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 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.
Atomic force microscopy (AFM) belongs to the scanning probe microscopy (SPM) family, where a tip with a radius of usually a few nanometers scans the surface of a sample. The detection of a surface is not achieved via optical or electron-based methods, but via the interaction forces between the tip and the sample surface. Thus, this technique is not limited to topographic characterization of a sample surface (3D resolution which can go down to a few nanometers), but also allows the measurement of any type of interaction forces such as electrostatic, van der Waals or contact forces. Furthermore, the tip can be used to apply forces at the surface of a biological sample and measure the resulting deformation, the so-called "indentation", in order to determine its mechanical properties (e.g., Young's modulus, viscoelastic properties).
Mechanical properties of plant cell walls are essential to be taken into account when trying to understand mechanisms underlying developmental processes1,2,3. Indeed, these properties are tightly controlled during development, in particular since cell wall softening is required to allow cells to grow. AFM can be used to measure these properties and study the way they change between organs, tissues or developmental stages.
In this paper, we describe how we use AFM to measure both cell wall mechanical properties and turgor pressure. These two applications are demonstrated on two different AFM microscopes and are detailed here after.
1.Measure of Cell Wall Mechanical Properties
NOTE: Example of the developing gynoecium of Arabidopsis is presented.
2. Measure of Turgor Pressure
NOTE: An example of the oryzalin-treated inflorescence meristem of Arabidopsis is presented.
Figure 1A and Figure 1B show a screenshot illustrating the result of the steps 1.3.4 to 1.3.6 of the protocol, used to locate a region of interest where to acquire the QI map. It is worth mentioning that the region of interest has been chosen in order not to be on a tilted surface (i.e., as flat as possible). Actually, as noticed by Routier et al.5, if the indentation axis is not perpendicular to the surface, the Young's modulus measured can be underestimated. This effect is visible on the upper-right corner of C or D panels of Figure 1, where part of the cells' border looks softer than the rest of the cell. The artefacts induced by this effect will be far stronger in the case of meristems, where the local tilt angle can easily overcome 40° over 50 x 50 µm2 scan areas. In our laboratory, we are currently developing an algorithm, based on the approach described by Routier et al.5, to correct those artefacts (paper in preparation). Figure 1C and Figure 1D show Young's modulus maps obtained after the analysis detailed in the fourth paragraph of the protocol. In particular, panel C represents the Young's modulus obtained analyzing the whole indentation, up to the user-defined force setpoint, while panel D shows the result of the analysis of the first 100 nm of indentation (as described in step 1.4.9). Here, the 2 maps look highly similar, because during the experiment set-up, the setpoint has been chosen in order to obtain average indentations around 100 nm. The variation of the analyzed indentation can sometimes lead to better highlight sample heterogeneities, that can be helpful for identifying the location or for providing information on the behavior of internal structures (e.g., Costa et al.10)
The force curve in Figure 2 shows two effects that are worth mentioning. First of all, it can be noticed that if the approach part of the force curve actually ends at the setpoint force of 500 nN, the downward tip movement keeps on going, meaning the final force applied by the tip is higher than expected (here 1,000 nN). This is due to the fact that the feedback loop monitoring the cantilever deflection does not act instantaneously, so its finite reaction time introduces a lag between deflection threshold detection and piezo movement stop. Furthermore, especially when using CellHesion which moves the sample holder, inertia of the moving parts comes into play, making this time lag longer. This issue can be limited by using the head piezo, in which case the user must be really careful in the case of measurements on very rough or tilted samples, or by reducing the ramp velocity, which anyway limits the resolution and/or the number of samples scanned (furthermore for too slow maps, the sample growing can become an issue). Generally, if the force curves are far enough from one another (based on the indentation model of choice, the radius of the indented area can be calculated given the indentation depth and used as minimum separation distance), the measurements performed in different positions along the sample should not be correlated and if the analysis is performed on the approach curve, overcoming the force threshold should not be an issue. Anyway, if the sample is delicate or if repeated scans are done on the same area, the scientist may want to try to minimize this overshoot. Unfortunately, there is not a rule of thumb to determine the amount of this overshoot, since it depends on the piezo used, on the ramp velocities, but also on material properties: the stiffer the material, the faster the variation of the deflection signal in time and, given a finite feedback response time, the higher the overshoot.
