The authors thank Dr. Soucy Patricia and Betty Nunn at the University of Louisville for assistance with the confocal microscope. This work was funded by the National Science Foundation (1456884 to M.P.R.) and by a National Science Foundation Cooperative Agreement (1849213 to M.P.R.).

NOTE: See the Table of Materials for the list of materials and equipment and Table 1 for the list of solutions to be used in this protocol.
1. Preparation of plants for glass bottom dishes
<ol>
	<li>Grow the moss protonemal tissue in BCDAT agar medium in a 13 cm Petri dish under constant white light (~50 &#181;mol m-2s-1) for 7 days at 25 &#176;C in the growth chamber.</li>
	<li>Filter-sterilize the calcofluor white and store at 4 &#176;C until use. The reagent is stable for several months.</li>
	<li>Disperse the 7-day-old moss tissue into a 1.5 mL microtube containing 20 &#181;L of sterilized water. Ensure the moss protonemal tissue is sufficiently dispersed as small pieces in the water. For wild type (WT), use forceps to separate the protonemata; for the ggb mutant, use a P-1000 pipette.</li>
	<li>Add 10 &#181;L of the sterilized calcofluor white into the 1.5 mL microtube and leave the microtube upright in the hood for 2 min for staining.</li>
	<li>Immediately after staining, mix the plants with 200 &#181;L of cooled BCDATG, containing BCDAT medium supplemented with 0.5% (w/v) glucose and 0.4% (w/v) gellan gum, by pipetting five times with a P-1000 pipette. 
	NOTE: Ensure the BCDATG medium is cooled enough (just before the medium solidifies) to avoid stressing the plants.</li>
	<li>Transfer the mixture containing the plants and the BCDATG medium into a 27 mm diameter glass base dish. Ensure that the volume of BCDATG for mixing the cells is no more than 200 &#181;L and that the 200 &#181;L of BCDATG with samples is evenly distributed on the surface of the 7 mm glass bottom dish to produce a thin layer of film, so that the samples are within the objective working distance during imaging.</li>
	<li>After solidifying for 2 min at room temperature, cover the thin layer with 3 mL of cooled BCDATG and let it set for 10 min to solidify again. 
	NOTE: Ensure that steps 1.2-1.5 are conducted in a sterile hood to avoid contamination.</li>
	<li>On the bottom of the dish, draw nine lines each in parallel and perpendicular directions (Figure 2). Mark the rows with characters from a to h, and mark the columns with numbers from 1 to 8. Use upper, lower, right, middle, and left to mark the positions within each square. For example, if a plant is in a square in the second row and the sixth column and is positioned in the upper left part of the square, then this plant is given the name of &#34;b6_upper_left&#34;.</li>
	<li>Preculture the glass bottom dish containing the plants under red light for 5 days at 25 &#176;C in the growth chamber, and then under white light for 1-2 days before being subjected to time-lapse imaging every day for up to 3 days. For WT, preculture the plants in weak red light (0.5 &#181;mol m-2s-1) from the above for 5 days to induce the phototropic response of the protonemata, so that the plants can attach to the bottom of the dishes11. 
	NOTE: The weak red light is provided by passing a white light through a 3 mm thick red acrylic plastic filter into light-proof boxes. For ggb mutants, preculturing in red light is optional, as the ggb mutants do not have a phototropic response3.</li>
</ol>
2. Imaging and 3-D reconstruction
<ol>
	<li>Place the glass bottom dish containing plants onto the stage of an inverted confocal microscope, with the glass bottom facing the objective. Use a confocal microscope for the visualization of calcofluor with a 20x objective lens. Take images with the 405 nm laser, the pinhole at 1.0, HV-gain at 100, offset-background adjustment at 0, Laser Power at 5%-7%, a scan size of 1,024 x 1,024, and a scan speed of 1/2 frame/s. Collect 17-30 z-series optical sections with a step size of 1.0 &#181;m. Name the images using the code in step 1.6, such as &#34;b6_upper_left-day0&#34;, and save.
	<ol>
		<li>Select the DAPI channel in Acquisition and check the Z box in ND Acquisition. To set the lower and upper limits for the Z-stack, click the Top and Bottom buttons.</li>
	</ol>
	</li>
	<li>To reconstruct the 3-D images, open the .nd2 file (Figure 3A; see Table of Materials) and click the Volume (Figure 3B).</li>
</ol>
3. Imaging the same plants at different time points
<ol>
	<li>After each imaging, put the glass base dish back into the growth chamber.</li>
	<li>Repeat from step 2.1 for imaging and 3-D reconstruction. 
	NOTE: To find the same plants, use the code mentioned in step 1.6 and give the name &#34;b6_upper_left-day1&#34;, or &#34;b6_upper_left-day2&#34;, according to the age of plants.</li>
</ol>

