We would like to acknowledge Dr. Dmitry Suslov (Saint Petersburg State University, Saint Petersburg, Russia) and Prof. Mira Ponomareva (Tatar Scientific Research Institute of Agriculture, FRC KazSC RAS, Kazan, Russia) for providing maize and rye seeds, respectively. The presented method was developed within the framework of the Russian Science Foundation Project No. 18-14-00168 awarded to LK. The part of the work (obtaining of the results presented) was performed by AP with the financial support of the government assignment for the FRC Kazan Scientific Center of RAS.

1. Sample preparation for AFM measurements
<ol>
	<li>Plant material: Sterilize the seeds of maize (Zea mays L.) and rye (Secale cereale L.) with a 0.35% NaOCl solution for 10 min, wash 3x with distilled water, and then grow hydroponically in the dark at 27 &#176;C for 4 days and 2 days, respectively. Primary roots were used for the experiment.</li>
	<li>Preparation of solutions and sample for vibratome sectioning
	<ol>
		<li>Prepare agarose solution for root embedding by dissolving 3% (w/w) low-melting-point agarose in water using a microwave oven. 
		NOTE: All experiments were performed in water. If buffer or any other type of medium is required for the study, it is better to use the same medium for agarose preparation, during sectioning, and in the liquid cell of the AFM to avoid damaging the sample.</li>
		<li>Pour a 4 mm layer of melted 3% agarose on the bottom of the Petri dish (diameter = 35 mm), and let it cool slightly to prevent thermal damage to the sample.</li>
		<li>Place three or four small pieces (about 5 mm long) of the studied plant organ horizontally on the agarose. 
		NOTE: Samples should be half-immersed but not submerged in the agarose layer. If the sample is not immersed, it may break out of the agarose during vibratome sectioning. If the sample sinks to the bottom, it will be too close to the edge of the section and become inconvenient for further steps.</li>
		<li>When a thin, semi-solid film appears on top of the first layer of agarose (30-60 s), carefully pour a second layer on top. 
		NOTE: The bottom layer of agarose should not completely solidify. Otherwise, the two layers may separate from each other during further work.</li>
		<li>After the agarose is completely solidified, cut out the block containing the specimen. Shape the block into a hexagonal truncated pyramid to ensure its stability during further sectioning (Figure 1A). 
		NOTE: The sample can be oriented vertically or horizontally within this block depending on the type of section required. When preparing the block, leave 2-3 mm of agarose around the specimen. In this case, the sample section will be retained by the layer of 3% agarose, which will facilitate its further immobilization (Figure 1B).</li>
	</ol>
	</li>
	<li>Sectioning the sample with the vibratome
	<ol>
		<li>Glue the block to the vibratome stage with cyanoacrylate adhesive. Place the stage in the vibratome so that one of the pyramid corners faces the blade of the vibratome. Pour water into the vibratome bath.</li>
		<li>Set the sectioning parameters (section thickness, blade speed, and vibration frequency), and cut the sample. 
		NOTE: Use the section thicknesses between 200 and 400 &#181;m. A section that is too thin can introduce errors in the evaluation of mechanical properties, while a section that is too thick is prone to breakage. A blade speed of 1.3 mm&#183;s-1 and an oscillation frequency of 90 Hz were used for rye and maize roots.</li>
		<li>Using a fine brush, move the section from the water bath onto a glass slide, and place a drop of water on the section to prevent it from drying out. Check the quality of the section under a light microscope. 
		NOTE: Oblique sections cannot be used to measure the modulus of elasticity. Cell walls must be perpendicular to the section plane, otherwise they may bend or buckle under the AFM tip.</li>
	</ol>
	</li>
	<li>Immobilizing the section for AFM measurements (Figure 1B)
	<ol>
		<li>Pour a layer of molten 1% (w/w) agarose (~1 mL) on the bottom of the Petri dish cap using a pipette. 
		NOTE: Make sure that the sides of the Petri dish cap do not prevent the tip from approaching the sample. The agarose layer should be 1 mm thick and should cover the entire bottom of the cap.</li>
		<li>After the 1% agarose has solidified, remove excess water from the section by bringing filter paper to its edge. 
		NOTE: Do not touch the plant section with the paper to avoid damage and complete drying of the sample.</li>
		<li>Carefully transfer the section from the slide to the center of the Petri dish cap using a brush. Using a 20 &#181;L pipette, carefully add 1% agarose around the section (Figure 1B). 
		NOTE: The 1% agarose should not get on the specimen itself. It should only cover the edges of the 3% agarose layer that retains the plant specimen. The 1% agarose should form small knolls on the edges of this 3% agarose layer. Large knolls may prevent the cantilever from approaching the sample.</li>
