Name: visualizing adhesion formation in cells by means of advanced spinning disk-total internal reflection fluorescence microscopy
Uploaded: 2019-01-21T14:00:00
Description: Watch this Scientific Journal Video about Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy at JoVE.com

We greatly thank the scientific community of the University Medical Center Hamburg-Eppendorf for supporting us with samples for evaluation. Namely, we thank Sabine Windhorst for NIH3T3 cells, Andrea Mordhorst for YFP-Vinculin and Maren Rudolph for RFP-Lifeact.

1. Preparation of cells
<ol>
	<li>Two days prior to the experiment, seed 3*105 HeLa or NIH3T3 cells in 2 mL of full growth medium per well of a 6-well cell-culture plate. Ensure that cells are handled in a laminar flow hood throughout this protocol.</li>
	<li>One day prior to the experiment, prepare the transfection reagents according to the manufacturer&#8217;s recommendations or an empirically determined protocol, e.g.:
	<ol>
		<li>Dilute 1 &#181;g of RFP-Lifeact and 1 &#181;g of YFP-Vinculin in a...

<ol>
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E., Unser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+pyramid+approach+to+subpixel+registration+based+on+intensity.">A pyramid approach to subpixel registration based on intensity.</a> IEEE Transactions on Image Processing. 7 (1), 27-41 (1998).</li><li>Partridge, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initiation+of+attachment+and+generation+of+mature+focal+adhesions+by+integrin-containing+filopodia+in+cell+spreading.">Initiation of attachment and generation of mature focal adhesions by integrin-containing filopodia in cell spreading.</a> Molecular Biology of the Cell. 17 (10), (2006).</li><li>Dubin-Thaler, B. J., Giannone, G., D&#246;bereiner, H. G., Sheetz, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanometer+analysis+of+cell+spreading+on+matrix-coated+surfaces+reveals+two+distinct+cell+states+and+STEPs.">Nanometer analysis of cell spreading on matrix-coated surfaces reveals two distinct cell states and STEPs.</a> Biophysical Journal. 86 (3), 1794-1806 (2004).</li><li>Choi, C. K., Vicente-Manzanares, M., Zareno, J., Whitmore, L. A., Mogilner, A., Horwitz, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin+and+&#945;-actinin+orchestrate+the+assembly+and+maturation+of+nascent+adhesions+in+a+myosin+II+motor-independent+manner.">Actin and &#945;-actinin orchestrate the assembly and maturation of nascent adhesions in a myosin II motor-independent manner.</a> Nature Cell Biology. 10 (9), 1039-1050 (2008).</li><li>Bachir, A. I., Zareno, J., Moissoglu, K., Plow, E. F., Gratton, E., Horwitz, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrin-associated+complexes+form+hierarchically+with+variable+stoichiometry+in+nascent+adhesions.">Integrin-associated complexes form hierarchically with variable stoichiometry in nascent adhesions.</a> Current Biology. 24 (16), 1845-1853 (2014).</li><li>Sun, Z., Guo, S. S., F&#228;ssler, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrin-mediated+mechanotransduction.">Integrin-mediated mechanotransduction.</a> Journal of Cell Biology. 215 (4), 445-456 (2016).</li><li>Shao, L., Kner, P., Rego, E. H., Gustafsson, M. G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super-resolution+3D+microscopy+of+live+whole+cells+using+structured+illumination.">Super-resolution 3D microscopy of live whole cells using structured illumination.</a> Nat. Methods. 8 (12), 1044-1046 (2011).</li><li>Chen, B. -. C., 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=Lattice+light-sheet+microscopy:+Imaging+molecules+to+embryos+at+high+spatiotemporal+resolution.">Lattice light-sheet microscopy: Imaging molecules to embryos at high spatiotemporal resolution.</a> Science. 346 (6208), 1257998-1257998 (2014).</li><li>Fu, Y., Winter, P. W., Rojas, R., Wang, V., McAuliffe, M., Patterson, G. 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In this paper was presented the first successful implementation of SD and TIRF microscopy in a configuration suitable for performing live cell imaging experiments, i.e. high acquisition rates such as 2 SD-TIRF image stacks per minute at 3 different stage positions, corresponding to a total of 168 frames (circa 3 frames per second), were acquired. The few SD-TIRF microscopes that were described previously12,13, mainly lack of sufficiently high imaging spe...

