Skip Navigation


AOBPreview originally published online on May 21, 2003
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Content Snapshot
Right arrow Content Select
Right arrow All Versions of this Article:
92/1/73    most recent
mcg115v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (10)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Agricola
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

Annals of Botany 92: 73-77, 2003
© 2003 Annals of Botany Company

Fracture of Plant Tissues and Walls as Visualized by Environmental Scanning Electron Microscopy

A. M. DONALD1, F. S. BAKER1, A. C. SMITH2 and K. W. WALDRON2

1 Cambridge University, Cavendish Laboratory, Madingley Road, Cambridge CB3 0HE, UK and 2 Institute of Food Research, Norwich Research Park, Norwich NR4 7UA, UK

* For correspondence. Fax +44 1603 507723, e-mail andrew.smith{at}bbsrc.ac.uk

Received: 9 July 2002; Returned for revision: 1 November 2002; Accepted: 24 March 2003    Published electronically: 21 May 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 LITERATURE CITED
 

The environmental scanning electron microscope (ESEM) provides a highly relevant and controllable environment in which to study hydrated systems without the artefacts of other highly prepared specimens. The instrument facilitates control of turgor through hydration using different chamber vapour pressures. Deformation of a simple plant tissue—upper epidermal layers in Allium cepa (onion)—was observed at the scale of the two principal failure mechanisms: cell breakage; and cell separation induced by treatment with a chelating agent. Cell rupture and release of contents occurred at cellular junctions ahead of an imposed growing notch, indicating that disruption of cells occurred remotely from the creation of a new surface. Cells that separated usually maintained their turgor and the separation process took place through progressive failure of middle lamellar material seen as strands between separating cells. These mechanisms were compared with the rupture of excised Chara corallina walls that occurred by formation and breakage of strands between separating wall layers. This study provides in situ visual characterization of wall rupture and cell separation at the microscopic level in hydrated plant material.

Key words: Environmental scanning electron microscope, ESEM, onion, Allium cepa L., alga, Chara corallina, fracture, cell wall, tissue.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 LITERATURE CITED
 
The role played by cellular structure in determining mechanical properties of plant tissues has been a topic of great research interest for several years (e.g. Waldron et al., 1997). In particular, one aim has been to image structural details at high resolution with minimal specimen preparation artefacts.

Experimental methodologies for studying in situ deformation are limited. Optical microscopy does not have sufficient resolution or depth of focus for studying cell separation and breakage. Scanning electron microscopy (SEM) overcomes this problem but, in the conventional instrument, tissue cannot be studied in its native state because of the requirements of high vacuum (leading to dehydration) and coating with a conductive layer. It is always questionable, therefore, whether the images gained by this technique truly represent the structure of the fully hydrated ‘live’ specimen. Nevertheless, many publications document considerable achievements. McGarry (1995) measured cell diameters from structures imaged by SEM, for which carrots had been prepared by freezing in liquid nitrogen and coating with gold.

Problems associated with conventional SEM are readily overcome in the environmental scanning electron microscope (ESEM). This instrument permits the presence of a gas, in this case water vapour, in the sample chamber. Through appropriate control of this gas pressure (e.g. Cameron and Donald, 1994) tissues can be maintained in their native state (Uwins et al., 1993). Furthermore, the gas molecules are themselves ionized by electrons traversing the chamber, generating positive ions which drift back towards the sample, thereby alleviating the build-up of charge which would otherwise occur. This charge compensation obviates the need for a conductive coating. These two factors together mean that in the ESEM, biological tissue can be examined in its native state and at a limited range of vapour pressures, covering controlled water contents, with minimal sample preparation, other than producing a sample of the correct size (which may be 10 mm in length). Particles in the 100 nm size range may be imaged in an aqueous environment (Miller and Cooper, 2002). Of relevance to plant cell walls, swelling of cellulose fibres has been observed by Jenkins and Donald (1996), and cellulose ribbons have been imaged pulled out from bacterial cellulose–pectin composites (Astley et al., 2001).

