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Physiology and Biochemistry |
Max Planck Institute of Colloids and Interfaces, Department of Biomaterials, Research Campus Golm, 14424 Potsdam, Germany
Received for publication August 21, 2006. Accepted for publication March 16, 2006.
ABSTRACT
Plants are hierarchically organized in a way that their macroscopic properties emerge from their micro- and nanostructural level. Hence, micromechanical investigations, which focus on the mechanical design of plant cell walls, are well suited for elucidating the details of the relationship between plant form and function. However, due to the complex nature of primary and secondary cell walls, micromechanical tests on the entire structure cannot provide exact values for polymer properties but must be targeted at the general mechanisms of cell wall deformation and polymer interaction. The success of micromechanical examinations depends on well-considered specimen selection and/or sample pretreatment as well as appropriate experimental setups. Making use of structural differences by taking advantage of the natural variability in plant tissue and cell structure, adaptation strategies can be analyzed at the micro- and nanoscale. Targeted genetic and enzymatic treatments can be utilized to specifically modify individual polymers without degrading the structural integrity of the cell wall. The mechanical properties of such artificial systems reveal the functional roles of individual polymers for a better understanding of the mechanical interactions within the cell wall assembly. In terms of testing methodology, in situ methods that combine micromechanical testing with structural and chemical analyses are particularly well suited for the study of the basic structure–property relationships in plant design. The micromechanical approaches reviewed here are not exhaustive, but they do provide a reasonably comprehensive overview of the methodology with which the general mechanisms underlying the functionality of plant micro- and nanostructure can be explored without destroying the entire cell wall.
Key Words: genetic modification • in situ methods • micromechanics • microtensile tests • natural variability • primary cell wall • secondary cell wall • structure–function relationships
Plants adapt the geometry of their organs and their component material properties to cope with external and internal stresses. Consequently, the structure and functionality of their tissues and cells are closely interdependent, resulting in fascinating mechanical designs and performance. Researchers in plant biomechanics aim to relate plant form and function to better understand the evolutionary process, particularly adaptive evolution (Niklas, 1992
). The field also provides a venue with which to extract biomimetic principles for the design of new materials (Vincent, 1990
, 2002
; Jeronimidis, 2000
; Fratzl et al., 2004a
; see Milwich, 2006
, in this issue).
One characteristic of biological materials is their hierarchical organization (Speck et al., 1996
; Fratzl, 2003
; Aizenberg et al., 2005
), which enables plants to structurally adapt with high efficiency on every hierarchical level, because all levels of organization are tightly integrated. Hence, macroscopic properties of the plant mainly originate from the cell wall organization (Salmén, 2000
) making the nano- and microstructural scale particularly relevant for biomechanical approaches. At these levels of hierarchy, plants manifest a wide variety of adaptable parameters such as cell shape, thickness, and arrangement of cell wall layers, the orientation of cellulose microfibrils within cell walls, and chemical composition of individual cell wall layers. The cell wall macromolecules form a complex network in which they act cooperatively and thereby determine the deformation behavior of the entire cell wall. Thus although the elastic properties of isolated cell wall components of secondary cell walls (e.g., cellulose, hemicelluloses, and lignin) are partly known (Salmén, 2001
), the native function of a single molecule can hardly be inferred from its properties in isolated conditions after the native assembly has been decomposed.
The micromechanical testing protocols discussed in the present article retain the structural integrity of the cell wall by targeting the general mechanisms of cell wall deformation and polymer interaction and thereby shedding light on the interdependency between plant structure, property, and functionality. The crucial role of specimen selection and preparation is emphasized because structural and biochemical differences among tissue and cell types are the starting point for micromechanical studies. Testing setups and approaches are highlighted, with particular emphasis on combined in situ methods, and recent progress in understanding plant form and function at the nano- and microstructural level is reviewed.
APPROACHING THE MECHANICAL DESIGN OF PLANT CELL WALLS
Selecting from the multitude of meaningful micromechanical test protocols for elucidating the mechanical design of plant cell walls, I highlight some approaches for better understanding the mechanical design of plant cell walls in a comprehensive but not exhaustive compilation. Rewarding combinations of microtensile testing techniques and material preparation and/or selection are exemplified.
For clarity, the structure of the article is presented in an organizational chart (Fig. 1), which can serve as a guide for the manuscript.
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; Fratzl et al., 2004a
). The modulus of elasticity of the homopolymer cellulose is approximately 134 GPa in the axial direction under moist conditions. In contrast, the matrix polymers, hemicellulose and lignin, have a modulus of elasticity of 40 MPa and 2 GPa, respectively (Salmén, 2001
). Despite a principle analogy in the fiber and matrix composition, primary and secondary cell walls differ significantly according to their function.
Primary walls have to cope with two conflicting goals, they have to be rigid to withstand the internal and external stresses and at the same time have to be pliant to allow cell wall expansion during growth (Cleland, 1971
; Taiz, 1984
). The almost transverse orientation of cellulose microfibrils facilitates longitudinal extension and stabilizes the cell in the circumferential direction (Baskin, 2005
). The matrix of primary walls consists mainly of hemicelluloses (e.g., xyloglucan, arabinoxylan), pectins, and structural cell wall proteins, which build a complex assembly with the cellulose microfibrils. Even though the individual cell wall polymers and their architecture are well characterized (McNeil et al., 1984
; McCann and Roberts, 1991
; McCann et al., 1992
; Carpita and Gibeaut, 1993
; Vincken et al., 1997
; Bacic et al., 1998
; Schindler, 1998
), little is known about their intrinsic spatial orientation and interplay during mechanical deformation. Various models of the cell wall assembly have been proposed in the last decades, which mainly differ in the structural organization of the cellulose–xyloglucan network and the bonding characteristics of the matrix polymers (Keegstra et al., 1973
; Hayashi, 1989
; Talbott and Ray, 1992
; Fry, 1995
; Ha et al., 1997
).
