Journal of Experimental Botany

Tripathi, Durgesh, Kumar;Singh, Vijay, Pratap;Lux,, Alexander;Vaculik,, Marek

doi: 10.1093/jxb/eraa448pmid: 33264414

Abiotic stress, aquaporins, apoplasmic barriers, beneficial elements, metals and metalloids, pests and diseases, phytoliths, silicon, Si-transport pathways, stress alleviation Silicon (Si) is the second most abundant element in the Earth’s crust after oxygen, and it has wide implications in plant biology. The effects of Si range from regulation of development to protection of plants from various types of stresses. In recent years, much progress has been made in research on the uptake of Si at the root surface, its loading into the xylem, and its transport to various tissues in Si-accumulating plants. However, much still remains to be discovered. For instance, there is currently no direct evidence for the participation of Si in plant metabolism, and little detail is known of the mechanisms by which Si functions in plants. In this special issue, we present a collection of papers that attempt to answer such questions, but we are still a long way from fully understanding the role of Si in plant biology. Silicon research in past years It has long been known that Si has beneficial effects on plants, and research in the early 20th century identified its potential for use as a plant growth supplement. Prior to that, Lupton (1880) noted its accumulation in wheat, corn, and grasses and suggested its presence was benificial to the plants. Sommer (1926) demonstrated enhanced growth of rice and millet in the presence of Si, and found that seed heads of millet grown in its absence were more severely infected by pathogenic fungi. Beet production was found to increase by 44% in acid soils with high aluminum content when treated with calcium silicate slag as a liming agent as compared with calcium hydroxide (Raleigh, 1939), and it was also observed that roots became necrotic and were infected with fungus in the absence of Si. In the second half of the 20th century, several studies using either pot-grown plants or hydroponics documented positive effects of Si in increasing yields and improving plant resistance to biotic and abiotic stresses (e.g. Woolley, 1957; Bollard and Butler, 1966; Lewin and Reimann, 1969). Jones and Handreck (1967) provided a detailed account of the importance of Si in agriculture, and plant physiology and pathology; indeed, there was a discussion as to whether it should be included in the list of essential elements, if not for all plants in general then at least for certain species such as horsetail and sugarcane (Chen and Lewin, 1969; Fox and Silva, 1978). The last decades of the 20th century through to the present time might be regarded as a ‘golden era’ of Si research. Several studies have demonstrated the importance of Si in crop resistance to pathogens, pests, and various kinds of abiotic stresses, and there has been a focus of research on Si management in sustainable crop production and its roles in plant life (e.g. Epstein, 1994; Birchall, 1995; Hodson and Evans, 1995; Datnoff et al., 1997; Hattori et al., 2005; Mitani and Ma, 2005). This interest in Si is reflected by the increasing number of papers that have been published on the topic in recent decades (Fig.1). This has been driven by a desire to better understand its role in plants combined with the increasing availability of improved technical methods and approaches for its detailed study. Fig. 1. Open in new tabDownload slide The number of papers focused on Si research related to plants in the last 25 years (A) and their countries of origin according to corresponding author (B), as listed in the Web of Science (data from August 2020). Fig. 1. Open in new tabDownload slide The number of papers focused on Si research related to plants in the last 25 years (A) and their countries of origin according to corresponding author (B), as listed in the Web of Science (data from August 2020). Recent progress and breakthroughs in silicon research Recent decades have brought a considerable increase in our knowledge regarding the uptake and translocation of Si in plant tissues. The breakthrough discovery of the first Si transport-channel protein, Lsi1, in rice published by Ma et al. (2006) prompted a search for Si transport mechanisms in other plant species. Our knowledge of Si-uptake pathways has thus been considerably improved in the last 15 years, and this progress is reviewed by Mandlik et al. (2020) in this special issue. Silicon is taken up from the soil solution in the form of silicic acid (Epstein, 1994) and enters cells through channel-type membrane proteins called aquaporins (Ma and Yamaji, 2015; Deshmukh and Bélanger, 2016). Recent novel findings have been made regarding aquaporins belonging to the nodulin 26-like intrinsic protein group III (NIP-III), which are considered to play a major role in plant Si uptake. Deshmukh et al. (2020) suggest that the aromatic/arginine (ar/R) selectivity filter of the NIP-IIIs does not need to be exclusively composed of the sequence Glycine-Serine-Glycine-Arginine (G-S-G-R), and some amino acids positions seem to be less conserved. However, they suggest that the change of NPA to NPV or an alternative motif structure does not play such an important role as has previously been believed. Instead, the spacing of 108 amino acids between two NPA motifs appears to be one of the most conserved and important features with respect to Si permeability. Deshmukh et al. (2020) also conduct a comprehensive genomic analysis of aquaporins from over a thousand plants species, which reveals that the evolution of NIP-IIIs as primary Si influx transporters dates back as early as 515 million years ago. This confirms previous assumptions that Si accumulation has been an important feature of land plants right from the time at which they left the aquatic environment. They also rule out the traditional accepted opinion that monocots generally take up more Si than dicots. Based on their results, species from several dicot families including the Cucurbitaceae, Fabaceae, and Ateraceae, accumulate considerably higher amount of Si compared with many monocots outside the Poaceae family. In recent years, channels and proteins involved in Si membrane permeability have been recorded in many monocot as well as dicot species (e.g. Yamaji et al., 2012; Bokor et al., 2019; Zellner et al., 2019). In this special issue, Noronha et al. (2020) report that there is an active Si uptake transport system in a species of agricultural importance, namely grapevine. They find that VvNIP2;1 is mostly expressed in roots and also in green berries, it is localized in the plasma membrane, and it codes for functional aquaporin that is able to transport Si as well as arsenite. It has been a matter of debate as to how silicic acid is transported within plants, but it is believed that Si is transported via the xylem to the aerial parts. In common with other compounds and elements, Si needs to cross (so-called) apoplasmic barriers (mostly Casparian bands and suberin lamellae) to enter the central cylinder and xylem veins. In this regard, it has been shown that Si can influence the deposition of lignin and suberin within the root exo- and endodermis and thereby modify plant metal(loid) uptake (e.g. Vaculik et al., 2020). However, Kreszies et al. (2020) note that controversies exist, as both enhanced as well as delayed formation of root apoplasmic barriers in response to Si have been described in the literature. In their own experiments with barley grown under control and osmotic stress conditions, they found no direct effect of Si and suggest that enhanced stress tolerance of plants after Si treatment is due to other responses. Silicon is deposited as amorphous silica in various organs and cells (Mandlik et al., 2020). With regards to the roots, the endodermis has been identified as a site of elevated deposition in various species, and particularly in sorghum (e.g. Lux et al., 2020). This special issue includes a series of studies describing the mechanisms of Si phytolith formation and silica precipitation in sorghum roots and leaves. Soukup et al. (2020) find that the process of Si phytolith deposition within the root endodermis is strictly associated with living and fully metabolic active cells, whilst Zexer and Elbaum (2020) suggest that polymerization of silicic acid occurs only at specific locations in the tangential endodermis cell walls where modification of lignin is present in the form of a lack of ferulic acid. Finally, Kumar et al. (2020) describe a unique protein in sorghum leaves that is responsible for silica precipitation, which they name as Siliplant1 (Slp1). They find that Slp1 is present in developing silica cells and that it is transported to the cell wall during silicification. The ameliorative effects of Si on metal toxicity are a well-known phenomenon, and many papers documenting its positive role have been previously published. Much research has focused on Al, which follows Si as the third most abundant element in the Earth’s crust (Epstein, 1994). In this special issue, Hodson and Evans (2020) review the last quarter-century of research in plant Si–Al interactions, providing an update to their review on the same topic that was published in this journal 25 years ago (Hodson and Evans, 1995). As they note, our knowledge has greatly improved in certain areas, especially in our understanding of Si and Al uptake and transport mechanisms at the molecular level, and some hypothesis regarding the amelioration of Al toxicity by binding with Si in the apoplasm have now been confirmed. However, a comprehensive understanding of a number of processes is still lacking, and the authors question why so little effort has been made in investigating the amelioration of Al toxicity in edible plants. The interactions of Si with other metals as well as metalloids is also the focus of another review by Vaculik et al. (2020). Whilst some elements already interact with Si in the soil and in the rhizosphere, important mitigation processes also occur on various levels inside the plant. Co-deposition in the apoplasm, as occurs with Al, has also been documented for cadmium. Vaculik et al. (2020) identify a number of gaps in our knowledge that need to be addressed if we are to properly understand how Si-based mechanisms alleviative metal(loid) toxicity. More broadly, Ahanger et al. (2020) review the role of Si in stress tolerance in affecting the synthesis of plant defense enzymes and in the integration of secondary metabolites. Another beneficial application of Si might be its use in increasing resistance against various pathogens and herbivores. It is well known that plants that actively take up relatively high levels of Si are more resistant to pests and diseases. In their review in this special issue, Singh et al. (2020) consider both direct aspects of resistance to plant herbivory that are induced by Si, such as strengthening mechanical protection and the synthesis of antioxidant enzymes, and indirect modes of resistance, such as the release of volatiles and alterations in the synthesis of defense-related hormones. A better understanding of the role of Si in host–pathogen interactions is clearly desirable. Rasoolizadeh et al. (2020) show that Si might help to create an unfavorable environment for pathogens in the apoplasm and it probably interferes with host-recognition and/or limits receptor–effector interactions, leading to an incompatible interaction between the pathogen and the host. We currently live in an era in which more environmentally friendly solutions are sought for agricultural problems, and in this context Si might provide a good tool for the improvement of resistance to pests and diseases, not only in cereals but also in other crop species. Perspectives and future challenges for silicon in plant biology Whilst the papers included in this special issue highlight our current knowledge about the uptake, translocation, deposition, and functioning of Si in plants, there are still many questions that remain to be answered, some of which are very fundamental. For instance, we do not know or have only limited knowledge about how Si is loaded into the xylem or the identity of the transporter that pumps Si to silica cells, and at a basic level we do not really know how Si provides such versatile benefits to plants. To date, there have been no reports showing an active role of Si in any biochemical and metabolic pathway that can define the benefits that the plant receives from supplementation with Si. Similarly, we have only limited information regarding the optimum quantities of Si that are needed for better plant growth at individual developmental stages. Hundreds of studies have reported beneficial effects of supplying Si to poor Si-accumulator species, which is hard to explain with our current level of understanding. As well as our lack of knowledge about the structural and functional characteristics of Si transport proteins, other unanswered questions include the role of Si in interactions with signaling molecules under normal as well as stress conditions, its impact on the uptake of nutrients, its influence on the photosynthetic machinery, and its role in the integration of phytohormones. A better understanding of Si biology will provide benefits in many different fields, including agriculture, industrial applications, and ecology. Collaborative research through organizations such as the Society for Silicon in Agriculture and Related Disciplines (ISSAG) should be encouraged to achieve this aim. 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Mandlik,, Rushil;Thakral,, Vandana;Raturi,, Gaurav;Shinde,, Suhas;Nikolić,, Miroslav;Tripathi, Durgesh, K;Sonah,, Humira;Deshmukh,, Rupesh

doi: 10.1093/jxb/eraa301pmid: 32592476

Abstract Numerous studies have shown the beneficial effects of silicon (Si) for plant growth, particularly under stress conditions, and hence a detailed understanding of the mechanisms of its uptake, subsequent transport, and accumulation in different tissues is important. Here, we provide a thorough review of our current knowledge of how plants benefit from Si supplementation. The molecular mechanisms involved in Si transport are discussed and we highlight gaps in our knowledge, particularly with regards to xylem unloading and transport into heavily silicified cells. Silicification of tissues such as sclerenchyma, fibers, storage tissues, the epidermis, and vascular tissues are described. Silicon deposition in different cell types, tissues, and intercellular spaces that affect morphological and physiological properties associated with enhanced plant resilience under various biotic and abiotic stresses are addressed in detail. Most Si-derived benefits are the result of interference in physiological processes, modulation of stress responses, and biochemical interactions. A better understanding of the versatile roles of Si in plants requires more detailed knowledge of the specific mechanisms involved in its deposition in different tissues, at different developmental stages, and under different environmental conditions. Cell wall stability, phytoliths, silicon transport, specialized cells, stress tolerance, xylem loading Introduction Deposition of silicon (Si) in plant tissues has frequently been associated with stress tolerance mechanisms and better resilience under stress conditions (Epstein, 1994; Ma, 2004; Coskun et al., 2019). Despite numerous reports indicating its beneficial effects, Si has never been considered as an essential element for higher plants. To be categorized as such, an element needs to fulfill specific criteria, the most important of which is that the plant would not be able to complete its life cycle in the absence of the element (Kirkby, 2011). Hence, except for the Equisetaceae and some algae (Pontigo et al., 2015), Si has been categorized separately as a ‘beneficial element’ (Epstein, 1994; Ma et al., 2001). Silicon-based fertilizers are being increasingly used worldwide as a result of the accumulating evidence of the protective effects of Si application, and its promotion as a beneficial element by agencies including the International Plant Nutrition Institute (www.ipni.net/topic/silicon-si). Thousands of published papers have indicated beneficial roles of Si in plants and several review articles have provided summaries of these studies (Ma et al., 2001; Fauteux et al., 2005; Bhat et al., 2019; Coskun et al., 2019; Zargar et al., 2019). However, most research efforts have largely failed to explain the basis of Si-derived benefits to plants. To date, there are no significant reports explaining the distinct roles of Si in plants, and we need a better understanding of its uptake at the root level, subsequent transport, and accumulation in different tissues. The beneficial effects of Si supply have largely been attributed to its deposition in plants (Ma et al., 2001; Sangster et al., 2001; Gong et al., 2006), and considerable variation between species in terms of quantity and deposition patterns have been observed (Guerriero et al., 2019, 2020). High levels of accumulation in some dicots and grasses led to the hypothesis that Si plays a role as a mechanical barrier, but recently Coskun et al. (2019) have proposed a working model termed the ‘apoplastic obstruction hypothesis’ that explains many roles of Si beyond just acting as a mechanical obstruction. However, defining all the observed Si-derived benefits to plants with any single model will be challenging. A simple mechanical role looks most obvious in species that accumulate high levels of Si and in those that have specialized silica cells. In monocots, silica cells are almost filled with solid silica. Mature silica cells are also referred as phytoliths, plant opal, or silica bodies (Kumar et al., 2017, 2020). The shape of silica cells varies among species (Piperno and Pearsall, 1998; Piperno et al., 1999; Dabney et al., 2016). For example, in rice leaves they are found in two different forms, dumbbell-shaped and completely silica-filled bulliform-shaped (Zhang et al., 2013). Mature silica cells impart enhanced abrasiveness to grass leaf blades, which deters herbivores, and they have been found to be associated with photosynthetic activities (Massey et al., 2007; Dabney et al., 2016). Cells filled with Si at the base of wheat awns respond to humidity to provide propulsion that is essential for seed dispersion (Elbaum et al., 2007). Many other specific Si deposition patterns with important roles have been reported for a variety of species (Sangster et al., 2001). Beneficial effects of Si are not only restricted to species capable of high uptake, similar effects have been observed in species with poor accumulation, which appears puzzling. However, recent studies have shown how Si deposition in cell walls can help improve nutrient transport across the membrane (Sheng et al., 2018). Optimum requirements for Si across different species are not yet well defined, and similarly the physiological effects of different amounts of deposition in different tissues are largely unknown. Over the last 15 years, significant progress has been made towards understanding Si uptake in roots and its subsequent transport to different tissues. However, the transporters at the connecting tissues within specific cell types and organs need further study and characterization. This review discusses our current understanding of the roles of Si, the molecular mechanisms involved in its transport, the pattern of its deposition, and the effects of its accumulation in different tissues. In doing so, we highlight gaps in our knowledge that need to be addressed. Phylogenetic variation in silicon uptake and accumulation Wide variation in accumulation of Si has been observed, with concentrations ranging from 0.1% to 10% on a dry-weight basis (Epstein, 1999; Ma and Takahashi, 2002). Bryophyta, Lycopsida, and Equisetopsida within the Pteridophyta showed higher accumulation of Si as compared to most of the angiosperms (Ma and Takahashi, 2002). In angiosperms, species belonging to the Poaceae and Cyperaceae are known to accumulate relatively high amounts of silicon, whilst the Commelinaceae, Urticaceae, and Cucurbitaceae are intermediate in accumulation (Ma and Takahashi, 2002; Hodson et al., 2005). The Brassicaceae and Solanaceae are well documented as being poor accumulators or Si excluders (Hodson et al., 2005; Sonah et al., 2017). Rice (Poaceae) can accumulate up to 10% Si and is considered as a model species to study its biological roles (Ma et al., 2002). Meta-analysis by Hodson et al. (2005) showed the following order for Si concentrations in various groups (high to low): liverworts > horsetails > clubmosses > mosses > angiosperms > gymnosperms > ferns. Early speculation that considered monocots as strong Si accumulators and dicots as poor accumulators were ruled out by studies conducted by Epstein (1999). The variations in Si uptake and accumulation can be attributed to specific transporter proteins in roots. Silicon uptake and transport Silicon is present in the soil as an inert element. Its uptake and transportability mainly depend on the plant roots and its chemical composition in the soil. Silicon is taken up by roots as monosilicic acid [Si(OH)4], which is the soluble form present in soil at pH<9 and concentration below 2 mM Si. After uptake and transportation to the shoots, as a result of transpiration, Si concentrates and polymerizes into colloidal silica gel (SiO2.·nH2O) (Yoshida et al., 1962), although various other biomolecules are also thought to play a role (Harrison, 1996). Plant species are categorized as accumulators, intermediate, or non-accumulators/excluders (Ma et al., 2001). Uptake of silicic acid in rice is much faster than that of water (Ma et al., 2001), indicating the possibility of an active transport mechanism facilitating Si transport across the cell membrane. Apoplastic pathway in rice roots are blocked by the development of Casparian strip and hence translocation of Si needs to be through the symplastic pathway. Tamai and Ma (2003) studied the uptake kinetics of Si and predicted the involvement of transporter proteins in the uptake mechanism, and numerous subsequent studies have helped to further reveal the molecular mechanisms involved in uptake (e.g. Ma et al., 2006, 2011; Markovich et al., 2019). A seminal study by Ma et al. (2006) using an induced mutagenesis approach identified the first Si transporter protein in rice. A mutant genotype lacking Si uptake, named as the lsi1 mutant, was used to identify the transporter gene Lsi1. Lsi1 is a Si influx transporter belonging to the aquaporin subfamily nodulin 26-like intrinsic proteins (NIPs). All aquaporins have a conserved hourglass structure with six alpha helix transmembrane domains (H1–H6) that are joined by five loops (LA–LE). The loops LB and LE contain a conserved motif, the NPA domain. A pore passing through the aquaporin has two constrictions that have a significant role in defining solute specificity and transport kinetics. One of the constriction is formed by two conserved NPA domains and the second constriction is formed by an ar/R selectivity filter consisting of four amino acids, which are from the helices H2 and H5, and loop LE. Lsi1, which transports uncharged silicic acid in particular, belongs to the NIP III group, comprising a unique selectivity filter Gly (G), Ser (S), Gly (G), and Arg (R) (Mitani-Ueno et al., 2011). Shortly after the discovery of Lsi1, a second Si transporter gene, Lsi2, was identified by the same group using the same mutagenesis approach (Ma et al., 2007). Lsi2 is an efflux transporter that actively pumps Si out of the cell with the help of a proton gradient. As an active transporter, its homology and mechanism are completely different from Lsi1. Lsi2 is an anion transporter coupled with a proton antiport having 9–12 transmembrane domains (Ma et al., 2007, 2011). The identification of the Lsi1 and Lsi2 transporters has largely explained the molecular mechanism involved in the uptake of Si by roots. Rice takes up Si as monosilicic acid through Lsi1 present on the distal side of the exodermis and pumps it in a de-protonated form out in aerenchyma by Lsi2 at the proximal side of the cell (Fig. 1a). Silicic acid then passes through the apoplast of aerenchyma. In the endodermis, the apoplastic pathway is blocked by the Casparian strip, which forces Si to be transported via the symplastic pathway through Lsi1 and then Lsi2 to the xylem. The silicic acid moves upwards to the shoot via the transpiration stream, where Lsi6 (an influx transporter, homolog of Lsi1) unloads Si from the xylem and facilitates its transport to the different aerial parts of the plant. Fig. 1. Open in new tabDownload slide Schematic representation of silicon transport and xylem loading. (a) Silicon (Si) transport through roots. Lsi1, which is a passive transporter, facilitates the entry of Si (as silicic acid) into the exodermis. Silicic acid exits the exodermis and enters the cortex through the active transporter Lsi2, and it then moves apoplastically through aerenchyma to reach the endodermis. Lsi1 and Lsi2 transport silicic acid through the endodermis and it is loaded into the xylem. (b) The transport of Si in node I. Lsi6, a polar transporter, transfers the silicic acid into the xylem transfer cells, from where it moves through the plasmodesmata to the bundle sheath cells. Lsi2 located at the distal ends of these cells transports some of the silicic acid into the apoplast, from where it moves into the xylem of diffused vascular bundles, whilst the rest of the Si is exported and loaded into the xylem by Lsi3. EVB, enlarged vascular bundle; BS, bundle sheath; XTC, xylem transfer cells; NVA, nodal vascular anastomosis; DVB, diffuse vascular bundle. (c) Xylem unloading into silica cells in leaves. Silicon is pumped into the xylem parenchyma cells by the Lsi6 transporter and is transported through them to be deposited into the silica cells. The transporter involved in transferring Si against a concentration gradient into the silica cells remains unknown. Fig. 1. Open in new tabDownload slide Schematic representation of silicon transport and xylem loading. (a) Silicon (Si) transport through roots. Lsi1, which is a passive transporter, facilitates the entry of Si (as silicic acid) into the exodermis. Silicic acid exits the exodermis and enters the cortex through the active transporter Lsi2, and it then moves apoplastically through aerenchyma to reach the endodermis. Lsi1 and Lsi2 transport silicic acid through the endodermis and it is loaded into the xylem. (b) The transport of Si in node I. Lsi6, a polar transporter, transfers the silicic acid into the xylem transfer cells, from where it moves through the plasmodesmata to the bundle sheath cells. Lsi2 located at the distal ends of these cells transports some of the silicic acid into the apoplast, from where it moves into the xylem of diffused vascular bundles, whilst the rest of the Si is exported and loaded into the xylem by Lsi3. EVB, enlarged vascular bundle; BS, bundle sheath; XTC, xylem transfer cells; NVA, nodal vascular anastomosis; DVB, diffuse vascular bundle. (c) Xylem unloading into silica cells in leaves. Silicon is pumped into the xylem parenchyma cells by the Lsi6 transporter and is transported through them to be deposited into the silica cells. The transporter involved in transferring Si against a concentration gradient into the silica cells remains unknown. The molecular mechanism of Si uptake varies amongst species. Homologs of both Lsi1 and Lsi2 have been identified in monocot crops such as wheat, barley, sorghum, and maize, with variations in their localization (Mitani et al., 2009). In maize, Si deposition is mediated by two genes, ZmLsi1 and ZmLsi6. ZmLsi1 is responsible for the uptake of Si through roots, whilst ZmLsi6 is located in the parenchyma cells of the leaves and is responsible for xylem unloading (Bokor et al., 2015). Homologs of OsLsi2 responsible for efflux have been found in barley (HvLsi2) and maize (ZmLsi2) and functions in a similar manner, but they are present only in the endodermis and do not show polar localization (Mitani et al., 2009). Another Si efflux transporter, Lsi3, which is a homolog of Lsi2, has been identified in rice. An explanation for the high accumulation of Si in rice leaves and inflorescences was provided by Yamaji et al. (2015), who found localization of Lsi2, Lsi3, and Lsi6 at the nodes of the stem and demonstrated the involvement of these three transporters in intervascular Si transport (Fig. 1b). Such transport is needed for preferential distribution. Although the identification of Lsi1, Lsi2, Lsi3, and Lsi6 has defined the path of Si uptake at the root level and its subsequent transport to the aerial tissues, the transporter involved in Si loading in the xylem is not yet known (Fig. 1c). Similarly, the transporters for silica cells, in which very high amounts of Si accumulate, are not known. These represent large gaps in our knowledge that need to be filled to have a full understanding of the transport and deposition of Si in different plant tissues. Silicon deposition in different parts of the plant The accumulation and deposition of Si in plants have been extensively studied. Sites of silicification include the cell wall, wholly or partially filled cell lumens, the intercellular spaces of the roots and shoots, and in specialized silica cells. Silicification mainly occurs in sclerenchyma, fibers, storage tissues, epidermis, and vascular tissues. The pattern of Si deposition, the amount, and its role drastically vary among tissue types. Silicon deposition in the roots Silicon is known to be present in the roots of several members of the Poaceae, with the endodermis being most common site of silicification. The endodermal deposition of Si in rice roots was first reported by Parry and Soni (1972), and it is found to be localized as a ring in endodermal cell walls with no differences in the proximal and distal parts of tissue (Moore et al., 2011). The heaviest deposition is found in the inner tangential wall (ITW) and sometimes in the endodermis radial walls (Fig. 2). Silicon deposition is also found in the ITW of endodermis in the proximal end of the seminal root in barley, oats, and wheat (Bennett, 1982). In sorghum, two distinct types of silica deposition were found (Sangster and Parry (1976). One is similar to rice in that it is associated with the ITW of the endodermis. The other type, a phytolith (Metcalfe, 1960), is a discrete dome-shaped structure, normally attached to the ITW and protruding into the cell lumen. Silica deposition in sorghum starts simultaneously with the secondary wall formation of the endodermal cell, and these silica aggregates penetrate deep into the secondary walls (Sangster and Parry, 1976). Hodson (1986) examined adventitious roots of Phalaris canariensis and found that whilst silica was deposited in both the ITW and radial walls of the endodermis in underground roots, it was absent in the endodermis of adventitious aerial roots. Silica deposition is observed in intracellular spaces just outside the outer tangential wall of the root endodermis of Molinea ceruleae (Montgomery and Parry, 1979). In date palm (Phoenix dactylifera), Si deposition occurs in specialized cells known as stegmata cells, which are arranged in rows surrounding the sclerenchyma bundles in the roots (Bokor et al., 2019) (Fig. 3). As roots age, deposition can also occur in other parts such as the stele, sclerenchyma, and conductive tissues (Parry and Kelso, 1975). No expression of Lsi1 and Lsi2 is observed in the root hairs of rice, implying that the hairs have no role in Si uptake (Ma et al., 2006). Fig. 2. Open in new tabDownload slide Schematic representation of silicon deposition on the inner tangential wall of the root endodermis. Fig. 2. Open in new tabDownload slide Schematic representation of silicon deposition on the inner tangential wall of the root endodermis. Fig. 3. Open in new tabDownload slide Scanning electron microscopy–energy-dispersive X-ray imaging for silicon in Phoenix dactylifera root tissues. (A) Cross-section of adventitious root. The white arrowheads indicate deposition of silicon (Si) in stegmata cells (specialized cells in palm species) and the red arrowhead indicates the position of a fiber band. (B) Silicon deposition (violet colour) in multiple phytoliths in a cross-section of an adventitious root. (C) Longitudinal section through a fiber band of an adventitious root. The white arrowheads indicate Si deposition in disrupted stegmata cells and the red arrowhead indicates a fiber. (D) Silicon deposition (violet) in a longitudinal section of an adventitious root. The figure is reproduced from Bokor et al. (2019). Fig. 3. Open in new tabDownload slide Scanning electron microscopy–energy-dispersive X-ray imaging for silicon in Phoenix dactylifera root tissues. (A) Cross-section of adventitious root. The white arrowheads indicate deposition of silicon (Si) in stegmata cells (specialized cells in palm species) and the red arrowhead indicates the position of a fiber band. (B) Silicon deposition (violet colour) in multiple phytoliths in a cross-section of an adventitious root. (C) Longitudinal section through a fiber band of an adventitious root. The white arrowheads indicate Si deposition in disrupted stegmata cells and the red arrowhead indicates a fiber. (D) Silicon deposition (violet) in a longitudinal section of an adventitious root. The figure is reproduced from Bokor et al. (2019). Silicon loading into the xylem and unloading at aerial tissues The silicon content is generally higher in transpirational organs such as leaves and lower in absorptive organs such as roots, indicating that deposition is influenced by upward flow of the evapotranspiration stream. Following primary uptake in the roots, Si is transported to the shoots via the xylem. Uptake in the roots involves at least two major processes. First, Si from external solutions is transferred via transporters and passive diffusion into the cortical cells, and, second, it is released into the transpiration stream via xylem loading. Silicon accumulation in the aerial parts of the plant mainly depends on these two processes. Interestingly, rapid release of Si into the xylem is observed against a concentration gradient. In rice, Mitani et al. (2005) found that the concentration in the xylem sap reached 6.0 mM within 30 mins of exposure to only 0.5 mM Si in the growth medium (Fig. 4). Furthermore, the concentration increased to 18 mM in the next 8.5 h, indicating that xylem loading is a rapid process that is accomplished against a concentration gradient. However, the rate of xylem loading varies in plants and also depends on Si uptake and xylem loading processes (Mitani et al., 2005). In the aerial tissues, Si is unloaded from the xylem and transported to peripheral tissues in the leaves and inflorescences. In hemp (Cannabis sativa), Si is observed in various tissues, but in the bast fiber cells deposition occurs explicitly at the distal end of the cell wall (Guerriero et al., 2019). These Si-impregnated walls are thought to strengthen the plant and provide resistance to lodging (Fig. 5). Fig. 4. Open in new tabDownload slide Schematic representation of uptake, transport, and deposition of silicon. Plants take up silicon as orthosilicic acid from the soil where the concentration ranges between 0.2–0.6 mM. On uptake, orthosilicic acid is transported from the roots to the xylem, where its level transiently reaches up to 6–18 mM. The orthosilicic acid moves from the xylem to leaves where it undergoes