The author wish to thank Prof. Elliot M. Meyerowitz for his support, and comments on the manuscript, as well as Ann Lavanway at Dartmouth College and Dr. Andres Collazo at the Biological Imaging Facility at Caltech for their help in solving technical issues with the live confocal imaging. Nathana&#235;l Prunet&#39;s work is supported by the US National Institutes of Health through Grant R01 GM104244 to Elliot M. Meyerowitz.

1. Media and Dishes Preparation
<ol>
	<li>Prepare dissecting dishes by filling round plastic boxes (approximately 6 cm wide, 2 cm deep) to 0.5 cm with 2% agarose.</li>
	<li>Prepare imaging dishes.
	<ol>
		<li>For an upright confocal microscope, fill rectangular plastic hinged boxes (approximately 7 cm long, 4.5 cm wide, 3 cm deep) to 0.5 cm with imaging medium (see section 1.3, &#34;Imaging medium&#34;; Figure 1F).</li>
		<li>For an inverted confocal microscope: fill small Petri dish (approximately 3.5 cm wide, 1 cm deep) exactly to the brim with imaging medium (see section 1.3, &#34;Imaging medium&#34;; Figure 1G).</li>
	</ol>
	</li>
	<li>Prepare imaging medium.
	<ol>
		<li>For single point imaging, use 1% agarose.</li>
		<li>For time-lapse experiments, use apex growth medium (0.5x Murashige and Skoog basal salt mixture without vitamin, 1% sucrose, 0.8% agarose, pH 5.8 with potassium hydroxide solution, supplemented with vitamin [0.01% myo-inositol, 0.0001% nicotinic acid, 0.0001% pyridoxine hydrochloride, 0.001% thiamine hydrochloride, 0.0002% glycine] and cytokinin [500 nM N6-benzyladenine])15.</li>
	</ol>
	</li>
</ol>
2. Plant Growth
<ol>
	<li>Sow sterilized seeds on MS plates (0.5 or 1x Murashige and Skoog basal salt mixture without vitamin, 0.8% agar, pH 5.8 with potassium hydroxide solution) with appropriate selection. Place plates in long day (16 h light) or continuous day, 16-22 &#176;C conditions for two weeks.</li>
	<li>Transplant seedlings onto soil with sufficient spacing to allow robust development. Place plants in short day, 16-22 &#176;C conditions for three weeks. 
	NOTE: Growing plants in long day or continuous day conditions directly after transplanting results in premature flowering and inflorescences that are less vigorous, and much harder to dissect. Similarly, leaving plants under short day conditions for more than three weeks results in day length-independent flowering and inflorescences that are less vigorous.</li>
	<li>Transfer plants to long day (16 h light) or continuous day, 16-22 &#176;C conditions until they flower. Shoot apices are easiest to dissect when the inflorescence is 2-10 cm long.</li>
</ol>
3. Dissection of the Shoot Apex
<ol>
	<li>Optional: Using a fine sharpening stone and a drop of light oil, sharpen forceps under the stereomicroscope to make them blade-like, not point-like.</li>
	<li>Remove siliques, older flower buds and secondary inflorescences from the primary inflorescence by pushing the base of the peduncles with the forceps until they break. Remove as many flowers as possible without magnification (Figure 1, compare A and B).</li>
	<li>Using forceps, pierce a vertical hole in the agarose of a dissecting dish, cut off the last 0.5 cm of the inflorescence and stick it vertically in the agarose. The remaining flower buds must be above the agarose surface.</li>
	<li>Fill the imaging dish with sterile, de-ionized water so that the shoot apex is fully immersed. Place the dissecting dish under the stereomicroscope, and remove the air trapped around the shoot apex by creating water jets with a 1,000 &#181;L pipette (Figure 1, compare C and D).</li>
	<li>Under the stereomicroscope, use the forceps to remove flower buds that are not to be imaged by pushing on the base of the peduncle or the top of the bud until the peduncle breaks (Figure 1E). It is important that the peduncles break cleanly at the junction with the stem, as leftover peduncles hinder the removal of the young flower buds.</li>
	<li>If imaging only flower buds stage 5 and younger (Figure 2A), remove the water before dissecting stage 6-8 flower buds. If imaging flower buds stage 5 and older, proceed to sepal ablation (see section 5.1). Otherwise, proceed to step 3.7.</li>
	<li>Using forceps, pierce a vertical hole in the medium of an imaging dish, and stick the dissected apex upright in the medium, so that only the shoot apical meristem and surrounding flower buds are above the surface of the medium. Depending on which flower bud(s) are to be imaged, it may be necessary to slightly tilt the sample, as flower buds are not necessarily oriented exactly like the stem (Figure 2A).</li>
	<li>Proceed to staining the sample if needed. If not staining and/or imaging the sample right away, add water and close the imaging dish to prevent dehydration. If using an inverted microscope, place the imaging dish in a transparent plastic box and close it.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/55156/55156fig1.jpg" /> 
Figure 1: Preparation of the shoot apices for live confocal imaging. (A-E) Dissection of a shoot apex for imaging. Inflorescence before (A) and after (B) removal of siliques and older flowers. (C-D) Shoot apex immersed in water in the dissecting dish, with an air bubble trapped at the tip (C) and after removal of the air bubble (D). (E) Shoot apex in the dissecting dish, after dissection of flower buds older than stage 5. (F-G) View of shoot apices in the imaging dish, on the stage of an upright (F) and inverted (G) confocal microscope. In (F), a 40X water-dipping lens is positioned above one of the apices, with the tip of the lens immersed in water. In (G), the shoot apex is positioned upside down above the 40X water-dipping lens, with a water column connecting the imaging medium to the tip of the lens. The smaller panel in (F) shows a higher magnification view of the area in the red rectangle, with a shoot apex inserted in imaging medium; red and blue lines indicate the surface of the medium and water, respectively. (H-I) Examples of custom-made devices that allow adding more water at the tip of the water-dipping lens: a silicon rubber sleeve made from a spark plug boot (H), and a makeshift sleeve made from the finger of a powderless latex glove (G). Scale bars = 0.5 cm in (A) and (B), 0.1 cm in (C) and (D), and 100 &#181;m in (E). This figure was initially published in reference14. <a href="http://cloudfront.jove.com/files/ftp_upload/55156/55156fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Staining
NOTE: Cell walls or plasma membranes can be stained with propidium iodide or FM4-64, respectively, to achieve cellular resolution during imaging. Alternatively, reporter lines with fluorescent proteins tagged to the plasma membrane can be used6,16.
<ol>
	<li>Remove water from the imaging dish, and ensure the surface of the medium is dry to avoid diluting the dye.</li>
	<li>Under the stereomicroscope, apply 20 to 30 &#181;L of either 1 mg/mL propidium iodide solution, or 80 &#181;g/mL FM4-64 solution to the dissected apex with a 10 &#181;L pipette. Make sure the whole sample is covered in dye to ensure a homogeneous staining.</li>
