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Snap-, CLIP- and Halo-Tag Labelling of Budding Yeast Cells.
PUBLISHED: 01-01-2013
Fluorescence microscopy of the localization and the spatial and temporal dynamics of specifically labelled proteins is an indispensable tool in cell biology. Besides fluorescent proteins as tags, tag-mediated labelling utilizing self-labelling proteins as the SNAP-, CLIP-, or the Halo-tag are widely used, flexible labelling systems relying on exogenously supplied fluorophores. Unfortunately, labelling of live budding yeast cells proved to be challenging with these approaches because of the limited accessibility of the cell interior to the dyes. In this study we developed a fast and reliable electroporation-based labelling protocol for living budding yeast cells expressing SNAP-, CLIP-, or Halo-tagged fusion proteins. For the Halo-tag, we demonstrate that it is crucial to use the 6-carboxy isomers and not the 5-carboxy isomers of important dyes to ensure cell viability. We report on a simple rule for the analysis of (1)H NMR spectra to discriminate between 6- and 5-carboxy isomers of fluorescein and rhodamine derivatives. We demonstrate the usability of the labelling protocol by imaging yeast cells with STED super-resolution microscopy and dual colour live cell microscopy. The large number of available fluorophores for these self-labelling proteins and the simplicity of the protocol described here expands the available toolbox for the model organism Saccharomyces cerevisiae.
Authors: Danette L. Daniels, Jacqui Méndez, Hélène Benink, Andrew Niles, Nancy Murphy, Michael Ford, Richard Jones, Ravi Amunugama, David Allen, Marjeta Urh.
Published: 07-12-2014
Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous proteomes, including those from viruses, bacteria, and yeast.  In comparison, analysis of the human proteome lags behind, partially due to the sheer number of proteins which must be studied, but also the complexity of networks and interactions these present. To specifically address the challenges of understanding the human proteome, we have developed HaloTag technology for protein isolation, particularly strong for isolation of multiprotein complexes and allowing more efficient capture of weak or transient interactions and/or proteins in low abundance.  HaloTag is a genetically encoded protein fusion tag, designed for covalent, specific, and rapid immobilization or labelling of proteins with various ligands. Leveraging these properties, numerous applications for mammalian cells were developed to characterize protein function and here we present methodologies including: protein pull-downs used for discovery of novel interactions or functional assays, and cellular localization. We find significant advantages in the speed, specificity, and covalent capture of fusion proteins to surfaces for proteomic analysis as compared to other traditional non-covalent approaches. We demonstrate these and the broad utility of the technology using two important epigenetic proteins as examples, the human bromodomain protein BRD4, and histone deacetylase HDAC1.  These examples demonstrate the power of this technology in enabling  the discovery of novel interactions and characterizing cellular localization in eukaryotes, which will together further understanding of human functional proteomics.              
21 Related JoVE Articles!
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Nanogold Labeling of the Yeast Endosomal System for Ultrastructural Analyses
Authors: Muriel Mari, Janice Griffith, Fulvio Reggiori.
Institutions: University Medical Center Utrecht.
Endosomes are one of the major membrane sorting checkpoints in eukaryotic cells and they regulate recycling or destruction of proteins mostly from the plasma membrane and the Golgi. As a result the endosomal system plays a central role in maintaining cell homeostasis, and mutations in genes belonging to this network of organelles interconnected by vesicular transport, cause severe pathologies including cancer and neurobiological disorders. It is therefore of prime relevance to understand the mechanisms underlying the biogenesis and organization of the endosomal system. The yeast Saccharomyces cerevisiae has been pivotal in this task. To specifically label and analyze at the ultrastructural level the endosomal system of this model organism, we present here a detailed protocol for the positively charged nanogold uptake by spheroplasts followed by the visualization of these particles through a silver enhancement reaction. This method is also a valuable tool for the morphological examination of mutants with defects in endosomal trafficking. Moreover, it is not only applicable for ultrastructural examinations but it can also be combined with immunogold labelings for protein localization investigations.
Cellular Biology, Issue 89, positively charged nanogold, silver enhancement, Tokuyasu procedure, electron microscopy, immunogold labeling, yeast
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Production and Targeting of Monovalent Quantum Dots
Authors: Daeha Seo, Justin Farlow, Kade Southard, Young-wook Jun, Zev J. Gartner.