		<li>Pour the water or other solution to be used for the AFM into the Petri dish cap with the immobilized section.</li>
	</ol>
	</li>
</ol>
2. AFM preparation and calibration
NOTE: The force-volume method of AFM generates a spatially resolved array of force-distance curves obtained at each point of the area studied. Obtain all parameters for the force-volume mode (cantilever stiffness, IOS, etc.) in contact mode. Similar procedures for instruments from other manufacturers have been described previously10,20.
<ol>
	<li>Select the appropriate type of cantilever. 
	NOTE: The cantilever stiffness must be comparable to the specimen stiffness21. A cantilever that is much stiffer than the specimen will not deflect much, while a cantilever that is too soft will not deform the sample enough. A typical resonance frequency should be sufficient to swing the cantilever in the liquid (i.e., be at least a few tens of kHz). The contact area should be small compared to the size of the cell wall since the sample under study in Young&#39;s modulus calculations is assumed to be an infinite half-space. The following cantilevers were used for rye and maize roots: sharp cantilevers with a typical resonance frequency of 60 kHz, an average spring constant of 1.5 N&#183;m-1, and an apex radius of 10 nm, or spherical cantilevers with a typical resonance frequency of 75 kHz, an average spring constant of 2.8 N&#183;m-1, and an apex radius of 150 nm.</li>
	<li>Switch on the AFM device and the associated software (Table of Materials). Mount the cantilever on the tip holder for the liquid cell. Place a drop of liquid on the tip to avoid air bubbles forming on the tip while immersing it in the liquid. 
	NOTE: In case of bubble formation, the liquid can be removed with a precision wipe, and then a new drop can be placed.</li>
	<li>Mount the tip holder on the scanning head.</li>
	<li>Place a hard sample (fresh glass slide) into a Petri dish cap. Pour the liquid into it, ensuring it covers the slide. Place the scanning head on the stage and raise the specimen so that the liquid covers the cantilever.</li>
	<li>Choose the Heads tab in the drop-down Tools menu. Make sure that the scanning head for the liquid cell is selected.</li>
	<li>Click on the Aiming button to open the Laser Aiming window. Click on the Camera button to open the Optical Microscope window. Position the laser at the tip of the cantilever, using the screws on the AFM head and the Optical Microscope window for orientation.</li>
	<li>Choose the Semicontact mode from the drop-down menu in the main window of the program.</li>
	<li>Open the Resonance tab and choose the type of cantilever being used in the Probes menu. Click on the Auto button to determine the resonance frequency. 
	NOTE: If the cantilever being used is not listed, open Tools &#62; Probe Passport. Create a new file, enter the cantilever parameters, and save it. Then choose it in the Probes drop-down menu.</li>
	<li>Choose the Contact mode from the drop-down menu in the main window of the program.</li>
	<li>Click on the N_Force Cal button to open cantilever calibration window. Choose the cantilever being used and click on the Sweep button, and then the Spect Meas button to determine the cantilever spring constant using the thermal-tune procedure. Close the window.</li>
	<li>Set the SetPoint to 1 nA. Open the Approach tab and click on the Landing button to approach the sample.</li>
	<li>Open the Scanning tab. Set the scan Rate to 0.5 Hz. Click on the Area button and set the scan Size to 10 &#181;m x 10 &#181;m and the scan Point to 256 x 256. Click on the Run button and scan about 20 lines to check that there is no contamination on the glass. Click on the Stop button to end the scan without losing data.</li>
	<li>Open the Curves tab. Set parameters for retract and approach to 500 nm and -100 nm, respectively.</li>
	<li>Choose the last scan in the drop-down menu Select Frame to observe the surface.</li>
	<li>Find a clean area on the glass and indicate the point where the force-deformation curve should be taken, and click on the Run button to get the curve. Record three to five force curves in different places on the scan.</li>
	<li>Click on the Data button to open the analysis window and select the frame with the force-deformation curve.</li>
	<li>Click on the Pair Markers button and indicate the linear part of the retraction curve to calculate the ratio of the DFL signal to the displacement, which is the Inverse Optical Sensitivity (IOS, nA nm-1) or deflection sensitivity.</li>
	<li>Repeat step 2.17 for all recorded curves and write down all calculated values of the DFL signal to displacement ratio. They should be the same.</li>
	<li>Close the analysis window, click on the Approach tab, and then the Remove button to retract the probe from the sample.</li>
</ol>
3. Data acquisition
<ol>
	<li>Guide the sample under the AFM cantilever using the optical microscope. Click on the Approach tab, and then the Landing button to approach the sample in contact mode with a SetPoint of 1 nA.</li>