	<li>Stain for 2 min if using propidium iodide, or 20 min if using FM4-64.</li>
	<li>Rinse twice with sterile, deionized water.</li>
</ol>
5. Sepal Ablation
NOTE: At stage 4, sepal primordia start covering the FM. As they grow, they filter out the fluorescence from underlying tissue, and hinder the imaging process. Sepals can either be removed manually using a metal pin mounted on a pin-vise, or prevented to grow using laser ablation on emerging sepal primordia. Alternatively, it is possible in some cases to use mutants such as apetala1-1, in which sepals are missing or replaced by leaf-like organs that do not cover the FM while the central part of the flower develops normally (Figure 2F)17.
<ol>
	<li>Sepal ablation on flower buds stage 5 and older using a pin-vise holding a straight metal needle (Figure 2, E1-E2)
	<ol>
		<li>Place the dissecting dish with the dissected apex under the stereomicroscope, and set the magnification to maximum. Immerse the shoot apex in sterile, deionized water, or alternatively, use a 1,000 &#181;L pipette to apply water to the shoot apex regularly while dissecting the sepals.</li>
		<li>With the pin-vise, position the pin on top of the abaxial sepal, tangentially relative to the shoot apex. Gently push the sepal away from the shoot apex until it breaks away from the flower bud.</li>
		<li>Proceed similarly with the adaxial sepal, but push the sepal towards the shoot apex.</li>
		<li>Proceed similarly with the lateral sepals, but position the pin radially relative to the shoot apex, and push the sepals to the side, away from the flower bud.</li>
		<li>Immerse the shoot apex in sterile, deionized water for a few minutes to prevent dehydration.</li>
	</ol>
	</li>
	<li>Laser ablation of sepal primordia at stage 3-4 (Figure 2, C5-D5)
	<ol>
		<li>Place the imaging dish with the stained, dissected apex on the confocal microscope stage. Locate and focus on the apex as if preparing to image it (see section 6, &#34;imaging setup&#34;).</li>
		<li>Using the laser ablation system software, define the ablation zone, which corresponds to the crest and tip of the emerging sepal primordia before they start covering the flower meristem (typically, at stage 3 for abaxial and adaxial sepals, and at late stage 3/early stage 4 for lateral sepals).</li>
		<li>Set laser power and dwelling time to appropriate parameters to ablate enough cells without inflicting too much damage to the underlying tissues. Typically, depending on the laser ablation system used, proper parameters initially need to be identified through a trial-and-error process to ensure that enough cells are removed to prevent the sepals to subsequently grow over the FM, without affecting the rest of the flower bud. Using too much laser power and dwelling time results in damages to the center of the flower, affecting its growth and/or survival. Once the proper parameters have been identified, they can be reused for subsequent experiments. 
		NOTE: If the initial ablation proves insufficient, it is possible to proceed to a second ablation on the following days (Figure 2, C4 and D2).</li>
	</ol>
	</li>
</ol>
6. Imaging Setup
<ol>
	<li>Use an upright microscope.
	<ol>
		<li>Fill the imaging dish with the dissected apices with sterile, deionized water so that the surface of the medium is covered in 2.5-5 mm water (Figure 1F). Completely immerse the samples. 
		NOTE: When imaging with an upright microscope, several apices can be placed in the same imaging dish. However, if the imaging process lasts more than an hour, propidium iodide tends to be diluted, and some samples might need re-staining prior to imaging.</li>
		<li>Place the imaging dish on the microscope stage (Figure 1F). Lower the water-dipping lens and raise the stage so that the tip of the lens dips in the water. Proceed cautiously so as not to crush the dissected apices or dip the lens into the imaging medium.</li>
		<li>If an air bubble is trapped at the tip of the lens, make water jets with a 1,000 &#181;L pipette to remove it.</li>
		<li>Under epifluorescence illumination, position one of the dissected apices into the lens field using the XY controller. Look through the eyepieces and focus on the sample using the Z controller. Proceed cautiously in order not to crush the sample.</li>
		<li>Using the confocal microscope software, zoom on the flower bud to be imaged, and proceed to imaging your sample.</li>
	</ol>
	</li>
	<li>Use an inverted microscope.
	<ol>
		<li>Using a 1,000 &#181;L pipette, put a drop of sterile, deionized water on the tip of the lens.</li>
		<li>Hold the imaging dish upside-down, and with a 1,000 &#181;L pipette, add a drop of sterile, deionized water to the dissected apex.</li>
		<li>Place the imaging dish upside-down on the microscope stage (Figure 1G). Proceed cautiously so as not to crush the sample.</li>
		<li>Under epifluorescence illumination, position the dissected apex over the tip of the lens using the XY controller. With the Z controller, carefully lower the stage until the sample reaches the drop of water at the tip of the lens. A water column should form between the tip of the lens and the medium.</li>
		<li>If the water column does not form, carefully add a drop of water to the sample with a 1,000 &#181;L pipette. Add makeshift sleeves to the lens in order to add more water at its tip, which facilitates the establishment and maintenance of the water column (Figure 1H and 1I).</li>
		<li>Under epifluorescence illumination, look through the eyepieces and focus on the sample using the Z controller. Proceed cautiously so as not to crush the sample.</li>
		<li>Using the confocal microscope software, zoom on the flower bud to be imaged, and proceed to imaging the sample.</li>
	</ol>
	</li>
	<li>If performing time-lapse experiments, pour the water out of the imaging dish and close it to prevent dehydration in between time points. Place the imaging dish with the samples in long day (16 h light) or continuous day, 16-22 &#176;C conditions. Re-stain samples before each time point. 
	NOTE: While cells in developing floral organs can divide faster, cells in the floral meristem divide once or twice every second day, and cell lineages are easy to track with time points every 24 h. However, if need be, it is possible to image the samples every six hours.</li>
</ol>
7. Considerations on the Imaging Parameters
NOTE: How to set up the imaging parameters depends a lot on the confocal system used. Below are suggestions for some of these parameters that can be used with any confocal microscope. For more considerations on the imaging parameters, see the Discussion section and reference14.
<ol>
	<li>For the XY resolution: use 1,024 x 1,024 for a good cellular resolution. Alternatively, use 512 x 512 to reduce imaging time, at the expense of the resolution.</li>
	<li>For the Z resolution: set the step-size to 0.5-1.5 &#181;m. Lowering the step-size increases the Z resolution but also the imaging time.</li>
