Cell Biology

Accidental wounding of the peripheral nervous system leads to acute neural dysfunction. Normally, chronic deficits are overcome because peripheral nerves naturally regenerate. However, various genetic and metabolic defects can impair their natural regenerative capacity, which may be due to neuron-extrinsic mechanisms. Therefore, characterizing the behavior of multiple cells during nerve injury and repair in vivo is a pressing need in regenerative medicine. Here, we detail a method for precise wounding of sensory axons in zebrafish, followed by high-resolution in toto long-term quantitative videomicroscopy of neurons, Schwann cells, and macrophages. This protocol can be easily adapted to study the effects of targeted genetic or metabolic disruptions in zebrafish and other suitable organisms, as well as for screening pharmacological agents with therapeutic potential.

Graphical overview

The mammalian Golgi complex consists of laterally connected Golgi stacks, each comprising close-packed and flattened membrane sacks called cisternae. However, the convoluted spatial organization of Golgi stacks and limited resolution of light microscopy prevent us from resolving the cisternal organization of the Golgi. Here, we describe our recently developed side-averaging approach coupled with Airyscan microscopy to visualize the cisternal organization of nocodazole-induced Golgi ministacks. First, the nocodazole treatment greatly simplifies the organization of Golgi stacks by spatially separating the crowded and amorphous Golgi complex into individual disk-shaped ministacks. The treatment also makes it possible to identify en face and side-views of Golgi ministacks. Next, after manually selecting the side-view Golgi ministack images, they are transformed and aligned. Finally, the resulting images are averaged to enhance the common structural features and suppress the morphological variations among individual Golgi ministacks. This protocol describes how to image and analyze the intra-Golgi localization of giantin, GalT-mCherry, GM130, and GFP-OSBP in HeLa cells by side-averaging.

Graphical overview

The cell surfaceome is of vital importance across physiology, developmental biology, and disease states alike. The precise identification of proteins and their regulatory mechanisms at the cell membrane has been challenging and is typically determined using confocal microscopy, two-photon microscopy, or total internal reflection fluorescence microscopy (TIRFM). Of these, TIRFM is the most precise, as it harnesses the generation of a spatially delimited evanescent wave at the interface of two surfaces with distinct refractive indices. The limited penetration of the evanescent wave illuminates a narrow specimen field, which facilitates the localization of fluorescently tagged proteins at the cell membrane but not inside of the cell. In addition to constraining the depth of the image, TIRFM also significantly enhances the signal-to-noise ratio, which is particularly valuable in the study of live cells. Here, we detail a protocol for micromirror TIRFM analysis of optogenetically activated protein kinase C-ϵ in HEK293-T cells, as well as data analysis to demonstrate the translocation of this construct to the cell-surface following optogenetic activation.

Graphic abstract

Nucleotide excision repair (NER) removes a wide variety of structurally unrelated lesions from the genome, including UV-induced photolesions such as 6–4 pyrimidine–pyrimidone photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs). NER removes lesions by excising a short stretch of single-stranded DNA containing the damaged DNA, leaving a single-stranded gap that is resynthesized in a process called unscheduled DNA synthesis (UDS). Measuring UDS after UV irradiation in non-dividing cells provides a measure of the overall NER activity, of which approximately 90% is carried out by the global genome repair (GGR) sub pathway. Here, we present a protocol for the microscopy-based analysis and quantification of UDS as a measurement for GGR activity. Following local UV-C irradiation, serum-starved human cells are supplemented with the thymidine analogue 5-ethynyl-2'-deoxyuridine (EdU), which is incorporated into repair patches following NER-dependent dual incision. The incorporated nucleotide analogue is coupled to a fluorophore using Click-iT chemistry, followed by immunodetection of CPD photolesions to simultaneously visualize both signals by fluorescence microscopy. Accompanying this protocol is a custom-built ImageJ plug-in to analyze and quantify UDS signals at sites of CPD-marked local damage. The local UDS assay enables an effective and sensitive fluorescence-based quantification of GGR activity in single cells with application in basic research to better understand the regulatory mechanism in NER, as well as in diagnostics to characterize fibroblasts from individuals with NER-deficiency disorder.

Graphical abstract

Utilizingresources available from the mother's body to guarantee healthy offspring growth is the fundamental reproductive strategy. Recently, we showed that a class of the largest extracellular vesicles known as exophers, which are responsible for the removal of neurotoxic components from neurons (Melentijevic et al., 2017) and damaged mitochondria from cardiomyocytes (Nicolás-Ávila et al., 2020), are released by the Caenorhabditis elegans hermaphrodite body wall muscles (BWM), to support embryonic growth (Turek et al., 2021). Employing worms expressing fluorescent reporters in BWM cells, we found that exopher formation (exophergenesis) is sex-specific and fertility-dependent. Moreover, exophergenesis is regulated by the developing embryo in utero, and exophers serve as transporters for muscle-generated yolk proteins, which can be used to nourish the next generation. Given the specific regulation of muscular exophergenesis, and the fact that muscle-generated exophers are much larger than neuronal ones and have different targeting, their identification and quantification required a modified approach from that designed for neuronal-derived exophers (Arnold et al., 2020). Here, we present a methodology for assessing and quantifying muscle-derived exophers that can be easily extended to determine their function and regulation in various biological contexts.

