Live Confocal Imaging of Developing Arabidopsis Flowers

The overall goal of this procedure is to prepare Arabidopsis thaliana inflorescences for imaging live developing flower buds with a confocal microscope. This method can help answer key questions in the flower development field about the formation and positioning of flower organs and the dynamics of flower stem cells. The main advantage of this technique is that it allows live imaging of flowers as they form which is critical to understanding what is quintessentially a dynamic process.

To expose the flower buds for imaging, use forceps to remove the secondary inflorescences, and older flower buds from the primary inflorescence. Next, use the forceps to pierce a vertical hole in the agarose of a dissecting dish. Now cut off the last 0.5 centimeters of the inflorescence and install it upright into the agarose with the remaining flower buds above the agarose surface.

Fill the dish with sterile deionized water so that the shoot apex is fully immersed and use a 1, 000 microliter pipette to flush any air trapped around the shoot apex. Then place the sample under a stereo microscope. Use the forceps to remove the flower buds that are not to be imaged from the immersed sample.

When reaching stage seven flower buds, remove the water from the dissecting dish, replace the dissecting dish under the microscope and finish the dissection. The apex can then be placed upright into the medium of an imaging dish such that only the shoot apical meristem and surrounding flower buds are above the surface of the medium. If not staining the apex immediately, use a 1, 000 microliter pipette to add a drop of water to the sample to keep it hydrated.

To stain the flower buds, first remove the water from the surface of the imaging dish with a piece of tissue paper. Then place the dish on its side to drain any remaining water from the medium surface. Wait a minute and then remove the pooled water with a new tissue.

Then under the microscope, use a 10 microliter pipette to apply 20 to 30 microliters of the appropriate stain, taking care that the entire sample is covered to ensure a homogenous staining. After the appropriate incubation period, rinse the buds two times with sterile deionized water. For imaging on an inverted microscope, use a 1, 000 microliter pipette to place a drop of sterile deionized water onto the tip of the lens.

Then turning the imaging dish upside down, add a drop of sterile deionized water to the dissecting apex and place the sample upside down onto the microscope stage, taking care not to crush the sample. Use the X, Y, and Z controllers to put the dissected apex in contact with the drop at the tip of the lens. A water column will form.

For imaging on an upright microscope, fill the imaging dish with sterile deionized water so that the surface of the medium and the samples are completely immersed in 2.5 to five millimeters of water. Next, place the dish on the upright microscope stage and adjust the water dipping lens in the stage so that the tip of the lens dips into the water. Then use a 1, 000 microliter pipette to flush away any air bubble trapped at the tip of the lens.

For either type of microscope, when the sample is ready, use the X, Y controller to position one of the dissected apices within the lens field and the Z controller to focus on the sample. When a sample has been located, use the confocal microscope software to zoom onto the flower bud of interest to obtain an image. Set the imaging parameters and define the top and bottom of the Z-stack and then proceed to imaging the flower bud.

For laser ablation of the sepal primordia, switch the light path to the camera associated with the laser ablation system. Next, use the laser ablation system software to define the ablation zone. Then set the laser power and dwelling time to the appropriate parameters for ablating enough cells without inflicting too much damage to the underlying tissues and then begin the ablation.

If timelapse experiments are to be performed, decant the water and cover the dish to prevent dehydration between the imaging time points. Then place the samples at the appropriate culture conditions. Most confocal systems allow the imaging of two fluorophores with non-overlapping emission spectra together such as GFP or YFP with either propidium iodide or FM4-64.

The best confocal systems are also able to separate multiple fluorophores with partially overlapping emission spectra in the same samples such as GFP or YFP, DsRed and propidium iodide as illustrated in this representative image. Here, examples of timelapse experiments with flower buds that developed normally after sepal ablation with a laser ablation system or manually with a pin vise and a metal pin are shown. Laser ablation of the emerging sepal primodia of this flower bud prevented them from covering the flower meristem which could still be imaged at stage five.

Conversely, in this flower bud, the laser ablation only delayed the sepal growth, although the sepals eventually grew to cover most of the flower meristems by stage five. After watching this video, you should have a good understanding of how to prepare Arabidopsis for the confocal imaging of live flower buds. Following this procedure, both qualitative and quantitative data can be extracted from your confocal images using different image analyzing software.

This technique will help researchers in the plant development field explore the dynamics of gene expression, cell division and organ formation in live developing flowers.