<ol>
	<li>Anderson, C. T., Kieber, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+construction,+perception,+and+remodeling+of+plant+cell+walls.">Dynamic construction, perception, and remodeling of plant cell walls.</a> Annual Review of Plant Biology. 71, 39-69 (2020).</li><li>Vaahtera, L., Schulz, J., Hamann, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+wall+integrity+maintenance+during+plant+development+and+interaction+with+the+environment.">Cell wall integrity maintenance during plant development and interaction with the environment.</a> Naure Plants. 5 (9), 924-932 (2019).</li><li>Bao, L., 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=The+cellular+function+of+ROP+GTPase+prenylation+important+for+multicellularity+in+the+moss+Physcomitrium+patens.">The cellular function of ROP GTPase prenylation important for multicellularity in the moss Physcomitrium patens.</a> Development. 149 (12), (2022).</li><li>Colin, L., 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=Imaging+the+living+plant+cell:+From+probes+to+quantification.">Imaging the living plant cell: From probes to quantification.</a> The Plant Cell. 34 (1), 247-272 (2022).</li><li>Fang, Y., Spector, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+cell+imaging+of+plants.">Live cell imaging of plants.</a> Cold Spring Harbor Protocols. 2010 (2), (2010).</li><li>Goh, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live-cell+imaging+approaches+to+study+lateral+root+formation+in+Arabidopsis+thaliana.">Long-term live-cell imaging approaches to study lateral root formation in Arabidopsis thaliana.</a> Microscopy. 68 (1), 4-12 (2019).</li><li>Hamant, O., Das, P., Burian, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-lapse+imaging+of+developing+shoot+meristems+using+a+confocal+laser+scanning+microscope.">Time-lapse imaging of developing shoot meristems using a confocal laser scanning microscope.</a> Methods in Molecular Biology. 1992, 257-268 (2019).</li><li>Rigal, A., Doyle, S. M., Robert, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+cell+imaging+of+FM4-64,+a+tool+for+tracing+the+endocytic+pathways+in+Arabidopsis+root+cells.">Live cell imaging of FM4-64, a tool for tracing the endocytic pathways in Arabidopsis root cells.</a> Methods in Molecular Biology. 1242, 93-103 (2015).</li><li>Yagi, N., 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=Advances+in+synthetic+fluorescent+probe+labeling+for+live-cell+imaging+in+plants.">Advances in synthetic fluorescent probe labeling for live-cell imaging in plants.</a> Plant &amp; Cell Physiology. 62 (8), 1259-1268 (2021).</li><li>Whitewoods, C. D., 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=CLAVATA+was+a+genetic+novelty+for+the+morphological+innovation+of+3D+growth+in+land+plants.">CLAVATA was a genetic novelty for the morphological innovation of 3D growth in land plants.</a> Current Biology. 28 (15), 2365-2376 (2018).</li><li>Bao, L., 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+PSTAIRE-type+cyclin-dependent+kinase+controls+light+responses+in+land+plants.">A PSTAIRE-type cyclin-dependent kinase controls light responses in land plants.</a> Science Advances. 8 (4), 2116 (2022).</li></ol>

The authors declare no competing or financial interests.

Time-lapse 3-D reconstruction, or 4-D imaging, is a powerful tool for observing the dynamics of cellular morphology during developmental processes. In this protocol, by mixing the calcofluor white in the medium, the dynamics of 3-D cellular morphology can be observed in the moss P. patens. Using this method, we observed that the surface of cell walls in ggb mutants are torn apart during development3. Moreover, the reduced thickening of cell walls is correlated with the cell expan...

