Bio-protocol最新文献

Flagellate stages of green microalgae such as Trebouxia are only partially characterised, with recent evidence suggesting that they are involved in both sexual and asexual reproduction. Conventional methods based on fixed samples in light, confocal, or electron microscopy provide only static observations and prevent real-time monitoring of living cells. To overcome this limitation, we have developed a simple and cost-effective protocol for observing Trebouxia flagellate cells over several days by coating microscopy slides with Bold's basal medium. The method preserves cell viability and allows repeated imaging of motile cells in the same areas so that their behaviour and development can be continuously observed. In this way, qualitative observations, such as flagellate cell release, motility, and gamete fusion, can be combined with quantitative analyses of cell morphology. The protocol has proven to be robust and reproducible and was applied to several Trebouxia species. Compared to existing techniques, it allows the monitoring of dynamic processes and provides a powerful tool to study specific life stages not only in Trebouxia but also in other unicellular and colonial green algae. Key features • This protocol allows real-time monitoring over several days of Trebouxia flagellate cells with standard light microscopy. • This protocol preserves cell viability and motility for repeated daily observations of the same cell groups. • This protocol is simple, low-cost, and adaptable to other motile algal cells. • This protocol is based on the methodology described in [1], where it was originally applied and validated.

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. Key features • Optimized nuclei isolation using methanol fixation and lipid removal enables efficient nuclei extraction from the fragile, lipid-rich fat body tissue of Anopheles gambiae. • We validated this procedure in sugar-fed and blood-fed samples, yielding high-quality single-nucleus RNA sequencing data with high gene counts and low mitochondrial RNA content. • Robust quality metrics allow effective filtering of ambient RNA, enhancing transcriptomic accuracy across different physiological states.

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. Key features • Quantification of HSV-1 nuclear egress by flow cytometry using a double fluorescent reporter system. • The assay is suitable for large-scale high-throughput screens, e.g., CRISPR/Cas9. • The assay can be adapted for use with other herpesviruses, provided a mature capsid-specific antibody is available.

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. Key features • A beginner-friendly protocol for users without access to a cryo-EM core/suite at their institution that utilizes basic equipment. • This protocol focuses on optimizing cell seeding and blotting to yield grids with thin ice and evenly distributed cells. • Grids prepared using this protocol can be used for focused ion beam milling.

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. Key features • The protocol presents the design of a compact Schlieren optics device (CSOD), with image boundaries enhanced by a slight shift in two overlapping, virtual images. • The CSOD captures high-resolution images of a transparent medium with variation in thickness or index of refraction. • The CSOD can detect transparent samples with thickness in the order of 1 µm; it is simple to build, user-friendly, and portable. • As a cheaper and portable complement to a phase contrast microscope, the device can image large samples more conveniently.

Since its introduction, the CRISPR/Cas9 system has been used in many organisms for precise and rapid genome editing, as well as for editing multiple genes at once. This targeted mutagenesis makes it easy to analyze the function of a gene of interest (goi). The standard method for genetic manipulation of the model organism Neurospora crassa has been homologous recombination. It is well established and widely used to create knock-out or overexpression mutants. The recently developed CRISPR/Cas9 system is an addition to the toolkit for genetically manipulating N. crassa. For this protocol, a strain stably expressing the Cas9 endonuclease is required. After designing the gRNA with the online tool CHOP-CHOP, a synthetic gRNA is used to transform macroconidia via electroporation. Combining the goi-gRNA with a gRNA targeting the csr-1 gene as a selection marker allows for easy identification of colonies with mutations at the target site of the goi, since the obtained resistance to Cyclosporin A (CsA) allows for selecting editing events. The mutation type can be detected by PCR of the edited gene region followed by Sanger sequencing. This system is fast and easy to handle, offering an attractive alternative to homologous recombination, especially for targeting multiple genes simultaneously. Key features • This protocol allows the use of CRISPR/Cas9 in Neurospora crassa to create loss-of-function mutants. • It can be used to create loss-of-function mutants of multiple genes in one round of transformation. • Time-saving mutagenesis without the need for vector construction. • In combination with csr-1 as a selection marker, the screening for successfully edited genes of interest is reduced.

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. Key features • Enables live, single-cell resolution imaging of ER-phagy dynamics across intact tissues in mice. • Features a Cre-recombinase-activated knock-in model (CA-ER-TRG) for spatiotemporally controlled ER-phagy studies in specific cell types. • Quantifies ER-phagy flux via pH-sensitive RFP-GFP signal ratiometry and lysosomal co-localization in vivo.

The protochlorophyllide (Pchlide) level is a crucial indicator of plant fitness. Precise quantification of Pchlide content is necessary not only in studies of flu-related mutants that over-accumulate Pchlide in the dark but also for research on plants suffering from environmental stresses. Due to its low content and interference of chlorophylls, quantitative determination of Pchlide content is a challenge. Here, we describe an optimized protocol for Pchlide extraction from Arabidopsis thaliana seedlings and subsequent analysis using high-performance liquid chromatography (HPLC) coupled with fluorescence detection. Divinyl-Protochlorophyllide (DV-Pchlide, the major form of Pchlide in plants) quantification is achieved by interpolating fluorescence peak areas against an experimentally derived standard curve. This protocol provides a reliable workflow for Pchlide quantification, facilitating the deciphering of the underlying mechanism of plant environmental resilience. Key features • This method adopts acetone as a solvent for both Pchlide extraction and HPLC run. • This protocol adopts a gradient HPLC system equipped with a fluorescence detector. • This protocol applies an experimentally derived standard calibration curve using synthetic DV-Pchlide.

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. Key features • Visualizes the activation of small GTPases in living cells as mNeonGreen fluorescence signal. • Can be applied to small GTPases whose effectors have not yet been identified. • SAIYAN exploits the intrinsic property of small GTPases to associate with cellular membranes upon activation.

Autonomic regulation of heart and respiratory rates is essential for understanding brain-body interactions in health and disease. Preclinical cardiovascular recordings are often performed under anesthesia or via telemetry, both of which introduce physiological confounds such as stress or impaired recovery due to the need for acute or chronic implantation of sensors. Here, we present a minimally invasive protocol for simultaneous acquisition of high-quality electrocardiography and respiratory signals in awake mice. Using an in-house-modified physiological monitor in awake, head-fixed mice that were briefly habituated to experimental conditions, we ultimately enable stable, long-term physiological recordings alongside in vivo microscopy. This protocol provides a robust, low-stress method for acquiring physiological signals, enabling the simultaneous study of cardiovascular-cerebral dynamics in awake head-fixed mice, thereby enhancing the translational relevance of preclinical measurements. Key features • Minimally invasive electrocardiogram and respiration rate acquisition in awake, head-fixed mice suitable for long-term physiological recordings. • Custom-built setup integrates physiological monitoring with in vivo imaging without surgical implantation or telemetry. • Rapid habituation protocol ensures low-stress conditions and high-quality signal acquisition in conscious mice. • Enables correlation of cardiovascular dynamics with brain activity and cerebrospinal fluid flow in translational neuroscience studies.