The second thing to notice is the waving on retract curve highlighted by the ellipse. Such waving can be an indicator of sample moving/vibrating under the action of the tip. In this case, the user can try to change the scanned area (sometimes other part of the microscope but the tip can touch the sample, or the sample can be insufficiently well fixed to its support) or the sample if several of them have been prepared on the same support. If the feature is always there, it is better to change the fixation method, in this case switching from the double-sided tape fixation to biocompatible glues.
Figure 3 depicts the determination of key parameters for turgor pressure deduction. Every parameter, except for cell wall thickness11 which was separately determined by electron microscopy to be ~740 nm, can be retrieved from AFM scans and indentations. Following the fitting process in step 2.4.10, the turgor pressure is deduced to be 1.50 MPa for this force curve. Pooling all eighteen force curves in this cell yields average values of E = 6.77 ± 1.02 MPa, k = 14.18 ± 2.45 N/m and P = 1.45 ± 0.29 MPa (mean ± standard deviation).
Figure 1: Topography maps (Height Measured) typically acquired during an experiment. (A) A first low resolution/low force map is recorded to find a suitable region to scan. (B) A smaller high resolution/higher force map is acquired for the calculation of Young's modulus (on the area highlighted by the black dotted square). (C) Young's modulus maps calculated from B map, as detailed in step 1.4. Here, the Young's modulus map has been obtained analyzing the whole indentation on each force curve. (D) Young's modulus map obtained analyzing only the first 100 nm of indentation. Please click here to view a larger version of this figure.
Figure 2: Example of force curve. In blue the approach, in red the retract segment. Force maximum (located on retract curve) is different from the force setpoint because of finite feedback loop reaction time and because of inertia. The waving in retract curve can be representative of a deficiency in sample fixation to its support. Please click here to view a larger version of this figure.
Figure 3: AFM measurements for turgor pressure deduction. (A) DMT modulus map shows clear cell contour to supervise the positioning of deep-indentation sites near cell barycenter. Indentation sites are marked by crosses. (B) Force curve of deep indentation at the red cross position in A. Cell wall Young's modulus E (green) and sample apparent stiffness k (red) are fitted at different regimes of the curve as detailed in step 2.4. R2 denotes coefficient of determination of the fits. (C) Height map with manually determined long and short axes of a, depicted by white lines, of the same cell analyzed in panels A and B. (D) Height profile of the long and short axes of the cell in Panel C (blue, long axis red, short axis) and the fitted radii of curvature (r1 and r2). Please click here to view a larger version of this figure.
The emergence of shapes in plants is mainly determined by the coordinated rate and direction of growth during time and space. Plant cells are encased in a rigid cell wall made of a polysaccharidic matrix, which glues them together. As a result, cell expansion is controlled by the equilibrium between turgor pressure pulling on the cell wall, and stiffness of the cell wall resisting to this pressure. In order to understand the mechanisms underlying development, it is important to be able to measure both cell wall mechanical properties as well as turgor pressure in different tissues or cells of a given organ. As demonstrated in this paper, AFM is the method of choice in this context.
There are several critical steps in the protocol. The first one is the tissue preparation which should be fast enough to avoid dehydration. This can be critical in particular if we want to measure turgor pressure. Very important is also a correct fixation of the sample to its substrate: an unstable sample can lead to very obvious effects like a macroscopic movement that can be easily detected optically, or a strong deformation of force curves. Anyway, sample vibration or bending can be subtle to detect, introducing artefacts into the results. Finally, another critical step concerns the choice of the indentation parameters. This is determinant for the quality of the results.
When measuring mechanical properties on large areas (over 40-50 µm scan sizes), height differences can be high, making it difficult to properly track the surface and realizing force curves without reaching the limits of Z piezo range. In order to overcome this issue, a CellHesion module has been used. This module adds an additional Z piezo, located into the sample stage, having a 100 µm range, which allows large areas acquisition without approaching dangerously Z piezo limits.
