细胞生物学

The mammalian kinetochore is a multi-layered protein complex that forms on the centromeric chromatin. The kinetochore serves as the attachment hub for the plus ends of microtubules emanating from the centrosomes during mitosis. For karyokinesis, bipolar kinetochore-microtubule attachment and subsequent microtubule depolymerization lead to the development of inter-kinetochore tension between the sister chromatids. These events are instrumental in initiating a signaling cascade culminating in the segregation of the sister chromatids equally between the new daughter cells. Of the hundreds of conserved proteins that constitute the mammalian kinetochore, many that reside in the outermost layer are loaded during early mitosis and removed around metaphase-anaphase. Dynamically localized kinetochore proteins include those required for kinetochore-microtubule attachment, spindle assembly checkpoint proteins, various kinases, and molecular motors. The abundance of these kinetochore-localized proteins varies at prometaphase, metaphase, and anaphase, and is thus considered diagnostic of the fidelity of progression through these stages of mitosis. Here, we document detailed, state-of-the-art methodologies based on high-resolution fluorescence confocal microscopy followed by quantification of the levels of kinetochore-localized proteins during mitosis. We also document methods to accurately measure distances between sister kinetochores in mammalian cells, a surrogate readout for inter-kinetochore tension, which is essential for chromosome segregation.

Phosphoinositides are rare membrane lipids that mediate cell signaling and membrane dynamics. PI(4)P and PI(3)P are two major phosphoinositides crucial for endolysosomal functions and dynamics, making them the lipids of interest in many studies. The acute modulation of phosphoinositides at a given organelle membrane can reveal important insights into their cellular function. Indeed, the localized depletion of PI(4)P and PI(3)P is a viable tool to assess the importance of these phosphoinositides in various experimental conditions. Here, we describe a live imaging method to acutely deplete PI(4)P and PI(3)P on endolysosomes. The depletion assay utilizes the GAI-GID1 or the FRB-FKBP inducible dimerization system to target the catalytic domain of the PI(4)P phosphatase, Sac1, or the PI(3)P phosphatase domain of MTM1 to the endolysosome for localized depletion of these phosphoinositides. By using the fluorescently tagged biosensors, 2xP4M and PX, we can validate and monitor the depletion of PI(4)P and PI(3)P, respectively, on endolysosomes in real-time. We discuss a method for normalizing the fluorescence measurements to appropriate the relative amount of these phosphoinositides in the organellar membranes (endolysosomes), which is required for monitoring PI(4)P or PI(3)P levels during the acute depletion assay. Since the localization of the dimerization partners is specified by the membrane targeting signal, our protocol will be useful for studying the signaling and functions of phosphoinositides at any membrane.

The process of T-lymphocyte migration involves a complex interplay of chemical and mechanical signals. Mechanotransduction mechanisms in T lymphocytes enable them to efficiently navigate through diverse architectural and topographical features of the dynamic tissue macro- and micro-niches encountered during immune responses. Piezo1 mechanosensors are crucial for driving optimal T-cell migration by driving actin-cytoskeletal remodeling. Chemokine-stimulated T lymphocytes demonstrate significant asymmetry or polarity of Piezo1 and actin along the cell axis. The establishment and maintenance of polarity in migrating cells are paramount for facilitating coordinated and directional movements along gradients of chemokine signals. Live-cell imaging techniques are widely employed to study the trajectories of migrating cells. Our approach expands upon current methodologies by not only tracking migrating cells but also imaging fluorescently labeled cellular components. Specifically, our method enables measurement of protein enrichment in the front and rear halves of the moving cell by analyzing the temporal direction of cell trajectories, subsequently bisecting the cell into front-back halves, and measuring the intensities of the fluorescent signals in each cell half at each time frame. Our protocol also facilitates the quantification of the angular distribution of fluorescent signals, enabling visualization of the spatial distribution of signals relative to the direction of cell migration. The protocol describes the examination of polarity in chemokine-treated Jurkat cells transfected with Piezo1-mCherry and actin-GFP constructs. This approach can be extended to live-cell imaging and polarity assessment of other fluorescently labeled proteins.

