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Imaging and quantitative data acquisition of biological cell walls with
Atomic Force Microscopy and Scanning Acoustic Microscopy
B. R. Tittmann and X. Xi
Department of Engineering Science and Mechanics, the Pennsylvania State University, University Park, PA, 16802 USA
This chapter demonstrates the feasibility of Atomic Force Microscopy (AFM) and High Frequency Scanning Acoustic
Microscopy (HF-SAM) as tools to characterize biological tissues. Both the AFM and the SAM have shown to provide
imaging (with different resolution) and quantitative elasticity measuring abilities. Plant cell walls with minimal
disturbance and under conditions of their native state have been examined with these two kinds of microscopy. After
descriptions of both the SAM and AFM, their special features and the typical sample preparation is discussed. The sample
preparation is focused here on epidermal peels of onion scales and celery epidermis cells which were sectioned for the
AFM to visualize the inner surface (closest to the plasma membrane) of the outer epidermal wall. The nm-wide cellulose
microfibrils orientation and multilayer structure were clearly observed. The microfibril orientation and alignment tend to
be more organized in older scales compared with younger scales. The onion epidermis cell wall was also used as a test
analog to study cell wall elasticity by the AFM nanoindentation and the SAM V(z) feature. The novelty in this work was to
demonstrate the capability of these two techniques to analyze isolated, single layered plant cell walls in their natural state.
AFM nanoindentation was also used to probe the effects of Ethylenediaminetetraacetic acid (EDTA), and calcium ion
treatment to modify pectin networks in cell walls. The results suggest a significant modulus increase in the calcium ion
treatment and a slight decrease in EDTA treatment. To complement the AFM measurements, the HF-SAM was used to
obtain the V(z) signatures of the onion epidermis. These measurements were focused on documenting the effect of
pectinase enzyme treatment. The results indicate a significant change in the V(z) signature curves with time into the
enzyme treatment. Thus AFM and HF-SAM open the door to a systematic nondestructive structure and mechanical
property study of complex biological cell walls. A unique feature of this approach is that both microscopes allow the
biological samples to be examined in their natural fluid (water) environment.
Keywords: Atomic Force Microscopy (AFM); High Frequency Scanning Acoustic Microscopy (HF SAM);
nanoindentation; V(z) signature curves; plant cell walls
1. Plant Cell walls
Plant cell walls serve a variety of important biological functions for plants, such as providing mechanical support and
determining their size and shape. They have also been an interesting subject for engineering applications. Plant cell
walls are essential in various commercial products such as paper, textile, plastic, etc. Most recently, they have gained
intensive attention as potential sources for biofuel. Also, the sophisticated composite structure, naturally occurring in
cell walls, inspires innovative designs of engineering materials. All these applications require understanding of the
composite structure of cell walls and their mechanical properties.
Primary cell walls are formed during the growth and division of plant cells. They provide mechanical support for the
cells and can expand to allow cells to grow. Primary plant cell walls are polysaccharide-rich complex structures. The
wall contains three main components: cellulose, hemicelluloses and pectin. Cellulose microfibrils are embedded in a
highly hydrated polysaccharide matrix which consists of hemicelluloses and pectin.[1-3] Cellulose is comprised of
many parallel chains of 1,4-glucan which make cellulose crystalline, strong and indigestible. Cellulose microfibrils
serve as reinforcement material of cell wall composite. Cellulose microfibrils are several nm in thickness and have
varied length. Hemicellulose is also a polysaccharide, but it is typically made up of chains of xylose interspersed with
side chains containing arabinose, galactose, mannose, glucose, acetyl, and other sugar groups, depending on the plant
type. [4-6] Hemicelluloses separate microfibrils from each other but may tether them together into a cohesive network.
