We would like to acknowledge the advice and guidance from Dr. Vytas Bindokas at the University of Chicago. We thank the Integrated Light Microscopy Core Facility at the University of Chicago for supporting and maintaining the lattice light-sheet microscope. This work was supported by NIH New Innovator Award 1DP2AI144245 and NSF Career Award 1653782 (To J.H.). J.R. is supported by the NSF Graduate Research Fellowships Program.

5C.C7 TCR-transgenic RAG2 knockout mice in B10.A background were used in this study according to a protocol approved by the Institutional Animal Care and Use Committee of the University of Chicago.
1. Harvest and Activate T Cells
NOTE: This part of the protocol is based on previous protocols. See citations for further detail20,21.
<ol>
	<li>Euthanize a 10-12 week-old 5C.C7 transgenic mouse of either sex (~...

<ol>
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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PD-L1+Binds+to+B7-1+Only+In+Cis+on+the+Same+Cell+Surface.">PD-L1 Binds to B7-1 Only In Cis on the Same Cell Surface.</a> Cancer Immunology Research. 6 (8), 921-929 (2018).</li><li>Forbes, C. A., Scalzo, A. A., Degli-Esposti, M. A., Coudert, J. 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(116), e54596 (2016).</li><li>Huang, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Single+Peptide-Major+Histocompatibility+Complex+Ligand+Triggers+Digital+Cytokine+Secretion+in+CD4++T+Cells.">A Single Peptide-Major Histocompatibility Complex Ligand Triggers Digital Cytokine Secretion in CD4+ T Cells.</a> Immunity. 39 (5), 846-857 (2013).</li><li>Irvine, D. J., Purbhoo, M. A., Krogsgaard, M., Davis, M. 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The presented protocol was optimized for the usage of CD4+ T cells isolated from 5C.C7 transgenic mice on the LLSM instrument used, and therefore other cell systems and LLSMs may need to be optimized differently. However, this protocol shows the power of 4D imaging, as it can be used to quantify the dynamics of a surface receptor on an entire cell with the least distortion in physiological conditions. Therefore, there are many possible future applications of this technique.
A critic...

