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.

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

confocal-live-imaging-shoot-apical-meristems-from-different-plant

Universidad de Cartagena UDEC

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Developmental Biology

12.2K 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 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. ...

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the flexible and efficient GreenGate cloning system with the proven and benchmarked LhGR system (here termed GR-LhG4) for the inducible expression, we have generated a set of transgenic Arabidopsis lines that can drive the expression of an effector cassette in a range of specific cell types in the three main plant meristems. To this end, we chose the previously developed GR-LhG4 system based on a chimeric transcription factor and a cognate pOp-type promoter ensuring tight control over a wide range of expression levels. In addition, to visualize the expression domain where the synthetic transcription factor is active, an ER-localized mTurquoise2 fluorescent reporter under control of the pOp4 or pOp6 promoter is encoded in driver lines. Here, we describe the steps necessary to generate a driver or effector line and demonstrate how cell type specific expression can be induced and followed in the shoot apical meristem, the root apical meristem and the cambium of Arabidopsis. By using several or all driver lines, the context specific effect of expressing one or multiple factors (effectors) under control of the synthetic pOp promoter can be assessed rapidly, for example in F1 plants of a cross between one effector and multiple driver lines. This approach is exemplified by the ectopic expression of VND7, a NAC transcription factor capable of inducing ectopic secondary cell wall deposition in a cell autonomous manner.

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Here, we describe the generation and application of a set of transgenic Arabidopsis thaliana lines enabling inducible, tissue-specific expression in the three main meristems, the shoot apical meristem, the root apical meristem, and the cambium.

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the ...

Genetics

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 ...

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 ...

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.

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.

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

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Biology

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is regulated spatially and temporally. The use of a fluorescent gene under the control of an immunity-related promoter in combination with an automated fluorescence microscopy is a simple way to understand spatiotemporal regulation of plant immunity. In contrast to the root tissues that have been used for a number of various intravital fluorescent imaging experiments, there exist few fluorescent live-imaging examples for the leaf tissues that encounter an array of airborne microbial infections. Therefore, we developed a simple method to mount leaves of Arabidopsis thaliana plants for live-cell imaging over an extended period of time. We used transgenic Arabidopsis plants expressing the yellow fluorescent protein (YFP) genefused to the nuclear localization signal (NLS) under the control of the promoter of a defense-related marker gene, Pathogenesis-Related 1 (PR1). We infiltrated a transgenic leaf with Pseudomonas syringae pv. tomato DC3000 (avrRpt2) strain (Pst_a2) and performed in vivo time-lapse imaging of the YFP signal for a total of 40 h using an automated fluorescence stereomicroscope. This method can be utilized not only for studies on plant immune responses but also for analyses of various developmental events and environmental responses occurring in leaf tissues.

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

We report a simple and versatile method for performing fluorescent live-imaging of Arabidopsis thaliana leaves over an extended period of time. We use a transgenic Arabidopsis plant expressing a fluorescent reporter gene under the control of an immunity-related promoter as an example for gaining spatiotemporal understanding of plant immune responses.

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is ...

Deep Fluorescence Observation in Rice Shoots via Clearing Technology

The recently developed clearing technology that eliminates refractive index mismatches and diminishes auto-fluorescent material has made it possible to observe plant tissues in three dimensions (3D) while preserving their internal structures. In rice (Oryza sativa L.), a monocot model plant and a globally important crop, clearing technology has been reported in organs that are relatively easy to observe, such as the roots and leaves. Applications of clearing technology in shoot apical meristem (SAM) and stems have also been reported, but only to a limited degree because of the poor penetration of the clearing solution (CS) in these tissues. The limited efficiency of the clearing solutions in these tissues has been attributed to auto-fluorescence, thickening, and hardening of the tissues in the stem as the vascular bundles and epidermis develop and layering of the SAM with water-repellent leaves. The present protocol reports the optimization of a clearing approach for continuous and 3D observation of gene expression from the SAM/young panicle to the base of the shoots during development. Fixed tissue samples expressing a fluorescent protein reporter were trimmed into sections using a vibrating micro-slicer. When an appropriate thickness was achieved, the CS was applied. By specifically targeting the central tissue, the penetration rate and uniformity of the CS increased, and the time required to make the tissue transparent decreased. Additionally, clearing of the trimmed sections enabled the observation of the internal structure of the whole shoot from a macro perspective. This method has potential applications in deep imaging of tissues of other plant species that are difficult to clear.

Deep Fluorescence Observation in Rice Shoots via Clearing Technology

The present protocol describes a clearing technique for rice shoots, which are difficult to prepare for internal structural observations owing to the hard, thick, or layered nature of the tissues. This method facilitates continuous and deep fluorescence observations, even in adult rice plants.

The recently developed clearing technology that eliminates refractive index mismatches and diminishes auto-fluorescent material has made it possible to ...

Improved Methods for Imaging Maize Leaf Primordia

Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study early leaf development. Here, we describe methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging. The first method uses a wire stripper to remove the upper portions of older leaves, exposing the tip of the leaf primordium and allowing its measurement for more accurate transverse section sampling. The second method uses clear, double-sided nano tape to unroll and mount whole-leaf primordia for imaging. We show the utility of the two methods in visualizing and analyzing fluorescent protein reporters in maize. These methods provide a solution to the challenges presented by the distinctive morphology of maize leaf primordia and will be useful for visualizing and quantifying leaf anatomical and developmental traits in maize and other grass species.

Author Spotlight: Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging  

Maize leaf primordia are deeply ensheathed and rolled, making them difficult to study. Here, we present methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging.

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study ...

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

Summary

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

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

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

Deep Fluorescence Observation in Rice Shoots via Clearing Technology

Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging