Cell Biology

Histological techniques to study muscle are crucial for assessing skeletal muscle health. To preserve tissue morphology, samples are usually fixed in formaldehyde or cryopreserved immediately after excision from the body. Freezing samples in liquid nitrogen, using isopentane as a mediator for efficient cooling, preserves the tissue in its natural state. However, this method is highly susceptible to freeze-fracture artifacts, which alter or destroy tissue architecture. Isopentane is most commonly used in a semi-frozen/liquid state that is visually assessed by the experimenter, which can pose a challenge when freezing multiple tissues at a time or maintaining a consistent temperature. Furthermore, tissue size is also a confounding factor; depending on the size, freezing times can vary. In this study, we compare two different options for using isopentane while cryopreserving tissue. We also present an easy and reproducible method of freezing the soleus tissue of mice using frozen isopentane. This method decreased the occurrence of freeze-fractures by an order of magnitude, to ~4%, whereas the traditional method of cryopreservation resulted in ~56% freeze-fracturing.

Neurons are highly polarized cells, with axons that may innervate distant target regions. In the brain, basal forebrain cholinergic neurons (BFCNs) possess extensive axons that project to several target regions such as the cortex, hippocampus, and amygdala, and may be exposed to a specific microenvironment in their axon targets that may have retrograde effects on neuronal health. Interestingly, BFCNs express the pan-neurotrophin receptor p75NTR throughout life while also concomitantly co-expressing all Trk receptors, making them capable of responding to both mature and precursor neurotrophins to promote survival or apoptosis, respectively. Levels of these trophic factors may be modulated in the BFCN axon or soma microenvironment under neurodegenerative conditions such as seizure and brain injury. In this protocol, BFCNs are established in microfluidic devices for compartmental culture, with the aim of studying the effects of axon- or soma-specific stimulation of BFCNs for an in vitro representation of distal axon vs. soma environments as seen in vivo. This study further establishes a novel method of tracing and imaging live BFCNs exposed to stimuli in their distal axons with the aim of assessing retrograde cell death. The in vitro compartmental culture system of BFCNs that allows live imaging may be applied to investigate various effects of axon- or soma-specific stimuli that affect BFCN health, maintenance, and death, to model events that occur in the context of brain injury and neurodegenerative disorders.

Mitochondrial cristae, formed by folding the mitochondrial inner membrane (IM), are essential for cellular energy supply. However, the observation of the IM is challenging due to the limitations in spatiotemporal resolution offered by conventional microscopy and the absence of suitable in vitro probes specifically targeting the IM. Here, we describe a detailed imaging protocol for the mitochondrial inner membrane using the Si-rhodamine dye HBmito Crimson, which has excellent photophysical properties, to label live cells for imaging via stimulated emission depletion (STED) microscopy. This allows for STED imaging over more than 500 frames (approximately one hour), with a spatial resolution of 40 nm, enabling the observation of cristae dynamics during various mitochondrial processes. The protocol includes detailed steps for cell staining, image acquisition, image processing, and resolution analysis. Utilizing the superior resolution of STED microscopy, the structure and complex dynamic changes of cristae can be visualized.

The bone is a highly dynamic organ that undergoes continuous remodeling through an intricate balance of bone formation and degradation. Hyperactivation of the bone-degrading cells, the osteoclasts (OCs), occurs in disease conditions and hormonal changes in females, resulting in osteoporosis, a disease characterized by altered microarchitecture of the bone tissue, and increased bone fragility. Thus, building robust assays to quantify OC resorptive activity to examine the molecular mechanisms underlying bone degradation is critical. Here, we establish an in vitro model to investigate the effect of estrogen withdrawal on OCs derived from the mouse macrophage RAW 264.7 cell line in a bone biomimetic microenvironment. This simple and robust model can also be adapted to examine the effect of drugs and genetic factors influencing OC resorptive activity in addition to being compatible with fluorescent imaging.

