The authors would like to thank Travis Wheeler and the machine shop at the Department of Cell Biology and Physiology, University of Pittsburgh for construction of the cryo-fluorescence sample stage, Changlu Tao and Cheng Xu at the University of Science and Technology of China for technical assistance, and Dr. Teresa Brosenitsch for critical reading of the manuscript. This work was supported by the National Institutes of Health (GM082251 &amp; GM085043).

1. HeLa Cell Culture on Carbon-coated, Gold EM Finder Grids
<ol>
	<li>Glow discharge the carbon-side of a 200-mesh R2/2 Quantifoil gold EM finder grid under 25 mA for 25 sec.</li>
	<li>Coat the EM grid with fibronectin, by floating it, carbon-side down, onto a 40 &mu;l droplet of 50 &mu;g/ml fibronectin solution, then disinfect it under UV light for 2 hr in a tissue culture hood.</li>
	<li>Plate HeLa cells onto the grid at a density of 2 x 104 cells/ml (total 2 ml culture) in a glass-bottom culture dish and grow the cells at 37 &deg;C, with 5% CO2, in DMEM supplemented with 4.5 g/L L-glutamine and glucose, 10% heat-inactivated fetal calf serum, 100 units/ml penicillin, and 100 &mu;g/ml streptomycin, for approximately 18 hr. HeLa cells are infected after O/N culture.</li>
</ol>
2. HIV-1 Infection and Live-cell Imaging
<ol>
	<li>Add a fluorescent cell tracker (Red CMTPX, 1 &mu;l/2 ml) into the glass-bottom culture dish (from step 1.3) and incubate the dish at 37 &deg;C for 10 min, to allow up-take of the fluorescent dye.</li>
	<li>Wash the cells with PBS and add 50 &mu;l pre-warmed fresh medium.</li>
	<li>Place the dish onto the live-cell chamber, at 37 &deg;C, of a Swept Field Confocal scanner microscope. Select multiple fields (10-15 positions) that contain 1-3 cells/square, with the cells between two mitosis phases when they are most spread-out and flat (see Figures 2c and 2d), since cryoEM requires relatively thin regions of the sample. Store these positions for future imaging (step 2.5).</li>
	<li>Infect the cells with VSV-G pseudo-typed HIV-1 particles that contain GFP-Vpr (we use 40 &mu;l of a sample containing 40 ng/ml of p24). When adding the virus particles into the bottom of the dish, be careful to not disturb the EM grid, since the positions for imaging have already been selected and stored.</li>
	<li>Immediately after addition of virions, collect time-lapse high-speed 3D confocal images, at the previously selected positions (from step 2.3) for 20-40 min.</li>
	<li>Confocal images were acquired using MetaMorph and analyzed by automated 3D particle tracking for single particle dynamics using Imaris software (See Figure 1). Since we are correlating the dynamic behavior of diffraction-limited HIV-1 particles with cellular ultrastructure, time-lapse live-cell imaging and 3D particle tracking are critical to the results. However, for a large and static structure, a simple fluorescence image (without live-cell) would be sufficient for correlative analysis.</li>
</ol>
3. Frozen-hydrated EM Sample Preparation
To minimize the time delay between collection of the last confocal image and cryo-fixation of the sample, the FEI Vitrobot (or other vitrification device) should be initiated and prepared for plunge-freezing during collection of fluorescence confocal live-cell images.
<ol>
	<li>Turn on the vitrification device and set its climate temperature to 22 &deg;C, the target humidity to 100%, the blotting time to 7 sec, and the wait and drain time to 1 sec.</li>
	<li>Place the grid storage box in the plunger dewar and prepare cold liquid ethane. Mount the dewar onto the vitrification device.</li>
	<li>Immediately after confocal live-cell imaging (section 2), place the culture dish onto an ice-cooled copper block and transfer it to the cryoEM sample preparation room. Load the EM grid onto the specialized tweezers and quickly blot away any media on the grid. The blotting is done manually, with filter paper, after which 4 &mu;l of 15 nm gold bead solution mixed with 0.2 &mu;m fluorescent microspheres is immediately placed on the grid. The gold beads are used as fiducial markers to aid tomographic data collection and alignment. The fluorescent microspheres are used to aid correlation between fluorescent and cryoEM images (See Figures 2c-2f).</li>
	<li>Load the tweezers onto the vitrification device and instruct the device to blot and plunge freeze in liquid ethane16. The blotting parameters optimized to achieve best result are: blot time, 7 sec; blot offset, 0; drain time, 1 sec; temperature, 22 &deg;C; and humidity, 100%.</li>
	<li>Transfer the frozen-hydrated grid to a cryoEM holder for immediate cryoET imaging, or to a liquid nitrogen storage dewar for later use.</li>
</ol>
4. Cryo-fluorescence Light Microscopy
A home-built, cryo-fluorescence sample chamber and stage system (Figure 3) is required to carry out steps 4.1- 4.10. The detailed specifications and description of the stage may be found in Jun et al.15.
<ol>
	<li>Fill a self-pressurized liquid nitrogen dewar with liquid nitrogen at least 2 hr before starting cryo-fluorescence light microscopy (See Figure 3a).</li>
	<li>Mount a homebuilt cryo-fluorescence sample stage15 (Figure 3) onto an inverted fluorescence light microscope, such as an Olympus IX 71.</li>