auto-polymerization and is deposited as silica. Fig. 4. Open in new tabDownload slide Schematic representation of uptake, transport, and deposition of silicon. Plants take up silicon as orthosilicic acid from the soil where the concentration ranges between 0.2–0.6 mM. On uptake, orthosilicic acid is transported from the roots to the xylem, where its level transiently reaches up to 6–18 mM. The orthosilicic acid moves from the xylem to leaves where it undergoes auto-polymerization and is deposited as silica. Fig. 5. Open in new tabDownload slide High-resolution secondary ion mass spectrometry (Nano-SIMS) analysis of xylem cells and bast fibers of hemp plants with or without silicon (Si) treatment. The arrowheads in the treated samples indicate the deposition of Si at the distal end of the fibers. Increasing intensity of deposition is indicated by colour scale, which ranges from black to red. Reprinted from Guerriero et al. (2019) with permission from Elsevier. Fig. 5. Open in new tabDownload slide High-resolution secondary ion mass spectrometry (Nano-SIMS) analysis of xylem cells and bast fibers of hemp plants with or without silicon (Si) treatment. The arrowheads in the treated samples indicate the deposition of Si at the distal end of the fibers. Increasing intensity of deposition is indicated by colour scale, which ranges from black to red. Reprinted from Guerriero et al. (2019) with permission from Elsevier. Silicon deposition in leaves The leaves are the major site of Si deposition among the shoot tissues. In the rice leaf blade, deposition results in the formation of a 2.5-μm thick double layer of Si beneath the cuticle (Yoshida et al., 1962c; Ma and Yamaji, 2006). The formation of this layer is believed to occur by an active process in which Si particles are attracted by the ionic forces of the membrane surface. The layer continues to thicken by deposition of monomeric silicic acid from a supersaturated solution, which is the result of active metabolic processes rather than being caused by evaporation (Kaufman et al., 1981; Sangster et al., 2001). Silicification occurs in almost all the cells of the leaf blade and more than 90% of the total Si in the leaf is deposited in the epidermis, as a result of the transpirational stream (Yoshida et al., 1962b). Silicon is deposited in the epidermal cell walls, middle lamella, and intercellular spaces of the subepidermal tissues of rice (Kim et al., 2002). Variations in Si content occur in different leaves of the same plant, as well as at different positions within a leaf. The older leaves have more deposition and it gradually decreases from the apex to the base of the leaves (Sangster, 1970). In young leaves, Si is only detected in specialized silica cells and in bulliform or ‘motor cells’ whereas in senescing leaves it is present in virtually all cell types (Sangster et al., 2001), thus showing that Si deposition increases as the plant ages. Deposition differs between the adaxial and abaxial surfaces of the leaves. In bread grass (Brachiaria brizantha) Si accumulates mainly in the upper epidermis (de Melo et al., 2010), whilst in other species deposition is observed in both the adaxial and abaxial surfaces. For example, bamboo has accumulation in cork cells, bulliform cells, silica cells, long cells, and guard cells on both sides of the leaf (Motomura et al., 2002). In Pleioblastus chino the densest deposition is in the epidermis and the least dense is in the mesophyll and vascular bundles (Motomura et al., 2006). Deposition in silica cells Understanding the deposition of Si in the silica cells, as well as the evolution of such specialized cells, is of great importance in revealing its role in plants. Prat (1948) divided the epidermal silica-depositing cells of grasses into three subgroups, namely differentiated elements, fundamental elements, and bulliform elements. Differentiated elements include silica cells and exodermic components, for example micro- and macrohairs, trichomes, cork cells, and stomata. Fundamental elements include epidermal cells that are greatly elongated. Bulliform cells are present between the vascular bundles and on the adaxial surfaces of the leaf blade. In grasses, cell division in leaf epidermal tissues is restricted to the base of the growing leaf. The newly formed cells elongate and in doing so push the older cells upwards and out of the leaf sheath (Skinner and Nelson, 1995). In rice, silica cells are usually present in pairs that are tandemly repeated and run parallel to each other (Kaufman et al., 1985). Silica cells and cork cells located close by are thought to originate from a single mother cell in Avena (Kaufmian et al., 1969). After disintegration of the protoplasm, the lumens of the silica cells become filled with Si, eventually becoming a mass of solid, hydrated, amorphous silica that results in cell death (Kumar et al., 2017). Suberized cork cells, which remain unsilicified, may be involved in the metabolism of silica cells. The two are connected to each other and to neighboring cells by plasmodesmata (Lawton, 1980). Silica cells are the first cells that undergo silicification in the tissue, even before the tissue is exposed to the outer atmosphere (Kaufmian et al., 1969; Ma and Takahashi, 2002). Deposition in silica cells only occurs during leaf development (Sangster, 1970), and initially transpiration was thought to be responsible for silicification. However, Kumar et al. (2017) demonstrated that while transpiration is required to pull Si up and into the leaves, silicification in silica cells is an active process and does not depend upon evapotranspiration of water. The mechanism of silicification in silica cells involves the presence of specific material in the apoplast that enhances Si deposition. This material may include proteins, peptides, or sugars that are capable of polymerizing soluble silicic acid to solid silica. Polymerization and deposition of silicic acid at the cell wall possibly creates a concentration gradient within the leaf, thereby drawing silicic acid towards the silica cell (Gallagher et al., 2015). Silicon deposition in trichomes Deposition of Si is also found in the cells surrounding the base of trichome hairs and in the trichomes themselves. Abe (2019) studied deposition in the trichomes of six different species of Cucurbitaceae, namely cucumber, pumpkin, melon, watermelon, sponge gourd, and bottle gourd. They found that in watermelon leaves, Si is located in both the trichomes and the cells surrounding their bases, whilst in cucumber, pumpkin, and melon Si is deposited only in the cells at the bases and calcium is present in the hairs themselves. This combined deposition of calcium and Si gives rigidity to the trichomes, which in turn makes the plants more rigid. In contrast, SEM–energy-dispersive X-ray imaging of wheat has shown high levels of Si accumulation in trichomes but no differences in calcium compared with the surrounding cells (Fig. 6). An earlier study by Samuels et al. (1991) also reported deposition of Si in the bases of the trichomes and they concluded that these base cells differ from surrounding cells. Their results suggested that cells at the base of the trichomes transform in a way that supports the polymerization of Si. Silicon accumulated in trichomes is found to efficiently propagate far-infrared light inside the trichomes and leaves, which can be helpful in warming the tissues (Takeda et al., 2013). However, the precise roles of Si-filled trichomes are not well understood. Similarly, the transporters involved in the accumulation of Si in trichomes have not yet been identified. Fig. 6. Open in new tabDownload slide Energy-dispersive X-ray (EDX) assisted SEM images of the adaxial surface of leaves from 1-month-old wheat plants grown in soil with silicon supplementation. (A) Image showing bulliform cells. (B) Silicon (Si) deposition in bulliform cells, indicated by the white arrowheads. Si detected with EDX-SEM is highlighted by the red colour (C) Image showing calcium deposition in the leaf; however, no deposition was observed together with silicon. Calcium detected with EDX-SEM is highlighted by the orange colour. (D) EDX results showing the relative concentrations of various elements in the leaves. (E) Trichomes on the adaxial surface of the leaf (F) Deposition of Si in the trichomes, indicated by the white arrowheads. (G) Calcium deposition in the trichomes; no deposition was observed together with silicon. H) EDX results showing the relative concentrations of various elements in the trichomes. Fig. 6. Open in new tabDownload slide Energy-dispersive X-ray (EDX) assisted SEM images of the adaxial surface of leaves from 1-month-old wheat plants grown in soil with silicon supplementation. (A) Image showing bulliform cells. (B) Silicon (Si) deposition in bulliform cells, indicated by the white arrowheads. Si detected with EDX-SEM is highlighted by the red colour (C) Image showing calcium deposition in the leaf; however, no deposition was observed together with silicon. Calcium detected with EDX-SEM is highlighted by the orange colour. (D) EDX results showing the relative concentrations of various elements in the leaves. (E) Trichomes on the adaxial surface of the leaf (F) Deposition of Si in the trichomes, indicated by the white arrowheads. (G) Calcium deposition in the trichomes; no deposition was observed together with silicon. H) EDX results showing the relative concentrations of various elements in the trichomes. Silicon deposition that affects physiological plasticity under stress conditions Biotic stress Pathogens need to overcome the physical and chemical barriers presented by plant cells for successful infection (Ferreira et al., 2006), and Si is known to enhance these barriers, thereby helping to mitigate the biotic stress (Epstein, 1999). Onodera (1917) was the first to correlate Si content in rice with resistance to bacterial blight, caused by Pyriculria oryza. This triggered further such studies, most of which revolved around the development of puncture resistance due to Si deposition. Yoshida et al. (1962c) demonstrated the deposition of a layer of Si under the leaf cuticle, on which basis the hypothesis of Si forming a mechanical barrier was proposed. Plant pathogens secrete molecules known as effectors, which help them in colonization (Snelders et al., 2018). According to the apoplast obstruction hypothesis proposed by Coskun et al. (2019), Si deposition interferes with the host–pathogen specificity by forming barriers that prevent the movement of effectors within the plant. Silicon is thought to function in biotic stress resistance in two ways, first by forming mechanical barriers and second by acting as a modulator for host responses (Ma and Yamaji, 2006). Using a hydroponic nutrient solution, it has been shown that cucumber (Cucumis sativus) plants grown in a Si-containing medium have increased leaf rigidity, roughness, and dry weight, and enhanced resistance to downy mildew fungus (Adatia and Besford, 1986). Amending soil with Si fertilizers results in increased content in rice tissues, which in turn increases resistance to infection by brown spot and blast (Magnaporthe grisea) (Datnoff et al., 1992). Microscopic examination of the adaxial leaf surface indicates that increased Si content reduces appressorial penetration of the rice blast fungus into the epidermis (Hayasaka et al. 2008). Fungal infection is also prevented by Si-mediated formation of papillae and deposition of callose in wheat (Bélanger et al., 2003). Similarly, rose plants showed Si-induced resistance against powdery mildew due to the formation of papillae and fluorescent epidermal cells, which are interpreted as an indication of the hypersensitive response (Shetty et al., 2012). Deposition of callose and accumulation of H2O2 was also observed at the sites of infection. Leaf sheaths of rice plants supplied with Si show higher puncture resistance compared to plants without supplementation; for example, Si deposition in the sheaths results in increased resistance to infection by Rhizoctonia solani (Schurt et al., 2012). Silicon strengthens the cell wall by associating with the lignin component, and this strengthening helps in blast disease resistance in rice (Kim et al., 2002). A pioneer study by Chérif et al. (1992) demonstrated that Si leads to increased fungal resistance in cucumber as the result of deposition of phenolic material in the infected tissues and in primary and secondary cell walls, which damages the fungal hyphae. This study helped to reveal the role of Si other than as a purely mechanical barrier. Similarly, Si-induced deposition of a dense, amorphous material in rice leaves has been found to be associated with enhanced resistance against blast infection that can be attributed to both physical and cytochemical factors (Rodrigues et al., 2003). Cucumber plants grown with Si supplementation show antifungal activity against Pythium infection in the roots as a result of stimulation of chitinase activity followed by activation of peroxidases, polyphenol oxidases, and glycosidically bound phenolics (Chérif et al., 1994). Accumulation of phytoalexins induced by Si has been found to be responsible for increased resistance to powdery mildew in cucumber (Fawe et al., 1998). Wheat plants supplemented with Si show increased defence against Blumeria graminis f. sp. tritici as the result of releasing glycosylated phenolics, which affect the haustoria (Bélanger et al., 2003). Studies such as these changed the idea that resistance developed by Si is merely the result of formation of physical barriers. Another example is the restriction of blast infection in susceptible rice cultivars by Si supplementation, which results in increased accumulation of glucanase, peroxidase, and pathogenesis-related protein 1 (PR-1; Rodrigues et al., 2005). Sun et al. (2010) also found that Si induces host resistance mechanisms to blast in rice by enhancing the activities of catalase and peroxidase, and it also leads to lipid degradation and accumulation of H2O2. Rose plants supplemented with Si show resistance to powdery mildew as a result of a several-fold increase in content of the antifungal phenolic compounds chlorogenic acid and rutin (Shetty et al., 2011). Thus, Si plays an active role in mounting resistance against biotic stresses in plants. The role of Si in biotic stress has been further confirmed by studies of the rice mutant lsi1, which shows lower accumulation of Si in leaves compared to the wild-type. When inoculated with blast fungus, symptoms rapidly become apparent in the mutant, whereas the wild-type shows resistance (Ueno et al., 2007), which clearly suggests that the inability to take up Si is responsible for the susceptibility to the disease. Silicon accumulated in epidermal cells acts as a mechanical barrier that protects plants from herbivores, either directly or passively. Silicon treatment in sugarcane hardens the external rind and reduces penetration and stalk damage caused by larvae of the borer insect Eldana saccharina (Kvedaras and Keeping, 2007). The reduced ability to penetrate the stalk increases larval mortality as they are exposed to harsh environmental conditions for longer. Silicon is known to enhance physiological and biochemical processes in plants and as a consequence increase physiological resistance to herbivores. Despite being considered as a non-accumulator, even small amount of Si deposition in leaves of collard (Brassica oleracea) hardens them, thereby affecting the nutrition and performance of larvae of Plutella xylostella (Teixeira et al., 2017). The presence of Si at relatively high levels in various grass species impedes herbivory by Schistocerca gregaria and Spodoptera exempta (Massey et al., 2006), although no effect is seen on the phloem-feeding Sitobion avenae. Silicon supplementation in rice increases its concentration in the stem and this enhances resistance to the brown planthopper by reducing successful probing and phloem-sap ingestion, with the result that the pest shows reduced preference for settling on the plants (Yang et al., 2017). Mechanical barriers formed by Si in rice also help to resist the leaf folder Cnaphalocrocis medinalis, which is known to cause heavy damage (Han et al., 2015). Relatively high concentrations of Si in grasses cause wear of the mandibles within the first instar of S. exempta, resulting in reduced growth and nitrogen absorption by the larvae (Massey and Hartley, 2009). Similarly, increased Si concentration in Bromus catharticus increases the frequency of phytoliths, thereby helping resistance to the grasshopper Oxya grandis as a result of mandible wear (Mir et al., 2019). Abiotic stress Drought stress Silicon enhances tolerance to various abiotic stresses including drought, salinity, heavy metal toxicity, chilling, freezing, high-temperatures, high radiation, and waterlogging (Ma, 2004; Liang et al., 2015). Despite an abundance of information about the role of Si in alleviating abiotic stress, knowledge of its specific mechanisms of action is lacking. The deposition of a layer of Si below the cuticle reduces transpiration during water deficit and thus provides tolerance against drought stress (Ma et al., 2001). Drought conditions intensify the production of reactive oxygen species (ROS), which have deleterious effects on physiological and biochemical processes in plants (Cruz de Carvalho, 2008). Silicon enhances the activities of the ROS-scavenging enzymes catalase, superoxide dismutase, and glutathione reductase in wheat under drought stress whilst the oxidative stress is decreased (Gong et al., 2005). Silicon also aids in osmotic adjustment by enhancing the accumulation of proline and glycine betaine during drought conditions (Ahmad and Haddad, 2011), and it prevents membrane damage in sunflower by increasing the relative water content (Gunes et al., 2008). Silicon helps to maintain photosynthetic rates in cucumber under drought by reducing stomatal conductance, improving the water holding capacity, stabilizing the transpiration rate, and reducing chlorophyll decomposition (Ma et al., 2004). Silicon improves water uptake and hydraulic conductance during drought stress in sorghum (Sonobe et al., 2009). Silicon deposition in the leaves of Poa pratensis aids in maintaining them in an erect position, which improves photosynthesis through increased light penetration in the canopy and reduced transpiration (Saud et al., 2014), a change in morphology that can be important in alleviating drought stress. It has been demonstrated that sorghum plants treated with Si show enhanced accumulation of polyamines together with a decline in content of the ethylene precursor 1-aminocyclopropane-1-carboxylic acid, which results in delayed leaf senescence and an increase in the root to shoot ratio (Yin et al., 2014). Thus, Si enhances drought tolerance in plants by modulating a number of different physiological processes. Salinity stress Deposition of Si in the root endodermis and exodermis hampers apoplastic flow during salinity stress and reduces the uptake of sodium ions (Na+), thereby helping to avoid their deleterious effects (Yeo et al., 1999; Gong et al., 2006). Silicon supply also enhances the formation of the Casparian strip during salinity stress (Fleck et al., 2015), and the resulting blocking of apoplastic flow may be important in imparting tolerance. Treatment with Si during salinity stress improves dry matter production and chlorophyll content in wheat (Tuna et al., 2008), whilst in maize it enhances the sequestration of Na+ into leaf vacuoles, thereby preventing its excessive accumulation in the chloroplasts (Bosnic et al., 2018). Similar to drought stress, Si nutrition has been found to increase the activity of antioxidant enzymes such as catalase, ascorbate peroxidase, and guaiacol peroxidase during salinity stress in a cucumber cultivar displaying salinity tolerance (Khoshgoftarmanesh et al., 2014). Silicon supplementation confers salinity tolerance in mung bean plants by enhancing the accumulation of K+ and Ca+ ions, and the osmoprotectants proline and glycine betaine (Ahmad et al., 2019), whilst in pepper it increases the expression of adenylosuccinate synthase and E3 ubiquitin ligase, which aid in biomass accumulation, floral development, and senescence (Manivannan et al., 2016). Expression of Rubisco and oxygen-evolving enhancer proteins are also increased. Heavy metal stress Williams and Vlamis (1957) found that Si alleviates manganese toxicity in barley by causing it to be evenly distributed across the leaves, rather than accumulating to toxic levels in localized zones. In cucumber subjected to excess copper, Si supply increases both its deposition in the cell walls in the roots and its chelation by plastocyanin and acconitate in the leaves, thus helping to maintain cytosolic copper concentrations at relatively low levels (Bosnic et al., 2019). Silicon treatment decreases the transport of cadmium from roots to shoot in wheat, and decreases its uptake into leaf protoplasts (Greger et al., 2016). Heavy metal uptake in rice is also hampered by the blockage of the apoplastic pathway by substantial deposition of Si in the endodermal region, thus resulting in a reduction in their translocation (Shi et al., 2005). Silicon-mediated enhancement of the Casparian strip and suberin lamellae may also further reduce uptake of toxic heavy metals in plants. Silicon can also function to reduce the uptake of heavy metals from the soil. Heavy metal uptake is favoured by low pH and hence it can be reduced by increasing the pH (Bolan et al., 2003), which can be achieved by addition of Si, leading to immobilization of the metals (Bhat et al., 2019). Silicon also reduces the phytoavailibity of heavy metals by forming silicate complexes with them, resulting in changes in speciation and conversions from toxic to non-toxic forms (Emamverdian et al., 2018). Flavanoid-phenolics and organic acid chelation of heavy metals are other Si-induced heavy metal stress mitigation mechanisms. Secretion of an organic acid with chelating ability aids in detoxifying aluminum in higher plants (Ma, 2000). Silicon treatment enhances exudation of aluminum-chelating phenols from the root tips in an aluminum-tolerant maize variety (Kidd et al., 2001). Accumulation of heavy metals in plants triggers ROS production, which induces oxidative stress (Jalmi et al., 2018), and Si supply enhances the activities of antioxidant enzymes such as superoxide dismutase, catalase, and ascorbate peroxidase whilst reducing levels of malondialdehyde and H2O2 in rice and Brassica chinensis when they are stressed with zinc and cadmium, respectively (Song et al., 2009, 2011). Application of Si and nitric oxide to wheat decreases cadmium accumulation and reduces toxicity by up-regulation of the antioxidant defence system (Singh et al., 2020). Morphology, composition, and distribution of phytoliths The term phytolith was introduced by Ruprecht (1866) and is mainly used to indicate siliceous deposits. Many other terms are also used, including opaline silica, biogenic silica, and plant opal, and they essentially distinguish plant-derived silica from inorganic silica. Silica deposition occurs in different types of plant cells including silicified cells, micro-hairs, macro-hairs, long cells, and short cells. Phytoliths derived from different cells differ morphologically. The morphology of silica mainly depends on two factors, namely the type of cell in which it is formed and its precise location within the plant tissue; however, there are exceptions where the shape of the phytolith is not in accordance with the cell type. Phytoliths may be regularly shaped (i.e. spherical, globular, cylindrical, hexagonal, and cubical) or irregularly shaped (i.e. dumbbell, saddle, bowl, boat-shaped, bulliform, and polylobate). Phytoliths remain in the environment after plant death and the decay of organic matter, and their accumulation and persistence as microfossils can assist in archaeological studies that examine the agricultural origins of crop plants around the world (Ball et al., 2016). Morphological characterization is the only approach that can be efficiently used for such studies, as phytoliths exist as a distinct entity rather than as a fragment of any plant tissue. Phytoliths are not always exclusively composed of Si, and the chemical composition varies according to species, climatic conditions, pH, temperature, and soil composition (Kameník et al., 2013; Nawaz et al., 2019). The cellular environment in which development of the phytolith takes place may also have a profound effect on its composition; for example, phytoliths from the carbohydrate matrix in the cell wall differ from phytoliths deposited in the cell lumen in terms of their organic content (Hodson, 2016). The elemental composition of phytoliths can also be affected by the presence of pollutants in the soil, such as those caused by the mining industry (Buján, 2013). The elemental composition of phytoliths can vary between species grown in the same soil and environment (Hart, 2001). A survey by Carnelli et al. (2002) suggested that aluminum deposition in phytoliths was specific to woody species, but Hodson and Sangster (2002) determined that this might not be the case and instead it could be dependent on the pH of the substrate. The composition of phytoliths is also known to vary from organ to organ within the same plant (Hodson et al., 2008). Phytoliths are formed in almost all parts of plants, including the roots, shoots, stems, and leaves, and a single species can produce different types of phytoliths (Golokhvast et al., 2014). Hence, phytoliths can be a tool to identify plant species and to place them within the taxonomic hierarchy (Metcalfe, 1960). Epidermal phytoliths are mainly utilized for identification of grasses at the subfamily and tribe levels. Some types are common in all tribe members of a family, but some short-cell phytoliths are mainly specific for a particular tribe and can be used as indicators of individual subfamilies, tribes, and genera (Piperno and Pearsall, 1998). Different subfamilies of Poaceae have been characterized on the basis of the type of phytoliths present. For example, bilobate and saddle-shaped phytoliths are a characteristic feature for the Panicoideae and Chloridoideae subfamilies, respectively (Shakoor and Bhat, 2014). Maize contains phytoliths of both the festucoid and panicoid type and they have anatomical and morphological characters that are similar to the Andropogoneae and Panacea, so accordingly the species was placed in the subfamily Panicoideae (Prat, 1948). The evolutionary path of the specific chemical composition of phytoliths and its relevance to Si-derived benefits to plants are not yet understood. Biogeocycling of phytoliths Transport and recycling of Si within earth systems forms a biogeochemical cycle that is also known as the silica cycle. Bartoli (1983) was first to suggest a role for phytoliths in the biogeochemical cycle of Si. Preliminary studies suggested that the amount of soluble silica in water and soil is chiefly controlled by the dissolution of quartz (Rimstidt, 1997). Alexandre et al. (1997) studied the silica cycle in rainforests and found that phytoliths restore the silica in soil and are responsible for the soil silica dynamics. Meunier et al. (2001) also highlighted the importance of phytoliths as a Si reservoir in soil. Meunier et al. (2001) confirmed the importance of phytoliths in the silica cycle with the finding that a 15-cm deep layer of biogenic silica was formed by forest fires in bamboo stands. It has been observed that phytoliths are major contributors to soluble silica as compared to the organic matrix during plant decomposition (Fraysse et al., 2006). Polymerization of silicon Monosilicic acid is known to undergo polymerization once its concentration exceeds 2 mM (Ma and Yamaji, 2006). Its concentration in soil is much lower than this, and hence polymerization is restricted to within plants. In wheat and rice, the concentration of Si in the xylem sap can reach up to 8 mM and 6 mM, respectively. Although monosilicic acid is the major form, this excess concentration is transient and it is rapidly transported to the leaf tissue, where polymerization takes place (Casey et al., 2004; Mitani et al., 2005). Silicon polymerization consists of the condensation of [Si(OH)4] to Si-O-Si, which takes place in three steps: nuclei formation by monomer polymerization, growth of polymer particles, and formation of branched-chains by linkage of the particles, which leads to formation of a gel (Iler, 1979). Polymerization of silicon takes place by oxolation involving an SN2 nucleophilic substitution, with the rate increasing with increasing pH. The oxidation process proceeds with the concentrating of silanol units and H2O being released. Inanaga and Okasaka (1995) demonstrated that cell wall extracts from rice shoots contain Si-binding compounds. Cellulose assists in the formation of ordered octahedral Si aggregates, with unordered aggregates being formed in its absence (Perry and Lu, 1992). Protein-containing biomolecules are known to aid polymerization of monosilicic acid and have been studied in protein extracts from plants that accumulate silicon such as Phalaris, Equisetum, and Phragmites (Harrison, 1996; Perry and Keeling-Tucker, 2003). These extracts are found to be rich in proline, aspartic acid, and glutamic acid. Belton et al. (2004) studied the effects of 11 amino acids and glycine and lysine homopeptides on silicification, and found interactions between negatively charged Si and amino acids with positively charged side-chains. Increase in the length of the L-lysine homopeptide increased monomer formation and aggregation of Si. Coradin and Livage (2001) demonstrated that peptides and the amino acids serine, proline, lysine, and aspartic acid interact with silicate and aid in its polymerization to polysilicates, with the effects of the peptides being the more prominent. Differences in amino acid properties such as the isoelectric point and hydrophobicity affect the surface area of the polymers (Belton et al., 2004). In cucumber, proline-rich protein 1 (PRP1), a systemic acquired resistance protein with high numbers of lysine and arginine residues at the C-terminal end, is known to function in the polymerization of monosilicic acid at the site of fungal infection (Kauss et al., 2003), and the polymerization activity of PRP1 is attributable to its high density of positive charges. Kumar et al. (2020) identified the siliplant1 (Slp1) protein in sorghum, which has seven repeat units rich in proline, lysine, and glutamic acid, and functions to precipitate in silica cells. Slp1 interacts with silica by binding through the lysine amino group. Vesicles release Slp1 into the paramural space where it precipitates silicic acid. Other biomolecules have also been studied for their roles in biosilicification. For example, Belton et al. (2005a, 2005b) found that putrescine homologs positively affect condensation and aggregation, whilst analogs of spermine and spermidine positively affect aggregation. The effects of spermine and spermidine are related to their chain lengths. Significance of silicon deposition in cell walls in poor-accumulator species Numerous reports have suggested that supplementation of Si has beneficial effects even in poor-accumulator species such as tomato, canola, and Arabidopsis (Fauteux et al., 2006; Romero-Aranda et al., 2006; Hashemi et al., 2010; Khandekar and Leisner, 2011). This is puzzling since most studies over the last century have correlated Si-derived benefits with high accumulation. However, experiments performed with the rice lsi1 mutant may provide an explanation. Isa et al. (2010) used the mutant to examine the role of silica bodies in rice leaves and found that plants benefited even without forming Si-filled silica cells. The physiological role played by Si deposited in the cell wall seems to be different to that of Si deposited in the silica cells and silicon bodies. The growth enhancement observed in the lsi1 mutant in the absence of a high level of Si deposition might be the result of the formation of complexes of Si with polysaccharides in the cell wall. Evaluation of isolated cell walls from rice cell suspension cultures has shown crosslinking of Si with hemicellulose polysaccharides (He et al., 2015), which also suggests that very low amounts of silicon can change the cell wall physiology, leading to different observed effects in poor accumulators. This notion is further supported by a recent study in which a biophysical evaluation of single, isolated cells was carried out. Sheng et al. (2018) performed in situ micro-testing of ammonium (NH4+) ion fluxes in conjunction with atomic force microscopy of the cell wall and single-cell proteomics, and found that Si deposition clearly altered the cell wall structure, resulting in uptake of NH4+ that was twice that of cells cultured without Si. This indicates the role of Si in providing stability to the cell wall, which leads to optimized nutrient uptake and hence enhanced the growth and development with Si supplementation. A similar mechanism might be expected in species that do not accumulate Si in the aerial tissue because they lack a functional transporter, where Si interactions in the roots could still take place. Results from studies such as this are helpful in understanding the beneficial effects of Si in poor accumulators, but we need to add to the very limited information that is currently available. Summary Silicon is found in varying quantities in different tissues and it clearly imparts benefits to plant species. Among the numerous mechanisms explaining these benefits, the most commonly observed include alterations in physiological processes, modulation of stress responses, and biochemical interactions, such as those related to cell wall stability and the compositional diversity of phytoliths. A single mechanism or mode of action cannot account for the versatile roles that Si has been shown to perform (Fig. 7), and more studies at all scales of detail are therefore required if we are to gain a better understanding of how it benefits plants. We hope that the information provided in this review on the known molecular mechanisms, patterns of deposition, and roles in biotic and abiotic stress will be helpful in guiding future studies aimed at exploiting Si for agriculture applications. Fig. 7. Open in new tabDownload slide Biotic and abiotic stress tolerance mechanisms induced by silicon. Silicon is taken up by plant in the form of silicic acid present in soil, and is subsequently transported to other tissues where it is deposited in different cells and intracellular spaces. The uptake of silicon and its deposition result in numerous benefits to the plants, as indicated. Fig. 7. Open in new tabDownload slide Biotic and abiotic stress tolerance mechanisms induced by silicon. Silicon is taken up by plant in the form of silicic acid present in soil, and is subsequently transported to other tissues where it is deposited in different cells and intracellular spaces. The uptake of silicon and its deposition result in numerous benefits to the plants, as indicated. Acknowledgements RD and HS are thankful to the Government of India Department of Biotechnology for Ramalingaswami Fellowships, and the Science and Engineering Research Board (SERB) for financial support in the form of a grant (CRG/2019/006599). 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All rights reserved. For permissions, please email: [email protected] This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Hodson, Martin J; Evans, David E

doi: 10.1093/jxb/eraa024pmid: 31950161

Abstract Aluminium (Al) and silicon (Si) are abundant in soils, but their availability for plant uptake is limited by low solubility. However, Al toxicity is a major problem in naturally occurring acid soils and in soils affected by acidic precipitation. When, in 1995, we reviewed this topic for the Journal of Experimental Botany, it was clear that under certain circumstances soluble Si could ameliorate the toxic effects of Al, an effect mirrored in organisms beyond the plant kingdom. In the 25 years since our review, it has become evident that the amelioration phenomenon occurs in the root apoplast, with the formation of hydroxyaluminosilicates being part of the mechanism. A much better knowledge of the molecular basis for Si and Al uptake by plants and of Al toxicity mechanisms has been developed. However, relating this work to amelioration by Si is at an early stage. It is now clear that co-deposition of Al and Si in phytoliths is a fairly common phenomenon in the plant kingdom, and this may be important in detoxification of Al. Relatively little work on Al–Si interactions in field situations has been done in the last 25 years, and this is a key area for future development. Aluminium toxicity, co-deposition, hydroxyaluminosilicates, silicon, phytolith, X-ray microanalysis Introduction Silicon (Si) is, at 27.7%, the second most abundant element after oxygen in the Earth’s crust, whilst aluminium (Al) is the third most abundant at 8.2% (Exley, 1998). In many soils, aluminosilicates form a very significant component of the solid mineral matter. However, both Si and Al have relatively low solubility, with Si at around a maximum of 2000 µmol l−1, and the solubility of Al is highly affected by pH. The uptake of both is dependent on molecular form. Uptake of Si is as Si(OH)4 in aqueous solution (Raven, 1983; Epstein, 1994). It is understood to travel through the plant as Si(OH)4 in the transpiration flow and until deposited (usually as SiO2: Casey et al., 2003; Mitani et al., 2005; Ma and Yamaji, 2015; Coskun et al., 2019). Silica is deposited in the lumen, cell wall, and intercellular spaces, and the deposits are known as phytoliths. Al uptake is strongly pH dependent and is predominantly as Al3+ which is only available below pH 4.5 (Marion et al., 1976). There is a wide variation in the extent to which different species take up Si. It has been suggested that Si accumulators should be defined as those species whose Si content is >1.0% and where the Si/Ca ratio is >1.0 (Ma and Takahashi, 2002). Hodson et al. (2005) conducted a meta-analysis of data for Si content in leaves and non-woody material in 735 species. The horsetails and non-vascular plant species accumulated more Si than the ferns, gymnosperms, and angiosperms. Within the angiosperms, dicots tended to be low Si accumulators, while the commelinid monocot orders Poales (including the cereals and grasses) and Arecales accumulated substantially more Si than other monocots. In the heavy accumulators, some tissues and organs (e.g. the inflorescence bracts of the grasses) accumulate Si to very high levels. For most plants, Si is usually seen as a beneficial element, rather than essential, but it has important roles in defence against pathogens and grazers, and in decreasing abiotic stresses. As with Si, there is a wide range of uptake of Al. For most plants, Al is toxic, and it is an important factor in acid soils around the world, where it often seriously limits plant growth. In most plants, it is excluded from the root (for a recent review, see Bojorquez-Quninal et al., 2017). There are also a small number of Al accumulators, for which Al is beneficial (e.g. tea, Camellia sinensis; Matsumoto et al., 1976). The exclusion of Al from the root is most frequently achieved by the secretion of organic acids through transporters induced in the presence of Al. The Al accumulator species transport Al to the shoot, where it is chelated, often with organic acids (Ma et al., 1997a, b). We last reviewed this topic 25 years ago (Hodson and Evans, 1995), and it seems timely to bring it up to date now. Considerable progress has been made during this time. We have a much better knowledge of the solution chemistry underlying Al–Si interactions, and of the mechanism whereby Al toxicity is ameliorated in root systems. We now have a good understanding of the molecular basis of Al and Si transport, and of co-deposition within the plant. In this review we will consider first the state of knowledge of the field when we last reviewed it in 1995, and then the advances made in the study of Al–Si interactions and evidence for molecular mechanisms. Finally we will re-evaluate the significance of this for agriculture and understanding of plant responses to Al in soils. What was known in 1995 When we reviewed the topic in 1995, the amelioration of Al toxicity by Si had already been investigated in eight species, all in hydroponic solutions. Of these, half of the investigations (for rice, wheat, cotton, and pea) showed little or no amelioration, while the other half (for sorghum, barley, teosinte, and soybean) showed a marked effect. At the time it was unclear why this was the case, and there seemed to be both plant species effects and solution effects. It was highly uncertain whether amelioration effects were to be found entirely in bulk solutions, within the plants, or a combination of the two. In 1995 it was very well known that Al was more available for plant uptake, and more toxic, at low pH. It was not then clear, however, that pH also affected the interactions between Al and Si in solution, impacting plant growth. At that time, Baylis et al. (1994) had only recently published their work on amelioration of Al toxicity by Si in soybean. They showed that higher concentrations of Si were required for amelioration at lower pH, and concluded that, ‘These results support the hypothesis that the pH-dependent affinity of Si for Al in dilute solutions, and the consequent formation of sub-colloidal inert hydroxyaluminosilicate species, is the basis for the alleviation of Al toxicity by Si.’ Although we were aware that pH had an effect on hydroxyaluminosilicate (HAS) formation, it was only after this work (Baylis et al., 1994) that plant scientists began to investigate the phenomenon. Birchall, Exley, and their colleagues had already investigated the chemistry of Al–Si interactions in solution, and particularly the formation of HAS (Exley and Birchall, 1992, 1993). HAS formation involved the inhibition of aluminium hydroxide nucleation. This resulted from silicic acid exchanging with hydroxylated Al at growth sites on the lattices of aluminium hydroxide. The effect depended on solution pH and the concentration of silicic acid. It was already clear that there were several distinct groups of plants with respect to Al–Si interactions (Hodson and Evans, 1995). The grasses and cereals transported large amounts of Si to the shoot (see ‘Si transport mechanisms’ below), but mostly prevented Al transport beyond the root. Thus, most of the interest in Al–Si interactions in these species was focused on the roots. Most of the dicots excluded both elements from the shoots, and there was already some evidence (Baylis et al., 1994) that Si could ameliorate Al toxicity under some conditions in these plants. Again this pointed to the roots as the main site of any amelioration effect. The phenomenon of Al hyperaccumulation in the shoot was already well recognized in some dicots and ferns, but it appeared that these plants largely excluded Si from the shoot, and that Al tolerance in the shoot mainly involved chelation with organic acids. It seemed that the conifers could transport moderate amounts of both Al and Si to the shoot. The first microanalytical study that demonstrated Al–Si co-deposition in conifer needles was that of Godde et al. (1988). They investigated spruce that was suffering from dieback in Germany, and found that Al and Si were co-located in the transfusion tissue of the needles. By 1995 it was already evident that there was no relationship between the uptake of Si and tolerance to Al, and that a variety of mechanisms and processes were involved. After 1995 we chose to work on Al–Si interactions in the cereals and the conifers. Both are economically important groups, and they appeared to cope with Al toxicity, and to use Si in amelioration of the toxicity, in somewhat different ways. Other scientists joined us in these investigations, and yet others continued the work on Al hyperaccumulators. Progress since 1995 has focused on three main areas: the amelioration of Al toxicity by Si in root systems; Al and Si transport at the cellular level; and the co-deposition of Al and Si in phytoliths. The amelioration of Al toxicity by Si in root systems Since our 1995 paper there have been many reports of Al toxicity being ameliorated by Si (e.g. Hammond et al., 1995; Corrales et al., 1997; Hara et al., 1999; Liang et al., 2001; Singh et al., 2011; Pontigo et al., 2017) and we will not catalogue these in detail here. Rather we will highlight work that elucidates the mechanism(s) behind the amelioration effect. The crucial effect of pH It was soon found that even relatively small changes in solution pH can have a very marked effect on amelioration of Al toxicity by Si. Cocker et al. (1997), working on wheat, observed amelioration at pH 4.6, but not at 4.2 (with 100 µmol l−1 Al and 2000 µmol l−1 Si). Similarly, Ryder et al. (2003) found that Si (1000 µmol l−1 or 2000 µmol l−1) did not ameliorate toxic effects of Al (with 100 µmol l−1) on root growth of spruce at pH 4.00, 4.25, and 4.50, while apparently complete amelioration was found at pH 4.75 and 5.00. It became obvious that the early papers that reported no amelioration by Si were almost certainly using a solution pH that was too low for the effect to be observed. The importance of pH change of culture solutions during plant growth experiments was also recognized, with later work mostly being carried out in highly controlled flowing culture systems. Solution chemistry It is now recognized that the chemistry of the interaction between Al and Si in solution is complex, and more complex than we had thought 25 years ago. Computer modelling of speciation has helped, and the constants used have been continually improved. Ryder et al. (2003) used an equilibrium speciation model (EQ3NR) to predict the behaviour of Al and Si in growth solutions over the pH range 4.0–5.0. As might be expected, addition of Si (1000 µmol l−1) to Al (100 µmol l−1) solutions caused a decrease in Al3+. At pH 4.00, Al3+ decreased from 92.4% to 83.3%, and at pH 5.00 the fall was from 54.6% to 17.7%. The decline in Al3+ was considered to be due to the formation of HAS species. One of the complexities in using such equilibrium speciation models is that they assume that solutions have come to equilibrium, and this may not always be the case. The most detailed recent explanation of the formation of HAS was provided by Beardmore et al. (2016). The authors used computational chemistry to recognize and describe the significant reaction steps. Density-functional theory combined with solvent continuum models were used to confirm that the reactants were an aluminium hydroxide dimer and silicic acid. The reaction products were two HAS species, HASA and HASB. According to Beardmore et al. (2016), HASA dominates in solutions where the concentration of Si(OH)4 is less than or equal to the total Al. However, HASB is dominant when the concentration of Si(OH)4 is at least double that of Al. This means that we would expect HASB to be the dominant species in almost all of the experiments conducted on plants, as in nearly all cases Si exceeds Al by a factor of 10. HASA and HASB are then precursors of much more insoluble HAS species. A solution effect or in planta? It is now certain that at least some of the ameliorative effects of Si on Al toxicity occur in solution, with the formation of non-toxic HAS species. However, evidence has accumulated since 1995 that part of the amelioration occurs within the plant (‘in planta’). Later we will consider cases where Al is co-deposited with Si as a solid, decreasing the availability of Al within the plant. Here we will only look at situations where there is no obvious co-deposition, mostly concerning experimental work on root systems. Corrales et al. (1997), working on maize, showed that pre-treatment of roots with 1000 µmol l−1 Si markedly increased growth of plants that were then exposed to Al (using a range of concentrations up to 100 µmol l−1). Obviously, this effect could not be due to HAS formation in the bulk solutions as Al and Si were not present in the solutions at the same time. The authors showed that the presence of Si decreased the uptake of Al into the root, and suggested that this was the reason for the better growth in the plants pre-treated with Si. A second example where solution effects could be ruled out was provided by the work of Cocker et al. (1998a) who studied the Al-ultrasensitive wheat cultivar, Scout 66. A low concentration of 1.5 µmol l−1 Al inhibited the root growth, but amelioration was observed with only 5 µmol l−1 Si. These low concentrations of Al and Si are well below those where the formation of HAS species has been observed. Again this is highly suggestive that in this case in planta effects are involved. Cocker et al. (1998a) also worked on the Atlas 66 cultivar of wheat, where Al-induced malate exudation from roots was already known to be involved in Al tolerance. It was found that 2000 µmol l−1 Si ameliorated the toxicity of Al (100 µmol l−1) in this cultivar at pH 4.6. Under these conditions, malate exudation was very similar to that found in plants treated with just 100 µmol l−1 Al. However, addition of citrate, a well-known chelator of Al, decreased malate exudation at 5–40 µmol l−1 and completely inhibited it at 100 µmol l−1. These findings strongly indicate that the formation of HAS in the bulk solution was not the reason for the amelioration of Al toxicity, and that this must take place in planta. We were then able to put together all of the available data and to form a hypothesis that the root cell walls were the main sites of Al detoxification, and that HAS species or aluminosilicates formed there as the primary mechanism (Cocker et al., 1998b). A number of factors were believed to be responsible: high apoplastic pH; organic substances such as malate; and local concentrations of Al and Si on or within the cell wall. The most detailed investigation of the amelioration phenomenon published so far was that of Wang et al. (2004), and this provided further confirmation of the role of HAS formation in the root cell wall. They worked on an Al-sensitive maize cultivar, Lixis, using short exposure times (25 µmol l−1 Al with or without 1400 µmol l−1 Si). Monomeric Al concentrations were not reduced by Si, implying that there were no interactions in the bulk solutions, and yet significant amelioration was observed. Greater than 85% of Al was bound within the cell wall in 1 cm root apices. Al content of root apices, which were treated with Si, were no different from those with no treatment for the apoplastic sap, the symplastic sap, and the cell wall. Confirming the results of Cocker et al. (1998a), Si had no effect on exudation of organic acids and phenols that were induced by Al treatment. The major effect of Al treatment was to increase accumulation of Si in the cell wall fraction. This decreased mobility of Al in the apoplast, and appeared to be the reason for the amelioration effects. Additional evidence that HAS formation in the root cell wall was involved in the amelioration of Al toxicity was provided by Kopittke et al. (2017). These authors used low energy X-ray fluorescence (LEXRF) to examine Al and Si distribution in the sorghum root apex. They found that Al and Si were co-localized in the mucigel and outer apoplast of the root. It seems clear that the hypothesis that we (Cocker et al., 1998b) proposed has been confirmed by further work over nearly 20 years. In planta co-deposition of Al with Si and the formation of HAS within the root apoplast does appear to be the mechanism behind amelioration. Very recently, Coskun et al. (2019) proposed their ‘apoplastic obstruction hypothesis’, suggesting that Si in the apoplast is involved in the reduction of a wide range of abiotic and biotic stresses. Although they did not consider the topic in detail, it does appear that work on the amelioration of Al toxicity by Si in plants fits reasonably well with their ideas. Al and Si transport at the cellular level One of the greatest advances since our previous review has been in the identification and characterization of transporters for Si and Al at the molecular level. The presence of membrane proteins (usually modified water channels, aquaporins) facilitates uptake and transport. Exley and Guerriero (2019) suggest these are channels, rather than active transporters (though there is a suggestion that at least one Si transporter is a proton-linked antiporter; Ma et al., 2007; Coskun et al., 2019; see below). They therefore suggest that directionality of transport is due to the concentration gradients of soluble Si(OH)4 created by water flow and the removal of Si from solution, usually as SiO2. As different species take up Si to different extents (see above), it would be expected that uptake would correlate with the presence and activity of transporters. Bélanger et al. (2016) demonstrated that Si uptake was dependent on root stock rather than transpiration rates in a soybean grafting experiment, suggesting that the characteristics of root cells were more important than the transpiration rate. Exley (2015) also noted the significance of guttation—the active exudation of aqueous solutes—which may include Si(OH)4, from the plant surface in some circumstances, though this may be considered as a variant of passive movement in the water flow. Si transport mechanisms Si transporters to date have been characterized as influx and efflux carriers suggested to provide directional transport to regions of high accumulation. First to be identified (Ma et al., 2006) was an Si influx protein, named Lsi1 (for low silicon protein 1, due to its absence in a low Si-accumulating rice), an aquaporin. Lsi1 is localized to the distal side of endodermal and exodermal cells of the root. This was followed by identification of an Si efflux protein, Lsi2 (Ma et al., 2007), located in the root exodermis and endodermis, but, in contrast to Lsi1, on the proximal side of these two layers. Ma et al. (2007) therefore suggested that movement of Si in rice is through a transport pathway traversing the endodermis and exodermis, to the xylem and the shoot. In silico modelling reveals that their location at the endodermis and exodermis coupled with the presence of the impermeable Casparian strip is required for effective movement of Si to the shoot (Skurai et al., 2015). Lsi1 is a member of the membrane intrinsic [MIP (major intrinsic protein) and NIPIII (Nodulin 26-like intrinsic protein 3-1)] family of aquaporins. Specificity for Si is achieved by an hourglass-like configuration of the protein, with six transmembrane domains and two half helices projecting into a central pore. This constriction, with two NPA (asparagine, proline, and alanine) domains is close to a second constriction, the selectivity filter, which determines specificity for Si. The key amino acids for Si are glycine–serine–glycine–arginine (GSGR; Hove and Bhave, 2011). The spacing of the two NPA domains is important for transport, as this renders the pore permeable to Si(OH)4; high Si accumulators have a 108 amino acid separation (Deshmukh et al., 2015). Lsi1 also transports methylated arsenic species (Li et al., 2009). Lsi2 resembles a bacterial arsenite transporter and is thought to be an Si(OH)4/H+ antiporter, though direct evidence for this is lacking (Ma et al., 2007; Coskun et al., 2019). It belongs to the arsenite–antimonite (ArsB) efflux (TC 2.A.45) family. Si transport activity was demonstrated using Xenopus oocytes pre-loaded with Si, which showed Si efflux activity (Ma et al., 2007). Pommerrenig et al. (2020) have recently provided evidence that the NIP family of transporters originated from bacterial arsenic efflux channels through a horizontal gene transfer followed by evolution to higher plant Si and boron transporters. The tissue location of Si transport proteins has been shown to be important for movement to different areas of the shoot and contributes to areas of high accumulation (Yamaji et al., 2015). Rice accumulates >10% Si (dry weight basis) in the husk, which protects the grain from biotic and abiotic stress. In addition to Lsi1 and Lsi2, Lsi6 is a plasma membrane-localized Si transporter expressed in roots and shoots, and is located at nodes involved in Si distribution from xylem to leaf tissue (Yamaji et al., 2008). In these and subsequent experiments, Yamaji et al. (2015) observed that Lsi2 is polarly localized under bundle sheath cells around enlarged vascular bundles (VBs). Lsi6 is localized in a xylem transfer cell layer adjacent to this; Lsi3 is in the parenchyma between VBs. Knockout of these two reduces Si in panicles and increases it in the flag leaf. Yamaji et al. (2015) modelled this to show that an apoplastic barrier at the bundle sheath and development of the enlarged VBs in the node are needed for Si hyperaccumulation in the husk. While rice Si transport is the best studied system, Si transporters have been identified in a number of other species based on sequence homology and, in some instances, function. These are summarized in Table 1. They include Si influx transporters in wheat, Triticum aestivum, TaLsi1 (Montpetit et al., 2012); barley, HvLsi1 (Chiba et al., 2009; Mitani et al., 2009), and Si influx and efflux channels in cucumber, CmLsi1 and CmLsi3 (Mitani et al., 2011); maize, NIP2-1 and ZmLsi2 (Mitani et al., 2009), perennial ryegrass, Lolium perenne, Lsi1 and Lsi2; (Pontigo et al., 2017), and a barley homologue of Lsi6 involved in Si distribution, HvSi6 (Yamaji et al., 2012). Channels predicted to allow Si movement have also been identified in castor bean, wild soybean, mung bean, date palm, red clover, pearl millet, sorghum, grape vine, and barrel medic (see Supplementary Table S1 at JXB online). While maize and barley show similar Si transporters to rice, the work of Mitani et al. (2009) suggests a different transport mechanism, as the Lsi2 homologue is located at the endodermis but immunostaining does not reveal the polarity exhibited by Lsi1 and Lsi2 in rice. The importance of Lsi2 in transport, however, is demonstrated by a correlation between expression levels and Si uptake and its ability to recover uptake in a low Si rice mutant. Studies on the Si influx transporter in two cultivars of pumpkin (Cucurbita sativus), one a high Si accumulator, the other a low accumulator, reveal that a change of one amino acid—Pro242 to lysine—results in loss of Si accumulation and relocation of the transporter from the plasma membrane to the endoplasmic reticulum (Mitani et al., 2011). Sun et al. (2019), in a study of tomato root Si transporters, demonstrate that while there are two Si transporter homologues present (SILsi1, a homologue of the rice LSi1 influx transporter; and SILsi2, a homologue of the rice LSi2 efflux transporter), only SILsi1 is active. They suggest that the absence of active SILsi2 explains the low levels of Si accumulation in this species. Table 1. Plant Si transporters in plant membranes with supporting references (Uniprot Consortium, 2019) Entry . Protein names . Gene names . Organism . Length . Reference . Q6Z2T3 Aquaporin NIP2-1 (Low silicon protein 1) (NOD26-like intrinsic protein 2-1) (OsNIP2;1) (Silicon influx transporter LSI1) NIP2-1 LSI1 SIIT1 Os02g0745100 LOC_ Os02g51110 OJ1118_ G04.16 OJ1734_E02.43 OsJ_008085 Oryza sativa subsp. japonica (rice) 298 Ma et al. (2006) Q10SY9 Silicon efflux transporter LSI2 (Low silicon protein 2) LSI2 SIET1 Os03g0107300 LOC_Os03g01700 OJ1384D03.1 OsJ_09099 Oryza sativa subsp. japonica (rice) 472 Ma et al. (2007) Q67WJ8 Aquaporin NIP2-2 (Low silicon protein 6) (NOD26-like intrinsic protein 2-2) (OsNIP2;2) NIP2-2 LSI6 Os06g0228200 LOC_Os06g12310 OsJ_019836 P0425F05.28-1 Oryza sativa subsp. japonica (rice) 298 Yamaji et al. (2008) Q9AV23 Silicon efflux transporter LSI3 (Low silicon protein 3) LSI3 Os10g0547500 LOC_Os10g39980 OSJNBa0001O14.19 Oryza sativa subsp. japonica (rice) 485 Yamaji et al. (2015) G0WXH5 Silicon transporter protein TaLsi1 Triticum aestivum (wheat) 295 Montpetit et al. (2012) B9X078 NIP2;1 (NOD26-like intrinsic protein) (Silicon transporter) HvLsi1 HvNIP2;2 NIP2;1 Hordeum vulgare (barley) 295 Chiba et al. (2009) Q19KC1 Aquaporin NIP2-1 (NOD26-like intrinsic protein 2-1) (ZmNIP2-1) (ZmNIP2;1) NIP2-1 LSI1 Zea mays (maize) 295 Mitani et al. (2009) C6KYS1 Silicon transporter HvLsi6 Hordeum vulgare (barley) 300 Yamaji et al. (2012) F1SX51 Silicon transporter 1 CmLsi1(B-) Cucurbita moschata 288 Mitani et al. (2011) C7G3B4 Silicon transporter ZmLsi2 100502546 ZEAMMB73_ Zm00001d027305 Zea mays (maize) 477 Mitani et al. (2009) F1SX50 Silicon transporter 1 CmLsi1(B+) Cucurbita moschata 288 Mitani et al. (2011) I4IY30 Aquaporin silicon transporter EaNIP3,1 Equisetum arvense (field horsetail) (common horsetail) 248 Grégoire et al. (2012) I4IY32 Aquaporin silicon transporter EaNIP3,3 Equisetum arvense (field horsetail) (common horsetail) 259 Grégoire et al. (2012) I4IY33 Aquaporin silicon transporter EaNIP3,4a Equisetum arvense (field horsetail) (common horsetail) 260 Grégoire et al. (2012) XP_008802606 Aquaporin silicon transporter PdNIP2-1 Phoenix dactylifera (date palm) 290 Bokor et al. (2019) XP_004240725.1 Si influx transporter (active) SILs2-L1 Solanum esculentum (tomato) 528 Sun et al. (2019) XP_010317628.1 Si efflux transporter (inactive) SILs2_L2 Solanum esculentum (tomato) 516 Sun et al. (2019) Entry . Protein names . Gene names . Organism . Length . Reference . Q6Z2T3 Aquaporin NIP2-1 (Low silicon protein 1) (NOD26-like intrinsic protein 2-1) (OsNIP2;1) (Silicon influx transporter LSI1) NIP2-1 LSI1 SIIT1 Os02g0745100 LOC_ Os02g51110 OJ1118_ G04.16 OJ1734_E02.43 OsJ_008085 Oryza sativa subsp. japonica (rice) 298 Ma et al. (2006) Q10SY9 Silicon efflux transporter LSI2 (Low silicon protein 2) LSI2 SIET1 Os03g0107300 LOC_Os03g01700 OJ1384D03.1 OsJ_09099 Oryza sativa subsp. japonica (rice) 472 Ma et al. (2007) Q67WJ8 Aquaporin NIP2-2 (Low silicon protein 6) (NOD26-like intrinsic protein 2-2) (OsNIP2;2) NIP2-2 LSI6 Os06g0228200 LOC_Os06g12310 OsJ_019836 P0425F05.28-1 Oryza sativa subsp. japonica (rice) 298 Yamaji et al. (2008) Q9AV23 Silicon efflux transporter LSI3 (Low silicon protein 3) LSI3 Os10g0547500 LOC_Os10g39980 OSJNBa0001O14.19 Oryza sativa subsp. japonica (rice) 485 Yamaji et al. (2015) G0WXH5 Silicon transporter protein TaLsi1 Triticum aestivum (wheat) 295 Montpetit et al. (2012) B9X078 NIP2;1 (NOD26-like intrinsic protein) (Silicon transporter) HvLsi1 HvNIP2;2 NIP2;1 Hordeum vulgare (barley) 295 Chiba et al. (2009) Q19KC1 Aquaporin NIP2-1 (NOD26-like intrinsic protein 2-1) (ZmNIP2-1) (ZmNIP2;1) NIP2-1 LSI1 Zea mays (maize) 295 Mitani et al. (2009) C6KYS1 Silicon transporter HvLsi6 Hordeum vulgare (barley) 300 Yamaji et al. (2012) F1SX51 Silicon transporter 1 CmLsi1(B-) Cucurbita moschata 288 Mitani et al. (2011) C7G3B4 Silicon transporter ZmLsi2 100502546 ZEAMMB73_ Zm00001d027305 Zea mays (maize) 477 Mitani et al. (2009) F1SX50 Silicon transporter 1 CmLsi1(B+) Cucurbita moschata 288 Mitani et al. (2011) I4IY30 Aquaporin silicon transporter EaNIP3,1 Equisetum arvense (field horsetail) (common horsetail) 248 Grégoire et al. (2012) I4IY32 Aquaporin silicon transporter EaNIP3,3 Equisetum arvense (field horsetail) (common horsetail) 259 Grégoire et al. (2012) I4IY33 Aquaporin silicon transporter EaNIP3,4a Equisetum arvense (field horsetail) (common horsetail) 260 Grégoire et al. (2012) XP_008802606 Aquaporin silicon transporter PdNIP2-1 Phoenix dactylifera (date palm) 290 Bokor et al. (2019) XP_004240725.1 Si influx transporter (active) SILs2-L1 Solanum esculentum (tomato) 528 Sun et al. (2019) XP_010317628.1 Si efflux transporter (inactive) SILs2_L2 Solanum esculentum (tomato) 516 Sun et al. (2019) See Supplementary Table S1 for a full list including putative Si transporters identified from genome database annotation. Open in new tab Table 1. Plant Si transporters in plant membranes with supporting references (Uniprot Consortium, 2019) Entry . Protein names . Gene names . Organism . Length . Reference . Q6Z2T3 Aquaporin NIP2-1 (Low silicon protein 1) (NOD26-like intrinsic protein 2-1) (OsNIP2;1) (Silicon influx transporter LSI1) NIP2-1 LSI1 SIIT1 Os02g0745100 LOC_ Os02g51110 OJ1118_ G04.16 OJ1734_E02.43 OsJ_008085 Oryza sativa subsp. japonica (rice) 298 Ma et al. (2006) Q10SY9 Silicon efflux transporter LSI2 (Low silicon protein 2) LSI2 SIET1 Os03g0107300 LOC_Os03g01700 OJ1384D03.1 OsJ_09099 Oryza sativa subsp. japonica (rice) 472 Ma et al. (2007) Q67WJ8 Aquaporin NIP2-2 (Low silicon protein 6) (NOD26-like intrinsic protein 2-2) (OsNIP2;2) NIP2-2 LSI6 Os06g0228200 LOC_Os06g12310 OsJ_019836 P0425F05.28-1 Oryza sativa subsp. japonica (rice) 298 Yamaji et al. (2008) Q9AV23 Silicon efflux transporter LSI3 (Low silicon protein 3) LSI3 Os10g0547500 LOC_Os10g39980 OSJNBa0001O14.19 Oryza sativa subsp. japonica (rice) 485 Yamaji et al. (2015) G0WXH5 Silicon transporter protein TaLsi1 Triticum aestivum (wheat) 295 Montpetit et al. (2012) B9X078 NIP2;1 (NOD26-like intrinsic protein) (Silicon transporter) HvLsi1 HvNIP2;2 NIP2;1 Hordeum vulgare (barley) 295 Chiba et al. (2009) Q19KC1 Aquaporin NIP2-1 (NOD26-like intrinsic protein 2-1) (ZmNIP2-1) (ZmNIP2;1) NIP2-1 LSI1 Zea mays (maize) 295 Mitani et al. (2009) C6KYS1 Silicon transporter HvLsi6 Hordeum vulgare (barley) 300 Yamaji et al. (2012) F1SX51 Silicon transporter 1 CmLsi1(B-) Cucurbita moschata 288 Mitani et al. (2011) C7G3B4 Silicon transporter ZmLsi2 100502546 ZEAMMB73_ Zm00001d027305 Zea mays (maize) 477 Mitani et al. (2009) F1SX50 Silicon transporter 1 CmLsi1(B+) Cucurbita moschata 288 Mitani et al. (2011) I4IY30 Aquaporin silicon transporter EaNIP3,1 Equisetum arvense (field horsetail) (common horsetail) 248 Grégoire et al. (2012) I4IY32 Aquaporin silicon transporter EaNIP3,3 Equisetum arvense (field horsetail) (common horsetail) 259 Grégoire et al. (2012) I4IY33 Aquaporin silicon transporter EaNIP3,4a Equisetum arvense (field horsetail) (common horsetail) 260 Grégoire et al. (2012) XP_008802606 Aquaporin silicon transporter PdNIP2-1 Phoenix dactylifera (date palm) 290 Bokor et al. (2019) XP_004240725.1 Si influx transporter (active) SILs2-L1 Solanum esculentum (tomato) 528 Sun et al. (2019) XP_010317628.1 Si efflux transporter (inactive) SILs2_L2 Solanum esculentum (tomato) 516 Sun et al. (2019) Entry . Protein names . Gene names . Organism . Length . Reference . Q6Z2T3 Aquaporin NIP2-1 (Low silicon protein 1) (NOD26-like intrinsic protein 2-1) (OsNIP2;1) (Silicon influx transporter LSI1) NIP2-1 LSI1 SIIT1 Os02g0745100 LOC_ Os02g51110 OJ1118_ G04.16 OJ1734_E02.43 OsJ_008085 Oryza sativa subsp. japonica (rice) 298 Ma et al. (2006) Q10SY9 Silicon efflux transporter LSI2 (Low silicon protein 2) LSI2 SIET1 Os03g0107300 LOC_Os03g01700 OJ1384D03.1 OsJ_09099 Oryza sativa subsp. japonica (rice) 472 Ma et al. (2007) Q67WJ8 Aquaporin NIP2-2 (Low silicon protein 6) (NOD26-like intrinsic protein 2-2) (OsNIP2;2) NIP2-2 LSI6 Os06g0228200 LOC_Os06g12310 OsJ_019836 P0425F05.28-1 Oryza sativa subsp. japonica (rice) 298 Yamaji et al. (2008) Q9AV23 Silicon efflux transporter LSI3 (Low silicon protein 3) LSI3 Os10g0547500 LOC_Os10g39980 OSJNBa0001O14.19 Oryza sativa subsp. japonica (rice) 485 Yamaji et al. (2015) G0WXH5 Silicon transporter protein TaLsi1 Triticum aestivum (wheat) 295 Montpetit et al. (2012) B9X078 NIP2;1 (NOD26-like intrinsic protein) (Silicon transporter) HvLsi1 HvNIP2;2 NIP2;1 Hordeum vulgare (barley) 295 Chiba et al. (2009) Q19KC1 Aquaporin NIP2-1 (NOD26-like intrinsic protein 2-1) (ZmNIP2-1) (ZmNIP2;1) NIP2-1 LSI1 Zea mays (maize) 295 Mitani et al. (2009) C6KYS1 Silicon transporter HvLsi6 Hordeum vulgare (barley) 300 Yamaji et al. (2012) F1SX51 Silicon transporter 1 CmLsi1(B-) Cucurbita moschata 288 Mitani et al. (2011) C7G3B4 Silicon transporter ZmLsi2 100502546 ZEAMMB73_ Zm00001d027305 Zea mays (maize) 477 Mitani et al. (2009) F1SX50 Silicon transporter 1 CmLsi1(B+) Cucurbita moschata 288 Mitani et al. (2011) I4IY30 Aquaporin silicon transporter EaNIP3,1 Equisetum arvense (field horsetail) (common horsetail) 248 Grégoire et al. (2012) I4IY32 Aquaporin silicon transporter EaNIP3,3 Equisetum arvense (field horsetail) (common horsetail) 259 Grégoire et al. (2012) I4IY33 Aquaporin silicon transporter EaNIP3,4a Equisetum arvense (field horsetail) (common horsetail) 260 Grégoire et al. (2012) XP_008802606 Aquaporin silicon transporter PdNIP2-1 Phoenix dactylifera (date palm) 290 Bokor et al. (2019) XP_004240725.1 Si influx transporter (active) SILs2-L1 Solanum esculentum (tomato) 528 Sun et al. (2019) XP_010317628.1 Si efflux transporter (inactive) SILs2_L2 Solanum esculentum (tomato) 516 Sun et al. (2019) See Supplementary Table S1 for a full list including putative Si transporters identified from genome database annotation. Open in new tab Horsetails are primitive plants with a very high Si content (Hodson et al., 2005), that require Si to complete their life cycle (Chen and Lewin, 1969; Miyake and Takahashi, 1976) and provide evidence of a distinct family of Si transporters (Grégoire et al., 2012). These are members of the NIP2 subfamily and, in place of the GSGR Ar/R filter, possess a STAR (serine–threonine–alanine–arginine) filter predicted to have a pore large enough to permit passage of silicic acid (Grégoire et al., 2012). Three horsetail transporter subgroups were identified, and EaNIP3;1, EaNIP3;3, and EaNIP3;4 are all efficient Si transporters in the Xenopus oocyte system and showed strong expression in roots. When EaNIP3;1 was expressed in Arabidopsis, it resulted in accumulation of Si in Arabidopsis shoots. A study of date palm (Bokor et al., 2019) resulted in characterization of an aquaporin Si transporter, also possessing the GSGR Ar/R selectivity filter, PdNIP2-1. Si transport was demonstrated using Xenopus oocytes. Al transport mechanisms Al transporters have also been characterized for the first time in plants since we wrote our previous review, though this remains a much less well explored area than Si transporters and much of the progress has been in identifying transporters for compounds such as organic acids induced under Al stress. Table 2 presents a listing of proteins suggested to be Al transporters, together with associated references. Zhang et al. (2019) recently reviewed evidence for Al transporters in plants, and readers are directed to this review for detailed consideration; evidence for Al transport has been associated with plasma membrane and tonoplast proteins and with proteins of the ATP-binding cassette (ABC), aquaporin, and natural resistance-associated macrophage protein (Nramp) families. These include ABC partial homologues AtALS3 and AtSTAR1 in Arabidopsis with homologues OsSTAR1 and OsSTAR2 in rice (Larsen et al., 2005, 2007; Huang et al., 2009, 2012). AtALS3 encodes a single domain homologous to a bacterial ABC protein, while the two rice proteins contain the nucleotide-binding domain (OsSTAR1) and transmembrane domain (OsSTAR2) also of a bacterial-type ABC transporter (Huang et al., 2009). However, when co-expressed in the oocyte transport assay system, OsSTAR1 and OsSTAR2 form a functional UDP-glucose transporter. The transporter Nrat1 (Nramp-natural resistance-associated macrophage protein aluminium transporter 1) was described by Xia et al. (2010) in rice and provided evidence that Nrat1 is specific for trivalent Al, localized to the plasma membrane of root tip cells other than the epidermis, and has a role in detoxifying Al in the vacuole; Nrat1 knockouts have enhanced Al sensitivity and increased cell wall Al. Providing evidence for aquaporin family transporters, Negishi et al. (2012, 2013) explored Al transport in the Al hyperaccumulator hydrangea where they identified both plasma membrane (PALT1) and tonoplast (VALT) Al transporters in the sepals. Both PALT1 and VALT are members of the aquaporin family. Expression in arabidopsis confers Al sensitivity (PALT) and tolerance (VALT), suggesting they are part of a pathway of Al detoxification in the vacuole. Table 2. Putative aluminium transporters with supporting publications listed (UniProt Consortium, 2019) Entry . Entry name . Protein names . Gene names . Organism . Length . Reference . Q9ZUT3 ALS3_ARATH Protein ALUMINUM SENSITIVE 3 ABC family member, but lacking ATP binding cassette. ALS3 ABCI16, At2g37330, F3G5.12 Arabidopsis thaliana (mouse-ear cress) 273 Larsen et al. (2005) K0ITY9 K0ITY9_HYDMC Plasma membrane aluminium transporter Aquaporin family PALT1 Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 304 Negishi et al. (2012) Q0WML0 AB27B_ARATH ABC transporter B family member ABCB27 ALS1, TAP2, At5g39040, MXF12.50 Arabidopsis thaliana (mouse-ear cress) 644 Larsen et al. (2007) Q9FNU2 AB25B_ORYSJ ABC transporter B family member ABCB25 ALS1, Os03g0755100, LOC_Os03g54790, OSJNBb0081K01.19 Oryza sativa subsp. japonica (rice) 641 Verrier et al. (2008) K0IVT1 K0IVT1_HYDMC Vacuolar aluminium transporter Aquaporin family VALT Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 252 Negishi et al. (2012) Q0D9V6 STAR1_ORYSJ Protein STAR1 Partial ABC transporter; when expressed with STAR2 transports UDP glucose. STAR1 ABCI12, ALS1, Os06g0695800, LOC_ Os06g48060, P0622F03.26 Oryza sativa subsp. japonica (rice) 346 Huang et al. (2009) Q5W7C1 STAR2_ORYSJ UPF0014 membrane protein STAR2 partial ABC transporter STAR2 Os05g0119000, LOC_ Os05g02750, P0496H07.22 Oryza sativa subsp. japonica (rice) 285 Huang et al. (2009) Q6ZG85 NRAT1_ORYSJ Metal transporter NRAT1 NRAT1 Os02g0131800, LOC_ Os02g03900, OJ1007_D04.24, OsJ_05257 Oryza sativa subsp. japonica (rice) 545 Xia et al. (2010) Entry . Entry name . Protein names . Gene names . Organism . Length . Reference . Q9ZUT3 ALS3_ARATH Protein ALUMINUM SENSITIVE 3 ABC family member, but lacking ATP binding cassette. ALS3 ABCI16, At2g37330, F3G5.12 Arabidopsis thaliana (mouse-ear cress) 273 Larsen et al. (2005) K0ITY9 K0ITY9_HYDMC Plasma membrane aluminium transporter Aquaporin family PALT1 Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 304 Negishi et al. (2012) Q0WML0 AB27B_ARATH ABC transporter B family member ABCB27 ALS1, TAP2, At5g39040, MXF12.50 Arabidopsis thaliana (mouse-ear cress) 644 Larsen et al. (2007) Q9FNU2 AB25B_ORYSJ ABC transporter B family member ABCB25 ALS1, Os03g0755100, LOC_Os03g54790, OSJNBb0081K01.19 Oryza sativa subsp. japonica (rice) 641 Verrier et al. (2008) K0IVT1 K0IVT1_HYDMC Vacuolar aluminium transporter Aquaporin family VALT Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 252 Negishi et al. (2012) Q0D9V6 STAR1_ORYSJ Protein STAR1 Partial ABC transporter; when expressed with STAR2 transports UDP glucose. STAR1 ABCI12, ALS1, Os06g0695800, LOC_ Os06g48060, P0622F03.26 Oryza sativa subsp. japonica (rice) 346 Huang et al. (2009) Q5W7C1 STAR2_ORYSJ UPF0014 membrane protein STAR2 partial ABC transporter STAR2 Os05g0119000, LOC_ Os05g02750, P0496H07.22 Oryza sativa subsp. japonica (rice) 285 Huang et al. (2009) Q6ZG85 NRAT1_ORYSJ Metal transporter NRAT1 NRAT1 Os02g0131800, LOC_ Os02g03900, OJ1007_D04.24, OsJ_05257 Oryza sativa subsp. japonica (rice) 545 Xia et al. (2010) Open in new tab Table 2. Putative aluminium transporters with supporting publications listed (UniProt Consortium, 2019) Entry . Entry name . Protein names . Gene names . Organism . Length . Reference . Q9ZUT3 ALS3_ARATH Protein ALUMINUM SENSITIVE 3 ABC family member, but lacking ATP binding cassette. ALS3 ABCI16, At2g37330, F3G5.12 Arabidopsis thaliana (mouse-ear cress) 273 Larsen et al. (2005) K0ITY9 K0ITY9_HYDMC Plasma membrane aluminium transporter Aquaporin family PALT1 Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 304 Negishi et al. (2012) Q0WML0 AB27B_ARATH ABC transporter B family member ABCB27 ALS1, TAP2, At5g39040, MXF12.50 Arabidopsis thaliana (mouse-ear cress) 644 Larsen et al. (2007) Q9FNU2 AB25B_ORYSJ ABC transporter B family member ABCB25 ALS1, Os03g0755100, LOC_Os03g54790, OSJNBb0081K01.19 Oryza sativa subsp. japonica (rice) 641 Verrier et al. (2008) K0IVT1 K0IVT1_HYDMC Vacuolar aluminium transporter Aquaporin family VALT Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 252 Negishi et al. (2012) Q0D9V6 STAR1_ORYSJ Protein STAR1 Partial ABC transporter; when expressed with STAR2 transports UDP glucose. STAR1 ABCI12, ALS1, Os06g0695800, LOC_ Os06g48060, P0622F03.26 Oryza sativa subsp. japonica (rice) 346 Huang et al. (2009) Q5W7C1 STAR2_ORYSJ UPF0014 membrane protein STAR2 partial ABC transporter STAR2 Os05g0119000, LOC_ Os05g02750, P0496H07.22 Oryza sativa subsp. japonica (rice) 285 Huang et al. (2009) Q6ZG85 NRAT1_ORYSJ Metal transporter NRAT1 NRAT1 Os02g0131800, LOC_ Os02g03900, OJ1007_D04.24, OsJ_05257 Oryza sativa subsp. japonica (rice) 545 Xia et al. (2010) Entry . Entry name . Protein names . Gene names . Organism . Length . Reference . Q9ZUT3 ALS3_ARATH Protein ALUMINUM SENSITIVE 3 ABC family member, but lacking ATP binding cassette. ALS3 ABCI16, At2g37330, F3G5.12 Arabidopsis thaliana (mouse-ear cress) 273 Larsen et al. (2005) K0ITY9 K0ITY9_HYDMC Plasma membrane aluminium transporter Aquaporin family PALT1 Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 304 Negishi et al. (2012) Q0WML0 AB27B_ARATH ABC transporter B family member ABCB27 ALS1, TAP2, At5g39040, MXF12.50 Arabidopsis thaliana (mouse-ear cress) 644 Larsen et al. (2007) Q9FNU2 AB25B_ORYSJ ABC transporter B family member ABCB25 ALS1, Os03g0755100, LOC_Os03g54790, OSJNBb0081K01.19 Oryza sativa subsp. japonica (rice) 641 Verrier et al. (2008) K0IVT1 K0IVT1_HYDMC Vacuolar aluminium transporter Aquaporin family VALT Hydrangea macrophylla (bigleaf hydrangea) (Viburnum macrophyllum) 252 Negishi et al. (2012) Q0D9V6 STAR1_ORYSJ Protein STAR1 Partial ABC transporter; when expressed with STAR2 transports UDP glucose. STAR1 ABCI12, ALS1, Os06g0695800, LOC_ Os06g48060, P0622F03.26 Oryza sativa subsp. japonica (rice) 346 Huang et al. (2009) Q5W7C1 STAR2_ORYSJ UPF0014 membrane protein STAR2 partial ABC transporter STAR2 Os05g0119000, LOC_ Os05g02750, P0496H07.22 Oryza sativa subsp. japonica (rice) 285 Huang et al. (2009) Q6ZG85 NRAT1_ORYSJ Metal transporter NRAT1 NRAT1 Os02g0131800, LOC_ Os02g03900, OJ1007_D04.24, OsJ_05257 Oryza sativa subsp. japonica (rice) 545 Xia et al. (2010) Open in new tab The exclusion of Al from the root is most frequently achieved by the secretion of organic acids through transporters induced in the presence of Al. Two families of transporters have been identified, designated MATEs (multidrug and toxic compound extrusion) and ALMTs (aluminium-activated malate transporters) (Bojorquez-Quninal et al., 2017; Yang et al., 2019; Zhang et al., 2019). As well as preventing Al uptake, other species transport Al to the shoot, where it is chelated. In high Al accumulators, such as hydrangea, buckwheat, and tea (Ma et al., 1997a; Carr et al., 2003; Shen et al., 2004), chelation by small organic compounds is key to detoxification; in buckwheat, this is oxalate, with Al-citrate being the mobile form in the xylem (Ma et al., 1997b; Shen et al., 2004). In hydrangea sepals, the chelator is delphinidin 3-glucoside and is 3-caffeoylquinic and citrate in the leaves (Ma et al., 1997a). The chelator in tea has been shown to be catechin (Nagata et al., 1992). Evidence for a role for Si and Al transporters in amelioration While Si involvement in the amelioration of Al toxicity could be the result of altered transporter activity, evidence for direct effects of Si on Al transporters (and vice versa) is limited. In an examination of this topic, Pontigo et al. (2017) explored the effect of Si on Si and Al transport in ryegrass, using a hydroponic system. Plants were cultivated in varying concentrations of Al and Si. Al uptake was decreased in the presence of Si; and Lsi1 and Lsi2 genes were both down-regulated in roots in the presence of Al and Si, applied singly. However, application of Si to Al-treated plants resulted in an up-regulation of both transporter genes. The authors suggest that this is indicative of an increased Si requirement by the tissue in Al toxicity. Bhat et al. (2019) have recently reviewed the importance of Si in ameliorating heavy metal toxicity. In rice, down-regulation of heavy metal transporter genes (OsHMA2 and OsHMA3), and up-regulation of Si transporter genes (OsLSi1 and OsLSi2) has been observed with Si supplementation (Kim et al., 2014). Similarly, the enhanced expression of phytochelatin synthase 1 (PCS1) and decreased expression of the metallothionein gene (MT1a) was associated with Si supplementation in Arabidopsis under copper stress (Khandekar and Leisner, 2011). Likewise, Si up-regulates OsLsi1 and down-regulates the cadmium (Cd) transporter Nramp5 in Cd stress (Ma et al., 2015). While results like these for heavy metals may shed some light on the role of transporters in amelioration, it is clear additional data are needed before their role is fully understood. Co-deposition of aluminium and silicon We have seen that the formation of HAS in the root apoplast appears to be part of the mechanism whereby Al toxicity is ameliorated by Si. However, as Beardmore et al. (2016) point out, HASA and HASB are precursors of much more insoluble HAS species. It would therefore seem quite likely that we would find solid deposits containing both Al and Si in plants. When we wrote our previous review of this topic (Hodson and Evans, 1995), relatively little was known about the co-deposition of Al and Si in plants. We considered two methodologies for studying this: chemical analysis and microanalysis. There has been less work since that time using chemical analysis, as the difficulties in obtaining samples of phytoliths that were not contaminated with minerals from non-mineralized plant tissue have become apparent. For example, Kameník et al. (2013) used three different extraction methodologies to isolate phytoliths from barley: acid digestion, dry ashing, and acid digestion followed by incineration. They did not recommend just using acid digestion as it left organic matter in the phytoliths, but even the other two methods gave somewhat different results. In all cases, terrigenous elements, including Al, were enriched in the phytoliths compared with the plant material. For the rest of this discussion, we will only consider microanalytical work. In 1995 we could only locate five microanalytical investigations that had concerned this topic. We now have a much better understanding of the scale of Al and Si co-deposition, although it is still far from clear how significant this is in the amelioration of Al toxicity. We will now assess the progress that has been made in this area under four headings: roots; conifers; broad-leaved woody plants (including Al accumulators); and others. Roots As we have seen, the grasses and cereals seem to largely exclude Al from their shoots, and so we would expect most of the interest in Al–Si interactions in these species to be in the roots. That does appear to be the case (Cocker et al., 1998b), but these interactions seem mostly to involve soluble HAS complexes, and there have only been two cases where Al and Si co-deposition has been observed. In both sorghum (Hodson and Sangster, 1993) and wheat (Cocker et al., 1997) plants grown in hydroponic culture, deposits containing Al and Si were located in the root epidermal cell walls. The authors speculated that this co-deposition may prevent some Al from penetrating further into the root cortex. As we noted above, Kopittke et al. (2017) used LEXRF and found that Al and Si were co-localized in the mucigel and outer apoplast in the sorghum root apex, a similar location to that observed by Hodson and Sangster (1993) for the same species. LEXRF is a more sensitive technique than X-ray microanalysis, and it is possible that it was detecting more soluble HAS in addition to deposited material. Until recently, the only example of Al and Si co-deposition in roots that we are aware of outside the grasses was from Norway spruce (Hodson and Wilkins, 1991) where the phenomenon was observed in the cortical cell walls. One slightly unusual example of Al and Si co-deposition comes from the work of Feng et al. (2019) working on the root border cells of pea. They were able to produce an extracellular silica nanocoat formed by layer-by-layer self-assembly on the surface of these cells. When they were then treated with Al, the coating adsorbed Al on its surface preventing it from penetrating into the cells, and thereby decreasing toxicity. The authors suggested that this approach might be used to solve the global Al toxicity problem, but we suspect that much more research will be needed before it can be applied in field situations. Conifer needles Carnelli et al. (2002) included five conifers in their study of Al in the phytoliths isolated from plants growing in the Valaisan Swiss Alps. All of the phytoliths from wood contained Al, and the element was also detected in the needles of three species (Juniperus nana, Pinus cembra, and Pinus mugo). Phytoliths containing Al were also common in woody dicots, but very rarely in grasses and other monocots. These findings led the authors to suggest that Al and Si co-deposition in phytoliths could be used as a marker for woody species in palaeoecological investigations. Following our review in 1995, one of us (MJH) was involved in a whole series of studies investigating mineral localization in the needles of conifers. The species investigated were: white spruce (Hodson and Sangster, 1998); balsam fir (Hodson and Sangster, 1999); American larch and European larch (Sangster et al., 2001); white pine (Hodson and Sangster, 1999, 2000, 2002); Douglas fir (Sangster et al., 2007); and Eastern hemlock (Sangster et al., 2009). These studies gave us a much better idea of the extent of Al and Si co-deposition, its locations, and the environmental factors that affect it. The co-deposition of Al and Si occurred in all the species investigated, but the phenomenon was more pronounced in white pine, Eastern hemlock, balsam fir, and the larch species, and less so in white spruce and Douglas fir. The major locations for Al and Si co-deposition in the needles were the epidermis and the transfusion tissue. Epidermal deposition was most pronounced in white spruce, balsam fir, and the larch species. Al and Si co-deposition in the transfusion tissue was noted particularly towards the tips of the needles in white pine and Eastern hemlock. It was also evident from these investigations that Al and Si co-deposition was only found when the plants were growing in acidic soils. This was most clearly shown in white pine (Table 3) where both Al and Si accumulate in the transfusion tissue near the tips of the needles, but more of both elements are present in soils with a pH below 4.2. It appears that the idea of Carnelli et al. (2002) that Al in phytoliths could be used as a marker for woody species is only likely to work if all of the plants are growing on acidic soils with reasonable Al availability. Table 3. A comparison between microanalysis results from three sites for Al and Si in the transfusion cells of 2-year-old white pine needles Needle zone . Al (mmol kg–1) . Si (mmol kg–1) . . Glendon pH 6.7a . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Glendon pH 6.7 . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Base 9.6 12.7 14 29.2 13 16.6 Middle 13.9 30.2 11.1 13.1 19.5 25 Tip 9.1 44.5 61.4 611 5838 4282 Needle zone . Al (mmol kg–1) . Si (mmol kg–1) . . Glendon pH 6.7a . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Glendon pH 6.7 . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Base 9.6 12.7 14 29.2 13 16.6 Middle 13.9 30.2 11.1 13.1 19.5 25 Tip 9.1 44.5 61.4 611 5838 4282 Data from Hodson and Sangster (1999, 2000, 2002). a Soil pH at 40 cm. Open in new tab Table 3. A comparison between microanalysis results from three sites for Al and Si in the transfusion cells of 2-year-old white pine needles Needle zone . Al (mmol kg–1) . Si (mmol kg–1) . . Glendon pH 6.7a . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Glendon pH 6.7 . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Base 9.6 12.7 14 29.2 13 16.6 Middle 13.9 30.2 11.1 13.1 19.5 25 Tip 9.1 44.5 61.4 611 5838 4282 Needle zone . Al (mmol kg–1) . Si (mmol kg–1) . . Glendon pH 6.7a . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Glendon pH 6.7 . Muskoka pH 4.2 . Sudbury pH 3.5–4.0 . Base 9.6 12.7 14 29.2 13 16.6 Middle 13.9 30.2 11.1 13.1 19.5 25 Tip 9.1 44.5 61.4 611 5838 4282 Data from Hodson and Sangster (1999, 2000, 2002). a Soil pH at 40 cm. Open in new tab Broad-leaved woody plants As we saw above, Carnelli et al. (2002) included seven woody shrub species in their study of Al and Si co-deposition in phytoliths from plants growing in the Swiss Alps. In all cases, Al was detected in phytoliths from leaves and wood. Turnau et al. (2007) conducted a detailed investigation of metal uptake in Erica andevalensis growing on a highly acidic pyrite mine in southeastern Portugal. Using microanalysis they showed that both iron and Al were co-localized with Si in the epidermis and glandular hairs of the leaves. They also found the elements associated with a fungal root endosymbiont, Hymenoschyphus ericae. The authors suggested that the endosymbiont is involved in restricting the entry of toxic metals into the roots, while co-deposition of Al with Si in the leaf epidermis was seen as a detoxification mechanism. In our 1995 review, we wrote that, ‘It would appear that heavy Al accumulation and heavy Si accumulation are mutually exclusive.’ This statement has proven to be largely correct, but it does now appear that some Al accumulator species potentially use Si in detoxification. The best example of this phenomenon is the Al accumulator, Faramea marginata (Rubiaceae), which was studied by Britez et al. (2002). This species is not an Si accumulator, and the authors described it as ‘no more than an intermediate or weak Si accumulator’, and it does not form phytoliths. However, Britez et al. presented very persuasive evidence that Al and Si form complexes, probably including HAS, in their shoot tissues. Most Al accumulators use organic ligands and form complexes with Al (see above), and this was thought to be the first example where an inorganic ligand was used for detoxification. So this appears to be a case where Al and Si co-deposition does not occur, but Si is involved in reducing the availability of Al in the shoot. More recently, Malta et al. (2016) investigated Rudgea viburnoides, another Al accumulator in the Rubiaceae, and using X-ray microanalysis found that Al and Si were co-localized in the roots, stems, and leaves, mostly in the epidermis. Probably the most famous Al accumulator plant is tea (Camellia sinensis) and, according to Matsumoto et al. (1976), it can accumulate >3% Al in its older leaves. These authors used light microscopy and X-ray microanalysis, and located Al in the epidermal cells, but did not consider Si. It appears that there is some uncertainty with regards to Al and Si co-localization in this species. Whilst Tolrà et al. (2011) reported that the two elements were not co-deposited, Haruyama et al. (2019) found that this occurred in the leaf epidermis. The latter suggested that the reason for the different distributions was possibly due to the conditions under which the plants grew and the ages of the leaves. Other plants For completeness, we should also point out that Al and Si co-localization has been observed in species beyond the Angiosperms. Liu et al. (2019) worked on Dicranopteris linearis, a fern which is an Al hyperaccumulator in China. They showed co-deposition of Si with Al and rare Earth elements (REEs). This is the one case we are aware of where a species is both an Al accumulator (up to 9660 mg kg−1) and an Si accumulator (up to 20 300 mg kg−1). Dicranopteris linearis also accumulates high concentrations of REEs (up to 3830 mg kg−1). How this species is capable of doing something that few, if any, others can do is a mystery. Finally, Pressel et al. (2011) found what they considered was the first example of Al and Si co-localization in the bryophytes. Working on the thalloid liverwort, Mizutania riccardioides, they found that the outer part of the wall consisted of irregular blocks of dense material, containing high concentrations of Al and Si. What questions remain? Having assessed 25 years of research on Al–Si interactions in plants what are the topics that still need further research? In the key area of the amelioration of Al toxicity by Si in the root, it seems that our hypothesis stated in Cocker et al. (1998b) is largely correct, and the formation of HAS within the apoplast is at least a major part of the mechanism. Although there has been considerable progress in determining the molecular basis for both Al and Si transport, we have only just begun to investigate the effect of Al on Si transport or of Si on Al transport (Pontigo et al., 2017). It will be some time before we can determine how important these processes are in the amelioration effect. Is the in planta amelioration, often observed, solely due to phenomena in the apoplast which prevent Al uptake, or is it more complex than that? A number of studies have shown that Si has a general effect on plant responses to abiotic stress (for reviews, see Kim et al., 2017; Etesami and Jeong, 2018). Application of Si increases stress tolerance by reducing the generation of reactive oxygen species (ROS). These effects were seen for a wide range of abiotic stresses, including the heavy metals Cd (Shi et al., 2010), manganese (Maksimović et al., 2007), and chromium (Tripathi et al., 2015), but evidence for Al/Si is limited. In a study of the effect of Si nanoparticles (SNPs) on Al toxicity in maize growing in an artificial soil system, de Sousa et al. (2019) demonstrated (amongst other effects) that while Al uptake remained the same as in the absence of SNPs, ROS production was reduced, as was oxidative damage. Antioxidant enzymes and metabolites were enhanced in leaves and roots. In a similar way, Dorneles et al. (2019) showed that Si was partially able to ameliorate Al toxicity in two potato genotypes. The authors related this to increased activity of antioxidant enzymes and mitigation of Al-induced damage to membrane lipids. One of the more disappointing aspects of our review of Al–Si interactions 25 years on is how little research on agricultural systems is recorded in the literature. In fact there have been very few investigations carried out in soil systems. Morikawa and Saigusa (2002) studied the impacts of Si fertilization on Al toxicity in barley plants growing in two andosol soils. They found that addition of silica gel and sodium metasilicate to the soils was ineffective in amelioration of Al toxicity despite the treatments causing an increase in soil Si concentration. Waste porous hydrated calcium silicate did lead to amelioration of Al toxicity but the authors considered that this was probably due to the increase in soil pH. Although the paper by Morikawa and Saigusa has, according to Google Scholar, been cited 31 times (in January 2020), none of the citations refer specifically to further experiments conducted in soil to investigate Al–Si interactions. In the study by de Sousa (2019; see above) on the effect of silicon dioxide nanoparticles on Al toxicity, amelioration but not a reduction in Al accumulation was observed. The overall effect was suggested to result from stimulation of organic exudation in the roots. It may be that the paucity of studies in this area is due to the complexities of the chemistry, particularly in soil. So at present we have definite ameliorative effects in hydroponics, but uncertainty as to whether the findings can be transferred to field situations and under what conditions. The overall beneficial effect of Si on stress and Si soil treatments on pH suggest that this is worth further exploration. We now know much more about Al and Si co-deposition than we did, particularly in the conifers. However, it is unclear whether this is part of a detoxification mechanism or is just coincidental. Al as Al3+ is toxic to plant cells, and precipitation in a solid form should decrease toxicity, but we have no direct experimental evidence that proves this. General conclusions Our knowledge of Al–Si interactions in plants has markedly increased in the last 25 years. We have determined a mechanism for the amelioration effect in roots that has withstood detailed investigation. This mechanism shows the importance of the apoplast in the amelioration phenomenon. We now have greatly increased knowledge of Al and Si transporters but have yet to ascertain whether these have a role in amelioration. It appears that Al and Si co-deposition is quite common in some plant groups, but its functional significance is not certain. 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This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited. © The Author(s) 2020. Published by Oxford University Press on behalf of the Society for Experimental Biology.

