This research was funded by CSIC I+D 2018, grant No. 95 (Mariana Sotelo Silveira).; CSIC Grupos (Omar Borsani) and PEDECIBA.

1. Preparation of the plant material and growth conditions
<ol>
	<li>To generate the needed plant material, sterilize the seeds of Arabidopsis wild type and mutant lines of interest. 
	NOTE: In this protocol, we used the following: ttl1: T-DNA insertion lines Salk_063943 (for TTL1; AT1G53300) - Columbia-0 (Col-0) wild-type; the Procuste1 (prc1-1) mutant, which consists of a knock-out mutation (Q720stop) in the CESA6 gene (AT5G64740), as previously described in10,11; and ttl1 x prc1-1 double mutant.
	<ol>
		<li>Put an approximate volume of 50 &#956;L of seeds in a microtube. Add 500 &#956;L of 70% ethanol + 0.01% Tween solution at room temperature. Mix and incubate for 7 min.</li>
		<li>Pour the ethanol solution and add 500 &#956;L of 20% sodium hypochlorite. Mix and incubate for 7 min.</li>
		<li>Pour the sodium hypochlorite solution and wash the seeds three to four times with sterile water.</li>
	</ol>
	</li>
	<li>Plating
	<ol>
		<li>Prepare in advance basal Murashige and Skoog (MS) medium12 + 1.5% sucrose + 1% agar (pH 5.7) (see Table of Materials). Autoclave the medium. Prepare a laminar flow hood to work in a sterile environment and label the square Petri dishes needed for the experiment. Pour the medium into the Petri dishes and allow the medium to solidify.</li>
		<li>Plate the sterilized seeds using sterile pipette tips onto square Petri dishes containing the medium. Distribute the seeds evenly in two rows. When the seeds are dry, close the lids and seal the plates. Stratify the seeds at 4 &#176;C in the dark for 4 days. 
		NOTE: Stratification is needed to synchronize germination.</li>
		<li>Place the plates vertically in the growth chamber in a long-day regime (16 h of light/8 h of dark) with 25 &#956;mol m2 s&#8722;1 light intensity and 23 &#176;C (day/night). Grow the seedlings for 7 days.</li>
	</ol>
	</li>
</ol>
2. Osmotic stress treatment (optional)
NOTE: This section provides details on the growth of Arabidopsis&#160;roots in osmotic potential of -1.2 MPa as estimated by cryoscopic osmometer (Table of Materials). This part can be omitted or changed depending on the experimental question at hand.
<ol>
	<li>Prepare basal MS medium + 1.5% sucrose + 400 mM mannitol + 1% agar (pH 5.7). Autoclave the medium. Pour the medium in square Petri dishes in the laminar flow hood and allow the medium to solidify.</li>
	<li>With tweezers (Table of Materials), place 5-day old seedlings pre-grown in basal MS + 1.5% sucrose medium in square Petri dishes containing the medium supplemented with 400 mM mannitol. Place the plates vertically in the growth chamber in the same conditions as in step 1.2.3. Grow the seedlings under this osmotic stress condition for 7 days. 
	NOTE: To evaluate root growth adaptation of the primary root during severe osmotic stress, it is suggested to use seedlings 5 days after germination grown in controlled conditions to ensure that the root meristem size is already established13. Alternatively, the analysis can be performed at every stage of development to determine the developmental effect of osmotic stress action.</li>
</ol>
3. Atomic force microscopy (AFM) nanoindentation experiments
<ol>
	<li>Preparation of the sample for nanoindentation experiments 
	NOTE: This is a crucial step for applying AFM to the study of live biological samples. The sample must be firmly attached to the substrate to be mechanically stable during indentation measurements. At the same time, sample structural qualities should be preserved. An effective strategy to stabilize a big sample for AFM is to use adhesives. The adhesive must dry quickly and must not be toxic or reactive with the surrounding medium. In this protocol, polystyrene Petri dishes were used as the substrate and non-acidic silicone glue (Table of Materials) was used to bind the Arabidopsis seedlings.
	<ol>
		<li>Spread a thin layer of silicone glue into the Petri dish with a coverslip. Leave the glue in the air for 45 s.</li>
		<li>With tweezers, place the seedling on the glue, orienting it in a direction that avoids contact between the seedling&#39;s protruding parts and the cantilever. Then, press the root gently to the silicone glue layer to bind it firmly. Leave the seedling in contact with the glue for 45 s before continuing with the next steps of the protocol.</li>
		<li>Incubate the attached seedlings with 2 &#956;g/mL propidium iodide (PI; Table of Materials) for 5 min in dark and carefully wash them thrice with 1x phosphate-buffered saline (PBS) solution (Table of Materials).</li>
		<li>Place the PI-incubated seedlings into an inverted fluorescence microscope coupled with the AFM. To observe fluorescence, set the excitation and emission wavelengths at 572 nm and 617 nm, respectively. The absence of fluorescence inside the epidermal cells confirms the viability of the roots.</li>
		<li>Cover the seedling with 1x phosphate-buffered saline (PBS) solution.</li>
	</ol>
	</li>
	<li>Indentation protocol 