	<li>Connect the liquid nitrogen inlet of the cryo-sample stage to the self-pressurized dewar and place the liquid nitrogen overflow protection outlet into an appropriate container. Connect a dry nitrogen gas line to a sleeve placed over the objective lens to keep the lens warm and free of frost.</li>
	<li>Place a copper block platform (black arrow in Figure 3b) into the cryo-stage chamber for loading the frozen-hydrated sample grid. Cool down and fill the copper chamber of the cryo-stage with liquid nitrogen through the inlet line from the self-pressurized filling dewar.</li>
	<li>When the cryo-stage reaches liquid nitrogen temperature (in ~4-6 min), transfer the grid storage box (from step 3.5) to the cryo-stage.</li>
	<li>Place the frozen-hydrated sample grid into the EM specimen cartridge on the copper block and clip the grid in place using a C-ring (if using a Polara microscope cartridge), or place a pre-cooled copper ring on top (black arrowhead in Figure 3d), to keep the grid in place and also for easy retrieval of the grid after cryo-fluorescence imaging (if for a non-Polara cryo-electron microscope).</li>
	<li>Place the cartridge (from step 4.6) in the inner chamber of the cryo-stage (white arrowhead in Figure 3b).</li>
	<li>Search and find the same virus particles from the live-cell imaging data. Since the EM grid has an index, it is straightforward to localize the same particle on the grid in both live-cell images and cryo-florescence images.</li>
	<li>Acquire cryo-DIC and GFP images at the identified positions under cryo-condition using a light microscope with a long working (2.7-4 mm) objective lens. During the cryo-fluorescence imaging, periodically check the liquid nitrogen level in the cryo-stage and refill it, as needed, to keep the sample stage below -170 &deg;C.</li>
	<li>Upon completion of cryo-fluorescence imaging, carefully return the specimen cartridge to the copper block platform and store the cartridge in a liquid nitrogen dewar for future cryoET analysis with a Polara system. If using a non-Polara cryo-electron microscope, remove the copper ring, retrieve the sample grid, place it in a grid storage box, and store the specimen in a liquid nitrogen dewar until the specimen is examined by cryoET.</li>
	<li>For data analysis, overlap the acquired cryo-DIC and GFP images (from step 4.9, see Figure 4b) and correlate the positions of the GFP signals in the cryo-fluorescence image with those obtained in the live-cell imaging series.</li>
</ol>
5. Cryo-electron Tomography
<ol>
	<li>Load the sample cartridge into the cryo-transfer station of an electron microscope equipped with a field emission gun, such as Polara G2 microscope.</li>
	<li>Under low-dose-search mode at a magnification of 140X, identify and save the associated grid squares (from step 4.11) into a stage file.</li>
	<li>Under low-dose-search mode at a magnification of 3,500X, insert a 100 &mu;m objective aperture, search and save all the positions correlated with GFP signals into a second stage file.</li>
	<li>Under low-dose-exposure mode, find a blank area to adjust beam intensity to a dose of 1 or 2 e-/&Aring;2 and refine the tilt axis, using the stage y function, by burning away the ice in an unimportant carbon region with several gold beads.</li>
	<li>Under low-dose-focus mode, adjust the focus distance to ~3 &mu;m and adjust the angle to align the direction of focus to be parallel to the tilt axis.</li>
	<li>Under low-dose-search mode, recall saved positions (from step 5.3), adjust the specimen eucentric height and check the tilt range for the area of interest. Acquire a projection image with very low electron-dose (~1 e-/&Aring;2) at 8 to 10 &mu;m defocus, in exposure mode, to confirm the correlated virus particles for cryo-electron tomography.</li>
	<li>Switch to the low-dose-focus mode. In TEM auto functions menu, precede the automated eucentric height and focus functions on the focus area.</li>
	<li>Set up the parameters for acquiring a tilt-series. For Figure 4 in this paper, tilt angles ranging &plusmn;70&deg;, below and above 45&deg;, at 3&deg; and 2&deg; tilt increments, respectively, were used, with a 6 &mu;m defocus and approximately 70 e-/&Aring;2 total dose.</li>
	<li>Repeat steps 5.7 and 5.8 for all the saved positions. Batch tomography software can be used for automated data collection at multiple positions to increase the throughput.</li>
</ol>
6. Three-dimensional Reconstruction using IMOD17
<ol>
	<li>Inspect the raw tilt series to adjust the minimum and maximum density values and determine if there is any view to be excluded due to large image shift.</li>
	<li>Start eTomo and setup tomogram panel using axis type, pixel size, fiducial diameter, etc. Then create com scripts.</li>
	<li>Configure the main window such that several stages of the tomogram computation can be adjusted/followed simultaneously. Modify the necessary parameters and execute the specific programs that are required by each processing step (See eTomo tutorial, <a href="http://bio3d.colorado.edu/imod/doc/etomoTutorial.html" target="_blank">http://bio3d.colorado.edu/imod/doc/etomoTutorial.html</a>).</li>
	<li>At the final stage, create a full aligned stack (with a mean residual error less than 0.6) and reconstruct tomograms using a weighted back-projection algorithm in IMOD.</li>
	<li>Denoise potential areas of interest using a 3D nonlinear anisotropic diffusion edge enhancing program implemented in IMOD with appropriate parameters to enhance the contrast and clarity.</li>
</ol>

