细胞生物学

Single-cell and single-nucleus RNA sequencing are revolutionizing our understanding of cellular biology. The identification of molecular markers, single-cell transcriptomic profiling, and differential gene expression at the cellular level has revealed key functional differences between cells within the same tissue. However, tissue dissociation remains challenging for non-model organisms and for tissues with unique biochemical properties. For example, the mosquito fat body, which serves functions analogous to mammalian adipose and liver tissues, consists of trophocytes—large, adipocyte-like cells whose cytoplasm is filled with lipid droplets. Conventional enzymatic dissociation methods are often too harsh for these fragile cells, and their high lipid content can interfere with reagents required for single-cell transcriptomic analysis. Single-nucleus RNA sequencing (snRNA-seq) offers an alternative strategy when intact cells with high-quality RNA cannot be obtained by enzymatic or mechanical dissociation. Here, we present an optimized reproducible methodology for nuclei isolation from the fat body of Anopheles gambiae mosquitoes, enabling high-quality snRNA-seq. Our approach involves tissue fixation and lipid removal, followed by cell lysis and nuclei purification using a sucrose cushion. We validated this protocol on both sugar-fed and blood-fed samples, established quality metrics to remove potential ambient RNA contamination, and demonstrated that snRNA-seq using this method yields high-quality sequencing results.

Most viruses extensively remodel their host cells to establish productive infection. Visualization of virus-induced cellular remodeling by electron microscopy (EM) has been revolutionized in recent years by advances in cryo-focused ion beam (cryo-FIB) milling paired with cryo-electron tomography (cryo-ET). As cryo-FIB/ET becomes more widely available, there is a need for beginner-friendly guides to optimize the preparation of virus-infected mammalian cells on EM grids. Here, we provide an in-house protocol for new users for preparing samples of cells infected with herpes simplex virus 1 (HSV-1) for cryo-FIB/ET. This protocol guides users in how to seed infected cells onto grids, blot, and plunge-freeze grids using basic, manual equipment. It also provides tips on how to screen and prioritize grids for efficient milling and data collection.

ER-phagy, a selective autophagy process crucial for maintaining cellular homeostasis by targeting the endoplasmic reticulum (ER), has been challenging to study in vivo due to the lack of suitable spatiotemporal quantification tools. Existing methods like electron microscopy, biochemical assays, and in vitro reporters lack resolution, scalability, or physiological relevance. Here, we present a detailed protocol for generating two transgenic mouse models: ER-TRG (constitutively expressing an ER lumen-targeting tandem RFP-GFP tag) and CA-ER-TRG (Cre-recombinase-activated ER-TRG). Additionally, we outline procedures for quantitative imaging of ER-phagy in vivo, covering tissue preparation, confocal microscopy, and signal analysis. This protocol offers a robust and reproducible tool for investigating ER-phagy dynamics across various tissues, developmental stages, and pathophysiological conditions, facilitating both fundamental and translational research.

Small GTPases function as molecular switches in cells, and their activation triggers diverse cellular responses depending on the GTPase type. Therefore, visualizing small GTPase activation in living cells is crucial because their activity is tightly regulated in space and time, and this spatiotemporal pattern of activation often determines their specific cellular functions. Various biosensors, such as relocation-based sensors and fluorescence resonance energy transfer (FRET)-based sensors, have been developed. However, these methods rely on interactions between activated GTPases and their downstream effectors, which limits their applicability for detecting activation of GTPases with unknown or atypical effectors. Recently, we developed a novel method utilizing split fluorescence technology to detect membrane recruitment of small GTPases upon activation, designated the Small GTPase ActIvitY ANalyzing (SAIYAN) system. This approach offers a new strategy for monitoring small GTPase activation based on membrane association and is potentially applicable to a wide range of small GTPases, including those with uncharacterized effectors.

During herpesvirus replication, capsids are assembled inside the nucleus and translocated into the cytosol by a non-canonical nucleocytoplasmic export process termed nuclear egress. Traditional methods of measuring nuclear egress rely on imaging-based technologies such as confocal and electron microscopy. These techniques are labor-intensive, limited by the number of cells that can be examined, and may not accurately represent the entire population, generating a potential bias during data interpretation. To overcome these problems, we have developed a flow cytometry–based method to measure HSV-1 nuclear egress that we termed FLARE (FLow cytometry–based Assay of nucleaR Egress). This assay uses a double fluorescent reporter system, utilizing HSV-1-tdTomato to identify infected cells and an Alexa Fluor-488-conjugated, capsid-specific antibody to detect cytosolic capsids, thereby distinguishing infected cells with nuclear egress from those without it. This assay provides more quantitative results than traditional methods, enables large-scale high throughput, and can be adapted for use with other herpesviruses.