The precise dynamics of molecules trafficking and diffusing on the three-dimensional cell surface in real time have been an enigma to solve. Microscopy has always been a balance of speed, sensitivity, and resolution; if any one or two are maximized, the third is minimized. Therefore, due to the small size and immense speed with which surface receptors move, tracking their dynamics has remained a major technological challenge to the field of cell biology. For example, many studies have been conducted using total internal reflection fluorescence (TIRF) microscopy1,2,3, which has high temporal resolution, but can only image a very thin slice of the T-cell membrane (~100 nm), and therefore misses events happening farther away in the cell. These TIRF images also only showing a two-dimensional section of the cell. By contrast, super-resolution techniques, such as stochastic optical reconstruction microscopy (STORM)4, photoactivated localization microscopy (PALM)5, and stimulated emission depletion microscopy (STED)6, can overcome the Abbe diffraction limit of light. These techniques have high spatial resolution (~20 nm resolution)4,5,6,7, but they often take many minutes to acquire a full two-dimensional (2D) or three-dimensional (3D) image, and therefore the temporal resolution is lost. In addition, techniques such as STORM and PALM that rely on blinking signals may have inaccuracies in counting8,9. Electron microscopy has by far the highest resolution (up to 50 pm resolution)10; it can even be conducted three-dimensionally with focused ion beam scanning electron microscopy (FIB-SEM), resulting in up to 3 nm XY and 500 nm Z resolution11. However, any form of electron microscopy requires harsh sample preparation and can only be conducted with fixed cells or tissues, eliminating the possibility of imaging live samples over time.
Techniques to obtain the high spatiotemporal resolution required to identify the dynamics of surface and intracellular molecules in live cells in their true physiological 3D nature is only being recently developed. One of these techniques is Lattice Light-Sheet Microscopy (LLSM)12, which utilizes a structured light sheet to drastically lower photobleaching. Developed in 2014 by Nobel Laureate Eric Betzig, the high axial resolution, low photobleaching and background noise, and ability to simultaneously image hundreds of planes per field of view make LLS microscopes superior to widefield, TIRF and confocal microscopes12,13,14,15,16,17,18,19. This four-dimensional (x, y, z and time) imaging technique, while still diffraction limited (~200 nm XYZ resolution), has incredible temporal resolution (we have achieved a frame rate of about 100 fps, resulting in a 3D reconstructed cell image with 0.85 seconds per frame) for 3D spatial acquisition.
LLSM can be generally used to track real-time dynamics of any molecules within any cell at the single-molecule and single-cell level, particularly those in highly motile cells such as immune cells. For example, we show here how to use LLSM to visualize T-cell receptor (TCR) dynamics. T cells are the effector cells of the adaptive immune system. TCRs are responsible for recognizing peptide-MHC (pMHC) ligands displayed on the surface of antigen-presenting cells (APC), which determines the selection, development, differentiation, fate, and function of a T cell. This recognition occurs at the interface between T cells and APCs, resulting in localized receptor clustering to form what is called the immunological synapse. While it is known that TCRs at the immunological synapse are imperative for T-cell effector function, still unknown are the underlying mechanisms of real-time TCR trafficking to the synapse. LLSM has allowed us to visualize in real time the dynamics of TCRs before and after trafficking to the synapse with the resultant pMHC-TCR interaction (Figure 1). LLSM can therefore be used to solve current questions of the formative dynamics of TCRs and provide insights to understand how a cell distinguishes between self and foreign antigens.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL Syringe</td>
			<td>BD</td>
			<td>309659</td>
			<td>For T cell harvest</td>
		</tr>
		<tr>
			<td>2-Mercaptoethanol</td>
			<td>Sigma-Aldrich</td>
			<td>M3148-25ML</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>5 mm round coverslips</td>
			<td>World Precision Instruments</td>
			<td>502040</td>
			<td>For Imaging</td>
		</tr>
		<tr>
			<td>70um Sterile Cell Strainer</td>
			<td>Corning</td>
			<td>7201431</td>
			<td>For T cell harvest</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 anti-mouse TCR &#946; chain Antibody</td>
			<td>BioLegend</td>
			<td>109215</td>
			<td>For Imaging</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>X&#38;Y Cell Culture</td>
			<td>FBS-500</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>Ficoll</td>
			<td>GE Healthcare</td>
			<td>17-1440-02</td>
			<td>Denisty gradient reagent for T cell harvest</td>
		</tr>
		<tr>
			<td>Fluorescein sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>F6377</td>
			<td>For microscope alignment</td>
		</tr>
		<tr>
			<td>FluoSpheres Carboxylate-Modified Microspheres</td>
			<td>Thermo Fisher Scientific</td>
			<td>F8810</td>
			<td>For microscope alignment</td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Bitplane</td>
			<td>N/A</td>
			<td>Tracking Software; Other options for tracking software include Amira or Trackmate (Fiji).</td>
		</tr>
		<tr>
			<td>Lattice Light-Sheet Microscope</td>
			<td>3i</td>
			<td>N/A</td>
			<td>Microscope Used</td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium, no phenol red</td>
			<td>Thermo Fisher Scientific</td>
			<td>21083027</td>
			<td>For Imaging</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Thermo Fisher Scientific</td>
			<td>25030-081</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>LLSpy</td>
			<td>Janelia Research Campus</td>
			<td>N/A</td>
			<td>LLSpy was used under license from Howard Hughes Medical Institute, Janelia Research Campus. Contact innovation@janelia.hhmi.org for access. Other deconvolution and deksewing methods are available in image processing softwares such as Fiji, Slidebook, Amira, and others. <a href="https://llspy.readthedocs.io/en/latest/" target="_blank">https://llspy.readthedocs.io/en/latest/</a></td>
		</tr>
		<tr>
			<td>Moth Cytochrome C (MCC), sequence ANERADLIAYLKQATK</td>
			<td>Elimbio</td>
			<td>Custom Synthesis</td>
			<td>For T cell harvest</td>
		</tr>
		<tr>
			<td>Penacillin/Streptamycin</td>
			<td>Life Technologies</td>
			<td>15140122_3683884612</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>Poly-L-Lysine</td>
			<td>Phenix Research Products</td>
			<td>P8920-100ML</td>
			<td>For Imaging</td>
		</tr>
		<tr>
			<td>RBC Lysis Buffer</td>
			<td>eBioscience</td>
			<td>00-4300-54</td>
			<td>For T cell harvest</td>
		</tr>
		<tr>
			<td>Recombinant mouse IL-2</td>
			<td>Sigma-Aldrich</td>
			<td>I0523</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>RPMI 1640 Medium</td>
			<td>Corning</td>
			<td>MT10040CV</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>Slidebook</td>
			<td>3i</td>
			<td>N/A</td>
			<td>LLSM imaging software</td>
		</tr>
		<tr>
			<td>Surgical Dissection Tools</td>
			<td>Nova-Tech International</td>
			<td>DSET10</td>
			<td>For T cell harvest</td>
		</tr>
		<tr>
			<td>T-25 Flasks</td>
			<td>Eppendorf</td>
			<td>2231710126</td>
			<td>For T cell culture</td>
		</tr>
		<tr>
			<td>Thermo Scientific Pierce Fab Micro Preparation Kits</td>
			<td>Thermo Fisher Scientific</td>
			<td>44685</td>
			<td>For preparing Fab</td>
		</tr>
	</tbody>
</table>

