Live Confocal Imaging of Developing Arabidopsis Flowers

1. Media and Dishes Preparation

1. Prepare dissecting dishes by filling round plastic boxes (approximately 6 cm wide, 2 cm deep) to 0.5 cm with 2% agarose.
2. Prepare imaging dishes.
   1. 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, "Imaging medium"; Figure 1F).
   2. 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, "Imaging medium"; Figure 1G).
3. Prepare imaging medium.
   1. For single point imaging, use 1% agarose.
   2. 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.

2. Plant Growth

1. 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 °C conditions for two weeks.
2. Transplant seedlings onto soil with sufficient spacing to allow robust development. Place plants in short day, 16-22 °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.
3. Transfer plants to long day (16 h light) or continuous day, 16-22 °C conditions until they flower. Shoot apices are easiest to dissect when the inflorescence is 2-10 cm long.

3. Dissection of the Shoot Apex

1. 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.
2. 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).
3. 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.
4. 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 µL pipette (Figure 1, compare C and D).
5. 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.
6. 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.
7. 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).
8. 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.

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 µm in (E). This figure was initially published in reference¹⁴. Please click here to view a larger version of this figure.

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 used^6,16.

1. Remove water from the imaging dish, and ensure the surface of the medium is dry to avoid diluting the dye.
2. Under the stereomicroscope, apply 20 to 30 µL of either 1 mg/mL propidium iodide solution, or 80 µg/mL FM4-64 solution to the dissected apex with a 10 µL pipette. Make sure the whole sample is covered in dye to ensure a homogeneous staining.
3. Stain for 2 min if using propidium iodide, or 20 min if using FM4-64.
4. Rinse twice with sterile, deionized water.

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.

1. Sepal ablation on flower buds stage 5 and older using a pin-vise holding a straight metal needle (Figure 2, E1-E2)
   1. 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 µL pipette to apply water to the shoot apex regularly while dissecting the sepals.
   2. 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.
   3. Proceed similarly with the adaxial sepal, but push the sepal towards the shoot apex.
   4. 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.
   5. Immerse the shoot apex in sterile, deionized water for a few minutes to prevent dehydration.
2. Laser ablation of sepal primordia at stage 3-4 (Figure 2, C5-D5)
   1. 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, "imaging setup").
   2. 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).
   3. 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).

6. Imaging Setup

1. Use an upright microscope.
   1. 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.
   2. 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.
   3. If an air bubble is trapped at the tip of the lens, make water jets with a 1,000 µL pipette to remove it.
   4. 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.
   5. Using the confocal microscope software, zoom on the flower bud to be imaged, and proceed to imaging your sample.
2. Use an inverted microscope.
   1. Using a 1,000 µL pipette, put a drop of sterile, deionized water on the tip of the lens.
   2. Hold the imaging dish upside-down, and with a 1,000 µL pipette, add a drop of sterile, deionized water to the dissected apex.
   3. Place the imaging dish upside-down on the microscope stage (Figure 1G). Proceed cautiously so as not to crush the sample.
   4. 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.
   5. If the water column does not form, carefully add a drop of water to the sample with a 1,000 µL 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).
   6. 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.
   7. Using the confocal microscope software, zoom on the flower bud to be imaged, and proceed to imaging the sample.
3. 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 °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.

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.

1. 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.
2. For the Z resolution: set the step-size to 0.5-1.5 µm. Lowering the step-size increases the Z resolution but also the imaging time.
3. 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.

8. Visualization of Confocal Data

1. Turn still images of time points of flower development into a movie.
   1. Open the still images (in jpeg or tif format) in the software (e.g., FiJi).
   2. Go to Image > Stacks > Images to Stack. The open images will be grouped in a stack.
   3. If the order of the images in the stack does not correspond to the chronological order, use the Stack Sorter plugin (Plugins > Stacks > Stack Sorter) to reorder them.
   4. To turn the stack into a movie, go to File > Save As > AVI.