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The authors declare no competing financial interests.

We have presented a straightforward set of protocols to provide an advanced correlative approach to analyze dynamic cellular events using time-lapse confocal, live-cell fluorescence imaging followed by cryoET. Our methodological development to correlate 3D live-cell imaging with high-resolution cryoET is critical to investigate many challenging biological problems, such as visualizing rare, dynamic (not static, as previously reported), and diffraction limited viral particles and their interactions with host cells. The sp...

Cryo-electron tomography (cryoET) is a powerful imaging technique that allows three-dimensional (3D) visualization of cells and tissues and provides insights into the organization of native organelles and cellular structures at molecular resolution in a close-to physiological state1. However, the inherently low contrast of unstained frozen-hydrated specimen, combined with their radiation sensitivity, makes it difficult to locate areas of interest inside a cell and subsequently performing the tilt series successfully without damaging the target area. In order to overcome these problems, a correlative approach that combines light and electron microscopy is necessary. Specific features highlighted by fluorescent labeling are identified and located by fluorescence light microscopy, and then their coordinates are transferred to the electron microscope for acquisition of high resolution 3D structural data. This correlative method helps locating the target areas of interest to be addressed. Due to the limitation on sample thickness with cryoEM (&lt;300 nm), currently only the peripheral regions of the cell are suitable for 3D structural analysis by CryoET. Further reducing the thickness of frozen-hydrated specimens by vitreous sectioning8 or by cryo-focused ion beam (FIB) milling9 would expand the capability of correlative imaging.
Previously, correlative methods were primarily used to facilitate cryoET data acquisition for large and static structures10-13. In these studies, cryo-stages have been implemented to accept cryoEM grids and fit onto either an upright microscope or an inverted microscope10,11,14. Although switching grids seems fairly straightforward in their designs, there are additional transferring steps involved for the EM grid, increasing the chance that the grid may be deformed, damaged and contaminated. We recently demonstrated a technical advance in correlative microscopy that allows us to directly visualize dynamic events that are by nature difficult to capture, such as HIV-1 and host cell interactions at early stages of infection15. We accomplished this by designing and implementing a cryo-light microscopy sample stage that adapts a cartridge system to minimize the specimen damage due to grid handling, thus facilitating correlation. Our design includes an integrated specimen cartridge holder, allowing both cryo-light and cryo-electron microscopy to be performed, sequentially, on the same specimen holder, without sample transfer, thus streamlining the correlative process. In addition, we also implemented an accurate and reliable correlation procedure using fluorescent latex beads as fiducial markers.