Our days, light microscopy techniques providing high/super resolution imaging in fixed and living specimen are developing rapidly. Super-resolution techniques such as stimulated emission depletion (STED), structured illumination microscopy (SIM) and photo-activation localization microscopy (PALM) or direct stochastic optical reconstruction microscopy (STORM), respectively, are commercially available and enable imaging of subcellular structures showing details almost on the molecular scale1,2,3,4,5,6. However, these approaches still have limited applicability for live imaging experiments in which large volumes need to be visualized with multiple frames per second acquisition speed. Varieties of highly dynamic processes regulated via the plasma membrane, e.g. endo-/exocytosis, adhesion, migration or signaling, occur with high speed within large cellular volumes. Recently, in order to fill up this gap, an integrated microscopy technique was proposed called spinning disk-TIRF (SD-TIRF)7. In detail, TIRF microscopy permits to specifically isolate and localize the plasma membrane8,9, while SD microscopy is one of the most sensitive and fast live imaging techniques for the visualization and tracking of subcellular organelles in the cytoplasm10,11. The combination of both imaging techniques in a single setup has already been realized in the past12,13, however, the microscope presented here (Figure 1) finally meets the criteria to perform live imaging SD-TIRF experiments of the aforementioned processes at 3 frames per second speed. Since this microscope is commercially available, the goal of this manuscript is to describe in details and provide open source tools and protocols for image acquisition, registration, and visualization associated with SD-TIRF microscopy.
The setup is based on an inverted microscope connected to two scan units via independent ports - the left port is linked to the SD unit and the back port to scanner unit for TIRF and photo-activation/-bleaching experiments. Up to 6 lasers (405/445/488/515/561/640 nm) can be used for excitation. For excitation and detection of the fluorescence signal, either a 100x/NA1.45 oil or 60x/NA1.49 oil TIRF objective, respectively, have been employed. The emitted light is split by a dichroic mirror (561 nm long-pass or 514 nm long-pass) and filtered by various band-pass filters (55 nm wide centered at 525 nm, 54 nm wide centered at 609 nm for green and red fluorescence, respectively) placed in front of the two EM-CCD cameras. Please note that more technical details about the setup are listed in Zobiak et al.7. In TIRF configuration, the SD unit is moved out of the light path within circa 0.5 s so that the same two cameras can be used for detection, allowing faster switching between the two imaging modalities compared to circa 1 s that was reported in the past13. This feature enables dual-channel simultaneous acquisition, thus 4 channels SD-TIRF imaging at previously unmatched speed and accuracy can be performed. Moreover, alignment between SD and TIRF images is unnecessary. Image alignment between the two cameras, however, has to be checked before starting the experiment and corrected if necessary. In the following protocol, a registration correction routine was implemented in a self-written ImageJ macro. Moreover, the macro was mainly designed to allow a simultaneous visualization of SD- and TIRF datasets despite their different dimensionality. The acquisition software itself did not provide these features.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Microscope and accessories</td>
			<td ></td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >SD-TIRF microscope</td>
			<td >Visitron Systems</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Ti with perfect focus system</td>
			<td >Nikon</td>
			<td ></td>
