Cell Biology

As an essential process for the maintenance of cellular homeostasis and function, autophagy is responsible for the lysosome-mediated degradation of damaged proteins and organelles; therefore, dysregulation of autophagy in humans can lead to a variety of diseases. The link between impaired autophagy and disease highlights the need to investigate possible interventions to address dysregulations. One possible intervention is hyperthermia, which is described in this protocol. To investigate these interventions, a method for absolute quantification of autophagosomal compartments is required that allows comparison of autophagosomal activity under different conditions. Existing methods such as western blotting and immunohistochemistry for analysing the location and relative abundance of intracellular proteins associated with autophagy, or transmission electron microscopy (TEM), which are either very time-consuming, expensive, or both, are less suitable for this purpose. The method described in this protocol allows the absolute quantification of autophagosomes per cell in human fibroblasts using the CYTO-ID^® Autophagy Detection Kit after heat therapy compared to a control. The Cyto-ID^® assay is based on the use of a specific dye that selectively stains autophagic compartments, combined with an additional Hoechst 33342 dye for nuclear staining. The subsequent recognition of these stained compartments by the Cytation Imager enables the software to determine the number of autophagosomes per nucleus in living cells. Additionally, this absolute quantification uses an image-based method, and the protocol is easy to use and not time-consuming. Furthermore, the method is not only suitable for heat therapy but can also be adapted to any other desired therapy or substance.

Diseases caused by trypanosomatid parasites remain a significant unmet medical need for millions of people globally. Trypanosomatid parasites such as Trypanosoma cruzi and subspecies of Trypanosoma brucei cause Chagas disease and human African trypanosomiasis (HAT), respectively. Although efforts to find novel treatments have been successful for HAT, Chagas disease is still treated with decades-old therapies that suffer from long treatment durations and severe safety concerns. We recently described the identification and characterization of the cyanotriazole compound class that kills trypanosomes, in vitro and in vivo, by selective inhibition of the trypanosome nuclear topoisomerase II enzyme. To evaluate whether inhibition of the topoisomerase II enzyme led to parasite death due to lethal double-strand DNA breaks, we developed assays for detecting DNA damage in both intracellular amastigotes of T. cruzi and bloodstream-form T. brucei by using the canonical DNA damage marker γH2A. Herein, this article describes the protocols for detecting DNA damage using an immunofluorescence assessment of γH2A by microscopy in trypanosome parasites.

All aerial organs in plants originate from the shoot apical meristem, a specialized tissue at the tip of a plant, enclosing a few stem cells. Understanding developmental dynamics within this tissue in relation to internal and external stimuli is of crucial importance. Imaging the meristem at the cellular level beyond very early stages requires the apex to be detached from the plant body, a procedure that does not allow studies in living, intact plants over longer periods. This protocol describes a new confocal microscopy method with the potential to image the shoot apical meristem of an intact, soil-grown, flowering Arabidopsis plant over several days. The setup opens new avenues to study apical stem cells, their interconnection with the whole plant, and their responses to environmental stimuli.

Cells need to migrate along gradients of chemicals (chemotaxis) in the course of development, wound healing, or immune responses. Neutrophils are prototypical migratory cells that are rapidly recruited to injured or infected tissues from the bloodstream. Their chemotaxis to these inflammatory sites involves changes in cytoskeletal dynamics in response to gradients of chemicals produced therein. Neutrophil chemotaxis has been largely studied in vitro; few assays have been developed to monitor gradient responses in complex living tissues. Here, we describe a laser-wound assay to generate focal injury in zebrafish larvae and monitor changes in behaviour and cytoskeletal dynamics. The first step is to cross adult fish and collect and rear embryos expressing a relevant fluorescent reporter (for example, Lifeact-mRuby, which labels dynamic actin) to an early larval stage. Subsequently, larvae are mounted and prepared for live imaging and wounding under a two-photon microscope. Finally, the resulting data are processed and used for cell segmentation and quantification of actin dynamics. Altogether, this assay allows the visualisation of cellular dynamics in response to acute injury at high resolution and can be combined with other manipulations, such as genetic or chemical perturbations.

Leishmaniasis, a neglected tropical disease, is caused by the intracellular protozoan parasite Leishmania. Upon its transmission through a sandfly bite, Leishmania binds and enters host phagocytic cells, ultimately resulting in a cutaneous or visceral form of the disease. The limited therapeutics available for leishmaniasis, in combination with this parasite’s techniques to evade the host immune system, call for exploring various methods to target this infection. To this end, our laboratory has been characterizing how Leishmania is internalized by phagocytic cells through the activation of multiple host cell signaling pathways. This protocol, which we use routinely for our experiments, delineates how to infect mammalian macrophages with either promastigote or amastigote forms of the Leishmania parasite. Subsequently, the number of intracellular parasites, external parasites, and macrophages can be quantified using immunofluorescence microscopy and semi-automated analysis protocols. Studying the pathways that underlie Leishmania uptake by phagocytes will not only improve our understanding of these host–pathogen interactions but may also provide a foundation for discovering additional treatments for leishmaniasis.