Conventional Schlieren optics equipment typically operates on a large optical table, which is inconvenient for imaging small samples or thin layers of transparent materials. We describe an imaging device based on Schlieren optics, aided by a slight shift in light reflected from two surfaces. The device is designed to place the sample between a thick concave mirror and a camera next to a point-light source located at the spherical origin of the concave mirror. The compact device is portable and convenient. It is similarly capable of sensitively detecting patterns in gaseous or liquid media created by a density gradient when the optical effect is too subtle to be detectable by regular cameras and scanners. The new device is particularly suitable for detecting translucent samples, including thin fluid films on the order of micrometers, tissue slices, and other biological samples. We show two examples of how our device can be applied to imaging biological samples. The first compares images acquired using several techniques of a bacterial swarm spread over an agar plate; the second is a set of images of human cells grown on a tissue culture plate.

The tissue explant culture (histoculture) is a method that involves maintaining small pieces taken from an organ ex vivo or post mortem in a controlled laboratory setting. Such a technique has a number of advantages: unlike the 2D, organoid, or on-chip cultures, tissue explants preserve the whole complexity of the original tissue in vivo, its structure, extracellular matrix, and the diverse cell populations, including resident immune cells. The explant culture method can be applied to human tissue specimens obtained from biopsies or autopsies, provided that proper ethical protocols are followed. This avoids the difficulties that may arise in translating results obtained on animal models into biomedical research for humans. This advantage makes histocultures especially desirable for studying human pathogenesis in the course of infectious diseases. The disadvantage of the method is the limited lifespan of the cultured tissues; however, a number of approaches allow extending tissue viability to a period sufficient for observing the infection onset and development. Here, we provide a protocol for lung explant maintenance that allows tracing the local effects of infection with SARS-CoV-2 in humans. Further applications of the lung tissues cultured according to this protocol include, but are not limited to, histochemical and immunohistochemical studies and microscopy, FACS, qPCR, and ELISA-based analysis of the conditioned culture media.

The pancreatic islet, the only type of tissue that secretes insulin in response to elevated blood glucose, plays a vital role in diabetes development and treatment. While various islet vascularization strategies have been developed, they have been hindered by major limitations such as relying on pre-patterning and the inability to span long distances. Furthermore, few strategies have demonstrated robust enough vascularization in vivo to support therapeutic subcutaneous islet transplantation. Using adaptive endothelial cells (ECs) reprogrammed by transient expression of the ETS Variant Transcription Factor 2 (ETV-2) gene, we have physiologically vascularized human islets within a generic microchamber and have achieved functional engraftment of human islets in the subcutaneous space of mice. Such adaptive ECs, which we term reprogrammed vascular ECs (R-VECs), have been proven to be a suitable tool for both in vitro disease modeling and in vivo transplantation of not only islets but also other organoids.

Expansion microscopy (ExM) enables nanoscale imaging of biological structures using standard fluorescence microscopes. Accurate labeling of cytoskeletal filaments, such as microtubules, remains challenging due to structural distortion and labeling inaccuracy during sample preparation. This protocol describes an optimized method combining detergent extraction and NHS-ester labeling for high-precision visualization of microtubules in expanded samples. Cytoplasmic components and membranes are selectively removed, preserving the ultrastructure of the microtubule network. Microtubules are digested into peptides during expansion and subsequently labeled at their N-termini using NHS-ester dyes, eliminating the need for antibodies. Effective fluorophore displacement of ~1 nm or lower is achieved, depending on the applied expansion factor. The protocol is compatible with both in vitro and cellular samples and can be integrated into a wide range of ExM workflows. Labeled microtubules can serve as internal reference standards for correcting expansion factors in ExM datasets.

Primary cilia are evolutionarily conserved organelles that play critical roles in brain development. In the developing cortex, neural progenitors extend their primary cilia into the ventricular surface, where the cilia act as key signaling hubs. However, visualizing these cilia in a systematic and intact manner has been challenging. The commonly used cryostat sectioning only provides a limited snapshot of cilia on individual sections, and this process often disrupts the ciliary morphology. By contrast, the previously established whole-mount technique has been shown to preserve ciliary architecture in the adult mouse cortex. Here, we adapt and optimize the whole-mount approach for embryonic and neonatal brain, allowing robust visualization of ciliary morphology at the ventricular surface during development. This protocol describes step-by-step procedures for whole-mounting and immunostaining delicate embryonic and neonatal mouse cortices, enabling direct visualization of cilia in neural progenitors in the developing brain.