The first limitation of the method is that we can only measure superficial cell layers. However, recently authors have used AFM on tissue sections12,13,14. Although this must be done on fixed material, it can give access to mechanical properties of deep cell layers. Alternatively, deeper indentations have also been performed to infer mechanical properties of internal tissues on living samples, albeit with bigger sacrifice on spatial resolution15. A second limitation can be linked to sample geometry, as a surface with a steep slope can be problematic since the tip is no more perpendicular to the sample. We need to restrict the range of angles within which measures are reliable and we are currently developing an algorithm to correct potential artefacts linked to this phenomenon. Also, some tissues are covered with trichomes which can block the measurements. Finally, indentation measures mechanical properties in a perpendicular direction regarding the cell surface and we can wonder if this can be easily linked to cell wall extensibility associated to growth capacity. Several studies suggest it is the case15,16.
It is worth doing here a more general observation about the measurement/application of forces by AFM. As described in the protocol section, in order to be able to transform laser displacements on the photodiode to forces, a calibration has to be performed. In this calibration one parameter, the deflection sensitivity, allows to convert laser displacements to actual cantilever deflection (generally measured in nm), so this factor calibrates photodiode output voltages (along the vertical axes) to displacements in nm. The second factor is the spring constant K, measured in N/m, which converts vertical cantilever deflections in force units (generally nN), since the flexible cantilevers behave like springs. Even if the procedure appears simple, a robust and reproducible calibration of K for AFM force spectroscopy measurements is still an issue, because of different error sources that can affect the different steps of the procedure. For example, the measurement of the deflection sensitivity realizing a force curve on a stiff surface, can lead to incorrect values if the tip slips on the surface, or if the surface is not perfectly cleaned or also if surface charge induce electrostatic repulsion. In all of the previous cases, the contact part of the force curve will be deformed (further tilted or curved instead of linear).
The contact procedure described in the protocol section is only one of several existing calibration procedures. There are at least 2 more methods that can be used in a relatively easy and inexpensive way: the Sader method (also called non-contact when implemented in some AFM software) or the reference cantilever method. Sader method17 has been originally developed by John Sader for V-shaped cantilevers and then for rectangular ones18 and does not need the acquisition of a force curve on a stiff substrate. Instead, the user needs to specify (out of the temperature as in the contact method), the length and width of the cantilever and the density and viscosity of the fluid where the cantilever is (generally air, water or a water-like fluids). Then a thermal spectrum is acquired and both deflection sensitivity and spring constant are measured. This method is advantageous when using very sharp or functionalized tips that may be damaged doing a force curve on a stiff substrate.
Both the previous methods relate to the acquisition of the oscillation spectrum of a thermally-excited cantilever. When stiff cantilevers are used at room temperature, the mean oscillation amplitude can be lower than 0.1 nm, making the resonance peak smaller and more difficult to detect. Anyway, at least for both instruments used in this paper, the resonance peak can actually be detected in air and in water and fitted for measuring the spring constant (considering that when the measurement is done in liquid, the resonance frequency shifts toward lower values and the peak broadens).
A third method doesn't use a thermal tune, but instead a reference cantilever or a reference elastic structure, with a known spring constant (generally with an uncertainty around or below 5%), in order to determine the cantilever K. In this case a first force curve needs to be acquired on a stiff substrate in order to measure the deflection sensitivity (which comes again along with all possible artefacts affecting the shape of curve). Then the stiff substrate is replaced by the reference cantilever (generally a tip-less cantilever with a calibrated spring constant) and another (or few others) force curve is performed, recording again the value of the deflection sensitivity. The cantilever spring constant is then calculated as:
where Kref is the spring constant of the reference cantilever, Sref the deflection sensitivity measured on it, L is its length and Sstiff is the deflection sensitivity measured on the stiff substrate. ΔL is the offset between the tip and the end of the reference cantilever and depends on the alignment done by the user and finally α is the tilt angle of the cantilever, specified by the AFM manufacturer. The same method can be used on elastic structures specifically designed for cantilever or indenter calibration. The advantage of those structures (that are generally circular) is that K at their center is constant and doesn't depend on any geometrical parameter or on the precise positioning of the AFM tip on them, so removing L and ΔL from the previous formula.
