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

The overall goal of this imaging system is to use confocal fluorescence microscopy to observe the development of plant organs over several days at a sub-cellular resolution. This method can help answer key questions in plant morphogenesis, such as how do cells of multicellular organisms coordinate their patterns of growth and proliferation during organogenesis? The main advantage of this technique is that it is simple to set up and does not rely on the specialist equipment often used in time lapse imaging, like profusion systems.

We first had the idea for this method when we read the publication by Littlejohn and Love, in New Phytologist that describes the advantages of perfluorodecalin for imaging within leaf tissues. To begin, use a glass cutter to make three millimeter wide glass strips from a one millimeter thick slide. Then, with cyanoacrylate glue or double sided tape, secure the strips approximately 45 millimeters apart across the width of a new slide.

Make an extra slide with the same dimensions to serve as a mold. Now, make a gasket of the same height from gas permeable PDMS gum. Wet a second slide with a little absolute ethanol and flatten the PDMS onto the slide to the height of the glass strips.

Next, wet a blade with absolute ethanol and trim off any excess PDMS. This is critical as tools wet with ethanol will not stick to the PDMS. Then, reflatten the PDMS down to the height of the strip.

Then, cut a cavity into the PDMS that will hold the seedling and agar slab. To produce a consistent cavity size, we use a bespoke homemade cutter built from perspecs but a razor blade could be used instead. Next, trim around the outside edge to make the gasket about two millimeters thick.

Now, onto a slide without the PDMS gasket, rest a coverslip on both glass strips. Then into the space below the coverslip, pipette about one milliliter of warm liquefied agar medium enough to fill the space. The agar can contain experimental compounds of interest.

Next, air equilibrate some PFD by shaking a small volume of PFD in a tube. Once the agar has set, load approximately 200 microliters of the PFD into the PDMS gasket, but do not fill it completely. Remove the coverslip from the agar slab, and cut it into a shape that will fit the well of the PDMS gel gasket.

Leave a two to four millimeter gap between the agar and the gasket wall. Then, completely fill the chamber with air equilibrated PFD. To image gravitropic primary roots, a cellulosic membrane is used as a physical barrier on the surface of the agar slab.

The agar slab must also be softer with 0.8%agar rather than 1.5%agar. Cut a piece of the membrane that is approximately the same dimensions as the agar block. Then, sterilize it with 80%ethanol and allow to air dry.

Once dried, soak the membrane in liquid growth medium and then transfer it to the agar surface. To begin, carefully place up to three seedlings onto the agar. Their positioning is critical.

The cotyledons and hypocotyl must hang over the agar edge, floating in PFD. Space must be provided between the seedlings to allow for growth. Seedlings can be curled if they are too large to fit into the chamber.

Then, close the chamber using a coverslip with a thickness that matches the chosen objective. Press gently along the edge so contact is made with the glass strips. Use another glass slide to gently press down the coverslip so it rests evenly on the two glass strips.

Then, secure the coverslip using micropore surgical tape cut in half lengthwise. Now, allow the specimen to settle for about half an hour before imaging. To prepare the arabidopsis thaliana seedlings for the chambers, first sterilize the seeds with 70%ethanol.

And then plant them on supplemented medium. Stratify the planted seeds for two or four days at four degrees Celsius. Then, transfer the planted seeds to 22 degrees Celsius, and grow them vertically oriented under a 16 hour daily light cycle.

To image the lateral roots, grow the plants for seven to 10 days and then transfer them to imaging chambers. Lateral roots growing at physiological rates in imaging chambers were documented every half an hour. Lateral roots were also grown on standard petri plates in the same medium for comparison.

The growth rate of individual roots varied a lot in both setups, but the range was similar under either condition. Overall, the rate of growth increased with length. The growth rate of plants in imaging chambers or on plates was indistinguishable over the first 27 hours of growth.

Both groups started with the same average lateral root length. Lateral roots co-expressing a plasma membrane marker and a micro tubule marker were imaged on one hour intervals in the imaging chambers. The roots were highly stable in their positions, allowing for several hours of confocal imaging.

Visual indicators of proliferation were observed throughout the experiment. In the meristematic cells, pre prophase bands mitotic spindles, and phragmoplasts could all be identified. Over three days, lateral root growth was not significantly different in imaging chambers compared to petri plates during the first 48 hours, but by 72 hours the mean growth rate dropped.

However, a significant proportion of roots in the chambers still grew at comparable rates to roots on plates over 72 hours. Once mastered, two slides can be made in 15 minutes if everything goes smoothly. It is possible to use this imaging system to study the effect of pharmacological treatments, although other more complex methods like perfusion systems are required for washout experiments.

After its development, this technique provided a cheap and simple means to follow cellular events underpinning plant morphogenesis, in particular to explore the role of cell edges, of spatial demands sharing organogenesis in the lateral roots of arabidopsis thaliana.