Imaging of Biological Nano-Composite Plant Cell Wall at the Micro- and Nano-Scales

Transcription

1 13th International Symposium on Nondestructive Characterization of Materials (NDCM-XIII), May 2013, Le Mans, France Imaging of Biological Nano-Composite Plant Cell Wall at the Micro- and Nano-Scales B. R. TITTMANN 1, S. LOUYEH-MAGHSOUDY 2, J. KIM 1, and X. XI 1 1 Department of Engineering Science and Mechanics, the Pennsylvania State University, University Park, PA USA Phone: ; brt4@psu.edu; juk252@psu.edu; xzx104@psu.edu 2 The Aerospace Corporation, El Segundo, CA USA; Sahar.Maghsoudy-Louyeh@aero.org Abstract This report is on the imaging and characterization of the nano-composite structure of native primary cell walls. In particular, the structure of primary celery epidermis cell walls was imaged with sub-nanometer resolution using the Atomic Force Microscope (AFM). The high-frequency acoustic microscope was used to image onion cell (Allium cepa) wall epidermis at the micro-scale at 600 MHz at 1 micron resolution. V(z) signatures were obtained and used to estimate the bulk modulus in good agreement with destructive measurements. The combined results reveal a surprisingly fine but strong nano-composite for the lignocellulosic biomass. Keywords: scanning acoustic microscopy, atomic force microscopy, plant cell walls, microfibrils, celery (Apium graveolens L.) parenchyma 1. Introduction The hypothesis of this study was to test the feasibility of imaging plant cell walls at both the macroscale and nanoscale. Scanning Acoustic Microscopy was used to image onion (Allium cepa) cell walls with micrometer resolution and Atomic Force Microscopy was used to image celery (Apium graveolens L.) parenchyma cell walls [1] with sub-nanometer resolution. This paper is a review and an extension of the previous works by the authors [2-3]. 2. Background Plant cells are bound by thin, yet mechanically strong, cell walls containing structural proteins, enzymes, phenolic polymers, and other materials which transform their chemical and physical characteristics [4,5]. Cell wall structures consist of a complex mixture of cellulose fiber, polysaccharides, and other polymers that are linked together by both covalent and noncovalent bonds. Cell walls adhere the cells together to prevent them from sliding and determine the mechanical strength of plants. They have many fundamental roles during the growth and development of a plant. This polymeric network (the cell wall) is enlarged by a process of stress relaxation and slipping (or creep) of the polysaccharides. The plant cell wall structure consists of three different layers: the middle lamella, the primary cell wall, and plasma membrane. The middle lamella (plural lamellae) is a layer high in pectin, which forms the interface between adjacent plant cells and glues them together [5]. The primary cell wall is a thin, flexible, and a fiberglass-like structure, with crystalline cellulose microfibrils embedded in a highly hydrated polysaccharide matrix [6]. The secondary cell wall also forms in some cells after expansion ceases and comprises a thick layer preventing collapse of the water-conducting cells during periods of high water tension due to high transpiration [7]. The Plasma membrane (cell membrane or plasmalemma) is a layer which regulates the movement of substances in and out of cells, and contains a wide range of biological molecules, lipids, and primary proteins that are involved in cell adhesion and ion channel conductance [8]. Cellulose, the most common biopolymer in nature, is present in a wide variety of living species including plants. Cellulose is the structural component of plant cell walls that acts as a

