The authors acknowledge Purdue Bindley Bioscience Center Imaging Facility for accessing the laser scanning confocal microscope and for the technical support, and the authors appreciate the help from Andy Schaber in the Purdue Bindley Imaging Facility. This activity was funded by Purdue University as part of AgSEED Crossroads funding to support Indiana&#8217;s Agriculture and Rural Development.

1. Media and imaging dishes preparation
<ol>
	<li>MS plates: Add 0.5x Murashige &#38; Skoog MS medium, 1% agar into deionized water and then adjust pH to 5.8 using potassium hydroxide solution (Optional: Add 1% sucrose for the long-term plant growing). Autoclave and pour plates.</li>
	<li>Imaging dishes: Fill plastic Petri dishes (6 cm wide, 1.5 cm depth) to 0.5-0.8 cm with 1.5% molten agarose.</li>
</ol>
2. Plant growth
<ol>
	<li>Arabidopsis growth
	<ol>
		<li>Sow sterilized seeds on MS plates and place plates under 4 &#176;C for two days. Then, move MS plates to a short day (8 h light/ 16 h dark), at 22 &#176;C for two weeks.</li>
		<li>Transplant seedlings to soil and grow them in a short day (8 h light/ 16 h dark) at 22 &#176;C for four weeks.</li>
		<li>Transfer plants to continuous light, at 22 &#176;C to induce the transition from the vegetative to the reproductive stage and for imaging the inflorescence SAMs.</li>
	</ol>
	</li>
	<li>Tomato and soybean growth
	<ol>
		<li>Incubate the seeds covered with wet filter paper under 28 &#176;C till they germinate.</li>
		<li>Transplant seedlings to soil and grow them in a long day (16 h light/ 8 h dark) at 25 &#176;C for one week or longer for imaging the vegetative SAMs.</li>
	</ol>
	</li>
</ol>
3. Dissection of the shoot apex
<ol>
	<li>Dissection of the inflorescence shoot apex
	<ol>
		<li>Cut the inflorescence shoot apex together with 1-2 cm main stem from the bolted Arabidopsis plants with a razor blade. Hold the basal part of the main stem and remove as many older flower organs as possible from the main stem by dissecting out the peduncles with jewelry forceps. 
		CAUTION: Avoid cutting fingers when using a razor blade. Dispose of the used razor blades to a proper sharps&#8217; container.</li>
		<li>Hold the attached stem of the shoot apex with jewelry&#160;forceps or fingers in the field of the stereomicroscope, continue removing the rest of flowers till nearly the whole SAM can be viewed from the eyepieces. Remove peduncles clearly at the junction of the main stem.</li>
	</ol>
	</li>
	<li>Dissection of the vegetative shoot apex
	<ol>
		<li>To view the vegetative SAMs from either tomato or soybean, dissect out the cotyledons, leaves, and roots from the plants.</li>
		<li>Hold the hypocotyls of the plants under the stereomicroscope and further dissect out the leaf primordia covering the vegetative SAMs using jewelry forceps.</li>
	</ol>
	</li>
</ol>
4. Staining
<ol>
	<li>To directly visualize cells in SAMs, use freshly prepared propidium iodide (PI) solution to stain cell walls. Dissolve PI powder in sterile, deionized water, make 1mL PI staining solution at the concentration of 1 mg/mL and store the PI solution in microcentrifuge tube covered with aluminum foil.</li>
	<li>Pipette 50 &#181;L PI solution in a clean and empty Petri dish and dip the whole dissected shoot apex into dye for 2 min. Rinse the stained shoot apex twice in sterile, deionized water. During the staining process, immerse the whole inflorescence SAM or vegetative SAM into the PI solution to achieve the uniform staining.</li>
</ol>
5. Image collection
NOTE: For this method, all the SAMs are imaged using an upright confocal microscope and with a 20x water-dipping lens. As described in other protocols9,12,13,15, it is also feasible to image SAMs using an inverted microscope. In addition, the live imaging can be achieved using different brands of confocal microscopes, with the same sample preparation steps. In this study, the imaging steps are described in detail as an example.
<ol>
	<li>Pierce a hole at the center of an imaging dish using forceps and stick the stained shoot apex upright in the medium.</li>
	<li>Fill the imaging dish with sterile, deionized water to completely immerse the sample. Viewing from the stereo microscope, pipette up and down to remove air bubbles trapped around the meristem. Then, adjust the angle of the stem in the agar to make sure that the SAM is fully visible from directly above.</li>
	<li>Place the imaging dish on the sample stage of the confocal microscope. Lower water-dipping lens and raise the microscope sample stage to let the tip of lens dip into the water.</li>
	<li>Open the confocal microscope software and locate the SAM in the brightfield in the eyepieces. Move the SAM sample right below the objective lens through adjusting the XY controller, then focus on the SAM from eyepieces through cautiously adjusting the Z controller.</li>
	<li>Operate the acquisition function in the confocal microscope software (see Table of Materials), start the Live mode to view the sample from the computer screen, and set up all parameters for the laser scanning experiment.
	<ol>
		<li>When adjusting the parameters, use Range Indicator function to define whether the signal is saturated or not.</li>
		<li>Optionally, apply the Reuse function to reload all the parameter settings from the selected confocal file. 
		NOTE: Suggested imaging parameters: Laser line (excitation): 515 nm or 561 nm; Emission 570-650 nm; pinhole: 1 airy unit (AU); Gain: 600-750, scan mode: Frame; Frame size: either 512 X 512 or 1024 X 1024; scanning speed: from 7 to Max; Scanning direction: bi-direction; Averaging number: 2-4; Averaging method: mean; Bit depth: 16 Bit; Scanning Interval: 0.5-1 &#181;M. Further, optimize all these parameters based on the nature of different plant samples and the specific imaging needs.</li>
	</ol>
	</li>
</ol>
6. Image processing
<ol>
	<li>For visualizing optical orthogonal and transverse section views, use the same commercial software for the imaging acquisition. Open the original confocal file, click ortho menu, select ortho. Then select either x position, y position and z position of the image, and save the images as the tiff files.
	<ol>
		<li>Optionally select 3D distance function to define the physical distance between two points that have been selected from the stack of the confocal images.</li>
		<li>Alternatively, use Fiji/Image J, the open resource image processing package to visualize the orthogonal and transverse section views.</li>
		<li>Open the original confocal file with Fiji, click image menu, select Stacks and then select Orthogonal views.</li>
		<li>Select XY, YZ, and XZ planes in the middle position and save as Tiff format images.</li>
	</ol>
	</li>
	<li>For visualizing a 3D transparent projection, use the same software.
	<ol>
		<li>Open the original confocal file, click 3D menu, and select Transparent to generate a 3D projection view.</li>
		<li>Optionally click 3D menu, select Appearance, and then select Transparency to adjust three parameters of the projection including Threshold, Ramp and Maximum for the transparency of the 3D image.</li>
		<li>Click 3D menu, select Appearance, and select Light to adjust the brightness of the 3D image.</li>
		<li>Export the projected images and save them as the Tiff files.</li>
	</ol>
	</li>
	<li>For visualizing a 3D maximum intensity projection, open the confocal files with the same software, and click the 3D menu.
	<ol>
		<li>Select 3D menu and select Maximum.</li>
		<li>Alternatively, use Fiji/Image J to visualize a 3D maximum intensity projection.</li>
		<li>Open the original confocal file with Fiji, click Image menu and select stacks.</li>
		<li>Select 3D project and save as Tiff format images.</li>
	</ol>
	</li>
	<li>For visualizing the depth coding view of the 3D images, use the same software.
	<ol>
		<li>Click 3D menu and select Appearance.</li>
		<li>Select Special and select Depth Coding.</li>
		<li>Alternatively, use Fiji/Image J to visualize the depth coding view.</li>
		<li>Open the confocal files, click image menu, select Hyperstack.</li>
		<li>Select Temporal-Color code and save as Tiff format images. 
		NOTE: The depth coding z-stack also can be achieved through the plugin Z Code Stack for Fiji.</li>
	</ol>
	</li>
	<li>For visualizing the 3D rotation view as shown (Movie 1 and Movie 2), use the same software (See the Table of Materials).
	<ol>
		<li>Open the confocal files, click 3D menu, and select Series.</li>
		<li>Select Render series, and select one of the four options including Turn around x, Turn around y, Start and end, and Position list.</li>
		<li>Save the render series as the AVI files.</li>
		<li>Alternatively, use Fiji/Image J to visualize the 3D rotation view.</li>
		<li>Open the original confocal file with Fiji, click Image menu, and select Stacks.</li>
		<li>Select 3D project and save as AVI format videos.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a simple imaging method that can be applied to the study of shoot apical meristems from different plants with minor modification, opening a new avenue to study the meristem regulation at both vegetative and reproductive stages in model plants and crops. In contrast to the SEM and histological staining methods, this protocol can help reveal both surface view and internal cellular structures of the SAMs, without the need for labor-intensive sample fixation and/or tissue sectioning steps. This protocol is ...