	<li>Click on the Scanning tab, and then the Area button. Select the Area Size of 70 &#181;m x 70 &#181;m to scan.</li>
	<li>Click on the Move Probe button and check the entire scan area by moving the scanner over it. Based on the degree of scanner protrusion, find the highest point.</li>
	<li>Open the Approach tab, then click on the Remove button to retract from the sample. Using the highest point as a target, click on the Landing button to approach the sample again. Then, check the surface again by clicking on the Move Probe button and moving the scanner over it. None of the points in the area should require the scanner to be fully raised. 
	NOTE: Ideally, the scanner z-range should exceed the height difference of the sample. However, it is acceptable if some points on the scan area are below the scanner&#39;s capacity. At the same time, there should not be many such points. Large areas which cannot be reached by the scanner may cause the scanning abortion.</li>
	<li>Set the scan Rate to 0.5 Hz. Set the scan Size to 70 &#181;m x 70 &#181;m and the scan Point to 64 x 64. Click on the Run button and scan to check the surface of the sample and its possible contamination with agarose. 
	NOTE: If the sample has cells with a large lumen, the size of the scan area can be reduced, or it should be moved so that there are more cell walls on the scan.</li>
	<li>Click on the On button at the top of the main window to switch off the feedback loop.</li>
	<li>Choose the HDPlus mode (force-volume method) in the drop-down menu in the main window of the program. An additional window (HD window) will appear.</li>
	<li>Set the SetPoint to 0.1 nA in the main program window.</li>
	<li>In the Main tab of the HD window, set the scanning parameters (amplitude and frequency of sinusoidal cantilever movement) appropriate to the studied sample. 
	NOTE: Each type of sample and cantilever requires the selection of scanning parameters. Some preliminary experiments are necessary. For maize or rye roots, sharp and spherical cantilevers were used at an amplitude of 400 nm and a frequency of 300 Hz.</li>
	<li>Open the Noises tab in the HD window and enter the resonance frequency of the cantilever.</li>
	<li>Open the Quant tab of the HD window and enter the IOS, cantilever Stiffness, and tip radius and angle. Select the model of contact which will be used for calculations depending on tip geometry. 
	NOTE: DMT model takes into account the adhesion force existing between the probe and the sample, and is most commonly used in mechanobiology21.</li>
	<li>Open the Scan tab of the HD window and select the signals to be recorded. Select the direction in which the signal is recorded. Tick the Force volume box to get a record of all force curves. 
	NOTE: Record the elasticity modulus signal in both forward and backward directions. It will provide more points to calculate.</li>
	<li>Click on the Off button at the top of the main program window to switch on the feedback loop.</li>
	<li>Click on the PhaseCorr button in the main HD window to correct the sensitivity of the optical system.</li>
	<li>The vs Time tab of the main HD window presents the function of the DFL signal vs. time in real-time; select the parts of this function that will be used for baseline level determination and to fit the contact model for further calculations.</li>
	<li>Set the scan Point value to 256 x 256 in the main window of the program. Set the scan Rate to 0.2 Hz and click on the Run button to scan the sample.</li>
	<li>After scanning stops, click on the On button at the top of the main window to switch off the feedback loop.</li>
	<li>Choose Contact mode in the drop-down menu, open the Approach tab, and click on the Remove button to retract from the sample.</li>
	<li>Click on the Data button to open the analysis software and save the output.</li>
	<li>At the end of the measurement day, remove the cantilever tip holder from the head and carefully rinse it with ultrapure water several times. Each time, remove the water with a precision wipe.</li>
</ol>
4. Data evaluation and post-processing
NOTE: Do not rely on elastic modulus values calculated automatically. Since the surface varies greatly in height, many artifact curves should be expelled.
<ol>
	<li>Open the saved file in the analysis software.</li>
	<li>Select the HDForceVolume frame.</li>
	<li>Hold down the Ctrl key and select one visual frame obtained in the same direction of scanning. Click on the Load External Map button to see where the cell walls are located. 
	NOTE: Any signal map can be used as a visual frame, but using the DFL signal map (different instruments may refer it to as a deflection signal or error signal) may be the easiest way.</li>
	<li>Check InvOptSens (IOS) and cantilever Stiffness values in the Main tab.</li>
	<li>Open the Additional tab and check the tip parameters and contact model.</li>
	<li>Click on various points on the cell walls on the visual frame and select only those curves that are well described by the model. See Representative Results for details.</li>
	<li>Write down the obtained values.</li>
</ol>

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The authors have no conflicts of interest.

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth, and therefore the future size and shape of the plant. The AFM-based method presented here complements existing techniques which are used to study the properties of plant cell walls. It allows the elasticity of cell walls, which belong to the inner tissues of the plant, to be investigated. Using the presented method, the mechanical properties of cell walls in different tissues of the growing maize root were mapped, a...

The mechanical properties of the plant cell wall determine the shape of the cell and its ability to grow. For example, the growing tip of the pollen tube is softer than the non-growing parts of the same tube1. The primordia formation on Arabidopsis meristem is preceded by a local decrease in cell wall stiffness at the site of the future primordium2,3. The cell walls of Arabidopsis hypocotyl, which are parallel to the main growth axis and grow faster, are softer than those that are perpendicular to this axis and grow slower4,5. In the maize root, the transition of cells from division to elongation was accompanied by a decrease in elastic moduli in all tissues of the root. The moduli remained low in the elongation zone and increased in the late elongation zone6.
Despite the availability of various methods, the large arrays of biochemical and genetic information on cell wall biology obtained annually are rarely compared with the mechanical properties of cell walls. For example, mutants on cell wall-related genes often have altered growth and development4,7,8, but are rarely described in terms of biomechanics. One of the reasons for this is the difficulty of conducting measurements at the cellular and subcellular levels. Atomic force microscopy (AFM) is currently the primary approach for such analyses9.
In recent years, numerous AFM-based studies on plant cell wall biomechanics have been carried out. The mechanical properties of cell walls of the outer tissues of Arabidopsis2,3,4,5,10,11 and onion12, as well as of cultured cells13,14,15, have been investigated. However, the superficial cells of a plant may have cell walls whose mechanical properties differ from those of the inner tissues6. In addition, plant cells are pressurized by turgor which makes them stiffer. To get rid of the influence of turgor pressure, researchers have to use plasmolyzing solutions2,3,4,5,10,11 or decompose the values obtained into turgor and cell wall contributions12. The first approach leads to sample dehydration and changes the thickness and properties of the cell wall16, while the second approach requires additional measurements and complicated mathematics, and applies only to cells of relatively simple shape12. The cell wall properties of internal tissues can be evaluated on cryosections17 or sections of plant material impregnated with resin8. However, both methods involve dehydration and/or impregnation of samples, which inevitably leads to changes in properties. The properties of isolated or cultured cells are difficult to relate to the physiology of the whole plant. Both cultivation and isolation of plant cells can affect the mechanical properties of their cell walls.
The method presented here complements the aforementioned approaches. Using it, the primary cell walls of any tissue and at any stage of plant development can be examined. Sectioning and AFM observations were performed in liquid which avoids sample dehydration. The problem of turgor was solved as the cells are cut. The protocol describes work with maize and rye roots, but any other sample can be examined if it is suitable for vibratome sectioning.
The AFM studies described here were performed using the force-volume technique. Different instruments use different names for this method. However, the basic principle is the same; a force-volume map of the sample is obtained by a sinusoidal or triangular motion of the cantilever (or sample) to achieve a certain loading force at each analyzed point, while recording the cantilever deflection18. The result combines a topographic image of the surface and the array of force-distance curves. Each curve is used to calculate the deformation, stiffness, Young&#39;s modulus, adhesion, and energy dissipation at a specific point. Similar data can be obtained by point-by-point force-spectroscopy after scanning in contact mode19, although it is more time-consuming.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose, low melting point</td>
			<td>Helicon</td>
			<td>B-5000-0.1</td>
			<td>for sample fixation</td>
		</tr>
		<tr>
			<td>Brush</td>
			<td>-</td>
			<td>-</td>
			<td>for section moving</td>
		</tr>
		<tr>
			<td>Cantilevers</td>
			<td>NanoTools, Germany</td>
			<td>NT_B150_v0020-5</td>
			<td>Model: Biosphere B150-FM</td>
		</tr>
		<tr>
			<td>Cantilevers</td>
			<td>NT-MDT, Russia</td>
			<td>FMG01/50</td>
			<td>Model: FMG01</td>
		</tr>
		<tr>
			<td>Cyanoacrylate adhesive</td>
			<td>-</td>
			<td>-</td>
			<td>for vibratomy</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>Heinz Herenz</td>
			<td>1042000</td>
			<td>for vibratomy and AFM calibration</td>
		</tr>
		<tr>
			<td>ImageAnalysis P9 Software</td>
			<td>NT-MDT, Russia</td>
			<td>-</td>
			<td>for data analysis</td>
		</tr>
		<tr>
			<td>Leica DM1000 epifluorescence microscope</td>
			<td>Leica Biosystems, Germany</td>
			<td>11591301</td>
			<td>for section check</td>
		</tr>
		<tr>
			<td>NaOCl</td>
			<td>-</td>
			<td>-</td>
			<td>for seed sterilization</td>
		</tr>
		<tr>
			<td>Nova PX 3.4.1 Software</td>
			<td>NT-MDT, Russia</td>
			<td>-</td>
			<td>for experiments conducting</td>
		</tr>
		<tr>
			<td>NTEGRA Prima microscope with HD controller</td>
			<td>NT-MDT, Russia</td>
			<td>-</td>
			<td>for AFM and data acquisition</td>
		</tr>
		<tr>
			<td>Petri dish 35 mm</td>
			<td>Thermo Fisher Scientific</td>
			<td>153066</td>
			<td>for sample fixation</td>
		</tr>
		<tr>
			<td>Tip pipette 1000 &#181;L</td>
			<td>Thermo Fisher Scientific</td>
			<td>4642092</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Tip pipette 2-20 &#181;L</td>
			<td>Thermo Fisher Scientific</td>
			<td>4642062</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Ultrapure water</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Vibratome Leica VT 1000S</td>
			<td>Leica Biosystems, Germany</td>
			<td>1404723512</td>
			<td>for sample sectioning</td>
		</tr>
	</tbody>
</table>