Institutions: University of California, San Francisco, University of California, Berkeley, Lawrence Berkeley National Laboratory, University of California, San Francisco, University of California, San Francisco, University of California, San Francisco, University of California, San Francisco.
The multivalent nature of commercial quantum dots (QDs) and the difficulties associated with producing monovalent dots have limited their applications in biology, where clustering and the spatial organization of biomolecules is often the object of study. We describe here a protocol to produce monovalent quantum dots (mQDs) that can be accomplished in most biological research laboratories via a simple mixing of CdSe/ZnS core/shell QDs with phosphorothioate DNA (ptDNA) of defined length. After a single ptDNA strand has wrapped the QD, additional strands are excluded from the surface. Production of mQDs in this manner can be accomplished at small and large scale, with commercial reagents, and in minimal steps. These mQDs can be specifically directed to biological targets by hybridization to a complementary single stranded targeting DNA. We demonstrate the use of these mQDs as imaging probes by labeling SNAP-tagged Notch receptors on live mammalian cells, targeted by mQDs bearing a benzylguanine moiety.
Bioengineering, Issue 92, monovalent quantum dots, single particle tracking, SNAP tag, steric exclusion, phosphorothioate, DNA, nanoparticle bioconjugation, single molecule imaging
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Growth Assays to Assess Polyglutamine Toxicity in Yeast
Authors: Martin L. Duennwald.
Institutions: Boston Biomedical Research Institute.
Protein misfolding is associated with many human diseases, particularly neurodegenerative diseases, such as Alzheimer’s disease, Parkinson's disease, and Huntington's disease 1. Huntington's disease (HD) is caused by the abnormal expansion of a polyglutamine (polyQ) region within the protein huntingtin. The polyQ-expanded huntingtin protein attains an aberrant conformation (i.e. it misfolds) and causes cellular toxicity 2. At least eight further neurodegenerative diseases are caused by polyQ-expansions, including the Spinocerebellar Ataxias and Kennedy’s disease 3. The model organism yeast has facilitated significant insights into the cellular and molecular basis of polyQ-toxicity, including the impact of intra- and inter-molecular factors of polyQ-toxicity, and the identification of cellular pathways that are impaired in cells expressing polyQ-expansion proteins 3-8. Importantly, many aspects of polyQ-toxicity that were found in yeast were reproduced in other experimental systems and to some extent in samples from HD patients, thus demonstrating the significance of the yeast model for the discovery of basic mechanisms underpinning polyQ-toxicity. A direct and relatively simple way to determine polyQ-toxicity in yeast is to measure growth defects of yeast cells expressing polyQ-expansion proteins. This manuscript describes three complementary experimental approaches to determine polyQ-toxicity in yeast by measuring the growth of yeast cells expressing polyQ-expansion proteins. The first two experimental approaches monitor yeast growth on plates, the third approach monitors the growth of liquid yeast cultures using the BioscreenC instrument. Furthermore, this manuscript describes experimental difficulties that can occur when handling yeast polyQ models and outlines strategies that will help to avoid or minimize these difficulties. The protocols described here can be used to identify and to characterize genetic pathways and small molecules that modulate polyQ-toxicity. Moreover, the described assays may serve as templates for accurate analyses of the toxicity caused by other disease-associated misfolded proteins in yeast models.
Molecular Biology, Issue 61, Protein misfolding, yeast, polyglutamine diseases, growth assays
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Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast
Authors: Pernilla Bjerling, Ida Olsson, Xi'nan Meng.
Institutions: University of Uppsala, Swedish University of Agricultural Sciences.
Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.
Molecular Biology, Issue 59, Fission yeast, fluorescence microscopy, nuclear organisation, chromatin, GFP
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Cryosectioning Yeast Communities for Examining Fluorescence Patterns
Authors: Babak Momeni, Wenying Shou.
Institutions: Fred Hutchinson Cancer Research Center.
