Biophysics

Lipid droplets (LD), triglycerides and sterol esters among them, are well known for their capacity as lipid storage organelles. Recently, they have emerged as critical cytoplasmic structures involved in numerous biological functions. LD storage is generated de novo by the cell and provides an energy reserve, lipid precursors, and cell protection. Moreover, LD accumulation can be observed in some pathologies as obesity, atherosclerosis, or lung diseases. Fluorescence imaging techniques are the most widely used techniques to visualize cellular compartments in live cells, including LD. Nevertheless, presence of fluorophores can damage subcellular components and induce cytotoxicity, or even alter the dynamics of the organelles. As an alternative to fluorescence microscopy, label-free techniques such as stimulated Raman scattering and coherent anti-stokes Raman scattering microscopy offer a solution to avoid the undesirable effects caused by dyes and fluorescent proteins, but are expensive and complex. Here, we describe a label-free method using live-cell imaging by 3D holotomographic microscopy (Nanolive) to visualize LD accumulation in the MH-S alveolar macrophage cell line after treatment with oleic acid, a monounsaturated fatty acid that promotes lipid accumulation.

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.

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Single-particle electron cryo-microscopy (cryo-EM) is an effective tool to determine high-resolution structures of macromolecular complexes. Its lower requirements for sample concentration and purity make it an accessible method to determine structures of low-abundant protein complexes, such as those isolated from native sources. While there are many approaches to protein purification for cryo-EM, attaining suitable particle quality and abundance is generally the major bottleneck to the typical single-particle project workflow. Here, we present a protocol using budding yeast (S. cerevisiae), in which a tractable immunoprecipitation tag (3xFLAG) is appended at the endogenous locus of a gene of interest (GOI). The modified gene is expressed under its endogenous promoter, and cells are grown and harvested using standard procedures. Our protocol describes the steps in which the tagged proteins and their associated complexes are isolated within three hours of thawing cell lysates, after which the recovered proteins are used directly for cryo-EM specimen preparation. The prioritization of speed maximizes the ability to recover intact, scarce complexes. The protocol is generalizable to soluble yeast proteins that tolerate C-terminal epitope tags.

Graphical abstract

Overview of lysate-to-grid workflow. Yeast cells are transformed to express a tractable tag on a gene of interest. Following cell culture and lysis, particles of interest are rapidly isolated by co-immunoprecipitation and prepared for cryo-EM imaging (created with BioRender.com).

The human immunodeficiency virus 1 (HIV-1) consists of a viral membrane surrounding the conical capsid. The capsid is a protein container assembled from approximately 1,500 copies of the viral capsid protein (CA), functioning as a reaction and transport chamber for the viral genome after cell entry. Transmission electron microscopy (TEM) is a widely used technique for characterizing the ultrastructure of isolated viral capsids after removal of the viral membrane, which otherwise hinders negative staining of structures inside the viral particle for TEM. Here, we provide a protocol to permeabilize the membrane of HIV-1 particles using a pore-forming toxin for negative staining of capsids, which are stabilized with inositol hexakisphosphate to prevent premature capsid disassembly. This approach revealed the pleomorphic nature of capsids with a partially intact membrane surrounding them. The permeabilization strategy using pore-forming toxins can be readily applied to visualize the internal architecture of other enveloped viruses using TEM.

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The ribosome is a complex cellular machinery whose solved structure allowed for an incredible leap in structural biology research. Different ions bind to the ribosome, stabilizing inter-subunit interfaces and structurally linking rRNAs, proteins, and ligands. Besides cations such as K⁺ and Mg²⁺, polyamines are known to stabilize the folding of RNA and overall structure. The bacterial ribosome is composed of a small (30S) subunit containing the decoding center and a large (50S) subunit devoted to peptide bond formation. We have previously shown that the small ribosomal subunit of Staphylococcus aureus is sensitive to changes in ionic conditions and polyamines concentration. In particular, its decoding center, where mRNA codons and tRNA anticodons interact, is prone to structural deformations in the absence of spermidine. Here, we report a detailed protocol for the purification of the intact and functional 30S, achieved through specific ionic conditions and the addition of spermidine. Using this protocol, we obtained the cryo-electron microscopy (cryo-EM) structure of the 30S–mRNA complex from S. aureus at 3.6 Å resolution. The 30S–mRNA complex formation was verified by a toeprinting assay. In this article, we also include a description of toeprinting and cryo-EM protocols. The described protocols can be further used to study the process of translation regulation.

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In eukaryotic cells, RNA Polymerase II (RNAP2) is the enzyme in charge of transcribing mRNA from DNA. RNAP2 possesses an extended carboxy-terminal domain (CTD) that gets dynamically phosphorylated as RNAP2 progresses through the transcription cycle, therefore regulating each step of transcription from recruitment to termination. Although RNAP2 residue-specific phosphorylation has been characterized in fixed cells by immunoprecipitation-based assays, or in live cells by using tandem gene arrays, these assays can mask heterogeneity and limit temporal and spatial resolution. Our protocol employs multi-colored complementary fluorescent antibody-based (Fab) probes to specifically detect the CTD of the RNAP2 (CTD-RNAP2), and its phosphorylated form at the serine 5 residue (Ser5ph-RNAP2) at a single-copy HIV-1 reporter gene. Together with high-resolution fluorescence microscopy, single-molecule tracking analysis, and rigorous computational modeling, our system allows us to visualize, quantify, and predict endogenous RNAP2 phosphorylation dynamics and mRNA synthesis at a single-copy gene, in living cells, and throughout the transcription cycle.