Calcium signalling in the endocardium is critical for heart valve development. Calcium ion pulses in the endocardium are generated in response to mechanical forces due to blood flow and can be visualised in the beating zebrafish heart using a genetically encoded calcium indicator such as GCaMP7a. Analysing these pulses is challenging because of the rapid movement of the heart during heartbeat. This protocol outlines an imaging analysis method used to phase-match the cardiac cycle in single z-slice movies of the beating heart, allowing easy measurement of the calcium signal.

Lipid nanoparticle (LNP)-based drug delivery systems (DDSs) are widely recognized for their ability to enhance efficient and precise delivery of therapeutic agents, including nucleic acids like DNA and mRNA. Despite this acknowledgment, there is a notable knowledge gap regarding the systemic biodistribution and organ accumulation of these nanoparticles. The ability to track LNPs in vivo is crucial for understanding their fate within biological systems. Fluorescent labeling of LNPs facilitates real-time tracking, quantification, and visualization of their behavior within biological systems, providing valuable insights into biodistribution, cellular uptake, and the optimization of drug delivery strategies. Our prior research established reversely engineered LNPs as an exceptional mRNA delivery platform for treating ulcerative colitis. This study presents a detailed protocol for labeling interleukin-22 (IL-22) mRNA-loaded LNPs, their oral administration to mice, and visualization of DiR-labeled LNPs biodistribution in the gastrointestinal tract using IVIS spectrum. This fluorescence-based approach will assist researchers in gaining a dynamic understanding of nanoparticle fate in other models of interest.

The cation-independent mannose 6-phosphate receptors (CI-M6PR) bind newly synthesized mannose 6-phosphate (Man-6-P)-tagged enzymes in the Golgi and transport them to late endosomes/lysosomes, providing them with degradative functions. Following the cargo delivery, empty receptors are recycled via early/recycling endosomes back to the trans-Golgi network (TGN) retrogradely in a dynein-dependent motion. One of the most widely used methods for studying the retrograde trafficking of CI-M6PR involves employing the CD8α-CI-M6PR chimera. This chimera, comprising a CD8 ectodomain fused with the cytoplasmic tail of the CI-M6PR receptor, allows for labeling at the plasma membrane, followed by trafficking only in a retrograde direction. Previous studies utilizing the CD8α-CI-M6PR chimera have focused mainly on colocalization studies with various endocytic markers under steady-state conditions. This protocol extends the application of the CD8α-CI-M6PR chimera to live cell imaging, followed by a quantitative analysis of its motion towards the Golgi. Additionally, we present an approach to quantify parameters such as speed and track lengths associated with the motility of CD8α-CI-M6PR endosomes using the Fiji plugin TrackMate.

Precision-cut lung slices (PCLS), ex vivo 3D lung tissue models, have been widely used for various applications in lung research. PCLS serve as an excellent intermediary between in vitro and in vivo models because they retain all resident cell types within their natural niche while preserving the extracellular matrix environment. This protocol describes the TReATS (TAT-Cre recombinase-mediated floxed allele modification in tissue slices) method that enables rapid and efficient gene modification in PCLS derived from adult floxed animals. Here, we present detailed protocols for the TReATS method, consisting of two simple steps: PCLS generation and incubation in a TAT-Cre recombinase solution. Subsequent validation of gene modification involves live staining and imaging of PCLS, quantitative real-time PCR, and cell viability assessment. This four-day protocol eliminates the need for complex Cre-breeding, circumvents issues with premature lethality related to gene mutation, and significantly reduces the use of animals. The TReATS method offers a simple and reproducible solution for gene modification in complex ex vivo tissue-based models, accelerating the study of gene function, disease mechanisms, and the discovery of drug targets.

Cultured mammalian cells are a common model system for the study of epithelial biology and mechanics. Epithelia are often considered as pseudo–two dimensional and thus imaged and analyzed with respect to the apical tissue surface. We found that the three-dimensional architecture of epithelial monolayers can vary widely even within small culture wells, and that layers that appear organized in the plane of the tissue can show gross disorganization in the apical-basal plane. Epithelial cell shapes should be analyzed in 3D to understand the architecture and maturity of the cultured tissue to accurately compare between experiments. Here, we present a detailed protocol for the use of our image analysis pipeline, Automated Layer Analysis (ALAn), developed to quantitatively characterize the architecture of cultured epithelial layers. ALAn is based on a set of rules that are applied to the spatial distributions of DNA and actin signals in the apical-basal (depth) dimension of cultured layers obtained from imaging cultured cell layers using a confocal microscope. ALAn facilitates reproducibility across experiments, investigations, and labs, providing users with quantitative, unbiased characterization of epithelial architecture and maturity.

Key features

• This protocol was developed to spatially analyze epithelial monolayers in an automated and unbiased fashion.

• ALAn requires two inputs: the spatial distributions of nuclei and actin in cultured cells obtained using confocal fluorescence microscopy.

• ALAn code is written in Python3 using the Jupyter Notebook interactive format.

• Optimized for use in Marbin-Darby Canine Kidney (MDCK) cells and successfully applied to characterize human MCF-7 mammary gland–derived and Caco-2 colon carcinoma cells.

• This protocol utilizes Imaris software to segment nuclei but may be adapted for an alternative method. ALAn requires the centroid coordinates and volume of nuclei.

Graphical overview