Microbes typically live in communities. The spatial organization of cells within a community is believed to impact the survival and function of the community1. Optical sectioning techniques, including confocal and two-photon microscopy, have proven useful for observing spatial organization of bacterial and archaeal communities2,3. A combination of confocal imaging and physical sectioning of yeast colonies has revealed internal organization of cells4. However, direct optical sectioning using confocal or two-photon microscopy has been only able to reach a few cell layers deep into yeast colonies. This limitation is likely because of strong scattering of light from yeast cells4. Here, we present a method based on fixing and cryosectioning to obtain spatial distribution of fluorescent cells within Saccharomyces cerevisiae communities. We use methanol as the fixative agent to preserve the spatial distribution of cells. Fixed communities are infiltrated with OCT compound, frozen, and cryosectioned in a cryostat. Fluorescence imaging of the sections reveals the internal organization of fluorescent cells within the community. Examples of yeast communities consisting of strains expressing red and green fluorescent proteins demonstrate the potentials of the cryosectioning method to reveal the spatial distribution of fluorescent cells as well as that of gene expression within yeast colonies2,3. Even though our focus has been on Saccharomyces cerevisiae communities, the same method can potentially be applied to examine other microbial communities.
Microbiology, Issue 70, Molecular Biology, Cellular Biology, Basic Protocols, Yeasts, Saccharomyces cerevisiae, Clinical Laboratory Techniques, Cytological Techniques, Environmental Microbiology, Investigative Techniques, Life Sciences, cryosectioning, sectioning, cryotome, fixing, microbial community, yeast colonies, Saccharomyces cerevisiae, community interactions
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Bimolecular Fluorescence Complementation
Authors: Katy A. Wong, John P. O'Bryan.
Institutions: University of Illinois at Chicago.
Defining the subcellular distribution of signaling complexes is imperative to understanding the output from that complex. Conventional methods such as immunoprecipitation do not provide information on the spatial localization of complexes. In contrast, BiFC monitors the interaction and subcellular compartmentalization of protein complexes. In this method, a fluororescent protein is split into amino- and carboxy-terminal non-fluorescent fragments which are then fused to two proteins of interest. Interaction of the proteins results in reconstitution of the fluorophore (Figure 1)1,2. A limitation of BiFC is that once the fragmented fluorophore is reconstituted the complex is irreversible3. This limitation is advantageous in detecting transient or weak interactions, but precludes a kinetic analysis of complex dynamics. An additional caveat is that the reconstituted flourophore requires 30min to mature and fluoresce, again precluding the observation of real time interactions4. BiFC is a specific example of the protein fragment complementation assay (PCA) which employs reporter proteins such as green fluorescent protein variants (BiFC), dihydrofolate reductase, b-lactamase, and luciferase to measure protein:protein interactions5,6. Alternative methods to study protein:protein interactions in cells include fluorescence co-localization and Förster resonance energy transfer (FRET)7. For co-localization, two proteins are individually tagged either directly with a fluorophore or by indirect immunofluorescence. However, this approach leads to high background of non-interacting proteins making it difficult to interpret co-localization data. In addition, due to the limits of resolution of confocal microscopy, two proteins may appear co-localized without necessarily interacting. With BiFC, fluorescence is only observed when the two proteins of interest interact. FRET is another excellent method for studying protein:protein interactions, but can be technically challenging. FRET experiments require the donor and acceptor to be of similar brightness and stoichiometry in the cell. In addition, one must account for bleed through of the donor into the acceptor channel and vice versa. Unlike FRET, BiFC has little background fluorescence, little post processing of image data, does not require high overexpression, and can detect weak or transient interactions. Bioluminescence resonance energy transfer (BRET) is a method similar to FRET except the donor is an enzyme (e.g. luciferase) that catalyzes a substrate to become bioluminescent thereby exciting an acceptor. BRET lacks the technical problems of bleed through and high background fluorescence but lacks the ability to provide spatial information due to the lack of substrate localization to specific compartments8. Overall, BiFC is an excellent method for visualizing subcellular localization of protein complexes to gain insight into compartmentalized signaling.
Cellular Biology, Issue 50, Fluorescence, imaging, compartmentalized signaling, subcellular localization, signal transduction
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Continuous High-resolution Microscopic Observation of Replicative Aging in Budding Yeast
Authors: Daphne H. E. W. Huberts, Georges E. Janssens, Sung Sik Lee, Ima Avalos Vizcarra, Matthias Heinemann.
Institutions: University of Groningen, University Medical Centre Groningen, University of Groningen, ETH Zurich, ETH Zurich, ETH Zurich.