Mechanical test rigs for subjecting samples to strain and measuring forces have been included in the ESEM in studies of polymer films (He and Donald, 1997; Thomas and Wollenden, 1998) and wood fibres (Mott et al., 1995, 1996). Thiel and Donald (1998, 1999) used these advantages to initiate studies into the in situ deformation of carrot parenchyma tissue using a specially constructed mechanical stage. This stage permitted cooling to sub-ambient temperatures via a Peltier chip to facilitate retention of the tissue in a fully hydrated state. The lower the temperature, the lower the saturated vapour pressure, and therefore the easier it is to maintain a working pressure in the chamber that is low, yet sufficient to prevent dehydration. These authors examined various modes of testing, including cutting, and the effects of ageing and cooking. They showed that it was possible to image the distortions produced by the stress distribution in the tissue, and follow the changing distortions of the cell walls and crack propagation in real time. Furthermore, this work has already built a bridge to theory on the effect of turgor on mechanical properties (Warner and Edwards, 1988; Warner et al., 2000).

This study describes how the ESEM has been used to probe failure of a well-defined hydrated tissue, the onion upper epidermal layer (which is more easily excised and prepared as a thin layer than carrot tissues). Previous work has examined the structure and mechanical properties of upper epidermal layers of onion (Ng et al., 2000) and the molecular orientation and intermolecular association in its cell walls when stretched (Wilson et al., 2000). The ability to control hydration, and thereby turgor, allowed investigation of failure at the cell wall level using stretching apparatus developed previously (Thiel and Donald, 1998). The aim was to characterize cell separation and cell breakage events in situ in hydrated tissue and compare the latter with failure in excised cell walls of the charophyte alga Chara corallina. The extremely large size of the individual cells of charophytes such as C. corallina makes it possible to obtain material to study the mechanical properties of plant cell walls. These giant algae consist of a linear series of nodes and internodes. The nodes are multicellular, with lateral cells of limited and unlimited growth and the internodes are single cylindrical cells with a central vacuole surrounded by a thin layer of cytoplasm. These cells are large enough to be opened up to give intact sheets of primary cell wall suitable for mechanical testing (Toole et al., 2001). Cleland (1971) stated that primary cell walls from the tissues of higher plants show mechanical properties that are similar to those determined for algal internode walls.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS AND DISCUSSION
 LITERATURE CITED
 
Plant material
Brown onion (Allium cepa L.) bulbs were obtained from a local supplier. The outer brown papery leaf bases and underlying outermost fleshy leaf bases were removed. Squares of tissue approx. 25 mm x 25 mm were excised from the next fleshy leaf base using a scalpel blade. The upper epidermal layer from the inner (concave) surface of the tissue was excised carefully using a pair of forceps and placed immediately into de-ionized water. The epidermis was spread out on a glass plate beneath a low-power binocular microscope and cut into pieces of the required size using a combination of razor and scalpel blades. Typically, a section approx. 15 mm x 5 mm was prepared for analysis. For stretching experiments, a notch of approx. 1–2 mm was cut into the section at approximately the halfway point, prior to attachment to the stretching apparatus. Epidermal sections were mounted on the tensometer stage of the ESEM and covered with a few drops of de-ionized water.

Orientation of the epidermal tissue (upper or lower surface) was checked by the behaviour of water on the surface. If the cuticle was uppermost, surface water formed globules on the waterproof, waxy cuticle. If the cuticle was downmost, the water dispersed immediately across the section.

To facilitate separation of epidermal cells during stretching, the epidermal tissue was extracted in CDTA (cyclohexane-trans-1,2-diamine-N,N,N',N'-tetraacetate) (0·05 M, pH 6·5) for 4–16 h prior to analysis by ESEM.

Chara corallina was grown in 20-l glass tanks in an artificial pond solution of 1 mM NaCl, 0·1 mM KCl and 0·1 mM CaCl2 (Dainty and Hope, 1959; Smith and West, 1969). The tanks contained a thin layer of sand on the bottom covered with local autoclaved (Gertel and Green, 1977) soil to a depth of 10 cm. Artificial light at an illuminance of 4000 lux was provided from above for 12 h d–1 (black plastic around the tank prevented exposure to excess light) and the temperature was maintained at 24 °C using water heaters. Branches of algae were taken from the tanks on the day of use and stored in distilled water. Sheets of cell wall several centimetres in length and 2–4 mm in width were excised from the giant cells. These were notched at the midpoint of one side and then stretched using the ESEM tensometer stage. These samples were not treated with CDTA.

Environmental scanning electron microscopy apparatus
Experiments were performed in an ElectroScan model 2010 ESEM with a tungsten filament. An accelerating voltage of 20 kV, a beam diameter of 20 nm and a working distance of 8·5 mm were used in all experiments. Procedures to reduce beam damage (Jenkins and Donald, 1996; Royall et al., 2001) were followed.

