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/pubmed?term=((Dynamic+construction%2C+perception%2C+and+remodeling+of+plant+cell+walls%5BTitle%5D)+AND+%22Annual+Review+of+Plant+Biology%22%5BJournal%5D)">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/pubmed?term=((Cell+wall+integrity+maintenance+during+plant+development+and+interaction+with+the+environment%5BTitle%5D)+AND+%22Naure+Plants%22%5BJournal%5D)">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/pubmed?term=((The+cellular+function+of+ROP+GTPase+prenylation+important+for+multicellularity+in+the+moss+Physcomitrium+patens%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">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/pubmed?term=((Imaging+the+living+plant+cell%3A+From+probes+to+quantification%5BTitle%5D)+AND+%22The+Plant+Cell%22%5BJournal%5D)">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. Live cell imaging of plants. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aFang%2C+Y+%22Cold+Spring+Harbor+Protocols%22">Cold Spring Harbor Protocols</a>. 2010 (2), (2010).</li>
<li>Goh, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+live-cell+imaging+approaches+to+study+lateral+root+formation+in+Arabidopsis+thaliana%5BTitle%5D)+AND+%22Microscopy%22%5BJournal%5D)">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/pubmed?term=((Time-lapse+imaging+of+developing+shoot+meristems+using+a+confocal+laser+scanning+microscope%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">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/pubmed?term=((Live+cell+imaging+of+FM4-64%2C+a+tool+for+tracing+the+endocytic+pathways+in+Arabidopsis+root+cells%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">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/pubmed?term=((Advances+in+synthetic+fluorescent+probe+labeling+for+live-cell+imaging+in+plants%5BTitle%5D)+AND+%22Plant+%26+Cell+Physiology%22%5BJournal%5D)">Advances in synthetic fluorescent probe labeling for live-cell imaging in plants.</a> Plant & Cell Physiology. 62 (8), 1259-1268 (2021).</li>
<li>Whitewoods, C. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CLAVATA+was+a+genetic+novelty+for+the+morphological+innovation+of+3D+growth+in+land+plants%5BTitle%5D)+AND+%22Current+Biology%22%5BJournal%5D)">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/pubmed?term=((A+PSTAIRE-type+cyclin-dependent+kinase+controls+light+responses+in+land+plants%5BTitle%5D)+AND+%22Science+Advances%22%5BJournal%5D)">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 expansion/branching sites in wild type, allowing for prediction of the expansion/branching sites. Further, because calcofluor white can be used with other dyes, such as microspheres, additional information about cell expansion can be observed3. 
There are two reasons that could explain how the calcofluor dye can bind for days to plants without affecting/damaging the molecular target. First, the low working concentration (10 &#181;L in 200 &#181;L of BCDATG medium, or 5%) may be sufficiently low for the moss protonemata to not be affected. Second, the calcofluor may stain the mature stage of cellulose, because the staining signal is much stronger in the basal part of wild type (which is older tissue) than the apical region of the apical cell (which is younger tissue), and the broken cell wall surface of ggb mutants (which is older cellulose) is stronger than the cells (which is younger cellulose) beneath the broken cell wall surface.
This protocol has been used for moss tissue, which is simple in structure. The current protocol may also be applied to other systems with simple structures, such as root hairs and trichomes in flowering plants. However, it remains to be seen if the calcofluor white staining works for other systems with more cell layers, such as stems. Moreover, because the calcofluor white is mixed with the medium, the tissue to be stained must be immersed within the medium. Therefore, optimization may be required in order to apply this protocol to other systems.