We demonstrate the use of a simple microfluidic setup, in which single budding yeast cells can be tracked throughout their entire lifespan. The microfluidic chip exploits the size difference between mother and daughter cells using an array of micropads. Upon loading, cells are trapped underneath these micropads, because the distance between the micropad and cover glass is similar to the diameter of a yeast cell (3-4 μm). After the loading procedure, culture medium is continuously flushed through the chip, which not only creates a constant and defined environment throughout the entire experiment, but also flushes out the emerging daughter cells, which are not retained underneath the pads due to their smaller size. The setup retains mother cells so efficiently that in a single experiment up to 50 individual cells can be monitored in a fully automated manner for 5 days or, if necessary, longer. In addition, the excellent optical properties of the chip allow high-resolution imaging of cells during the entire aging process.
Bioengineering, Issue 78, Microbiology, Cellular Biology, Molecular Biology, Biochemistry, Biomedical Engineering, Genetics, Anatomy, Physiology, Lifespan analysis, Live-cell imaging, Microfluidics, Aging, Microscopy, Saccharomyces cerevisiae, microfluidic chip, yeast, cell culture, cells, imaging
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Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS
Authors: Christopher J. Zopf, Narendra Maheshri.
Institutions: Massachusetts Institute of Technology.
Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.
Microbiology, Issue 77, Cellular Biology, Molecular Biology, Genetics, Biophysics, Saccharomyces cerevisiae, Microscopy, Fluorescence, Cell Biology, microscopy/fluorescence and time-lapse, budding yeast, gene expression dynamics, segmentation, lineage tracking, image tracking, software, yeast, cells, imaging
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Genetic Manipulation in Δku80 Strains for Functional Genomic Analysis of Toxoplasma gondii
Authors: Leah M. Rommereim, Miryam A. Hortua Triana, Alejandra Falla, Kiah L. Sanders, Rebekah B. Guevara, David J. Bzik, Barbara A. Fox.
Institutions: The Geisel School of Medicine at Dartmouth.
Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene function and phenotype(s). The development of mutant strains with targeted gene deletions, targeted mutations, complemented gene function, and/or tagged genes provides powerful strategies to address gene function, particularly if these genetic manipulations can be efficiently targeted to the gene locus of interest using integration mediated by double cross over homologous recombination. Due to very high rates of nonhomologous recombination, functional genomic analysis of Toxoplasma gondii has been previously limited by the absence of efficient methods for targeting gene deletions and gene replacements to specific genetic loci. Recently, we abolished the major pathway of nonhomologous recombination in type I and type II strains of T. gondii by deleting the gene encoding the KU80 protein1,2. The Δku80 strains behave normally during tachyzoite (acute) and bradyzoite (chronic) stages in vitro and in vivo and exhibit essentially a 100% frequency of homologous recombination. The Δku80 strains make functional genomic studies feasible on the single gene as well as on the genome scale1-4. Here, we report methods for using type I and type II Δku80Δhxgprt strains to advance gene targeting approaches in T. gondii. We outline efficient methods for generating gene deletions, gene replacements, and tagged genes by targeted insertion or deletion of the hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT) selectable marker. The described gene targeting protocol can be used in a variety of ways in Δku80 strains to advance functional analysis of the parasite genome and to develop single strains that carry multiple targeted genetic manipulations. The application of this genetic method and subsequent phenotypic assays will reveal fundamental and unique aspects of the biology of T. gondii and related significant human pathogens that cause malaria (Plasmodium sp.) and cryptosporidiosis (Cryptosporidium).
Infectious Diseases, Issue 77, Genetics, Microbiology, Infection, Medicine, Immunology, Molecular Biology, Cellular Biology, Biomedical Engineering, Bioengineering, Genomics, Parasitology, Pathology, Apicomplexa, Coccidia, Toxoplasma, Genetic Techniques, Gene Targeting, Eukaryota, Toxoplasma gondii, genetic manipulation, gene targeting, gene deletion, gene replacement, gene tagging, homologous recombination, DNA, sequencing
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Isolation of Cellular Lipid Droplets: Two Purification Techniques Starting from Yeast Cells and Human Placentas
Authors: Jaana Mannik, Alex Meyers, Paul Dalhaimer.
Institutions: University of Tennessee, University of Tennessee.
Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of neutral lipids surrounded by a phospholipid monolayer. One of the most useful techniques in determining the cellular roles of droplets has been proteomic identification of bound proteins, which can be isolated along with the droplets. Here, two methods are described to isolate lipid droplets and their bound proteins from two wide-ranging eukaryotes: fission yeast and human placental villous cells. Although both techniques have differences, the main method - density gradient centrifugation - is shared by both preparations. This shows the wide applicability of the presented droplet isolation techniques. In the first protocol, yeast cells are converted into spheroplasts by enzymatic digestion of their cell walls. The resulting spheroplasts are then gently lysed in a loose-fitting homogenizer. Ficoll is added to the lysate to provide a density gradient, and the mixture is centrifuged three times. After the first spin, the lipid droplets are localized to the white-colored floating layer of the centrifuge tubes along with the endoplasmic reticulum (ER), the plasma membrane, and vacuoles. Two subsequent spins are used to remove these other three organelles. The result is a layer that has only droplets and bound proteins. In the second protocol, placental villous cells are isolated from human term placentas by enzymatic digestion with trypsin and DNase I. The cells are homogenized in a loose-fitting homogenizer. Low-speed and medium-speed centrifugation steps are used to remove unbroken cells, cellular debris, nuclei, and mitochondria. Sucrose is added to the homogenate to provide a density gradient and the mixture is centrifuged to separate the lipid droplets from the other cellular fractions. The purity of the lipid droplets in both protocols is confirmed by Western Blot analysis. The droplet fractions from both preps are suitable for subsequent proteomic and lipidomic analysis.
Bioengineering, Issue 86, Lipid droplet, lipid body, fat body, oil body, Yeast, placenta, placental villous cells, isolation, purification, density gradient centrifugation
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ReAsH/FlAsH Labeling and Image Analysis of Tetracysteine Sensor Proteins in Cells
Authors: Sevgi Irtegun, Yasmin M. Ramdzan, Terrence D. Mulhern, Danny M. Hatters.
Institutions: Bio21 Molecular Science and Biotechnology Institute.
Fluorescent proteins and dyes are essential tools for the study of protein trafficking, localization and function in cells. While fluorescent proteins such as green fluorescence protein (GFP) have been extensively used as fusion partners to proteins to track the properties of a protein of interest1, recent developments with smaller tags enable new functionalities of proteins to be examined in cells such as conformational change and protein-association 2, 3. One small tag system involves a tetracysteine motif (CCXXCC) genetically inserted into a target protein, which binds to biarsenical dyes, ReAsH (red fluorescent) and FlAsH (green fluorescent), with high specificity even in live cells 2. The TC/biarsenical dye system offers far less steric constraints to the host protein than fluorescent proteins which has enabled several new approaches to measure conformational change and protein-protein interactions 4-7. We recently developed a novel application of TC tags as sensors of oligomerization in cells expressing mutant huntingtin, which when mutated aggregates in neurons in Huntington disease 7. Huntingtin was tagged with two fluorescent dyes, one a fluorescent protein to track protein location, and the second a TC tag which only binds biarsenical dyes in monomers. Hence, changes in colocalization between protein and biarsenical dye reactivity enabled submicroscopic oligomer content to be spatially mapped within cells. Here, we describe how to label TC-tagged proteins fused to a fluorescent protein (Cherry, GFP or CFP) with FlAsH or ReAsH in live mammalian cells and how to quantify the two color fluorescence (Cherry/FlAsH, CFP/FlAsH or GFP/ReAsH combinations).
Cell Biology, Issue 54, tetracysteine, TC, ReAsH, FlAsH, biarsenical dyes, fluorescence, imaging, confocal microscopy, ImageJ, GFP
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Identification of Growth Inhibition Phenotypes Induced by Expression of Bacterial Type III Effectors in Yeast
Authors: Dor Salomon, Guido Sessa.
Institutions: Tel Aviv University.