Any secondary cell wall that may form on the primary wall is deposited after the differentiating cell has reached its final shape and size. The thick secondary walls make cells stiff and strong and stabilize cells even when they die at the end of their differentiation. The secondary walls of fibers of monocotlyedons are multilamellar, whereas secondary walls of the secondary xylem of trees consist of three layers (S1, S2, S3) with different inclinations of the parallel-oriented cellulose fibrils toward the cell axis. The cellulose microfibril angle (MFA) is a measure of this inclination. The microfibrils are 2.5 nm thick and are embedded in a matrix of hemicelluloses and lignin (Fengel and Wegener, 1984
). Several models of the spatial orientation of the cell wall polymers have been proposed (Fengel, 1970
; Kerr and Goring, 1975
; Sell and Zimmermann, 1993
; Fahlén and Salmén, 2005
). For a comprehensive review of the micro- and ultrastructural organization of secondary cell walls, particularly in spruce (Picea abies [L.] Karst.) I refer readers to an article recently written by Brandström (2001)
. Due to the spatial organization and structural variety of the hemicelluloses, they are not only part of the matrix but also function as a coupling agent between the matrix and the stiff cellulose fibrils. For instance, in wood cell walls of gymnosperms, glucomannans are closely associated with cellulose, whereas xylans are associated with lignin (Salmén and Olsson, 1998
).
Micromechanical tests
To study the interrelation of cell wall structure and mechanical performance, micromechanical investigations are needed to determine the mechanical properties and to elucidate deformation behavior. In Table 1, the main mechanical terms used in this article are listed.
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An important concern is the manner in which different cell or tissue types are tested mechanically. Biaxial tests closely mimic the natural loading conditions of many organ types and therefore are the most appropriate technique to measure mechanical properties of tissue samples (Chanliaud et al., 2002
; Bargel et al., 2004
; Mackenzie-Helnwein et al., 2005
). However, biaxial tests require that tissue samples be prepared in ways that limit their applicability in terms of cell and cell wall mechanics. As a consequence, uniaxial tensile testing techniques are more generally used for this purpose (Hepworth and Vincent, 1998
; Reiterer et al., 1999
; Spatz et al., 1999
; Köhler et al., 2000
; Ryden et al., 2003
; Burgert and Jungnikl, 2004
). Moreover, uniaxial tensile testing techniques are particularly well suited for investigating the mechanical properties of individual fibers and tracheids (Jayne, 1959
; Page and El-Hosseiny, 1983
; Groom et al., 2002a
; Burgert et al., 2003
). Compression tests, however, are an appropriate alternative for the study of single parenchymatous cells with spherical shape (Wang and Thomas, 2000
). Microindentation provides valuable information on tissue and cell wall properties with an even higher spatial resolution at the microscale (Hiller et al., 1996
; Geitmann et al., 2004
; Parre and Geitmann, 2005b
), whereas nanoidentation allows for hardness and stiffness to be directly measured at the submicron level (Wimmer et al., 1997
; Gindl et al., 2004
). However, the orthotropic nature of the cell wall should be considered when using this method to determine the longitudinal modulus of elasticity of the wall (Gindl and Schoberl, 2004
). Arguably however, the most powerful approach is to use a combination of techniques that can shed light on the nano- and microstructural deformation of plant materials upon mechanical loading, e.g., in situ tensile tests combined with scanning electron microscopy, Raman microscopy, or X-ray diffraction methods (Mott et al., 1996
; Eichhorn et al., 2003
; Frühmann et al., 2003
; Keckes et al., 2003
; Kölln et al., 2005
). A combination of experimental data with theoretical treatments proved helpful to better understand general mechanisms of cell wall deformation and the principles of structure–function relationships (Preston, 1974
; Niklas and Paolillo, 1997
; Köhler et al., 2000
; Zhu and Melrose, 2003
; Fratzl et al., 2004b
; Hepworth and Bruce, 2004
).
MAKING USE OF STRUCTURAL DIFFERENCES
Native systems
An important concern in micromechanical test protocols lies in the selection of specimens and in the biological variation evident both within and across tissue types. Structure–function relationships can be elucidated either by studying different tissue types with various structural features or by investigating different ontogenetic stages of a plant or developmental stages of an organ as well as thigmomorphogenesis (see Telewski, 2006, in this issue).
Ontogenetic stages
When plants have to change their mechanical properties in order to cope with variable environmental conditions or to change their growth strategies, tissues can be modulated by a multitude of micro- and nanostructural features. Hoffmann et al. (2003)
found variations in the mechanical, structural, and chemical features during ontogeny of two tropical lianas when they switched from a self-supporting to a nonself-supporting phase. For the early ontogeny of two lianescent Aristolochia species, Köhler et al. (2000)
measured an increase in stiffness, which was closely related to changes in density, lignification, and microfibril angle. Bargel and Neinhuis (2005)
investigated distinct ripening stages of tomato fruit exocarps. Uniaxial tensile tests on enzymatically isolated cuticular membranes showed an increase in stiffness from the immature to the fully ripe state (Fig. 2), which could be related to morphological changes during fruit ripening.
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Specific tissue types
Plants manifest a wide variety of adaptable parameters at the nano- and microscale that enable them to form specific tissue types with cells of various shape, thickness, chemical composition, arrangement of cell wall layers, and orientation of cellulose microfibrils. This natural variability of plant materials can be utilized to study given microstructural variables, but one has to be aware of the variation of additional parameters across tissue types besides the targeted one. While the mechanical role of a few parameters is fairly obvious (e.g., cell wall thickness), little of the role of others is known (e.g., chemical composition of hemicelluloses and lignin). Here, the crucial role of cellulose fibril orientation and the interaction of cellulose fibrils and matrix polymers are highlighted.
Cellulose fibril orientation
The microfibril angle as measure of the cellulose fibril orientation in the secondary wall layer, particularly the S2 layer, plays a crucial role in the mechanical properties of plant tissues and cells (Preston, 1974
). Among all cell wall features, this angle is probably the most effective structural parameter for plants to control and adjust their mechanical performance. The comparison of tensile properties of different tissue types at the tissue level clearly indicates that mechanical properties depend on the cellulose fibril orientation in the secondary cell wall (Fig. 3).