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><tbody><tr><td>3-mm-thick red plastic light filter</td><td>&amp;nbsp;Mitsubishi</td><td>&amp;nbsp;no.102</td><td></td></tr><tr><td>27 mm diameter glass base dish&amp;nbsp;</td><td>Iwaki</td><td>3930-035</td><td></td></tr><tr><td>Agar</td><td>&amp;nbsp;Sigma</td><td>A6924</td><td></td></tr><tr><td>Calcofluor white&amp;nbsp;</td><td>&amp;nbsp;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&amp;nbsp;</td><td>Nikon&amp;nbsp;</td><td>A1</td><td> &lt;p&gt;NIS element software; .nd2 file in NIS-elements Viewer, download from https://www.microscope.healthcare.nikon.&lt;br/&gt; com/en_AOM/products/software/nis-elements/viewer&lt;/p&gt; </td></tr><tr><td>Fluorescence microscope&amp;nbsp;</td><td>Nikon&amp;nbsp;</td><td>TE200&amp;nbsp;</td><td>Equipped with a DS-U3 camera;</td></tr><tr><td>Gellan gum&amp;nbsp;</td><td>Nacali Tesque</td><td>12389-96</td><td></td></tr><tr><td>Plant Growth Chambers</td><td>SANYO&amp;nbsp;</td><td>Sanyo MLR-350H</td><td></td></tr><tr><td>Sterilized syringe 0.22 &amp;mu;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 expansion3 (Figure 1B). Moreover, the staining signal of the broken surface of ggb mutants (which is older cellulose) is stronger than the cell surface (which is younger cellulose) beneath the broken cell wall surface.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64651/64651fig01.jpg" /> 
Figure 1: 3-D time series of moss development. (A,B) Dynamic changes in cross-walls were shown by calcofluor white staining (which stains cellulose) in WT (A) and ggb mutants (B). For ggb mutants (B), white dash lines below each image indicate the broken surface cell walls on the expanding cells, and orange lines indicate the cells beneath the cell wall surface in ggb. Panel B is reprinted with permission from3. Note that the differences in calcofluor white signal can be detected on the surface positions of the cells, and less dense staining regions (arrowheads) are correlated to the cell expansion/branching sites. Scale bars = 20 &#181;m. Abbreviations: d = days; WT = wild type; ggb = geranylgeranyltransferase-I beta. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64651/64651fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64651/64651fig02.jpg" /> 
Figure 2: Marking location of plants. On the bottom of the dish, nine lines each were drawn in parallel and perpendicular directions to mark the location of squares. Characters a to h were used for marking rows, and numbers 1 to 8 were used for marking columns. Upper, lower, right, middle, and left were used for marking positions within each square. For example, if a plant (denoted by a green dot) was located in a square at the second row and the sixth column, and positioned in the upper left part, then the plant was named b6_upper_left. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64651/64651fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64651/64651fig03.jpg" /> 
Figure 3: Reconstruction of 3-D images using a nd2 z-stack file. (A)&#160;Open an nd2 file in the NIS-elements Viewer. Click LUTs (circle) to adjust the contrast and brightness of images and click Volume (square) to reconstruct the 3-D image. (B) Reconstructed 3-D image from A. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64651/64651fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: List of solutions used in this protocol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64651/64651_R1_Table 1.xlsx" target="_blank">Please click here to download this Table.</a>

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

Nanyang Technological University

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Developmental Biology

1.5K Views.  University of Louisville. Time-lapse beeldvorming met fluorescentiemicroscopie maakt observatie van de dynamische veranderingen van groei en ontwikkeling op cellulair en subcellulair niveau mogelijk. Dit eenvoudige protocol voor 3D time-lapse beeldvorming kan de celwanddynamiek over een lange periode direct bestuderen. Dit protocol maakt gebruik van calcofluorkleurstof om de dynamiek van de celwanddikte te markeren en vereist daarom geen transformatie van een fluorescerend eiwit.De calcofluorsignalen zijn stabi...

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

Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls are composed of cellulose and non-cellulosic polysaccharides together with proteins, phenolic compounds and water. 90% of the cell wall is composed of&#160;polysaccharides (e.g., pectins) and arabinogalactan proteins (AGPs). The fluorescent immunolocalization of specific glycan epitopes in plant histological sections remains a key tool to uncover remodeling of wall polysaccharide networks, structure and components.
Here, we report an optimized fluorescent immunolocalization procedure to detect glycan epitopes from AGPs and pectins in plant tissues. Paraformaldehyde/glutaraldehyde fixation was used along with LR-White embedding of the plant samples, allowing for a better preservation of the tissue structure and composition. Thin sections of the embedded samples obtained with an ultra-microtome were used for immunolocalization with specific antibodies. This technique offers great resolution, high specificity, and the chance to detect multiple glycan epitopes in the same sample. This technique allows subcellular localization of glycans and detects their level of accumulation in the cell wall. It also permits the determination of spatio-temporal patterns of AGP and pectin distribution during developmental processes. The use of this tool may ultimately guide research directions and link glycans to specific functions in plants. Furthermore, the information obtained can complement biochemical and gene expression studies.