	NOTE: Perform all AFM measurements within 1 h after the immobilization of the sample on its support at room temperature. The room where the AFM is located should be maintained at 25 &#176;C with an air conditioner. The PBS solution should be at the same temperature.
	<ol>
		<li>Mount a standard silicon nitride cantilever with a pyramidal tip (see Table of Materials) into the AFM probe holder for fluid (Table of Materials).</li>
		<li>Align the laser on the cantilever close to the position of the tip. Next, move the photodiode to place the laser spot at the center of the detector.</li>
		<li>To calibrate the deflection sensitivity, position a polystyrene Petri dish with 1x PBS under the AFM. 
		NOTE: This step is needed to calculate the indentation. When a force measurement is made on a hard surface, there is no indentation into the surface, so any change in the z scanner displacement corresponds to the cantilever deflection. Therefore, it is very important to calibrate the deflection sensitivity on a hard surface. Using a soft surface will overestimate the deflection sensitivity, which will result in spring constants that are too low.</li>
		<li>Adjust the photodetector such that the non-contact deflection is near 0 V. Calibrate the deflection sensitivity by performing an indentation (force curve), with a ramp size of 3 &#956;m, an indentation-and-retraction rate of 0.6 &#956;m/s, and a trigger threshold of 0.5 V.</li>
		<li>To calibrate the spring constant of the cantilever, use the Thermal Tune utility of the AFM software (Table of Materials) recommended for probes with spring constants less than approximately 5 N/m or use the method described in reference14. Before activating Thermal Tune in the software, ensure that the probe does not interact with the sample.
		<ol>
			<li>Enter the cantilever temperature. Click on Calibrate &#62; Thermal Tune or on the Thermal Tune icon in the NanoScope toolbar. Select a frequency range (1-100 Hz).</li>
			<li>Click on Acquire Data in the Thermal Tune panel. Wait for the AFM to acquire data, and then click on the Simple Harmonic Oscillator (Fluid) button.</li>
			<li>Adjust Median Filter Width to 3. Adjust PSD Bin Width to reduce the noise in the acquired data by averaging. Set fit boundaries around the first resonance peak.</li>
			<li>Click on Calculate Spring Constant k, and then, click on Yes in the pop-up window asking if the user wants to use this value.</li>
			<li>Repeat the steps described above three times and manually take an average of spring constant values. Enter this average value in the Spring Constant box in the Ramp parameter list in the PicoForce view. The calibration ends at this point.</li>
		</ol>
		</li>
		<li>Using the inverted optical microscope at 10x, 20x, and 40x eyepiece magnification, position the AFM probe on the surface of the fourth elongated epidermal cell of the primary root, ensuring to position it in the center of the cell.</li>
		<li>With the calculated spring constant value (step 3.2.5.5), obtain force curves with a ramp size of 3 &#181;m, a trigger threshold of 11 nN, and an indentation-and-retraction rate of 0.6 &#956;m/s at selected points. 
		&#8203;NOTE: Previous work15,16 found that low-frequency of indentation-and-retraction helps to minimize hysteresis and drag force.</li>
		<li>Obtain force curves from three cells per root for each treatment (use three biological replicates per treatment). Capture at least 150 force curves for each root.</li>
	</ol>
	</li>
</ol>
4. Measure the apparent Young&#39;s modulus
<ol>
	<li>Fit each force curve to the following model for pyramidal indenters17: <img alt="Equation 1" src="/files/ftp_upload/63533/63533eq01.jpg" style="margin: auto;" />, where E is the apparent Young&#39;s modulus of the sample, <img alt="Equation 2" src="/files/ftp_upload/63533/63533eq02.jpg" style="margin: auto;" /> is the Poisson modulus of the sample, and &#945; is the semi-angle to the vertex. Discard the fits with a correlation coefficient (r2) &#60; 0.99. Consider the Poisson&#39;s ratio for totally incompressible material: <img alt="Equation 2" src="/files/ftp_upload/63533/63533eq02.jpg" style="margin: auto;" /> = 0.5 and the geometry of the indenter given by the manufacturers. 
	NOTE: Use the first 100 nm from the contact point of the loading curve for the fitting to be as sensitive as possible to cell wall mechanics and to reduce the impact of turgor pressure on the apparent Young&#39;s modulus values8. The fitting has been done using a custom MATLAB program (available upon personal request by writing to J. C. B) that allows setting a specific indentation depth from the contact point of the loading curve.</li>
	<li>Create a normalized histogram with the apparent Young&#39;s modulus data and fit it with a Gaussian distribution. Discard data points outside the 95% confidence interval and recalculate both the histogram and the Gaussian fit. Calculate and report the mean and standard deviation of the apparent Young&#39;s modulus.</li>
	<li>Determine the significance of the comparisons between groups by multiple comparison tests like ANOVA.</li>
</ol>