Deshmukh,, Rupesh;Sonah,, Humira;Belanger, Richard, R

doi: 10.1093/jxb/eraa342pmid: 32710120

Abstract Understanding the evolution events defining silicon (Si) uptake in plant species is important for the efficient exploration of Si-derived benefits. In the present study, Si accumulation was studied in 456 diverse plant species grown in uniform field conditions, and in a subset of 151 species grown under greenhouse conditions, allowing efficient comparison among the species. In addition, a systematic analysis of nodulin 26-like intrinsic proteins III (NIP-III), which form Si channels, was performed in >1000 species to trace their evolutionary path and link with Si accumulation. Significant variations in Si accumulation were observed among the plant species studied. For their part, species lacking NIP-IIIs systematically showed low Si accumulation. Interestingly, seven NIP-IIIs were identified in three moss species, namely Physcomitrella patens, Andreaea rupestris, and Scouleria aquatica, indicating that the evolution of NIP-IIIs dates back as early as 515 million years ago. These results were further supported from previous reports of Si deposition in moss fossils estimated to be from around the Ordovician era. The taxonomical distribution provided in the present study will be helpful for several other disciplines, such as palaeoecology and geology, that define the biogeochemical cycling of Si. In addition to the prediction of Si uptake potential of plant species based on sequence information and taxonomical positioning, the evolutionary path of the Si uptake mechanism described here will be helpful to understand the Si environment over the different eras of land plant evolution. Aquaporins, genomics, mosses, nodulin 26-like intrinsic proteins, plant evolution, silicon Introduction Silicon (Si) plays an important role in plant biology by providing protection against many biotic and abiotic stresses in crop plants (Coskun et al., 2019). While not considered a nutrient per se (Epstein, 2009), Si has recently been classified as a beneficial element by the International Plant Nutrition Institute (IPNI, www.ipni.net/nutrifacts-northamerican) on the basis of hundreds of reports describing its positive role. However, plant species do not respond similarly to Si because of their different ability to absorb the element, as a wide range of concentrations, from 0.1% to as high as 10% Si (DW basis), has been observed (Epstein, 1994). Initially, based on the concentrations measured, plants were categorized as low, intermediate, and high accumulators (Jones and Handreck, 1967). In general, there is a positive correlation between the ability of a species to absorb Si and the benefits it derives from the element, so it is important to properly assess the properties of a plant to take up Si. In this context, Hodson et al. (2005) conducted an exhaustive analysis of 735 plant species from 125 different studies and normalized the data based on measurements from at least two independent studies for each species, in order to classify plants according to their ability to accumulate Si. At the time when Si transporters had not yet been discovered, this dataset provided a valuable resource of phylogenetic distribution within the plant kingdom for Si accumulation. Even though the data normalization can sometimes be misleading considering the different conditions under which the experiments are conducted, the approach is convenient to determine variation due to genetic and environmental factors. More recently, Trembath-Reichert et al. (2015), using Hodson et al.’s dataset to normalize their own Si data from plant collections, drew conclusions from these renormalized data to establish a history of Si uptake in plants. As a result of this normalization, the study obtained overestimation or underestimation of Si in many species. The discrepancy related to Si accumulation data in plants among different studies can be explained by many factors such as variable Si content in the growing medium, plant-available Si (PAS) in soil, age of the plant, Si quantification methods, etc. For instance, PAS in the growing medium or soil was rarely considered in the previous studies while comparing Si in plant tissues. The same plant species grown in different media or sampled at different ages can present large variations in Si content, such as a range of 0.2–3% observed in strawberry (Ouellette et al., 2017). Therefore, to better categorize plant species in terms of their ability to accumulate Si, standard methods need to be implemented together with the integration of the molecular mechanisms involved in Si uptake by plants. The seminal discoveries of Si transporters in rice by Ma et al. (2006, 2007) have laid the foundation to exploit the molecular mechanisms inherent to Si uptake to better understand how and what plants can accumulate the element. According to Ma et al. (2006, 2007), Si will enter the plant from the outside environment in the form of silicic acid through specific Si influx transporters (termed Lsi1), while efflux transporters (termed Lsi2) will translocate Si into the xylem which takes it through xylem flow to the aerial parts of the plants where it will deposit as amorphous Si (SiO2). The Si influx proteins belong to the large family of aquaporins (AQPs) (Ma et al., 2006). AQPs are a class of channel-forming proteins that facilitate the transport of water and many other small solutes across the cell membrane (Deshmukh et al., 2016). They have a characteristic hourglass-like structure made up of six transmembrane (TM) domains, and two half TM helices protruding from opposite sides towards the center of the pore (Murata et al., 2000). The two half TM helices form a constrict hosting two NPA motifs at the center of the pore. The pore forms another constrict, often referred to as a selectivity filter (SF), and is composed mostly of four amino acids. The amino acids at the SF are usually highly conserved and involved in the solute specificity of a given AQP. Interestingly, the phylogenetic distribution of Si influx transporters identified in different crop plants, including monocots and dicots, showed a specific clustering of all known Si influx transporters within the nodulin 26-like intrinsic protein III (NIP-III) group (Deshmukh and Bélanger, 2016). Moreover, all Si influx transporters in crop plants identified to date have an SF composed of a glycine–serine–glycine–arginine (GSGR) conserved sequence (Deshmukh and Bélanger, 2016). Therefore, plant AQPs belonging to the NIP-III group with a GSGR SF, six TM domains, and two NPA motifs can be categorized as candidate Si transporters. While much is known about the structural and functional features of Lsi1 transporters and their influence on tissue Si content, very little is known about Lsi2 in comparison. No crystallographic structures have ever been resolved for Lsi2, which creates a bottleneck to understand how exactly Lsi2 actively transports Si (Ma et al., 2006, 2007). To date only a few homologs have been reported in a limited number of plant species (Vatansever et al., 2017). With new genomic data becoming available for an ever-expanding number of plant species, this should support future research to decipher with greater precision the properties of Si efflux transporters. Currently, while both influx and efflux Si transporters contribute to Si uptake from soil and subsequent transport to aerial tissues, it is clear that Lsi1s are essential by providing the primary entry of Si into the plant roots. Confirming the presence of AQPs bearing all the molecular signatures of an Si influx transporter in a plant should theoretically lead to an accurate prediction of a plant’s ability to take up Si. In this context, the rapidly increasing sequence resources for plants provide a perfect opportunity to mine putative Si transporters in many crop plants and model species and determine if their presence can be correlated with Si accumulation in plants. In an effort to provide a definitive and precise classification of Si-accumulating plant species, the objectives of this study were: (i) to define the effects of PAS in soil, plant age, plant organs, and plant genotypes on the phenotypes of Si accumulation; (ii) to identify molecular signatures associated with phenotypes; and (iii) to understand the evolution of NIP-IIIs in the plant lineage. On the basis of >1000 plant species’ transcriptomes analyzed, our results show that only plant species carrying NIP-III AQPs with specific molecular signatures can accumulate significant Si. We conclude that plants can be classified directly as an Si accumulator or not, based on the acquisition/presence of these specific NIP-IIIs, a finding that is also useful to better understand the evolution of Si uptake in plant lineages. Materials and methods Plant material and growth conditions Plant propagation material such as seeds, cuttings, and bulbs maintained at the Department of Horticulture, University Laval was used for growing plants on raised bed garden soil. The same genetic stock was used for the greenhouse experiment. For the evaluation of closely related species and intraspecies variation, seed material was obtained from the Germplasm Resources Information Network (GRIN), the United States Department of Agriculture (USDA). In the greenhouse experiment, a total of 151 plant species were grown in 8 inch plastic pots with three replications. Potting mix was prepared from the Fafard AGRO Mix G10 (https://fafardpro.ca), garden topsoil, and washed fine sand at the ratio of 10:2:1 (v/v/v). Plants were irrigated with 1.7 mM Si provided in the form of potassium silicate (Kasil6, 23.6% SiO2, National Silicates, Etobicoke, Toronto, Ontario, Canada). Growth conditions in the greenhouse were maintained at ~22 °C day/18 °C night temperature, 80% humidity, and 16 h/8 h photoperiod cycle ensured with the artificial light source. In the field experiment, raised beds prepared from uniform garden topsoil, farmyard manure (FYM), and compost were used to grow 456 plant species. The plants were irrigated through a precisely regulated drip irrigation system. Recommended seed treatment and agronomical practices were applied to raise healthy and disease-free plants in the greenhouse as well as in the field experiment. Silicon quantification in plant and soil samples Healthy and mature leaves were collected from 30-day-old plants grown in a greenhouse with Si supplementation and from plants grown on uniform raised beds in the field. Leaf tissues were directly collected in polypropylene tubes and dried in a hot-air incubator at 65 °C for 24 h. Dried leaves were powdered by using a bead homogenizer (Omni Bead Ruptor, Omni International). The dried leaf powder was then compressed into a 5 mm diameter size pellet using a manual compressor at a uniform pressure. The pellets were subjected to Si quantification using a portable X-ray fluorescence spectrometer (Niton XL3t900 GOLDD XRF analyzer; Thermo Scientific), according to the methods of Reidinger et al. (2012). The PAS was quantified using the calcium chloride extraction method (Liang et al., 2015). Soil samples collected from seven different fields in Iowa, USA were air-dried and then digested with 0.01 M calcium chloride, centrifuged, and finally the dissolved amount of Si in the supernatant was quantified colorimetrically. At the same time, leaf samples from soybean genotype Majesta grown in the same fields were harvested in mid-July. Si concentration was then quantified in soybean leaves as described above. Identification and characterization of aquaporins Whole-genome sequence annotation information was retrieved from different databases (Supplementary Table S1 at JXB online). A local database of transcript and protein sequences of predicted gene models was created for each species using command-line BLAST utilities provided in the BioEdit software tool (version 7.0.9.0; Hall, 2011). To identify putative AQPs, a BLAST search was performed against the local database using query sequences of 342 known AQP genes (Supplementary Dataset S1). To claim a significant match, the cut-off of the e-value <10–5 and bit-score >100 was used. Protein tertiary structure modeling and pore characterization Protein tertiary structure was developed using the I-TASSER (Iterative Threading ASSEmbly Refinement) server (https://zhanglab.ccmb.med.umich.edu/I-TASSER/). Subsequently, the MOLE online server was used to predict the pore and pore-lining residues (https://mole.upol.cz/). Identification of NIP-IIIs in transcriptomic sequences Transcriptome data available for 1000 diverse plant species (1KP) belonging to different orders was used to identify NIP-III homologs (Carpenter et al., 2019; One Thousand Plant Transcriptomes Initiative, 2019). As described above, query sequences of known AQPs were used to perform BLAST search against the 1KP database (Supplementary Dataset S1). The NIP-IIIs in the AQP homologs were further identified based on the phylogenetic distribution. Phylogenetic analysis A phylogenetic tree was made by using the RAxML method provided in The CIPRES Science Gateway to identify genes specific to the NIP-III group. Similarly, the taxonomical tree was developed based on information retrieved from the NCBI taxonomy browser and visualized through the PhyloT (https://phylot.biobyte.de/). Results Plant age and tissue type affect Si accumulation To evaluate the effects of plant age and tissue type on Si accumulation, soybean (dicot) and barley (monocot) plants were tested for Si uptake over a period of 5 weeks. Both species gradually accumulated Si over time, and the concentration in leaf tissues saturated after 4 weeks (Fig. 1a). The highest concentration of Si was observed in the leaves, whereas smaller amounts were detected in the stem and root tissues of both species (Fig. 1b). Fig. 1. Open in new tabDownload slide Effect of plant age, tissue type, and plant-available silicon (PAS) in soil on silicon (Si) accumulation in plants. (a) Si accumulation in soybean and barley over a span of 5 weeks; (b) Si concentration in different tissues of soybean and barley; (c) univariate relationship between PAS and Si concentrations in soybean leaves. Fig. 1. Open in new tabDownload slide Effect of plant age, tissue type, and plant-available silicon (PAS) in soil on silicon (Si) accumulation in plants. (a) Si accumulation in soybean and barley over a span of 5 weeks; (b) Si concentration in different tissues of soybean and barley; (c) univariate relationship between PAS and Si concentrations in soybean leaves. Silicon accumulation depends on plant-available silicon in the soil Soybean, previously described as a good Si-accumulating dicot plant species (Deshmukh et al., 2016), was grown on different soils to evaluate the relationship between Si accumulation and PAS in soil. Si concentrations ranging from 0.4% to 1.5% were quantified in leaves of soybean genotype cv. Majesta grown in different fields across Iowa with PAS varying from 13 ppm to 139 ppm. The results showed a strong positive correlation between Si accumulation in soybean leaves and PAS in the soil despite possible variations in environmental conditions across the fields (Fig. 1c). Silicon accumulation in 456 plant species grown in soil A set of 456 diverse plant species (658 total genotypes) belonging to 73 families representing 32 different taxonomical orders were grown on raised beds prepared with uniform soil. Plant species belonging to orders Poales, Zingiberales, Boraginales, and Asterales showed the highest Si accumulation (Fig. 2). In contrast, members of the Solanaceae and Brassicaceae families showed very little Si accumulation (Supplementary Table S2). Overall, Si accumulation observed in the different species was lower than amounts previously reported in greenhouse studies (Deshmukh et al., 2014), as a result of the low PAS (20 ppm) present in the soil. For instance, well-known Si accumulators such as maize and soybean contained as little as 0.5% Si (DW), while several Poaceae species recorded <1% Si (DW). Fig. 2. Open in new tabDownload slide Phylogenetic distribution of silicon (Si) accumulation in species belonging to diverse taxonomical plant orders. The entire set of genotypes representing 456 diverse plant species grown on soil supplemented with 20 ppm plant-available Si was evaluated for Si accumulation. The plant orders from the monocot clade are shown in blue text. Fig. 2. Open in new tabDownload slide Phylogenetic distribution of silicon (Si) accumulation in species belonging to diverse taxonomical plant orders. The entire set of genotypes representing 456 diverse plant species grown on soil supplemented with 20 ppm plant-available Si was evaluated for Si accumulation. The plant orders from the monocot clade are shown in blue text. Silicon accumulation in plant species grown under optimal silicon feeding A wide range of Si accumulation was observed in a set of 151 diverse plant species fed with 1.7 mM Si under greenhouse conditions over a 4 week period. Plant species belonging to Poaceae, Fabaceae, and Cucurbitaceae families accumulated in general well over 1% Si (Fig. 3; Supplementary Table S3). Surprisingly, several species found to be low Si accumulators under field conditions (Fig. 2) displayed a high affinity for the element under greenhouse conditions. Notable examples were soybean, maize, and Brachypodium, which contained >2% Si (DW) under a 1.7 mM treatment compared with ~0.5% Si (DW) in soil experiments (Supplementary Tables S2, S3). Fig. 3. Open in new tabDownload slide Phylogenetic distribution of silicon (Si) accumulation in species belonging to diverse taxonomical plant orders grown under greenhouse conditions. The entire set of genotypes representing 151 diverse plant species grown with supplementation of 1.7 mM Si. The plant orders from the monocot clade are shown in blue text. Fig. 3. Open in new tabDownload slide Phylogenetic distribution of silicon (Si) accumulation in species belonging to diverse taxonomical plant orders grown under greenhouse conditions. The entire set of genotypes representing 151 diverse plant species grown with supplementation of 1.7 mM Si. The plant orders from the monocot clade are shown in blue text. Several families within the monocot clade, which are mostly considered as high Si accumulators, were found to contain concentrations as low as 0.01%. For instance, very low Si concentrations were recorded for the plant members of the Asparagales and the Cannaceae families (Supplementary Tables S2, S3). Low accumulation in the latter family is more surprising since it belongs to the order Zingiberales which is taxonomically close to Poales. Intraspecies variation for silicon accumulation Si content among high and low accumulator species was analyzed on different genotypes of eight species to determine intraspecies variation. As shown in Fig. 4, regardless of the predisposition of a given species to accumulate Si, all genotypes within a species were remarkably similar in Si content. All genotypes of high-accumulating species had Si concentrations well above 1%, while those of low accumulating species were significantly below 0.5% when grown under greenhouse conditions. Fig. 4. Open in new tabDownload slide Intraspecies variation in leaf silicon (Si) content in different genotypes of four species reported to be high Si accumulators (a), and four species reported to be low accumulators (b). The coefficient of variation for Si accumulation in these species are: Hordeum vulgaris 0.02, Vigna mungo 0.15, Phaseolus vulgaris 0.10, Glycine max 0.11, Solanum lycopersicum 0.23, Beta vulgaris 0.09, Brassica oleracea 0.11, and Tropaeolum majus 0.15. Fig. 4. Open in new tabDownload slide Intraspecies variation in leaf silicon (Si) content in different genotypes of four species reported to be high Si accumulators (a), and four species reported to be low accumulators (b). The coefficient of variation for Si accumulation in these species are: Hordeum vulgaris 0.02, Vigna mungo 0.15, Phaseolus vulgaris 0.10, Glycine max 0.11, Solanum lycopersicum 0.23, Beta vulgaris 0.09, Brassica oleracea 0.11, and Tropaeolum majus 0.15. Phylogenetic distribution and evolution of silicon influx transporter in plant lineages A total of 4170 AQPs were identified in the set of 116 plant genomes through a BLAST search performed by using previously reported AQPs as query sequences (Supplementary Dataset S1). Subsequently, the identification of conserved motifs, TM domains, and the tertiary structure was evaluated to confirm the AQPs (Fig. 5a). Among those, a total of 280 AQPs with either both or one NPA motif missing were excluded from further analysis. The removed 280 AQPs only had partial sequences and were very short in length. A total of 140 NIP-IIIs (Si influx transporter) representing 81 plant genomes were identified based on the information including amino acid constitution, top hit against previously reported AQP genes, and clustering in the phylogenetic tree of 3890 AQPs (Supplementary Table S4). Similarly, a total of 16 875 AQPs initially identified using transcriptome data were filtered to 11 551 AQPs based on the presence of two NPA motifs. Finally, phylogenetic analysis of 1544 NIPs showed three distinct groups, namely NIP-I, NIP-II, and NIP-III. A total of 349 proteins belonging to 236 plant species were identified as NIP-III with the transcriptome data (Supplementary Table S5; Supplementary Fig. S1). Fig. 5. Open in new tabDownload slide Characterization and phylogenetic analysis of aquaporins. (a) Simplified 2-D structure of a typical NIP-III showing conserved amino acid residues and motifs known to have functional roles in silicon permeability. (b) The tertiary structure of rice NIP-III (OsNIP2-1) showing pore morphology and constricts formed by the Ar/R selectivity filter and NPA motifs. Fig. 5. Open in new tabDownload slide Characterization and phylogenetic analysis of aquaporins. (a) Simplified 2-D structure of a typical NIP-III showing conserved amino acid residues and motifs known to have functional roles in silicon permeability. (b) The tertiary structure of rice NIP-III (OsNIP2-1) showing pore morphology and constricts formed by the Ar/R selectivity filter and NPA motifs. Presence of NIP-IIIs and associated silicon uptake in angiosperms Among the sequences of 715 angiosperm species analyzed in the present study, only 264 species harbored NIP-IIIs (Supplementary Tables S4, S5). Most of them displayed characteristic G-S-G-R Ar/R SF, with a few exceptions. For instance, an S-S-G-R SF was observed in two monocots, Phalaenopsis equestris and Dendrobium catenatum, and five dicots, Schlegelia parasitica, Cota tinctoria, Matricaria matricarioides, Solidago canadensis, and Bituminaria bituminosa. Another notable exception was an A-S-G-R amino acid motif present in a few species of the dicot families such as the Fabaceae, Chenopodiaceae, and Papaveraceae. Surprisingly, the Fabaceae species Phaseolus vulgaris, Vigna angularis, and Vigna radiata, which accumulated Si well above 1% (Supplementary Table S3), harbored NIP-IIIs with an A-S-G-R SF (Supplementary Tables S4, S5). Another exception was specifically observed in nine species (Philoxerus vermicularis, Alternanthera spp., Alternanthera brasiliana, Alternanthera sessilis, Aerva persica, Alternanthera sessilis, Alternanthera caracasana, Amaranthus retroflexus, and Amaranthus cruentus) of the Amaranthaceae family that carried a V-S-[A/G]-R SF. Among them, A. brasiliana only accumulated 0.69% Si under greenhouse conditions. (Supplementary Table S3). Apart from A. brasiliana, four other members of the Amaranthaceae (Amaranthus caudatus, Amaranthus paniculatus, Amaranthus tricolor, and Gomphrena globosa) were also found to be low Si accumulators. Based on the premise of a conserved G-S-G-R SF, the first position, G, located in helix 2 (H2) was found to be the least conserved (Supplementary Table S4; Fig. 5a). Similarly, the fifth position of the Froger’s residue (Froger et al., 1998) present in transmembrane helix 6 (H6) was found to be the most variable across the NIP-IIIs from different species, but no association was observed with Si accumulation (Supplementary Fig. S2). Mitani’s residue (Mitani et al., 2011), earlier found to be associated with the genotypic variation for Si uptake in cucumber, was found to be the most conserved feature in NIP-IIIs. In the case of NPA domains, NPV sometimes replaced NPA in NIP-IIIs. Interestingly, Cucurbitaceae species such as Cucumis melo and Cucumis sativus carried NPV in loop E but were nevertheless considered high accumulators (Supplementary Table S4; Fig. 5a). Analysis of the NPA motifs also revealed that a spacing of 108 amino acids between two NPA motifs was a conserved feature of Si-permeable NIP-IIIs. Indeed, among 20 species that were common between the greenhouse experiment and the sequencing dataset, all were found to carry an NPA spacing of 108 amino acids (Supplementary Tables S4, S5). In contrast, species having NIP-IIIs with an NPA spacing different from 108 showed consistently low Si accumulation. For instance, Tragopogon porrifolius, a species from the Asteraceae family, carrying two NIP-IIIs with G-S-G-R and G-I-G-R SFs, and NPA spacings of 105 and 136 amino acids, respectively, had a low Si content (Supplementary Tables S3, S5). Similarly, three species of the Solanaceae family, Ipomoea purpurea, Solanum lycopersicum, and S. tuberosum with NPA spacing of 110, 109, and 109 amino acids, respectively, were all low Si accumulators (Supplementary Tables S3, S4). Another low accumulator, A. brasiliana from the Amaranthaceae family, was found to have a V-S-A-R SF and an NPA spacing >108 amino acids. Absence of NIP-IIIs in angiosperm families The absence of NIP-IIIs from a plant genome is challenging to confirm with transcriptomic data because it represents only a portion of the total genes that are expressed. Therefore, only well-annotated genomic data were used to investigate the absence of NIP-IIIs within the angiosperm species. Out of 116 whole-genome sequenced plants, 35 plant species did not show the presence of NIP-IIIs (Fig. 6). Interestingly, the nine species within the Brassicaceae family showed complete absence of NIP-IIIs. Accordingly, other Brassicaceae species such as Arabidopsis thaliana, Brassica napus, Brassica oleracea, Brassica rapa, and Camelina sativa, tested for Si uptake, contained Si concentrations <0.2% (Supplementary Table S3). In the same manner, two species of the Solanaceae family, Nicotiana tomentosiformis and Capsicum annuum, lacked NIP-IIIs and were found to be low Si accumulators (Supplementary Table S3). Within the Solanaceae family, some species were found to carry NIP-IIIs, but all had an NPA–NPA spacing >108 amino acids and tested to be low Si accumulators (Supplementary Table S3). Fig. 6. Open in new tabDownload slide Taxonomical phylogeny of 116 plant species based on whole-genome sequence data showing presence or absence of nodulin 26-like intrinsic protein-IIIs (NIP-IIIs). Presence of NIP-IIIs is denoted with a red circle, whereas a blue triangle represents the absence of NIP-IIIs. Fig. 6. Open in new tabDownload slide Taxonomical phylogeny of 116 plant species based on whole-genome sequence data showing presence or absence of nodulin 26-like intrinsic protein-IIIs (NIP-IIIs). Presence of NIP-IIIs is denoted with a red circle, whereas a blue triangle represents the absence of NIP-IIIs. Interspecies variation in Asteraceae A surprisingly high level of interspecies variation was observed within the Asteraceae. For instance, sunflower contained the highest Si concentration among all dicot species analyzed in this study, whereas other Asteraceae species such as Aster alpinus, Cladanthus arabicus, Erigeron speciosus, Hieracium villosum, Jacobaea maritima, Osteospermum ecklonis, Tragopogon porrifolius, and Cynara cardunculus recorded <0.2% Si (Fig. 3; Supplementary Table S3). Among the 29 Asteraceae species evaluated in the greenhouse experiment for Si accumulation, only 11 species from the Heliantheae alliance showed Si accumulation >1% (Fig. 7). Further analysis of the very recently made available genome sequence of Asteraceae member Cynara cardunculus var. scolymus (http://www.artichokegenome.unito.it/) showed the absence of NIP-IIIs. In addition, Tragopogon porrifolius having a NIP-III with NPA spacing >108 amino acids is also present in the same clade of C. cardunculus. All the species taxonomically closer to C. cardunculus and T. porrifolius showed low Si content compared with the species from the Heliantheae alliance group (Fig. 7). Fig. 7. Open in new tabDownload slide Phylogeny based on the taxonomical distribution of species from the Asteraceae family showing silicon (Si) determined in the leaf tissue. Plants were analyzed for Si uptake under optimal Si supplementation in the greenhouse. Fig. 7. Open in new tabDownload slide Phylogeny based on the taxonomical distribution of species from the Asteraceae family showing silicon (Si) determined in the leaf tissue. Plants were analyzed for Si uptake under optimal Si supplementation in the greenhouse. NIP-IIIs in gymnosperms In gymnosperms, NIP-IIIs were observed in 16 out of 76 species (Supplementary Tables S4, S5). There was a clear delineation on the basis of order where NIP-IIIs were found in species within Cupressales and Pinales, and absent in species belonging to the Araucariales, Cycadales, Ginkgoales, Gnetales, and Welwitschiales. All NIP-IIIs in gymnosperms carried a very specific S-[S/A]-G-R SF, all with the specific 108 amino acid NPA spacing. Interestingly, as mentioned above, an S-S-G-R SF was also observed in seven angiosperms. Unlike angiosperms, where the second NPA motif in loop E sometimes showed an NPV variation, a similar variation was also observed in gymnosperms, but only in the first NPA motif (NP[A/V]) located in loop B (Supplementary Table S4, Fig. 8). Fig. 8. Open in new tabDownload slide Evolution of the Ar/R selectivity filter (SF) and NPA motifs in NIP-IIIs identified in plant lineages. Fig. 8. Open in new tabDownload slide Evolution of the Ar/R selectivity filter (SF) and NPA motifs in NIP-IIIs identified in plant lineages. NIP-IIIs in ferns In ferns, NIP-IIIs were identified in nine out of 60 species studies. Fern orders such as Equisetales, Ophioglossales, Osmundales, and Polypodiales have NIP-IIIs with a very distinct Ar/R SF. For instance, Equisetales, known as one of the highest Si-accumulating plants, has NIP-IIIs with T-N-A-R and F-A-A-R SFs, whereas Polypodiales and Osmundales have a G-I-G-R SF, all with conserved 108 amino acid NPA spacing. Similar to previously reported Si influx transporters in Equisetum arvense (Grégoire et al., 2012), NIP-IIs comprising S-T-A-R Ar/R SFs with characteristic 108 NPA spacing were identified in Equisetum hyemale and E. diffusum, and also in a fern species Mapania palustris. Surprisingly, NIP-IIs with a S-T-A-R SF were also observed in two monocot species, Dipteris conjugate and Lepidosperma gibsonii from the Cyperaceae (Supplementary Table S6). Clubmosses, mosses, and liverwort We identified NIP-IIIs in a total of 22 clubmosses. The majority of them had F-A-A-R and N-N-A-R Ar/R SFs. Furthermore, a NIP-III with a G-S-G-R SF along with 108 amino acid NPA spacing and a conserved Mitani’s residue, as observed in flowering plants, was also present in Phylloglossum drummondii, a species from the clubmoss order Lycopodiales. Since this finding was unusual, the NIP-III sequence from P. drummondii was further confirmed by a BLAST search performed against the 1KP database (Supplementary Table S5). As expected, the sequence matched perfectly with the sequence from P. drummondii (self-match) but, surprisingly, the second top hit was the NIP-III sequence from Cana sp., a member of the flowering plants. There was no sequence similarity with other clubmoss sequences. Therefore, a BLAST search was also performed by using the NIP-III with the F-A-A-R SF from P. drummondii as a query sequence, and the top hit after the self-match was, as expected, a NIP-III sequence from Huperzia selago, a member of the Lycopodiaceae family. A total of seven NIP-IIIs were identified in three moss species (Physcomitrella patens, Andreaea rupestris, and Scouleria aquatic). They carried F-A-A-R and G-V-A-R SFs with NPA spacing of 108 and 107/106, respectively. In liverwort, two NIP-IIIs from Pellia cf. Epiphylla with a F-A-A-R/P SF and with NPA spacing of 116 and 107 amino acids were observed (Supplementary Table S5). Algae were devoid of NIP-IIIs. In addition, the NIPs identified in algae can be considered as the closest possible homolog of NIP-IIIs and have an NPA spacing >108 amino acids. Discussion Si is now widely accepted as a beneficial element for plant growth because of its protective role observed under stress conditions (Coskun et al., 2019). In general, high Si accumulator plant species are found to attain more benefits compared with low accumulating speicies (Coskun et al., 2019). However, ascertaining the ability of a given species to accumulate Si has led to conflicting reports (Hodson et al., 2005; da Silva Lobato et al., 2013; Ouellette et al., 2017). Earlier efforts to classify the Si-accumulating properties of plants have painstakingly compiled data from over a hundred studies, and did not have access to genomic resources (Hodson and Sangster, 2002; Hodson et al., 2005). In the present study, Si uptake in >450 diverse plant species was evaluated under uniform growth conditions, extensive sequence analysis was performed in >1000 plant species to identify NIP-IIIs, and the information was used to corroborate with Si uptake data. As a result, we have identified conserved features of NIP-IIIs explaining the evolutionary history associated with the functionality of NIP-IIIs with respect to Si permeability, and factors affecting inter- and intraspecies variations related to Si uptake at the genetic and phenotypic levels. Classifying the ability of a plant to accumulate Si has always been challenging because of the inconsistencies associated with PAS in growing media, plant age, plant tissue, and plant physiology (Rosen and Weiner, 1994; Hodson et al., 2005; da Silva Lobato et al., 2013; McLarnon et al., 2017; Ouellette et al., 2017). To draw the correct inference from the comparison of Si uptake in different species, controlled experiments with uniform growing conditions is a prerequisite. In the present study, we have shown that plant age, tissue type, and PAS can all affect Si concentrations. In terms of plant age, a 4 week growth period was necessary to reach saturation in both barley and soybean (Fig. 1a), although other species such as strawberry and white spruce have shown continued accumulation well beyond that period, probably because of simultaneous accumulation in the multiple branch crowns and needles, respectively (Hodson and Sangster, 1998; Ouellette et al., 2017). Regarding Si accumulation in different plant tissues, the comparisons made in the present study clearly showed the highest level of Si in the leaves (Fig. 1b), a result consistent with the absorption transport model described by Ma et al. (2006). As expected, PAS had a significant effect on Si uptake (Fig. 1c). Under low soil Si concentrations (<20 ppm), soybean plants accumulated <0.4% Si which would classify them as low accumulators, while concentrations above 60 ppm yielded nearly 1% Si. These results explain why soybean has been considered a low accumulator in some work (Van der Vorm, 1980; Arsenault-Labrecque et al., 2012) and a moderate to high accumulator in other works (Deshmukh et al., 2013). While large variations in Si accumulation exist among species, it was quite noteworthy to observe more uniform Si accumulation among genotypes within a species (Fig. 4). This indicates that the phenotype is under strong genetic control and that it is highly conserved within a species. As a general rule, monocots are considered high Si accumulators and dicots low accumulators (Hodson et al., 2005; da Silva Lobato et al., 2013; Ouellette et al., 2017). In the present study, several dicot families, including the Cucurbitaceae, Fabaceae, and Asteraceae, showed high Si accumulation, while only monocot species within the Poaceae family did the same. Therefore, the concept that monocots and dicots belong in different classes of Si accumulators appears inappropriate. For instance, the dicot species sunflower accumulated as much Si as rice, a monocot species often referred to as the highest accumulator of commercial crops. Evolution of NIP-IIIs in plant lineages Understanding the evolution of NIP-IIIs is important to provide insights into why some plant species evolved with this trait, whereas others lost or lacked it. In this regard, the seminal discovery of Si influx transporters belonging to NIP-IIIs provided the basis to track the evolution of the gene (Ma et al., 2006). At the same time, recent advances in sequencing technologies helped to develop useful resources for that purpose (Matasci et al., 2014). In this study, by exploiting >1000 transcriptomic data available for diverse plant species, we were able to make phylogenetic inferences to analyze the evolution of the NIP-IIIs. Along with the transcriptomic data, we also used whole-genome sequencing information available for >00 plant species representing all the major clades of land plants (Fig. 6). Identification and in-depth analysis of NIP-IIIs performed here highlighted several interesting facts. For instance, unlike the earlier assumption that the NIP-III Ar/R SFs are exclusively composed of G-S-G-R SF (Trembath-Reichert et al., 2015), our results suggested a more diversified pattern. In angiosperms, the first position appears to be the least conserved, suggesting a minimal role in functionality. In legumes such as Phaseolus vulgaris, Vigna angularis, and Vigna mungo, Si accumulation was recorded at >2% (DW) in spite of an A-S-G-R SF (Supplementary Table S3). Interestingly, results of Mitani-Ueno et al. (2011) supported our findings by showing that substitution of the first amino acid of the SF in the rice NIP-III (OsLsi1) did not affect Si permeability. Nevertheless, the predominance of G at the first position may indicate specificity for other solutes or be attributable to confounding effects of surrounding convergent amino acid sequences constituting specific structural features. Similarly, the change of NPA to NPV does not seem to have an impact on the functionality of the NIP-IIIs. Our results showed high Si uptake in Cucurbitaceae species in spite of having NPV instead of NPA (Supplementary Tables S3, S4). While most AQPs contain NPA motifs, several variations, including NPV, NPS, NPL, NPC, and NPT, were reported in different plant species (Sonah et al., 2017). The NPA motif is also common to AQPs in fungi, insects, and mammals with few exceptions, as observed in plants (Ikeda et al., 2011; Xu et al., 2013; Lu et al., 2017). In the mammalian AQP11, Ikeda et al. (2011) found that the change in a wild-type NPC to NPA had no effect on the subcellular localization but affected its oligomerization. In addition, the change from NPC to NPA was shown to reduce the water permeability of AQP11 (Ikeda et al., 2011). Similar mutagenesis experiments are required to understand the effect of sequence variation at the NPA motif in plant AQPs. Notwithstanding the structure of NPA motifs, their spacing appears to be one of the most conserved and important features with respect to Si permeability of NIP-IIIs (Supplementary Tables S3, S4). In the greenhouse experiments, all species carrying a NIP-III with 108 amino acid NPA spacing recorded high Si accumulation (Supplementary Tables S3, S4). In contrast, species with an NPA spacing deviating from 108 amino acids showed lower Si accumulation. Some dicot families such as the Solanaceae and Amaranthaceae, in particular, appear to have evolved an NPA spacing different from 108, which correlates with their low Si accumulation (Supplementary Tables S3, S4). Earlier, Deshmukh et al. (2015) provided evidence that a 109 amino acid spacing in tomato NIP-III (SlNIP2-1) significantly reduced Si permeability compared with a 108 spacing. Interestingly, NIP-IIs cloned from Equisetum arvense with an NPA spacing of 108 amino acids were observed to have high Si transport activity in the oocyte assay in spite of having a S-T-A-R SF instead of the common G-S-G-R (Grégoire et al., 2012). Sequence analysis performed here also showed NIP-IIs with an S-T-A-R SF in two more species from the Equisetales, suggesting its conservation across the order (Supplementary Table S6). The presence of similar NIP-IIs with an S-T-A-R SF in the monocot species Lepidosperma gibsonii and the fern species Dipteris conjugata is more surprising. It is not clear whether most of the angiosperm species lost the NIP-IIs with an S-T-A-R SF or if the feature has evolved specifically in these two species. In any event, the strong association of a 108 amino acid spacing between NPA motifs with high Si uptake in plants suggests the importance of this trait for the permeability of Si influx proteins. By using whole-genome sequencing information available for >100 plant species, we uncovered new characteristics of NIP-III evolution. The ubiquitous presence of NIP-IIIs in the 21 monocot genomes analyzed combined with their absence in only a few specific dicot families suggests a recent loss of this specific AQP (Fig. 6). This is further supported by the failure of some monocots to efficiently take up Si, which is associated with the loss of functionality of the NIP-IIIs. This may be explained by several factors, including variation in conserved features, improper folding of the protein, improper subcellular localization, transcriptional activity, post-translational modifications, etc. Presently, we know that the constitution of the Ar/R SF, NPA motifs, and NPA spacing are linked with the ability for Si uptake in some species. However, limited information is available to explain how other factors affect protein functionality in NIP-IIIs. Recently, Coskun et al. (2019) identified a single-residue conformational change in tobacco NIP-III that affected Si permeability in spite of the presence of the other essential features. In addition, NIP-IIIs are known to be efficient transporters of many solutes such as Si, boric acid, arsenic, urea, and water (Mitani-Ueno et al., 2011), and the variation observed in the conserved features may not necessarily be associated with the transport activity of other solutes. Based on our phylogenetic analyses (Fig. 7; Supplementary Table S3), we can conclude that NIP-IIIs are not necessarily well conserved within families, as evidenced in the case of Asteraceae. The variations range from complete loss of NIP-IIIs to structural variations in the conserved attributes, suggesting that selection pressure for Si uptake does not extend to all members within a family. In contrast, our results revealed that Si uptake, and by association NIP-IIIs, showed very limited genotypic variation within a species (see Fig. 4). In gymnosperms, only a few studies have suggested a potential role for Si (Hodson and Sangster, 1999; Prabagar et al., 2011; Hogan et al., 2018). In our study, none of the identified NIP-IIIs contained a G-S-G-R SF, which would suggest an inability to take up high concentrations of Si and explain the scarcity of reports linking Si with benefits in gymnosperms. In ferns, different orders such as Equisetales, Ophioglossales, Osmundales, and Polypodiales have NIP-IIIs with distinct SFs. Earlier reports showed a wide range of Si concentrations from 0.1% to 3.9% among 27 fern species (Höhne and Richter, 1981). Within the Equisetales, known for their very high Si uptake, NIP-IIIs with very specific T-N-A-R SFs were observed, but is it unknown if such SFs allow Si permeability. In previous work, Grégoire et al. (2012) showed that Equisetales had evolved Si influx transporters belonging to NIP-IIs, a rare feat probably linked to their atypical plant anatomy dependent on Si. However, based on limited sequence availability at the time, the authors did not identify NIP-IIIs, so it remains to be determined if the latter also contribute to Si uptake in Equisetales. Liverwort and mosses depend greatly on transcellular AQP transport because of their undeveloped vascular system. This explains the larger number of AQP subfamilies observed in the mosses (Danielson and Johanson, 2008). In general, gymnosperms and angiosperms have five distinct AQP subfamilies, while seven subfamilies were reported in mosses (Danielson and Johanson, 2008). Our results identified for the first time the presence of NIP-IIIs in mosses and liverwort, which redefines the evolutionary origin of the NIP-IIIs. Ma et al. (2001) found >1% Si in several species belonging to the bryophytes, clubmosses, and Equisetopsida. Similarly, >3% Si was observed in Ceratophyllum demersum a hornwort species of the Ceratophyllales order (Schoelynck et al., 2010). In an ultrastructural study of the liverwort (Mizutania riccardioides), Si deposition was noted on the cell wall lining (Pressel et al., 2011). Based on the previous studies and our sequence analysis results, it can be inferred that Si uptake and specific patterns of Si distribution have evolved millions of years ago in non-vascular plants. Land plants (Embryophytes) are estimated to have evolved during the middle Cambrian–early Ordovician era, dating back to 515 million years ago (Ma) to 470 Ma (Morris et al., 2018). The occurrence of NIP-IIIs in today’s Bryophyta (first land plants) is not sufficient alone to claim the existence of Si uptake mechanisms in Cambrian Bryophyta. For this reason, fossil records have enormous importance to connect today’s information with historical records. Recently, Si deposition was observed in moss fossils estimated to date from the Ordovician era around 455–454 Ma (Cardona-Correa et al., 2016). The moss fragment fossils described by Cardona-Correa et al. (2016) were claimed to be the oldest fossils presenting distinctive features that helped to link them with modern vascular plants. These findings also correlate well with molecular evidence that estimated the peat moss evolution dating back 607–460 Ma (Laenen et al., 2014; Morris et al., 2018). We identified NIP-IIIs in 22 clubmosses, a seemingly logical finding since NIP-IIIs were also observed in mosses and liverwort. However, the NIP-III with a G-S-G-R SF identified in the clubmoss species P. drummondii is very surprising. To better understand this exception, sequence analysis in other species related to P. drummondii is required. Earlier, two clubmoss species were found to accumulate >3% Si (Ma and Takahashi, 2002). Although some species within the bryophytes and primitive vascular plants are found to be high Si accumulators and carry NIP-IIIs, their mode of Si uptake remains largely unknown. The advent of conserved features such as the Ar/R SF and NPA motifs was shown to occur through stepwise changes over the course of plant evolution (Fig. 8). In the bryophytes, the first two positions of the Ar/R SF and the third position of both NPA motifs were less conserved. However, in clubmosses and ferns, only the first position of the Ar/R SF is less conserved while the second position became conserved. Similarly, both NPAs became more conserved. In general, there was a clear trend of selection for conserved sequences during each new era of plant evolution for the Ar/R SF and NPA motifs. As expected, bryophytes and primitive vascular plants displayed higher diversity at the Ar/R SF when compared with gymnosperms and angiosperms. The present study exploited the most comprehensive genomic resources to date in an attempt to explain the origin and evolution of Si uptake ability in plants. Phenotypic evaluations further showed that factors such as plant age, tissue type, plant physiology, and PAS could all significantly affect Si accumulation in plants and their classification as accumulators or non-accumulators. Our results further showed that monocots and dicots are not as distinctly separated in terms of Si uptake as previously reported. The taxonomical distribution provided in the present study will be helpful for several other disciplines such as palaeoecology and geology that define the biogeochemical cycling of Si. In addition to the prediction of Si uptake potential of plant species based on sequence information and taxonomical positioning, the evolutionary path of the Si uptake mechanism described here will be helpful to understand the Si environment over the different eras of land plant evolution. Results presented here clearly indicate that the evolution of NIP-IIIs as primary Si influx transporters dates back as early as 515 Ma. Supplementary data The following Supplementary data are available at JXB online. Table S1. List of 116 whole-genome sequenced plant species used for NIP-III identification. Table S2. Silicon (Si) accumulation observed in leaves of diverse plant species from different orders and familiess. Table S3. Silicon (Si) accumulation observed in the leaves of diverse plant species grown under greenhouse conditions with 1.7 mM Si supplementation. Table S4. Details of 140 NIP-IIIs identified in 81 whole-genome sequenced plant species. Table S5. Details of 349 NIP-IIIs identified using 1KP transcriptome data. Table S6. Details of NIP-IIIs with S-T-A-R Ar/R selectivity filters identified in Equisetales, ferns, and angiosperms. Fig. S1. Phylogenetic tree depicting three groups of nodulin 26-like intrinsic proteins identified by transcriptomic and genomic sequence data of 1133 plant species. Fig. S2. Evolution of the Ar/R selectivity filter (SF), NPA motifs, Froger’s residue, and other features in NIP-IIIs identified in plant lineages. Dataset S1. Sequences of known aquaporins from diverse plant species used as query sequences for BLAST search. Acknowledgements This work was supported by a grant from the Natural Sciences and Engineering Research Council of Canada (NSERC) and the Canada Research Chairs to RRB. We thank the Department of Biotechnology, Government of India for granting the Ramalingaswami Fellowship to RD and HS. We thank Mrs Claudette Roy for assistance with field experiments, and Mrs Caroline Labbé and Mr Maxime de Ronne for soil and leaf sample collection in Iowa fields. We are also grateful to Mr Jean Martin, Mrs Ariane Belzile, and Mrs Gowsica Ramakrishnan for their help in sample collection and silicon quantification. Author contributions RD and HS designed the study, conducted the experiments, and prepared the first draft of the manuscript. 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Kreszies,, Tino;Kreszies,, Victoria;Ly,, Falko;Thangamani, Priya, Dharshini;Shellakkutti,, Nandhini;Schreiber,, Lukas