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). For instance, cell walls of the wood of young trees have a rather high microfibril angle (Lindström et al., 1998
), which makes their wood flexible and enables the plant to reduce wind loads by bending. Cell walls of adult wood, however, have only a minor inclination toward the cell axis, making their wood much stiffer in order to withstand wind loads (Lichtenegger et al., 1999
). Consistently, also in studies on the mechanical properties of single fibers of a tree trunk, juvenile and adult wood differed significantly (Groom et al., 2002b
; Mott et al., 2002
). Moreover, branches are particularly suited for studying the highly sophisticated system of cellulose orientation adjustment and its influence on mechanical properties (Färber et al., 2001
; Burgert and Jungnikl, 2004
) because severe reaction wood tissues (e.g., compression wood, tension wood) with extreme values of microfibril angles are regularly formed (Wardrop, 1965
).
Fiber and matrix interaction
The interaction between the stiff cellulose fibrils and the pliant matrix polymers in the cell wall is one of the key issues in understanding the mechanical performance of plants (Fratzl, 2003
). By way of an analogy with technical fiber composites, the plant cell wall can be subdivided into fiber and matrix components (Kerstens et al., 2001
; Fratzl et al., 2004b
; Fahlén and Salmén, 2005
). While the prominent role of cellulose fibril orientation for the mechanical performance is well established, the influence of fiber–matrix interactions remains more elusive. Currently, we know most about the mechanical interaction of hemicelluloses with cellulose in the secondary cell walls (Akerholm and Salmén, 2001
; Salmén, 2004
), but its crucial role in the mechanical properties of primary walls is still controversial (Cosgrove, 2001
, 2005
; Thompson, 2005
).
Cyclic loading tests are particularly well suited for studying the specific interactions between fibers and matrix in the entire cell wall. For tissues and individual cells in particular when characterized by rather high fibrillar angles, a biphasic (or triphasic) stress–strain curve has been typically observed (Page et al., 1971
; Bodig and Jane, 1993
; Navi et al., 1995
; Köhler et al., 2000
; Köhler and Spatz, 2002
; Keckes et al., 2003
). A typical stress–strain curve for a tissue with a high microfibril angle [spruce: Picea abies (L.) Karst.] is shown in Fig. 4.
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). An explanation for the deformation behavior beyond the yield point can be given by the slippage of the matrix (Köhler and Spatz, 2002
), supposing that a large number of hydrogen bonds transmit shear stresses between cellulose fibrils. When a certain shear stress is exceeded, hydrogen bonds break, which results in viscous flow of the matrix. When stresses are removed, hydrogen bonds reform and lock the fibrils into new positions. The plant biomaterial thus shows permanent plastic deformation without significant mechanical damage through a molecular stick–slip mechanism operated by some sort of "velcro" connection (Keckes et al., 2003
). A similar effect was found by Hogan and Niklas (2004)
when they examined the viscoelastic behavior of green and frozen wood.
In terms of primary walls, the mechanisms of cell wall deformation by means of cellulose fibril and matrix interaction remain more elusive and cannot be discussed in greater detail here. Primary walls not only have to be rigid to withstand internal and external forces, but they also have to be flexible to be able to expand during growth (Jarvis and McCann, 2000
; Cosgrove, 2005
). The mechanisms that trigger the necessary modification of the fiber and matrix network (cell wall loosening) and allow for a substantial change in deformation behavior are still disputed.
It is generally accepted that the cellulose-xyloglucan interaction is of crucial relevance with respect to cell wall deformation. However, whether xyloglucan and cellulose form a tethered network (Cosgrove, 2005
) or xyloglucan chains act as spacers or struts between the cellulose microfibrils rather than as tethers (Thompson, 2005
) is open to question. In terms of a mechanical characterization of the composite, in vivo and in vitro assays have shown the crucial role of growth hormones, pH, and temperature on the mechanical properties of the primary cell wall (Kutschera and Schopfer, 1986a
, b
; Cosgrove, 1989
, 1993
; O'Looney and Fry, 2005
). Interestingly, Nolte and Schopfer (1997)
found in cyclic loading tests a viscoelastic deformation of the cell wall, while for cell growth a plastic deformation of the wall is needed. However, recently, Suslov and Verbelen (2006)
argued that elastic and plastic deformations can be distinguished when different stress states of the cell wall are compared.
Artificial systems
Genetic and enzymatic treatments
In enzymatic treatments or genetic modifications of plant cell walls, the individual components are specifically suppressed or abolished without disintegrating the entire composite. In comparison to natural variability in plant tissues, such modifications change individual structural parameters and not a multitude. Hence, the basic idea of these approaches is to alter the functioning of individual polymers in the primary and secondary cell walls to reveal the crucial mechanical role of the targeted component. For instance, microindentation experiments on enzymatically treated pollen tubes revealed that cellular stiffness was reduced and viscoelasticity was increased when both pectin and callose were attacked by enzymes (Parre and Geitmann, 2005a
, b).
Genetic modification is likely to be the most effective and precise way for modification aimed at gaining insight into the mechanical interaction of individual cell wall polymers. Micromechanical tests have been carried out on the entire shoots of Arabidopsis with genetically modified cell walls (Köhler and Spatz, 2002
) and on Arabidopsis hypocotyls (Ryden et al., 2003
; Pena et al., 2004
). Pena studied primary walls with an altered xyloglucan side chain structure by investigating the Arabidopsis mutants mur2 and mur3 with a well-defined and characterized modification (Reiter et al., 1997
; Pena et al., 2004
). Mur2 has a defect in xyloglucan-specific fucosyltransferase (Vanzin et al., 2002
), whereas mur3 has a defect in the galactosyltransferase (Madson et al., 2003
). Tensile tests on the hypocotyls revealed only minor changes for mur2 but a drastic decrease in stiffness and strength for mur3 (Ryden et al., 2003
; Pena et al., 2004
). In Fig. 5, the effect of a side-chain modification in xyloglucan on ultimate stress and stiffness is shown in comparison to the mechanical properties of the wild type (Fig. 5).