2 reinforcement material which has a high tensile strength, equivalent to steel [6]. Cellulose is a polysaccharide consisting of a linear chain of β (1, 4)-linked β-d-glucan units from several hundred to over ten thousand units [9]. These parallel glucans form a crystalline microfibril that is chemically stable, insoluble, and relatively resistant to chemical and enzymatic attack These properties make cellulose an excellent candidate to build a strong and complex cell wall. A fundamental understanding of the cellulosic structure of plant cell walls will provide the insight needed to create a new generation of biorenewable, nanocomposites with novel properties tailored to diverse applications. 3. Microscale Imaging with Scanning Acoustic Microscopy The Scanning acoustic microscope is 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 reflective wave maps the structures based on contrast of the elastic properties of the sample. The first scanning acoustic microscope, operated in the transmission mode, was created by Lemons and Quate at Stanford University in 1973 and was effectively used for phase imaging in reflection [10]. The acoustic microscopy has been improved since then and is generally in reflective mode now. As a convenient and non-destructive method, acoustic microscopy has been successfully used to map microstructures and show contrast of in vivo elastic properties of soft bio- tissues [11-17]. In this work, the Olympus UH3 reflective scanning acoustic microscope will be used to measure the in situ mechanical properties and anisotropy in plant cell walls as well as the enzyme treatment and tensile effect on these properties. Working principle of the SAM can be described as below: As shown in Figure 1, a transmitter generates an electric signal (usually a tone burst wave) which travels into a piezoelectric transducer located on the top of a sapphire buffer rod. A circulator makes sure that the wave signal only travels in 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 acoustic plane wave by the transducer, 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 transmitted through the fluid buffer usually deionized water into the 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 transverses the 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 cause changes in the amplitude and phase of the reflected signal from the sample and generate contrasted features on the grey-scale image. The operating frequencies of SAMs are between 100 MHz and 2 GHz. Higher frequency lens provides more accurate measurement results with a resolution of up to 1 µm at a depth of 10 µm.

3 Transmitter RF Input RF Output Receiver + Transducer Z Scanning Buffer Rod - Plane Wave Coupling Medium AARC Spherical Wave Specimen Soda-lime Glass Substrate X-Y Y Scanning Stage Figure 1. Schematic diagram of the SAM [18] Other than imaging variations of elastic properties of materials, another important usage of high frequency SAM is measuring the velocity of surface acoustic waves. Atalar, Quate, and Wickramasinghe found out that the amplitude of the output voltage V as a function of lens-tosample distance z ''has a characteristic response that is dependent upon the elastic properties of the reflecting surface'' [10]. Later Weglein and Wilson reported the periodicity of dips in the V(z) curves [19]. The periodicity of the V(z) curve was related to surface wave propagation. Parmon and Bertoni [20] proposed a simple formula for determining the SAW velocity from V(z) curves measurement using a ray model. The V(z) signature was further studied [21,22]. In 1981 Kushibiki and coworkers invented the line focus acoustic microscope (LFAM), which can measure the anisotropy of surface acoustic waves in anisotropic materials [23]. The material characterization by the inversion of V(z) was also studied by Endo et al. [24] and Kulik at al [25]. Wave velocity in the sample derived from the V(z) curve can be calculated according to Atalar s model [21]. The sample material longitudinal velocity V is related to the elastic modulus with the relationships below: V =... (1) where V 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:

4 K= E... (2) where ν is the material Poisson ratio. Then the elastic modulus E can be presented as E= ρv... (3) With this technique, mechanical properties of plant cell walls as an intact composite structure will be evaluated in its natural state or with different enzyme treatments. With the help of line focus lens, anisotropy of plant cell walls related to cellulose fibers orientation and alignment will be examined. 4. SAM measurement of elastic properties of onion cell walls Fresh onion epidermis were laid smoothly on a clean (100) silicon wafer with edges glued. The samples were dipped into 1 phosphate buffered saline (ph 7.4) containing 0.1% Tween 20 for 1 hour and then rinsed with water. During experiments, water was used as the buffer liquid so the samples were kept hydrated. Acoustic images, such as those shown in Figure 2, were obtained first. The positions to measure Vz curves were chosen based on these images. Figure 2. Typical acoustic images of onion epidermal walls, f= 600 MHz Acoustic lens model used in the Vz curve measurement is AL4M350 (f= 400 MHz). Figure 3 shows a typical experiment Vz curved generated on onion epidermal wall, as well as the matching curve from simulation. The longitudinal wave velocity in epidermal wall was calculated as 1628 m/s. Ten curves were measured in different positions. The average wave velocity after simulation is 1626 m/s. Figure 3. A typical experimental Vz curve shown in blue line and the simulated Vz curve in red line. The curve was obtained at room temperature. According to Equation (9), the elastic modulus E can be calculated as