doi: 10.1093/jxb/eraa203pmid: 32333766

Abstract Plant roots are the major organs that take up water and dissolved nutrients. It has been widely shown that apoplastic barriers such as Casparian bands and suberin lamellae in the endo- and exodermis of roots have an important effect on regulating radial water and nutrient transport. Furthermore, it has been described that silicon can promote plant growth and survival under different conditions. However, the potential effects of silicon on the formation and structure of apoplastic barriers are controversial. A delayed as well as an enhanced suberization of root apoplastic barriers with silicon has been described in the literature. Here we review the effects of silicon on the formation of suberized apoplastic barriers in roots, and present results of the effect of silicon treatment on the formation of endodermal suberized barriers on barley seminal roots under control conditions and when exposed to osmotic stress. Chemical analysis confirmed that osmotic stress enhanced barley root suberization. While a supplementation with silicon in both, control conditions and osmotic stress, did not enhanced barley root suberization. These results suggest that enhanced stress tolerance of plants after silicon treatment is due to other responses. Apoplast, barley, Casparian bands, root, silicon, suberin Introduction Silicon (Si) is one of the most abundant elements in the earth’s crust. It has been described as being beneficial for plants under various biotic and abiotic stresses (Epstein, 1999; Coskun et al., 2019). In the earth’s crust it is usually present in the form of silicate mineral as silicon dioxide (SiO2). When SiO2 is dissolved in water (approx. up to 2 mM) it forms orthosilic acid (Si(OH4)), which is accessible to plants. Silicon can then be taken up together with water by plant roots (Exley, 1998). The transport of water and nutrients, including Si, is described by the composite transport model, in which they are taken up via the apoplastic and the cell-to-cell (symplastic and transcellular) pathway. These pathways can be regulated by transport proteins in the plasma membrane or by suberin depositions in the endo- and exodermis of plant roots (Steudle and Peterson, 1998; Kim et al., 2018; Kreszies et al., 2018). Symplastic Si transporters that can facilitate Si uptake have been described in a wide range of plant species. These transporters belong to the aquaporin protein family or to an uncharacterized anion transporter protein family (Ma and Yamaji, 2015). Also the interaction of Si with several types of plant cell walls has been observed (Guerriero et al., 2016). Here we specifically focus on the effect of Si on specialized cell wall components: the apoplastic barriers, including Casparian bands and the suberin lamellae (Soukup and Tylová, 2018). Suberin is deposited in the form of Casparian bands made of lignin (Schreiber, 1996), suberin, and other cell wall components (Schreiber et al., 1999) in the primary cell walls of the endo- and exodermis of plant roots and as suberin lamellae on the inner surface of cell walls, separating them from the plasma membrane (Ranathunge et al., 2011). The suberin biopolyester is composed of an aromatic domain consisting mainly of phenolic acids and an aliphatic domain made of α,ω-bifunctional fatty acids, which are cross-linked via ester bonds (Graça, 2015). Enhanced formation of suberin lamellae has been described as a response to a wide range of biotic and abiotic stresses such as pathogens, osmotic, salt, and drought stress, and high nutrient concentrations (Ranathunge et al., 2011; Kreszies et al., 2018). As mentioned above, Si generally is described as a beneficial substance for plants under abiotic and biotic stresses. In contrast, recent reports of the effect of Si specifically on root apoplastic barriers are quite controversial. It was described that barrier formation was enhanced, unaffected, or delayed. Here we summarize the recent literature on the effects of Si on suberized root barriers and critically discuss these results, which were obtained by different experimental methodological approaches. Finally, we report our own results of the effect of Si on the development of the suberin lamellae in the endodermis of barley seminal roots during osmotic stress. Here we compare results obtained from histochemical detection via microscopy and from chemical analysis via gas chromatography–mass spectrometry (GC-MS). The effect of silicon on suberized apoplastic barriers Most of the studies describing the effect of Si on suberized barriers investigated the interaction between Si and different abiotic or biotic stress factors of plants. In all of the studies the formation of apoplastic barriers in roots with only Si treatment was investigated as well, and the results are quite inconsistent. While in some of these studies Si treatment enhanced the staining of suberin lamellae and suberin gene expression, in others it did not have an effect or even resulted in delayed formation of suberin lamellae (Table 1). By histochemical investigations staining with Fluorol Yellow 088, it was reported that Si treatment (i) enhanced the formation of the suberin lamellae in rice, maize, onion, wheat, canola, and Indian mustard (Fleck et al., 2011, 2015; Vatehová et al., 2012; Lukačová et al., 2013; Vulavala et al., 2016; Hinrichs et al., 2017; Wu et al., 2019), (ii) had no effect on the suberin development in maize and wheat (Vaculík et al., 2009; Wu et al., 2019), or even (iii) delayed the suberin development in maize and sorghum (Vaculík et al., 2012; Bathoova et al., 2018). These very different results between control grown plants and plants treated with Si illustrate that comparison and explanation of different experimental results is not straightforward. Reasons for these different results might be different growth conditions, such as Si concentration and hydroponics versus soil; plant age; or obvious anatomical, morphological, and genotypic differences between plant species. For the interpretation of experimental data, it is essential to be aware of the limitations (advantages and disadvantages) of the methods used. Typically, the above-mentioned studies used Fluorol Yellow 088, a widely accepted method for the detection of suberin lamellae. Fluorol Yellow 088 specifically stains lipids bright yellow and produces very high contrast images of suberin lamellae and other hydrophobic structures (Brundrett et al., 1991). The main advantage of using Fluorol Yellow 088 is that it is cheap and easy to handle, in contrast to chemical-analytical and metabolomic analyses. In addition, suberized tissues (e.g. endo- and exodermis) can be stained by Fluorol Yellow 088 and thereby anatomically identified by microscopy, which is not possible with chemical analysis alone. The disadvantages of using Fluorol Yellow 088 are that it is a qualitative method, as are all staining methods; it needs a certain threshold to bind and show a signal; and it rapidly fades when exposed to UV light—and thus different intensities cannot be quantified easily. Another disadvantage is that the exact binding mechanism of Fluorol Yellow 088 to hydrophobic fatty cell wall modifications is not completely understood, and it might also bind to other lipidic or aromatic cell wall deposits that are not necessarily part of the suberin lamellae. Moreover, it has been shown that Si aggregates in the endodermal cell walls of many plant species, such as rice and sorghum (Parry and Soni, 1972; Sangster and Parry, 1976; Soukup et al., 2017), where it might, but must not, interfere with the staining mechanism. Thus, an under- or overestimation of the suberin amounts, due to visual interpretation of microscopic pictures, is easily possible. Alternatively, a chemical analysis by GC-MS, which is more sensitive for the detection and quantification of suberin monomers, is helpful in parallel to microscopic studies to get a more detailed quantitative answer on suberization of apoplastic barriers in roots. For rice, maize, and onion it was, for example, described that Si enhanced the formation of the suberin lamellae and the Casparian band formation in the exodermis. This observation is based on histochemistry and microscopy. But chemical analyses of apoplastic barriers of onion, rice, and maize roots showed that amounts of aliphatic suberin monomers were unaffected (Fleck et al., 2015; Hinrichs et al., 2017). The enhanced formation of Casparian bands by Si treatment found by Fleck and colleagues (2015) can best be explained by the fact that they are mostly composed of lignin (Schreiber, 1996; Schreiber et al., 1999), and it was described that aromatics are more responsive towards biotic stress. For example, it has been described for Sorghum bicolor roots that infection with the fungus Alternaria alternata could be suppressed by Si treatment, and this is largely attributed to an enhanced deposition of phenolic components to the root exodermis (Bathoova et al., 2018). The potential enhancement of the aliphatic suberin domain by Si treatment should be more relevant in the case of drought stress. For example, in potato tubers an enhanced skin suberization was observed when plants were treated with Si in parallel to their exposure to drought stress (Vulavala et al., 2016). Table 1. Overview of studies investigating the effect of Si to apoplastic suberized barriers Species . Plant Age . Growth conditions . Treatments . Methods . Effect on the SL . Reference . Maize 10 d Hydroponics Cadmium 5 µM + Si 35 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si: less SL than with Cd only; Si alone had no effect Vaculík et al. (2009) Maize 10 d Hydroponics Cadmium 5 or 50 µM + Si 5 mM (sodium silicate solution) Phloroglucinol HCl, berberin hemisulphate, Fluorol Yellow 088 Cd 5 and 50 mM enhanced SL; Si alone led to further SL at root apex; Cd 5 mM + Si: less than Cd 5 mM; Cd 50 mM + Si: no effect compared with 50 mM Cd alone Vaculík et al. (2012) Maize 7 d Hydroponics Cadmium 5, 10, and 100 µM + Si 0.08 and 5 mM (sodium silicate solution) Phloroglucinol HCl, Fluorol Yellow 088 Si enhanced SL; Cd enhanced SL; Cd+Si enhanced SL too Lukačová et al. (2013) Maize 10 d Hydroponics Metalloid antimony (Sb) 5, 10, and 30 mg l−1 + Si 2.5 mM (sodium silicate solution) Fluorol Yellow 088 Sb enhanced SL; Si reduced the effect of 5 and 10 mg l−1 Sb, but not with 30 mg l−1 Sb Vaculíková et al. (2016) Maize 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice 28 d Hydroponics Si 1.78 mM (50 p.p.m. silica gel) Fluorol Yellow 088, microarray and qPCR Si enhanced SL and CBs; suberin genes are up-regulated Fleck et al. (2011) Rice 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice WT, OsABCG25 KO and OE 28 d Hydroponics Low Si 3 mg l−1 and high Si 30 mg l−1 (silica gel) Berberin hemisulphate, qPCR, GC Suberin genes are up-regulated in high Si compared with low Si; exodermis development enhanced; GC: no effect on suberin amounts Hinrichs et al. (2017) Wheat 14 d Hydroponics Cd 5 µM (short 6 h and long 7 d) + Si 1 mM (Na2SiO3) Fluorol Yellow 088, qPCR Short term: Si alone had no effect; Cd enhanced SL; Cd+Si delayed suberization compared with Cd alone; suberin genes were down-regulated in Si treatment. Long term: Si enhanced suberization; Cd alone had no effect; Cd+Si enhanced suberization; suberin genes were up-regulated in Si treatment Wu et al. (2019) Barley 12 d Hydroponics Si 1 mM (Na2SiO3) and osmotic stress (0.8 MPa) Fluorol Yellow 088, GC Si delayed suberization; osmotic stress enhanced suberization; GC: osmotic stress enhanced suberization; Si had no statistically significant effect This study Brassica napus (Canola) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Brassica juncea (Indian Mustard) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Onion 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB and SL formation; GC: no effect on suberin amounts Fleck et al. (2015) Potato (tuber skin) 8 weeks and 11 weeks Perlite Si 3.57 mM (sodium silicate solution) and drought (withholding water for 8 weeks) qPCR, Raman spectroscopy Si led to up-regulation of suberin genes; Si enhances skin suberization Vulavala et al. (2016) Sorghum bicolor 12 d Petri dishes (0.5 MS medium) Si 1 mM (sodium silicate solution) and inoculation with Alternaria alternata Fluorol Yellow 088, phosphoglucinol HCl, Raman spectroscopy In non-inoculated plants Si delayed suberization; in inoculated plants Si enhanced suberization; Raman spectroscopy showed higher phenolic components in Si treatments Bathoova et al. (2018) Species . Plant Age . Growth conditions . Treatments . Methods . Effect on the SL . Reference . Maize 10 d Hydroponics Cadmium 5 µM + Si 35 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si: less SL than with Cd only; Si alone had no effect Vaculík et al. (2009) Maize 10 d Hydroponics Cadmium 5 or 50 µM + Si 5 mM (sodium silicate solution) Phloroglucinol HCl, berberin hemisulphate, Fluorol Yellow 088 Cd 5 and 50 mM enhanced SL; Si alone led to further SL at root apex; Cd 5 mM + Si: less than Cd 5 mM; Cd 50 mM + Si: no effect compared with 50 mM Cd alone Vaculík et al. (2012) Maize 7 d Hydroponics Cadmium 5, 10, and 100 µM + Si 0.08 and 5 mM (sodium silicate solution) Phloroglucinol HCl, Fluorol Yellow 088 Si enhanced SL; Cd enhanced SL; Cd+Si enhanced SL too Lukačová et al. (2013) Maize 10 d Hydroponics Metalloid antimony (Sb) 5, 10, and 30 mg l−1 + Si 2.5 mM (sodium silicate solution) Fluorol Yellow 088 Sb enhanced SL; Si reduced the effect of 5 and 10 mg l−1 Sb, but not with 30 mg l−1 Sb Vaculíková et al. (2016) Maize 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice 28 d Hydroponics Si 1.78 mM (50 p.p.m. silica gel) Fluorol Yellow 088, microarray and qPCR Si enhanced SL and CBs; suberin genes are up-regulated Fleck et al. (2011) Rice 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice WT, OsABCG25 KO and OE 28 d Hydroponics Low Si 3 mg l−1 and high Si 30 mg l−1 (silica gel) Berberin hemisulphate, qPCR, GC Suberin genes are up-regulated in high Si compared with low Si; exodermis development enhanced; GC: no effect on suberin amounts Hinrichs et al. (2017) Wheat 14 d Hydroponics Cd 5 µM (short 6 h and long 7 d) + Si 1 mM (Na2SiO3) Fluorol Yellow 088, qPCR Short term: Si alone had no effect; Cd enhanced SL; Cd+Si delayed suberization compared with Cd alone; suberin genes were down-regulated in Si treatment. Long term: Si enhanced suberization; Cd alone had no effect; Cd+Si enhanced suberization; suberin genes were up-regulated in Si treatment Wu et al. (2019) Barley 12 d Hydroponics Si 1 mM (Na2SiO3) and osmotic stress (0.8 MPa) Fluorol Yellow 088, GC Si delayed suberization; osmotic stress enhanced suberization; GC: osmotic stress enhanced suberization; Si had no statistically significant effect This study Brassica napus (Canola) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Brassica juncea (Indian Mustard) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Onion 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB and SL formation; GC: no effect on suberin amounts Fleck et al. (2015) Potato (tuber skin) 8 weeks and 11 weeks Perlite Si 3.57 mM (sodium silicate solution) and drought (withholding water for 8 weeks) qPCR, Raman spectroscopy Si led to up-regulation of suberin genes; Si enhances skin suberization Vulavala et al. (2016) Sorghum bicolor 12 d Petri dishes (0.5 MS medium) Si 1 mM (sodium silicate solution) and inoculation with Alternaria alternata Fluorol Yellow 088, phosphoglucinol HCl, Raman spectroscopy In non-inoculated plants Si delayed suberization; in inoculated plants Si enhanced suberization; Raman spectroscopy showed higher phenolic components in Si treatments Bathoova et al. (2018) CB, Casparian band; Cd, cadmium; GC, gas chromatography; Si, silicon; SL, suberin lamellae. Open in new tab Table 1. Overview of studies investigating the effect of Si to apoplastic suberized barriers Species . Plant Age . Growth conditions . Treatments . Methods . Effect on the SL . Reference . Maize 10 d Hydroponics Cadmium 5 µM + Si 35 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si: less SL than with Cd only; Si alone had no effect Vaculík et al. (2009) Maize 10 d Hydroponics Cadmium 5 or 50 µM + Si 5 mM (sodium silicate solution) Phloroglucinol HCl, berberin hemisulphate, Fluorol Yellow 088 Cd 5 and 50 mM enhanced SL; Si alone led to further SL at root apex; Cd 5 mM + Si: less than Cd 5 mM; Cd 50 mM + Si: no effect compared with 50 mM Cd alone Vaculík et al. (2012) Maize 7 d Hydroponics Cadmium 5, 10, and 100 µM + Si 0.08 and 5 mM (sodium silicate solution) Phloroglucinol HCl, Fluorol Yellow 088 Si enhanced SL; Cd enhanced SL; Cd+Si enhanced SL too Lukačová et al. (2013) Maize 10 d Hydroponics Metalloid antimony (Sb) 5, 10, and 30 mg l−1 + Si 2.5 mM (sodium silicate solution) Fluorol Yellow 088 Sb enhanced SL; Si reduced the effect of 5 and 10 mg l−1 Sb, but not with 30 mg l−1 Sb Vaculíková et al. (2016) Maize 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice 28 d Hydroponics Si 1.78 mM (50 p.p.m. silica gel) Fluorol Yellow 088, microarray and qPCR Si enhanced SL and CBs; suberin genes are up-regulated Fleck et al. (2011) Rice 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice WT, OsABCG25 KO and OE 28 d Hydroponics Low Si 3 mg l−1 and high Si 30 mg l−1 (silica gel) Berberin hemisulphate, qPCR, GC Suberin genes are up-regulated in high Si compared with low Si; exodermis development enhanced; GC: no effect on suberin amounts Hinrichs et al. (2017) Wheat 14 d Hydroponics Cd 5 µM (short 6 h and long 7 d) + Si 1 mM (Na2SiO3) Fluorol Yellow 088, qPCR Short term: Si alone had no effect; Cd enhanced SL; Cd+Si delayed suberization compared with Cd alone; suberin genes were down-regulated in Si treatment. Long term: Si enhanced suberization; Cd alone had no effect; Cd+Si enhanced suberization; suberin genes were up-regulated in Si treatment Wu et al. (2019) Barley 12 d Hydroponics Si 1 mM (Na2SiO3) and osmotic stress (0.8 MPa) Fluorol Yellow 088, GC Si delayed suberization; osmotic stress enhanced suberization; GC: osmotic stress enhanced suberization; Si had no statistically significant effect This study Brassica napus (Canola) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Brassica juncea (Indian Mustard) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Onion 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB and SL formation; GC: no effect on suberin amounts Fleck et al. (2015) Potato (tuber skin) 8 weeks and 11 weeks Perlite Si 3.57 mM (sodium silicate solution) and drought (withholding water for 8 weeks) qPCR, Raman spectroscopy Si led to up-regulation of suberin genes; Si enhances skin suberization Vulavala et al. (2016) Sorghum bicolor 12 d Petri dishes (0.5 MS medium) Si 1 mM (sodium silicate solution) and inoculation with Alternaria alternata Fluorol Yellow 088, phosphoglucinol HCl, Raman spectroscopy In non-inoculated plants Si delayed suberization; in inoculated plants Si enhanced suberization; Raman spectroscopy showed higher phenolic components in Si treatments Bathoova et al. (2018) Species . Plant Age . Growth conditions . Treatments . Methods . Effect on the SL . Reference . Maize 10 d Hydroponics Cadmium 5 µM + Si 35 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si: less SL than with Cd only; Si alone had no effect Vaculík et al. (2009) Maize 10 d Hydroponics Cadmium 5 or 50 µM + Si 5 mM (sodium silicate solution) Phloroglucinol HCl, berberin hemisulphate, Fluorol Yellow 088 Cd 5 and 50 mM enhanced SL; Si alone led to further SL at root apex; Cd 5 mM + Si: less than Cd 5 mM; Cd 50 mM + Si: no effect compared with 50 mM Cd alone Vaculík et al. (2012) Maize 7 d Hydroponics Cadmium 5, 10, and 100 µM + Si 0.08 and 5 mM (sodium silicate solution) Phloroglucinol HCl, Fluorol Yellow 088 Si enhanced SL; Cd enhanced SL; Cd+Si enhanced SL too Lukačová et al. (2013) Maize 10 d Hydroponics Metalloid antimony (Sb) 5, 10, and 30 mg l−1 + Si 2.5 mM (sodium silicate solution) Fluorol Yellow 088 Sb enhanced SL; Si reduced the effect of 5 and 10 mg l−1 Sb, but not with 30 mg l−1 Sb Vaculíková et al. (2016) Maize 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice 28 d Hydroponics Si 1.78 mM (50 p.p.m. silica gel) Fluorol Yellow 088, microarray and qPCR Si enhanced SL and CBs; suberin genes are up-regulated Fleck et al. (2011) Rice 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB formation and SL; GC: aromatic suberin decreased Fleck et al. (2015) Rice WT, OsABCG25 KO and OE 28 d Hydroponics Low Si 3 mg l−1 and high Si 30 mg l−1 (silica gel) Berberin hemisulphate, qPCR, GC Suberin genes are up-regulated in high Si compared with low Si; exodermis development enhanced; GC: no effect on suberin amounts Hinrichs et al. (2017) Wheat 14 d Hydroponics Cd 5 µM (short 6 h and long 7 d) + Si 1 mM (Na2SiO3) Fluorol Yellow 088, qPCR Short term: Si alone had no effect; Cd enhanced SL; Cd+Si delayed suberization compared with Cd alone; suberin genes were down-regulated in Si treatment. Long term: Si enhanced suberization; Cd alone had no effect; Cd+Si enhanced suberization; suberin genes were up-regulated in Si treatment Wu et al. (2019) Barley 12 d Hydroponics Si 1 mM (Na2SiO3) and osmotic stress (0.8 MPa) Fluorol Yellow 088, GC Si delayed suberization; osmotic stress enhanced suberization; GC: osmotic stress enhanced suberization; Si had no statistically significant effect This study Brassica napus (Canola) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Brassica juncea (Indian Mustard) 7 d Hydroponics Cadmium 40 µM + Si 5 mM (sodium silicate solution) Fluorol Yellow 088 Cd enhanced SL; Cd+Si enhanced SL more; Si alone enhanced SL Vatehová et al. (2012) Onion 21 d Hydroponics Si 1.07 mM (silica gel) Berberin hemisulphate, Fluorol Yellow 088, GC Si enhanced CB and SL formation; GC: no effect on suberin amounts Fleck et al. (2015) Potato (tuber skin) 8 weeks and 11 weeks Perlite Si 3.57 mM (sodium silicate solution) and drought (withholding water for 8 weeks) qPCR, Raman spectroscopy Si led to up-regulation of suberin genes; Si enhances skin suberization Vulavala et al. (2016) Sorghum bicolor 12 d Petri dishes (0.5 MS medium) Si 1 mM (sodium silicate solution) and inoculation with Alternaria alternata Fluorol Yellow 088, phosphoglucinol HCl, Raman spectroscopy In non-inoculated plants Si delayed suberization; in inoculated plants Si enhanced suberization; Raman spectroscopy showed higher phenolic components in Si treatments Bathoova et al. (2018) CB, Casparian band; Cd, cadmium; GC, gas chromatography; Si, silicon; SL, suberin lamellae. Open in new tab Most of the Si studies in the literature with the main focus on apoplastic barriers have investigated the interaction of Si with abiotic stress caused by heavy metals, e.g. Cd (Table 1). Despite different experimental approaches and different Cd concentrations, a typical consistent reaction of plant roots to Cd stress was the enhancement of the suberin lamellae in order to reinforce the apoplastic barrier in roots and to prevent passive diffusional apoplastic Cd uptake (Table 1). However, as already mentioned above, a wide range of responses has been obtained by combining Si treatment with Cd stress. It looks like the effect of Si treatment during Cd stress depends on the plant species, the Cd concentrations and the plant developmental state. In maize, at low Cd concentrations of 5 mM, the formation of suberin lamellae was delayed in combined Cd and Si treatment, compared with Cd treatment only (Vaculík et al., 2009, 2012). On the other hand, at high Cd concentrations of 50 mM, Si treatment did not delay the suberin development in maize, compared with Cd treatment alone (Vaculík et al., 2012). A similar observation in maize roots was made by treatment with metallic antimony. Addition of Si reduced the suberin development at low concentrations and had no effect at higher concentrations (Vaculíková et al., 2016). In contrast to this, Lukačová and colleagues observed enhanced suberin lamellae in maize roots at much lower Cd concentrations in the micromolar range during Si treatment (Lukačová et al., 2013). Similar to this, in other plant species, such as canola and Indian mustard, additional Si enhanced the suberin lamellae compared with a single Cd treatment (Vatehová et al., 2012). However, it must kept in mind that all these studies used Fluorol Yellow 088 for the detection of suberin lamellae and thus might to some extent be affected by the above-describe advantages and disadvantages of this histochemical/microscopical method. However, it must also be considered that an enhanced or delayed development of suberin lamellae during abiotic stress, such as Cd exposure, can be advantageous as well as disadvantageous to the plant. Clearly, enhanced apoplastic barriers prevent the passive radial uptake of toxic Cd in the plant root. In parallel, an enhanced apoplastic barrier might also reduce the uptake of water and especially uptake of dissolved nutrients, which are crucial for plant survival. At low Cd concentrations it might be of advantage to have a reduced suberized barrier. With this scenario, plants would still gain the positive effects of higher nutrient uptake, even under the negative effect of a higher Cd uptake as long as they can deal with it. Contrary to that, at high or toxic Cd concentrations it might be essential to enhance the apoplastic barriers to efficiently prevent Cd uptake. The enhancement of apoplastic barriers represents a coarse long-term reaction of plant roots regulating radial transport (Kreszies et al., 2019), while the translational (e.g. gene expression) or post-translational (e.g. phosphorylation/dephosporylation) regulation of membrane transporters and aquaporins allows a short-term and fine regulation of radial root uptake of nutrients and water (Kaneko et al., 2015). In wheat a short-period treatment for 6 h with Cd and Si delayed suberization, compared with a treatment with Cd alone. However, long-term treatment for 7 d of wheat roots with Cd and Si enhanced the formation of suberin lamellae (Wu et al., 2019). This shows that for a short time interval, within hours, the plant can handle more Cd, while in the long term, within days, the accumulation of Cd is toxic. Material and methods Plant material and growth conditions Barley seeds (Hordeum vulgare cv. Scarlett) were stratified at 4 °C for 1 week. They were germinated for 3 d at 25 °C in the dark, covered with wet filter paper. The 3-day-old seedlings were then transferred into an aerated hydroponic system containing half-strength Hoagland nutrient solution, placed in a climatic chamber under long day conditions (16 h:8 h, light:dark), an air temperature of 23:20 °C (day:night) and a relative humidity of 50–65%. When the plants were 6 d old (3 d of germination and 3 d of growth) they were transferred to their respective treatment solutions for further 6 d until they were 12 d old. Four different treatments were tested: (i) control without Si, (ii) control with 1 mM Si (NaSiO3), (iii) osmotic stress −0.8 MPa without Si (the water potential of the nutrient solution was reduced by adding 25.5% (w/w) PEG8000), and (iv) osmotic stress −0.8 MPa with 1 mM Si. For all treatments the pH was adjusted to 6 using HCl to prevent precipitation of Si. Treatments without Si application were supplied with NaCl to compensate for the potential effect of Na+ ions supplied by NaSiO3. Histochemical detection of Casparian bands and suberin lamellae in roots Seminal root cross-sections were made using a cryostat microtome (Microm HM 500M, Microm International, Walldorf, Germany). Casparian bands were stained with 0.1% (w/v) berberine hemisulfate for 1 h and 0.5% (w/v) aniline blue for 30 min (Brundrett et al., 1988). Suberin lamellae were stained with 0.01% (w/v) lipophilic Fluorol Yellow 088 for 1 h (Brundrett et al., 1991). Cross-sections were studied by epifluorescence microscopy using an ultraviolet (UV) filter set (excitation filter BP 365, dichroic mirror FT 395, barrier filter LP 397; Zeiss, Oberkochen, Germany). Pictures were taken with a Canon EOS 600D camera at ISO 200 or 400 with an exposure time for 1 s to 2 s. Chemical analysis of barley root suberin The 12-day-old barley seminal roots were divided into three zones, A, B, and C, based on the previous microscopic investigations as described earlier (Kreszies et al., 2019, 2020). The youngest part of the root (0–25% of total root length), including the root apex, was designated zone A. This zone characteristically has only a few Casparian bands in the endodermis and no microscopically detectable suberin lamellae. The middle part of the root (25–50% of total root length) was assigned to the transition zone or zone B. Here all endodermal cells have Casparian bands and some cells are suberized. In the oldest part of the root (50–100% of total root length), designated zone C, all endodermal cells have Casparian bands as well as a suberin lamellae. For one replicate, 10 segments, from each of the three root zones, were pooled. Next, the root segments were enzymatically digested with 0.5% (w/v) cellulase and 0.5% (w/v) pectinase for 3 weeks at room temperature under continuous shaking (Zeier and Schreiber, 1997). Within these 3 weeks the enzyme solution was replaced about four times. Prior to the continuous shaking of the roots when immersed in the enzyme solution, they were vacuum infiltrated. Afterwards the remaining isolated cell walls were washed in borate buffer and transferred to 1:1 (v/v) chloroform:methanol to extract soluble lipids, at room temperature under continuous shaking for another 2 weeks. The chloroform:methanol solution was replaced four times within the 2 weeks. In the final step the root samples were dried on polytetrafluoroethylene in a desiccator, containing activated silica gel. The dried samples were subjected to transesterification with BF3–methanol to release suberin monomers (Kolattukudy and Agrawal, 1974). Further GC-MS sample preparation was performed as described previously (Zeier and Schreiber, 1997, 1998). A 1 μl aliquot of the sample dissolved in chloroform was injected in splitless mode at injector temperature of 250 °C. The column oven temperature was held at 50 °C for 1 min and then increased by 25 °C per minute from 50 to 200 °C, followed by a 1 min hold. Finally the temperature was increased 10 °C per minute to a final temperature of 320 °C, which was held for 8 min (Delude et al., 2017). Suberin amounts were referenced to the endodermal surface area. The endodermal area was calculated for each root zone with: A=2πrL, where r is the endodermis radius and L is the length of the individual root zone. Three biological replicates were used for each experiment. Statistical analysis Data analysis and statistical tests were performed with Origin Pro 9. A normal distribution of the data was tested with the Shapiro–Wilk test. Significant differences between means were tested with one-way ANOVA (Fisher’s LSD) at a significance level of 0.05. Results Root morphology and anatomy The mean seminal root length decreased in response to osmotic stress of −0.8 MPa. Under control conditions the plants with applied Si had marginally longer seminal roots, while at osmotic stress conditions there was no effect on the seminal root length with and without Si application (Fig. 1). In roots grown under control conditions there was no suberin lamellae detected in the younger root zone up to 25% of the relative root length (Fig. 2A), while at 50% most of the endodermis cells were suberized with only a few passage cells left (Fig. 2E). In the basal part of the root at 90% of the relative root length, all endodermis cells were suberized (Fig. 2I). A similar suberin development pattern was observed when Si was applied (Fig. 2B, F, J). For plants grown under osmotic stress conditions of −0.8 MPa without Si application an enhanced formation of suberin lamellae was observed in all root zones compared with the control treatments, especially at 25% and 50% of the relative root length (Fig. 2C, G, K). In contrast, when seminal roots were grown under osmotic stress with additional Si application, they showed a tendency to have a reduced number of suberized cells in the endodermis at 25% and 50%, compared with osmotic stress conditions without additional Si application. (Fig. 2D, H, L). Under all conditions tested, we detected no exodermis in our hydroponic growth system. Fig. 1. Open in new tabDownload slide Root lengths of 12-day-old barley plants grown under control conditions or at a water potential of −0.8 MPa induced with PEG8000, with or without external silicon application. The bars represent the mean values with standard deviation; n=40 seminal roots. Different letters indicate significant differences between means at a significance level of 0.05 by one-way ANOVA (Fisher’s LSD test). Fig. 1. Open in new tabDownload slide Root lengths of 12-day-old barley plants grown under control conditions or at a water potential of −0.8 MPa induced with PEG8000, with or without external silicon application. The bars represent the mean values with standard deviation; n=40 seminal roots. Different letters indicate significant differences between means at a significance level of 0.05 by one-way ANOVA (Fisher’s LSD test). Fig. 2. Open in new tabDownload slide Development of the suberin lamellae in the endodermis of barley seminal roots grown under control conditions or at a water potential of −0.8 MPa induced with PEG8000, with or without external Si application. All cross-sections were stained with Fluorol Yellow 088. The presence of suberin lamellae is indicated by a bright yellow fluorescence. Scale bars: 50 µm. Fig. 2. Open in new tabDownload slide Development of the suberin lamellae in the endodermis of barley seminal roots grown under control conditions or at a water potential of −0.8 MPa induced with PEG8000, with or without external Si application. All cross-sections were stained with Fluorol Yellow 088. The presence of suberin lamellae is indicated by a bright yellow fluorescence. Scale bars: 50 µm. Chemical analysis of suberin in response to osmotic stress and silicon treatment In the chemical analysis the quantitative and qualitative suberin amounts in all three root zones (A, 0–25% of the total root length; B, 25–50% of the total root length; and C, 50–100% of the total root length) were measured. In all zones chain lengths of the aliphatic suberin monomers ranged from C16 to C26 with C18:1 α,ω-dicarboxylic acid and C18:1 and C24 ω-hydroxy acids being the most prominent monomers. There was no significant difference in single monomer composition between control and osmotic stress as well as with or without Si application (see Supplementary Fig. S1 at JXB online). In total aliphatic suberin amounts, there was a pronounced increase over the length of the root from the tip (zone A) to the basal part (zone C), reflecting the root development over the length. In zone A no significant differences in total aliphatic suberin amounts between the different treatments were detectable. In zone B the osmotic stress significantly enhanced the suberin lamellae between control and stress of −0.8 MPa. However, there was no difference detectable between Si treatments in zone B. In the oldest part, zone C, the osmotic stress further increased the total suberin amount. Similar to zone B, in zone C there were no significant differences detectable between the Si treatments (Fig. 3). Fig. 3. Open in new tabDownload slide Total amount of aliphatic suberin in barley seminal roots grown under control conditions or at osmotic stress of −0.8 MPa induced with PEG8000, with and without Si application. The roots were divided into the three root zones: A, 0–25% of the total root length, from the root tip; B, 25–50% of the total root length, the transition zone; and C, 50–100% of the total root length, the basal part. The bars represent mean values with standard deviation of three independent biological replicates (n=3); each biological replicate was obtained from 10 individual root segments. Different letters indicate significant differences between means at a significance level of 0.05 by one-way ANOVA (Fisher’s LSD test). Fig. 3. Open in new tabDownload slide Total amount of aliphatic suberin in barley seminal roots grown under control conditions or at osmotic stress of −0.8 MPa induced with PEG8000, with and without Si application. The roots were divided into the three root zones: A, 0–25% of the total root length, from the root tip; B, 25–50% of the total root length, the transition zone; and C, 50–100% of the total root length, the basal part. The bars represent mean values with standard deviation of three independent biological replicates (n=3); each biological replicate was obtained from 10 individual root segments. Different letters indicate significant differences between means at a significance level of 0.05 by one-way ANOVA (Fisher’s LSD test). Discussion and conclusion Suberized apoplastic barriers can (i) control the radial fluxes of water and nutrients, (ii) prevent uptake of toxicants, and (iii) protect against microbiological aggression (Ranathunge et al., 2011). In the Introduction we reviewed the effect of Si, which is accepted as a beneficial element for plants especially under biotic and abiotic stress (Coskun et al., 2019), on suberized apoplastic barriers (Table 1). In the Results, we expanded the previous data by our own experiments on barley seminal roots, dealing with a water deficit induced by the osmoticum PEG8000. Our histochemical analyses staining Casparian bands with berberine–aniline blue did not show any differences between our treatments (data not shown). This is similar to what we have observed with the same barley cultivar earlier under osmotic stress (Kreszies et al., 2019). Staining with Fluorol Yellow 088 along the root length showed a gradual increase of the suberin lamellae according to root development. In our experiments with barley seminal roots, no differences were detectable when additional Si was supplied to the roots in the control nutrient solution (Fig. 2). Similar results have been obtained in other Gramineae species such as maize and wheat (Vaculík et al., 2009; Wu et al., 2019). Direct quantification of suberin by GC-MS confirmed the results obtained by histochemistry for control plants with and without Si treatment. Indeed, there was a developmental gradient along the root length and there were no prominent differences in the suberin amount (Fig. 3). After applying osmotic stress to the plant roots there was a significant increase in aliphatic suberin. The aliphatic suberin domain is the main hydrophobic barrier against uncontrolled passive radial transport of water and solutes during water deficit, especially in the transition zone B, as we have described earlier under the same experimental conditions, with the same and different modern barley cultivars (Kreszies et al., 2019, 2020). Aliphatic suberin of endodermal cell walls quantified via GC-MS was not significantly altered, neither enhanced nor delayed, when additional Si was applied. These chemical-analytical data fit with the results that have been obtained in rice, maize, and onion (Fleck et al., 2015; Hinrichs et al., 2017). Furthermore Si did not change the suberin monomer composition between the treatments, and it was the same as measured before with the same barley cultivar (Kreszies et al., 2019). However, in fluorescence microscopy with Fluorol Yellow 088 staining, the formation of the endodermal suberin lamellae seemed to be delayed in the presence of Si under osmotic stress conditions. This might be a consequence of the interaction of Si with the endodermal cell walls, possibly, for example, through silica aggregates, which could interfere with the binding intensity of Fluorol Yellow 088. In summary, these results obtained from barley seminal root analyses are in line with the results presented in the literature (Table 1). By histochemistry and microscopy we obtained a delayed suberin formation in response to Si treatment, whereas results obtained by analytical chemistry showed no difference in suberin development when Si was applied. In principle, it seems that plant species, morphology, age, experimental design (e.g. stress concentrations or short versus long term), and how Si is applied play an important role in the formation of suberized apoplastic barriers. With microscopy alone it is very difficult to correctly quantify suberin amounts, while gas chromatography alone does not provide important information on anatomy and morphology. Therefore, it is necessary to combine omics technologies with plant physiology and microscopic approaches to understand plant responses to Si and draw the correct conclusions. Since it was shown in the past that osmotic stress induces root suberization, which enhances stress tolerance of cultivated barley (Kreszies et al., 2019, 2020), we initially hypothesized that Si could mediate such an induced root suberization directly, leading to enhanced stress tolerance even without an endogenous stress signal. However, since our chemical analysis clearly showed that there was no difference in suberization, we conclude that an enhanced stress tolerance of plants described as occurring when there is Si supplementation must be due to other effects. These effects could be regulation of symplastic transport, instead of apoplastic transport, as this has been described for Sorghum bicolor and tomato, in which Si enhanced aquaporin activity during osmotic stress induced by PEG6000 or NaCl (Liu et al., 2014, 2015; Shi et al., 2016), or the enhancement of osmotic adjustment in the presence of Si (Rizwan et al., 2015; Turner, 2018). Supplementary data Supplementary data are available at JXB online. Fig. S1. Aliphatic suberin monomer amounts in zone C (50–100% root length) of barley seminal roots grown under control conditions or at a water potential of −0.8 MPa induced with PEG8000, with or without external silicon application. Acknowledgements Financial support by the Deutsche Forschungsgemeinschaft (DFG) to LS is highly appreciated. This manuscript is dedicated to Ole. References Bathoova M , Bokor B, Soukup M, Lux A, Martinka M. 2018 . Silicon-mediated cell wall modifications of sorghum root exodermis and suppression of invasion by fungus Alternaria alternata . Plant Pathology 67 , 1891 – 1900 . Google Scholar Crossref Search ADS WorldCat Brundrett MC , Enstone DE, Peterson CA. 1988 . A berberine-aniline blue fluorescent staining procedure for suberin, lignin, and callose in plant tissue . Protoplasma 146 , 133 – 142 . Google Scholar Crossref Search ADS WorldCat Brundrett MC , Kendrick B, Peterson CA. 1991 . 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Soukup, Milan; Rodriguez Zancajo, Victor M; Kneipp, Janina; Elbaum, Rivka