Many Gram-negative pathogenic bacteria use a type III secretion system to translocate a suite of effector proteins into the cytosol of host cells. Within the cell, type III effectors subvert host cellular processes to suppress immune responses and promote pathogen growth. Numerous type III effectors of plant and animal bacterial pathogens have been identified to date, yet only a few of them are well characterized. Understanding the functions of these effectors has been undermined by a combination of functional redundancy in the effector repertoire of a given bacterial strain, the subtle effects that they may exert to increase virulence, roles that are possibly specific to certain infection stages, and difficulties in genetically manipulating certain pathogens. Expression of type III effectors in the budding yeast Saccharomyces cerevisiae may allow circumventing these limitations and aid to the functional characterization of effector proteins. Because type III effectors often target cellular processes that are conserved between yeast and other eukaryotes, their expression in yeast may result in growth inhibition phenotypes that can be exploited to elucidate effector functions and targets. Additional advantages to using yeast for functional studies of bacterial effectors include their genetic tractability, information on predicted functions of the vast majority of their ORFs, and availability of numerous tools and resources for both genome-wide and small-scale experiments. Here we discuss critical factors for designing a yeast system for the expression of bacterial type III effector proteins. These include an appropriate promoter for driving expression of the effector gene(s) of interest, the copy number of the effector gene, the epitope tag used to verify protein expression, and the yeast strain. We present procedures to induce expression of effectors in yeast and to verify their expression by immunoblotting. In addition, we describe a spotting assay on agar plates for the identification of effector-induced growth inhibition phenotypes. The use of this protocol may be extended to the study of pathogenicity factors delivered into the host cell by any pathogen and translocation mechanism.
Microbiology, Issue 37, type III secretion system, type III effector proteins, Gram-negative bacteria, Saccharomyces cerevisiae, yeast expression system
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Budding Yeast Protein Extraction and Purification for the Study of Function, Interactions, and Post-translational Modifications
Authors: Eva Paige Szymanski, Oliver Kerscher.
Institutions: The College of William & Mary.
Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously hard to disrupt. Here we describe the use of a bead mill homogenizer for the extraction of proteins into buffers optimized to maintain the functions, interactions and post-translational modifications of proteins. Logarithmically growing cells expressing the protein of interest are grown in a liquid growth media of choice. The growth media may be supplemented with reagents to induce protein expression from inducible promoters (e.g. galactose), synchronize cell cycle stage (e.g. nocodazole), or inhibit proteasome function (e.g. MG132). Cells are then pelleted and resuspended in a suitable buffer containing protease and/or phosphatase inhibitors and are either processed immediately or frozen in liquid nitrogen for later use. Homogenization is accomplished by six cycles of 20 sec bead-beating (5.5 m/sec), each followed by one minute incubation on ice. The resulting homogenate is cleared by centrifugation and small particulates can be removed by filtration. The resulting cleared whole cell extract (WCE) is precipitated using 20% TCA for direct analysis of total proteins by SDS-PAGE followed by Western blotting. Extracts are also suitable for affinity purification of specific proteins, the detection of post-translational modifications, or the analysis of co-purifying proteins. As is the case for most protein purification protocols, some enzymes and proteins may require unique conditions or buffer compositions for their purification and others may be unstable or insoluble under the conditions stated. In the latter case, the protocol presented may provide a useful starting point to empirically determine the best bead-beating strategy for protein extraction and purification. We show the extraction and purification of an epitope-tagged SUMO E3 ligase, Siz1, a cell cycle regulated protein that becomes both sumoylated and phosphorylated, as well as a SUMO-targeted ubiquitin ligase subunit, Slx5.
Basic Protocol, Issue 80, Life Sciences (General), budding yeast, protein extracts, bead beating, sumo, Ubiquitin, post-translational modifications, 6xHis affinity tag
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High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy
Authors: Titiwat Sungkaworn, Finn Rieken, Martin J. Lohse, Davide Calebiro.
Institutions: University of Würzburg, Germany.
Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., β2-adrenergic and γ-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.
Bioengineering, Issue 89, pharmacology, microscopy, receptor, live-cell imaging, single-molecule, total internal reflection fluorescence, tracking, dimerization, protein-protein interactions
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Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy
Authors: Nikki M. Curthoys, Michael J. Mlodzianoski, Dahan Kim, Samuel T. Hess.
Institutions: University of Maine.
Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.
Basic Protocol, Issue 82, Microscopy, Super-resolution imaging, Multicolor, single molecule, FPALM, Localization microscopy, fluorescent proteins
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Test Samples for Optimizing STORM Super-Resolution Microscopy
Authors: Daniel J. Metcalf, Rebecca Edwards, Neelam Kumarswami, Alex E. Knight.
Institutions: National Physical Laboratory.