	<li>For the pinhole: set the pinhole to 1-1.5 airy units. Increasing the pinhole increases the intensity of the signal, but also the noise from non-focal planes.</li>
</ol>
8. Visualization of Confocal Data
<ol>
	<li>Turn still images of time points of flower development into a movie.
	<ol>
		<li>Open the still images (in jpeg or tif format) in the software (e.g., FiJi).</li>
		<li>Go to Image &#62; Stacks &#62; Images to Stack. The open images will be grouped in a stack.</li>
		<li>If the order of the images in the stack does not correspond to the chronological order, use the Stack Sorter plugin (Plugins &#62; Stacks &#62; Stack Sorter) to reorder them.</li>
		<li>To turn the stack into a movie, go to File &#62; Save As &#62; AVI.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Prunet, N., Jack, T. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flower+Development+in+Arabidopsis:+There+Is+More+to+It+Than+Learning+Your+ABCs.">Flower Development in Arabidopsis: There Is More to It Than Learning Your ABCs.</a> Methods Mol. Biol. 1110, 3-33 (2014).</li><li>Stewart, D., Graciet, E., Wellmer, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+and+regulatory+mechanisms+controlling+floral+organ+development.">Molecular and regulatory mechanisms controlling floral organ development.</a> FEBS J. , (2016).</li><li>Heisler, M. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterns+of+auxin+transport+and+gene+expression+during+primordium+development+revealed+by+live+imaging+of+the+Arabidopsis+inflorescence+meristem.">Patterns of auxin transport and gene expression during primordium development revealed by live imaging of the Arabidopsis inflorescence meristem.</a> Curr. Biol. 15 (21), 1899-1911 (2005).</li><li>Reddy, G. V., Meyerowitz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stem-cell+homeostasis+and+growth+dynamics+can+be+uncoupled+in+the+Arabidopsis+shoot+apex.">Stem-cell homeostasis and growth dynamics can be uncoupled in the Arabidopsis shoot apex.</a> Science. 310 (5748), 663-667 (2005).</li><li>Yadav, R. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=WUSCHEL+protein+movement+mediates+stem+cell+homeostasis+in+the+Arabidopsis+shoot+apex.">WUSCHEL protein movement mediates stem cell homeostasis in the Arabidopsis shoot apex.</a> Genes &amp; Dev. 25 (19), 2025-2030 (2011).</li><li>Reddy, G. V., Heisler, M. G., Ehrhardt, D. W., Meyerowitz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+lineage+analysis+reveals+oriented+cell+divisions+associated+with+morphogenesis+at+the+shoot+apex+of+Arabidopsis+thaliana.">Real-time lineage analysis reveals oriented cell divisions associated with morphogenesis at the shoot apex of Arabidopsis thaliana.</a> Development. 131 (17), 4225-4237 (2004).</li><li>Chandler, J. W., Jacobs, B., Cole, M., Comelli, P., Werr, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DORNROSCHEN-LIKE+expression+marks+Arabidopsis+floral+organ+founder+cells+and+precedes+auxin+response+maxima.">DORNROSCHEN-LIKE expression marks Arabidopsis floral organ founder cells and precedes auxin response maxima.</a> Plant Mol. Biol. 76 (1-2), 171-185 (2011).</li><li>Urbanus, S. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+planta+localisation+patterns+of+MADS+domain+proteins+during+floral+development+in+Arabidopsis+thaliana.">In planta localisation patterns of MADS domain proteins during floral development in Arabidopsis thaliana.</a> BMC Plant Biol. 9, (2009).</li><li>Hong, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variable+Cell+Growth+Yields+Reproducible+OrganDevelopment+through+Spatiotemporal+Averaging.">Variable Cell Growth Yields Reproducible OrganDevelopment through Spatiotemporal Averaging.</a> Dev. Cell. 38 (1), 15-32 (2016).</li><li>Roeder, A. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variability+in+the+control+of+cell+division+underlies+sepal+epidermal+patterning+in+Arabidopsis+thaliana.">Variability in the control of cell division underlies sepal epidermal patterning in Arabidopsis thaliana.</a> PLoS Biol. 8 (5), e1000367 (2010).</li><li>Heisler, M. G., Ohno, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-imaging+of+the+Arabidopsis+inflorescence+meristem.">Live-imaging of the Arabidopsis inflorescence meristem.</a> Methods Mol. Biol. 1110, 431-440 (2014).</li><li>Tobin, C. J., Meyerowitz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-Time+Lineage+Analysis+to+Study+Cell+Division+Orientation+in+the+Arabidopsis+Shoot+Meristem.">Real-Time Lineage Analysis to Study Cell Division Orientation in the Arabidopsis Shoot Meristem.</a> Methods Mol. Biol. 1370, 147-167 (2016).</li><li>Smyth, D. R., Bowman, J. L., Meyerowitz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+flower+development+in+Arabidopsis.">Early flower development in Arabidopsis.</a> Plant Cell. 2 (8), 755-767 (1990).</li><li>Prunet, N., Jack, T. P., Meyerowitz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+confocal+imaging+of+Arabidopsis+flower+buds.">Live confocal imaging of Arabidopsis flower buds.</a> Dev. Biol. , (2016).</li><li>Fernandez, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+plant+growth+in+4D:+robust+tissue+reconstruction+and+lineaging+at+cell+resolution.">Imaging plant growth in 4D: robust tissue reconstruction and lineaging at cell resolution.</a> Nat. Methods. , (2010).</li><li>Cutler, S. R., Ehrhardt, D. W., Griffitts, J. S., Somerville, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Random+GFP::cDNA+fusions+enable+visualization+of+subcellular+structures+in+cells+of+Arabidopsis+at+a+high+frequency.">Random GFP::cDNA fusions enable visualization of subcellular structures in cells of Arabidopsis at a high frequency.</a> Proc. Natl. Acad. Sci. U S A. 97 (7), 3718-3723 (2000).</li><li>Irish, V. F., Sussex, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Function+of+the+apetala-1+gene+during+Arabidopsis+floral+development.">Function of the apetala-1 gene during Arabidopsis floral development.</a> Plant Cell. 2 (8), 741-753 (1990).</li><li>Vernoux, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+auxin+signalling+network+translates+dynamic+input+into+robust+patterning+at+the+shoot+apex.">The auxin signalling network translates dynamic input into robust patterning at the shoot apex.</a> Mol. Systems Biol. 7, 508 (2011).</li><li>Zhou, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+plant+stem+cell+function+by+conserved+interacting+transcriptional+regulators.">Control of plant stem cell function by conserved interacting transcriptional regulators.</a> Nature. 517 (7534), 377-380 (2015).</li><li>Barbier de Reuille, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MorphoGraphX:+A+platform+for+quantifying+morphogenesis+in+4D.">MorphoGraphX: A platform for quantifying morphogenesis in 4D.</a> Elife. 4, 05864 (2015).</li><li>Barbier de Reuille, P. B., Robinson, S., Smith, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantifying+cell+shape+and+gene+expression+in+the+shoot+apical+meristem+using+MorphoGraphX.">Quantifying cell shape and gene expression in the shoot apical meristem using MorphoGraphX.</a> Methods Mol. Biol. 1080, 121-134 (2014).</li></ol>