doi: 10.1093/jxb/erz387pmid: 31504726

Abstract Silica deposition in plants is a common phenomenon that correlates with plant tolerance to various stresses. Deposition occurs mostly in cell walls, but its mechanism is unclear. Here we show that metabolic processes control the formation of silica aggregates in roots of sorghum (Sorghum bicolor L.), a model plant for silicification. Silica formation was followed in intact roots and root segments of seedlings. Root segments were treated to enhance or suppress cell wall biosynthesis. The composition of endodermal cell walls was analysed by Raman microspectroscopy, scanning electron microscopy and energy-dispersive X-ray analysis. Our results were compared with in vitro reactions simulating lignin and silica polymerization. Silica aggregates formed only in live endodermal cells that were metabolically active. Silicic acid was deposited in vitro as silica onto freshly polymerized coniferyl alcohol, simulating G-lignin, but not onto coniferyl alcohol or ferulic acid monomers. Our results show that root silica aggregates form under tight regulation by endodermal cells, independently of the transpiration stream. We raise the hypothesis that the location and extent of silicification are primed by the chemistry and structure of polymerizing lignin as it cross-links to the wall. Cell wall, lignin, root endodermis, SEM–EDX, silica, Sorghum bicolor (L.) Moench Introduction Amorphous hydrated silica (SiO2·nH2O, herein silica) is one of the most common biominerals in the plant kingdom (Bauer et al., 2011; He et al., 2014; Coskun et al., 2019). Its deposition in plant cell walls, cell lumen, and intercellular spaces modifies the properties of plant tissues and supports their resistance against various biotic and abiotic stresses (Currie and Perry, 2007; Yamanaka et al., 2009; Liu et al., 2013; He et al., 2015; Ma et al., 2015). The chemical form of Si available for plant absorption is mono-silicic acid, Si(OH)4. Silicic acid is taken up by roots either passively or actively (Ma and Yamaji, 2015), distributed to target locations in the plant, and mineralized into silica. The silicified structures are often referred to as silica phytoliths (Bauer et al., 2011). Although silicification is a well-known phenomenon, the mineralization process and its regulation are unknown (Guerriero et al., 2016; Kumar et al., 2017b). Many studies indicate interactions between the deposition of silica and lignin/phenolic compounds (Inanaga and Okasaka, 1995; Fang and Ma, 2006; Schoelynck et al., 2010; Fleck et al., 2011, 2015; Suzuki et al., 2012; Yamamoto et al., 2012; Zhang et al., 2013; Hinrichs et al., 2017; Klotzbücher et al., 2018). Furthermore, silica deposits are frequently associated with lignified tissues (Scurfield et al., 1974; Piperno et al., 2002; Zhang et al., 2013). However, a direct link between plant tissue silicification and lignification is missing. In lignifying plant cell walls, a variety of phenolic compounds participate in radical coupling reactions and incorporate into the growing lignin polymer. The fundamental subunits to form lignin are coniferyl alcohol (CA), p-coumaryl alcohol, and sinapyl alcohol, producing the G-, H- and S-subunits of the lignin polymer, respectively. The proportions of the lignin subunits vary between species, with G-lignin as the most abundant in grasses (Barros et al., 2015). Ferulic acid (FA) is not one of the three canonical monolignols. Nonetheless, ferulates are a natural component of lignins in grasses and could be considered a monolignol as well (Ralph et al., 2008). In addition, FA cross-links the non-lignified cell walls via FA–arabinoxylan (AX) complexes. In the roots of sorghum (Sorghum bicolor (L.) Moench; Poaceae), silica is deposited in large quantities as discrete silica aggregates, anchored within the inner tangential cell walls (ITCWs) of the root endodermis (Sangster and Parry, 1976a,b,c; Hodson and Sangster, 1989; Lux et al., 2002, 2003). Our previous work shows that the silica aggregates are associated with the thickened cell walls, which appear only after the ITCW lignification initiates, and contain traces of the FA-AX complexes (Soukup et al., 2017). These observations indicate a possible interconnection between the deposition of silica and phenolic compounds. Thus, in this study we aimed to elucidate the relationship between silica aggregate formation, cell wall deposition, and lignification in the root endodermal cell walls. Material and methods Plant material Grains of Sorghum bicolor (L.) Moench, line BTx623, were surface-sterilized with 2.5% sodium hypochlorite for 15 min, washed with distilled water, and imbibed in distilled water for 24 h. Grains were then placed into wet paper rolls and germinated for 72 h. Hydroponic cultivation (also referred to as in planta cultivation) After germination, the seedlings were grown hydroponically for 6 d. The first 3 d they grew in distilled water supplied with either sodium silicate (Na2O(SiO2)x·xH2O) at final concentration 1 mM (Si+ medium) or NaCl at final concentration 1 mM (Si− medium) to preserve similar ionic balance of the media. Final pH of the solutions was adjusted to 5.8 with HCl. After 3 d of cultivation, the growth solutions were changed and the seedlings were grown for additional 3 d to obtain the following growth conditions: (i) seedlings grown in Si− medium throughout the entire cultivation (Si− treatment), (ii) seedlings grown in Si+ medium throughout the entire cultivation (Si+ treatment), (iii) seedling grown for 3 d in Si− medium and for 3 d in Si+ medium (Si−/Si+ treatment), or (iv) 3 d in Si+ medium and 3 d in Si− medium (Si+/Si− treatment). The cultivations were performed in a growth chamber, under controlled conditions with photoperiod 16 h–8 h (light–dark) illuminated with photosynthetically active radiation (PAR) of approximately 200 μmol m−2 s−1, 28 °C–22 °C (light–dark) temperature, and 70% air humidity. Root samples were then collected from primary root region 1–3.5 cm from the primary root–shoot junction. Ex planta cultivation of detached primary root segments (based onSoukup et al., 2017) Silica is a common contaminator of many salts, because of its omnipresence. To avoid exposure to silicic acid we chose to grow our samples in a minimal medium, taking into account the important nutritional supply of the seed. Therefore, after germination, seedlings were precultivated hydroponically in distilled water for 72 h under Si− conditions as described above for the hydroponic cultivations. Afterwards, seedlings of similar morphology, with emerged lateral roots and primary root length exceeding 6 cm were selected and used for root segment preparation. Segments were collected from the primary root region 1–3.5 cm from the root–shoot junction. Lateral roots emerging from the segments were cut off. Some segments were pretreated before the ex planta cultivation (Table 1). Prepared segments were placed in Erlenmeyer flasks containing 50 ml of the cultivation medium (Table 1). The pH of all cultivation media containing sodium silicate was adjusted to 5.8 before adding other components. The flasks were sealed with Parafilm and cultured for 72 h (unless stated otherwise) in the dark at 24 °C, with permanent shaking at 60 rpm. After cultivation, the segments were fixed in FAA solution (3.7% formaldehyde, 50% ethanol, 5% glacial acetic acid, 41.3% distilled water; v/v). Table 1. The pretreatments and treatments applied to detached sorghum root segments cultivated ex planta Treatment ID . Segment pretreatment . Compositiona . Si+ 1 mMb Sodium silicate 1 mmol dm−3 Si+ 2 mM Sodium silicate 2 mmol dm−3 Si+ 2 mM sucrose Sodium silicate 2 mmol dm−3, sucrose 1% (w/v) Si− NaCl 1 mmol dm−3 Removed cortex Rhizodermis and cortical tissues peeled-off mechanically Sodium silicate 1 mmol dm−3 Liquid nitrogen Segments kept in liquid nitrogen for 5 min Sodium silicate 1 mmol dm−3 DNPc Sodium silicate 1 mmol dm−3, 2,4-dinitrophenol 0.01 or 0.05 mmol dm−3 Low temperaturec Erlenmeyer flask with the segments cultivated in ice-filled Styrofoam box (temperature 2–4 °C) Sodium silicate 1 mmol dm−3 Sucroseb Sodium silicate 1 mmol dm−3, Sucrose 1% (w/v) Arabinoseb Sodium silicate 1 mmol dm−3, arabinose 1% (w/v) ATP Sodium silicate 1 mmol dm−3, ATP 1 or 5 mmol dm−3 Brefeldin Ab Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v), brefeldin A 1, 5 or 10 µmol dm−3 SHAMb Sodium silicate 1 mmol dm−3, salicylhydroxamic acid 0.2 or 2 mmol dm−3 KIb Sodium silicate 1 mmol dm−3, potassium iodide 0.5 or 5 mmol dm−3 Ascorbic acidb Sodium silicate 1 mmol dm−3, ascorbic acid 1 or 5 mmol dm−3 Ferulic acidb Sodium silicate 1 mmol dm−3, ferulic acid 0.05 or 0.2 mmol dm−3 H2O2b Sodium silicate 1 mmol dm−3, hydrogen peroxide 1 mmol dm−3 H2O2 pretreatment Segments cultivated in distilled water supplied with 5 mmol dm−3 H2O2 for 8h (dark, constant 60 rpm shaking) Sodium silicate 1 mmol dm−3 Ethanolb Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v) Treatment ID . Segment pretreatment . Compositiona . Si+ 1 mMb Sodium silicate 1 mmol dm−3 Si+ 2 mM Sodium silicate 2 mmol dm−3 Si+ 2 mM sucrose Sodium silicate 2 mmol dm−3, sucrose 1% (w/v) Si− NaCl 1 mmol dm−3 Removed cortex Rhizodermis and cortical tissues peeled-off mechanically Sodium silicate 1 mmol dm−3 Liquid nitrogen Segments kept in liquid nitrogen for 5 min Sodium silicate 1 mmol dm−3 DNPc Sodium silicate 1 mmol dm−3, 2,4-dinitrophenol 0.01 or 0.05 mmol dm−3 Low temperaturec Erlenmeyer flask with the segments cultivated in ice-filled Styrofoam box (temperature 2–4 °C) Sodium silicate 1 mmol dm−3 Sucroseb Sodium silicate 1 mmol dm−3, Sucrose 1% (w/v) Arabinoseb Sodium silicate 1 mmol dm−3, arabinose 1% (w/v) ATP Sodium silicate 1 mmol dm−3, ATP 1 or 5 mmol dm−3 Brefeldin Ab Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v), brefeldin A 1, 5 or 10 µmol dm−3 SHAMb Sodium silicate 1 mmol dm−3, salicylhydroxamic acid 0.2 or 2 mmol dm−3 KIb Sodium silicate 1 mmol dm−3, potassium iodide 0.5 or 5 mmol dm−3 Ascorbic acidb Sodium silicate 1 mmol dm−3, ascorbic acid 1 or 5 mmol dm−3 Ferulic acidb Sodium silicate 1 mmol dm−3, ferulic acid 0.05 or 0.2 mmol dm−3 H2O2b Sodium silicate 1 mmol dm−3, hydrogen peroxide 1 mmol dm−3 H2O2 pretreatment Segments cultivated in distilled water supplied with 5 mmol dm−3 H2O2 for 8h (dark, constant 60 rpm shaking) Sodium silicate 1 mmol dm−3 Ethanolb Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v) a Final concentration of the substance in distilled water. b Treatments analysed by Raman spectroscopy and SEM-EDX for modelling the contributions of cell wall components to silica deposition. c After 4 h cultivation the segments were either collected immediately, or they were rinsed in distilled water for 10 min and cultured for additional 68 h in Si+ medium. Open in new tab Table 1. The pretreatments and treatments applied to detached sorghum root segments cultivated ex planta Treatment ID . Segment pretreatment . Compositiona . Si+ 1 mMb Sodium silicate 1 mmol dm−3 Si+ 2 mM Sodium silicate 2 mmol dm−3 Si+ 2 mM sucrose Sodium silicate 2 mmol dm−3, sucrose 1% (w/v) Si− NaCl 1 mmol dm−3 Removed cortex Rhizodermis and cortical tissues peeled-off mechanically Sodium silicate 1 mmol dm−3 Liquid nitrogen Segments kept in liquid nitrogen for 5 min Sodium silicate 1 mmol dm−3 DNPc Sodium silicate 1 mmol dm−3, 2,4-dinitrophenol 0.01 or 0.05 mmol dm−3 Low temperaturec Erlenmeyer flask with the segments cultivated in ice-filled Styrofoam box (temperature 2–4 °C) Sodium silicate 1 mmol dm−3 Sucroseb Sodium silicate 1 mmol dm−3, Sucrose 1% (w/v) Arabinoseb Sodium silicate 1 mmol dm−3, arabinose 1% (w/v) ATP Sodium silicate 1 mmol dm−3, ATP 1 or 5 mmol dm−3 Brefeldin Ab Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v), brefeldin A 1, 5 or 10 µmol dm−3 SHAMb Sodium silicate 1 mmol dm−3, salicylhydroxamic acid 0.2 or 2 mmol dm−3 KIb Sodium silicate 1 mmol dm−3, potassium iodide 0.5 or 5 mmol dm−3 Ascorbic acidb Sodium silicate 1 mmol dm−3, ascorbic acid 1 or 5 mmol dm−3 Ferulic acidb Sodium silicate 1 mmol dm−3, ferulic acid 0.05 or 0.2 mmol dm−3 H2O2b Sodium silicate 1 mmol dm−3, hydrogen peroxide 1 mmol dm−3 H2O2 pretreatment Segments cultivated in distilled water supplied with 5 mmol dm−3 H2O2 for 8h (dark, constant 60 rpm shaking) Sodium silicate 1 mmol dm−3 Ethanolb Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v) Treatment ID . Segment pretreatment . Compositiona . Si+ 1 mMb Sodium silicate 1 mmol dm−3 Si+ 2 mM Sodium silicate 2 mmol dm−3 Si+ 2 mM sucrose Sodium silicate 2 mmol dm−3, sucrose 1% (w/v) Si− NaCl 1 mmol dm−3 Removed cortex Rhizodermis and cortical tissues peeled-off mechanically Sodium silicate 1 mmol dm−3 Liquid nitrogen Segments kept in liquid nitrogen for 5 min Sodium silicate 1 mmol dm−3 DNPc Sodium silicate 1 mmol dm−3, 2,4-dinitrophenol 0.01 or 0.05 mmol dm−3 Low temperaturec Erlenmeyer flask with the segments cultivated in ice-filled Styrofoam box (temperature 2–4 °C) Sodium silicate 1 mmol dm−3 Sucroseb Sodium silicate 1 mmol dm−3, Sucrose 1% (w/v) Arabinoseb Sodium silicate 1 mmol dm−3, arabinose 1% (w/v) ATP Sodium silicate 1 mmol dm−3, ATP 1 or 5 mmol dm−3 Brefeldin Ab Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v), brefeldin A 1, 5 or 10 µmol dm−3 SHAMb Sodium silicate 1 mmol dm−3, salicylhydroxamic acid 0.2 or 2 mmol dm−3 KIb Sodium silicate 1 mmol dm−3, potassium iodide 0.5 or 5 mmol dm−3 Ascorbic acidb Sodium silicate 1 mmol dm−3, ascorbic acid 1 or 5 mmol dm−3 Ferulic acidb Sodium silicate 1 mmol dm−3, ferulic acid 0.05 or 0.2 mmol dm−3 H2O2b Sodium silicate 1 mmol dm−3, hydrogen peroxide 1 mmol dm−3 H2O2 pretreatment Segments cultivated in distilled water supplied with 5 mmol dm−3 H2O2 for 8h (dark, constant 60 rpm shaking) Sodium silicate 1 mmol dm−3 Ethanolb Sodium silicate 1 mmol dm−3, 0.01% ethanol (v/v) a Final concentration of the substance in distilled water. b Treatments analysed by Raman spectroscopy and SEM-EDX for modelling the contributions of cell wall components to silica deposition. c After 4 h cultivation the segments were either collected immediately, or they were rinsed in distilled water for 10 min and cultured for additional 68 h in Si+ medium. Open in new tab Raman microspectroscopy Cross-sections were prepared from the root segments fixed in FAA either before the ex planta cultivation, after the cultivation, or at both time points. Several cross-sections from each root segment were placed on microscope slides, washed three times and mounted in distilled water, covered with coverslips, and sealed with nail polish to avoid water evaporation. Raman spectra were collected with an InVia spectrometer (Renishaw, UK) equipped with 532 nm laser, utilizing WIRE3.2 software (Renishaw, UK). Measurements were performed with 100 kW cm−2 laser intensity, 0.1 s acquisition time and 150 accumulations per spectrum. At least five spectra from each root were collected, using at least three different roots per treatment. Collected spectra were smoothed (Savitzky–Golay algorithm, 9-point interval, polynomial order 4) and baseline corrected (adaptive baseline correction, coarseness 10%) using Spectragryph 1.0.7 (F. Menges ‘Spectragryph – optical spectroscopy software,’ Version 1.0.7, 2017, http://www.effemm2.de/spectragryph/). Scanning electron microscopy and energy dispersive X-ray analysis From each root segment fixed in FAA, we collected several hand cross-sections and a 0.5 cm-long piece, from which rhizodermis and outer cortical tissues were peeled-off (referred to as peeled-off segments; Lux et al., 2002). The samples were stuck onto a carbon tape and placed on a metal stand. Observations and energy dispersive X-ray spectroscopy (EDX) analyses were performed with a scanning electron microscope (JSM-IT 100 InTouchScopeTM; JEOL, Japan) under low vacuum (30 Pa), with accelerating voltage of 20 kV. Images were collected in back-scattered electron imaging mode. The EDX analyses were acquired from the peeled-off segments as point measurements with 20 s acquisition time per aggregate location, obtained from minimally 50 endodermal cells per treatment, from at least three different roots. Based on the EDX analyses the Si:C mass ratios were calculated. Raman–EDX data analysis Raman and EDX data collected in the ex planta experiments were used to estimate the contributions of cell wall components to silica deposition. Raman analysis is described in detail in Supplementary Dataset S1 at JXB online. Considering all the Raman spectra collected, peak positions (p) were identified (np=30; for assignments see Supplementary Dataset S1). Intensities of all the identified peaks were then extracted from the 72 h Si+ ex planta treatments to create a matrix of independent variables (the treatments used are indicated in Table 1, footnote b). For each variable, a mean value was calculated from the Si+ 1 mM treatment and used as a reference value. All observations in all treatments were afterwards expressed relative to corresponding reference values. For each treatment, a median of Si:C ratios (estimated based on EDX analyses) was then assigned as a dependent variable. The matrix of these relative peak intensities (independent variables) and relative Si:C ratios (dependent variable) was analysed using multiple linear regression with the weighted least squares estimation method. The number of independent variables was afterwards iteratively reduced using the backward elimination procedure, with maximal significance level for keeping a variable in the model being 0.005. The analyses and their visual representations were performed using the Python programming language (version 3.5, Python Software Foundation, https://www.python.org/). The Si:C ratios were plotted using box plots showing interquartile ranges (IQR) with 95% confidence intervals indicated as notches and Tukey style whiskers (1.5×IQR) (Krzywinski and Altman, 2014). Identified outliers outside the 1.5×IQR whiskers are not displayed in the plotted data. In vitro lignin synthesis by oxidative coupling and infrared spectroscopic analysis An oxidative coupling reaction was carried out in several different substrate conditions: (i) CA alone, (ii) a 10/90 mixture of FA/CA, (iii) a 50/50 mixture of FA/CA, (iv) a 10/90 mixture of AX/CA, and (v) a 10/10/80 mixture of AX/FA/CA. For a reaction, 3 mg of substrate (equivalent to 4 mM of CA concentration in the reaction medium) was dissolved in 100 μl acetone and then added to phosphate buffer (3.7 ml, 0.1 M, pH 7.4), containing horseradish peroxidase (20 units, 20 μl) and hydrogen peroxide (30% solution, 4 μl). The solution was stirred at room temperature in the dark. Time points for ending the reaction were 3 and 18 h. The resulting suspension was rinsed with distilled water and centrifuged. The insoluble polymer was finally suspended in distilled water and dried, turning into an amorphous powder. Reactions were repeated with the addition of silicic acid (20 mM) for conditions in which a polymer was obtained. The reaction using CA only was also repeated with silicic acid concentrations of 10, 15, 20, 35, 50, and 80 mM. Silicic acid was obtained by adding 150 μl of tetramethyl orthosilicate to 850 μl of HCl 1 mM and stirred for 5 min. Samples were weighed and prepared for infrared spectroscopic analysis by crushing the products to a fine powder in a mortar, mixing thoroughly with powdered KBr (about 1 part sample to 100 parts KBr by weight), and pressed to obtain a transparent disk. Transmission spectra were measured with a Nicolet 6700 FTIR spectrometer, in the range 400–4000 cm−1, with 4 cm−1 spectral resolution, collecting 40 scans. Spectra were visualized and analysed using OriginPro 2018 (64-bit) SR1 (OriginLab Corp., Northampton, MA, USA). Results Silica aggregates are formed along with cell wall deposition In order to understand the relationship between silica supply and deposition, Si was provided in limited time intervals during seedling development (Fig. 1). Scanning electron micrographs collected at the back-scattered electrons mode could not detect silica deposits in plants grown without Si supply (hydroponics Si−). In plants supplied with Si throughout the entire cultivation (hydroponics Si+), massive silica aggregates formed, traversing most of the ITCWs, with tips protruding into the cell lumen. If Si was supplied only during the initial 3 d of cultivation (hydroponics Si+/Si−), the aggregates were substantially smaller, sunk within the ITCWs and covered with non-silicified cell wall layers. In plants supplied with Si only for the last 3 d of cultivation (hydroponics Si−/Si+), silica deposition occurred only within the inner (younger) portions of the ITCWs. The aggregates protruded to the cell lumen, similarly to aggregates in the hydroponics Si+ treatment, but with less cell wall material covering their surface. To quantify silicification we calculated the Si:C ratios of the aggregates, based on EDX analyses. In accordance with the scanning electron microscopy (SEM) images, the highest values of Si:C ratios were detected in the Si−/Si+ treatment, followed by Si+ and Si+/Si− treatment. No Si signal was detected in the Si− treatment (Fig. 1E). In the following experiments, we used the Si:C ratios accordingly, to estimate the silicification levels of silica aggregates. Fig. 1. Open in new tabDownload slide Patterns of silica deposition in hydroponically grown seedlings supplied with Si in controlled time intervals. (A–D) Scanning electron micrographs of the endodermis in primary root cross-sections: (A) Si− treatment; (B) Si+ treatment; (C) Si+/Si− treatment; (D) Si−/Si+ treatment. E, endodermis; ITCW, inner tangential cell wall of endodermis; P, pericycle; C, cortical cell; arrowheads, silica aggregates. (E) SEM-EDX silicon to carbon ratios of silica aggregates exposed by removing the cortex tissue. Measurements were recorded from the radial direction, using peeled-off segments. Different letters above the boxes indicate statistically significant differences at P≤0.05. The formation of silica aggregates requires metabolic activity Based on Soukup et al. (2017), we established an ex planta silicification system as follows. Segments of seedling roots that were grown hydroponically without silicon for 3 d were cultivated ex planta for another 3 d in a 1 mM silicic acid solution. Bright regions in SEM images representing high silicon content were detected already 4 h after exposure to Si+ solution (Fig. 2). The deposition of the mineral was detected only in the inner, younger parts of the wall, similarly to intact roots developed in planta under Si−/Si+ treatment (Fig. 1D). After 3 d of cultivation, the Si:C ratios were also similar to aggregates developed in planta under Si−/Si+ growth, reaching 0.47±0.01 (Fig. 1E). This result shows that silica deposition is independent of the plant transpiration stream. Fig. 2. Open in new tabDownload slide The time dynamics of endodermal silicification in detached root segments cultivated ex planta. Scanning electron micrographs of root segment cross-sections showing the endodermis before (A) and after (B–E) ex planta cultivation in Si+ 1 mM for (B) 4 h, (C) 24 h, (D) 48 h, and (E) 72 h. E, endodermis; ITCW, inner tangential cell wall of endodermis; P, pericycle; arrowheads, silica aggregates. We further used ex planta cultivation to test whether silica aggregates form spontaneously as silicic acid reacts with cell wall materials. Root segments were collected from Si− plants, and their outer cortical tissues were removed mechanically (including the endodermal outer tangential wall). No silica deposition was detected after 72 h cultivation in Si+ 1 mM solution. Similarly, we found no mineral deposition in segments frozen in liquid nitrogen prior to ex planta cultivation. We thus concluded that silica is not forming as a simple reaction with cell wall constituents. Next, we tested whether silica deposition involves metabolic activity of the root. We added to the ex planta growth medium 2,4-dinitrophenol (DNP), which is an ATP synthesis inhibitor. During a 4-h ex planta cultivation in 0.01 mM DNP, the aggregate formation was inhibited substantially. No silica deposition was detected under 0.05 mM DNP (Fig. 3). The segments were afterwards transferred to fresh Si+ medium without DNP and cultivated for an additional 68 h to reach the 72-h cultivation time in total. The silica deposition recovered after the transfer, suggesting that the components involved in silica deposition were not damaged by the DNP treatments. Decreasing the cultivation temperature to 4 °C hindered the silica deposition within a 4-h treatment as well (Fig. 3). Similarly to DNP treatment, transferring the segments back to regular cultivation temperature recovered the aggregate formation. Our results show that silica aggregates form only in live and active root tissue. Fig. 3. Open in new tabDownload slide The effects of metabolic inhibitors on the aggregate Si:C ratio in ex planta cultivation. The treatment (DNP or low temperature) lasted 4 h (white symbols), after which the samples were moved to Si+ medium at normal cultivation conditions for additional 68 h (gray symbols). Different letters above the columns indicate statistically significant differences at P≤0.05. Aiming to supply the segments with available energy and/or polysaccharide monomers we added to the growth solution sucrose, arabinose, or ATP (Fig. 4). As controls we used growth media containing only Si, at a concentration of 1 or 2 mM. Addition of sucrose resulted in faster silica deposition, as indicated by higher Si:C ratio along the aggregate formation (Fig. 4G). Moreover, the aggregates Si:C ratios were significantly higher in segments supplemented with sucrose as compared with segments cultivated without sucrose (Fig. 4H). In contrast, neither exogenous ATP nor arabinose increased the Si:C ratios (Fig. 4H). Our results thus propose that the sucrose provides glucose for the synthesis of the cell wall, which is a bottleneck for silica deposition. Fig. 4. Open in new tabDownload slide Endodermal silicification in root segments cultivated in 1 mM Si (control) solution, supplemented with additional silicon (final concentration of 2 mM), sucrose, arabinose, or ATP. Scanning electron micrograph were collected after 72 h in vitro cultivation in control medium (A, B) and Si+ 1 mM and 1% sucrose (suc) (C, D) or Si+ 2 mM and 1% sucrose (suc Si 2mM) (E, F) medium. Panels (A, C, E) present cross-sections and panels (B, D, F) present longitudinal views (cortical tissues removed after cultivation). E, endodermis; ITCW, inner tangential cell wall of endodermis; P, pericycle; arrowheads silica aggregates. (G) The dynamics of silicification in detached root segments cultivated ex planta for 72 h, expressed as EDX Si:C mass ratios that were collected from the silica aggregates in longitudinal views, as shown in the panels (B, D, F). The shaded area represents 95% confidence intervals indicating statistically significant differences between the treatments, and the bars represent SD. (H) EDX Si:C mass ratios of silica aggregates after 72 h ex planta cultivation in control medium (1 mM Si), or medium containing 2 mM Si and 1% sucrose (suc Si 2mM), 1 mM Si and 1% sucrose (suc), 1 mM Si and 1% arabinose (arab), or 1 mM Si and ATP (1 or 5 mM). Different letters above the boxes indicate statistically significant differences at P≤0.05. Silica deposition occurs together with the deposition of phenolic compounds Raman microspectroscopy was used to characterize the endodermal ITWC. We found that the concentration of cell wall matrix polymers (lignin and hemicelluloses) were reduced significantly under ex planta as comparison to in planta growth. Nonetheless, Si supplementation did not change the mean Raman signal significantly. (Supplementary Dataset S1). To test whether silica deposition is linked to specific components in the ITCW, we aimed to reduce specifically hemicelluloses or lignin, by treating root segments with a variety of drugs and chemicals (brefeldin A, ethanol, FA, salicylhydroxamic acid (SHAM), KI, ascorbic acid, and H2O2; Supplementary Dataset S1). A multilinear regression analysis was used to estimate the correlation between sets of ITCW Raman bands representing individual cell wall components and EDX Si:C ratios. An iterative backward elimination procedure (Pmax=0.005) allowed us to reduce the model to the major components affecting silica EDX signals. The final model (R2=0.878, Prob(F)<0.001) identified 10 peaks that change significantly with the aggregate Si:C ratio (Fig. 5). A positive correlation was associated with vibrational bands of G-lignin and CA (1173, 1268, 1575, 1656 cm−1), and negative correlations were associated with those attributed to H-lignin (643, 1205 cm−1), S-lignin (1140, 1338 cm−1), and phenolic aldehydes (1618 cm−1). A positive effect was attributed also to the peak at 492 cm−1, typically associated with a hemicellulosic C–O–C vibration. However, this contribution cannot be unequivocally assigned, since the characteristic frequencies of Si–O–Si vibrations are also located in this region. Fig. 5. Open in new tabDownload slide Assessment of the contribution of cell wall components to the aggregate Si:C ratio based on Raman–EDX correlation analysis. A visual representation of multilinear regression coefficients after the backward elimination procedure with Pmax=0.005. The combination of Raman peaks showing positive correlation with Si:C ratio indicates a dominant positive contribution of coniferyl alcohol (G-lignin constituent). The peaks exhibiting negative correlation are ascribed to phenolic aldehydes and H- and S-lignin. Coniferyl alcohol polymers establish nucleation sites for silica deposition in vitro To assess the potential relationship between silica and lignin deposition, we studied an in vitro lignin-like polymer production. Based on the Raman results we tested the hydrogenation of CA alone, or in mixtures together with FA and AX, using peroxidase and H2O2. No solid product was collected in the absence of CA, peroxidase, or H2O2. When the reaction was performed using CA alone, a dehydrogenated polymer was obtained with a reaction yield of 68% (Table 2). The infrared (IR) spectrum of the CA polymerization product (with bands at 1031, 1139, 1214, 1267, 1350, and 1598 cm−1, band assignment in Table 3) is characteristic of lignin (Agarwal and Atalla, 2016). The presence of AX in the reaction strongly reduced the yield and did not allow for infrared analysis. Surprisingly, a 1:1 mixture of FA/CA also failed to produce any polymer (Table 2). However, conducting the reaction with a CA/FA ratio of 9:1 produced a lignin polymer modified by FA (Fig. 6A). The signal at 1750 cm−1, assigned to the ester bond carbonyl stretching vibration, appeared as a consequence of the incorporation of FA into the polymer. Other spectral features, attributed to aromatic ring stretching, C–H stretching, and deformation vibrations were also found, evidenced by the bands at 1598, 1346, 1214, and 1139 cm−1 (see Table 3 for assignments). When silicic acid was added at supersaturation (20 mM) to the oxidative coupling reaction of either pure CA or CA+FA, we identified infrared absorption bands typical of silica (Fig. 6B; Table 3). The chosen above-saturation concentration probably reflects the Si concentration at its deposition sites in the plant (Casey et al., 2004). During the reaction di- and tri-silicic acid as well as small oligomeric chains of silica may form in addition to lignin and silica particles. However, these small chemical species stay in solution, and cannot be recovered with the reaction product by centrifugation, in contrast to the lignin and bigger silica molecules. Table 2. Percentage yield of the in vitro lignin precipitation after 18 h Substrate used . Yield (%) . CA 68 CA/FA (9:1) 65 CA+Si (20 mmol) 129 CA/AX (9:1) 22 CA/FA (9:1)+Si (20 mmol) 106 CA/FA/AX (8:1:1) 30 CA/FA/AX (8:1:1)+Si (20 mmol) 17 Substrate used . Yield (%) . CA 68 CA/FA (9:1) 65 CA+Si (20 mmol) 129 CA/AX (9:1) 22 CA/FA (9:1)+Si (20 mmol) 106 CA/FA/AX (8:1:1) 30 CA/FA/AX (8:1:1)+Si (20 mmol) 17 Yield of 100% equals the theoretical weight of polymerized phenolic (coniferyl alcohol, CA; ferulic acid, FA) and arabinoxylan (AX) precursors. Open in new tab Table 2. Percentage yield of the in vitro lignin precipitation after 18 h Substrate used . Yield (%) . CA 68 CA/FA (9:1) 65 CA+Si (20 mmol) 129 CA/AX (9:1) 22 CA/FA (9:1)+Si (20 mmol) 106 CA/FA/AX (8:1:1) 30 CA/FA/AX (8:1:1)+Si (20 mmol) 17 Substrate used . Yield (%) . CA 68 CA/FA (9:1) 65 CA+Si (20 mmol) 129 CA/AX (9:1) 22 CA/FA (9:1)+Si (20 mmol) 106 CA/FA/AX (8:1:1) 30 CA/FA/AX (8:1:1)+Si (20 mmol) 17 Yield of 100% equals the theoretical weight of polymerized phenolic (coniferyl alcohol, CA; ferulic acid, FA) and arabinoxylan (AX) precursors. Open in new tab Table 3. Assignment of the infrared bands in spectra obtained from the lignin-silica in vitro polymerization reactions Peak position (cm−1) . Assignment . 470 Si–O rocking mode 803 Si–O–Si symmetrical stretching mode 817 C–H out-of-plane in positions 2, 5, and 6, aromatic 854 C–H out of-plane in position 2, 5, and 6, aromatic 921 C–H out-of-plane, aromatic 966 –HC=CH out-of-plane deformation. 968 Si–OH stretching 1031 Aromatic C–H in-plane deformation; plus C–O deformation, in primary alcohols; plus C=O stretch (unconjugated) 1087 C–O deformation in secondary alcohols and aliphatic ethers 1093 Si–O asymmetrical stretching mode 1139 Aromatic C–H in-plane deformation; plus secondary alcohols plus C–O stretch 1214 C–C plus C–O plus C=O stretch 1267 C=O stretch 1330 Coniferyl ring 1367 Aliphatic C–H stretch in CH3, not in OMe; phenolic OH 1421 Aromatic skeletal vibrations plus C–H in-plane deformation 1463 C–H deformations in –CH3 and –CH2– 1506 Aromatic skeletal vibrations 1598 Aromatic skeletal vibrations plus C=O stretch 1650 H2O bending 1662 C=O stretch 2840–3000 2840 2871 C–H stretch in methyl and methylene groups 2935 3000 3399 O–H stretch Peak position (cm−1) . Assignment . 470 Si–O rocking mode 803 Si–O–Si symmetrical stretching mode 817 C–H out-of-plane in positions 2, 5, and 6, aromatic 854 C–H out of-plane in position 2, 5, and 6, aromatic 921 C–H out-of-plane, aromatic 966 –HC=CH out-of-plane deformation. 968 Si–OH stretching 1031 Aromatic C–H in-plane deformation; plus C–O deformation, in primary alcohols; plus C=O stretch (unconjugated) 1087 C–O deformation in secondary alcohols and aliphatic ethers 1093 Si–O asymmetrical stretching mode 1139 Aromatic C–H in-plane deformation; plus secondary alcohols plus C–O stretch 1214 C–C plus C–O plus C=O stretch 1267 C=O stretch 1330 Coniferyl ring 1367 Aliphatic C–H stretch in CH3, not in OMe; phenolic OH 1421 Aromatic skeletal vibrations plus C–H in-plane deformation 1463 C–H deformations in –CH3 and –CH2– 1506 Aromatic skeletal vibrations 1598 Aromatic skeletal vibrations plus C=O stretch 1650 H2O bending 1662 C=O stretch 2840–3000 2840 2871 C–H stretch in methyl and methylene groups 2935 3000 3399 O–H stretch Assignment of vibrations is based on Yoshino et al. (1990), Faix (1991), Gunde (2000), Landreau et al. (2012), and Agarwal and Atalla (2016). Open in new tab Table 3. Assignment of the infrared bands in spectra obtained from the lignin-silica in vitro polymerization reactions Peak position (cm−1) . Assignment . 470 Si–O rocking mode 803 Si–O–Si symmetrical stretching mode 817 C–H out-of-plane in positions 2, 5, and 6, aromatic 854 C–H out of-plane in position 2, 5, and 6, aromatic 921 C–H out-of-plane, aromatic 966 –HC=CH out-of-plane deformation. 968 Si–OH stretching 1031 Aromatic C–H in-plane deformation; plus C–O deformation, in primary alcohols; plus C=O stretch (unconjugated) 1087 C–O deformation in secondary alcohols and aliphatic ethers 1093 Si–O asymmetrical stretching mode 1139 Aromatic C–H in-plane deformation; plus secondary alcohols plus C–O stretch 1214 C–C plus C–O plus C=O stretch 1267 C=O stretch 1330 Coniferyl ring 1367 Aliphatic C–H stretch in CH3, not in OMe; phenolic OH 1421 Aromatic skeletal vibrations plus C–H in-plane deformation 1463 C–H deformations in –CH3 and –CH2– 1506 Aromatic skeletal vibrations 1598 Aromatic skeletal vibrations plus C=O stretch 1650 H2O bending 1662 C=O stretch 2840–3000 2840 2871 C–H stretch in methyl and methylene groups 2935 3000 3399 O–H stretch Peak position (cm−1) . Assignment . 470 Si–O rocking mode 803 Si–O–Si symmetrical stretching mode 817 C–H out-of-plane in positions 2, 5, and 6, aromatic 854 C–H out of-plane in position 2, 5, and 6, aromatic 921 C–H out-of-plane, aromatic 966 –HC=CH out-of-plane deformation. 968 Si–OH stretching 1031 Aromatic C–H in-plane deformation; plus C–O deformation, in primary alcohols; plus C=O stretch (unconjugated) 1087 C–O deformation in secondary alcohols and aliphatic ethers 1093 Si–O asymmetrical stretching mode 1139 Aromatic C–H in-plane deformation; plus secondary alcohols plus C–O stretch 1214 C–C plus C–O plus C=O stretch 1267 C=O stretch 1330 Coniferyl ring 1367 Aliphatic C–H stretch in CH3, not in OMe; phenolic OH 1421 Aromatic skeletal vibrations plus C–H in-plane deformation 1463 C–H deformations in –CH3 and –CH2– 1506 Aromatic skeletal vibrations 1598 Aromatic skeletal vibrations plus C=O stretch 1650 H2O bending 1662 C=O stretch 2840–3000 2840 2871 C–H stretch in methyl and methylene groups 2935 3000 3399 O–H stretch Assignment of vibrations is based on Yoshino et al. (1990), Faix (1991), Gunde (2000), Landreau et al. (2012), and Agarwal and Atalla (2016). Open in new tab Fig. 6. Open in new tabDownload slide Infrared spectra of synthetic lignin, silica gel, and silica formed with the synthetic lignin. (A) Comparison of spectra of synthetic lignin formed using CA alone (lower line) and CA+FA (upper line) as precursors in the polymerization reaction. The main difference in the spectra is the presence of a band at 1750 cm−1 that corresponds to a stretching vibration of the carbonyl (C=O) bond. This distinctive band indicates the presence of an ester bond and confirms the incorporation of FA into the polymer. (B) Infrared spectra of silica formed with the synthetic lignin produced using CA alone (lower line) and CA+FA (upper line) show typical silica bands. No differences were found between the spectra. (This figure is available in color at JXB online.) Addition of silicic acid to the reaction solution resulted in an increased yield (Fig. 7). The increased yield relative to the CA precursors started only above 10 mM silicic acid, growing linearly with increasing concentration of silicic acid (Fig. 7A). The higher the silicic acid concentrations in the reaction, the higher was the fraction that precipitated (Fig. 7B). These results indicate a catalytic deposition, enhanced by silicic acid availability, as opposed to a stoichiometric reaction with the polymerized lignin. In agreement, when the silicic acid concentration was lower than 15 mM, infrared spectra of the product showed only typical lignin bands (Fig. 7C). When using concentrations of 15 and 20 mM silicic acid, we identified both silica and lignin bands, in agreement with minute silica yields (assuming that the yield of the CA polymerized was similar in these reactions). When using higher silicic acid concentrations, silica bands dominated the spectra and masked the lignin polymer signals (Fig. 7C). Fig. 7. Open in new tabDownload slide Reaction yields of the in vitro lignin and silica formation. (A) Total yield calculated relative to the CA precursors added to the reaction. (B) Silica yield calculated by estimating a yield of the synthetic lignin of 68% (mean yield reaction with no addition of silicic acid). (C) Infrared spectra of the reaction product. In the products formed with concentrations of silicic acid higher than 20 mM, only the silica signals are visible and mask the lignin bands completely. (This figure is available in color at JXB online.) Silicic acid autopolymerized from supersaturated solutions in reaction mixtures missing the lignin polymerization precursors (product denoted as SiO2-Auto). This can be expected from the well-known ability of silicic acid to autopolymerize from solutions at concentrations above 2 mM (Iler, 1979). Nevertheless, the silica obtained together with CA polymer (denoted as SiO2-CAp) exhibited substantial spectral and morphological differences from SiO2-Auto (Fig. 8). Most prominently, a band that is assigned to the stretching vibration of non-bridging –SiOH (Wood et al., 1983) appeared in the spectra of SiO2-CAp at 968 cm−1 and in the SiO2-Auto spectra at 983 cm−1. SiO2-Auto formed a soft transparent gel, similar to gelatine, which was still very soft after 4 h of drying at 40 °C. The gel hardened only after several days, preserving its transparent appearance. In contrast, SiO2-CAp had whitish appearance with sandy texture, developing a solid consistency already after 4 h of drying (Fig. 8 inset). Fig. 8. Open in new tabDownload slide Comparison of silica obtained by silicic acid autopolymerization (SiO2-Auto) and silica precipitated together with the lignin like polymer (SiO2-CAp). Infrared spectra of SiO2-Auto (lower line) and SiO2-CAp (upper line) show differences in the position of the band corresponding to the –SiOH stretching (shifted from 968 cm−1 in SiO2-CAp to 983 cm−1 in SiO2-Auto), the shoulder of the band at 1093 cm−1 (asymmetric stretching Si–O–Si) and the relative intensity of the band at 803 cm−1 (Si–O–Si symmetrical stretching mode). Such spectral variations indicate differences in the molecular order between the two materials. The SiO2-Auto was formed at 20 mM concentration. The SiO2-Cap was formed at 80 mM silicic acid, in order to minimize signals of lignin. Inset: photographs of the SiO2-Auto (right) and SiO2-CAp (left) suggest that these are two different minerals. Scale bars, 500 µm. (This figure is available in color at JXB online.) Following the reaction in time we noted that while the lignin-like polymer started forming immediately, silica formation was delayed. We thus aimed to test whether the CA polymerization or the formed CA polymer caused silica deposition. After 3 h of CA polymerization in the presence of silicic acid, we could detect only the synthetic lignin and no silica (Fig. 9). SiO2-CAp formed when the synthetic lignin (product of the 3-h experiment) was centrifuged, rinsed, redissolved in phosphate buffer with 20 mM silicic acid, and stirred for 15 additional hours. This was shown by the presence of prominent silica bands in the IR spectrum (Fig. 9, iii), consistent with the SiO2-CAp IR spectra previously obtained (Figs 7C, 8). Our result indicates that the SiO2-CAp formed at a later stage, catalysed by the CA polymer. Fig. 9. Open in new tabDownload slide Infrared spectra of synthetic lignin and SiO2-CAp at 3 and 18 h. (i) CA polymerization product in absence of silicic acid after reaction of 18 h. Synthetic lignin was detected. (ii) CA polymerization product in the presence of silicic acid after reaction of 3 h. Synthetic lignin with no silica bands was observed. (iii) After a 3-h reaction CA polymerization product was redissolved in a 20 mM silicic acid solution and left to react for another 15 h. Silica and lignin bands are clearly visible. (This figure is available in color at JXB online.) Discussion Silicification in grasses typically exhibits three general patterns: silica is deposited at epidermal surfaces, cell walls of internal tissues, or intercellular spaces, including the cell lumen voids (Kumar et al., 2017b). Whereas the first type of silicification may be driven by water evaporation and considered as passive/spontaneous, evidence indicate that at least some silica deposition is controlled actively (Perry et al., 1987; Kumar and Elbaum, 2017; Kumar et al., 2017a). In our previous work we found a positive correlation between the diameter of the silica aggregates in sorghum root endodermis and the extent of the thickening of the endodermal cell wall (Fig. 4S in Soukup et al., 2017), suggesting that the aggregates form only during tertiary wall deposition. Here we show that the formation of silica aggregates takes place exclusively in the cell walls of living cells (Fig. 1). The deposition only in new cell walls and the enhanced silicification with supplementation of sucrose support a model in which silica is deposited only as an integral part of the wall, during its formation (Fig. 4). The dependence of the mineral deposition on metabolic activity at the wall, and specifically deposition of matrix polymers, can explain the restricted final dimensions of silica aggregates (Sangster and Parry, 1976a,b; Soukup et al., 2014). We adapted SEM–EDX combined with Raman microspectroscopy to correlate the extent of silica and lignin deposition. SEM–EDX is a very common method to map silica (e.g. Sangster and Parry, 1976b), and relatively quantify it in a plant tissue (e.g. Lux et al., 2002). The Si/C ratio allowed us to assess surface silica deposition in relation to the background carbon deposition that exists in the cell wall. Raman mapping of plant tissues and specifically cell wall is a commonly used methodology (Gierlinger et al., 2012), which was applied to silicified plant tissues (Sapei et al., 2007; Gierlinger et al., 2008; Blecher et al., 2012). The statistical analysis of the spectral dataset opened this field to a broad range of applications (Schulte et al., 2008; Chylińska et al., 2014; Felten et al., 2015). Utilizing statistical tools, we were able to relate increased CA peaks to increased EDX-detected silica (Fig. 5). Our in vitro experiments were conducted in supersaturated silicic acid solution. Above-saturation conditions are the standard conditions for studying silica bioproduction (Kröger et al., 1999; Dove et al., 2019). Supersaturation occurs in the xylem sap of members of the Poacae (grass family), including wheat (up to 8 mM; Casey et al., 2004), rice (up to 25 mM; Mitani and Ma, 2005), and sorghum (7–12 mM; our unpublished data). The formation of silica in the presence of polymerized CA suggests that the aromatic polymer stabilizes negative charges on oligomeric silicic acid (Dove et al., 2019). This is possible through hydrogen bonds forming between the aromatic hydroxyls and Si–OH groups. Bonds that remained active after the radical coupling may also stabilize negatively charged silicic acid. In contrast, the silica was not forming on CA monomers, nor during the CA polymerization, but only after most of the CA polymerized (Fig. 9). This result well conforms with the study by Fang et al. (2003) and Fang and Ma (2006), who demonstrate silica precipitation in vitro by synthetic or natural lignin, but not by lignin monomers, and raises the hypothesis that 3-D scaffolding is required for the mineral to grow. Our previous analyses show that the sites of silica aggregation are low in lignin and rich in FA in comparison with the cell wall surrounding them (Soukup et al., 2017). We therefore tested in vitro the possibility of precipitating silica onto a CA–FA polymer. Infrared spectra suggested that the FA units bind to the polymer via ester bonds, indicative by the appearance of a peak at 1750 cm−1 (Fig. 6). In contrast to the in vitro system, native silica aggregates autofluoresce blue under high pH (Soukup et al., 2014). This suggests that the FA in the silica aggregates is tethered to the wall via ether bonds (Leplé et al., 2007) and not through an ester. Silica formation may be induced similarly by the polymerization of monolignols in the apoplast of live cell walls. Root silica aggregates contain traces of FA and AX (Soukup et al., 2017) that may interact with the lignin to produce a silica deposition scaffold within the cell wall. We suggest a model in which negatively charged silicic acid is stabilized by newly synthesized cell wall lignin, accelerating silica nucleation and growth (Dove et al., 2019). Such interactions could be facilitated by hydrogen bonding between the hydroxyl groups on the surface of the lignin (on residues of CA, FA, other aromatic constituents, and possibly AX crosslinked to the lignin) and silica colloids or poly-silicic acid units. The stabilization may also involve bound H2O molecules. Further research is needed to identify the moieties that are trapped in the silica and can nucleate its deposition. From an evolutionary perspective, the role of silicification is still puzzling. Although many potential benefits were attributed either to the silicified cell walls (Hattori et al., 2003; Gong et al., 2006; He et al., 2014, He et al., 2015; Ma et al., 2015) or to the silica phytoliths (Massey and Hartley, 2009; Yamanaka et al., 2009; Hartley et al., 2015; Sato et al., 2016), a clear link to evolutional demands is missing (Strömberg et al., 2016). The interaction between silica and lignin we describe in this study recalls an important question – has the site-specific silicification evolved as a response to some environmental factors or as a protection against potential Si toxicity imposed by silica–lignin competition/co-precipitation? The question remains enigmatic and requires further studies. Supplementary data Supplementary data are available at JXB online. Dataset S1. Supplementary protocols. Abbreviations: Abbreviations: AX arabinoxylan CA coniferyl alcohol DNP 2,4-dinitrophenol EDX energy dispersive X-ray spectroscopy FA ferulic acid ITCW inner tangential cell wall SHAM salicylhydroxamic acid Acknowledgements This study was supported by Israel Science Foundation (534/14), ISF-ICORE grant 757/12, and Excellence Initiative of the German Research Foundation (DFG) GSC 1013 (SALSA). The authors would like to thank to Prof Alexander Lux and Dr Michal Martinka for support and comments on the manuscript. References Agarwal UP , Atalla RH. 2016 . Vibrational spectroscopy. In: Heitner C, Dimmel D, Schmidt J, eds. Lignin and lignans: advances in chemistry . Boca Raton, FL, USA : CRC Press/Taylor & Francis , 103 – 136 . Google Scholar Crossref Search ADS Google Preview WorldCat COPAC Barros J , Serk H, Granlund I, Pesquet E. 2015 . The cell biology of lignification in higher plants . Annals of Botany 115 , 1053 – 1074 . 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Do lignification and silicification of the cell wall precede silicon deposition in the silica cell of the rice (Oryza sativa L.) leaf epidermis? Plant and Soil 372 , 137 – 149 . Google Scholar Crossref Search ADS WorldCat © The Author(s) 2019. Published by Oxford University Press on behalf of the Society for Experimental Biology. This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact [email protected] © The Author(s) 2019. Published by Oxford University Press on behalf of the Society for Experimental Biology.