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Mitochondria are cellular organelles essential for the function and survival of eukaryotic cells. Nearly all mitochondrial proteins are nuclear-encoded and require mitochondrial import upon their synthesis in the cytosol. Various approaches have been described to study mitochondrial protein import, such as monitoring the entry of radiolabeled proteins into purified mitochondria or quantifying newly synthesized proteins within mitochondria by proteomics. Here, we provide a detailed protocol for a commonly used and straightforward assay that quantitatively examines mitochondrial protein import by monitoring the co-localization of mitochondrially targeted enhanced green fluorescent protein (eGFP) with the mitochondrial fluorescence dye MitoTracker ^TM Deep Red FM by live cell imaging. We describe the preparation and use of a stable mammalian cell line inducibly expressing a mitochondrial targeting sequence (MTS)-eGFP, followed by quantitative image analysis using an open-source ImageJ-based plugin. This inducible expression system avoids the need for transient transfection while enabling titration of MTS-eGFP expression and thereby avoiding protein folding stress. Overall, the assay provides a simple and robust approach to assess mitochondrial import capacity of cells in various disease-related settings.

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During an animal's development, a large number of cells undergo apoptosis, a suicidal form of death. These cells are promptly phagocytosed by other cells and degraded inside phagosomes. The recognition, engulfment, and degradation of apoptotic cells is an evolutionarily conserved process occurring in all metazoans. Recently, we discovered a novel event in the nematode Caenorhabditis elegans: the double-membrane autophagosomes are recruited to the surface of phagosomes; subsequently, the outer membrane of an autophagosome fuses with the phagosomal membrane, allowing the inner vesicle to enter the phagosomal lumen and accumulate there over time. This event facilitates the degradation of the apoptotic cell inside the phagosome. During this study, we developed a real-time imaging protocol monitoring the recruitment and fusion of autophagosomes to phagosomes over two hours during embryonic development. This protocol uses a deconvolution-based microscopic imaging system with an optimized setting to minimize photodamage of the embryo during the recording period for high-resolution images. Furthermore, acid-resistant fluorescent reporters are chosen to label autophagosomes, allowing the inner vesicles of an autophagosome to remain visible after entering the acidic phagosomal lumen. The methods described here, which enable high sensitivity, quantitative measurement of each step of the dynamic incorporation in developing embryos, are novel since the incorporation of autophagosomes to phagosomes has not been reported previously. In addition to studying the degradation of apoptotic cells, this protocol can be applied to study the degradation of non-apoptotic cell cargos inside phagosomes, as well as the fusion between other types of intracellular organelles in living C. elegans embryos. Furthermore, its principle of detecting the membrane fusion event can be adapted to study the relationship between autophagosomes and phagosomes or other intracellular organelles in any biological system in which real-time imaging can be conducted.

Understanding the molecular and structural mechanisms that govern the assembly and organization of higher-order actin architecture requires the use of in vitro actin binding and bundling assays. Crosslinking of actin filaments into bundles can be monitored in vitro via several techniques, including negative staining/electron microscopy, low-speed co-sedimentation assay/SDS-PAGE, and fluorescence staining/confocal microscopy. We and others have previously characterized the N-BAR domain of ASAP1, an ADP-ribosylation factor GTPase-activating protein, as an actin-bundling module; we further identified key lysine residues responsible for actin cross-linking. Here, we use the ASAP1 BAR domain as an example and describe a detailed procedure for observing the actin bundle formation by confocal microscopy. This protocol requires small reaction volumes and takes advantage of bright commercially available fluorescent phalloidins, making it an ideal choice for medium-throughput screening of mutants or domain truncations in their ability to bundle actin.

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In the human cell cycle, complete replication of DNA is a fundamental process for the maintenance of genome integrity. Replication stress interfering with the progression of replication forks causes difficult-to-replicate regions to remain under-replicated until the onset of mitosis. In early mitosis, a homology-directed repair DNA synthesis, called mitotic DNA synthesis (MiDAS), is triggered to complete DNA replication. Here, we present a method to detect MiDAS in human U2OS 40-2-6 cells, in which repetitive lacO sequences integrated into the human chromosome evoke replication stress and concomitant incomplete replication of the lacO array. Immunostaining of BrdU and LacI proteins is applied for visualization of DNA synthesis in early mitosis and the lacO array, respectively. This protocol has been established to easily detect MiDAS at specific loci using only common immunostaining methods and may be optimized for the investigation of other difficult-to-replicate regions marked with site-specific binding proteins.

Mitochondrial dysfunction is associated with perturbations in the cellular oxidative status, changes in energy production and metabolic rate, and the onset of pathological processes. Classic methods of assessing mitochondrial dysfunction rely on indirect measures, such as evaluating mitochondrial DNA copy numbers, or direct but more costly and skilled techniques, such as electron microscopy. The protocol presented here was recently implemented to evaluate mitochondrial dysfunction in response to insecticide exposure in Drosophila melanogaster larvae, and it relies on the use of a previously established MitoTimer mutant strain. MitoTimer is a genetically engineered mitochondrial protein that shows green fluorescence when newly synthetized, irreversibly turning into red as mitochondria age. The protocol described here allows for the easy and direct assessment of shifts in mitochondrial turnover, with tissue-specific accuracy. This protocol can be adapted to assess changes in mitochondrial turnover in response to drugs, rearing conditions, and/or mutations in larva, pupa, or adult fruit flies.