			<td>Inverted microscope stand</td>
		</tr>
		<tr >
			<td >CSU-W1 T2</td>
			<td >Yokogawa</td>
			<td ></td>
			<td>Spinning disk unit in dual-camera configuration</td>
		</tr>
		<tr >
			<td >iLAS2&#160;</td>
			<td >Roper Scientific</td>
			<td ></td>
			<td>TIRF/FRAP scanner</td>
		</tr>
		<tr >
			<td >Evolve&#160;</td>
			<td >Photometrix</td>
			<td ></td>
			<td>EM-CCD cameras</td>
		</tr>
		<tr >
			<td >PiezoZ stage</td>
			<td >Ludl Electronic Products</td>
			<td ></td>
			<td>Motorized Z stage</td>
		</tr>
		<tr >
			<td >Bioprecision2 XY stage</td>
			<td >Ludl Electronic Products</td>
			<td ></td>
			<td>Motorized XY stage</td>
		</tr>
		<tr >
			<td >Stage top incubation chamber</td>
			<td >Okolab</td>
			<td >Bold Line</td>
			<td>Temperature, CO2 and humidity supply</td>
		</tr>
		<tr >
			<td >Cell culture</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >HeLa cervical cancer cells</td>
			<td >DSMZ</td>
			<td >ACC-57</td>
			<td></td>
		</tr>
		<tr >
			<td >NIH3T3 fibroblasts</td>
			<td >DSMZ</td>
			<td >ACC-59</td>
			<td></td>
		</tr>
		<tr >
			<td >Dulbecco&#39;s phosphate buffered saline (PBS)</td>
			<td >Gibco</td>
			<td >14190144</td>
			<td></td>
		</tr>
		<tr >
			<td >Trypsin-EDTA 0.05%</td>
			<td >Gibco</td>
			<td >25300054</td>
			<td></td>
		</tr>
		<tr >
			<td >Dulbecco&#39;s Modified Eagle Medium + GlutaMAX-I (DMEM)</td>
			<td >Gibco</td>
			<td >31966-021</td>
			<td></td>
		</tr>
		<tr >
			<td >OptiMEM</td>
			<td >Gibco</td>
			<td >31985070</td>
			<td>Reduced serum medium</td>
		</tr>
		<tr >
			<td >Fetal calf serum (FCS)</td>
			<td >Gibco</td>
			<td >10500064</td>
			<td></td>
		</tr>
		<tr >
			<td >Penicillin/Streptomycin (PenStrep)</td>
			<td >Gibco</td>
			<td >15140148</td>
			<td></td>
		</tr>
		<tr >
			<td >Full growth medium (DMEM supplemented with 10% FCS and 1% PenStrep)</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >TurboFect</td>
			<td >ThermoFisher Scientific</td>
			<td >R0531</td>
			<td>Transfection reagent</td>
		</tr>
		<tr >
			<td >Ascorbic acid (AA)</td>
			<td >Sigma</td>
			<td >A544-25G</td>
			<td></td>
		</tr>
		<tr >
			<td >6-well cell culture plate</td>
			<td >Sarstedt</td>
			<td >83.392</td>
			<td></td>
		</tr>
		<tr >
			<td >Glass bottom dishes</td>
			<td >MatTek</td>
			<td >P35G-1.5-10-C</td>
			<td>35mm, 0.17mm glass coverslip</td>
		</tr>
		<tr >
			<td >Fibronectin, bovine plasma</td>
			<td >ThermoFisher Scientific</td>
			<td >33010018</td>
			<td></td>
		</tr>
		<tr >
			<td >Neubauer improved chamber</td>
			<td >VWR</td>
			<td >631-0696</td>
			<td></td>
		</tr>
		<tr >
			<td >TetraSpeck beads</td>
			<td >ThermoFisher Scientific</td>
			<td >T7279</td>
			<td></td>
		</tr>
		<tr >
			<td >Plasmids</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >RFP-Lifeact</td>
			<td ></td>
			<td ></td>
			<td>Maren Rudolph, Institute of Medical Microbiology, University Medical Center Hamburg Eppendorf, Germany</td>
		</tr>
		<tr >
			<td >YFP-Vinculin</td>
			<td ></td>
			<td ></td>
			<td>Andrea Mordhorst, Institute of Medical Microbiology, University Medical Center Hamburg Eppendorf, Germany</td>
		</tr>
		<tr >
			<td >Software and plugins</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >VisiView</td>
			<td >Visitron Systems</td>
			<td ></td>
			<td>Version 3</td>
		</tr>
		<tr >
			<td >ImageJ</td>
			<td ></td>
			<td ></td>
			<td>Version 1.52c</td>
		</tr>
		<tr >
			<td >Turboreg plugin</td>
			<td ></td>
			<td ></td>
			<td>http://bigwww.epfl.ch/thevenaz/turboreg/</td>
		</tr>
		<tr >
			<td >Macro &#34;SD-TIRF_helper_JoVE.ijm&#34;</td>
			<td >this publication</td>
			<td ></td>
			<td>https://github.com/bzobiak/ImageJ</td>
		</tr>
		<tr >
			<td >Volocity</td>
			<td >PerkinElmer</td>
			<td ></td>
			<td >Version 6.2.2</td>
		</tr>
	</tbody>
</table>

