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

Summary

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.

Abstract

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.

Introduction

Plant cells are surrounded by an extracellular matrix of polysaccharides and glycoproteins that mechanically resembles a fiber reinforced composite material capable of dynamically changing its mechanical properties¹. Growth in plant cells is driven by the uptake of water into the cell, which results in a concomitant buildup of tensile forces on the cell wall. In response to such forces, modifications to the physical state of the cell wall allows for cell expansion. Cells with primary walls are capable of undergoing rapid growth compared to secondary cell wall containing cells mainly due to differences in the chemical composition of the polysaccharides within. Primary wall cells are composed of cellulose, hemicellulose, and pectin in addition to glycoproteins, and lack lignin, a component that is present in the secondary cell wall². Cellulose, a glucose polymer linked via β-1,4 bonds, is the major component of the cell walls. It is organized into fibrillar structures that are capable of withstanding high tensile forces experienced during cell growth³. In addition to withstanding tensile forces, mechanical reinforcement along a preferential direction results in turgor-driven expansion along an axis perpendicular to the net orientation of the cellulose microfibril. The organization of the cellulose microfibrils is influenced by the cortical microtubule cytoskeleton, as they guide the directional movement of the cellulose-synthesizing complexes located at the plasma membrane⁴. Therefore, monitoring cortical microtubule organization using a microtubule-associated protein or tubulin fused with a fluorescent molecule serves as a proxy for the observation of overlying patterns of cellulose in plant cells.

The patterning of the cortical microtubule cytoskeleton is under the control of cell and tissue morphology derived mechanical forces. Cortical microtubule organization does not have any preferential organization over time in cells located at the apex of the SAM, whereas cells in the periphery and the boundary between the SAM and the emerging organ have a stable, highly organized supracellular array of cortical microtubules⁵. Several approaches have been developed to physically perturb the mechanical status of the cells. Changes to osmotic status, as well as treatment with pharmacological and enzymatic compounds that influence the stiffness of the cell wall can result in subsequent changes in the tensile forces experienced by the cell^6,7. The use of contraptions that allow for the gradual increase in compressive forces experienced by tissues is another alternative⁸. Application of centrifugal forces has also been shown to influence the mechanical forces without any physical contact with the cells⁹. However, the most widely used means of changing directional forces in a group of cells take advantage of the fact that all epidermal cells are under tension and physical ablation of cells will eliminate turgor pressure locally as well as disrupting cell-to-cell adhesion, thereby modifying the tensile forces experienced by the neighboring cells. This is performed either by targeting a high-powered pulsed ultraviolet laser or by means of a fine needle.

Here we elaborate on the process of imaging and assessing cortical microtubule behavior for mechanical perturbation at the SAM.

Protocol

1. Plant growth

1. Sow Arabidopsis seeds expressing microtubule binding domain fused with green fluorescent protein (MBD-GFP)¹⁰ on soil and keep in long day (16 h day /8 h night), 20 °C/6 °C conditions for 1 week for germination.
2. After germination, transfer seedlings to new pots with sufficient growth space to allow robust vegetative growth. Keep plants in short day (8 h day /16 h night), 20 °C/16 °C conditions for 3-5 weeks.
3. Transfer plants to long day (16 h day /8 h night), 20 °C/16 °C conditions and keep until plants bolt (2-3 weeks). Allow the inflorescence to grow up to 2-5 cm long.

2. Medium preparation

1. Prepare the dissecting dishes by filling small Petri dishes (approximately 5.5 cm wide, 1.5 cm deep) to approximately half their depth with 2% agarose. The dissecting dishes could also be used for single time point imaging.
2. Prepare the growth medium.
   1. Prepare 50 mL of a 1,000x vitamin stock solution with 5 g of myo-inositol, 0.05 g of nicotinic acid, 0.05 g of pyridoxine hydrochloride, 0.5 g of thiamine hydrochloride, and 0.1 g of glycine.
   2. Prepare the growth medium, composed of ½ Murashige and Skoog medium, 1% sucrose, 1.6% agar, 1x vitamins, pH = 5.8. Autoclave and allow it to cool.
   3. On a sterile, clean bench, sterilize hinged plastic boxes (approximately 5.2 cm x 5.2 cm x 3 cm) by immersing in 70% ethanol for 15 min.
   4. On a sterile, clean bench, once the medium is at bearable warmth, add N6-benzyladenine to a final concentration of 200 nM, and 1/1,000 PPM (plant preservative mixture) and mix well. Fill the sterile boxes to approximately half their depth with the medium.

3. Dissection of the SAM

1. Cut the inflorescence and remove the older flower buds with sharp forceps by peeling the flowers at the base of the peduncles until it is difficult to see them with the naked eye.
2. Create a slit in the agarose in the dissecting dish with forceps and plant the inflorescence base into the thick agar. This gives good solid support for further fine dissection of younger flowers to expose the SAM.
3. Remove the remaining flower buds by pushing them down with the forceps starting with the oldest and sequentially progressing to the younger stages under a dissecting microscope until the SAM is visible. The SAM is usually exposed when older flowers up to stage 6 to 7 are removed. Once the SAM is exposed, avoid dehydration of the sample by moving quickly to the next step.