Cell-generated forces play a critical role in driving and regulating complex biological processes, such as cell migration and division and cell and tissue morphogenesis in development and disease. Traction force microscopy (TFM) is an established technique developed in the field of mechanobiology used to quantify cellular forces exerted on soft substrates and internal mechanical tissue stresses. TFM measures cell-generated traction forces in 2D or 3D environments with varying mechanical and biochemical properties. This technique involves embedding fiducial markers in the substrate, imaging substrate deformations caused by the cells, and using mathematical models to infer forces. This protocol compiles procedures from various previously published studies and software packages and describes how to perform TFM on 2D micropatterned substrates. Although not the focus of this protocol, the methods and software packages shown here also allow to perform monolayer stress microscopy (MSM), a method to calculate internal mechanical stress within the cells by modeling them as a thin plate with linear and homogeneous material properties. TFM and MSM are non-invasive methods capable of yielding spatially and temporally resolved force and stress maps with high throughput. As such, they enable the generation of rich datasets, which can provide valuable insights into the roles of cell-generated forces in various physiological and pathological processes.

The human intestine plays a pivotal role in nutrient absorption and immune system regulation. Along the longitudinal axis, cell-type composition changes to meet the varying functional requirements. Therefore, our protocol focuses on the processing of the whole human intestine to facilitate the analysis of region-specific characteristics such as tissue architecture and changes in cell populations. We describe how to generate a biobank that can be used to isolate specific immune cell subtypes, generate organoid lines, and establish autologous immune cell-organoid co-cultures.

The human T-lymphotropic virus type-1 (HTLV-1) is an oncogenic retrovirus that predominantly spreads through cell-to-cell contact due to the limited infectivity of cell-free viruses. Among various modes of intercellular transmission, HTLV-1 biofilms emerge as adhesive structures, polarized at the cell surface, which encapsulate virions within a protective matrix. This biofilm is supposed to facilitate simultaneous virion delivery during infection. Yet, the molecular and functional intricacies of viral biofilms remain largely unexplored, despite their pivotal role in understanding retroviral pathogenesis. In this study, we optimized a protocol to isolate HTLV-1 biofilms from chronically infected T cells, facilitating their structural and molecular characterization using proteomic and super-resolution microscopy analyses. This protocol involves cultivating HTLV-1 chronically infected T cells at high density to facilitate the natural detachment of viral biofilms into the supernatant. Then, employing successive centrifugations, the cells are separated from the detached biofilms, and these structures are pelleted at medium speed (10,000× g). This method circumvents the need for mechanical, chemical, or enzymatic biofilm detachment, bypasses the use of ultracentrifugation, and enables us to resuspend the biofilms in the appropriate buffer for subsequent analyses such as western blotting or super-resolution microscopy imaging as presented.

The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.

Proteomics analysis is crucial for understanding the molecular mechanisms underlying muscle adaptations to different types of exercise, such as concentric and eccentric training. Traditional methods like two-dimensional gel electrophoresis and standard mass spectrometry have been used to analyze muscle protein content and modifications. This protocol details the preparation of muscle samples for proteomics analysis using ultra-high-performance liquid chromatography (UHPLC). It includes steps for muscle biopsy collection, protein extraction, digestion, and UHPLC-based analysis. The UHPLC method offers high-resolution separation of complex protein mixtures, providing more detailed and accurate proteomic profiles compared to conventional techniques. This protocol significantly enhances sensitivity, reproducibility, and efficiency, making it ideal for comprehensive muscle proteomics studies.

Sterol regulatory element binding proteins (SREBPs) are transcription factors that reside in the endoplasmic reticulum (ER) membrane as inactive precursors. To be active, SREBPs are translocated to the Golgi where the transcriptionally active N-terminus is cleaved and released to the nucleus to regulate gene expression. Nuclear SREBP levels can be determined by immunoblot analysis; however, this method can only determine the steady-state levels of nuclear SREBPs and does not capture the actual status of activation. The vesicle budding assay provides an alternative way to quantify the activation of SREBPs by monitoring the initiation of SREBP translocation from the ER to the Golgi through vesicles. Microsomal membranes isolated from the liver are incubated in a reaction buffer containing the necessary components to facilitate vesicle formation. Microsomal membranes and vesicles are isolated and SREBPs are quantified in each by immunoblot analysis. The amount of SREBPs found in the budded vesicles provides an assessment of the SREBP activation in the liver.