Using double-sided adhesive tape, a section of the sample was mounted onto the jaws of a small tensometer (Fig. 1) designed to fit onto the stage of the microscope (Thiel and Donald, 1998). The maximum jaw separation rate was 9 µm s–1, but the movement was stopped to record micrographs. A block of copper that could be heated or cooled was positioned between the jaws of the tensometer and the notched region of the specimen was held in contact with this stage to ensure adequate temperature control in the area of interest. The block was fitted with a platinum resistance thermometer, which not only measured the temperature of the block, but also acted as a sensor to a temperature controller driving a Peltier chip. The sample was stretched across the jaws and maintained in contact with the copper block.



View larger version (33K):
[in this window]
[in a new window]
  		Fig. 1. Simplified schematic view of the tensometer attachment showing components and position of sample. Movement direction is shown by bold arrows.

 
The temperature of the block and the pressure of water vapour in the chamber were arranged such that only a slight change in temperature or pressure would allow water to condense onto the sample (Stokes et al., 1998). These conditions were met when the pressure was 667 Pa and the temperature of the block was 2 °C.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
LITERATURE CITED
 
Initial attempts at visualizing the specimen highlighted the importance of precise temperature and humidity control. The most successful technique involved placing the specimen on the pre-cooled stage (2 °C) with a drop of water on the upper surface and excess water on the stage to ensure water was present during pumpdown. After evacuating the chamber, surface water was removed slowly through temperature and humidity control.

The susceptibility to beam damage varies with biological sample (Jenkins and Donald, 1996), but there was no visible electron beam-induced damage under the conditions used in this experiment with the beam rastered over an area of 300 x 300 nm. However, damage was induced in separate experiments in which the focused area was reduced to 50 x 50 nm at this accelerating voltage.

Cell rupture and release of contents may occur at cellular junctions ahead of the notch as it propagates
Initial studies involved stretching freshly prepared epidermal tissue (Fig. 2). The tissue fractured through the cells in a relatively straight line. However, the progression of the notch (‘n’, Fig. 2) was difficult to visualize owing to the release of intracellular contents from deflating cells. It was, however, possible to watch cells in the line of the notch lose their turgor. Often, cell rupture and release of contents occurred at cellular junctions ahead of the notch (see arrows, Fig. 2A and B). Figure 2 shows how cell ‘x’ lost its turgor by this route.



View larger version (76K):
[in this window]
[in a new window]
  		Fig. 2. Sequence of images with increasing time corresponding to stretching a notched sample of onion upper epidermis, mounted cuticle downmost. Release of cell contents (arrowed) and deflation of cell ‘x’ during fracture. n, Notch. Bars = 250 µm (A and B), 350 µm (C).

 
Cells that separated maintain their turgor
It has long been established that cell separation can be induced in some plant tissues by soaking in agents that chelate divalent cations such as calcium. However, there is little information regarding the dynamic fracture of tissues by cell separation at the cellular level, even though this forms the basis for textural softening during the ripening of fruits and thermal processing of vegetables (Van Buren, 1979; Redgewell and Selvendran, 1986).

Stretching of the extracted epidermis using the tensometer stage in the ESEM caused tissue rupture through cell separation (Fig. 3). Most cells that separated maintained their turgor, indicating that some mechanism was effectively sealing any plasmodesmata that might allow release of cell contents, but some occasionally deflated (e.g. ‘d’, Fig. 3A).



View larger version (58K):
[in this window]
[in a new window]
  		Fig. 3. Micrographs at different stages of stretching a notched sample of CDTA-treated onion upper epidermis, mounted cuticle downmost. A, Single deflated cell (d) adjacent to junction of separated cells for a notch being propagated parallel to the major axis of the cells. Bar = 300 µm. B, Higher magnification of an earlier stage of separating cells showing strands (s) of wall-based material between two separating cells. Bar = 100 µm. C, Shows failure perpendicular to the major axis of the cells. Instead of separation along a continuous almost linear cell–cell interface, the failure occurred at a cell–cell interface that is irregular. The scored brass substrate is shown in the lower half of picture. Bar = 200 µm.

 
Strands of material between separating onion cells and between the layers of rupturing algal wall
At high magnification, strands of material were visualized between separating onion cells (‘s’ in Fig. 3B) providing novel insight into pectic polymer behaviour during tissue fracture. This shows that separation of cells was not ‘clean’ or rapid at the cell wall–middle lamella interface, but within middle lamellar material which was highly deforming and adhered to both cells during the early stages of separation. These strands are likely to contain pectic polysaccharides that have become solubilized by the chelator. The weakening of adhesion facilitated cell separation even when the line of fracture was perpendicular to the orientation of the cells (Fig. 3C) to leave a ‘toothed’ fracture in which the separated cells retained their turgor. Thus, there is a different structural failure path brought about by the packing of the highly anisotropic cells. The fracture is dissipated over a large surface area perpendicular to the major axis of the cells. Mechanical anisotropy in the upper epidermal layer between the directions of major and minor axes of cells has been noted (Ng et al., 2000).

The rupture of turgid onion epidermal cells released cellular contents (Fig. 2) that prevented visualization of the point of wall fracture. Cell walls from Chara corallina were used to investigate this event more effectively. The results for notched walls of C. corallina (Fig. 4A) showed clearly the point of fracture (arrowed) of this fully hydrated cell wall, and revealed strands (‘s’) between separating layers (Fig. 4B) some of which were apparently similar to those between onion cells in Fig. 3B. Further deformation showed the tearing of the different layers (‘l’), comprising microfibrils oriented as a function of wall thickness (Richmond, 1983), leaving thinner protruding strands (Fig. 4C).



View larger version (80K):
[in this window]
[in a new window]
  		Fig. 4. Progressively stretched notched sample of excised cell wall of Chara corallina. A, Head of notch indicated by an arrow. Bar = 200 µm. B, Crack propagated showing strands (s) of material between fractured surfaces. Some condensed water droplets shown at top left of picture. The layered structure of the wall is apparent showing fragments at different depths. Bar = 100 µm. C, Notch has now moved to right of the micrograph. Broken edges of cell wall showing ruptured layered structure (l) and broken fibrous strands. Bar = 20 µm.

 
These are features of importance in relation to future modelling of plant structure, facilitating and building on current cell wall models. For example, the molecular network comprising the expanding primary cell wall featuring cleavage of xyloglucan and microfibril separation has been modelled by a number of authors, including Carpita and Gibeaut (1993), but the behaviour under larger strains and the microscopic nature of the failure have not been described. The next stage would be a molecular description of failure of multiple cells at a higher level of structure, whether by cell separation or breakage.

The present work has extended previous ESEM observations (Thiel and Donald, 1998) of propagation of failure in a fully hydrated bulk tissue to deformation at the scale of cell separation and breakage modes of failure in a tissue layer. In the case of separating cells, the formation and progressive breakage of strands has been observed between cells that largely retained their turgor. Cell breakage was characterized by loss of turgor and loss of cell contents remotely from the breakage zone or notch in the tissue. The simpler case of breakage within walls alone showed that in addition to the breaking strands there was a thinner strand breakage involving the layers of a large algal wall.


   ACKNOWLEDGEMENTS
 
A.C.S. and K.W.W. were funded by the Biotechnology and Biological Sciences Research Council Competitive Strategic Grant.


   LITERATURE CITED
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 LITERATURE CITED
 

    Astley O, Chanliaud E, Donald AM, Gidley MJ. 2001. Structure of Acetobacter cellulose composites in the hydrated state. International Journal of Biological Macromolecule 29: 193–202.[CrossRef]

    Cameron RE, Donald AM. 1994. Minimising sample evaporation in the environmental scanning electron microscope. Journal of Microscopy 173: 227–237.

    Carpita NC, Gibeaut DM. 1993. Structural models of primary-cell walls in flowering plants – consistency of molecular structure with the physiological properties of the wall during growth. Plant Journal 3: 1–30.[CrossRef][Web of Science][Medline]

    Cleland RE. 1971. Cell wall extension. Annuals Reviews in Plant Physiology 22: 197–222.

    Dainty J, Hope AB. 1959. Ionic relations of cells of Chara australis. Australian Journal of Biological Sciences 12: 395–411.

    Gertel ET, Green PB. 1977. Cell growth pattern and wall microfibrillar arrangement. Plant Physiology 60: 247–254.[Abstract/Free Full Text]

    He C, Donald AM. 1997. Morphology of a deformed rubber toughened poly (methyl methacrylate) film under tensile strain. Journal of Materials Science 32: 5661–5667.[CrossRef]

    Jenkins LM, Donald AM. 1996. Use of the environmental scanning electron microscope for the observation of the swelling behaviour of cellulosic fibres. Scanning 19: 92–97.

    McGarry A. 1995. Cellular basis of tissue toughness in carrot (Daucus carota L.) storage roots. Annals of Botany 75: 57–163.

    Miller AF, Cooper SJ. 2002. In situ imaging of Langmuir films of Nylon-6,6 polymer using environmental scanning electron microscopy. Langmuir 18: 1310–1317.[CrossRef]

    Mott L, Shaler SM, Groom LH. 1996. A technique to measure strain distributions in single wood pulp fibers. Wood and Fiber Science 28: 429–437.

    Mott L, Shaler SM, Groom LH, Liang B-H. 1995. The tensile testing of individual wood fibers using environmental scanning electron microscopy and video image analysis. Technical Association of the Pulp and Paper Industry Journal 78: 143–148.

    Ng A, Parr AJ, Parker ML, Saunders PK, Smith AC, Waldron KW. 2000. Structural and mechanical characteristics of onion (Allium cepa L). Journal of Agricultural and Food Chemistry 48: 5612–5617.[CrossRef][Web of Science][Medline]

    Redgewell RJ, Selvendran RR. 1986. Structural features of cell-wall polysaccharides of onion (Allium cepa). Carbohydrate Research 157: 183–199.[CrossRef]

    Richmond PA. 1983. Patterns of cellulose microfibril deposition and rearrangement in Nitella. In vivo analysis by a birefringence index. Journal of Applied Polymer Science: Applied Polymer Symposium 37: 107–122.

    Royall CP, Thiel BL, Donald AM. 2001. Radiation damage of water in environmental scanning electron microscopy. Journal of Microscopy 204: 185–195.[Web of Science][Medline]

    Smith FA, West KR. 1969. A comparison of the effects of metabolic inhibitors on chloride uptake and photosynthesis in Chara corallina. Australian Journal of Biological Sciences 22: 351–363.

    Stokes DJ, Thiel BL, Donald AM. 1998. Direct observation of water-oil emulsion systems in the liquid state by environmental SEM. Langmuir 14: 4402–4408.[CrossRef]

    Thiel BL, Donald AM. 1998. In-situ mechanical testing of fully hydrated carrots (Daucus carota L.) in the environmental SEM. Annals of Botany 82: 727–733.[Abstract/Free Full Text]

    Thiel BL, Donald AM. 1999. Microstructural failure mechanisms in cooked and aged carrots. Journal of Texture Studies 31: 437–455.

    Thomas V, Wollenden A. 1998. A tensile stage to study thin polymer films in an environmental scanning electron microscope. Reviews of Scientific Instruments 69: 463–465.[CrossRef]

    Toole GA, Gunning PA, Parker ML, Smith AC, Waldron KW. 2001. Fracture mechanics of the cell-wall of Chara corallina. Planta 212: 606–611.[CrossRef][Web of Science][Medline]

    Uwins PJR, Murray M, Gould RJ. 1993. Effects of four different processing techniques on the microstructure of potatoes – comparison with fresh samples in the ESEM. Microscopy Research and Technique 25: 412–418.[CrossRef][Web of Science][Medline]

    Van Buren JP. 1979. The chemistry and texture in fruits and vegetables. Journal of Texture Studies 10: 1–23.

    Waldron KW, Smith AC, Parr AJ, Ng A, Parker ML. 1997. New approaches to understanding and controlling the effects of cell separation on fruit and vegetable texture. Trends in Food Science and Technology 8: 213–221.

    Warner M, Edwards SF. 1988. A scaling approach to elasticity and flow in solid foams. Europhysics Letters 5: 623–628.

    Warner M, Thiel BL, Donald AM. 2000. The elasticity and failure of fluid-filled cellular solids – theory and experiment. Proceedings of the National Academy of Sciences of the USA 97: 1370–1375.[Abstract/Free Full Text]

    Wilson RH, Smith AC, Kacurakova M, Saunders PK, Wellner N, Waldron KW. 2000. The mechanical properties and molecular dynamics of plant cell wall biopolymers studied by novel infrared spectroscopic techniques Plant Physiology 124: 397–406.[Abstract/Free Full Text]


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?



This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Content Snapshot
Right arrow Content Select
Right arrow All Versions of this Article:
92/1/73    most recent
mcg115v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (10)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Agricola
Right arrow Articles by DONALD, A. M.
Right arrow Articles by WALDRON, K. W.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?