This protocol describes in detail how the plant material for immunolocalization of Arabinogalactan proteins and pectins is fixed, embedded in a hydrophilic acrylic resin, sectioned and mounted on glass slides. We show cell wall related epitopes will be detected with specific antibodies.

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls ...

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of plant leaves or vertebrate lungs further difficulties arise from multiple transitions in refractive index between cellular components, between cells and airspaces and between the biological tissue and the rest of the optical system. Moreover, refractive index mismatches lead to attenuation of fluorophore excitation and signal emission in fluorescence microscopy. We describe here the application of the perfluorocarbon, perfluorodecalin (PFD), as an infiltrative imaging medium which optically improves laser scanning confocal microscopy (LSCM) sample imaging at depth, without resorting to damaging increases in laser power and has minimal physiological impact2. We describe the protocol for use of PFD with Arabidopsis thaliana leaf tissue, which is optically complex as a result of its structure (Figure 1). PFD has a number of attributes that make it suitable for this use3. The refractive index of PFD (1.313) is comparable with that of water (1.333) and is closer to that of cytosol (approx. 1.4) than air (1.000). In addition, PFD is readily available, non-fluorescent and is non-toxic. The low surface tension of PFD (19 dynes cm-1) is lower than that of water (72 dynes cm-1) and also below the limit (25 - 30 dyne cm-1) for stomatal penetration4, which allows it to flood the spongy mesophyll airspaces without the application of a potentially destructive vacuum or surfactant. Finally and crucially, PFD has a great capacity for dissolving CO2 and O2, which allows gas exchange to be maintained in the flooded tissue, thus minimizing the physiological impact on the sample. These properties have been used in various applications which include partial liquid breathing and lung inflation5,6, surgery7, artificial blood8, oxygenation of growth media9, and studies of ice crystal formation in plants10. Currently, it is common to mount tissue in water or aqueous buffer for live confocal imaging. We consider that the use of PFD as a mounting medium represents an improvement on existing practice and allows the simple preparation of live whole leaf samples for imaging.

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

We describe the use of perfluorodecalin as an infiltrative mounting medium. This is a simple method for improving depth of imaging in Arabidopsis thaliana leaf tissue with minimal physiological impact.

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of ...

Biology

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 ...

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ binding proteins that initiate Ca2+ signaling cascades. However, we still know little about how stimulus specific Ca2+ signals are generated. The specificity of a Ca2+ signal may be attributed to the sophisticated regulation of the activities of Ca2+ channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+ signals at both the tissue and cellular levels. Genetically encoded Ca2+ indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+ signals. Here we describe instructions for the use of two Ca2+ detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+ imaging and case12 based live cell confocal fluorescence Ca2+ imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+ signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+ signals at a high resolution.

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Ca2+ signaling regulates diverse biological processes in plants. Here we present approaches for monitoring abiotic stress induced spatial and temporal Ca2+ signals in Arabidopsis cells and tissues using the genetically encoded Ca2+ indicators Aequorin or Case12.

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

A Strategy to Validate the Role of Callose-mediated Plasmodesmal Gating in the Tropic Response

The plant hormone auxin plays an important role in many growth and developmental processes, including tropic responses to light and gravity. The establishment of an auxin gradient is a key event leading to phototropism and gravitropism. Previously, polar auxin transport (PAT) was shown to establish an auxin gradient in different cellular domains of plants. However, Han et al. recently demonstrated that for proper auxin gradient formation, plasmodesmal callose-mediated symplasmic connectivity between the adjacent cells is also a critical factor. In this manuscript, the strategy to elucidate the role of particular genes, which can affect phototropism and gravitropism by altering the symplasmic connectivity through modulating plasmodesmal callose synthesis, is discussed. The first step is to screen aberrant tropic responses from 3-day-old etiolated seedlings of mutants or over-expression lines of a gene along with the wild type. This preliminary screening can lead to the identification of a range of genes functioning in PAT or controlling symplasmic connectivity. The second screening involves the sorting of candidates that show altered tropic responses by affecting symplasmic connectivity. To address such candidates, the movement of a symplasmic tracer and the deposition of plasmodesmal callose were examined. This strategy would be useful to explore new candidate genes that can regulate symplasmic connectivity directly or indirectly during tropic responses and other developmental processes.