visualizing surface t-cell receptor dynamics four-dimensionally using lattice light-sheet microscopy

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the structure-function relationship of these receptors in situ, we need to visualize and track them on the live cell surface with enough spatiotemporal resolution. Here we show how to use recently developed Lattice Light-Sheet Microscopy (LLSM) to image T-cell receptors (TCRs) four-dimensionally (4D, space and time) at the live cell membrane. T cells are one of the main effector cells of the adaptive immune system, and here we used T cells as an example to show that the signaling and function of these cells are driven by the dynamics and interactions of the TCRs. LLSM allows for 4D imaging with unprecedented spatiotemporal resolution. This microscopy technique therefore can be generally applied to a wide array of surface or intracellular molecules of different cells in biology.

Here, we describe the isolation, preparation, and imaging of primary mouse 5C.C7 T cells using a lattice light-sheet microscope. During section 3, it is imperative to align the microscope correctly, and to collect PSF daily with which to deconvolve the data after collection. In Figure 2, we show the correct alignment images that will be seen when aligning the microscope. Figure 2A and Figure 2B show...

Watch this Scientific Journal Video about Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy at JoVE.com

Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy

The goal of this protocol is to show how to use Lattice Light-Sheet Microscopy to four-dimensionally visualize surface receptor dynamics in live cells. Here T cell receptors on CD4+ primary T cells are shown.

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the ...

Lattice light-sheet microscopy is a powerful imaging technology, but few labs in the world are capable of using it. This protocol gives a detailed overview of how to use the commercially available lattice light-sheet microscope. It's an extremely powerful system capable of four-dimensional imaging of individual cells or embryos that allows for real-time tracking of molecules.The main advantage of this technique is that it can three-dimensionally image single live cells or organs with high spatiotemporal resolution and minimal photo bleaching. The lattice pattern of the light sheet allows us to high-speed image samples and obtain real-time videos of three-dimensional living cells and organs with molecular details that other techniques cannot achieve. This first video protocol showing how to us lattice light-sheet microscopy to image single cell four dimensionally.This is a very complicated technique due to alignment and sample preparation. And we believe that visual demonstration will improve accessibility by allowing more scientists to image many different molecules, cells, and organs in biology. To begin this procedure, incubate five millimeter round coverslips with 0.1%poly-L-lysine at room temperature for 10 minutes.Aspirate off the liquid and let the coverslips air dry naturally. Next, add three milliliters of density gradient to a 15 milliliter conical tube. Carefully add cells drop-wise to the edge of the tube.Centrifuge at 930 times g and at four degrees Celsius for 10 minutes. After this, carefully remove the thin middle layer of cells between the complete media and the density gradient reagent making sure to put each cell type into a separate conical tube. Centrifuge at 300 times g for five minutes to wash the cells.Discard the supernatant and add five milliliters of RPMI to the tubes of T-cells and CH27 cells. Repeat the wash with fresh RPMI until three total washes have been performed. Then, resuspend the cells in each tube with one milliliter of complete RPMI and count the cells with a hemocytometer.Resuspend one million antigen-presenting cells in 500 microliters of complete RPMI and add 10 micromolar MCC. Incubate at 37 degrees Celsius with 5%carbon dioxide for three hours. After this, resuspend one million T-cells in 500 microliters of complete RPMI.Add two micrograms of anti-TCR beta Alexa 488 labeled FAB and incubate at 37 degrees Celsius with 5%carbon dioxide for 30 minutes. Next, centrifuge both cells at 300 times g for five minutes and discard the supernatant. Add 500 microliters of complete RPMI and repeat the centrifuge to wash the cells.Repeat the wash with fresh RPMI until three total washes have been performed discarding the supernatant after each wash. After this, resuspend both cell types in 500 microliters of imaging media. First, add 10 milliliters of water and 30 microliters of fluorescein to the LLSM bath.Press Image Home to move the object to image position and look at a single Bessel laser beam pattern. Align the laser beam using the guide and preset region of interest to make the beam a thin pattern that is balanced in all directions. The beam should also appear focused in the finder camera.Use the two mirror tilt adjusters, the top focus micrometer, and the objective color to adjust the beam. Next, wash the bath and the objectives with at least 200 milliliters of water to completely remove any fluorescein. To image standard fluorescent beads, turn on dither by setting to three in the Xgalvo range box and press Live to view current field.Manually adjust the tilt mirror, objective color, and focus micrometer for highest gray values. Then, adjust as necessary to obtain proper patterns for objective scan, Z galvo, Z plus objective, and sample scan capture modes. After this, press Execute in sample scan mode to collect the sample point spread function for de-skewing and de-convolution.Change the lasers to three color mode and press Execute again. To begin, add 100, 000 anti-presenting cells to the prepared five millimeter diameter circular coverslip and allow them to settle for 10 minutes. Grease the sample holder and add the coverslip to it cell side up.Next, add a drop of imaging media to the back of the coverslip. Screw the sample holder onto the Piezo and press Image Home. Find an antigen-presenting cell to image and ensure that the LLSM and imaging software are functioning properly.Press Live to view the current image. Move along Z to find the coverslip and cells. Find the center of an antigen-presenting cell by moving in the Z direction, then press Stop to pause the laser.Check 3D and input the desired settings. Press Center and then press Execute to collect the data. Lower the stage to the load position and add 50 microliters of T-cells and imaging media drop-wise directly over the coverslip.It is best to let a drop form on the end of the pipette tip and then touch the tip to the bath liquid. Raise the stage back by clicking Image Return. Begin imaging.Be sure to set the desired Z stack and timelapse length. For example, image 60 Z stacks at a 0.4 micrometer step size and input 500 timeframes. Stop recording before 500 frames are reached to avoid photo bleaching.Use live mode to search for cell pairs and when ready and desired settings have been entered, press Execute to collect data. In this study, primary mouse 5CC7 T-cells are isolated and prepared and are then imaged using a lattice light-sheet microscope. Shown here are the correct beam path and beam alignment when imaged with fluorescein.The objective scan should show a large X shape in the XZ and YZ projections that is as symmetrical as possible. They should also be adjusted to be as small of an X as possible. The Z galvo scan should show an oval in XZ and XY with a single dot on either side.Finally, both the Z plus objective scan and the sample scan should show dots that look as round as possible. Using this protocol, the four-dimensional dynamics of the T-cell receptors on a T-cell surface could be seen. The main advantage of this microscope lies in the ability to track the visualized surface units of the T-cell receptors and to obtain quantified data from their size, motion, signal intensity, and excreta.The most important thing to remember is the quality of your alignment. The lattice light sheet must be aligned daily and not doing this properly will result in distorted imaging. Similarly, the quality of the PSF collected directly impacts the quality of the de-convolution which ultimately results in the quality of your final image.One of the reasons this method is so powerful is because each cell and label can be replaced to visualize many cell types and molecules. Therefore, this method can be used to answer any question related to four-dimensional tracking of molecules. We think that this technique will pave the way for researchers to explore new questions within the field of molecular trafficking.It is not yet being widely used due to the complicated system so we're hoping this Jove video will improve accessibility to the technique. The lasers used on the system are class four so absolute caution must be taken to ensure laser safety. Please take necessary laser safety training prior to using this method.