visualizing adhesion formation in cells by means of advanced spinning disk-total internal reflection fluorescence microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

In order to show the potential of SD-TIRF imaging, an assay was developed that should reveal the spatio-temporal organization of cell-matrix adhesion complexes and their interaction with the cytoskeleton during cellular adhesion. Therefore, adherent HeLa or, alternatively, NIH3T3 cells were transfected with YFP-Vinculin and RFP-Lifeact for 18-24 h, trypsinized and seeded onto fibronectin-coated glass bottom dishes. These cell lines were chosen for their pronounced cytoskeleton and higher ...

Watch this Scientific Journal Video about Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy at JoVE.com

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...

Spinning disk TIRF microscopy can be a useful tool for studying the role of protein during cytoskeletal network formation. This technique allowed three-dimensional imaging of fast cellular approaches that occur in the cell and at the same the precise localization of fluorescence molecule that are placed at the plasma membrane. This method can provide insight to research areas like microbiology, cell biology, and all those branches where it's important to study the interaction between the plasma membrane and the cellular environment.This method can be extended to those live imaging experiments where it's important to localize the structures in the plasma membrane and where you also want to maintain a high resolution of the remaining cellular volume. The crucial things one has to consider are the choice of the proper cell line to set up the TIRF illumination and the other imaging parameters and to find the focus of the transfected cells. The visual demonstration of this protocol would certainly help to avoid problems with the handling of the cells during image acquisition.Two days prior to the experiment, seed 3, 000 HELA or NIH 3T3 cells in two milliliters of full growth medium into each well of a six-well cell culture plate. The next day, prepare the transfection reagents. First, dilute one microgram of RFP Livact and one microgram of YFP Vinculin in a total of 200 microliters of reduced serum medium.Vortex the transfection reagent briefly. Then add four microliters of the mixture to 200 microliters of DNA. Vortex the reagents again and then incubate the transfection mix for 15 to 20 minutes at room temperature.After incubation, add the entire transfection mix drop-wise directly to the cells. Mix the wells by shaking the plate and then place back into the incubator. On the day of the experiment, prepare the sample for live imaging by first coating a 35 millimeter glass bottom dish with a 10 microgram per milliliter solution of Fibronectin in PBS.Leave the solution on the glass surface for 30 minutes at room temperature then remove it and let the dish air dry. Next, dilute a 0.1 micron diameter multi-fluorescent bead solution to a density of 1.8 billion particles per milliliter in distilled water. Add the solution to the Fibronectin-coated glass surface for 30 to 60 seconds.Then remove the solution and let the dish air dry. Precoating of the dishes with fluorescent beads is only necessary for two reasons, first if you want to find the TIRF plane before seeding the cells or to acquire a two-colored bead image for image registration. Now prepare an anti-oxidizing growth medium by adding 0.1 molar ascorbic acid solution at a final concentration of 0.1 millimolar in growth medium.Prewarm the media by placing it in a 37 degree Celsius water bath. Next, wash the cells with two milliliters of PBS and add 250 microliters of Trypsin EDTA to each well of the plate. Incubate the cells at 37 degrees Celsius and wait two to three minutes until the cells are fully detached.Resuspend the cells carefully in one milliliter of prewarmed anti-oxidizing growth medium and add them to an additional four milliliters of the medium in a 15 milliliter cell culture tube. Place the cell suspension with a slightly opened lid in an incubator set at 37 degrees Celsius buffered with 5%carbon dioxide. To begin, start the environmental control of the microscope to achieve stable cell culture conditions.Then place the glass bottom dish containing one milliliter of prewarmed anti-oxidizing growth medium into the sample holder of the preheated microscope. Next, in the imaging software, set up the acquisition settings at the microscope. Set the acquisition time interval to 30 seconds and the duration to between 60 and 90 minutes.Also, activate the autofocusing function of the hardware-based autofocus for every time point by setting the value to one. Now adjust the camera exposure to 200 