5 1 ν1+ν1 2νρVL2 (3) Using ν=0.3 and ρ=1.4 g/cm, the elastic modulus E = 3.30 GPa. Unlike AFM, SAM measure elastic properties at the macroscopic level. The macroscopic properties depend on the structure of the cell wall, which can be heavily affected by the cell age, type and other factors. Therefore, large variations are expected. Further study with both AFM and SAM will be conducted to link the macroscopic measurement results with microscopic structure and properties. 5. Nanoscale Atomic Force Microscopy In this study the Atomic Force Microscope was used to investigate celery (Apium graveolens L.) parenchyma cell walls in situ. The use of the AFM and its evolution and use is described in a recent publication [3]. Here we only briefly mention recent methods adapted to image the fragile microfibrils of plant cell walls. We used the Peak Force Tapping (PFT) mode, in which the probe and sample are intermittently brought together (similar to Tapping Mode) to contact the surface quickly, which eliminates lateral forces. Both normal forces and lateral forces exerted by the tip can cause damage to the sample and increase the contact area resulting in scan resolution reduction. Unlike Tapping Mode, where the feedback loop keeps the cantilever vibration amplitude constant, Peak Force Tapping controls the maximum force (Peak Force) on the tip, and protects the tip and sample from damage by decreasing the contact area. The level of force control in PFT can be in the range of pn, even when scanning in liquid environments. The two biggest challenges of force control in a liquid environment are nonlinear deflection variations and viscous forces when the tip and sample are not in contact. To eliminate this problem, the PFT mode uses the feedback to maintain a constant peak force for each tap with the force range from pn to µn, depending on the application. This makes the PFT mode significantly applicable to the imaging and measurement of plant cell walls, which are naturally sensitive to tip movement and associated damage to the structure. We use the Peak Force Quantitative Nano-mechanical Mapping (Peak Force QNM) a new AFM mode that uses tapping mode technology to record very fast force response curves at every pixel in the image, and uses the peak tip-sample interaction force as the feedback mechanism. Peak Force QNM is able to simultaneously obtain quantitative modulus, adhesion, dissipation, and deformation data while imaging topography at high resolution. Also, by maintaining control of direct force to a very low level (pn), the scanning can limit indentation depths to deliver a non-destructive and high-resolution imaging technique to sensitive samples. The celery epidermis was bathed in 1x PBS (Phosphate Buffered Saline) solution with 0.05% detergent, called Tween 20, to remove the proteins. Due to the significant amount of Pectin, in comparison to some other cell walls, the samples needed to be bathed in solution for six hours or even longer. Due to the different structure of celery microfibrils, the preparation of a monolayer cell profile in celery epidermis was necessary and required sample preparation that was more complex and time consuming than with other samples. The celery epidermis included both multi layers and a single layer of cell, and cell profile. Therefore, finding a good position on the mono layer cells was a challenge. The sides of celery sample were glued to the clean, glass slides while the internal part remained intact and free. The Peak Force Tapping method was used with ultra low force (piconewton) for tapping the cantilever during scanning process. Sharp Scan Asyst Fluid + tips were used in this study with

6 radius of curvature of 2nm. The tip was washed after each set of experiments by dipping in distilled water + 90% ethanol to remove the residue from the scanning. Figure 4 shows 500 nm size images of intact celery microfibrils in water. The right side images are topography, while the left side images are the Peak Force error signal images. Peak Force Error, also called error signal of deflection, is obtained by the subtraction of the set point force from detector signal (actual deflection). In general, the deflection image shows the edges of features in the topography image. In soft materials, deflection Image was clearer than the Topography Image. If the error signal was too large, which was not in our case, the tip is unable to track the sample accurately. The images show the cell wall microfibrils arranged in a linear pattern. The smallest diameter is about 3-5 nm. Figure 4: 500 nm scan of the celery fibrils, performed with Peak Force Tapping mode in fluid, with Dimension ICON. The top layer has an angle of 42 to 50 degrees and the second layer has an angle of 111 to 117 degrees. The right side image is topography, while the left side image is the Peak Force error signal image. Peak Force Error, also called error signal of deflection, is obtained by the subtraction of the set point force from the detector signal (actual deflection). In general, the deflection image shows the edges of features in the topography image. 7. Conclusions As fossil fuel resources approach an end, the need for alternative fuels becomes essential. Ethanol from lignocellulosic biomass is one source of alternative energy which can substitute for its fossil fuel counterpart, gasoline. This fuel, from renewable lignocellulosic material, represents a source of supply, with limited conflict with land use for food and feed production, and lower fossil fuel inputs. The difficulties associated with pretreatment of lignocellulosic materials are a major obstacle in the production of ethanol. Based on these difficulties, there is an important need to understand the polysaccharide nature of cellulose fibrils in plant cell walls which are insoluble, resistant to chemical attack, and cannot be readily fermented. In this work we report on the imaging and characterization of the nanocomposite structure of native primary cell wall. In particular we investigated the structure of primary celery (Apium graveolens L.) epidermis cell walls. We imaged with sub-nanometer resolution using the Atomic Force Microscope (AFM) in the Peak Force Tapping Mode. We were able to clearly distinguish the micro-fibrils making up the cell wall. We developed special software to characterize the surface and sub-surface at the nano-scale. We imaged the structure of native primary cell wall surfaces and we could identify and evaluate 5 layers of micro-fibrils in terms of fiber thickness, angular orientation and spacing. We conclude that the micro-fibril stucture is weakly anisotropic and shows evidence of both horizontal and vertical bundling of micro-fibrils. We also used the high-frequency acoustic microscope to image