visualizing-surface-t-cell-receptor-dynamics-four-dimensionally-using

Research

JoVE Journal

Biochemistry

7.8K ビュー.  The University of Chicago. ラティスライトシート顕微鏡は強力なイメージング技術ですが、それを使用できるラボは世界ではほとんどありません。このプロトコルは、市販の格子光シート顕微鏡の使用方法の詳細な概要を提供します。これは、分子のリアルタイム追跡を可能にする個々の細胞または胚の4次元イメージングが可能な非常に強力なシステムです。この技術の主な利点は、高い時...

ビデオ: Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy

Extracellular Protein Microarray Technology for High Throughput Detection of Low Affinity Receptor-Ligand Interactions

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling cascades during homeostasis and disease, and as such represent prime therapeutic targets. Despite their relevance, these interaction networks remain significantly underrepresented in current databases; therefore, most extracellular proteins have no documented binding partner. This discrepancy is primarily due to the challenges associated with the study of the extracellular proteins, including expression of functional proteins, and the weak, low affinity, protein interactions often established between cell surface receptors. The purpose of this method is to describe the printing of a library of extracellular proteins in a microarray format for screening of protein-protein interactions. To enable detection of weak interactions, a method based on multimerization of the query protein under study is described. Coupled to this microbead-based multimerization approach for increased multivalency, the protein microarray allows robust detection of transient protein-protein interactions in high throughput. This method offers a rapid and low sample consuming-approach for identification of new interactions applicable to any extracellular protein. Protein microarray printing and screening protocol are described. This technology will be useful for investigators seeking a robust method for discovery of protein interactions in the extracellular space.

Here, we present a protocol to screen extracellular protein microarrays for identification of novel receptor-ligand interactions in high throughput. We also describe a method to enhance detection of transient protein-protein interactions by using protein-microbead complexes.

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Screening for Phytoestrogens using a Cell-based Estrogen Receptor  &#946; Reporter Assay

Plants are a source of food for many animals, and&#160;they can produce thousands of chemicals. Some of these compounds affect physiological processes in the vertebrates that consume them, such as endocrine function. Phytoestrogens, the most well studied endocrine-active phytochemicals, directly interact with the hypothalamo-pituitary gonadal axis of the vertebrate endocrine system. Here we present the novel use of a cell-based assay to screen plant extracts for the presence of compounds that have estrogenic biological activity. This assay uses mammalian cells engineered to highly express estrogen receptor beta (ER&#946;) and that have been transfected with a luciferase gene. Exposure to compounds with estrogenic activity results in the cells producing light. This assay is a reliable and simple way to test for biological estrogenic activity. It has several improvements over transient transfection assays, most notably, ease of use, the stability of the cells, and the sensitivity of the assay.

Screening for Phytoestrogens using a Cell-based Estrogen Receptor  &amp;#946; Reporter Assay

We have optimized a commercially available estrogen receptor &#946; reporter assay for screening human and nonhuman primate foods for estrogenic activity. We validated this assay by showing that the known estrogenic human food soy registers high, while other foods show no activity.