Pectin is a gel phase that embeds the cellulose-hemicelluloses network. Pectin includes relatively simple
polysaccharides and becomes soluble with mild treatments such as boiling water or mildly acidic solutions. [7]
2. Atomic Force Microscopy
AFM was invented by Binnig, Quate and Gerber in 1986.[8] AFM can image topography and obtain structural and
mechanical information from samples’ surfaces at micrometer to nanometer levels. Most AFM can work in air, liquid or
vacuum environment to gather three dimension topographic images of the sample surface without complicated sample
preparations. This technique can be applied to different types of materials including semiconductors [9, 10], biological
systems [11-13]and polymers [10, 14-16]. Another important usage of AFM is to study surface properties as the force
spectroscopy by measuring forces as a function of distance.[17-20] AFM has been proven to be an effective tool for
quantitative studies of various surface properties, and has been used in some studies to measure surface adhesion
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forces.[21-24] AFM has become a powerful and promising tool in the development of micro-systems and
nanotechnology.
AFM has been used for more than a decade to examine the plant cell wall structures.[25-27] However, these studies
imaged dehydrated or partially dehydrated cell wall samples where surface structure during drying may greatly change.
In our work, fully hydrated epidermis cell walls were examined in peak force tapping mode in liquid environment. The
results reflect the properties of cell walls in their natural state.
2.1 AFM Imaging of hydrated cell walls
Onion abaxial epidermis walls and celery epidermis cell walls were prepared for AFM imaging as described by Zhang
et al [28]. Freshly peeled cell walls were fixed onto glass slides with edges glued. The epidermal strips were then
immersed in a phosphate buffered saline solution (pH 7.4) containing 0.1% Tween-20 for 1 h. After that, the samples
were rinsed with distilled water. Epidermal wall strips were probed by AFM (Veeco Dimension ICON from Bruker) in
Peak Force QNM fluid mode. Nanoscope (v 8.10b44) was used to control AFM operation and Nanoscope Analysis (v
1.40) was employed for image processing. Figure 1 shows a top view of AFM tip engaged in epidermis cell walls.
Fig. 1 A typical experimental view during imaging and nano-indentation of hydrated
onion cell with AFM. Magnification 50X.
Tip
Onion epidermis
Figure 2 shows the typical topography images of onion epidermis wall and celery (Apium graveolens L.)
parenchyma cell walls. In the topography images, the nm-wide cellulose microfibrils orientation and multilayer
structure were clearly observed. In our previous work of imaging epidermis cell wall of different onion scales, the AFM
images showed that cellulose microfibrils exposed at the innermost surface of the abaxial epidermis are oriented
perpendicular to the bulb axis in the outer scales and more dispersed in the inner scales of onion bulb.[29]
250 nm
500 nm
a)
b)m
Fig. 2 AFM images of the abaxial epidermal cell wall of a) the onion scale and b) celery epidermis walls. a) 1μm square and b)
500 nm square of the wall surface clearly show cellulose microfibril and matrix detail.
2.2 AFM based nanoindentation
Plant cell walls are complex composites which require delicate instruments and special techniques to study their
structures and properties. The analysis of micro- or nano- mechanical properties of plant cell walls has become
increasingly important in understanding cell wall structure and cell growth. Because of its micro-scale tip size,
indentation technique can investigate the mechanical properties of thin, small and heterogeneous materials.
Nanoindentation has been a new application tool on cellulose fibers and plant cell walls. Hardness and Young’s
modulus of spruce cell-wall, bamboo cell walls, individual wood fibers and crop stalks cell walls were examined.[3033] These experiments were done with add-on force transducers or special indenter testing systems. However, in
traditional nanoindentation experiments, samples were dehydrated and many were embedded in epoxy resin. Such
sample preparations may cause undesired modifications and influence the mechanical properties of the cell walls.
