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>

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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 border="0" cellpadding="0" cellspacing="0" width="858">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:272px;">Forceps&#160;</td>
			<td style="width:143px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">72701-D</td>
			<td style="width:325px;">Dumont Style 5 Inox, for apex dissection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Pin Vise</td>
			<td style="width:143px;">Ted Pella, Inc.</td>
			<td align="right" style="width:119px;">13560</td>
			<td>80 cm long, for sepal ablation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Straight Stainless Steel Needles&#160;</td>
			<td style="width:143px;">Ted Pella, Inc.</td>
			<td style="width:119px;">13561-50</td>
			<td>0.1 mm Diameter, 1.2 cm long, for sepal ablation</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:272px;">Round plastic boxes</td>
			<td style="width:143px;">Electron Microscopy Sciences</td>
			<td align="right" style="width:119px;">64332</td>
			<td>transparent, 6 cm diameter, 2 cm deep, to use as dissecting dishes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rectangular hinged boxes</td>
			<td>Durphy Packaging Co.&#160;</td>
			<td style="width:119px;">DG-0730</td>
			<td>transparent, 2-7/8&#8221; long, 2&#8221; wide, 1-1/4&#8221; deep, to use as imaging dishes with an upright microscope</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:272px;">Tissue Culture Dishes, Polystyrene, Sterile</td>
			<td style="width:143px;">VWR</td>
			<td style="width:119px;">25382-064</td>
			<td>transparent, 3.5 cm diameter, 1 cm deep,to use as imaging dishes with an inverted microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">P10 and P1000 pipettes&#160;</td>
			<td style="width:143px;"></td>
			<td style="width:119px;"></td>
			<td>with corresponding filter tips</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Stereomicroscope&#160;</td>
			<td style="width:143px;"></td>
			<td style="width:119px;"></td>
			<td>with an 80-90 x maximum magnification</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Laser ablation system</td>
			<td style="width:143px;"></td>
			<td style="width:119px;"></td>
			<td>for sepal ablation</td>
		</tr>
		<tr height="21">
			<td colspan="2" height="21" style="height:21px;">Laser scanning confocal microscope system</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">20 or 40 x water-dipping lens</td>
			<td style="width:143px;"></td>
			<td style="width:119px;"></td>
			<td>with a NA of 1 and a working distance of 1.7-2.5 mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Agarose</td>
			<td></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sucrose</td>
			<td style="width:143px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Murashige and Skoog basic salt mixture</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">M5524</td>
			<td>without vitamins</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Myo-inositol</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">I7508</td>
			<td>vitamin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Nicotinic acid</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">N0761</td>
			<td>vitamin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pyridoxine hydrochloride&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">P6280</td>
			<td>vitamin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Thiamine hydrochloride&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">T1270</td>
			<td>vitamin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Glycine</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">G8790</td>
			<td>vitamin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N6-Benzyladenine&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">B3408</td>
			<td>BAP, a cytokinin</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">Propidium iodide solution</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;">P4864</td>
			<td>1 mg/mL solution in water, to stain the cell walls</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:272px;">FM4-64</td>
			<td style="width:143px;">Invitrogen</td>
			<td style="width:119px;">F34653</td>
			<td>dilute in water to 80 &#956;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. ...

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14.7K 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

Live-imaging of the Drosophila Pupal Eye

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

Live-imaging of the Drosophila Pupal Eye

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

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

In Vitro Differentiation of Mature Myofibers for Live Imaging

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

In Vitro Differentiation of Mature Myofibers for Live Imaging

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

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

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

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

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

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

Long-term Live Imaging of Drosophila Eye Disc

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

Long-term Live Imaging of Drosophila Eye Disc

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

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

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

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

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

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

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

Live Imaging of Mouse Secondary Palate Fusion

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

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

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

Live Cell Imaging of Chromosome Segregation During Mitosis

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

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

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

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 Imaging and Analysis of Muscle Contractions in Drosophila Embryo

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

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

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

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

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...