Biological Engineering

Human neuromuscular diseases represent a diverse group of disorders with unmet clinical need, ranging from muscular dystrophies, such as Duchenne muscular dystrophy (DMD), to neurodegenerative disorders, such as amyotrophic lateral sclerosis (ALS). In many of these conditions, axonal and neuromuscular synapse dysfunction have been implicated as crucial pathological events, highlighting the need for in vitro disease models that accurately recapitulate these aspects of human neuromuscular physiology. The protocol reported here describes the co-culture of neural spheroids composed of human pluripotent stem cell (PSC)–derived motor neurons and astrocytes, and human PSC-derived myofibers in 3D compartmentalised microdevices to generate functional human neuromuscular circuits in vitro. In this microphysiological model, motor axons project from a central nervous system (CNS)–like compartment along microchannels to innervate skeletal myofibers plated in a separate muscle compartment. This mimics the spatial organization of neuromuscular circuits in vivo. Optogenetics, particle image velocimetry (PIV) analysis, and immunocytochemistry are used to control, record, and quantify functional neuromuscular transmission, axonal outgrowth, and neuromuscular synapse number and morphology. This approach has been applied to study disease-specific phenotypes for DMD and ALS by incorporating patient-derived and CRISPR-corrected human PSC-derived motor neurons and skeletal myogenic progenitors into the model, as well as testing candidate drugs for rescuing pathological phenotypes. The main advantages of this approach are: i) its simple design; ii) the in vivo–like anatomical separation between CNS and peripheral muscle; and iii) the amenability of the approach to high power imaging. This opens up the possibility for carrying out live axonal transport and synaptic imaging assays in future studies, in addition to the applications reported in this study.

Graphical abstract

Graphical abstract abbreviations: Channelrhodopsin-2 (CHR2+), pluripotent stem cell (PSC), motor neurons (MNs), myofibers (MFs), neuromuscular junction (NMJ).

In this study, we introduce a detailed protocol for the preparation of DNA-assembled GRS-DNA-copper sulfide (CuS) nanodandelion, a multifunctional theranostics nanoparticle. Using transmission electron microscope (TEM) and dynamic light scattering techniques, we characterize the physicochemical property of DNA-assembled GRS-DNA-CuS nanodandelions and their dissociation property after the first near-infrared (NIR) light irradiation. In addition, we systematically monitor the processes of tumor accumulation and uniform intratumoral distribution (UITD) of ultrasmall CuS photothermal agents (PAs), which are dissociated from GRS-DNA-CuS nanodandelions, by Raman imaging and photoacoustic imaging, respectively. The UITD of the dissociated ultrasmall CuS PAs can enhance the therapeutic efficiency of photothermal treatment under the second NIR light irradiation. Overall, this protocol provides a powerful tool to achieve UITD of PAs by explosively breaking the hydrogen bonds of DNA in GRS-DNA-CuS nanodandelions under NIR light irradiation. We expect DNA-assembled nanotheranostics to serve as a robust platform for a variety of biomedical applications related to photothermal therapy in the oncology field. This protocol can increase experimental reproducibility and contribute to efficient theranostics nanomedicine.

Traditional drug safety assessments often fail to predict complications in humans, especially when the drug targets the immune system. Rodent-based preclinical animal models are often ill-suited for predicting immunotherapy-mediated adverse events in humans, in part because of the fundamental differences in immunological responses between species and the human relevant expression profile of the target antigen, if it is expected to be present in normal, healthy tissue. While human-relevant cell-based models of tissues and organs promise to bridge this gap, conventional in vitro two-dimensional models fail to provide the complexity required to model the biological mechanisms of immunotherapeutic effects. Also, like animal models, they fail to recapitulate physiologically relevant levels and patterns of organ-specific proteins, crucial for capturing pharmacology and safety liabilities. Organ-on-Chip models aim to overcome these limitations by combining micro-engineering with cultured primary human cells to recreate the complex multifactorial microenvironment and functions of native tissues and organs. In this protocol, we show the unprecedented capability of two human Organs-on-Chip models to evaluate the safety profile of T cell–bispecific antibodies (TCBs) targeting tumor antigens. These novel tools broaden the research options available for a mechanistic understanding of engineered therapeutic antibodies and for assessing safety in tissues susceptible to adverse events.