Recent work has used an improved nanoindentation technique to measure the mechanical properties of single cells or
tissue. The indentation depth is comparable to or larger than the cell wall thickness. The indent force is in a range of 1–
100 μN.[34] [35] The nano-indentation with an AFM has been recently applied to living plant cells in conditions close
to natural. Mechanical properties of suspended grapevine cells cultured in liquid medium, the primary cell wall of shoot
apical meristems, rosette leaves, tomato cells and epidermal cells of living roots of Arabidopsis thaliana were measured
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using AFM based nanoindentation.[36-40] The indentation depth is much lower (about 100 nm) in most of these
studies. However, the experimental samples contain living cells with different turgor pressure which has large influence
on measured results. No conclusive results were reached on the cell mechanical properties with the effect of turgor
pressure.[41, 42] The aim of this mini-review is to focus on the elastic properties of cells walls isolated from onion
scales using AFM based nanoindentation and scanning acoustic microscopy in order to assess the feasibility of these
methods for further study of plant cell walls. In this review, the focus is further on the freshly peeled single-layer onion
abaxial epidermis wall tested in liquid mode. The samples are independent of the turgor pressure influence and reflect
only the properties of hydrated cell wall in its natural condition.
With its piconewton force sensitivity and nanometer displacement accuracy, the AFM has been recognized as a
promising tool to for studying materials properties. AFM with force-distance curve measurement for nanoindentation
has emerged as a useful tool measuring the elastic moduli of biological samples. An AFM force-distance (F-D) curve is
a plot of tip-to-sample forces versus the extension of the piezoelectric scanner measured with a position-sensitive photo
detector. The F-D curve provides information of important mechanical properties such as adhesion, contaminants,
viscosity, and local variations in the elastic properties of the surface.
In experiments, F-D curves are quite complex and vary especially with different samples. For the experiments
discussed here, the inner side of plant epidermis cell walls and the tip are both negatively charged in water, so there is
no adhesion in F-D curves. Figure 3 shows a representative F-D curve obtained on onion epidermis cell wall in liquid.
Fig. 3 A representative F-D curve (approach and retract) obtained in
the experiment on onion epidermis cell wall in liquid. The y axis is the
recorded force in nano-newtons and the x axis is the relative piezo
displacement in nm.
The Hertz model is used as the theoretical model for f-d curve analysis. The Hertz model assumes that the sample is
isotropic, elastic and occupies infinite half space. It also assumes that the indenter is not deformable and there is no
additional interaction between the tip and sample.[43] The elastic modulus E can be obtained by a fit to the
experimental F-D curve using the Hertz model. The F-D curve obtained by AFM nanoindentation is in fact a forcepiezo displacement curve. To understand the force – sample deformation relationship, several specific parameters and
equations are introduced.
Fig. 4
Sketch of indentation distance and tip geometry.
As shown in Fig. 4, when using the AFM tip to indent the sample, the recorded piezo displacement D is comprised of
two parts: the tip cantilever bending x and the sample indentation δ. As shown in Fig. 4, the cantilever is moved towards
the sample by a carefully measured distance D.
(1)
According to Hertz model, the measured force based on a four-sided pyramid can be represented as [44]:
(2)
√
where F is recorded force, E is sample’s elastic modulus, ν is samples’s passion ratio, α is geometry angle of the
indenter. In our analysis, ν takes the value 0.3 and α is estimated as 13 according to the parameters given by the
manufacturer.
Knowing that x= F/k, where k is the cantilever spring constant, from the above two equations, we can easily obtain
relationship between recorded force F and recorded piezo displacement D:
(3)
√
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By fitting the experimental F-D retrace curves, we can get the sample’s elastic modulus E. Figure 5 shows a sample
F-D curve fitting result. The experimental F-D retrace curve and fitted curve from the Hertz model are in good
agreement.
Fig. 5 A sample of F-D curve fitting. Experiment data in blue dots and fitted
data in the red curve from the Hertz model showed good agreement.
2.3 Thickness of hydrated cell walls
For nanoindentation experiments, substrate influence on the measured results is negligible when the sample deformation
is less than 10% of sample thickness. The cell wall thickness was measured from scales of three different bulbs to make
sure the accuracy of the nanoindentation result. The onions used in this study typically contained twelve scales. The
outermost fresh scale will be referred as the 1st scale with the inner scales numbered sequentially through this paper. In
the sample preparation, the never-dried cell wall thickness of the abaxial epidermal cells is typically determined with
optical microscopy. The onion scale is hand-sectioned tangentially under a dissecting microscope (Olympus). Sections
from each of the 4 scales are stained with 0.1 % toludiene blue for 1 min at room temperature on a slide followed by
rinsing with deionized water and imaged with an optical microscope (EVOS, AMG). The thickness of different scales is
shown in Table 1. The sample deformation is typically less than 150 nm so the substrate influence does not affect the
accuracy. The F-D curve generating positions are in the middle of the cells thus avoiding area with vertical adjacent
walls.
Table 1
Hydrated onion epidermis wall thickness measured by toluidine test.
Onion scales
2
5
8
11
Cell wall thickness (μm)
AVG±SEM (n=3)
11.5±0.7
6.1±0.9
3.4±0.7
2.4±0.7
2.4 Moduli of different onion scales
The scales of onion contain primary cell walls at different growing stage, with inner scales being younger and outer
scales being older. Elastic moduli (E) of the four scales (11th, 8th, 5th and 2nd scales) in different onions were
measured by AFM nanoindentation. 100 curves in 5 random cells in each scale were generated and analyzed. The
results are shown in Fig. 6. The error bar shows sample standard deviation. Changes in moduli are statistically
significant. Previous research of onion epidermis scales indicate the orientation of cellulose microfibrils variation
tendency along scales.[29] Our moduli results indicate the elastic properties increase from inner/younger to outer/older
onion scales correspondingly. The plant cell walls become stronger as they rearrange the cellulose microfibrils
alignment during growth.
As shown Fig. 6, the bulb-to-bulb moduli variation was larger than the scale-to-scale variation for the same bulb. It is
understandable that onion bulbs would have different properties due to different growth environments and status. In our
subsequent experiments, the “same” samples were measured avoid original modulus variation.
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Fig. 6 Elastic moduli of different onion scales, scale 11, scale 8,
scale5 and scale 2 in three different onions. The error bar shows
sample standard deviation. Changes in moduli are statistically
significant. The moduli increase from inner scale to outer scale.
2.5 Onion epidermis wall with pectin aimed chemical treatment
2.5.1 EDTA effect
In the first chemical treatment, 100 mM EDTA was added onto the cell wall sample taken from onion scale 5. This
treatment is known to partially remove the EDTA-soluble pectin from the epidermis walls.[45, 46] Topography images
were generated in the same area in water and after adding the EDTA solution. 80 F-D curves were probed in the sample
area. Topography images and elastic moduli results are shown in Figure 7.
1µm
1µm
1µm
a)
b)
1µm
c)
d)
e)
Fig. 7 Topography images of onion scale 5 in a) water, b) 20 minutes after dipping
in 100 mM EDTA, c) 40 minutes after dipping in 100 mM EDTA, d) 100 minutes
after dipping in 100 mM EDTA. e) Elastic moduli measured in water and after adding
EDTA solution in the same area of cell wall shown in topography images. The error
bar shows sample standard deviation.
Topography images became clearer with time after adding in 100 mM EDTA solution. Elastic modulus decreased
slightly and remained stable with time after adding EDTA solution. The variation is statistically significant (p=0.04).
Elastic modulus decrease with EDTA treatment is probably due to the removal of EDTA-soluble pectin.
2.5.2 Calcium ion effect
In the second chemical treatment, the Calcium ion effect was explored. 100 mM CaCl2 solution was added onto the cell
wall removed from onion scale 5. Topography images were generated in the same area in water and after adding CaCl2
solution. 120 F-D curves were generated in the same area. Topography images and elastic moduli results are shown in
Fig. 8.
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1 µm
1 µm
1 µm
a)
b)
c)
d)
Fig. 8 Topography images of scale 5 in a) water, b) 30 minutes after adding
100 mM CaCl2, c) 60 minutes dipping in CaCl2 solution, d) Elastic moduli of
onion cell wall from scale 5 in water and in CaCl2 solution with time. The
increase is statistically significant.
Topography images showed apparent changes after adding 100 mM CaCl2 solution. Pectin, especially
homoglacturonan (HG) is negatively charged under cell wall physiological pH (around 5.0), Ca2+ serve as a bridge that
cross-links the HG chains and increase the regional density of the cell wall surface, which adds rigidity to the cell
wall.[47, 48] Due to the formation of gelation structures, elastic modulus increased dramatically after adding CaCl2
solution and the standard deviation increased significantly on the calcium ion treated sample. The topography and
elastic modulus remained relatively stable with time.
3. High Frequency Scanning Acoustic Microscopy (HF SAM)
In contrast to Scanning Electron Microscopy (SEM), Optical or Scanning Laser Confocal Microscopy (SLCM), high
frequency scanning acoustic microscopy (HF SAM) is capable of imaging not only the surface but also the shallow
subsurface regions of materials with high resolution [1-9]. For example, at 500 MHz the resolution in water at Standard
Temperature and Pressure is about 1.5 um. For the focus below the surface of a biological specimen the resolution is
approximately 2 micron depending on the acoustical properties of the specimen. The image contrast is based on at least
two mechanisms: (1) the contrast resulting from the dissimilar acoustic impedances of the different polymers and (2)
the contrast due to absorption of acoustic waves in the material. Conventional acoustic microscopy typically operates in
the 20 MHz to 200 MHz range with the use of focused transducers operating in the pulse wave mode. By contrast, HF
SAM operates in the 0.4 GHz to 2 GHz range in the long tone burst mode which allows correspondingly much higher
resolution [49]. More importantly, the high numerical aperture lens when brought close to the surface produces leaky
surface acoustic waves which are sensitive to the presence of changes in structure and materials and mark them in high
intensity contrast. This paper provides a description of the HF SAM and a review of the technique’s contrast
mechanism and associated simulations involved in data analysis as it is applied to the mechanical characterization of
plant cell walls subjected to enzymatic removal of pectin.
3.1 HF SAM Imaging
The HF SAM uses a mechanically scanning, acoustic lens system operating in a pulse-echo reflection mode. The
Scanning acoustic microscope provides a non-destructive method for studying the surface/subsurface microstructure of
nontransparent solids or biological materials. In scanning acoustic microscopy, a sample is examined by ultrasound
waves, and the variation of wave propagation generates the contrast maps which are used to form the resulting image.
Mechanical wave propagation and scattering is affected by the structure and elastic properties of the sample.
The working principle of the HF SAM can be described as below: As shown in Fig.9, a transmitter generates an
electric signal (usually a tone burst waveform) which travels into a piezoelectric transducer located on the top of a
sapphire buffer rod. A circulator limits the signal transmission to one direction: from the transmitter to the transducer or
from the transducer to the receiver. The piezoelectric transducer is applied for the electro-acoustic conversion for the
transmitter and receiver. The outgoing electric signal is converted to an acoustic plane wave by the transducer operating
in transmission mode, and this plane wave is focused into an ultrasonic beam by a spherical or cylindrical lens at the
end of the buffer rod. The ultrasonic beam is then transmitted through the fluid buffer, usually de-ionized water, into the
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sample. The wave travels through the sample at the material's velocity; part of the signal is reflected by the sample and
travels back through the lens. The transducer, now in receiving mode, converts the reflected ultrasonic signal into an
electrical signal which is collected by the receiver. The returning signal’s amplitude or phase is recorded and modulated
on a monitor to show the image of the focal area. Variations in the mechanical properties elicit changes in wave
propagation which affect the amplitude and phase of the reflected signal from the sample forming the contrast for
features on the grey-scale image. The operating frequencies of HF SAMs are between 400 MHz and 1 GHz with most at
400 MHz. Higher frequency lenses provide more accurate measurement results with a resolution of up to 1 µm at a
depth of 10 µm. 600MHz was found to give optimum results for imaging. For the V(z) measurement the choice of 400
MHz as the operating frequency was based on experience and a practical compromise between resolution and depth of
penetration of the waves into the specimen. The parameters of the acoustic lens are shown in below in Fig. 9. The
specimen is located in a container filled with coupling medium. The temperature of the coupling medium (i.e., distilled
water) is monitored by a thermocouple. The temperature is stabilized at 20.0°C (change less than ±0.1°C) in our
experiments.
a)
b)
Fig. 9 a) Schematic diagram of the HF SAM and b) Diagram of 400 MHz Acoustic Lens (Olympus, Model: AL4M631). The
dimensions in the figure are in cm.