Zexer, Nerya; Elbaum, Rivka

doi: 10.1093/jxb/eraa127pmid: 32154874

Abstract Silicon dioxide in the form of hydrated silica is a component of plant tissues that can constitute several percent by dry weight in certain taxa. Nonetheless, the mechanism of plant silica formation is mostly unknown. Silicon (Si) is taken up from the soil by roots in the form of monosilicic acid molecules. The silicic acid is carried in the xylem and subsequently polymerizes in target sites to silica. In roots of sorghum (Sorghum bicolor), silica aggregates form in an orderly pattern along the inner tangential cell walls of endodermis cells. Using Raman microspectroscopy, autofluorescence, and scanning electron microscopy, we investigated the structure and composition of developing aggregates in roots of sorghum seedlings. Putative silica aggregation loci were identified in roots grown under Si starvation. These micrometer-scale spots were constructed of tightly packed modified lignin, and nucleated trace concentrations of silicic acid. Substantial variation in cell wall autofluorescence between Si+ and Si– roots demonstrated the impact of Si on cell wall chemistry. We propose that in Si– roots, the modified lignin cross-linked into the cell wall and lost its ability to nucleate silica. In Si+ roots, silica polymerized on the modified lignin and altered its structure. Our work demonstrates a high degree of control over lignin and silica deposition in cell walls. Endodermis, lignin, root, silica, silicic acid, silicon, Sorghum bicolor Introduction Silicon (Si) is a highly abundant mineral in specific plant tissues, yet its contribution to plant biology is poorly understood. It is well known that stressed plants lacking access to silicic acid suffer in many respects (Ma and Yamaji, 2006; Coskun et al., 2019). Therefore, although Si is not an essential element, it is extremely important. Si is present in most soils as various forms of Si dioxide, constituting the major environment of plant roots (Currie and Perry, 2007). Soil silicates dissolve and release monosilicic acid [Si(OH)4], which is available for root plant uptake. The silicic acid moves with water in the plant to target locations, mostly in the shoot epidermis. There, the acid deposits as solid hydrated silica (SiO2·nH2O), also called biogenic opal or silica (herein referred to as silica). The current paradigm is that the solid biomineral has little interaction with plant biochemistry because it is insoluble after deposition (Yoshida et al., 1962). Its effects are thus limited to its material properties. Some examples are reduction of cell wall digestibility (Van Soest and Jones, 1968; Massey and Hartley, 2009), fortification of prickles and hairs (Bauer et al., 2011; Mustafa et al., 2018), and filtering of UV light (Goto et al., 2003; Pierantoni et al., 2017). However, exposing plants to soluble silicic acid improves their performance under a wide variety of stresses (Epstein, 2009; Liang et al., 2015). The mechanism by which Si (as silica or silicic acid) influences plant physiology remains elusive (Detmann et al., 2012; Vivancos et al., 2015; Markovich et al., 2017; Coskun et al., 2019). Plants may take up Si passively or actively through channels and transporters termed Low silicon (Lsi1, Lsi2; Ma et al., 2006, 2007; Ma and Yamaji, 2015; Coskun et al., 2019). These proteins allow the passage of Si across the endodermis, which otherwise constitutes an impermeable barrier to Si diffusion (Ma and Yamaji, 2015). The Si is then translocated to the shoot via the xylem transpiration stream (Jones et al., 1963). The amount of silica deposited usually correlates with the transpiration rates of an organ (Jones et al., 1963; Trembath-Reichert et al., 2015; McLarnon et al., 2017), suggesting that silicic acid is mostly present in the apoplastic volume. Transpiration is also regarded as the main driving force for the deposition of silica. However, we and others have shown that leaf silicification in silica cells is not interrupted even in the absence of transpiration, and that silicification occurs only in live cells (Sangster and Parry, 1971; Kumar et al., 2017; Kumar and Elbaum, 2018). Sorghum (Sorghum bicolor (L.) Moench) silica cells express the silica-depositing protein Siliplant1 (Slp1) and export it to the cell wall, timed with cell mineralization. There, Slp1 interacts with the silicic acid in the apoplast, causing local silica precipitation (Kumar et al., 2019, Preprint). Silica may deposit within cellulosic cell walls (Hodson, 2016). Cell wall components interacting with silica may include cellulose (Perry and Lu, 1992), hemicelluloses (Perry et al., 1987; He et al., 2015), callose (Brugiére and Exley, 2017; Kulich et al., 2018), lignin (Fang and Ma, 2006), and ferulic acid (Soukup et al., 2017). Silicified structures are frequently associated with lignified tissues (Zhang et al., 2013; Fleck et al., 2015). Silica in rice straw is associated mostly with Klason extracted lignin, constituting ~80% of the total silica (Pan et al., 2017). Several studies have observed lignin–silica trade-offs in plants grown in Si-available soils or media (Goto et al., 2003; Schoelynck et al., 2010; Yamamoto et al., 2012). Considerable silicification occurs in the root endodermis of sorghum, in the inner tangential cell wall (ITCW) (Parry and Kelso, 1975; Soukup et al., 2017) (Fig. 1). The deposits, which are internal to the Casparian strips, precipitate from the apoplastic sap of the stele. They are constituted of silicic acid molecules that were taken up by the root transporters and reached the transpiration stream. However, the Si did not flow up to the shoot, but was bound to the thick cell wall at the boundary of the stele. The silica aggregates can form in detached root segments cultivated in silicic acid solution ex planta. The mineral forms without any transpiration, in an energy-dependent metabolic process (Soukup et al., 2019). Fig. 1. Open in new tabDownload slide Cross sections of primary roots of sorghum. (A) Light micrograph of a cross section of a Si+ root stained with toluidine blue. (B) Close-up of the endodermal area marked by an arrow in panel A. Silica aggregates (red arrows) form on the ITCW of endodermis cells. (C, D) Scanning electron micrographs in back-scattered electrons mode (SEM-BSE) of the endodermis of (C) Si+ and (D) Si– roots. Silica aggregates (white in appearance) form only in the ITCW of endodermis cells of Si+ roots. c, cortex; e, endodermis, p, pericycle. Scale bars: A=50 µm, B–D=10 µm. Partial acid digestion of sorghum roots leaves ferulic acid and hemicellulose associated with the silica aggregates, suggesting that these components are occluded in the mineral (Soukup et al., 2017). Blue fluorescence is associated with intact aggregates, which increases under high pH (Soukup et al., 2014). This enhancement in the fluorescence indicates the content of lignin associated with the mineral (Harris and Hartley, 1976). In grasses, ferulic acid is a common lignin monomer (Ralph et al., 2008) that may also crosslink hemicellulose. An indication of ferulic acid bound to hemicellulose may come from a shift from blue fluorescence under neutral pH to green fluorescence under higher pH (Harris and Hartley, 1976). This shift occurs as a result of ionization of the phenolic hydroxyl of ferulic acid (Rudall and Caddick, 1994). In this work we examine the first stages of silica deposition at the endodermis of sorghum roots. Using fluorescence microscopy combined with Raman microspectroscopy and scanning electron microscopy (SEM), we show that very local lignin modifications induce silica nucleation. These loci are poor in ferulic acid in comparison to their surroundings. Materials and methods Plant material and growing conditions Grains of Sorghum bicolor (L.) Moench, line BTx623, were surface-sterilized with 3% sodium hypochlorite for 10 min and washed twice with distilled water. Grains were then placed in petri dishes lined with wet filter papers and germinated for 72 h. After germination, seedlings were grown hydroponically for 1 week. All growth solutions were based on double-distilled water containing 1 mM CaCl2. Si+ medium was supplied with sodium silicate (Na2SiO3) at a final concentration of 2 mM. Si– medium was provided with NaCl at a similar final concentration in order to maintain equal ionic balance across all media. The final pH of all the solutions was adjusted to 5.8 with HCl. Seedlings were grown on plates of polystyrene foam mounted on top of darkened 1 litre beakers containing growing media, with 10 seedlings per beaker. Cultivation was done in a growth chamber under controlled conditions, with a photoperiod of 16 h:8 h (light:dark), illuminated with photosynthetically active radiation of approximately 200 μmol m−2 s−1, 28 °C:22 °C (light:dark) temperature, and 70% air humidity. Root tissue embedding and sectioning for light microscopy Complete primary roots were harvested and fixed in FAA [50% ethanol, 35% double-distilled water, 10% formaldehyde (37%), 5% glacial acetic acid; v/v]. The roots were then divided into four zones: zone i, lowest 4 cm from the root tip; zone ii: 4–8 cm from the root tip; zone iii, 8–12 cm from root tip; zone iv, 12–16 cm from the root tip. Root pieces 5 mm in length were taken from zone iii and embedded using a Leica Historesin embedding kit (Leica, Germany) according to the manufacturer’s instructions. Sections 4 µm thick were obtained using a tungsten carbide blade mounted on a Leica RM2165 microtome. Sections were stained with 0.1% (w/v) toluidine blue for 2 min and washed three times in double-distilled water. Specimens were visualized under a Leica DM500 light microscope equipped with a Leica ICC50W camera (Leica, Germany). Scanning electron microscopy and energy-dispersive X-ray analyses Complete primary roots were harvested and fixed in ethanol:acetic acid (9:1 v/v) under vacuum for 24 hours. Root samples were kept in the same fixative at 4 °C for an additional 48 hours. The fixing solution was then replaced with 70% ethanol and samples were stored at 4 °C until use. The roots were divided into the abovementioned four zones. Cross sections cut by hand were collected from all zones and mounted on aluminum stubs using carbon tape. Other root sections were carefully peeled out of their outer cortical tissues using finely serrated tweezers. Representative segments from all zones were mounted on aluminum stubs using carbon tape. Observations and scanning electron microscopy–energy-dispersive X-ray (SEM-EDX) analyses were performed with a JEOL JSM-IT 100 InTouchScopeTM scanning electron microscope (JEOL, Japan) under a low vacuum (30 Pa) and with an accelerating voltage of 20 kV. All micrographs were taken using the back-scattered electrons (BSE) mode (SEM-BSE). SEM-EDX analysis maps were collected at the K-edge using the JEOL integral detector with a dwell time of 0.1 ms and sweep count of 30. Confocal and fluorescence microscopy Peeled root segments were prepared as described for SEM analysis. Root sections 20 mm in length were mounted on glass microscope slides. Samples were immersed in double-distilled water and covered with cover slips. Observations and images were made using a Leica TCS SP8 confocal laser scanning microscope equipped with a ×63 water-immersed objective. The excitation wavelengths used were 405 nm and 488 nm; emission was collected at 400–500 nm and 500–550 nm, respectively. Epifluorescence and bright-field micrographs were collected by a Nikon Eclipse 80i microscope (Nikon, Japan); an X-cite 120Q (Lumen Dynamics, Canada) UV light source was used. Specimens were observed with the following set of Nikon filters: GFP (excitation: 450–490 nm; emission: 500–550) and DAPI (excitation: 400–418 nm; 450–465 nm). Images were taken with a CRI camera controlled by Abrio 1.4 software (PerkinElmer, USA). Raman microspectroscopic observation of the endodermal cell wall Peeled root segments were prepared as described for SEM analysis. Root sections were mounted on aluminum microscope slides using super glue. In cases where roots were chemically treated, root segments were gently shaken for 1 h in 1 M NaOH or 3% acidefid NaOCl (pH 4.5). The samples were then washed tree times with double-distilled water and mounted on aluminium slides. The samples were covered with a drop of double-distilled water and measurements were made using a ×63 water-immersed objective. Raman maps were collected with a Renishaw InVia spectrometer equipped with a 532 nm laser (45 mW maximum intensity, 2 μm2 beam), utilizing WiRE3.2 software (Renishaw, New Mills, UK). Measurements were performed using the Streamline mode with acquisition time of 30 s. Spectral analysis was done in WiRE3.2 (Renishaw), including normalization of the total signal to 1000 arbitrary scattering units, background subtraction, and peak picking. Co-localization of Raman, SEM, and fluorescence signal maps Raman maps of single endodermis cells were measured from the edge of a root segment extracted from zone iii. The areas measured were also photographed using the instrument’s internal reflected light microscope. For same-cell SEM-BSE analysis, a measured root segment was carefully transferred from the Raman aluminum slide onto a SEM aluminum stub covered by carbon tape and observed by SEM-BSE. For fluorescence observation of the Raman-mapped cells, the same root segment, still mounted on the aluminum slide was examined. The root segment was immersed in double-distilled water, covered with a cover slip, and imaged by epifluorescence. Identification of the single cells mapped with Raman spectroscopy in SEM-BSE and epifluorescence was achieved by superimposing and aligning the images using ImageJ software (Bethesda, MD, USA). Results Silica aggregates in the endodermis of sorghum roots are associated with a tertiary lignified cell wall that forms after the primary and secondary cell walls have completed their development (Sangster and Parry, 1976) (Fig. 1A, B). These can be identified by SEM-BSE in roots grown hydroponically with silicic acid (Si+). In this scanning mode, Si atoms scatter more electrons than carbon (C) and oxygen (O) atoms, and thus appear brighter than the organic tissue (Fig. 1C). We did not observe silica aggregates in roots grown without silicic acid (Si–) (Fig. 1D). In order to study lignin deposition, we examined the autofluorescence of the cross sections (Fig. 2). The ITCW of endodermis cells of Si+ roots exhibited blue fluorescence localized to the rims and base of the aggregate. Green autofluorescence was identified surrounding the silica body (Fig. 2A–C). In contrast, in Si– ITCW we observed a distinct green spot at the center of the cell wall. This feature was not identified in cross sections of Si+ roots (Fig. 2D–F). Fig. 2. Open in new tabDownload slide Autofluorescence emitted by cross sections of sorghum roots. (A) Blue, (B) green, and (C) merged image of both signals in Si+ roots. Arrows indicate silica aggregates. The blue fluorescence is more intense at the base and edges of a silica aggregate than in the cell wall itself and at the tip of the aggregate. Green fluorescence of the ITCW outlines the aggregate. (D) Blue, (E) green, and (F) merged image of both signals in Si– roots. Arrows mark the location of spots of green fluorescence in the ITCW. c, cortex; e, endodermis; p, pericycle. Scale bars=10 µm. To assess the developmental relationship between silica aggregation and the fluorescence pattern, we defined root zones 4 cm in length, starting at the root tip, which is the youngest part of the root (Fig. 3A). To identify the active silicification zone (ASZ) along roots of Si+ seedlings, we gently stripped roots from their cortex and exposed the developing endodermis. Using SEM-BSE, we identified silica aggregation at 4 cm above the root tip (Fig. 3B, zone ii). The silica aggregates increased in diameter along the developmental gradient of the root. Closer to the root–shoot junction, additional aggregates appeared between more developed ones, filling the ITCW with aggregates (Fig. 3B, zone iv). Fig. 3. Open in new tabDownload slide Identification of the active silicification zone (ASZ) in roots of sorghum seedlings. (A) Representative image of a primary root of a 10-day-old sorghum seedling cultivated in Si+ hydroponic solution. Red rectangles indicate the regions observed in panel B. Scale bar=1 cm. (B) To visualize the endodermis along the root, the cortical tissues were removed. SEM-BSE micrographs reveal the silica aggregates (white in appearance). Four distinct silicification regions were defined, starting at the young zone close to the root tip (bottom): (i) no silica aggregation; (ii) aggregates begin to form from approximately 4 cm above the root tip; (iii) aggregates grow in diameter; (iv) at the root base close to the root–shoot junction, new aggregates appear between developed ones. Scale bar=50 µm. A close-up of the silica aggregates in zones ii–iv shows that they form a pattern independent of the numerous pits (seen as black dots in Fig. 4A–C). In zone iii (Fig. 4B), we identified the initiation of small aggregates between more developed ones. In this region, silica deposition seems to be most intensive. Timed with the appearance of the aggregates, we detected a pattern of blue autofluorescent spots (Fig. 4D–F). The spatial distribution of the blue fluorescence was similar to the silicification pattern, suggesting that phenolic molecules are distributed in a similar pattern to the silica. Green fluorescence was detected mostly in the radial cell walls of the young root, in zone ii (Fig. 4I). In zones iii and iv, the fluorescence was also detected in the ITCW (Fig. 4G, H). Interestingly, green fluorescing rings circled the blue fluorescent spots (Fig. 4J, K). This complements our observations in the cross sections (Fig. 2B, C), suggesting that a ring of green fluorescence delineates the silica aggregates at the surface of the ITCW. Fig. 4. Open in new tabDownload slide Autofluorescence and SEM of the ITCW of the endodermis, imaged along roots of Si+ sorghum seedlings after removal of the cortical tissues. SEM-BSE micrographs of zones iv (A), iii (B), and ii (C), as defined in Fig. 3. Blue autofluorescence of zones iv (D), iii (E), and ii (F) creates a similar pattern to that of the aggregates. Green autofluorescence of the same samples in zones iv (G), iii (H), and ii (I) is limited mostly to longitudinal cell walls in zone ii; in zones iii and iv, fluorescence is extended to the ITCW, with the exception of non-fluorescent spots encircled by highly fluorescing green rings. (J–L) Merged view of the blue and green autofluorescence images confirms that the green fluorescence delineates the blue fluorescent spots. SEM-BSE and confocal micrographs are independent and do not represent the same cells. Scale bar=10 µm. To study the time course of silicification, we exposed Si– roots to Si+ growth solution, and imaged zone iii by SEM-EDX (Fig. 5). At time 0, Si– roots exhibited bright ring-like structures arranged in a pattern that resembled the silica aggregation. Importantly, no Si signal could be measured by SEM-EDX in these spots, and we detected no significant topography related to the spots (see Supplementary Fig. S1 at JXB online). The higher BSE signal was attributed to higher signals of C and O atoms, suggesting some differences in cell wall composition and density (Fig. 5A). Initial silica aggregation in distinct spots was detected after 2 h of exposure to Si+ media by both SEM-BSE and SEM-EDX (Fig. 5B). The density of the aggregates was similar to that detected in zone ii (Fig. 4C). At this stage, the silica was mostly embedded within the ITCW. After 4 h of exposure to Si+ solution, we detected small aggregates more or less equidistant from larger ones (Fig. 5C). The locations with high Si exhibited high O and low C, as we would expect with the formation of silica (SiO2). With 6 h of exposure, many of the larger aggregates protruded from the ITCW (Fig. 5D). After 24 h, the density of the aggregates remained similar to that at the 4 h exposure (Fig. 5E), and similar to that detected in Si+ grown roots (Fig. 4A). A similar rate of Si deposition was previously reported by Lux et al. (2003). Fig. 5. Open in new tabDownload slide SEM-BSE and EDX imaging of the ITCW of root endodermis in zone iii. Sorghum seedlings were cultivated in Si– medium for 1 week and then transplanted into Si+ medium. Samples were imaged by SEM-BSE at higher (top row) and lower (second row) magnification, and by SEM-EDX for carbon (C; red panels), oxygen (O; gray panels), and silicon (Si; pink panels). (A) Before exposure to Si+ medium (time 0), bright structures were identified by SEM-BSE (top two panels). These structures were richer in C and O, but not in Si. (B) After exposure to Si+ for 2 h, sporadic Si aggregation was detected by SEM-BSE (top two panels) and SEM-EDX. The Si-rich regions were rich in O and poor in C. (C) After exposure for 4 h, the density of the aggregates increased. (D) At 6 h exposure, the aggregates became larger. SEM-BSE at higher magnification (top panel) revealed that the aggregates started to project out of the ITCW. (E) At 24 h exposure, the aggregates were even larger and protruded more from the cell wall, while their density was similar to that at 4 h exposure to Si+ medium. Scale bar in E top panel, common to all top panels, represents 5 µm. Scale bar in E bottom panel, common to all but the top panels, represents 25 µm. We hypothesized that the non-silicified bright structures we detected in Si–deprived roots (Fig. 5A) are predictors for silica deposition. We thus aimed to study the chemistry of these structures. We first examined the autofluorescence and SEM-BSE-visible features along Si– roots (Fig. 6). The bright cell wall structures visible by SEM-BSE could be detected in zones iii and iv but not zone ii (Fig. 6A–C). Interestingly, we could not detect the blue spotted pattern that characterized the silica aggregation in Si+ roots (compare Fig. 6D–F with Fig. 4). Instead, green autofluorescent spots were apparent in zones iii and iv (Fig. 6G–I). These features can explain the localized green autofluorescence detected in cross sections of Si– root (Fig. 2E). Together, the results of our autofluorescence study suggest that in Si– ITCW a green fluorescent spot lies on top of a blue fluorescent cell wall. Fig. 6. Open in new tabDownload slide Autofluorescence and SEM of the ITCW of the endodermis, imaged along roots of Si– sorghum seedlings after removal of the cortical tissues. (A–C) SEM-BSE micrographs of zones iv (A), iii (B), and ii (C), as defined in Fig. 3. (D) Blue autofluorescence of zone iv creates a pattern of gaps (red arrows), suggestive of the silica distribution in Si+ roots. The blue autofluorescence pattern in zones iii (E) and ii (F) is irregular, with lines running along the endodermis ITCW long axis, in addition to the fluorescence of the radial cell walls. The pattern of green autofluorescence of the same sample in zone iv (G) complements the blue autofluorescence, creating spots that are distributed similarly to the silica aggregates in Si+ roots. This pattern is initiated in zone iii (H). (I) The green autofluorescence in zone ii is limited mostly to the radial cell walls. A merged image of the blue and green autofluorescence images in zone iv (J) confirms that the green fluorescent spots do not fluoresce in blue. Merged images of the blue and green autofluorescence images of zone iii (K) and zone ii (L) show that the green and blue autofluorescence are mostly separated, except for the radial cell walls, which show both green and blue autofluorescence. SEM-BSE and confocal micrographs are independent and do not represent the same cells. Scale bar=10 µm. We further studied the chemistry of the Si– ITCW by Raman microspectroscopy and chemical manipulation (Fig. 7). A spectrum typical of lignocellulose was collected from the native ITCW (Fig. 7A, i). We assigned the major scatterings at 1697, 1633, 1603, and 1171 cm−1 to lignin and aromatic molecules (Agarwal, 2014). We then treated the ITCW with a solution of 1 M NaOH (Fig. 7A, ii). Hemicelluloses are extracted with this treatment (Carpita, 1984). We found that the typical ferulic acid peaks at 1633 and 1171 cm−1 were reduced relative to the total aromatic scattering at 1603 cm−1 (Ram et al., 2003). Peaks assigned to cellulose, with the strongest scattering at 1122 and 1093 cm−1 (Agarwal, 2014), were smaller in relation to the band representing total aromatics at 1603 cm−1. We also examined the spectral changes as we removed lignin by treatment with acidic bleach (Carpita, 1984). This extraction resulted in a decrease in the ferulic acid and lignin-related peaks at 1697, 1633, 1603, and 1171 cm−1. The cellulose/hemicellulose strongest peaks at 1122 and 1093 cm−1 became the major spectral features (Fig. 7A, iii). Raman mapping of the peak intensity at 1630 cm−1, which represents ferulic acid, revealed the familiar spotted pattern, with low intensities marking the spots (Fig. 7B). After treatment with NaOH solution, the pattern at the ITCW was still visible, although less clear (Fig. 7C). The acidic bleach treatment removed the Raman pattern completely (Fig. 7D). We could not detect the pattern by plotting other peaks. In correlation, the blue and green autofluorescence pattern was also weaker after NaOH treatment and undetectable after acidic bleach treatment. Moreover, the acidic bleach treatment also removed the bright SEM-BSE structures (Supplementary Fig. S2). Fig. 7. Open in new tabDownload slide Raman microspectroscopy of the ITCW of root endodermis at zone iii in Si– sorghum seedlings. (A) Spectra of the native ITCW (i) and the cell wall after extraction with 1 M NaOH (ii) or acidic bleach (iii). Peaks were assigned according to Agarwal (2014). Specifically, peaks were identified at the strongest scattering of lignin (at 1171, 1603, 1633, and 1697 cm−1) and xylan (at 491 and 1091 cm−1), and the strong and medium scattering peaks of cellulose (at 379, 432, 460, 520, 899, 970, 996, 1057, 1096, 1122, 1152, 1294, 1340, and 1378 cm−1) and glucomannan (at 1089, 1121, 1264, 1374, and 1469 cm−1). Brown text represents lignin residues and black text represents cellulose/hemicellulose residues. The ITCW Raman map of the peak intensity at 1630 cm−1, assigned to ferulic acid, was plotted for (B) native ITCW, (C) ITCW treated by NaOH, and (D) ITCW treated by acidic bleach. A spotted pattern of low intensity at 1630 cm−1 was detected in the native and NaOH-treated cell walls, but absent in the cell walls treated with acidic bleach. Scale bar=5 µm. To link the spectral features to the autofluorescent spots and the structures visualized by SEM-BSE, we measured the same endodermis cell by both autofluorescence and Raman (Fig. 8A, B), and by SEM-BSE and Raman (Fig. 8C, D). Plotting the ferulic acid peak at 1630 cm−1, we mapped its lowest relative intensities to the locations of high green autofluorescence and to high SEM-BSE brightness. These results confirm that the spotted pattern observed by the three different methods co-localize (Fig. 8). Collectively, this observation suggests that the same material producing the spotted pattern is detected by the three methods of microscopy and is removed by acidic bleach. We propose that the spots are made of a lignin-related compound. Fig. 8. Open in new tabDownload slide Spatial correlation between autofluorescence, Raman, and SEM of the ITCW of the endodermis at zone iv, imaged in roots of Si– sorghum seedlings after removal of the cortical tissues. (A) Green autofluorescence of the ITCW of a single endodermis cell. The radial cell walls are seen as a frame to the spots. (B) Intensity of the Raman scattering at 1630 cm−1 of the same cell is plotted in red, on top of the fluorescence map. (C) SEM-BSE micrograph showing a pattern of bright structures within the ITCW (arrows). (D) Intensity of the Raman scattering at 1630 cm−1 of the same cell is plotted in red, on top of the SEM-BSE micrograph. Low Raman intensities are co-localized to both the autofluorescent green spots and the bright structures visible in SEM-BSE, suggesting that the SEM-BSE and fluorescence patterns are also co-localized. Scale bar=5 µm. To characterize lignin modifications at the ITCW, we compared autofluorescence under neutral and basic conditions (Fig. 9). Unmodified lignin has blue autofluorescence, which is enhanced under basic pH, while modified lignin may fluoresce in green, depending on specific chemical modifications (Abraham et al., 2018). Enhanced fluorescence and a shift to green fluorescence under high pH is a result of the ionization of lignin hydroxyl and/or carboxyl groups. However, if these groups are engaged in ester or ether bonds, the fluorescence is quenched (Meyer et al., 2003). Blue autofluorescence of the Si– ITCW was enhanced when observed immediately after mounting in 1 M NaOH. In contrast, the spots were less fluorescent (Fig. 9A, B). This indicated a fraction of non-modified lignin surrounding the spots. The spots themselves presented green autofluorescence, which disappeared with high pH only at the center of the spots (Fig. 9C, D). Superimposing the green and blue epifluorescence signals, we found that the radial cell walls and some of the ITCW changed their fluorescence from blue at pH 7 to green at pH 12 (Fig. 9E, F). This indicated the presence of ester-bound ferulic acid with a phenolic hydroxyl that was free to ionize, possibly modifying hemicellulose (Harris and Hartley, 1976). Under high pH, the green spot itself showed a radial variation in autofluorescence: dark at the center, surrounded by a green ring, which in turn was surrounded by a blue ring. Fig. 9. Open in new tabDownload slide Changes in the autofluorescence of the ITCW of root endodermis at zone iii in Si– sorghum seedlings with changes in pH. (A) Under neutral pH, blue epifluorescence shows a pattern spots of low fluorescence at the ITCW. (B) Under high pH, the low-intensity spots are enhanced, indicating that the fluorescent cell wall contains unmodified lignin. (C) Green epifluorescence shows a pattern of spots with high fluorescence at the ITCW. (D) Under high pH, the green spots are extended, and their center loses its fluorescence. (E) Merged image of the green and blue epifluorescence at neutral pH demonstrates that in the ITCW the two signals do not overlap. (F) Merged image of the green and blue epifluorescence at high pH reveals a ‘bagel’ structure, with a dark center encircled by a green ring, which in turn is encircled by a blue ring. Scale bar=10 µm. Interestingly, the very center of the green spots, which did not fluoresce under high pH in most spots, did fluoresce in blue in rare cases. Blue fluorescence was observable as a central dot under high pH, in the very mature parts of some of the roots (Fig. 10A). In the same regions scanned by SEM-BSE, we could also identify a bright dot in the middle of some of the bright structures (Fig. 10B). These dots were identified by SEM-EDX as silica (Fig. 10C). We explain the deposition of silica in Si– roots as the result of a very minute amount of silicic acid as a contaminant in the hydroponic solution. This finding assigns a role of silica nucleation to the lignin-based spots at the ITCW. Fig. 10. Open in new tabDownload slide Imaging of the ITCW of Si– root endodermis in zone iv. (A) Blue autofluorescence at high pH reveals a bright dot (arrow) at the center of some of the low-fluorescence spots. (B) In the center of some of the bright structures observable by SEM-BSE, a dot (arrow) brighter than the organic background can be identified. SEM-EDX spectra were collected at the + signs. (C) The SEM-EDX spectrum at the dot (blue +) suggested that silica aggregated in this location. The SEM-EDX spectrum adjacent to the dot (orange +) indicates that Si content was below the limit of detection. Scale bars=10 μm. This figure is available in color at JXB online. Discussion The pattern of silica deposition is strictly controlled Our results show that the tertiary thickening of the ITCW includes the deposition of modified lignin and silica in a pattern of spots. Lignin patterning can be achieved by patterning the binding of cell wall NADPH oxidases and peroxidases to the cell membrane through scaffold proteins, which creates local production of hydrogen peroxide (Lee et al., 2013). Laccases that produce radicals of monolignols can be linked specifically to secondary cell walls, possibly through their glycosyl modifications (Yi Chou et al., 2018). They can also create localized spots of lignin polymerization (Schuetz et al., 2014). The deposition of lignin at the ITCW starts together with the formation of the silica aggregates in Si+ roots (Soukup et al., 2017). In Si– roots, we detected a micron-scale patterning of lignin types that exhibit variation in their autofluorescence (Fig. 9). This highly regulated process must be actively controlled by living endodermis cells, in agreement with our previous results showing that the aggregation of silica depends on metabolic energy (Soukup et al., 2019). The spotted pattern starts to develop at ~4 cm from the root tip. In this region lateral roots are emerging, and the major role assigned to the endodermis is mechanical (Geldner, 2013). Silica and lignin may strengthen the ITCW. However, the biological role of the complex lignification–silicification patterning is unknown. The first spots, which form in zone ii (Fig. 3), are located distantly from one another, with a gap of about twice the width of the cell (Fig. 4). In zone iii, new spots appear between the older ones, creating a weak pattern of alternating larger and smaller aggregates. This variation diminishes as the aggregates complete their growth in zone iv. Under Si depletion, the appearance of the spotted pattern is delayed. This is in agreement with the delayed development of the endodermis in Si– roots of rice and maize (Fleck et al., 2011; Lukačová et al., 2013). In sorghum, the spots are established in Si– roots only in zone iii, where they assume a spatial distribution similar to the aggregates in zone iii of Si+ roots (Fig. 6). Our observations show that the root tissue prepares itself for the deposition of silica, and there are no silica deposits in young tissues (Fig. 3). In addition, in Si– roots that are exposed to silicic acid, a gradient of aggregate sizes can be observed (Soukup et al., 2017). The largest aggregates develop in zones iii and iv, and smaller aggregates are located in older tissues, close to the root–shoot junction. This suggests that there is a region where the deposition is most effective, in correlation with the formation of the tertiary cell wall. This is suggested to be the ASZ. The whole process occurs within hours, suggesting that transcription and translation of the relevant genes has already occurred in the younger parts of the root. The spots are possibly made of densely packed lignin The greater brightness of the non-silicified cell wall at the putative silica deposition spots observed by SEM-BSE (Fig. 6) could be related to a higher topography or density of the cell wall. However, the topography is not pronounced, as can be seen in root sections (Fig. 1C, Supplementary Fig. S1; see also Sangster and Parry, 1976; Lux et al., 2003; Soukup et al., 2017). Similarly, lignin, cellulose, and hemicellulose have the same dry density (~1.5 g ml–1; Raven et al., 2005) and atomic fraction of C and O to H (with a general formula of C5–6H10–12O4–7). Nonetheless, since the bright spots seen by SEM-BSE were eliminated with the removal of lignin (Supplementary Fig. S2), we conclude that they are made of lignin that was packed in a denser manner than the other cell wall constituents. A possible structure could be the stacking of aromatic rings of lignin monomers. This organization may also cause a red shift of the autofluorescence of unmodified lignin, explaining the distinct green autofluorescence we detected in the dense cell wall spots (Fig. 6). Careful autofluorescence analysis located ferulic acid modifying hemicellulose in the radial and bulk inner tangential cell walls. Fig. 9 shows that in these regions there was a shift from blue to green autofluorescence with a pH change from 7 to 12. This indicated the presence of free hydroxyl groups on ferulic acid moieties, typically bound to hemicellulose (Harris and Hartley, 1976). However, the blue to green shift was not observed in the silica nucleating spots. The absence of ester-bound ferulic acid at the center of the spots was supported by chemically removing hemicellulose with NaOH. We could still identify the green fluorescent spots in root tissue subjected to this treatment. In addition, mapping of the ferulic acid Raman peak revealed that the spotted pattern was still present after the removal of hemicellulose by NaOH (Fig. 7C). This, too, indicated that ferulic acid bound to hemicellulose is not localized uniquely to the silica-nucleating spots. Instead, the green fluorescence of the spot centers disappeared under high pH (Fig. 9D). This may be explained by the disruption of densely packed lignin. The aggregates form layer by layer only at their base Our observations are in contrast to our previous findings showing that root silica extracted by sulfuric acid contains hemicellulose and ferulic acid (Soukup et al., 2017). We can explain this by separating the nucleation process from the growth of a silica aggregate that may capture ferulic acid-bound hemicellulose. We suggest a sequence of events that may produce silica aggregates at the endodermis ITCW in sorghum (Fig. 11). According to our model, the endodermis cell produces a local spot at its cell wall, where a unique lignin polymer is synthesized. We term this polymer the ASZ lignin. The putative ASZ lignin has extreme affinity for silicic acid and the ability to catalyze the condensation of silicic acid to silica even under residual concentrations of silicic acid (Fig. 10). Under Si starvation, the ASZ lignin is incorporated into the cell wall, making it fluorescently green (Fig. 11B). In the presence of silicic acid, the ASZ lignin monomers/oligomers cross-link to the cell wall and at the same time interact with the apoplastic silicic acid. In this form, the ASZ lignin is fluorescently blue (Fig. 11C, bottom panel). As the polymerization proceeds in zone iii (Fig. 11C, middle panel), some ASZ lignin is captured in the growing mineral before cross-linking to the cell wall. Close to the location of ASZ lignin formation, silica polymerizes very rapidly, incorporating only little ASZ lignin and cell wall polymers. Some of the ASZ lignin units that are not interacting with silicic acid (possibly due to a lack of free acid) diffuse to the rim of the aggregate and create cross-links between the cell wall polymers, maybe in a similar fashion to the cross-links created by ferulic acid (Hatfield et al., 2016). This non-silicified ASZ lignin may be the source of the green fluorescence at the periphery of the aggregate, similar to the green spots in Si– roots. Binding to the cell wall changes these lignin units and they lose their ability to polymerize silicic acid. The final form of the aggregate includes a layer-by-layer deposition of cell wall and silica at its base and margins, and mostly silica at its top (Fig. 11C, top panel). A cross-sectional SEM-BSE image of an aggregate embedded in the endodermis ITCW shows the layers of (mostly) cell wall at the rim of the aggregate and (mostly) silica at its center (Fig. 11D). Fig. 11. Open in new tabDownload slide Suggested model for the formation of silica aggregates in sorghum root endodermis. (A) Scheme of a root section, showing the cortex (c) in gray, the stele (s) in brown, and the endodermis cell layer in yellow. The upper diagram shows a magnified view of the stele and endodermis, with one cell exhibiting silica aggregates (dark gray), viewed from the cytoplasm of the endodermis cell (marked by a rectangle) and in a cross section (marked by a circle). (B, C) Schemes showing a suggested developmental sequence of the endodermis ITCW in zones ii (bottom), iii (middle), and iv (top) in Si– (B) and Si+ (C) roots. The left panels show a longitudinal view, similar to the rectangle marked in (A). The right panels show cross sections, similar to the circle marked in (A). Arrows in zone ii indicate a putative initiation center of ASZ lignin. (D) SEM-BSE image of a cross section of a fresh adventitious root of sorghum grown in soil, showing an endodermis cell (e) and adjacent cortex cell (c). The developed silica aggregate in the ITCW (arrow) shows cell wall layers (darker areas). Scale bar=5 μm. Silica coupled to lignin polymerization may affect cell wall oxidative levels The variation in autofluorescence coupled with silicic acid availability and silica deposition indicates that Si influences the cell wall chemistry. Specifically, our work demonstrates a strong effect of Si on lignin structure or composition. Control over lignin composition and its incorporation into the cell wall are major aims in developing crops for biofuels and fodder. The polymerization of lignin monomers into the functional polymer occurs in the cell wall, as a result of the activity of reactive oxygen species (ROS) (Wang et al., 2013). The ROS are patterned by the localization of ROS-producing and manipulating enzymes on the cell membrane. Lignin monomers are suggested to be added to the growing polymer in a combinatorial manner, and the monolignols are suggested to diffuse quickly through the cell membrane and in the cell wall. This model suggests a uniform and random incorporation of varied monolignols into the cell wall. There are examples of variation in the composition of cell wall-lignin with time (Perkins et al., 2019). Our data demonstrate variation in lignin on a size scale of microns within a single cell wall layer. We found that at the aggregate formation site ASZ lignin accumulated, surrounded by non-modified lignin, which was linked to hemicellulose through ferulic acid (Figs 5 and 7). This requires a mechanism to control highly local spatial variations in monolignols. Our results support the existence of putative monolignol active transporters bound at specific locations on the cell membrane. ATP-binding cassette (ABC) transporters that may actively export specific monolignols (Perkins et al., 2019) could be involved in the formation of root silica aggregates. Silicic acid and silica availability correlate with a reduction of ROS activity in various systems (e.g. Liang et al., 2003; Gong et al., 2005; Fleck et al., 2011; Mehrabanjoubani et al., 2019). Our results suggest that Si also affects the chemistry of lignin. Further molecular research is needed to test whether the formation of silica in the cell wall affects ROS balance and results in changes in the level of oxidative stress in tissues. The control over the localized formation of silica aggregates uncovered in this work indicates an important biological role for silica in roots. Understanding silica deposition at the root is a key step towards understanding the role of Si in plant biology. Supplementary data Supplementary data are available at JXB online. Fig. S1. SEM-BSE micrograph of a cross-section of a Si– root stripped of its cortical tissues. Fig. S2. SEM-BSE micrograph of the ITCW of Si root, treated with acidic bleach. Abbreviations: Abbreviations: ASZ active silicification zone BSE back-scattered electrons EDX energy-dispersive X-ray ITCW inner tangential cell wall ROS reactive oxygen species Acknowledgements NZ thanks Robert H. Smith Program for a scholarship. 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