The plant meristem contains a pool of undifferentiated stem cells and continuously sustains the plant organ growth and development1. During the postembryonic development, almost all aboveground tissues of a plant are derived from the shoot apical meristem (SAM). In crops, the activity and size of the SAM and its derived floral meristems are tightly associated with many agronomic traits such as shoot architecture, fruit production, and seed yield. For example, in tomato, an enlarged SAM causes an increase in the shoot and inflorescence branching, and thus results in generating extra flower and fruit organs2. In maize, an increase in SAM size leads to a higher seed number and total yield3,4. In soybean, the meristem indeterminacy is also closely associated with the shoot architecture and yield5.
The morphology and anatomy of SAMs can be characterized by several different methods, including histological sectioning/staining and scanning electron microscopy (SEM)6, both of which have greatly advanced the meristem research through providing either the sectional view or a three-dimensional (3D) surface view of SAMs. However, both methods are time consuming, involving several experimental steps from the sample preparation to the data acquisition, and these methods mainly depend on fixed samples. Recent advances in laser scanning confocal microscopy technique have overcome these limitations and provide us with a powerful tool to investigate the cellular structure and developmental process of plant tissues and organs7,8. Through optical rather than physical tissue sectioning, confocal microscopy allows the collection of a series of z-stack images and the subsequent 3D reconstruction of the sample through image analysis software.
Here, we describe an efficient procedure for investigating both inside and surface structures of the living SAMs from different plant species using laser scanning confocal microscopy, which potentially allows researchers to accomplish all the experimental process within as short as 20 minutes. Different from other published methods for live confocal imaging of Arabidopsis inflorescence SAMs9,10,11,12,13,14,15 and Arabidopsis flowers12,13, here we demonstrate that this protocol is highly efficient for studying not only the inflorescence meristems of the model species but also the vegetative shoot apical meristems from different crops, such as tomato and soybean. This method does not rely on transgenic fluorescent markers, and potentially can be applied to study the shoot meristems from many different species and cultivars. In addition, we also introduce the simple image processing steps for viewing and analyzing different SAMs in a&#160;3D view. Taken together, this simple method will facilitate researchers better understanding both the structure and developmental process of the meristems from both model organisms and crops.

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	<tbody>
		<tr height="32">
			<td height="32" style="height:32px;width:283px;">Agar Phyto</td>
			<td style="width:176px;">Dot Scientific Inc.</td>
			<td style="width:119px;">DSA20300-1000</td>
			<td style="width:391px;"></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:35px;width:283px;">Agarose</td>
			<td style="width:176px;">Dot Scientific Inc.</td>
			<td style="width:119px;">AGLE-500</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:283px;">Forceps&#160;</td>
			<td style="width:176px;">ROBOZ</td>
			<td style="width:119px;">RS-4955</td>
			<td>Dumont #5SF Super Fine Forceps Inox Tip Size .025 X .005mm, for dissecting shoot apices.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:283px;">LSM 880 Upright Confocal Microscope&#160;</td>
			<td style="width:176px;">Zeiss</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Murashige &#38; Skoog MS medium</td>
			<td style="width:176px;">Dot Scientific Inc.</td>
			<td style="width:119px;">DSM10200-50</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Plan APO 20x/1.1 water dipping lens</td>
			<td style="width:176px;">Zeiss</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:283px;">Plastic petri dishes 100 mm X 15 mm</td>
			<td style="width:176px;">CELLTREAT Scientific Products</td>
			<td style="width:119px;">229694</td>
			<td style="width:391px;">Use as making MS plates</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Plastic petri dishes60 mm X 15</td>
			<td style="width:176px;">CELLTREAT Scientific Products</td>
			<td style="width:119px;">229665</td>
			<td>Use as imaging dishes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Propagation Mix</td>
			<td style="width:176px;">Sungro Horticulture</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Propidium iodide</td>
			<td style="width:176px;">Acros Organics</td>
			<td style="width:119px;">440300250</td>
			<td>1 mg/mL solution in water, to stain the cell walls</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Razor blade</td>
			<td style="width:176px;">PERSONNA&#160;</td>
			<td style="width:119px;">62-0179</td>
			<td style="width:391px;">For cutting shoot apex from plants</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:35px;width:283px;">Stereomicroscope</td>
			<td style="width:176px;">Nikon</td>
			<td style="width:119px;">SMZ1000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Tissue</td>
			<td style="width:176px;">VWR</td>
			<td style="width:119px;">82003-820</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Zen black</td>
			<td style="width:176px;">Zeiss</td>
			<td style="width:119px;"></td>
			<td>Image acquisition software</td>
		</tr>
	</tbody>
</table>