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, 1997
). However, the effect on the mechanical properties is only pronounced for galactosyltransferase (Ryden et al., 2003
; Pena et al., 2004
), which may also indicate that the side chain length influences the degree of hydrogen bonding of xyloglucan.
In vitro synthesis
Micromechanical approaches based on in vitro synthesis of the cell wall assembly aim to mimic the natural composite networks in the primary cell wall in order to improve our understanding of their mechanical design. The interaction of bacterial cellulose and combinations of various matrix polymers is molecularly and ultrastructurally analyzed, and the mechanical performance of the artificial cell wall can be revealed in micromechanical tests (Chanliaud and Gidley, 1999
). Results can not be discussed in greater detail here, but generally, the addition of matrix substances decreases cell wall stiffness in comparison to the pure cellulose system. Data indicate that the type of hemicellulose that interacts with cellulose plays an important role in the mechanical design of cell walls (Whitney et al., 1995
, 1999
).
MAKING USE OF COMBINED STUDIES
A powerful strategy to gain access to deformation mechanisms at the cell and cell wall level is to combine in situ methods for simultaneous deformation and monitoring. By watching materials deform, we can more precisely and directly detect structure–function relationships at the nano- and microstructural levels. Load-bearing components or networks can be examined, providing further knowledge on the underlying macromolecular interactions. Several approaches for simultaneous examinations have been used in the last decade. In micromechanical tests combined with scanning electron microscopy, microscale structural changes and fracture events have been observed during sample straining (Bodner et al., 1996
; Mott et al., 1996
). In a wet (environmental) mode, specimens can be both tested under moist conditions and examined without conductive coating. This combination is of particular advantage in microfracture studies when new surfaces are created in the deformation process (Mott et al., 1995
; Frühmann et al., 2003
).
The organization and mechanical interaction of cell wall polymers can be studied by combining micromechanical tests with nanostructural characterization. The interaction between cellulose and hemicelluloses has been successfully measured by dynamic Fourier transform infrared (FT-IR) spectroscopy (Akerholm and Salmén, 2001
). In the following, two examples of in situ approaches are given for the examination of nano-deformation at the cell wall level.
Microtensile tests combined with X-ray diffraction
Small-angle X-ray scattering (SAXS) and wide-angle X-ray diffraction (WAXD) are powerful techniques for characterizing plant cell walls at the nanoscale. The contrasting electron density between cellulose microfibrils and matrix polymers is the basis for small-angle X-ray scattering. Wide-angle X-ray diffraction provides information on periodic structures such as the atomic order in the crystalline parts of the cellulose microfibrils (Lichtenegger et al., 1998
). Both methods can be used to determine the microfibril angle in plant cell walls; however, the detection of the changes in the microfibril angle upon stretching demands combined in situ methods. Such molecular deformation mechanisms can be studied by straining the tissues or individual cells in tensile stages as shown by Köhler and Spatz (2002)
, who measured the change in cellulose microfibril angle against the applied strain in Aristolochia.
The data acquisition time for biological materials has to be short to exclude additional relaxation phenomena. Because microfibril angle measurements under laboratory conditions are time consuming, experiments are better done with a synchrotron source in order to collect X-ray signals while monitoring stress response during tensile straining (Keckes et al., 2003
; Kamiyama et al., 2005
; Kölln et al., 2005
) (Fig. 6).
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Microtensile tests combined with Raman spectroscopy
Raman spectroscopy is well suited to detect changes that occur at the molecular level in samples subjected to mechanical loads, which makes this technique suitable even for molecular stress mapping (Pezzotti, 2005
). In various assays, cellulose, cellulosic fibers, and wood have been examined during mechanical loading (Eichhorn et al., 2001
; Kong and Eichhorn, 2005
; Sturcova et al., 2005
; Peetla et al., 2006
). By relating changes in the Raman spectra with the specific stress–strain behavior, molecular deformation can be elucidated as a function of external strain. Spectra acquired during deformation show changes in peak intensity, peak shape, and peak position (data not shown). Wavenumber shifts upon tensile straining are shown for an individual spruce tracheid (Fig. 7).
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Summary
Plant biomechanics provides a powerful tool to gather insights into the relationship of plant form and function as an expression of plant strategy to survive under given environmental conditions and physical constraints. Because macroscopic properties of the plant mainly originate at the nano- and microscale, micromechanical approaches are well suited to gain better insights into both the optimization strategies of living plants and their material design. Micromechanical setups focus on the behavior of tissues and individual plant cell walls, their specific cell wall polymers, and the mechanical interaction of the different components in the polymer assembly. This focus sets a priori limit on the size and shape of tissue samples as well as on the methods used to measure mechanical properties.
The success in micromechanical studies is dependent on careful selection and preparation of materials with different structural and biochemical features as well as on modifications via enzymatic digestion and genetic engineering. To better understand the structure–function relationships of plants at the tissue, cell, and cell wall levels, we need to draw synergisms from the unique combination of micromechanical and nanostructural characterization. So far, targeted material modification and combined in situ methods have been barely employed conjointly. In particular, this combination may have the highest potential in gaining deeper insights into the structure–function relationships of plants at the micro- and nanoscale.
FOOTNOTES
1 The author thanks all those colleagues who contributed to the projects discussed, despite the lack of space to mention them all, H. Bargel for providing Fig. 2, and K. Niklas, H.-Ch. Spatz, and P. Fratzl for their helpful comments. The research was supported by the Max Planck Society and the Fonds zur Förderung der wissenschaftlichen Forschung (FWF), Austria. 
2 ingo.burgert{at}mpikg.mpg.de
; phone: +49-331-567-9432; fax: +49-331-567-9402
LITERATURE CITED
Aizenberg J. Weaver J. C. Thanawala M. S. Sundar V. C. Morse D. E. Fratzl P.. 2005. Skeleton of Euplectella sp.: structural hierarchy from the nanoscale to the macroscale. Science 309: 275-278.