<table border="1" fo:keep-together.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagents</td>
		</tr>
		<tr>
			<td>DMEM 4.5 g/L Glucose w/ L-Glutamine</td>
			<td>Atlanta Biologicals, Inc. Lawrenceville, GA</td>
			<td>12-604F</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma, St. Louis, MO</td>
			<td>F1141-1MG</td>
			<td>HeLa cell culture on EM grids</td>
		</tr>
		<tr>
			<td>Cell tracker</td>
			<td>Invitrogen Corporation, Carlsbad, CA</td>
			<td>C34552</td>
			<td>Red CMTPX</td>
		</tr>
		<tr>
			<td>Protein A gold conjugates</td>
			<td>Ted Pella, Inc., Redding, CA</td>
			<td>15822-1</td>
			<td>15 nm diameter</td>
		</tr>
		<tr>
			<td>0.2 &mu;m fluorescent microspheres</td>
			<td>Invitrogen Corporation, Carlsbad, CA</td>
			<td>F8811</td>
			<td>Yellow-green fluorescent</td>
		</tr>
		<tr>
			<td>Heat-inactivated fetal calf bovine serum</td>
			<td>Invitrogen Corporation, Carlsbad, CA</td>
			<td>10082-139</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Invitrogen Corporation, Carlsbad, CA</td>
			<td>15140-122</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glass-bottom culture dish</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-14-C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gold quantifoil finder EM grids</td>
			<td>Quantifoil Micro Tools, Jena, Germany</td>
			<td>R2/2 Au NH2 200 mesh</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Invitrogen Corporation, Carlsbad, CA</td>
			<td>70011-044</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Glow-discharge device 100X</td>
			<td>EMS, Hatfield, PA</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tecnai Polara G2 electron microscope with a Field Emission Gun</td>
			<td>FEI, Hillsboro, OR</td>
			<td>&nbsp;</td>
			<td>300 keV</td>
		</tr>
		<tr>
			<td>Vitrobot Mark III</td>
			<td>FEI, Hillsboro, OR</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Olympus IX 71 microscope</td>
			<td>Olympus America Inc., Center Valley, PA</td>
			<td>&nbsp;</td>
			<td>LUCPlanFLN 40X/0.6 NA (2.7-4 mm working distance) objective lens</td>
		</tr>
		<tr>
			<td>Nikon TiE microscope</td>
			<td>Nikon Instruments, Melville, NY</td>
			<td>&nbsp;</td>
			<td>using a 60X/1.35 NA oil immersion objective lens</td>
		</tr>
		<tr>
			<td>Sweptfield confocal microscope</td>
			<td>Prairie Technologies, Middleton, WI</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tokai Hit live cell chamber</td>
			<td>Tokyo, Japan</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cryo-fluorescence sample stage</td>
			<td>Home-made</td>
			<td>&nbsp;</td>
			<td>Homebuilt by machine shop, see reference 15 for the design.</td>
		</tr>
	</tbody>
</table>

correlative microscopy for 3d structural analysis of dynamic interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

To characterize the dynamic behavior of the virus particles, HeLa cells infected with HIV-1 were imaged by high-speed confocal live-cell microscopy and the particle movements were analyzed by automated 3D particle tracking (Figure 1). To avoid the time lapse of several minutes that can occur between collection of the last confocal live-cell image and plunge-freezing (which may be long enough to lose correlated HIV-1 particles), a cryo-fluorescence light microscopy stage (Figure 3) was de...

Watch this Scientific Journal Video about Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions at JoVE.com

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

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Research

JoVE Journal

Bioengineering

13.9K Views.  University of Pittsburgh School of Medicine. We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Video: Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Studying the Stoichiometry of Epidermal Growth Factor Receptor in Intact Cells using Correlative Microscopy

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative fluorescence microscopy and environmental scanning electron microscopy (ESEM) of whole cells in hydrated state. Fluorescent quantum dots (QDs) were coupled to EGFR via a two-step labeling protocol, providing an efficient and specific protein labeling, while avoiding label-induced clustering of the receptor. Fluorescence microscopy provided overview images of the cellular locations of the EGFR. The scanning transmission electron microscopy (STEM) detector was used to detect the QD labels with nanoscale resolution. The resulting correlative images provide data of the cellular EGFR distribution, and the stoichiometry at the single molecular level in the natural context of the hydrated intact cell. ESEM-STEM images revealed the receptor to be present as monomer, as homodimer, and in small clusters. Labeling with two different QDs, i.e.,&#160;one emitting at 655 nm and at 800 revealed similar characteristic results.