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The authors have no conflicts of interest. The MATLAB script used for fitting the data is available upon personal request by writing to J. C. B.

Cell and cell-wall mechanics are increasingly becoming relevant to gain insight into how mechanics affects growth processes. As physical forces propagate over considerable distances in solid tissues, the study of changes in the physical properties of the cell wall and how they are sensed, controlled, tuned, and impact the plant&#39;s growth are becoming an important field of study2,3,8.
A method is pr...

Plant cells are surrounded by a cell wall that is a complex structure composed of interacting networks of polysaccharides, proteins, metabolites, and water that varies in thickness from 0.1 to several &#181;m depending on the cell type and the phase of growth1,2. Cell wall mechanical properties play an essential role in the growth of plants. Low stiffness values of the cell wall have been proposed as a precondition for cell growth and cell-wall expansion, and there is increasing evidence that all cells sense mechanical forces to perform their functions. However, it is still debated whether changes in the physical properties of the cell wall determines cell fate2,3,4. Because plant cells do not move during development, the final shape of an organ depends on how far and in what direction a cell expands. Thus, Arabidopsis root is a good model to study the impact of cell wall physical properties in cell expansion because different types of expansion occur in different regions of the root. For example, anisotropic expansion is evident in the elongation zone and particularly noticeably in the epidermal cells5.
The method described here was used to characterize the physical properties of the cell wall of epidermal cells at the nanoscale of living Arabidopsis roots using an Atomic Force Microscope (AFM) coupled with an inverted fluorescence phase microscope6. For an extensive revision of the AFM technique, read7,8,9.
This protocol outlines a basic sample preparation method and a general method for AFM-based elasticity measurements of plant cell walls.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63533/63533fig01v2.jpg" /> 
Figure 1: Schematic overview of force-indentation experiment in Arabidopsis roots using atomic force microscopy (AFM). The scheme gives an overview of the steps of a Force-Indentation experiment from the preparation of the substrate to immobilize the root sample firmly (1-2), root viability confirmation through propidium iodide staining (3), cantilever positioning on the surface of an elongated epidermal cell of the primary root (4-5), force curves measurement (6), and force curve processing to calculate the apparent Young&#39;s modulus (7-8). EZ: elongation zone. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63533/63533fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>1 x Phosphate-Buffered Saline (PBS)</td><td></td><td></td><td>Include sodium chloride and phosphate buffer and is formulated to prevent osmotic shock and maintain water balance in living cells.</td></tr><tr><td>AFM software</td><td>Bruker, Billerica, MA, USA</td><td></td><td></td></tr><tr><td>Atomic force microscopy (AFM)</td><td>BioScope Catalyst, Bruker, Billerica, MA, USA</td><td></td><td></td></tr><tr><td>Catalyst Probe holder-fluid</td><td>Bruker, Billerica, MA, USA</td><td>CAT-FCH</td><td>A probe holder for the Bioscope Catalyst, designed for fluid operation in contact or Tapping Mode.&amp;nbsp; Also compatible with air operation.</td></tr><tr><td>Cryoscopic osmometer; model OSMOMAT 030</td><td>Gonotech, Berlin, Germany</td><td></td><td></td></tr><tr><td>Murashige &amp;amp; Skoog Medium</td><td>Duchess Biochemie</td><td>M0221</td><td>Original concentration, (1962)</td></tr><tr><td>Optical inverted microscope coupled to the AFM</td><td>Olympus IX81, Miami, FL, USA</td><td></td><td></td></tr><tr><td>PEGAMIL</td><td>ANAEROBICOS S.R.L., Buenos Aires, Argentina</td><td>100429</td><td>Neutral, non acidic silicone glue</td></tr><tr><td>Petri dishes</td><td>Deltalab</td><td>200201.B</td><td>Polystyrene, 55 x 14 mm, radiation sterile.</td></tr><tr><td>Propidium iodide</td><td>Sigma</td><td>P4170</td><td>For root viability test.</td></tr><tr><td>Silicon nitride probe, DNP-10, cantilever A</td><td>Bruker, Billerica, MA, USA</td><td>DNP-10/A</td><td>For force modulation microscopy in liquid operation. Probe tip radius of 20-60 nm. 175-&amp;mu;m-long triangular cantilever,&amp;nbsp; with a spring constant of 0.35 N/m.</td></tr><tr><td>Tweezers</td><td>Sigma</td><td>T4537</td><td></td></tr></tbody></table>

atomic force microscopy to study the physical properties of epidermal cells of live arabidopsis roots