Akerholm M. Salmén L.. 2001. Interactions between wood polymers studied by dynamic FT-IR spectroscopy. Polymer 42: 963-969.[CrossRef][Web of Science]
Bacic A. Harris P. J. Stone B. A.. 1998. Structure and function of plant cell walls. In J. Priess [ed.], The biochemistry of plants 297-371 Academic Press, New York, New York, USA.
Bargel H. Neinhuis C.. 2005. Tomato (Lycopersion esculentum Mill.) fruit growth and ripening as related to the biomechanical properties of fruit skin and isolated cuticle. Journal of Experimental Botany 56: 1049-1060.
Bargel H. Spatz H.-C. Speck T. Neinhuis C.. 2004. Two-dimensional tension tests in plant biomechanics: sweet cherry fruit skin as a model system. Plant Biology 6: 432-439.[CrossRef][Medline]
Baskin T. I.. 2005. Anisotropic expansion of the plant cell wall. Annual Review of Cell and Developmental Biology 21: 203-222.[CrossRef][Web of Science][Medline]
Bodig J. Jayne B. A.. 1993. Mechanics of wood and wood composites Krieger Publishing, Malabar, Florida, USA.
Bodner J. Grüll G. Schlag M. G.. 1996. In-situ fracturing of wood in the scanning electron microscope. Holzforschung 50: 487-490.[Web of Science]
Brandström J.. 2001. Micro- and ultrastructural aspects of Norway spruce tracheids: a review. International Association of Wood Anatomists Journal 22: 333-353.
Burgert I. Frühmann K. Keckes J. Fratzl P. Stanzl-Tschegg S. E.. 2003. Microtensile testing of wood fibres combined with video extensometry for efficient strain detection. Holzforschung 57: 661-664.[CrossRef][Web of Science]
Burgert I. Jungnikl K.. 2004. Adaptive growth of gymnosperm branches: ultrastructural and micromechanical examinations. Journal of Plant Growth Regulation 23: 76-82.
Burgert I. Keckes J. Fratzl P.. 2006. Mechanics of the wood cell wall. In D. D. Stokke, L. H. Groom [eds.], Characterization of the cellulosic cell wall 30-37 Blackwell, Oxford, UK.
Carpita N. C. Gibeaut D. M.. 1993. Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant Journal 3: 1-30.[CrossRef][Medline]
Chanliaud E. Burrows K. M. Jeronimidis G. Gidley M. J.. 2002. Mechanical properties of primary plant cell wall analogues. Planta 215: 989-996.[CrossRef][Web of Science][Medline]
Chanliaud E. Gidley M. J.. 1999. In vitro synthesis and properties of pectin/Acetobacter xylinus cellulose composites. Plant Journal 20: 25-35.[CrossRef][Web of Science][Medline]
Cleland R.. 1971. Cell wall extension. Annual Review of Plant Physiology 22: 179-222.
Cosgrove D. J.. 1989. Characterization of long-term extension of isolated cell walls from growing cucumber hypocotyls. Planta 177: 121-130.[CrossRef][Web of Science][Medline]
Cosgrove D. J.. 1993. Wall extensibility: its nature, measurement and relationship to plant cell growth. New Phytologist 124: 1-23.[CrossRef][Web of Science][Medline]
Cosgrove D. J.. 2001. Wall structure and wall loosening. A look backwards and forwards. Plant Physiology 125: 131-134.
Cosgrove D. J.. 2005. Growth of the plant cell wall. Nature Reviews 6: 850-861.[CrossRef]
Eichhorn S. J. Sirichaisit J. Young R. J.. 2001. Deformation mechanisms in cellulose fibres, paper, and wood. Journal of Materials Science 36: 3129-3135.[CrossRef][Web of Science]
Eichhorn S. J. Young R. J. Davies G. R. Riekel C.. 2003. Characterisation of the microstructure and deformation of high modulus cellulose fibres. Polymer 44: 5901-5908.[CrossRef][Web of Science]
Fahlén J. Salmén L.. 2005. Pore and matrix distribution in the fiber wall revealed by atomic force microscopy and image analysis. Biomacromolecules 6: 433-438.[CrossRef][Web of Science][Medline]
Färber J. Lichtenegger H. C. Reiterer A. Stanzl-Tschegg S. Fratzl P.. 2001. Cellulose microfibril angles in a spruce branch and mechanical implications. Journal of Materials Science 36: 5087-5092.[CrossRef][Web of Science]
Fengel D.. 1970. Ultrastructural behavior of cell wall polysaccharides. Tappi 53: 497-503.[Web of Science]
Fengel D. Wegener G.. 1984. Wood: chemistry, ultrastructure, reactions de Gruyter, Berlin, Germany.
Fratzl P.. 2003. Cellulose and collagen: from fibres to tissues. Current Opinion in Colloid and Interface Science 8: 32-39.[CrossRef]
Fratzl P. Burgert I. Gupta H. S.. 2004a. On the role of interface polymers for the mechanics of natural polymeric composites. Physical Chemistry, Chemical Physics 6: 5575-5579.[CrossRef][Web of Science]
Fratzl P. Burgert I. Keckes J.. 2004b. Mechanical model for the deformation of the wood cell wall. International Journal of Materials Research 95: 579-584.
Frühmann K. Burgert I. Stanzl-Tschegg S. E.. 2003. Detection of the fracture path under tensile loads through in situ tests in an ESEM chamber. Holzforschung 57: 326-332.[CrossRef][Web of Science]
Fry S. C.. 1995. Polysaccharide-modifiying enzymes in the plant cell wall. Annual Review of Plant Physiology and Plant Molecular Biology 46: 497-520.[CrossRef][Web of Science]
Geitmann A. McConnaughey W. Lang-Pauluzzi I. Franklin-Tong V. E. Emons A. M. C.. 2004. Cytomechanical properties of Papaver pollen tubes are altered after self-incompatibility challenge. Biophysical Journal 86: 3314-3323.[Web of Science][Medline]
Gierlinger N. Schwanninger M. Reinecke A. Burgert I.. 2006. Molecular changes during tensile deformation of single wood fibers followed by Raman microscopy. Biomacromolecules 7: 2077-2081.[CrossRef][Web of Science][Medline]
Gindl W. Gupta H. S. Schoberl T. Lichtenegger H. C. Fratzl P.. 2004. Mechanical properties of spruce wood cell walls by nanoindentation. Applied Physics A: Materials Science & Processing 79: 2069-2073.[Web of Science]
Gindl W. Schoberl T.. 2004. The significance of the elastic modulus of wood cell walls obtained from nanoindentation measurements. Composites Part A: Applied Science and Manufacturing 35: 1345-1349.[CrossRef][Web of Science]
Groom L. Mott L. Shaler S.. 2002a. Mechanical properties of individual southern pine fibers. Part I. Determination and variability of stress–strain curves with respect to tree height and juvenility. Wood and Fiber Science 34: 14-27.[Web of Science]
Groom L. Shaler S. Mott L.. 2002b. Mechanical properties of individual southern pine fibers. Part III. Global relationships between fiber properties and fiber location within an individual tree. Wood and Fiber Science 34: 238-250.[Web of Science]
Ha M. A. Apperley D. C. Jarvis M. C.. 1997. Molecular rigidity in dry and hydrated onion cell walls. Plant Physiology 115: 593-598.[Abstract]
Hayashi T.. 1989. Xyloglucans in the primary cell wall. Annual Review of Plant Physiology and Plant Molecular Biology 40: 139-168.[CrossRef][Web of Science]
Hepworth D. G. Bruce D. M.. 2004. Relationships between primary plant cell wall architecture and mechanical properties for onion bulb scale epidermal cells. Journal of Texture Studies 35: 586.[CrossRef][Web of Science]
Hepworth D. G. Vincent J. F. V.. 1998. The mechanical properties of xylem tissue from tobacco plants (Nicotiana tabacum ‘Samsun’). Annals of Botany 81: 751-759.
Hiller S. Bruce D. M. Jeronimidis G.. 1996. Micro-penetration technique for mechanical testing of plant cell walls. Journal of Texture Studies 27: 559-587.[CrossRef][Web of Science]
Hoffmann B. Chabbert B. Monties B. Speck T.. 2003. Mechanical, chemical and X-ray analysis of wood in the two tropical lianas Bauhinia guianensis and Condylocarpon guianense: variations during ontogeny. Planta 217: 32-40.[Web of Science][Medline]
Hogan J. C. Niklas K. J.. 2004. Temperature and water content effects on the viscoelastic behavior of Tilia americana (Tiliaceae) sapwood. Trees 18: 339-345.
Jarvis M. C. McCann M. C.. 2000. Macromolecular biophysics of the plant cell wall: concepts and methodology. Plant Physiology and Biochemistry 38: 1-13.[CrossRef][Web of Science]
Jayne B. A.. 1959. Mechanical properties of wood fibers. Tappi 42: 461-467.
Jeronimidis G.. 2000. Structure–property relationships in biological materials. In M. Elices [ed.], Structural biological materials: design and structure–property relationships 3-16 Pergamon, Amsterdam, Netherlands.
Kamiyama T. Suzuki H. Sugiyama J.. 2005. Studies of the structural change during deformation in Cryptomeria japonica by time-resolved synchrotron small-angle X-ray scattering. Journal of Structural Biology 151: 1-11.[CrossRef][Web of Science][Medline]
Keckes J. Burgert I. Frühmann K. Müller M. Kölln K. Hamilton M. Burghammer M. Roth S. V. Stanzl-Tschegg S. E. Fratzl P.. 2003. Cell-wall recovery after irreversible deformation of wood. Nature Materials 2: 810-814.[CrossRef][Web of Science][Medline]
Keegstra K. Talmadge K. W. Bauer W. D. Albersheim P.. 1973. The structure of plant cell walls. III. A model of the walls of suspension-cultured sycamore cells based on the interconnections of the macromolecular components. Plant Physiology 51: 188-196.
Kerr A. J. Goring D. A. I.. 1975. Ultrastructural arrangement of the wood cell wall. Cellulose Chemistry and Technology 9: 563-573.
Kerstens S. Decraemer W. F. Verbelen J. P.. 2001. Cell walls at the plant surface behave mechanically like fiber-reinforced composite materials. Plant Physiology 127: 381-385.
Köhler L. Spatz H.-C.. 2002. Micromechanics of plant tissues beyond the linear-elastic range. Planta 215: 33-40.[CrossRef][Web of Science][Medline]
Köhler L. Speck T. Spatz H.-C.. 2000. Micromechanics and anatomical changes during early ontogeny of two lianescent Aristolochia species. Planta 210: 691-700.[CrossRef][Web of Science][Medline]
Kölln K. Grotkopp I. Burghammer M. Roth S. V. Funari S. S. Dommach M. Müller M.. 2005. Mechanical properties of cellulose fibres and wood: orientational aspects in situ investigated with synchrotron radiation. Journal of Synchrotron Radiation 12: 739-744.[CrossRef][Web of Science][Medline]
Kong K. Eichhorn S. J.. 2005. Crystalline and amorphous deformation of process-controlled cellulose-II fibres. Polymer 46: 6380-6390.[CrossRef][Web of Science]
Kutschera U. Schopfer P.. 1986a. Effect of auxin and abscisic acid on cell wall extensibility in maize coleoptiles. Planta 167: 527-535.[CrossRef][Web of Science]
Kutschera U. Schopfer P.. 1986b. In-vivo measurement of cell-wall extensibility in maize coloeptiles: effects of auxin and abscisic acid. Planta 169: 437-442.[CrossRef][Web of Science]
Levy S. Maclachlan G. Staehelin L. A.. 1997. Xyloglucan side chains modulate binding to cellulose during vitro binding assays as predicted by conformational dynamics simulations. Plant Journal 11: 373-386.[CrossRef][Web of Science][Medline]
Levy S. York W. S. Stuike-Prill R. Meyer B. Staehelin L. A.. 1991. Simulations of the static and dynamic molecular conformations of xyloglucan: the role of the fucosylated sidechain in surface-specific sidechain folding. Plant Journal 1: 195-215.[Web of Science][Medline]
Lichtenegger H. Reiterer A. Stanzl-Tschegg S. E. Fratzl P.. 1999. Variation of cellulose microfibril angles in softwoods and hardwoods—a possible strategy of mechanical optimization. Journal of Structural Biology 128: 257-269.[CrossRef][Web of Science][Medline]
Lichtenegger H. Reiterer A. Tschegg S. Fratzl P.. 1998. Determination of spiral angles of elementary fibrils in the wood cell wall: comparison of small-angle X-ray scattering and wide-angle X-ray diffraction. In B. G. Butterfield [ed.], Microfibril angle in wood 140-156 University of Canterbury, Christchurch, New Zealand.
Lindström H. Evans J. W. Verrill S. P.. 1998. Influence of cambial age and growth conditions on microfibril angle in young Norway spruce (Picea abies [L.] Karst). Holzforschung 52: 573-581.[Web of Science]
Mackenzie-Helnwein P. Eberhardsteiner J. Mang H. A.. 2005. Rate-independent mechanical behaviour of biaxially stressed wood: experimental observations and constitutive modeling as an orthotropic two-surface elasto-plastic material. Holzforschung 59: 311-321.[CrossRef][Web of Science]
Madson M. Dunand C. Li X. M. Verma R. Vanzin G. F. Calplan J. Shoue D. A. Carpita N. C. Reiter W. D.. 2003. The MUR3 gene of Arabidopsis encodes a xyloglucan galactosyltransferase that is evolutionarily related to animal exostosins. Plant Cell 15: 1662-1670.
McCann M. C. Roberts K.. 1991. Architecture of the primary cell wall. In C. W. Lloyd [ed.], The cytoskeletal basis of plant growth and form 109-129 Academic Press, New York, New York, USA.
McCann M. C. Wells B. Roberts K.. 1992. Complexity in the spatial localization and length distribution of plant cell-wall matrix polysaccharides. Journal of Microscopy 166: 123-136.[Web of Science]
McNeil M. Darvill A. G. Fry S. C. Albersheim P.. 1984. Structure and function of the primary cell walls of plants. Annual Review of Biochemistry 53: 625-663.[CrossRef][Web of Science][Medline]
Milwich M. Speck T. Speck O. Stegmaier T. Planck H.. 2006. Biomimectics and technical textiles: solving engineering problems with the help of nature's wisdom. American Journal of Botany 93: 1455-1465.
Mott L. Groom L. Shaler S.. 2002. Mechanical properties of individual southern pine fibers. Part II. Comparison of earlywood and latewood fibers with respect to tree height and juvenility. Wood and Fiber Science 34: 221-237.[Web of Science]
Mott L. Shaler S. Groom L. H.. 1996. A technique to measure strain distributions in single wood pulp fibers. Wood and Fiber Science 28: 429-437.[Web of Science]
Mott L. Shaler S. Groom L. Liang B.-H.. 1995. The tensile testing of individual wood fibers using environmental scanning electron microscopy and video imaging analysis. Tappi 78: 143-148.
Navi P. Rastogi P. K. Gresse V. Tolou A.. 1995. Micromechanics of wood subjected to axial tension. Wood Science Technology 29: 411-429.
Niklas K. J.. 1992. Plant biomechanics University of Chicago Press, Chicago, Illinois, USA.
Niklas K. J. Paolillo Jr. D. J.. 1997. The role of the epidermis as a stiffening agent in Tulipa (Liliaceae) stems. American Journal of Botany 84: 735-744.[Abstract]
Nolte T. Schopfer P.. 1997. Viscoelastic versus plastic cell wall extensibility in growing seedling organs: a contribution to avoid some misconceptions. Journal of Experimental Botany 48: 2103-2107.
O'Looney N. Fry S. C.. 2005. A simple apparatus for measuring long-term extension of plant cell walls subjected to tensile stress. Plant Biosystems 139: 102-106.
Page D. H. El-Hosseiny F.. 1983. The mechanical properties of single wood pulp fibres. Part IV. Fibril angle and the shape of the stress-strain curve. Journal of Pulp and Paper Science 9: 1-2.
Page D. H. El-Hosseiny F. Winkler K.. 1971. Behaviour of single wood fibres under axial tensile strain. Nature 229: 252-253.[CrossRef][Medline]
Parre E. Geitmann A.. 2005a. More than a leak sealant. The mechanical properties of callose in pollen tubes. Plant Physiology 137: 274-286.
Parre E. Geitmann A.. 2005b. Pectin and the role of the physical properties of the cell wall in pollen tube growth of Solanum chacoense. Planta 220: 582-592.[CrossRef][Web of Science][Medline]
Peetla P. Schenzel K. C. Diepenbrock W.. 2006. Determination of mechanical strength properties of hemp fibers using near-infrared Fourier transform Raman microspectroscopy. Applied Spectroscopy 60: 682-691.[CrossRef][Web of Science][Medline]
Pena M. J. Ryden P. Madson M. Smith A. C. Carpita N. C.. 2004. The galactose residues of xyloglucan are essential to maintain mechanical strength of the primary cell walls in Arabidopsis during growth. Plant Physiology 134: 443-451.
Pezzotti G.. 2005. Raman piezo-spectroscopic analysis of natural and synthetic biomaterials. Analytical and Bioanalytical Chemistry 381: 577-590.[CrossRef][Web of Science][Medline]
Preston R.. 1974. The physical biology of plant cell walls Chapmann and Hall, London, UK.
Reiter W. D. Chapple C. Somerville C. R.. 1997. Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition. Plant Journal 12: 335-345.[CrossRef][Web of Science][Medline]
Reiterer A. Lichtenegger H. Tschegg S. Fratzl P.. 1999. Experimental evidence for a mechanical function of the cellulose microfibril angle in wood cell walls. Philosophical Magazine A 79: 2173-2184.[CrossRef]
Ryden P. Sugimoto-Shirasu K. Smith A. C. Findlay K. Reiter W.-D. McCann M. C.. 2003. Tensile properties of Arabidopsis cell walls depend on both a xyloglucan cross-linked microfibrillar network and rhamnogalacturonan II-borate complexes. Plant Physiology 123: 1033-1040.
Salmén L.. 2000. Structure-property relations of wood; from the cell-wall polymeric arrangement to the macroscopic behavior. In H.-C. Spatz, T. Speck [eds.], Proceedings of 3rd Plant Biomechanics Conference,Freiburg-Badenweiler, Germany,452-462 Georg Thieme Verlag, Stuttgart, Germany.
Salmén L.. 2001. Micromechanics of the wood cell wall: a tool for a better understanding of its structure. In P. Navi [ed.], Proceedings of 1st International Conference of the European Society for Wood Mechanics 385-398 EPFL, Lausanne, Switzerland.
Salmén L.. 2004. Micromechanical understanding of the cell-wall structure. Comptes Rendus Biologies 327: 873-880.[CrossRef][Web of Science][Medline]
Salmén L. Olsson A. M.. 1998. Interaction between hemicelluloses, lignin and cellulose: structure–property relationships. Journal of Pulp and Paper Science 24: 99-103.[Web of Science]
Schindler T. M.. 1998. The new view of the primary cell wall. Journal of Plant Nutrition and Soil Science 161: 499-508.
Sell J. Zimmermann T.. 1993. Radial fibril agglomerations of the S2 on transverse-fracture surfaces of tension-loaded spruce and white fir. Holz- als Roh- und Werkstoff 51: 384.[CrossRef][Web of Science]
Spatz H.-C. Köhler L. Niklas K. J.. 1999. Mechanical behaviour of plant tissues: composite materials or structures?. Journal of Experimental Biology 202: 3269-3272.[Abstract]
Speck T. Rowe N. Spatz H.-C.. 1996. Pflanzliche Achsen—Hochkomplexe Verbundstrukturen mit erstaunlichen mechanischen Eigenschaften. In W. Nachtigall, A. Wisser [eds.], Technische Biologie und Bionik 3, BIONA-report 10: 101-131 Fischer Verlag, Stuttgart, Germany.
Sturcova A. Davies G. R. Eichhorn S. J.. 2005. Elastic modulus and stress-transfer properties of tunicate cellulose whiskers. Biomacromolecules 6: 1055-1061.[CrossRef][Web of Science][Medline]
Suslov D. Verbelen J. P.. 2006. Cellulose orientation determines mechanical anisotropy in onion epidermis cell walls. Journal of Experimental Botany 57: 2183-2192.
Taiz L.. 1984. Plant cell expansion: regulation of cell wall mechanical properties. Annual Review of Plant Physiology 35: 585-657.[CrossRef][Web of Science]
Talbott L. D. Ray P. M.. 1992. Molecular size and separability features of pea cell wall polysaccharides. Plant Physiology 98: 357-368.
Thompson D. S.. 2005. How do cell walls regulate plant growth. Journal of Experimental Botany 56: 2275-2285.
Vanzin G. F. Madson M. Carpita N. C. Raikhel N. V. Keegstra K. Reiter W. D.. 2002. The mur2 mutant of Arabidopsis thaliana lacks fucosylated xyloglucan because of a lesion in fucosyltransferase AtFUT1. Proceedings of the National Academy of Sciences, USA 99: 3340-3345.
Vincent J. F. V.. 1990. Structural biomaterials Princeton University Press, Princeton, New Jersey.
Vincent J. F. V.. 2002. Survival of the cheapest. Materials Today 5: 28-41.
Vincken J. P. York W. S. Beldman G. Voragen A. G. J.. 1997. Two general branching patterns of xyloglucan, XXXG and XXGG. Plant Physiology 114: 9-13.[CrossRef][Web of Science][Medline]
Wang L. Thomas C. R.. 2000. pH affects the mechanical properties of single tomato cells. In H.-C. Spatz, T. Speck [eds.], Proceedings of 3rd Plant Biomechanics Conference, Freiburg-Badenweiler, Germany 368-373 Georg Thieme Verlag, Stuttgart, Germany.
Wardrop A. B.. 1965. The formation and function of reaction wood. In W.A. Coté [ed.], Cellular ultrastructure of woody plants 371-390 Syracuse University Press, New York, USA.
Whitney S. E. C. Brigham J. E. Darke A. H. Reid J. S. G. Gidley M. J.. 1995. In vitro assembly of cellulose/xyloglucan networks: ultrastructural and molecular aspects. Plant Journal 8: 491-504.[CrossRef][Web of Science]
Whitney S. E. C. Gothard M. G. E. Mitchell J. T. Gidley M. J.. 1999. Roles of cellulose and xyloglucan in determining the mechanical properties of primary plant cell walls. Plant Physiology 121: 657-663.
Wimmer R. Lucas B. N. Tsui T. Y. Oliver W. C.. 1997. Longitudinal hardness and Young's modulus of spruce tracheid secondary walls using nanoindentation. Wood Science and Technology 31: 131-141.[Web of Science]
Zhu H. X. Melrose J. R.. 2003. A mechanics model for the compression of plant and vegetative tissues. Journal of Theoretical Biology 221: 89-101.[CrossRef][Web of Science][Medline]
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of different wood tissues of spruce (Picea abies) measured in a tensile test as a function of cellulose microfibril angle. Data points show the means of a study by Reiterer et al. (1999)
< 0.001, comparing with the wild-type)
C-O-C (1907·cm–1) and 
C=C (1602·cm–1) recorded by Raman spectroscopy at different stages of a tensile experiment on an individual normal wood spruce tracheid (Picea abies) tested in dry condition (after Gierlinger et al., 2006