This article describes the methods for screening the genes controlling plasmodesmal permeability and hence auxin gradient during tropic response. This includes the measurement of the degree of tropic response in hypocotyl of Arabidopsis thaliana and checking plasmodesmal permeability by 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) loading and finally callose level assessment.

The plant hormone auxin plays an important role in many growth and developmental processes, including tropic responses to light and gravity. The ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

Application of Membrane and Cell Wall Selective Fluorescent Dyes for Live-Cell Imaging of Filamentous Fungi

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two decades ago and since then continues to contribute greatly to our understanding of the filamentous fungal lifestyle. This paper provides a practical guide for the utilization of the two membrane dyes FM 1-43 and FM 4-64 and the four cell wall stains Calcofluor White M2R, Solophenyl Flavine 7GFE 500, Pontamine Fast Scarlet 48 and Congo Red. The focus is on their low-dose application to ascertain artefact-free staining, their co-imaging properties, and their quantitative evaluation. The presented methods are applicable to all filamentous fungal samples that can be prepared in the described ways. The fundamental staining approaches can serve as starting points for adaptations to species that might require different cultivation conditions. First, biophysical and biochemical properties are reviewed as their understanding is essential for using these dyes as truly vital fluorescent stains. Secondly, step-by-step protocols are presented that detail the preparation of various fungal sample types for fluorescent live-cell imaging. Finally, example experiments illustrate different approaches to: (1) identify defects in the spatio-temporal organization of endocytosis in genetic mutants, (2) comparatively characterize shared and distinct co-localization of GFP-labeled target proteins in the endocytic pathway, (3) identify morphogenetic cell wall defects in a genetic mutant, and (4) monitor cell wall biogenesis in real time.

Vital fluorescent dyes are essential tools for live-cell imaging analyses in modern fungal cell biology. This paper details the application of established and lesser-known fluorescent dyes for tracking plasma membrane dynamics, endo-/exocytosis and cell wall morphogenesis in filamentous fungi.

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two ...

Experimental Screening Protocols, Immunocytochemistry and Microscopy-based Imaging Techniques for Penium margaritaceum

The cell wall is the first component of signal reception/transduction for a plant cell during development and when responding to environmental abiotic and biotic stressors. The cell constantly monitors the integrity of its cell wall and modulates it in response to stress. Elucidating the specific structural and biochemical modulations occurring in the cell wall is a difficult task especially when employing multicellular plants and their organs/tissues. This is due to limits as to what can be resolved in an individual cell that is part of a complex multicellular network. The unicellular streptophyte alga, Penium margaritaceum, has recently been used in investigations of pectin dynamics, cell wall-based phenotypic plasticity and multiple aspects of algal cell biology. Its simple phenotype, distinct cell wall that has many components notably similar to land plant cell walls, and ease in immunocytochemical and experimental studies make it a powerful model organism in plant cell wall biology. The goal of this study is to provide the basic techniques for culturing, experimental manipulation and screening of applied stressors. Screening protocols for immunocytochemistry, confocal laser scanning microscopy imaging and scanning electron microscopy imaging&#160;of cell wall structure. Likewise, many of the described techniques may be modified for a wide array of other cell and molecular studies.

This report describes the fundamental methods used to culture and experimentally manipulate the unicellular streptophyte alga, Penium margaritaceum. It also provides fundamental protocols of microscopy-based imaging, including live cell labeling with monoclonal antibodies and other fluorescent probes and scanning electron microscopy.

The cell wall is the first component of signal reception/transduction for a plant cell during development and when responding to environmental abiotic and ...

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

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