Plants are a source of food for many animals, and&#160;they can produce thousands of chemicals. Some of these compounds affect physiological processes in ...

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using &#967;CRAC

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be bound by many different RBPs that will regulate its production, stability, activity, and degradation. As such, much has been done to understand the dynamics that exist between these two types of molecules. A particularly important breakthrough came with the emergence of &#8216;cross-linking and immunoprecipitation&#8217; (CLIP). This technique allowed stringent investigation into which RNAs are bound by a particular RBP. In short, the protein of interest is UV cross-linked to its RNA substrates in vivo, purified under highly stringent conditions, and then the RNAs covalently cross-linked to the protein are converted into cDNA libraries and sequenced. Since its conception, many derivative techniques have been developed in order to make CLIP amenable to particular fields of study. However, cross-linking using ultraviolet light is notoriously inefficient. This results in extended exposure times that make the temporal study of RBP-RNA interactions impossible. To overcome this issue, we recently designed and built much-improved UV irradiation and cell harvesting devices. Using these new tools, we developed a protocol for time-resolved analyses of RBP-RNA interactions in living cells at high temporal resolution: Kinetic CRoss-linking and Analysis of cDNAs (&#967;CRAC). We recently used this technique to study the role of yeast RBPs in nutrient stress adaptation. This manuscript provides a detailed overview of the &#967;CRAC method and presents recent results obtained with the Nrd1 RBP.

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using &amp;#967;CRAC

Kinetic cross-linking and analysis of cDNA is a method that allows investigation of the dynamics of protein-RNA interactions in living cells at high temporal resolution. Here the protocol is described in detail, including the growth of yeast cells, UV cross-linking, harvesting, protein purification, and next generation sequencing library preparation steps.

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be ...

Visualizing Diffusional Dynamics of Gold Nanorods on Cell Membrane using Single Nanoparticle Darkfield Microscopy

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and provides a theoretical basis for the rational design of nano-medicine delivery. Single particle tracking (SPT) analysis could probe the position and orientation of individual nanoparticles on cell membrane, and reveal their translational and rotational states. Here, we show how to use traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on live cell membrane. We also show how to extract the location and orientation of AuNRs using ImageJ and MATLAB, and how to characterize the diffusive states of AuNRs. Statistical analysis of hundreds of particles show that single AuNRs perform Brownian motion on the surface of U87 MG cell membrane. However, individual long trajectory analysis shows that AuNRs have two distinctly different types of motion states on the membrane, namely long-range transport and limited-area confinement. Our SPT methods can be potentially used to study the surface or intracellular particle diffusion in different biological cells and can become a powerful tool for investigations of complex cellular mechanisms.

Here, we show the use of traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on cell membrane. The location and orientation of single AuNRs are detected using ImageJ and MATLAB, and the diffusive states of AuNRs are characterized by single particle tracking analysis.

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and ...

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This enables quantifications of ultrastructural aspects such as volume fractions, surface area to volume ratios, morphometry, and physical contact sites of different subcellular structures. In the 1970s, a protocol for enhanced staining of glycogen in cells was developed and paved the way for a string of studies on the subcellular localization of glycogen and glycogen particle size using transmission electron microscopy. While most analyses interpret glycogen as if it is homogeneously distributed within the muscle fibers, providing only a single value (e.g., an average concentration), transmission electron microscopy has revealed that glycogen is stored as discrete glycogen particles located in distinct subcellular compartments. Here, the step-by-step protocol from tissue collection to the quantitative determination of the volume fraction and particle diameter of glycogen in the distinct subcellular compartments of individual skeletal muscle fibers is described. Considerations on how to 1) collect and stain tissue specimens, 2) perform image analyses and data handling, 3) evaluate the precision of estimates, 4) discriminate between muscle fiber types, and 5) methodological pitfalls and limitations are included.

A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This ...

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...