milliseconds, the gain level to 500, and the laser power to 20%for every channel. High gain levels, low exposure time, and low laser power are recommended to reduce phototoxicity.Next, set the Z-stack for the spinning disk channels to 10 microns with 0.4 micron spacing, then set the bottom offset to zero so that the lowest plane will be the focus position of the hardware autofocus and deactivate the Z-stacks for the TIRF channels. Finally, activate the multi-point function stage positions. This will allow for up to three positions to be recorded over a 30-second interval.Now find the fluorescent beads with epifluorescent illumination, activate one TIRF channel and set the illumination angle to a value that denotes TIRF illumination. Next, activate the autofocus by pushing the button at the microscope panel and adjust the focus with the offset wheel. Once in focus, acquire a two-colored data set using TIRF 488 and TIRF 561 for subsequent bead-based image registration.Remove the cells from the incubator and mix the cell suspension by inverting the closed tube two to three times. Then apply one milliliter of the cells to the imaging dish. Quickly find double transfected cells with low level epifluorescent illumination.Once found, center the cells in the live camera preview using bright-field illumination and mark the position. Then find an additional one to two points of interest, save them to the positions list and begin data acquisition by clicking on the sequence button. In the beginning, the cells can easily detach due to the stage movement so it's better to set four to five positions and later on discard one or two.In order to generate a registration-free hyperstack in Fiji, first download and save the provided file SD-TIRF_helper_jove. ijm in the Fiji subfolder macros. Run the macro by clicking on the menu command starting with plugins then macros and finally run.If the color channels need registration correction, select the option and create a new bead-based registration reference known as a landmark file. Next, import the data with the bioformats importer and choose hyperstack as a viewing option. Load the image data set, select the spinning disk series in the first step and the TIRF series in the second step.At this point, Fiji will display the data sorted by channel and stage position. Apply the registration correction to the respective channels by loading the predetermined landmark file. Then select the desired color lookup table for every spinning disk and TIRF channel and merge them into a single multidimensional hyperstack.During the processing of the TIRF images, a number of zed planes is added to the bottom plane and this number matches with the number of the planes in the spinning disk data sets. That's very important for the visual representation of the final hyperstack. Using the setup described in this video, YFP and RFP-expressing cells were selected and their adhesion process observed during a 60-minute timelapse.As expected, cells were round shaped and only weakly adherent at the beginning with membrane protrusions sensing the environment and making contact with the substrate. Cell matrix contact strengthened quickly upon formation of so called nascent adhesions. Over the course of time, adhesion complexes enlarged and matured to focal adhesions.These elongated structures were predominantly apparent at the periphery of the cell. This resulted from forces that were exerted by actomyosin fibers which induced the strengthening of cell matrix adhesions as well as the bundling of actin fibers. The cell also flattened as a result of the actin network formation.The acquisition speed is dependent on the following parameters, the time interval, the exposure time, the number of zed planes and the number of positions. So it is very important to optimize these parameters for high acquisition rates. The implementation of the newest spinning disk technology will turn spinning disk TIRF microscopy into new super resolution microscopy technique that will allow imaging at high temporal rates.Spinning disk microscopy is a very powerful technique for studying processes that occur between the cell interior and the cell membrane. We have been shown that we can follow the dynamic of vesicle by acquiring very large image stacks within few seconds. Collimated laser light is hazardous for the eyes.Never look directly into the beam and use the safety precautions of the instrument to avoid direct light exposure.

visualizing-adhesion-formation-cells-means-advanced-spinning-disk

Research

JoVE Journal

Bioengineering

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Nanotopology of Cell Adhesion upon Variable-Angle Total Internal Reflection Fluorescence Microscopy (VA-TIRFM)