Plant cell walls undergo dynamic changes during cell expansion and development1,2,3. Maintaining cell wall integrity is critical for plant cell adhesion during growth and development, as well as for the response to environmental signals. Although visualizing cell wall dynamics of living cells over a long period of time is critical to understanding how cell adhesion is maintained during development and adaptation to environmental changes, current methods for directly observing cell wall dynamics are still challenging.
Time-lapse imaging of cellular changes can provide informative developmental dynamics of an organism using a high-resolution fluorescence microscope4,5,6,7. While time-lapse 3-D imaging has a great deal of potential for studying dynamic changes of cell shape during growth and development, the technique normally requires transformation of a fluorescent protein4,5,6,7. However, for most systems, genetic transformations are either time-consuming or technically challenging. As an alternative, fluorescent dyes that attach to cellular components have long been available. The fluorescent dyes can emit fluorescent light after irradiation with light of a certain wavelength. Common examples are Edu, DAPI, PI, FM4-64, and calcofluor white8,9,10. One major drawback, however, is that these dyes can typically only be used in fixed tissue or for short experiments, in part due to the harm they cause to the cell8,9,10.
With the protocol presented here, calcofluor signals are stable when calcofluor white is mixed within the medium during time-lapse experiments in the moss P. patens. Using this method, the detachment of cells in ggb mutants using 3-D time-lapse imaging was observed over a 3 day period3 (Figure 1). This method can be applied to many other systems that contain cell walls and that can be stained by calcofluor.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3-mm-thick red plastic light filter</td>
			<td>&#160;Mitsubishi</td>
			<td>&#160;no.102</td>
			<td></td>
		</tr>
		<tr>
			<td>27 mm diameter glass base dish&#160;</td>
			<td>Iwaki</td>
			<td>3930-035</td>
			<td></td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>&#160;Sigma</td>
			<td>A6924</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcofluor white&#160;</td>
			<td>&#160;Sigma</td>
			<td>18909-100ML-F</td>
			<td>Calcofluor White M2R, 1 g/L and Evans blue, 0.5 g/L</td>
		</tr>
		<tr>
			<td>Confocal microscope&#160;</td>
			<td>Nikon&#160;</td>
			<td>A1</td>
			<td>
			NIS element software; .nd2 file in NIS-elements Viewer, download from https://www.microscope.healthcare.nikon. 
			com/en_AOM/products/software/nis-elements/viewer
			</td>
		</tr>
		<tr>
			<td>Fluorescence microscope&#160;</td>
			<td>Nikon&#160;</td>
			<td>TE200&#160;</td>
			<td>Equipped with a DS-U3 camera;</td>
		</tr>
		<tr>
			<td>Gellan gum&#160;</td>
			<td>Nacali Tesque</td>
			<td>12389-96</td>
			<td></td>
		</tr>
		<tr>
			<td>Plant Growth Chambers</td>
			<td>SANYO&#160;</td>
			<td>Sanyo MLR-350H</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilized syringe 0.22 &#956;m filter</td>
			<td>Millipore</td>
			<td>SLGV033RS</td>
			<td></td>
		</tr>
	</tbody>
</table>

3-d time-lapse imaging of cell wall dynamics using calcofluor in the moss physcomitrium patens

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In general, for observations over a long period, the technique requires transformation of a fluorescent protein; however, for most systems, genetic transformation is either time-consuming or technically unavailable. This manuscript presents a protocol for 3-D time-lapse imaging of cell wall dynamics over a 3 day period using calcofluor dye (which stains cellulose in the plant cell wall), developed in the moss Physcomitrium patens. The calcofluor dye signal from the cell wall is stable and can last for 1 week without obvious decay. Using this method, it has been shown that the detachment of cells in ggb mutants (in which the protein geranylgeranyltransferase-I beta subunit is knocked out) is caused by unregulated cell expansion and cell wall integrity defects. Moreover, the patterns of calcofluor staining change over time; less intensely stained regions correlate with the future cell expansion/branching sites in the wild type. This method can be applied to many other systems that contain cell walls and that can be stained by calcofluor.

This method allows the observation of cell wall dynamics during development in wild type and ggb mutants (Figure 1). The results showed that regions with less thickening of the cell wall correlate with the cell expansion/branching sites, allowing for the prediction of expansion/branching sites in the wild type (Figure 1A). The surface of the cell walls in ggb mutants was torn apart during development due to uncontrolled cell expansion<sup class...

Watch this Scientific Journal Video about 3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens at JoVE.com

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

This manuscript presents a detailed protocol to image the 3-D cell wall dynamics of living moss tissue, allowing the visualization of the detachment of cell walls in ggb mutants and thickening cell wall patterns in the wild type during development over a long period.

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In ...

3-d-time-lapse-imaging-cell-wall-dynamics-using-calcofluor-moss

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Developmental Biology

1.3K Views.  University of Louisville. This manuscript presents a detailed protocol to image the 3-D cell wall dynamics of living moss tissue, allowing the visualization of the detachment of cell walls in ggb mutants and thickening cell wall patterns in the wild type during development over a long period.

Video: 3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of morphogenetic events, such as gastrulation2, neurulation3-5, somitogenesis6, heart bending7,8, and brain formation9-13, during early embryogenesis. Key to understanding morphogenetic processes is to follow them dynamically by time-lapse imaging. The acquisition of time-lapse movies of chick embryogenesis ex ovo has been limited either to short time windows or to the need for an incubator to control temperature and humidity around the embryo14. Here, we present a new technique to culture chick embryos ex ovo for high-resolution time-lapse imaging using transmitted light microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique (EC-culture)1 and allows for the culturing of chick embryos without the need for a climate chamber. The embryo is sandwiched between two identical filter paper carriers and is kept fully submerged in a simple, temperature-controlled medium covered by a layer of light mineral oil. Starting from the primitive streak stage (Hamburger-Hamilton stage 5, HH5)15 up to at least the 28-somite stage (HH16)15, embryos can be cultured with either their ventral or dorsal side up. This allows the acquisition of time-lapse movies covering about 30 hr of embryonic development. Representative time-lapse frames and movies are shown. Embryos are compared morphologically to an embryo cultured in the standard EC-culture.
The submerged filter paper sandwich provides a stable environment to study early dorsal and ventral morphogenetic processes. It also allows for live fluorescence imaging and micromanipulations, such as microsurgery, bead implantation, microinjection, gene silencing, and electroporation, and has a strong potential to be combined with immersion objectives for laser-based imaging (including light-sheet microscopy).

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Here, we present a new technique to culture chick embryos ex ovo for time-lapse imaging using transmitted light and fluorescence microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique1 that allows for the culturing of chick embryos fully-submerged in a liquid culture medium.

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of ...

Production and Administration of Therapeutic Mesenchymal Stem/Stromal Cell (MSC) Spheroids Primed in 3-D Cultures Under Xeno-free Conditions

Mesenchymal stem/stromal cells (MSCs) hold great promise in bioengineering and regenerative medicine. MSCs can be isolated from multiple adult tissues via their strong adherence to tissue culture plastic and then further expanded in vitro, most commonly using fetal bovine serum (FBS). Since FBS can cause MSCs to become immunogenic, its presence in MSC cultures limits both clinical and experimental applications of the cells. Therefore, studies employing chemically defined xeno-free (XF) media for MSC cultures are extremely valuable. Many beneficial effects of MSCs have been attributed to their ability to regulate inflammation and immunity, mainly through secretion of immunomodulatory factors such as tumor necrosis factor-stimulated gene 6 (TSG6) and prostaglandin E2 (PGE2). However, MSCs require activation to produce these factors and since the effect of MSCs is often transient, great interest has emerged to discover ways of pre-activating the cells prior to their use, thus eliminating the lag time for activation in vivo. Here we present protocols to efficiently activate or prime MSCs in three-dimensional (3D) cultures under chemically defined XF conditions and to administer these pre-activated MSCs in vivo. Specifically, we first describe methods to generate spherical MSC micro-tissues or &#39;spheroids&#39; in hanging drops using XF medium and demonstrate how the spheres and conditioned medium (CM) can be harvested for various applications. Second, we describe gene expression screens and in vitro functional assays to rapidly assess the level of MSC activation in spheroids, emphasizing the anti-inflammatory and anti-cancer potential of the cells. Third, we describe a novel method to inject intact MSC spheroids into the mouse peritoneal cavity for in vivo efficacy testing. Overall, the protocols herein overcome major challenges of obtaining pre-activated MSCs under chemically defined XF conditions and provide a flexible system to administer MSC spheroids for therapies.

Production and Administration of Therapeutic Mesenchymal Stem/Stromal Cell MSC Spheroids Primed in 3-D Cultures Under Xeno-free Conditions

The therapeutic potential of mesenchymal stem/stromal cells (MSCs) is well-documented, however the best method of preparing the cells for patients remains controversial. Herein, we communicate protocols to efficiently generate and administer therapeutic spherical aggregates or 'spheroids' of MSCs primed under xeno-free conditions for experimental and clinical applications.

Mesenchymal stem/stromal cells (MSCs) hold great promise in bioengineering and regenerative medicine. MSCs can be isolated from multiple adult tissues via ...

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune cells, and the coordination with lymphatic vessels and nerves. The multi-cell, multi-system interactions necessitate the investigation of angiogenesis in a physiologically relevant environment. Thus, while the use of in vitro cell-culture models have provided mechanistic insights, a common critique is that they do not recapitulate the complexity associated with a microvascular network. The objective of this protocol is to demonstrate the ability to make time-lapse comparisons of intact microvascular networks before and after angiogenesis stimulation in cultured rat mesentery tissues. Cultured tissues contain microvascular networks that maintain their hierarchy. Immunohistochemical labeling confirms the presence of endothelial cells, smooth muscle cells, pericytes, blood vessels and lymphatic vessels. In addition, labeling tissues with BSI-lectin enables time-lapse comparison of local network regions before and after serum or growth factor stimulation characterized by increased capillary sprouting and vessel density. In comparison to common cell culture models, this method provides a tool for endothelial cell lineage studies and tissue specific angiogenic drug evaluation in physiologically relevant microvascular networks.

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis involves multi-cell, multi-system interactions that need to be investigated in a physiologically relevant environment. The objective of this study is to demonstrate the ability of the rat mesentery culture model to make time-lapse comparisons of intact microvascular networks during angiogenesis.

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...