characterizing mechanical properties of primary cell wall in living plant organs using atomic force microscopy

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the plant. Many sophisticated techniques have been developed to measure these properties; however, atomic force microscopy (AFM) remains the most convenient for studying cell wall elasticity at the cellular level. One of the most important limitations of this technique has been that only superficial or isolated living cells can be studied. Here, the use of atomic force microscopy to investigate the mechanical properties of primary cell walls belonging to the internal tissues of a plant body is presented. This protocol describes measurements of the apparent Young&#39;s modulus of cell walls in roots, but the method can also be applied to other plant organs. The measurements are performed on vibratome-derived sections of plant material in a liquid cell, which allows (i) avoiding the use of plasmolyzing solutions or sample impregnation with wax or resin, (ii) making the experiments fast, and (iii) preventing dehydration of the sample. Both anticlinal and periclinal cell walls can be studied, depending on how the specimen was sectioned. Differences in the mechanical properties of different tissues can be investigated in a single section. The protocol describes the principles of study planning, issues with specimen preparation and measurements, as well as the method of selecting force-deformation curves to avoid the influence of topography on the obtained values of elastic modulus. The method is not limited by sample size but is sensitive to cell size (i.e., cells with a large lumen are difficult to examine).

Typical elastic modulus and DFL maps, as well as force curves obtained on rye and maize roots by the method described, are presented in Figure&#160;2. Figure&#160;2A shows elastic modulus and DFL maps obtained on the transverse section of rye primary root. The white areas in the modulus map (Figure&#160;2A, left) correspond to an erroneous overestimation of Young&#39;s modulus due to the scanner reaching its limit in the z-direction...

Watch this Scientific Journal Video about Characterizing Mechanical Properties of Primary Cell Wall in Living Plant Organs Using Atomic Force Microscopy at JoVE.com

Characterizing Mechanical Properties of Primary Cell Wall in Living Plant Organs Using Atomic Force Microscopy

Studies of cell wall biomechanics are essential for understanding plant growth and morphogenesis. The following protocol is proposed to investigate thin primary cell walls in the internal tissues of young plant organs using atomic force microscopy.