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Schematic of the steps for visualizing, quantifying, and predicting RNAP2 phosphorylation at a single-copy gene.

Kinetoplastids are unicellular eukaryotic parasites responsible for human pathologies such as Chagas disease, sleeping sickness or Leishmaniasis, caused by Trypanosoma cruzi, Trypanosoma brucei, and various Leishmania spp., respectively. They harbor a single large mitochondrion that is essential for the survival of the parasite. Interestingly, most of the mitochondrial gene expression machineries and processes present significant differences from their nuclear and cytosolic counterparts. A striking example concerns their mitochondrial ribosomes, in charge of translating the few essential mRNAs encoded by mitochondrial genomes. Here, we present a detailed protocol including the specific procedures to isolate mitochondria from two species of kinetoplastids, T. cruzi and L. tarentolae, by differential centrifugations. Then, we detail the protocol to purify mitochondrial ribosomal complexes from these two species of parasites (including ribosomal maturating complexes) by a sucrose gradient approach. Finally, we describe how to prepare cryo-electron microscopy (cryo-EM) grids from these two sorts of samples. This protocol will be useful for further studies aiming at analyzing mitochondrial translation regulation.

Polarized actin cables in S. cerevisiae are linear bundles of crosslinked actin filaments that are assembled by two formins, Bnr1 (localized to the bud neck), and Bni1 (localized to the bud tip). Actin is polymerized at these two sites, which results in cables extending along the cell cortex toward the back of the mother cell. These cables serve as polarized tracks for myosin-based transport of secretory vesicles and other cargo, from the mother cell to the growing daughter cell. Until recently, descriptions of actin cable morphology and architecture have largely been qualitative or descriptive in nature. Here, we introduce a new quantitative method that enables more precise characterization of actin cable length. This technological advance generates quantitative datasets that can be used to determine the contributions of different actin regulatory proteins to the maintenance of cable architecture, and to assess how different pharmacological agents affect cable arrays. Additionally, these datasets can be used to test theoretical models, and be compared to results from computational simulations of actin assembly.

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Illustration of actin cable length and morphology analysis.
(A) Representative maximum intensity projection image of S. cerevisiae fixed and stained with fluorescently-conjugated phalloidin to label F-actin (displayed in color), and fluorescently-conjugated Concanavalin A to label the cell wall (displayed in grey scale). Lengths of actin cables traced from the bud neck to their ends are indicated (dashed lines). (B) Inverted grey scale image of F-actin labelled with fluorescently-conjugated phalloidin and the cell wall traced in black. The length (purple) and end-to-end distance (green) of a single actin cable is indicated. Scale bar, 2 µm. (C–E) Actin cable length (C), end-to-end distance (D), and tortuosity (E) from hypothetical datasets, where each data point represents an individual cable and larger symbols represent the mean from each hypothetical experiment. Error bars, 95% confidence intervals.

Bsoft is a software package primarily developed for processing electron micrographs, with the goal of determining the structures of biologically relevant molecules, molecular assemblies, and parts of cells. However, it incorporates many ways to deal with images, from the mundane to very sophisticated algorithms. This article is an introduction into its use, illustrating that it is an extensive toolbox, for manipulating and understanding images. Bsoft has over 150 programs, allowing the user an infinite number of ways to process images. These programs can be executed on the command line, or through the interactive program called brun. The main visualization program is bshow, providing numerous ways to manipulate and interpret images. The primary aim is to provide the user with powerful capabilities, including processing large numbers of images. An important additional aim is to make it as accessible as possible, making it easier to deal with image formats and features, and enhance productivity.

Single molecule tracking (SMT) is a powerful technique to study molecular dynamics, and is particularly adapted to monitor the motion and interactions of cell membrane components. Assessing interactions among two molecular populations is classically performed by several approaches, including dual-color videomicroscopy, which allows monitoring of interactions through colocalization events. Other techniques, such as fluorescence recovery after photobleaching (FRAP), Förster resonance energy transfer (FRET), and fluorescence correlation spectroscopy (FCS), are also utilized to measure molecular dynamics.

We developed MTT2col, a set of algorithmic tools extending multi-target tracing (MTT) to dual-color acquisition (https://github.com/arnauldserge1/MTT2col). In this protocol, we used MTT2col to monitor adhesion molecules at the contact between leukemic stem cells and stromal cells, a process involved in cancer resistance to chemotherapy and in relapse. Our dual-color single molecule protocol includes the following steps: (i) labeling molecules of interest with fluorescent probes, (ii) video-acquisition, (iii) analyses using our MTT2col in-house software, to obtain positions and trajectories, followed by (iv) detailed analyses of colocalization, distribution, and dynamic motion modes, according to the issues addressed. MTT2col is a robust and efficient SMT algorithm. Both MTT and MTT2col are open-source software that can be adapted and further developed for specific analyses.

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