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through nanoindentations with an atomic force microscope (AFM) coupled with an optical inverted fluorescence microscope. The method consists of applying controlled forces to the sample while measuring its deformation, allowing quantifying parameters such as the apparent Young's modulus of cell walls at subcellular resolutions. It requires a careful mechanical immobilization of the sample and correct selection of indenters and indentation depths. Although it can be used only in external tissues, this method allows characterizing mechanical changes in plant cell walls during development and enables the correlation of these microscopic changes with the growth of an entire organ.

Force-Indentation experiments 
The following text presents some results expected when a force-indentation experiment is conducted to show the typical output to expect when the protocol is well executed.
Force-displacement curves 
Representative force indentation plots that were obtained indenting live samples at a position placed in the center of the cell of the root elongation zone are presented in Figure 2. When the AF...

Watch this Scientific Journal Video about Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots at JoVE.com

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

The atomic force microscopy indentation protocol offers the possibility to dissect the role of the physical properties of the cell wall of a particular cell of a tissue or organ during normal or constrained growth (i.e., under water deficit).

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through ...

atomic-force-microscopy-to-study-physical-properties-epidermal-cells

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

2.0K Views.  Universidad de la República (UdelaR). The atomic force microscopy indentation protocol offers the possibility to dissect the role of the physical properties of the cell wall of a particular cell of a tissue or organ during normal or constrained growth (i.e., under water deficit).

Video: Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Bioengineering

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 ...

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in indentation mode, we show how to measure an elastic modulus and use it to evaluate cell wall mechanical properties. In addition, we also explain how to evaluate turgor pressure. The main advantages of atomic force microscopy are that it is non-invasive, relatively rapid (5~20 min), and that virtually any type of living plant tissue that is superficially flat can be analyzed without the need for treatment. The resolution can be very good, depending on the tip size and on the number of measurements per unit area. One limitation of this method is that it only gives direct access to the superficial cell layer.

Here, we present atomic force microscopy (AFM), operated as a nano- and micro-indentation tool on cells and tissues. The instrument allows the simultaneous acquisition of 3D surface topography of the sample and its mechanical properties, including cell wall Young's modulus as well as turgor pressure.

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in ...

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

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.

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

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.

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of plant leaves or vertebrate lungs further difficulties arise from multiple transitions in refractive index between cellular components, between cells and airspaces and between the biological tissue and the rest of the optical system. Moreover, refractive index mismatches lead to attenuation of fluorophore excitation and signal emission in fluorescence microscopy. We describe here the application of the perfluorocarbon, perfluorodecalin (PFD), as an infiltrative imaging medium which optically improves laser scanning confocal microscopy (LSCM) sample imaging at depth, without resorting to damaging increases in laser power and has minimal physiological impact2. We describe the protocol for use of PFD with Arabidopsis thaliana leaf tissue, which is optically complex as a result of its structure (Figure 1). PFD has a number of attributes that make it suitable for this use3. The refractive index of PFD (1.313) is comparable with that of water (1.333) and is closer to that of cytosol (approx. 1.4) than air (1.000). In addition, PFD is readily available, non-fluorescent and is non-toxic. The low surface tension of PFD (19 dynes cm-1) is lower than that of water (72 dynes cm-1) and also below the limit (25 - 30 dyne cm-1) for stomatal penetration4, which allows it to flood the spongy mesophyll airspaces without the application of a potentially destructive vacuum or surfactant. Finally and crucially, PFD has a great capacity for dissolving CO2 and O2, which allows gas exchange to be maintained in the flooded tissue, thus minimizing the physiological impact on the sample. These properties have been used in various applications which include partial liquid breathing and lung inflation5,6, surgery7, artificial blood8, oxygenation of growth media9, and studies of ice crystal formation in plants10. Currently, it is common to mount tissue in water or aqueous buffer for live confocal imaging. We consider that the use of PFD as a mounting medium represents an improvement on existing practice and allows the simple preparation of live whole leaf samples for imaging.

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

We describe the use of perfluorodecalin as an infiltrative mounting medium. This is a simple method for improving depth of imaging in Arabidopsis thaliana leaf tissue with minimal physiological impact.

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of ...

Biology

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

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Chapters in this video

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

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

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

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

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions