New insight into silica deposition in horsetail (Equisetum arvense)
© Law and Exley; licensee BioMed Central Ltd. 2011
Received: 15 April 2011
Accepted: 29 July 2011
Published: 29 July 2011
The horsetails (Equisetum sp) are known biosilicifiers though the mechanism underlying silica deposition in these plants remains largely unknown. Tissue extracts from horsetails grown hydroponically and also collected from the wild were acid-digested in a microwave oven and their silica 'skeletons' visualised using the fluor, PDMPO, and fluorescence microscopy.
Silica deposits were observed in all plant regions from the rhizome through to the stem, leaf and spores. Numerous structures were silicified including cell walls, cell plates, plasmodesmata, and guard cells and stomata at varying stages of differentiation. All of the major sites of silica deposition in horsetail mimicked sites and structures where the hemicellulose, callose is known to be found and these serendipitous observations of the coincidence of silica and callose raised the possibility that callose might be templating silica deposition in horsetail. Hydroponic culture of horsetail in the absence of silicic acid resulted in normal healthy plants which, following acid digestion, showed no deposition of silica anywhere in their tissues. To test the hypothesis that callose might be templating silica deposition in horsetail commercially available callose was mixed with undersaturated and saturated solutions of silicic acid and the formation of silica was demonstrated by fluorimetry and fluorescence microscopy.
The initiation of silica formation by callose is the first example whereby any biomolecule has been shown to induce, as compared to catalyse, the formation of silica in an undersaturated solution of silicic acid. This novel discovery allowed us to speculate that callose and its associated biochemical machinery could be a missing link in our understanding of biosilicification.
Silicon is the second most abundant element of the Earth's crust after oxygen and, perhaps surprisingly, its essentiality in biota remains equivocal . The difficulty in ascribing true biochemical essentiality to silicon probably emanates from a lack of demonstration of any silicon-requiring biochemistry and specifically Si-C, Si-O-C, Si-N, et c. bonds in any form of extant life . However, in spite of such limitations the essentiality of silicon in plants remains the subject of rigorous debate [3, 4] as do elaborations of the underlying mechanisms. Biosilicification was recently defined as 'the movement of silicic acid from environments in which its concentration does not exceed its solubility (< 2 mM) to intracellular or systemic compartments in which it is accumulated for subsequent deposition as amorphous hydrated silica'  and a number of plants are known biosilicifiers . One of the best known of these are the horsetails, Equisetum sp., and silica deposition in the tissues of these plants has been studied extensively [6–12], perhaps the seminal work in the field being carried out by Perry and Fraser . In this work scanning and transmission electron microscopy was used to illuminate the elaborate and detailed micromorphology and ultrastructure of silicas extracted from different regions of the horsetail, Equisetum arvense. The images of silicified stomata and other silica sculptures are truly breathtaking and the level of organisation of silica in the tissues prompted the authors to speculate that 'the silica acts as an in vivo stain, faithfully reproducing the organic matrix skeleton at the microscopic and macroscopic levels without staining'. Perry and Lu (1992) suggested that the organic matrix in question might be made from polymers of carbohydrates, for example, cellulose , and this suggestion was reinforced recently by Fry and colleagues who speculated that the hemicellulose, callose, in horsetail cell walls might be a potential site of silica deposition . Many different biomolecules, often having originally been extracted from biogenic silica, have been shown to accelerate or catalyse silica deposition in saturated solutions of silicic acid . However, biosilicifiers, such as horsetails, harvest silicic acid from solutions which are far from saturation and deposit it as amorphous hydrated silica and it is the elucidation of this mechanism which remains the 'Holy Grail' of biological silicification research .
Herein we have taken inspiration from the work of Perry and Fraser  on horsetail and we have used fluorescence microscopy to investigate biosilicification in horsetail and to identify the organic matrix involved in templating silica deposition in this plant.
PDMPO as a fluorescent marker of biosilicification
PDMPO as a fluorescent indicator of silica formation in vitro
The fluor PDMPO has been used to identify silica deposition in horsetail and to provide new insight into silicification in this plant. It was observed that silica deposition in horsetail exactly mirrored the known deposition of callose in the related fern and other plants. Callose was shown to induce the formation and precipitation of silica in undersaturated solutions of silicic acid. This was the first time that this had been demonstrated for any biomolecule and it suggested that callose and perhaps other similar carbohydrates might be key molecules in biological silicification.
Hydroponic culture of horsetail
Horsetail (Equisetum arvense) rhizomes were collected locally, washed in ultrapure water (conductivity < 0.067 μS/cm) and subjected to hydroponic culture in 1/6th MS medium in the presence (2 mM) or absence of added silicic acid. The latter media included an additional 8 mM Na+ to account for Si addition as Na4SiO4. After 10-12 weeks of a 14 h light/10 h dark cycle at 25°C healthy horsetail plants had grown under both sets of conditions.
Digestion of horsetail materials
Horsetail plants, either collected locally or grown hydroponically, were washed in ultrapure water, allowed to air-dry, cut into discrete 1 cm sections of rhizome/root, basal stem, distal stem, nodal regions and leaves and ca 0.5 g of each placed in acid-washed 20 mL PFA teflon© vessels. The samples were then digested in a 1:1 mixture of 15.8M HNO3 and 18.4M H2SO4 using a Mars Xpress microwave oven (CEM Microwave Technology Ltd.). The acid digests were clear and, upon dilution with 8 mL of ultrapure water, were filtered and the residues washed several times with further volumes of ultrapure water. Filters were then placed in plastic Petri dishes in an incubator at 37°C to achieve dryness over several days. Dry residues off each filter were then collected into Bijoux tubes and stored in a dry, sealed, perspex cabinet.
PDMPO labelling of horsetail silica
Silica residues collected from filters were suspended in 20 mM PIPES at pH 7 and containing 0.125 μM 2-(4-pyridyl)-5-((4-(2-dimethylaminoethylaminocarbamoyl) -methoxy)phenyl)oxazole (PDMPO; LysoSensor Yellow/Blue DND-160, 1 mM in DMSO). This intracellular pH probe  has been shown to be bound by silica (but not silicic acid) and to emit 'green' fluorescence upon excitation at 338 nm [35–38]. Suspensions were left for 24 h to allow the reaction between silica surfaces and PDMPO to achieve completion after which 50 μL samples were transferred to a cavity slide and viewed using an Olympus BX50 fitted with a BXFLA fluorescent attachment using a U-MWU filter cube (Ex: 333-385 nm; Em: 400-700 nm). A ColourView III digital camera (OSIS FireWire Camera 3.0 digitizer) was used to capture images in conjunction with CELL* Imaging software (Olympus Cell* family, Olympus Soft Imaging solutions GmbH 3.0).
In vitro preparations of callose and silicic acid
Callose (β-D Glucan, Barley, Sigma, UK) was dissolved at 5% w/v in 20 mM PIPES buffer solutions at pH 7 and containing 0, 1, 2, 4 and 7 mM Si(OH)4 by warming each preparation in a water bath at 100°C for 60 seconds. Upon cooling to room temperature PDMPO was added to a concentration of 0.125 μM. Equivalent control solutions to which no callose had been added were treated in an identical manner. All solutions were then incubated at room temperature in the dark for 5 days before being examined by fluorescence microscopy, see above, or their emission spectra were determined by fluorimetry (Perkin-Elmer LS50B; Ex; 338 nm; Em: 400-650 nm) as previously described .
CL was in receipt of a NERC studentship.
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