The AFM user has to keep in mind that all those calibration methods are affected by different error sources (deflection sensitivity determination in first and third, the use of thermal tune in the case of stiff cantilevers for first and second and geometrical parameters and alignment for the third as well as calibration accuracy of Kref in the case reference cantilever is used) and that contact methods are potentially harmful for the tip (especially the third, where calibration imposes the use of two different samples). This implies that the spring constant will always be determined up to a certain precision and so will be the measured/applied forces (see Sikora19 for a comprehensive review of different calibration methods and their precision. This means that Young's moduli and turgor pressures will also be affected by cantilever calibration. Anyway, other even more important error sources are involved when calculating those quantities (like the use of simplified models for Young's modulus determination) so it will always be a challenge to obtain absolute values out of those kinds of measurements. What is important is to obtain values that are of the right order of magnitude and that are coherent with each other, meaning that if the experiment is repeated by the same user on the same sample, the values should be compatible. In order to obtain it, the calibration must be done carefully, and error sources must be minimized (for example limiting as much as possibly acoustic noise/vibrations). One possible strategy focused to increasing the coherence of repeated calibrations has been recently proposed in a paper by Schillers et al.20, where thanks to the collaboration of 11 European labs, a protocol (called SNAP) to compensate for errors in deflection sensitivity determination has been developed. In this paper the authors use directly calibrated cantilevers for their measurements, anyway the same protocol can be applied to non-calibrated cantilevers, simply calibrating them the first time with the method of choice and then considering the first spring constant as the reference ("calibrated") value.
The authors have nothing to disclose.
We would like to thank the PLATIM team for their technical support, as well as Arezki Boudaoud and members of the Biophysic team at the RDP lab for helpful discussions.
Growth medium | |||
1000x vimatin stock solution | used to make ACM, composition see Stanislas et al., 2017. Add to ACM after autoclaving, before pouring. | ||
1-N-Naphthylphthalamic acid (NPA) | Sigma-Aldrich/Merck | 132-66-1 | add to Arabidopsis medium, 10 μM. Add after autoclaving, before pouring. |
agar-agar | Sigma-Aldrich/Merck | 9002-18-0 | add to Arabidopsis medium, 1% w/v. |
agarose | Merck Millipore | 9012-36-6 | used to make solid ACM, 0.8% w/v. |
Arabidopsis medium | Duchefa Biochimie | DU0742.0025 | For in vitro arabidopsis culture, 11.82g/L. |
Calcium nitrate tetrahydrate | Sigma-Aldrich/Merck | 13477-34-4 | add to Arabidopsis medium, 2mM. |
MURASHIGE & SKOOG MEDIUM | Duchefa Biochimie | M0221.0025 | Basal salt mixture, used to make ACM, 2.2g/L. |
N6-benzyladenine (BAP) | Sigma-Aldrich/Merck | 1214-39-7 | used to make ACM, 555 nM. Add to ACM after autoclaving, before pouring. |
oryzalin | Sigma-Aldrich/Merck | 19044-88-3 | for oryzalin treatement, 10 μg/mL. |
plant preservation mixture (PPM) | Plant Cell Technology | used to make ACM, 0.1% v/v. Add to ACM after autoclaving, before pouring. | |
Potassium hydroxide | Duchefa Biochimie | 1310-58-3 | used to make Arabidopsis medium and ACM, both pH 5.8. |
sucrose | Duchefa Biochimie | 57-50-1 | used to make ACM, 1% w/v. |
Tools for AFM | |||
BioScope Catalyst BioAFM | Bruker | The AFM used for turgor pressure measurement in this protocol. | |
Nanowizard III + CellHesion | JPK (Bruker) | The AFM used for measuring mechanical properties. | |
Patafix | UHU | D1620 | |
Reference elasitic structure | NanoIdea | 2Z00026 | |
Reprorubber-Thin Pour | Flexbar | 16135 | biocompatible glue. |
Spherical AFM tips | Nanoandmore | SD-SPHERE-NCH-S-10 | Tips used for measuring mechanical properties. |