confocal live imaging of shoot apical meristems from different plant species

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic development. The activity and morphology of SAMs determine important agronomic traits, such as shoot architecture, size and number of reproductive organs, and most importantly, grain yield. Here, we provide a detailed protocol for analyzing both the surface morphology and the internal cellular structure of the living SAMs from different species through laser scanning confocal microscope. The whole procedure from the sample preparation to the acquisition of high resolution three-dimensional (3D) images can be accomplished within as short as 20 minutes. We demonstrate that this protocol is highly efficient for studying not only the inflorescence SAMs of the model species but also the vegetative meristems from different crops, providing a simple but powerful tool to study the organization and development of meristems across different plant species.

To evaluate the efficiency of our protocol and to explore the morphology of the meristems from different species, we have performed the confocal live imaging experiments on the inflorescence meristem from Arabidopsis and the vegetative meristems from both tomato and soybean. In this study, Arabidopsis ecotype Landsberg erecta, tomato cultivar Micro-Tom and soybean cultivar Williams 82 have been used as examples.
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Watch this Scientific Journal Video about Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species at JoVE.com

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

This protocol presents how to live image and analyze the shoot apical meristems from different plant species using laser scanning confocal microscopy.

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic ...

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12.1K Views.  Purdue University. This protocol presents how to live image and analyze the shoot apical meristems from different plant species using laser scanning confocal microscopy.

Video: Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

In Vitro Differentiation of Mature Myofibers for Live Imaging

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily conserved structures that compose it are assembled remains under investigation. Their size and physiological features often constrain manipulation and imaging applications. The culture of immortalized cell lines is widely used, but it can only replicate the early steps of differentiation.
Here, we describe a protocol that enables easy genetic manipulation of myofibers originating from primary mouse myoblasts. After one week of differentiation, the myofibers display contractility, aligned sarcomeres and triads, as well as peripheral nuclei. The entire differentiation process can be followed by live imaging or immunofluorescence. This system combines the advantages of the existing ex vivo and in vitro protocols. The possibility of easy and efficient transfection as well as the ease of access to all differentiation stages broadens the potential applications. Myofibers can subsequently be used not only to address relevant developmental and cell biology questions, but also to reproduce muscle disease phenotypes for clinical applications.

In Vitro Differentiation of Mature Myofibers for Live Imaging

Muscle cells are among the most complex eukaryotic cells. We present a protocol for the in vitro differentiation of highly mature myofibers that allows for genetic manipulation and clear imaging during all developmental stages.

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily ...

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in molecular and imaging technologies allowed us to gain insights into how biochemical signals influence morphogenetic events. However, it is increasingly evident that apart from biochemical signals, mechanical cues also impact several aspects of cell and tissue growth. The Arabidopsis shoot apical meristem (SAM) is a dome-shaped structure responsible for the generation of all aboveground organs. The organization of the cortical microtubule cytoskeleton that mediates apoplastic cellulose deposition in plant cells is spatially distinct. Visualization and quantitative assessment of patterns of cortical microtubules are necessary for understanding the biophysical nature of cells at the SAM, as cellulose is the stiffest component of the plant cell wall. The stereotypical form of cortical microtubule organization is also a consequence of tissue-wide physical forces existing at the SAM. Perturbation of these physical forces and subsequent monitoring of cortical microtubule organization allows for the identification of candidate proteins involved in mediating mechano-perception and transduction. Here we describe a protocol that helps investigate such processes.

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Here we describe a protocol for live cell imaging of the cortical microtubule cytoskeleton at the shoot apical meristem and monitoring its response to changes in physical forces.

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in ...

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Subtitles

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species