This study focuses on the primary cell wall of onion epidermis as specimens. Typically, the plant epidermis was
bathed in 1x PBS (Phosphate Buffered Saline) solution with 0.05% Tween 20 for 1 hour. The edges of the sample were
glued to a clean glass slide while the internal part remained intact and free. During experiments, water was used as the
buffer liquid so the samples were kept hydrated. Acoustic images were obtained before V(z) measurement. In Fig. 10
are displayed typical HF SAM images of onion epidermal walls at a frequency of f= 600 MHz and at two different
scales. The white lines in (a) are identified as cuticle structures located between adjacent cell walls making up the risers
in the “shoe-box-like” structure.
a)
b)
Fig. 10 Typical acoustic images of onion epidermal walls at a frequency of 600 MHz. The white lines are identified as cuticle
structures located between adjacent cell walls making up the risers in the “shoe-box” structure. In (a) the scan width is 1 mm,
whereas in (b) the scan width is 0.5 mm.
3.2 V(z) measurement with HF SAM of Plant Cell wall
In addition to contrast variations provided by elastic properties of materials, another important usage of high frequency
HF SAM is measuring the velocity of surface acoustic waves. Atalar, Quate, and Wickramasinghe established that the
amplitude of the output voltage V as a function of lens-to-sample distance z ''has a characteristic response that is
dependent upon the elastic properties of the reflecting surface''.[50] Later Weglein reported the periodicity of dips in the
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V(z) curves.[51] The periodicity of the V(z) curve was related to surface wave propagation. Parmon and Bertoni
proposed a simple formula for determining the SAW velocity from V(z) curves measurement using a ray model.[52]
Wave velocity in the sample derived from the V(z) curve can be calculated according to Atalar’s model.[53] Following
well-known procedures to obtain, measure, and interpret V(z) curves, the longitudinal wave velocities were obtained by
implementing a computer parameter-fitting technique. The algorithm of the V(z) curve simulation is described in the
following steps. First, initialize parameters of acoustic lens, specimen (thinly sectioned biological tissue), and soda-lime
substrate. Second, calculate parameters of acoustic field at the back focal plane, pupil function of the lens, and
reflectance function. Third, calculate and draw the V(z) curve. The velocity of the longitudinal wave velocity of water
was set as 1,487 m/s. Based on the previous research study, the speed of sound in most plant cells is quite constant and
close to that of water, with ultrasound velocities are in the range of about 1600 m/s. Therefore the longitudinal velocity
of the tissue (thickness: 8μm) was set to a range from 1540 m/s to 1650 m/s. Moreover, the plant cell wall density is set
within a range of 1.3 – 1.6 (103 kg m-3). A Soda-lime glass slide was chosen as the substrate, its velocities of
longitudinal and shear waves are 6,000 m/s and 3200 m/s respectively. The summarized physical parameters of
biological tissue for computer simulation are shown in Table 2.
Table 2
Parameters of experimental conditions and biological tissue.
Lens
Liquid
Temperature of Water
Substrate
Radius of Lens
Wave Length in Water
Longitudinal Velocity of
Tissue
Density of Tissue
Thickness of Tissue
AL4M350
De-ionized Water
20ºC
soda-lime glass
577.52 µm
3.725 µm
1540 – 1650 m/s
1.4 g / cm3
5.00 – 8.00 µm
Fig. 11
A typical experimental V(z) curve
shown in blue line and the simulated V(z) curve in
red line. The curve was obtained at room
temperature on hydrated onion epidermis. The
vertical axis is the intensity of the received signal.