PD-1 is an immune checkpoint on T cells. Antibodies to PD-1 or its ligand PD-L1 are gaining popularity as a leading immunotherapy approach. In the US, 40% of all cancer patients will be treated with anti-PD-1 or anti-PD-L1 antibodies but, unfortunately, only 30% will respond, and many will develop immune-related adverse events. There are nine FDA-approved anti-PD-1/PD-L1 antibodies, and approximately 100 are in different stages of clinical development. It is a clinical challenge to choose the correct antibody for a given patient, and this is critical in advanced malignancies, which often do not permit a second-line intervention. To resolve that, an in vitro assay to compare the performance of the different anti-PD-1/PD-L1 antibodies is not only a critical tool for research purposes but also a possible tool for personalized medicine. There are some assays describing the binding affinity and function of anti-PD-1/PD-L1 antibodies. However, a significant limitation of existing assays is that they need to consider the location of PD-1 in the immune synapse, the interface between the T cell and tumor cells, and, therefore, ignore a critical component in its biology. To address this, we developed and validated an imaging-based assay to quantify and compare the ability of different anti-PD-1/PD-L1 antibodies to remove PD-1 from the immune synapse. We correlated that with the same antibodies' ability to increase cytokine secretion from the targeted cells. The strong correlation between PD-1 location and its function in vitro and in vivo within the antibody treatment setting validates this assay's usability, which is easily recordable and straightforward.

In this protocol, we focused on analyzing internal branches of Drosophila class IV neurons. These neurons are characterized by their highly branched axons and dendrites and intricately tile the larval body. As Drosophila larvae progress through developmental stages, the dendritic arbors of Class IV neurons undergo notable transformations. As Drosophila larvae develop, their Class IV dendritic arbors grow. In the initial 24 h after egg laying (AEL), the dendrites are smaller than segments. During the subsequent 24 h of the first instar larval stage, dendritic arbors outpace segment growth, achieving tiling. After 48 h, arbors and segments grow concurrently. Epidermal cells near Class IV dendrites expand in proportion to segment growth. This observation suggested that Class IV cells might grow via branch dilation—uniformly elongating branches, akin to Class I cells [1,2]. To understand whether the class IV complex arbor structure is formed by dilation or simply from growing tips, we developed this protocol to introduce a systematic approach for quantitatively assessing the growth dynamics of internal branches.

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.

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.

Autophagy is a conserved homeostatic mechanism involved in cellular homeostasis and many disease processes. Although it was first described in yeast cells undergoing starvation, we have learned over the years that autophagy gets activated in many stress conditions and during development and aging in mammalian cells. Understanding the fundamental mechanisms underlying autophagy effects can bring us closer to better insights into the pathogenesis of many disease conditions (e.g., cardiac muscle necrosis, Alzheimer’s disease, and chronic lung injury). Due to the complex and dynamic nature of the autophagic processes, many different techniques (e.g., western blotting, fluorescent labeling, and genetic modifications of key autophagy proteins) have been developed to delineate autophagy effects. Although these methods are valid, they are not well suited for the assessment of time-dependent autophagy kinetics. Here, we describe a novel approach: the use of DAPRed for autophagic flux measurement via live cell imaging, utilizing A549 cells, that can visualize and quantify autophagic flux in real time in single live cells. This approach is relatively straightforward in comparison to other experimental procedures and should be applicable to any in vitro cell/tissue models.

Key features

• Allows real-time qualitative imaging of autophagic flux at single-cell level.

• Primary cells and cell lines can also be utilized with this technique.

• Use of confocal microscopy allows visualization of autophagy without disturbing cellular functions.

Neurons communicate via synapses—specialized structures that consist of a presynaptic terminal of one neuron and a postsynaptic terminal of another. As knowledge is emerging that mutations in molecules that regulate synaptic function underpin many neurological disorders, it is crucial to elucidate the molecular mechanisms regulating synaptic function to understand synaptic strength, plasticity, modulation, and pathology, which ultimately impact neuronal circuit output and behavior. The presynaptic calyx of Held is a large glutamatergic presynaptic terminal in the auditory brainstem, which due to its accessibility and the possibility to selectively perform molecular perturbations on it, is an ideal model to study the role of presynaptic proteins in regulating synaptic function. In this protocol, we describe the use of confocal imaging and three-dimensional reconstruction of the calyx of Held to assess alterations in gross morphology following molecular perturbation. Using viral-vector delivery to perform molecular perturbations at distinct developmental time points, we provide a fast and cost-effective method to investigate how presynaptic proteins regulate gross morphology such as surface area and synapse volume throughout the lifetime of a neuronal circuit.

Key features

• Confocal imaging and 3D reconstruction of presynaptic terminals.

• Used with a virus-mediated expression of mEGFP to achieve efficient, cell-type specific labeling of the presynaptic compartment.

• Protocol was developed with the calyx of Held but is suitable for pre- and postsynaptic compartments of various neurons across multiple mammalian and invertebrate species.