Graphical abstract

Figure 1. Graphical representation of the major steps in target-dependent T cell–bispecific antibodies engagement and immunomodulation, as performed in the Colon Intestine-Chip

The study and use of decellularized extracellular matrix (dECM) in tissue engineering, regenerative medicine, and pathophysiology have become more prevalent in recent years. To obtain dECM, numerous decellularization procedures have been developed for the entire organ or tissue blocks, employing either perfusion of decellularizing agents through the tissue’s vessels or submersion of large sections in decellularizing solutions. However, none of these protocols are suitable for thin tissue slices (less than 100 µm) or allow side-by-side analysis of native and dECM consecutive tissue slices. Here, we present a detailed protocol to decellularize tissue sections while maintaining the sample attached to a glass slide. This protocol consists of consecutive washes and incubations of simple decellularizing agents: ultrapure water, sodium deoxycholate (SD) 2%, and deoxyribonuclease I solution 0.3 mg/mL (DNase I). This novel method has been optimized for a faster decellularization time (2–3 h) and a better correlation between dECM properties and native tissue-specific biomarkers, and has been tested in different types of tissues and species, obtaining similar results. Furthermore, this method can be used for scarce and valuable samples such as clinical biopsies.

This protocol describes the recombinant expression of proteins in E. coli containing phosphoserine (pSer) installed at positions guided by TAG codons. The E. coli strains that can be used here are engineered with a ∆serB genomic knockout to produce pSer internally at high levels, so no exogenously added pSer is required, and the addition of pSer to the media will not affect expression yields. For “truncation-free” expression and improved yields with high flexibility of construct design, it is preferred to use the Release Factor-1 (RF1) deficient strain B95(DE3) ∆A ∆fabR ∆serB, though use of the standard RF1-containing BL21(DE3) ∆serB is also described. Both of these strains are serine auxotrophs and will not grow in standard minimal media. This protocol uses rich auto-induction media for streamlined and maximal production of homogeneously modified protein, yielding ~100–200 mg of single pSer-containing sfGFP per liter of culture. Using this genetic code expansion (GCE) approach, in which pSer is installed into proteins during translation, allows researchers to produce milligram quantities of specific phospho-proteins without requiring kinases, which can be purified for downstream in vitro studies relating to phosphorylation-dependent signaling systems, protein regulation by phosphorylation, and protein–protein interactions.

Graphical abstract:

Bispecific antibodies (BsAbs) are typically monoclonal antibody (mAb)–derived molecular entities engineered to bind to two distinct targets, including two antigens or two epitopes on the same antigen. When compared to parental monoclonal antibodies or combinational therapies, the generated BsAbs have the ability to bridge the two targets and thus may offer additional clinical benefits. Characterizing BsAbs’ ability to bind to both targets simultaneously is critical for their biotherapeutic development. A range of bi-functional quantitative bridging assays to enable target-specific capture and detection of binding properties include enzyme-linked immunosorbent assay (ELISA), surface plasmon resonance (SPR), and cell-based flow cytometry. Developing suitable and robust cell-based bioassays is more challenging than non-cell-based binding assays because cell-based assays with complex matrices can be inherently variable and often lack precision. Compared to SPR, ELISA has a rapid setup and readily available method, being widely and extensively applied in almost every laboratory. Here, we describe a dual-target bridging ELISA assay that characterizes the ability of a HER2(human epidermal growth factor receptor 2)/PD-L1(programmed cell death ligand 1) BsAb in binding to both HER2 and PD-L1 simultaneously, a prerequisite for its envisioned mode of action.