The author has nothing to disclose.

Here, we provide a protocol for the imaging of live, developing Arabidopsis flower buds with a confocal microscope and a water-dipping lens. While this can be done with either an upright or an inverted microscope, it is easier and faster to use the former. It is also worth noting that different confocal systems vary significantly in terms of speed, sensitivity, and ability to separate wavelengths. The best confocal microscopes now allow to image several different channels (e.g. GFP, YFP, dsRed and propidium iodide; Figure 2G) in the same samples. Some confocal microscopes are equipped with a spectral detector, which can collect the fluorescence from several fluorophores simultaneously and separate them afterwards. When using a regular detector, however, a set of lasers and filters must be used to excite and separate the fluorescence from different fluorophores. Some fluorophores have fully distinct emission spectra (e.g. CFP and YFP), and can be imaged simultaneously. Other fluorophores have partially overlapping emission spectra (e.g. GFP and YFP), and usually need to be imaged separately, which increases imaging time. Imaging close fluorophores also often requires restricting how much of the spectrum is collected for each channel to prevent fluorescence from one fluorophore from leaking into another channel. This causes a loss of intensity of the collected signal, which can be compensated by an increase in laser power and gain. However, prolonged imaging time and increased laser power may bleach and/or damage the sample. Increased laser power and/or gain may also increase background noise. Optimization of signal intensity, resolution and imaging time for each sample is done through a trial-and-error process, and involves permanent trade-offs.
This protocol explains how to perform live confocal imaging of flower buds developing on the flanks of a dissected shoot apex. It may be argued that the fact that the shoot apex is not connected to the rest of the plant and grows in a medium containing cytokinins might affect SAM and flower development. However, this medium was designed empirically through a trial-and-error process to ensure the dissected shoot apical meristems produce new flower buds at the normal plastochron, and that these flower buds develop normally15. Similarly, sepal dissection does not appear to affect gene expression patterns or the development of the rest of the flower. Other protocols detail how to perform live confocal imaging of the SAM still attached to the rest of the plant (e.g. reference11). Such protocols could be adapted to, and offer a useful alternative for the imaging of developing flowers, particularly when studying cytokinin-related processes. They present several disadvantages however: the stem elongates and twists, and changes the orientation of the flower buds; and while imaging a shoot apex attached to the rest of the plant works well with an upright microscope, it is much more complicated to do so with an inverted microscope.
Immersion lenses have a better numerical aperture (NA) than &#34;dry&#34; lenses (i.e. lenses that are separated from the sample by air), and therefore provide a finer resolution of the sample, which is critical when using a confocal microscope. Using a water-dipping lens thus provides a much better NA than a dry lens, while preventing the shoot apex from dehydrating during the imaging process. While glycerin and oil lenses have an even higher NA than water lenses, they require the use of a coverslip, which is very unpractical with a sample the size of a shoot apex, which also keeps growing during time-lapse experiments. Given the size of the samples (depending on the floral stages, stacks can be over 150 &#181;m thick), it is also important to use a lens with a long working distance. We typically use a 20 or 40X lens with a NA of 1 and a working distance of 1.7-2.5 mm.
Confocal microscope software offers several ways to visualize confocal data, including three-dimensional reconstructions (Figure 2, B1, B2 and B4) and slice views (Figure 2, B3). While the most commonly used three-dimensional views are maximum intensity projections of the confocal Z-stacks (Figure 2, B1 and B4), some software (e.g. ZEN and Imaris) also offer transparency views that filters out the signal from the deeper parts of the sample (Figure 2, B2), which can be useful when looking at processes taking place, or genes expressed in, the epidermal layer. Signal intensity can either be coded with a single color (Figure 2, B1), using pixel intensity, or using a color code (Figure 2, B4).
Live confocal imaging not only offers valuable qualitative insights into flower development, it also potentially provides developmental biologists with a wealth of quantitative data. Different software (e.g. Imaris, FiJi, MARS-ALT, MorphographX15,20,21) allows for the quantification of the number or the volume of cells expressing one or several reporters, or the level of expression of a reporter in different cells. Such software can also be used to perform automatic cell segmentation, track cell lineages and quantify growth. This different software offers similar tools, but also differs in many ways. MARS-ALT and MorphoGraphX were designed specifically for plants15,20,21, unlike Imaris and FiJi. MorphoGraphX only allows for the segmentation and analysis of the epidermal layer, while Imaris and MARS-ALT allow for the segmentation and analysis of the whole sample. However, MARS-ALT requires the prior imaging of the same sample from three different angles15, and Imaris only requires a single stack. Access to quantitative information is critical to further our understanding of flower development.