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative light- and electron microscopy of whole cells in hydrated state. The label contained fluorescent quantum dots. The protocol can be used to study the stoichiometry of EGFR at the single molecule level.

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative fluorescence ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Correlative Light- and Electron Microscopy Using Quantum Dot Nanoparticles

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using widefield fluorescence light microscopy and transmission electron microscopy (TEM). To demonstrate the protocol we have immunolabeled ultrathin epoxy sections of human somatostatinoma tumor using a primary antibody to somatostatin, followed by a biotinylated secondary antibody and visualization with streptavidin conjugated 585 nm cadmium-selenium (CdSe) quantum dots (QDs). The sections are mounted on a TEM specimen grid then placed on a glass slide for observation by widefield fluorescence light microscopy. Light microscopy reveals 585 nm QD labeling as bright orange fluorescence forming a granular pattern within the tumor cell cytoplasm. At low to mid-range magnification by light microscopy the labeling pattern can be easily recognized and the level of non-specific or background labeling assessed. This is a critical step for subsequent interpretation of the immunolabeling pattern by TEM and evaluation of the morphological context. The same section is then blotted dry and viewed by TEM. QD probes are seen to be attached to amorphous material contained in individual secretory granules. Images are acquired from the same region of interest (ROI) seen by light microscopy for correlative analysis. Corresponding images from each modality may then be blended to overlay fluorescence data on TEM ultrastructure of the corresponding region.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of epoxy embedded human pathology tissue. We employ commercial antibody fragment conjugated QDs that are visualized by widefield fluorescence light microscopy and transmission electron microscopy.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using ...

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly controlled, high-throughput engineered system for measuring the impact of arrayed biochemical signals on cell differentiation. Recent efforts using cell microarrays have demonstrated their utility for combinatorial studies in which many microenvironmental factors are presented in parallel. However, these efforts have focused primarily on investigating the effects of biochemical cues on cell responses. Here, we present a cell microarray platform with tunable material properties for evaluating both cell differentiation by immunofluorescence and biomechanical cell&#8211;substrate interactions by traction force microscopy. To do so, we have developed two different formats utilizing polyacrylamide hydrogels of varying Young&#39;s modulus fabricated on either microscope slides or glass-bottom Petri dishes. We provide best practices and troubleshooting for the fabrication of microarrays on these hydrogel substrates, the subsequent cell culture on microarrays, and the acquisition of data. This platform is well-suited for use in investigations of biological processes for which both biochemical (e.g., extracellular matrix composition) and biophysical (e.g., substrate stiffness) cues may play significant, intersecting roles.

Cell differentiation is regulated by a host of microenvironmental factors, including both matrix composition and substrate material properties. We describe here a technique utilizing cell microarrays in conjunction with traction force microscopy to evaluate both cell differentiation and biomechanical cell&#8211;substrate interactions as a function of microenvironmental context.

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly ...

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.

Correlative Light Electron Microscopy CLEM for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

A correlative light electron microscopy (CLEM) method is applied to image virus-induced intracellular structures via electron microscopy (EM) in cells that are previously selected by light microscopy (LM). LM and EM are combined as a hybrid imaging approach to achieve an integrated view of virus-host interactions.

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing ...

Construction of a Multilayered Mesenchymal Stem Cell Sheet with a 3D Dynamic Culture System

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low cell retention and poor survival within the target zone. However, during the in vitro construction process, a solution for maintaining stem cell bioactivity and increasing the cell amount within the cell sheet is urgently needed. Here, this protocol presents a method for constructing a multilayered cell sheet with favorable stem cell bioactivity and optimal operability. Decellularized porcine pericardium (DPP) is prepared by phospholipase A2 (PLA2) decellularization method as the cell sheet scaffold, and rat bone marrow mesenchymal stem cells (BMSCs) are isolated and expanded as the seeded cells. The temporary multilayered cell sheet structure is constructed by using RAD16-I peptide hydrogel. Finally, the cell sheet is cultured with a dynamic perfusion system to stabilize the three-dimensional (3D) structure, and the cell sheet could be obtained following a 48-hour culture in vitro. This protocol provides an efficient and feasible method for constructing a multilayered stem cell sheet, and the cell sheet could be developed as a favorable stem cell therapy product in the future.