Graphical abstract:

RNA binding proteins (RBPs) are critical regulators of cellular phenotypes, and dysregulated RBP expression is implicated in various diseases including cancer. A single RBP can bind to and regulate the expression of many RNA molecules via a variety of mechanisms, including translational suppression, prevention of RNA degradation, and alteration in subcellular localization. To elucidate the role of a specific RBP within a given cellular context, it is essential to first identify the group of RNA molecules to which it binds. This has traditionally been achieved using cross-linking-based assays in which cells are first exposed to agents that cross-link RBPs to nucleic acids and then lysed to extract and purify the RBP-nucleic acid complexes. The nucleic acids within the mixture are then released and analyzed via conventional means (e.g., microarray analysis, qRT-PCR, RNA sequencing, or Northern blot). While cross-linking-based ribonucleoprotein immunoprecipitation (RIP) has proven its utility within some contexts, it is technically challenging, inefficient, and suboptimal given the amount of time and resources (e.g., cells and antibodies) required. Additionally, these types of studies often require the use of over-expressed versions of proteins, which can introduce artifacts. Here, we describe a streamlined version of RIP that utilizes exclusion-based purification technologies. This approach requires significantly less starting material and resources compared to traditional RIP approaches, takes less time, which is tantamount given the labile nature of RNA, and can be used with endogenously expressed proteins. The method described here can be used to study RNA-protein interactions in a variety of cellular contexts.

Graphical abstract:

Cell bioprinting technologies aim to fabricate tissue-like constructs by delivering biomaterials layer-by-layer. Bioprinted constructs can reduce the use of animals in drug development and hold promise for addressing the shortage of organs for transplants. We recently introduced a laser-assisted drop-on-demand bioprinting technology termed Laser Induced Side Transfer (LIST). This technology can print delicate cell types, including primary neurons. This bioprinting protocol includes the following key steps: cell harvesting, bio-ink preparation, laser setup priming, printing, and post-printing analysis. This protocol includes a detailed description of the laser setup, which is a rather unusual setup for a biology lab. This should allow easy reproduction by readers with basic knowledge of optics. Although we have focused on neuron bioprinting, interested readers will be able to adapt the protocol to bioprint virtually any cell type.

Graphical abstract:

Type 1 regulatory T (Tr1) cells are an immunoregulatory CD4⁺ Foxp3^- IL-10^high T cell subset with therapeutic potential for various inflammatory diseases. Retroviral (RV) transduction has been a valuable tool in defining the signaling pathways and transcription factors that regulate Tr1 differentiation and suppressive function. This protocol describes a method for RV transduction of naïve CD4⁺ T cells differentiating under Tr1 conditions, without the use of reagents such as polybrene or RetroNectin. A major advantage of this protocol over others is that it allows for the role of genes of interest on both differentiation and function of Tr1 cells to be interrogated. This is due to the high efficiency of RV transduction combined with the use of an IL10^GFP/Foxp3^RFP dual reporter mouse model, which enables successfully transduced Tr1 cells to be identified and sorted for functional assays. In addition, this protocol may be utilized for dual/multiple transduction approaches and transduction of other lymphocyte populations, such as CD8⁺ T cells.

Currently, there are several in vitro protocols that focus on directing human induced pluripotent stem cell (hiPSC) differentiation into either the cardiac or pulmonary lineage. However, these systemsprotocols are unable to recapitulate the critical exchange of signals and cells between the heart and lungs during early development. To address this gap, here we describe a protocol to co-differentiate cardiac and pulmonary progenitors within a single hiPSC culture by temporal specific modulation of Wnt and Nodal signaling. Subsequently, human cardio-pulmonary micro-tissues (μTs) can be generated by culturing the co-induced cardiac and pulmonary progenitors in 3D suspension culture. Anticipated results include expedited alveolarization in the presence of cardiac cells, and segregation of the cardiac and pulmonary μTs in the absence of exogenous Wnt signaling. This protocol can be used to model cardiac and pulmonary co-development, with potential applications in drug testing, and as a platform for expediting the maturation of pulmonary cells for lung tissue engineering.