7 onion cell (Allium cepa) wall epidermis at the micro-scale at 600 MHz at 1 micron resolution. The images reveal an open shoe-box like structure. We were able to obtain the V(z) signatures and used them to estimate the bulk modulus in good agreement with destructive measurements. The combined results reveal a surprisingly fine but strong nano-composite for the lignocellulosic biomass. Acknowledgements The authors were partially 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 DE- SC References 1. S Maghsoudy-Louyeh, J Kim, M Kropf and B R Tittmann, Subsurface Image Analysis of Plant Cell Wall with Atomic Force Microscopy, J. of Advanced Microscopy Research, in press. 2. J C Thimm, D J Burritt, W A Ducker, L D Melton, Pectins Influence Microfibril Aggregation in Celery Cell Walls: An Atomic Force Microscopy Study, J Struct Biol. Vol 168, No 2, pp , November S Maghsoudy-Louyeh, M Kropf and B R Tittmann, Special Issue on Patents for Biological Imaging, Bentham Science Press, in press. 4. D J Cosgrove, Characterization of Long-Term Extension of Isolated Cell Walls from Growing Cucumber Hypocotyls, Plant, Vol 177, pp , D J Cosgrove, Loosening of Plant Cell Walls by Expansins, Nature, Vol 407, pp , D J Cosgrove, Growth of the Cell Wall, Nat. Rev. Mol. Cell Biol Vol 6, pp , D J Cosgrove, Cell Walls: Structure, Biogenesis, and Expansion, in Plant Physiology, 2nd ed. Lincoln Taiz and Eduardo Zeiger, eds. Sunderland, MA: Sinauer Associates, Chapter 15, B Alberts, A Johnson, J Lewis, M Raff, K Roberts, P Walter, Molecular biology of the cell (5th ed.). New York, NY: Garland Science Press, P Albersheim, A Darvill, K Roberts, R Sederoff, A Staehelin, Plant Cell Walls, New York, NY: Garland Science Press, A Atalar, C F Quate, H K Wickramasinge, Phase Imaging in Reflection with Acoustic Microscope, Appl. Phys. Lett., Vol 31, p. 791, L I Petrella, H A Valle, P R Issa, C J Martins, W C Pereira, J C Machado, Study of Cutaneous Cell Cardinomas Ex Vivo Using Ultrasound Biomicroscopy Images, Skin Pharmacol Appl. Skin Physiol. Vol 15, No 2, p , C Guittet, F Ossant, L Vaillant, M Berson, In Vivo High-Frequency Ultrasonic Characterization of Human Dermis, Trans Biomed Eng. Vol 46, No 6, pp , TM Nguyen, M Couade, J Bercoff, M Tanter, Assessment of Viscous and Elastic Properties of Sub-Wavelength Layered Soft Tissues Using Shear Wave Spectroscopy: Theoretical Framework and In-Vitro Experimental Validation, IEEE Trans Ultrason Ferroelectr Frequ Control, Vol 58, No 11, pp , E Maeva, F Severin, C Miyasaka, B R Tittmann, R Gr Maev, Acoustic Imaging of Thick Biological Tissue, IEEE Trans. Ultrason, and Freq. Control, Vol 56, No 7, pp , 2009.

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