The horizontal axis is the defocusing distance of
the lens. z=0 represents the position of the lens
when it is focused on the top surface of the
specimen. The curves are obtained as the lens is
moved deeper into the interior of the specimen
Changing the values of the longitudinal wave velocity and the thickness, the simulations were continued until the
periodicities in the interference pattern were nearly coincident with the experimental values. Figure 11 shows a typical
experimental V(z) curve generated on an onion epidermal wall, as well as the matching curve from simulation. The
vertical axis is the intensity of the received signal. The horizontal axis is the defocusing distance of the lens. z=0
represents the position of the lens when it is focused on the top surface of the specimen. The curves are obtained as the
lens is moved deeper and toward the interior of the specimen. The longitudinal wave velocity in epidermal wall was
calculated as 1628 m/s. Ten curves were measured in different positions along the sample surface. The wave velocity
resulting from simulation was found to be 1626 ±3 m/s.
The sample material longitudinal velocity VL is related to the elastic modulus with the relationships below:
(4)
where VL is the longitudinal wave velocity in the sample, K is the sample material bulk modulus which is related to the
Young’s modulus E by:
(5)
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where ν is the material Poisson ratio. Then the elastic modulus E can be presented as
(6)
According to Eq. (6), the elastic modulus E could be calculated as
3.3
using a standard assumed value for Poisson’s ratio and density, respectively, of ν=0.3and ρ=1.4 g/cm3.
Those values are in approximate agreement with the modulus values in the literature. HF SAM measures elastic
properties at the macroscopic level. The macroscopic properties incorporate the mechanical composition of a sample
over a distance proportionally large in comparison with the dimension of cellulose microfibrils and bundles. The V(z)
measurement is therefore, potentially more sensitive to visco-elasticity of the plant cell wall as a composite as compared
to for example AFM techniques. AFM techniques do have the ability to measure the viscoelastic response of a single
microfiber embedded in the plant cell wall matrix, however, the loss of the leaky surface acoustic wave resulting from
a V(z) measurement over millimeters of the surface of a sample is significantly more apparent.
3.3 Enzyme treatment
The V(z) curve was also used to study the influence to the mechanical properties of the plant cell wall by using
pectinase to remove the biopolymer pectin, which is one of the important structural components of a plant cell wall.
Ten to twenty units of Pectinase were mixed with 1mL of NaOAc buffer and 100 to 150 microliters of the mixed liquid
was applied to the prepared onion abaxial epidermis sample during the experiment. The V(z) measurements were taken
before the enzyme treatment, directly after the enzyme was applied, and at elapsed increments of 10, 30, 60, and 180
minutes after the enzyme was applied. Figure 12 shows the V(z) curves of onion skin under the enzyme treatment with
time. The gradual decrease of the amplitude at z=0 is obvious as a function of time, which means that less acoustic
energy is reflected back to the transducer from the sample surface. Under the action of the enzyme, the biopolymer
pectin was moved away from the cell wall, which changed the cell wall structurally. More acoustic energy was
absorbed by the cell wall loosened by the removal of the supporting pectin. To illustrate the peak amplitude change
more clearly, Fig. 13 shows the relative peak value of the V(z) curve at z=0 of as a function of time under enzyme
treatment.
Fig. 12
V(z) curve of onion skin under enzyme
treatment. Notice the gradual decrease of the surface
echo V(z=0) in amplitude as a function of time.
The control experiment in water was undertaken to compare with the enzyme treatment experiment. All the steps and
measurements were the same except instead of using enzyme, water was applied. Figure 13 shows the measured V(z)
curves peak value at z=0 of onion epidermis scale in water. It shows that the peak amplitude at z=0 of different times
remain at a static value with small amount of variation in water. This means the structure and mechanics of the cell wall
remained unchanged during the experiment.
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Fig. 13 Relative peak values of the V(z=0) amplitude as a
function of time into enzyme treatment. Also shown are the
results of the Control Experiment with water without enzyme
treatment as a function of time. Notice the relatively small
change with time of the V(z=0) peak amplitudes.