Most of the plant body forms post-embryonically from groups of stem cells situated at or near the tip &#8212; or apical meristems &#8212; of the shoots and roots. All the above-ground structures of the adult plant derive from the shoot apical meristem (SAM), which continuously produces lateral organs on its flanks: leaves during the vegetative phase, and flower meristems (FMs) after it transitions to the reproductive phase. FMs in turn develop into flowers. While the Arabidopsis SAM produces lateral organs one at a time, in an iterative, spiral pattern, FMs produce four types of floral organs within four whorls, in a partially synchronous manner, with multiple developmental programs unraveling simultaneously. The genetic networks underlying the specification of the identity of the different floral organs have been partially deciphered (for reviews, see references1,2), but many aspects of flower development, such as floral organ positioning and the definition of boundaries between whorls, remain poorly understood.
Early molecular genetic studies of plant development mostly relied on techniques such as in situ hybridization and GUS reporters to analyze gene expression. While these methods have provided a wealth of information and greatly contributed to our understanding of plant growth and flower development, there are important limitations: they lack good cellular resolution, do not allow for the easy observation of the expression pattern of multiple genes in the same samples, and importantly, can only be applied to dead, fixed tissues. Recent advances in confocal microscopy have overcome these limitations, and provide developmental biologists with a powerful tool to investigate the processes underlying plant morphogenesis. In particular, confocal microscopy allows for the observation of live tissues and organs throughout their formation, which is critical to fully understand a quintessentially dynamic process such as development.
Live confocal imaging has been extensively used to analyze the aerial growth of plants and the production of lateral organs by the SAM (e.g. references3,4,5,6), but with the exception of a few reports (e.g. references7,8,9,10), it has not been widely applied to the study of flower development. Protocols for live confocal imaging of the SAM are available (e.g. references11,12), and provide a good base for how to image the developing flower buds that surround the SAM. However, imaging flower buds presents specific challenges: for instance, flower buds quickly become bigger than the SAM, and at stage 4, sepals start covering the FM (stages as described in reference13), and dimming the fluorescence from underlying tissues (Figure 2A). Here, we provide a detailed protocol explaining how to perform live confocal imaging on developing Arabidopsis flower buds using either an upright or an inverted microscope and a water-dipping lens. We previously published this protocol in reference14.