Surface topology, e.g. of cells growing on a substrate, is determined with nanometer precision by Variable-Angle Total Internal Reflection Fluorescence Microscopy (VA-TIRFM). Cells are cultivated on transparent slides and incubated with a fluorescent marker homogeneously distributed in their plasma membrane. Illumination occurs by a parallel laser beam under variable angles of total internal reflection (TIR) with different penetration depths of the evanescent electromagnetic field. Recording of fluorescence images upon irradiation at about 10 different angles permits to calculate cell-substrate distances with a precision of a few nanometers. Differences of adhesion between various cell lines, e.g. cancer cells and less malignant cells, are thus determined. In addition, possible changes of cell adhesion upon chemical or photodynamic treatment can be examined. In comparison with other methods of super-resolution microscopy light exposure is kept very small, and no damage of living cells is expected to occur.

Nanotopology of Cell Adhesion upon Variable-Angle Total Internal Reflection Fluorescence Microscopy VA-TIRFM

Topology of cell adhesion on a substrate is measured with nanometre precision by variable-angle total internal reflection fluorescence microscopy (VA-TIRFM).

Surface topology, e.g. of cells growing on a substrate, is determined with nanometer precision by Variable-Angle Total Internal Reflection Fluorescence ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

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., &#946;2-adrenergic and &#947;-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.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

Visualizing Angiogenesis by Multiphoton Microscopy In Vivo in Genetically Modified 3D-PLGA/nHAp Scaffold for Calvarial Critical Bone Defect Repair

The reconstruction of critically sized bone defects remains a serious clinical problem because of poor angiogenesis within tissue-engineered scaffolds during repair, which gives rise to a lack of sufficient blood supply and causes necrosis of the new tissues. Rapid vascularization is a vital prerequisite for new tissue survival and integration with existing host tissue. The de novo generation of vasculature in scaffolds is one of the most important steps in making bone regeneration more efficient, allowing repairing tissue to grow into a scaffold. To tackle this problem, the genetic modification of a biomaterial scaffold is used to accelerate angiogenesis and osteogenesis. However, visualizing and tracking in vivo blood vessel formation in real-time and in three-dimensional (3D) scaffolds or new bone tissue is still an obstacle for bone tissue engineering. Multiphoton microscopy (MPM) is a novel bio-imaging modality that can acquire volumetric data from biological structures in a high-resolution and minimally-invasive manner. The objective of this study was to visualize angiogenesis with multiphoton microscopy in vivo in a genetically modified 3D-PLGA/nHAp scaffold for calvarial critical bone defect repair. PLGA/nHAp scaffolds were functionalized for the sustained delivery of a growth factor pdgf-b gene carrying lentiviral vectors (LV-pdgfb) in order to facilitate angiogenesis and to enhance bone regeneration. In a scaffold-implanted calvarial critical bone defect mouse model, the blood vessel areas (BVAs) in PHp scaffolds were significantly higher than in PH scaffolds. Additionally, the expression of pdgf-b and angiogenesis-related genes, vWF and VEGFR2, increased correspondingly. MicroCT analysis indicated that the new bone formation in the PHp group dramatically improved compared to the other groups. To our knowledge, this is the first time multiphoton microscopy was used in bone tissue-engineering to investigate angiogenesis in a 3D bio-degradable scaffold in vivo and in real-time.

Visualizing Angiogenesis by Multiphoton Microscopy In Vivo in Genetically Modified 3D-PLGA/nHAp Scaffold for Calvarial Critical Bone Defect Repair

Here, we present a protocol to visualize blood vessel formation in vivo and in real-time in 3D scaffolds by multiphoton microscopy. Angiogenesis in genetically modified scaffolds was studied in a murine calvarial critical bone defect model. More new blood vessels were detected in the treatment group than in controls.

The reconstruction of critically sized bone defects remains a serious clinical problem because of poor angiogenesis within tissue-engineered scaffolds ...

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic significance. Possible reasons for the lack of exploration are the experimental limitations and technical challenges that render the study of cell division in 3D culture inefficient. Here, we describe an imaging-based method to efficiently study mammalian cell division and cell-matrix interactions in 3D collagen matrices. Cells labeled with fluorescent H2B are synchronized using the combination of thymidine blocking and nocodazole treatment, followed by a mechanical shake-off technique. Synchronized cells are then embedded into a 3D collagen matrix. Cell division is monitored using live-cell microscopy. The deformation of collagen fibers during and after cell division, which is an indicator of cell-matrix interaction, can be monitored and quantified using quantitative confocal reflection microscopy. The method provides an efficient and general approach to study mammalian cell division and cell-matrix interactions in a physiologically relevant 3D environment. This approach not only provides novel insights into the molecular basis of the development of normal tissue and diseases, but also allows for the design of novel diagnostic and therapeutic approaches.

This protocol efficiently studies mammalian cell division in 3D collagen matrices by integrating synchronization of cell division, monitoring of division events in 3D matrices using live-cell imaging technique, time-resolved confocal reflection microscopy and quantitative imaging analysis.

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic ...

Assessing Collagen and Elastin Pressure-dependent Microarchitectures in Live, Human Resistance Arteries by Label-free Fluorescence Microscopy

The pathogenic contribution of resistance artery remodeling is documented in essential hypertension, diabetes and the metabolic syndrome. Investigations and development of microstructurally motivated mathematical models for understanding the mechanical properties of human resistance arteries in health and disease have the potential to aid understanding how disease and medical treatments affect the human microcirculation. To develop these mathematical models, it is essential to decipher the relationship between the mechanical and microarchitectural properties of the microvascular wall. In this work, we describe an ex vivo method for passive mechanical testing and simultaneous label-free three-dimensional imaging of the microarchitecture of elastin and collagen in the arterial wall of isolated human resistance arteries. The imaging protocol can be applied to resistance arteries of any species of interest. Image analyses are described for quantifying i) pressure-induced changes in internal elastic lamina branching angles and adventitial collagen straightness using Fiji and ii) collagen and elastin volume densities determined using Ilastik software. Preferably all mechanical and imaging measurements are performed on live, perfused arteries, however, an alternative approach using standard video-microscopy pressure myography in combination with post-fixation imaging of re-pressurized vessels is discussed. This alternative method provides users with different options for analysis approaches. The inclusion of the mechanical and imaging data in mathematical models of the arterial wall mechanics is discussed, and future development and additions to the protocol are proposed.

We describe simultaneous mechanical testing and 3D-imaging of the arterial wall of isolated, live human resistance arteries, and Fiji and Ilastik image analyses for the quantification of elastin and collagen spatial organization and volume densities. We discuss the use of these data in mathematical models of arterial wall mechanics.

The pathogenic contribution of resistance artery remodeling is documented in essential hypertension, diabetes and the metabolic syndrome. Investigations ...

Wet-spinning-based Molding Process of Gelatin for Tissue Regeneration

This article presents an inexpensive method to fabricate gelatin, as a natural polymer, into monofilament fibers or other appropriate forms. Through the wet spinning method, gelatin fibers are produced by smooth extrusion in a suitable coagulation medium. To increase the functional surface of these gelatin fibers and their ability to mimic the features of tissues, gelatin can be molded into a tube form by referring to this concept. Examined by in vitro and in vivo tests, the gelatin tubes demonstrate a great potential for application in tissue engineering. Acting as a suitable filling gap material, gelatin tubes can be used to substitute the tissue in the damaged area (e.g., in the nervous or cardiovascular system), as well as to promote regeneration by providing a direct replacement of stem cells and neural circuitry. This protocol provides a detailed procedure for creating a biomaterial based on a natural polymer, and its implementation is expected to greatly benefit the development of correlative natural polymers, which help to realize tissue regeneration strategies.

We developed and describe a protocol based on the wet spinning concept, for the construction of gelatin-based biomaterials used for the application of tissue engineering.

This article presents an inexpensive method to fabricate gelatin, as a natural polymer, into monofilament fibers or other appropriate forms. Through the ...