4. Discussion and Conclusions
In summary, this mini-review presents an educational description of two methods in advanced microscopy. The two
methods stand out from other microscopies in that they are used with water-immersed samples. The two methods are
acoustic microscopy for microscale resolution and atomic force microscopy for nanmeter resolution. The two methods
are not restricted to but are demonstrated for biological samples, in particular plant cell walls, which are currently of
particular interest. The interest arise in the desire to understand the composition and properties of plant cell walls which
are the fundamental building blocks for all natural plants ubiquitous around the globe and are therefore of great
importance to mankind and civilization.
In the feasibility demonstration of atomic force microscopy (AFM) measurements on single layered, fully hydrated
onion epidermis cell walls are used as test analogs. In the tests the AFM based nano-indentation method is described to
observe changes in the elastic properties of onion epidermis cell walls. In particular, the effects on the cell wall modulus
by different chemical treatments on pectins in the cell wall structure are shown. The results demonstrate that this
approach can effectively track variations in cell wall properties and thus show the feasibility of AFM based
nanoindentation as a tool to characterize - nondestructively – intact fully hydrated plant cell walls.
In the observations the AFM is shown to provide both imaging and quantitative elasticity measuring capabilities. The
influence on cell wall elasticity when pectin is chemically modified was examined. Pectin has long been suspected of
playing an important role in cell wall mechanics. Two chemical methods were used to modify the pectin network: Ca2+
to cross-link the pectin network and EDTA to partially remove pectin. Ca2+ treatment caused the formation of crosslinked pectin gelation structures, which can be clearly observed in the topography images. Correspondingly, the elastic
moduli were found to increase dramatically. On the other hand, the topography images after the EDTA treatment
became clearer as a result of partial pectin removal. Correspondingly, the moduli decreased, indicating the weakening
of pectin structure. This verifies that the pectin network makes significant contribution to cell wall property. Based on
these findings the conclusion can be drawn that the AFM based nanoindentation method is a viable technique and thus
opens the door to a systematic nondestructive examination and evaluation of complex cell wall structures in the future
for education and research.
The objective of this mini-review is also to demonstrate the feasibility of High Frequency Scanning Acoustic
Microscopy (HF SAM) as a bio-technological tool to characterize biological tissues, in particular primary plant cell wall
tissues. Currently, there is interest in the structure of cellulose micro-fibrils in primary cell walls from celery
collenchymas.[26, 54] A combination of x-ray and neutron scattering methods with vibration and nuclear magnetic
resonance spectroscopy were found useful in the elucidation of many details on the micro-fibril structure. These
observations were all obtained with a dry primary wall structure. In contrast, the approach using the HF SAM is shown
to provide both imaging capability and quantitative measurements of stiffness in the natural wet state of plant cell walls.
The plant cell wall is used as a test analog to demonstrate the influence on stiffness when one important structural
component is removed from a complex polymer structure. In particular, this work tested whether the biopolymer pectin
has a strong effect on the mechanical properties of primary plant cell walls. Onion epidermis were used as examples
because they are thought to contain about 42% of pectin. The mechanism for the effect on the elastic wave propagation
is thought to be the viscous absorption by the plant cell wall. The bio- technical approach is to use HF SAM to
document the effect of pectinase enzyme treatment to remove pectin from onion primary cell wall. The results indicate
a significant change in the V(z) signature with time after enzymatic pectin removal. Thus the HF SAM method opens
the door to a systematic study of complex polymer structures.
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Microscopy: advances in scientific research and education (A. Méndez-Vilas, Ed.)
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Acknowledgements The authors gratefully acknowledge Dr. Seong Kim for his help with AFM nanoindentation measurement and
Kabindra Kafle for his help with the cell wall thickness measurements and the helpful discussions with Dr. Sarah Kiemle. Authors
thank Mr. Fumio Uchino (formally of Olympus Corp.) for providing the information on the lens configuration, and Dr. Tatsuro
Kosaka (associate professor at Kochi Institute of Technology) for developing the V(z) simulation software program (bio application).
The authors were supported as part of the Center for Lignocellulose Structure and Formation (CLSF) an Energy Frontier Research
Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DESC0001090.
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