4. Transfer and growth of cultured SAMs

1. Plant the freshly dissected sample as detailed in section 1 into the growth medium in the rectangular plastic hinged culture box with the SAM just exposed above the medium surface. Add a few drops of sterile deionized water to the edges of the culture boxes and close the lid to maintain the humidity inside the box. Ensure that the added water does not cover the SAM.
2. Close the lid and wrap the box with micropore tape. Place the growth box in long day or continuous day conditions at 22 °C and grow for 12-24 h to allow the SAM to recover from the dissection procedure and adapt to the culture conditions.

5. Imaging of the SAM

1. Fill the culture box containing the SAM with sterile deionized water to cover the sample. Check under the dissecting microscope and remove any air bubbles by forcefully spraying water directed at the sample with a 1 mL pipette.
2. Place the culture box on an upright confocal microscope stage. Take care that the culture box does not contact the microscope objectives. Use a long distance 40x or 60x water dipping lens of numerical aperture 0.8-1 that is optimal for imaging without a cover glass.
3. Lower the objective into the water and check for air bubbles formed on the objective's front lens. Remove any bubbles by lowering the stage and gently wiping the lens with an optical tissue and adding a small amount of water to the front lens of the objective with a Pasteur pipette before reimmersion into the water.
4. Using the GFP filter and epi-illumination module of the confocal microscope, adjust the XY controller to locate the sample. Adjusting the position of the oculars, put the SAM directly under the light source and focus along the Z axis until the apex is located.
   NOTE: Do not look directly at the ultraviolet light.
5. Once the sample is found, illuminate it using a laser capable of exciting GFP (i.e., a 488 nm or 496 nm laser source). Adjust the optical zoom of the microscope so that the entire SAM and the stage 1 floral primordia are in the field of view. Further adjust the power of the laser output and gain settings to ensure optimal signal-to-noise ratio.
   NOTE: The high intensity of the laser will result in photo bleaching of the sample. The under- and overexposure palette help ensure better adjustment of these settings.
6. Allow the sample to settle down for 2-5 min. Acquire confocal Z stacks of the sample at 0.25 µm-0.5 µm Z slice intervals at a resolution of approximately 0.3 µm pixel size. Ensure that the total imaging time required for acquiring Z stacks for one sample does not exceed 10 min from the time the sample is immersed into water until the completion of the acquisition.
7. Immediately remove the water and transfer the culture box back to the growth chamber. Keeping the samples for a long time under water will influence the osmotic status of the cells.

6. Micromechanical perturbation of the SAM

1. Set up the imaging conditions as described in section 5 and acquire preablation image stacks of the cortical microtubule organization.
2. Decant the water in the culture dish. Perform the ablation with a clean syringe needle (0.4 mm x 20 mm).
   1. Using a dissecting microscope, carefully hold the needle and slowly approach the SAM.
      NOTE: Breath holding and handling the needle with a relaxed grip helps avoid shaking.
   2. Briefly contact the SAM dome with the needle tip to confirm that the ablation is done. Preferably perform the ablation on the periphery of the SAM, which allows visualization of the behavior and transitioning of unorganized cortical microtubules in the central region of the dome.
3. Refill the culture dish with sterile deionized water and add propidium idodide (10 µg/mL). In addition to monitoring the GFP cortical microtubule signal, visualizing the propidium iodide using a separate channel can clearly mark regions of ablated cells. Propidium iodide is illuminated using the 561 nm or any other suitable laser with emission recorded between 600-650 nm.
4. Acquire image stacks right after ablation (see section 6) and repeat the acquisition process every 2 h for a period of 6 h. Return the culture dish along with the sample into the incubation chamber after every time point. Ensure there is some water left on the culture media and wrap the culture dish with micropore tape to prevent desiccation.

7. Data visualization and quantification

1. Generate surface projections using freely available software such as MORPHOGRAPHX¹¹, FIJI¹², or macro SurfCut¹³.
2. Perform extraction of cortical microtubule anisotropy using FibrilTool¹⁴ macro in FIJI.
   NOTE: Detailed protocols for software use can be obtained from the respective citations.

Results

Figure 1 shows typical projection images obtained from MBD-GFP lines with cells at the center of the dome containing disorganized cortical microtubules, and cells at the periphery having a circumferential distribution (Figure 1A,B), whereas the boundary domain cells contain cortical microtubules aligned parallel to the cell's long axis. These observations show differences in the spatial distribution of cortic...

Discussion

The assessment of mechanical signal transduction events is crucial to identify molecular regulators involved in the mechano-perception and transduction pathways. The protocol described here provides a quantitative view of such events by using the cortical microtubule response as a readout for such a process in Arabidopsis SAMs. The procedure described here is routinely used in several studies in various tissue types^16,17,18

Materials

Name	Company	Catalog Number	Comments
FibrilTool			Boudaoud, A. et al., Nat Protoc. 2014
FIJI			Schindelin, J. et al., Nat Methods. 2012
glycine	Merck	1.04201.1000
Leica SP8 confocal microscope	Leica	DM6000 CS
MAP4-GFP			Marc, J. et al., Plant Cell 1998
micropore tape	Leukopor	02482-00
MorphographX			Strauss, S. et al., Methods Mol Biol. 2019
myo-inositol	Sigma	I5125
N6-benzyladenine	Sigma	B3408
nicotinic acid	Sigma	N4126
plastic hinged box	Electron microscopy sciences	64312
PPM (Plant Preservative Mixture)	Plant Cell Technology	PPM
Propidium iodide	Sigma	P4864
pyridoxine hydrochloride	Sigma	P9755
SURFCUT			Erguvan, O. et al., BMC Biol. 2019
thiamine hydrochloride	Sigma	T4625