<table><tbody><tr><td>Forceps&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>72701-D</td><td>Dumont Style 5 Inox, for apex dissection</td></tr><tr><td>Pin Vise</td><td>Ted Pella, Inc.</td><td>13560</td><td>80 cm long, for sepal ablation</td></tr><tr><td>Straight Stainless Steel Needles&amp;nbsp;</td><td>Ted Pella, Inc.</td><td>13561-50</td><td>0.1 mm Diameter, 1.2 cm long, for sepal ablation</td></tr><tr><td>Round plastic boxes</td><td>Electron Microscopy Sciences</td><td>64332</td><td>transparent, 6 cm diameter, 2 cm deep, to use as dissecting dishes</td></tr><tr><td>Rectangular hinged boxes</td><td>Durphy Packaging Co.&amp;nbsp;</td><td>DG-0730</td><td>transparent, 2-7/8&amp;rdquo; long, 2&amp;rdquo; wide, 1-1/4&amp;rdquo; deep, to use as imaging dishes with an upright microscope</td></tr><tr><td>Tissue Culture Dishes, Polystyrene, Sterile</td><td>VWR</td><td>25382-064</td><td>transparent, 3.5 cm diameter, 1 cm deep,to use as imaging dishes with an inverted microscope</td></tr><tr><td>P10 and P1000 pipettes&amp;nbsp;</td><td></td><td></td><td>with corresponding filter tips</td></tr><tr><td>Stereomicroscope</td><td></td><td></td><td>with an 80-90X maximum magnification</td></tr><tr><td>Laser ablation system</td><td></td><td></td><td>for sepal ablation</td></tr><tr><td>Laser scanning confocal microscope system</td><td></td><td></td><td></td></tr><tr><td>20 or 40X water-dipping lens</td><td></td><td></td><td>with a NA of 1 and a working distance of 1.7-2.5 mm</td></tr><tr><td>Agarose</td><td></td><td></td><td></td></tr><tr><td>Sucrose</td><td></td><td></td><td></td></tr><tr><td>Murashige and Skoog basic salt mixture</td><td>Sigma-Aldrich</td><td>M5524</td><td>without vitamins</td></tr><tr><td>Myo-inositol</td><td>Sigma-Aldrich</td><td>I7508</td><td>vitamin</td></tr><tr><td>Nicotinic acid</td><td>Sigma-Aldrich</td><td>N0761</td><td>vitamin</td></tr><tr><td>Pyridoxine hydrochloride&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P6280</td><td>vitamin</td></tr><tr><td>Thiamine hydrochloride&amp;nbsp;</td><td>Sigma-Aldrich</td><td>T1270</td><td>vitamin</td></tr><tr><td>Glycine</td><td>Sigma-Aldrich</td><td>G8790</td><td>vitamin</td></tr><tr><td>N6-Benzyladenine&amp;nbsp;</td><td>Sigma-Aldrich</td><td>B3408</td><td>BAP, a cytokinin</td></tr><tr><td>Propidium iodide solution</td><td>Sigma-Aldrich</td><td>P4864</td><td>1 mg/mL solution in water, to stain the cell walls</td></tr><tr><td>FM4-64</td><td>Invitrogen</td><td>F34653</td><td>dilute in water to 80&amp;nbsp;&amp;mu;g/mL, to stain the plasma membranes</td></tr></tbody></table>

