, primarily due to their unsatisfactory stability and inefficient intracellular assembly. We recently reported a new strategy, called artificial metalloenzymes in artificial sanctuaries (ArMAS) through liquid–liquid phase separation (LLPS), to enhance the performance of ArMs in cells by placing them in more friendly artificial microenvironments. Here, this protocol describes the detailed method for using this ArMAS–LLPS strategy, a robust way to create artificial compartments using an ArM protein scaffold through LLPS and construct ArMs within using self-labeling cofactor anchoring reactions. In detail, in Escherichia coli, membraneless protein condensates are formed by expressing a self-labeling fusion protein, HaloTag-SNAPTag (HS) and act as intracellular sanctuaries. Simultaneously, the HS scaffolds enable site-specific, bioorthogonal conjugation with synthetic metal cofactors, facilitating efficient ArM formation within the LLPS domains. This strategy can significantly enhance the intracellular catalytic activity and stability of the named HS-based ArMs, allowing whole-cell catalysis to be performed to enable abiotic transformations both in vitro and in vivo. The protocol provides a proof-of-concept approach for researchers aiming to develop stable ArM-based whole-cell catalytic systems for synthetic biology and therapeutic applications.

Lipid nanoparticles (LNPs) are powerful carriers for nucleic acid delivery, but plasmid DNA-loaded LNPs (pDNA-LNPs) have been limited by inflammation and toxicity. We showed that standard pDNA-LNPs activate the cGAS–STING pathway, leading to severe immune responses and mortality in mice. To overcome this, we co-loaded nitro-oleic acid (NOA), an endogenous STING inhibitor, into pDNA-LNPs. NOA-pDNA-LNPs mitigated inflammation, enabled safe in vivo delivery, and supported sustained gene expression for months. Here, we present a detailed protocol for producing and characterizing NOA-pDNA-LNPs to facilitate safer, long-term gene delivery applications.