STORM is a recently developed super-resolution microscopy technique with up to 10 times better resolution than standard fluorescence microscopy techniques. However, as the image is acquired in a very different way than normal, by building up an image molecule-by-molecule, there are some significant challenges for users in trying to optimize their image acquisition. In order to aid this process and gain more insight into how STORM works we present the preparation of 3 test samples and the methodology of acquiring and processing STORM super-resolution images with typical resolutions of between 30-50 nm. By combining the test samples with the use of the freely available rainSTORM processing software it is possible to obtain a great deal of information about image quality and resolution. Using these metrics it is then possible to optimize the imaging procedure from the optics, to sample preparation, dye choice, buffer conditions, and image acquisition settings. We also show examples of some common problems that result in poor image quality, such as lateral drift, where the sample moves during image acquisition and density related problems resulting in the 'mislocalization' phenomenon.
Molecular Biology, Issue 79, Genetics, Bioengineering, Biomedical Engineering, Biophysics, Basic Protocols, HeLa Cells, Actin Cytoskeleton, Coated Vesicles, Receptor, Epidermal Growth Factor, Actins, Fluorescence, Endocytosis, Microscopy, STORM, super-resolution microscopy, nanoscopy, cell biology, fluorescence microscopy, test samples, resolution, actin filaments, fiducial markers, epidermal growth factor, cell, imaging
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In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy
Authors: Michael Stoneman, Deo Singh, Valerica Raicu.
Institutions: University of Wisconsin - Milwaukee, University of Wisconsin - Milwaukee.
The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. Förster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 α-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.
Cellular Biology, Issue 47, Forster (Fluorescence) Resonance Energy Transfer (FRET), protein-protein interactions, protein complex, in vivo determinations, spectral resolution, two-photon microscopy, G protein-coupled receptor (GPCR), sterile 2 alpha-factor protein (Ste2p)
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From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope
Authors: Carmine Di Rienzo, Enrico Gratton, Fabio Beltram, Francesco Cardarelli.
Institutions: Scuola Normale Superiore, Instituto Italiano di Tecnologia, University of California, Irvine.
It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn’t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on µm-sized membrane regions in the micro-to-milli-second time range.
Bioengineering, Issue 92, fluorescence, protein dynamics, lipid dynamics, membrane heterogeneity, transient confinement, single molecule, GFP
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Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)
Authors: Lynne Turnbull, Michael P. Strauss, Andrew T. F. Liew, Leigh G. Monahan, Cynthia B. Whitchurch, Elizabeth J. Harry.
Institutions: University of Technology, Sydney.
Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques – stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 µm from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.
Molecular Biology, Issue 91, super-resolution microscopy, fluorescence microscopy, OMX, 3D-SIM, Blaze, cell division, bacteria, Bacillus subtilis, Staphylococcus aureus, FtsZ, Z ring constriction
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Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging
Authors: Christopher R. Provost, Luo Sun.
Institutions: New England Biolabs.
SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.
Cellular Biology, Issue 39, fluorescence, labeling, imaging, SNAP-tag, tag, microscopy, AGT, surface, intracellular, fusion
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A Rapid Technique for the Visualization of Live Immobilized Yeast Cells
Authors: Karl Zawadzki, James Broach.
Institutions: Princeton University.
We present here a simple, rapid, and extremely flexible technique for the immobilization and visualization of growing yeast cells by epifluorescence microscopy. The technique is equally suited for visualization of static yeast populations, or time courses experiments up to ten hours in length. My microscopy investigates epigenetic inheritance at the silent mating loci in S. cerevisiae. There are two silent mating loci, HML and HMR, which are normally not expressed as they are packaged in heterochromatin. In the sir1 mutant background silencing is weakened such that each locus can either be in the expressed or silenced epigenetic state, so in the population as a whole there is a mix of cells of different epigenetic states for both HML and HMR. My microscopy demonstrated that there is no relationship between the epigenetic state of HML and HMR in an individual cell. sir1 cells stochastically switch epigenetic states, establishing silencing at a previously expressed locus or expressing a previously silenced locus. My time course microscopy tracked individual sir1 cells and their offspring to score the frequency of each of the four possible epigenetic switches, and thus the stability of each of the epigenetic states in sir1 cells. See also Xu et al., Mol. Cell 2006.
Microbiology, Issue 1, yeast, HML, HMR, epigenetic, loci, silencing, cerevisiae
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