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.

Figure 2 presents different views of confocal Z-stacks of live Arabidopsis flower buds expressing different fluorescent reporter genes, and stained with either propidium iodide (Figure 2, A-C5 and G) or FM4-64 (Figure 2, E1-F) to provide a clear cellular resolution. Most confocal systems allow for the imaging of two fluorophores with non-overlapping emission spectra such as GFP or YFP together with either propidium iodide or FM4-64 (Figure 2A-2F). The best confocal systems are also able to separate multiple fluorophores with partially overlapping emission spectra in the same samples, such as GFP and YFP, and dsRed and propidium iodide (Figure 2G).
Three examples of time-lapse experiments are presented in Figure 2 and show flower buds developing normally after sepal ablation was performed either with a laser ablation system (Figure 2, C1-C5 and D1-D5) or manually with a pin-vise and a metal pin (Figure 2, E1-E2). Note that the flower bud shown in Figure 2, E1-E2 is from a superman-1 mutant plant, which is why carpel primordia are not observed at the center of the bud at stage 7. Laser ablation of emerging sepal primordia of the flower bud shown in Figure 2, C1-C5 prevents them from covering the FM, which can still be imaged at stage 5 (Figure 2, C5). Conversely, in the case of the flower bud shown in Figure 2, D1-D5, laser ablation only delayed sepal growth, but sepals eventually grew to cover most of the FM by stage 5 (Figure 2, D5). On the contrary, if too much laser power and dwelling time is used during laser ablation, damage spreads to the central part of the flower and affects its growth and sometimes results in the subsequent death of the whole flower bud.
<img alt="Figure 2" src="/files/ftp_upload/55156/55156fig2.jpg" /> 
Figure 2: Visualization of confocal Z-stacks of live flower buds. A and B2 are transparency views; B1 and B4-G are maximum intensity projections; B3 is an orthogonal slice view. (A-E2) flowers expressing a Venus reporter for the APETALA3 gene (green); cell walls were stained with propidium iodide (red, except for B4, green). (A) Inflorescence; numbers indicate floral stages; sepals in stage 4 and 5 flowers filter out the fluorescence of the Venus reporter, which normally forms a ring. Note that some flower buds appear tilted compared to the inflorescence. (B1-B4) Four different views of the same confocal Z-stack of a stage 5 flower bud: maximum intensity projections (B1 and B4) with Venus signal intensity coded with pixel intensity in the green color (B1) or with fire color code (B4; color code is shown at the bottom of the panel); transparency view (B2) and orthogonal slice view (B3; the XY, XZ and YZ orientation of the slices is indicated in the panel). (C1-C5) 4-day time-lapse of an individual flower bud from stage 3 to stage 5; laser ablations (marked as white traits) performed on day 1 and day 3 were sufficient to prevent the sepals from covering the center of the flower bud at stage 5. (D1-D5) 4-day time-lapse of an individual flower bud from stage 3 to stage 5; laser ablations performed on day 1, 2 and 3 were insufficient to prevent the sepals from covering the center of the flower bud. (E1-E2) Individual flower bud after manual removal of the abaxial and adaxial sepals (E1), and of all sepals (E2); white arrowheads indicate remaining sepals; white asterisks indicate scars resulting from the removal of the sepals. (F) stage 7 apetala1-1 flower expressing a DR5-3xVenusN7 reporter18 (green); plasma membranes were stained with FM4-64 (red); blue arrowheads indicate the leaf-like structures that replace sepals and do not cover the flower bud. (G) Stage 4 flower bud expressing a fluorescent GFP reporter for DORNROSCHEN-LIKE7 (green), a Venus reporter for SUPERMAN (red) and a dsRed reporter for CLAVATA319 (cyan); cells walls were stained with propidium iodide (grey). d: day; st: stage. Scale bars = 25 &#181;m; scale is identical in B1, B2 and B4, in C1-C5 and in D1-D5. This figure was modified from reference14. <a href="http://cloudflare.jove.com/files/ftp_upload/55156/55156fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Live Confocal Imaging of Developing Arabidopsis Flowers at JoVE.com