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the ...

characterizing-mechanical-properties-primary-cell-wall-living-plant

Research

JoVE Journal

Biology

2.0K Views.  Kazan Institute of Biochemistry and Biophysics, FRC Kazan Scientific Center of RAS. Studies of cell wall biomechanics are essential for understanding plant growth and morphogenesis. The following protocol is proposed to investigate thin primary cell walls in the internal tissues of young plant organs using atomic force microscopy.

Video: Characterizing Mechanical Properties of Primary Cell Wall in Living Plant Organs Using Atomic Force Microscopy

Measuring Plant Cell Wall Extension (Creep) Induced by Acidic pH and by Alpha-Expansin

Growing plant cell walls characteristically exhibit a property known as 'acid growth', by which we mean they are more extensible at low pH (&lt; 5) 1. The plant hormone auxin rapidly stimulates cell elongation in young stems and similar tissues at least in part by an acid-growth mechanism 2, 3. Auxin activates a H+ pump in the plasma membrane, causing acidification of the cell wall solution. Wall acidification activates expansins, which are endogenous cell wall-loosening proteins 4, causing the cell wall to yield to the wall tensions created by cell turgor pressure. As a result, the cell begins to enlarge rapidly. This 'acid growth' phenomenon is readily measured in isolated (nonliving) cell wall specimens. The ability of cell walls to undergo acid-induced extension is not simply the result of the structural arrangement of the cell wall polysaccharides (e.g. pectins), but depends on the activity of expansins 5. Expansins do not have any known enzymatic activity and the only way to assay for expansin activity is to measure their induction of cell wall extension. This video report details the sources and preparation techniques for obtaining suitable wall materials for expansin assays and goes on to show acid-induced extension and expansin-induced extension of wall samples prepared from growing cucumber hypocotyls.
 To obtain suitable cell wall samples, cucumber seedlings are grown in the dark, the hypocotyls are cut and frozen at -80 &deg;C. Frozen hypocotyls are abraded, flattened, and then clamped at constant tension in a special cuvette for extensometer measurements. To measure acid-induced extension, the walls are initially buffered at neutral pH, resulting in low activity of expansins that are components of the native cell walls. Upon buffer exchange to acidic pH, expansins are activated and the cell walls extend rapidly. We also demonstrate expansin activity in a reconstitution assay. For this part, we use a brief heat treatment to denature the native expansins in the cell wall samples. These inactivated cell walls do not extend even in acidic buffer, but addition of expansins to the cell walls rapidly restores their ability to extend.

Measuring Plant Cell Wall Extension Creep Induced by Acidic pH and by Alpha-Expansin

We demonstrate the use of a constant-force extensometer to measure long-term extension (creep) of plant cell wall specimens induced by acidic buffers and expansin protein.

Growing plant cell walls characteristically exhibit a property known as 'acid growth', by which we mean they are more extensible at low pH (&lt; 5) 1. The ...

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3. On the macro scale the stiffness of tissues and organs within the human body span several orders of magnitude4. Much less is known about how stiffness varies spatially within tissues, and what the scope and spatial scale of stiffness changes are in disease processes that result in tissue remodeling. To better understand how changes in matrix stiffness contribute to cellular physiology in health and disease, measurements of tissue stiffness obtained at a spatial scale relevant to resident cells are needed. This is particularly true for the lung, a highly compliant and elastic tissue in which matrix remodeling is a prominent feature in diseases such as asthma, emphysema, hypertension and fibrosis. To characterize the local mechanical environment of lung parenchyma at a spatial scale relevant to resident cells, we have developed methods to directly measure the local elastic properties of fresh murine lung tissue using atomic force microscopy (AFM) microindentation. With appropriate choice of AFM indentor, cantilever, and indentation depth, these methods allow measurements of local tissue shear modulus in parallel with phase contrast and fluorescence imaging of the region of interest. Systematic sampling of tissue strips provides maps of tissue mechanical properties that reveal local spatial variations in shear modulus. Correlations between mechanical properties and underlying anatomical and pathological features illustrate how stiffness varies with matrix deposition in fibrosis. These methods can be extended to other soft tissues and disease processes to reveal how local tissue mechanical properties vary across space and disease progression.

The stiffness of the extracellular matrix strongly influences multiple behaviors of adherent cells. Matrix stiffness varies spatially throughout a tissue, and undergoes modification in various disease conditions. Here we develop methods to characterize spatial variations in stiffness in normal and fibrotic mouse lung tissue using atomic force microscopy microindentation.