This article provides an efficient and feasible method for constructing multilayered stem cell sheets with favorable stem cell property.

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low ...

Environmental Dynamic Mechanical Analysis to Predict the Softening Behavior of Neural Implants

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior of these materials. In the past, it has not been possible to satisfactorily measure the softening of thin films under conditions mimicking body environment without substantial effort. This publication presents a new and simple method that allows dynamic mechanical analysis (DMA) of polymers in solutions, such as phosphate buffered saline (PBS), at relevant temperatures. The use of environmental DMA allows measurement of the softening effects of polymers due to plasticization in various media and temperatures, which therefore allows a prediction of the materials behavior under in vivo conditions.

To allow reliable predictions of the softening of polymeric substrates for neural implants in an in vivo environment, it is important to have a reliable in vitro method. Here, the use of dynamic mechanical analysis in phosphate buffered saline at body temperature is presented.

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior ...

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and chemical complexity of living tissues and mimic in vivo tissue functions. Recent advances in microfabrication technologies have facilitated the development of 3D in vitro environments in which cells can be integrated into a well-defined extracellular matrix (ECM) and a defined set of soluble or matrix associated biomolecules. However, technological barriers have limited their widespread use in research laboratories. Here, we describe a method to construct simple devices for 3D culture and experimentation with cells and multicellular organoids in 3D microenvironments with a defined chemoattractant gradient. We illustrate the use of this platform for analysis of the response of epithelial cells and organoids to gradients of growth factors, such as epidermal growth factor (EGF). EGF gradients were stable in the devices for several days leading to directed branch formation in breast organoids. This analysis allowed us to conclude that collective gradient sensing by groups of cells is more sensitive vs. single cells. We also describe the fabrication method, which does not require photolithography facilities nor advanced soft lithography techniques. This method will be helpful to study 3D cellular behaviors in the context of the analysis of development and pathological states, including cancer.

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and ...

Substructure Analyzer: A User-Friendly Workflow for Rapid Exploration and Accurate Analysis of Cellular Bodies in Fluorescence Microscopy Images

The last decade has been characterized by breakthroughs in fluorescence microscopy techniques illustrated by spatial resolution improvement but also in live-cell imaging and high-throughput microscopy techniques. This led to a constant increase in the amount and complexity of the microscopy data for a single experiment. Because manual analysis of microscopy data is very time consuming, subjective, and prohibits quantitative analyses, automation of bioimage analysis is becoming almost unavoidable. We built an informatics workflow called Substructure Analyzer to fully automate signal analysis in bioimages from fluorescent microscopy. This workflow is developed on the user-friendly open-source platform Icy and is completed by functionalities from ImageJ. It includes the pre-processing of images to improve the signal to noise ratio, the individual segmentation of cells (detection of cell boundaries) and the detection/quantification of cell bodies enriched in specific cell compartments. The main advantage of this workflow is to propose complex bio-imaging functionalities to users without image analysis expertise through a user-friendly interface. Moreover, it is highly modular and adapted to several issues from the characterization of nuclear/cytoplasmic translocation to the comparative analysis of different cell bodies in different cellular sub-structures. The functionality of this workflow is illustrated through the study of the Cajal (coiled) Bodies under oxidative stress (OS) conditions. Data from fluorescence microscopy show that their integrity in human cells is impacted a few hours after the induction of OS. This effect is characterized by a decrease of coilin nucleation into characteristic Cajal Bodies, associated with a nucleoplasmic redistribution of coilin into an increased number of smaller foci. The central role of coilin in the exchange between CB components and the surrounding nucleoplasm suggests that OS induced redistribution of coilin could affect the composition and the functionality of Cajal Bodies.

We present a freely available workflow built for rapid exploration and accurate analysis of cellular bodies in specific cell compartments in fluorescence microscopy images. This user-friendly workflow is designed on the open-source software Icy and also uses ImageJ functionalities. The pipeline is affordable without knowledge in image analysis.

The last decade has been characterized by breakthroughs in fluorescence microscopy techniques illustrated by spatial resolution improvement but also in ...

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...