Live Confocal Imaging of Developing Arabidopsis Flowers

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. ...

live-confocal-imaging-of-developing-arabidopsis-flowers

Research

JoVE Journal

Developmental Biology

14.8K Views.  California Institute of Technology. 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.

Video: Live Confocal Imaging of Developing Arabidopsis Flowers

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 Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

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.

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 ...

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 ...

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

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the Arabidopsis leaf is a complex organ made up of many different cell types and several structures. At the same time, the growing leaf contains cells at different stages of development, with the cells furthest from the petiole being the first to stop expanding and undergo senescence1. Different cells within the leaf are therefore dividing, elongating or differentiating; active, stressed or dead; and/or responding to stimuli in sub-sets of their cellular type at any one time. This makes genomic study of the leaf challenging: for example when analyzing expression data from whole leaves, signals from genetic networks operating in distinct cellular response zones or cell types will be confounded, resulting in an inaccurate profile being generated.
To address this, several methods have been described which enable studies of cell specific gene expression. These include laser-capture microdissection (LCM)2 or GFP expressing plants used for protoplast generation and subsequent fluorescence activated cell sorting (FACS)3,4, the recently described INTACT system for nuclear precipitation5 and immunoprecipitation of polysomes6.
FACS has been successfully used for a number of studies, including showing that the cell identity and distance from the root tip had a significant effect on the expression profiles of a large number of genes3,7. FACS of GFP lines have also been used to demonstrate cell-specific transcriptional regulation during root nitrogen responses and lateral root development8, salt stress9 auxin distribution in the root10 and to create a gene expression map of the Arabidopsis shoot apical meristem11. Although FACS has previously been used to sort Arabidopsis leaf derived protoplasts based on autofluorescence12,13, so far the use of FACS on Arabidopsis lines expressing GFP in the leaves has been very limited4. In the following protocol we describe a method for obtaining Arabidopsis leaf protoplasts that are compatible with FACS while minimizing the impact of the protoplast generation regime. We demonstrate the method using the KC464 Arabidopsis line, which express GFP in the adaxial epidermis14, the KC274 line, which express GFP in the vascular tissue14 and the TP382 Arabidopsis line, which express a double GFP construct linked to a nuclear localization signal in the guard cells (data not shown; Figure 2). We are currently using this method to study both cell-type specific expression during development and stress, as well as heterogeneous cell populations at various stages of senescence.

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

A method for producing Arabidopsis leaf protoplasts that are compatible with fluorescence activated cell sorting (FACS), allowing for studies of specific cell populations. This method is compatible with any Arabidopsis line that expresses GFP in a subset of cells.

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the ...

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+ ...

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 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 ...

Live Confocal Imaging of Developing Arabidopsis Flowers

Summary

Explore More Videos

Chapters in this video

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

A Method for Characterizing Embryogenesis in Arabidopsis

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

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

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

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

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

Measuring Spatial and Temporal Ca²⁺ Signals in Arabidopsis Plants

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

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