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3.  On the macro scale the stiffness of tissues and organs ...

Glycan Profiling of Plant Cell Wall Polymers using Microarrays

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g. wood, paper, textile and biofuel industries)1,2. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3. Furthermore, their composition is also modified during development1, or in response to environmental cues4.
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5. However, relatively few of the biosynthetic genes have been functionally characterized 4,5. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9 have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11.
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 &amp; 2)10,11 that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12 and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.

A technique called Comprehensive Microarray Polymer Profiling (CoMPP) for the characterisation of plant cell wall glycans is described. This method combines the specificity of monoclonal antibodies directed to defined glycan-epitopes with a miniature microarray analytical platform allowing screening of glycan occurrence in a broad range of biological contexts.

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell behavior such as cell function, differentiation, and motility. However, as these processes are not homogeneous throughout the whole organ, it has become increasingly important to understand changes in the mechanical properties of tissues on the cellular level.
To be able to monitor the stiffening of collagen-rich areas within the liver lobes, this paper presents a protocol for measuring liver tissue elastic moduli with high spatial precision by atomic force microscopy (AFM). AFM is a sensitive method with the potential to characterize local mechanical properties, calculated as Young&#39;s (also referred to as elastic) modulus. AFM coupled with polarization microscopy can be used to specifically locate the areas of fibrosis development based on the birefringence of collagen fibers in tissues. Using the presented protocol, we characterized the stiffness of collagen-rich areas from fibrotic mouse livers and corresponding areas in the livers of control mice.
A prominent increase in the stiffness of collagen-positive areas was observed with fibrosis development. The presented protocol allows for a highly reproducible method of AFM measurement, due to the use of mildly fixed liver tissue, that can be used to further the understanding of disease-initiated changes in local tissue mechanical properties and their effect on the fate of neighboring cells.

We present a protocol to measure the elastic moduli of collagen-rich areas in normal and diseased liver using atomic force microscopy. The simultaneous use of polarization microscopy provides high spatial precision for localizing collagen-rich areas in the liver sections.

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell ...

Investigating Zinc Transport Mechanism in Mammalian Cells with FluoZin-3 Assay

Characterizing Mammalian Zinc Transporters Using an In Vitro Zinc Transport Assay

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured indirectly by determining the expression level of the transporter under different concentrations of Zn2+. This was done by utilizing immunohistochemistry, measuring mRNA in the tissue, or determining the cellular Zn2+ levels. With the development of intracellular Zn2+ sensors, the activities of zinc transporters are currently primarily determined by correlating changes in intracellular Zn2+, detected using fluorescent probes, with the expression of the Zn2+ transporters. However, even today, only a few labs monitor dynamic changes in intracellular Zn2+ and use it to measure the activity of zinc transporters directly. Part of the problem is that out of the 10 zinc transporters of the ZnT family, except for ZnT10 (transports manganese), only zinc transporter 1 (ZnT1) is localized at the plasma membrane. Therefore, linking the transport activity to changes in the intracellular Zn2+ concentration is hard. This article describes a direct way to determine the zinc transport kinetics using an assay based on a zinc-specific fluorescent dye, FluoZin-3. This dye is loaded into mammalian cells in its ester form and then trapped in the cytosol due to cellular di-esterase activity. The cells are loaded with Zn2+ by utilizing the Zn2+ ionophore pyrithione. The ZnT1 activity is assessed from the linear part of the reduction in fluorescence following the cell washout. The fluorescence measured at an excitation of 470 nm and emission of 520 nm is proportional to the free intracellular Zn2+. Selecting the cells expressing ZnT1 tagged with the mCherry fluorophore allows for monitoring only the cells expressing the transporter. This assay is used to investigate the contribution of different domains of ZnT1 protein to the transport mechanism of human ZnT1, a eukaryotic transmembrane protein that extrudes excess zinc from the cell.

Author Spotlight: Exploring Cellular Zinc Regulation Through ZnT1 Functionality 

Zinc transport has proven challenging to measure due to the weak causal links to protein function and the low temporal resolution. This protocol describes a method for monitoring, with high temporal resolution, Zn2+ extrusion from living cells by utilizing a Zn2+ sensitive fluorescent dye, thus providing a direct measure of Zn2+ efflux.

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured ...