Plant and Soil

, Volume 355, Issue 1, pp 1–16

Roles of root border cells in plant defense and regulation of rhizosphere microbial populations by extracellular DNA ‘trapping’

Authors

    • Department of Soil, Water and Environmental SciencesUniversity of Arizona
  • Gilberto Curlango-Rivera
    • Department of Soil, Water and Environmental SciencesUniversity of Arizona
  • Zhongguo Xiong
    • Division of Plant Pathology and Microbiology, School of Plant SciencesUniversity of Arizona
  • John O. Kessler
    • Physics DepartmentUniversity of Arizona
Marschner Review

DOI: 10.1007/s11104-012-1218-3

Cite this article as:
Hawes, M.C., Curlango-Rivera, G., Xiong, Z. et al. Plant Soil (2012) 355: 1. doi:10.1007/s11104-012-1218-3

Abstract

Background

As roots penetrate soil, specialized cells called ‘border cells’ separate from root caps and contribute a large proportion of exudates forming the rhizosphere. Their function has been unclear. Recent findings suggest that border cells act in a manner similar to that of white blood cells functioning in defense. Histone-linked extracellular DNA (exDNA) and proteins operate as ‘neutrophil extracellular traps’ to attract and immobilize animal pathogens. DNase treatment reverses trapping and impairs defense, and mutation of pathogen DNase results in loss of virulence.

Scope

Histones are among a group of proteins secreted from living border cells. This observation led to the discovery that exDNA also functions in defense of root caps. Experiments revealed that exDNA is synthesized and exported into the surrounding mucilage which attracts, traps and immobilizes pathogens in a host-microbe specific manner. When this plant exDNA is degraded, the normal resistance of the root cap to infection is abolished.

Conclusions

Research to define how exDNA may operate in plant immunity is needed. In the meantime, the specificity and stability of exDNA and its association with distinct microbial species may provide an important new tool to monitor when, where, and how soil microbial populations become established as rhizosphere communities.

Keywords

Root border cellsMucilageRoot capExtracellular DNA (exDNA)Root exudatesRhizosphere colonization

Abbreviations

exDNA

Extracellular DNA

DAPI

4′,6-diamidino-2-phenylindole

One of the take-home messages is that spatial and temporal variability act to confound root research (Zobel and Wright 2005). There is an urgent need to develop new approaches and methods for probing rhizodeposition (Jones et al. 2009).

Background

Critical needs for sustainable practices in agriculture have been considered in many excellent articles (e.g. Brady and Weil 2010; Compant et al. 2005; Donato et al. 2010; Pinton et al. 2007; Sylvia et al. 1998; Zobel and Wright 2005). The root-soil interface is a target where positive changes can yield stable improvement in fertility, water use, and disease control leading to increased crop productivity with reduced damage to the environment (Bruehl 1987; Gilbert et al. 1996; Marschner et al. 2011; Rovira 1991; Schroth and Snyder 1961; Uren 2001). Efforts to apply biological control to root systems have been a focus of interest for decades with promising results and progress in understanding mechanisms (Handelsman and Stabb 1996; Hirsch 2004; Loh et al. 2002; Morris and Monier 2003; Pierson and Pierson 2007; Weller 1988; Zentmyer 1963). Of special interest are carbon allocation to the root and its delivery to the soil environment (Curl and Truelove 1986; Kuzyakov 2001; Lynch and Whipps 1990). If exudates control microbial growth, then controlling the composition, timing, and localization of root exudation would seem to be a reasonable approach to stimulate the growth of beneficial microorganisms at the expense of pathogens (Bednarek et al. 2010; Broeckling et al. 2008; Liu et al. 2005).

Unfortunately, despite ever-increasing precision in measuring carbon deposition and microbial colonization in the rhizosphere, the goal of developing predictive models, let alone controlling the process for crop improvement, has eluded researchers (Bowen and Rovira 1976; Cooper and Rao 2006; Darrah and Roose 2001; Handelsman 2004; Hinsinger 2001; Hinsinger et al. 2011; Luster et al. 2009). Apart from the extremes of environment and composition encountered in soils, the process of root exudation per se, as detailed below, is an intrinsically dynamic process that can be difficult to predict even under controlled conditions (Brady and Weil 2010; Lynch and Whipps 1990; Watt et al. 2006). Here we describe challenges and opportunities presented by the recent discovery that extracellular DNA (exDNA) is a component of exudates whose delivery into the rhizosphere is controlled by metabolically active cells at the root apex.

Lots of exudates at the root tip, not much microbial colonization: why?

Microbial growth in the rhizosphere, by definition, is increased relative to that in bulk soil (Rovira 1969). This phenomenon is attributed to the plant’s release of nutrient-rich exudates that can support the growth of diverse microbiota. Therefore, regions of the root that release more exudates might be predicted to support a corresponding increase in microbial growth relative to that in other regions. The root cap has been reported to be a primary source of exudate in experiments using diverse species and conditions (Dennis et al. 2010; Jones et al. 2009; Lundegardh and Stenlid 1944; Lynch and Whipps 1990; McDougall and Rovira 1970; Odell et al. 2008; VanEgeraat 1975; Wood 1967). In direct measurements from whole roots of young legume seedlings grown in hydroponic or plate culture under aseptic conditions, for example, more than 90 % of the total fresh or dry weight derives from the root cap (Griffin et al. 1975; Gunawardena et al. 2005). Therefore it would seem reasonable to predict that root exudate-stimulated microbial populations would predominate at the root cap under more complex conditions.

Instead, root caps of cereals, legumes, and other agronomically important species repeatedly have been found to be free of infection and colonization. In field-grown wheat Foster et al. (1983) reported that, ‘Unlike the rest of the root surface, the root cap as seen in scanning electron micrographs is generally quite devoid of microbial colonies.’ On tomato roots inoculated with Fusarium, ‘the root cap is not an important site of colonization’ (Lagopodi et al. 2002). On tomato inoculated with Pseudomonas fluorescens, ‘the root cap was always devoid of bacteria’ (Gamalero et al. 2005). Similar results occurred on maize root caps inoculated with P. fluorescens, but upon removal of root caps colonization of the apex developed (Humphris et al. 2005). On pea roots inoculated with spores of pathogenic fungi, then incubated in warm, moist conditions, the root cap remains sterile despite being ensheathed within a mantle of fungal hyphae (Gunawardena and Hawes 2002). Newly synthesized plant cells like those in the region of elongation are more susceptible to infection than older tissue with lignified cell walls (Hawes et al. 2000). Because root caps also are comprised of newly synthesized cells generated by meristems in the root apex, this was an especially surprising observation (Curlango-Rivera and Hawes 2011). New insight into the nature and function of root cap defense systems may yield an answer to this long-standing mystery: Sometimes, the carbon-based ‘exudates’ may act to trap, immobilize and inhibit microbial growth rather than serving as a passive nutrient base.

Extracellular DNA (exDNA) and protein in root tip defense

The recognition that exDNA is a key component of root exudates involved in border cell ‘extracellular trapping’ (Hawes et al. 2011) followed a long history of clues whose significance was overlooked until Brinkmann et al. (2004) documented the importance of exDNA in mammalian defense. VanEgeraat (1975) documented that the primary source of root exudates from young healthy seedlings under laboratory conditions is the root apex. Seedlings were placed onto damp filter paper for 24 h, then removed and the paper was dried and sprayed with ninhydrin (2,2-dihydroxyindane-1,3-dione) which reacts with lysine present in peptides and proteins. Positive reactions were limited to sites where root caps had been in contact with the filter paper. In older seedlings, an additional source is the site of lateral root emergence from the pericycle. However, chromatographic profiles of the material released from these natural wound sites are similar to those of root extracts, while profiles of material released from the root cap are distinct. As VanEgeraat (1975) recognized, ‘The process by which compounds are exuded from the root tip region is completely different from the release following damage of the root....Exudation by the root tip might be more selective so that certain specific compounds would be liberated.’

This prediction proved correct, despite the longstanding presumption that apart from a high molecular weight ‘slime’ or mucilage secreted from root caps, exudates from root tips primarily are the product of cytoplasmic contents leaking from dead ‘sloughed’ cells (Esau 1967; Levy-Booth et al. 2007; Voeller et al. 1964). Synonyms for ‘sloughed’ are ‘putrid’ and ‘gangrenous.’ Border cells, once termed ‘sloughed root cap cells,’ instead are metabolically active cells which exhibit host specific susceptibility and resistance to infection (Goldberg et al. 1989; Sherwood 1987). The border cell gene expression profile is distinct from that of progenitor cells in the root cap but parallel across diverse species (Brigham et al. 1998; Wen et al. 2008). Two-dimensional gel electrophoresis of proteins synthesized by the root cap during a 1-h test period (Fig. 1a) also yielded a profile markedly distinct from that of border cells (Fig. 1b) (Brigham et al. 1995). Most surprising was that the profile of proteins extracted from intact border cells (Fig. 1b) was markedly similar to that of a secretome with >100 proteins synthesized and exported during the same experiment (Fig. 1c). Extracellular proteins were found to play a key role in defense of the root tip: when treated with protease at the time of inoculation with spores of a pathogenic fungus, the normal resistance to root tip infection was abolished (Wen et al. 2007b). Among the proteins were antimicrobial enzymes long known to be associated with plant and mammalian defense (De-la-Pena and Vivanco 2010; Kwon et al. 2008). Therefore, it was perhaps not surprising that their destruction altered the normal root defense processes. Treatment with protease also resulted in disintegration of a surrounding mucilage layer and release of bacteria within the layer (Wen et al. 2007a). These data support the suggestion by Matsuyama et al. (1999) that proteins may play a role in the structural integrity of the matrix, even though protein comprises only a small fraction of the matrix composition (Bacic et al. 1986; Chaboud and Rougier 1990; Moody et al. 1988).
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Fig. 1

The root cap secretome. After a 1-h period of labelling, large differences in protein profiles from (a) root caps and (b) border cells of Pisum sativum L. are evident using two-dimensional gel electrophoresis. c More than 100 proteins are synthesized and exported from living cells during the test period (from Brigham et al. 1995). Examples of proteins common to root caps and border cells (black arrows), specific to root caps (open triangles), or specific to border cells (closed triangles) are denoted. Plant Physiol 143:773–783 (www.plantphysiol.org) “Copyright American Society of Plant Biologists”

The discovery that histone H4 was among the proteins synthesized and exported into the extracellular matrix was a surprise, given the long-established role of histones in assembly of genetic material inside the cell (Wen et al. 2007a). However, emerging research provided insights into alternative functions of histones, including potent antimicrobial activity in the extracellular environment (Bergsson et al. 2005; Kawasaki and Iwamuro 2008; Patat et al. 2004; Wang et al. 2009; Xu et al. 2009). Of special interest were reports of a role for histones in extracellular chemotaxis and ‘trapping’ of pathogens by neutrophils in the mammalian immune response, because a very similar process occurs in border cells in response to plant pathogens (Gochnauer et al. 1990; Goldberg et al. 1989; Gunawardena et al. 2005; Hawes and Pueppke 1987; Hawes et al. 1988; Zhu et al. 1997). ‘Neutrophil extracellular traps’ (NETs) were first described by Zychlinsky and coworkers (Brinkmann et al. 2004), and now have been implicated in defense against diverse pathogens and other aspects of immune responses in mammals (Abdallah et al. 2012; Amulic and Hayes 2011; Brinkmann and Zychlinsky 2007; Harding and Kubes 2012; Medina 2009; Mitroulis et al. 2011; Park et al. 2012; Urban et al. 2006; Wardini et al. 2010; Wen et al. 2012; Yost et al. 2009; Young et al. 2011). As with the border cell slime layer (Fig. 2), NET formation can occur rapidly in response to specific signals, in the absence of cell death (Pilszik et al. 2010). Experiments therefore were carried out to determine (1) whether the presence of extracellular histone surrounding border cells, like neutrophils, is associated with exDNA; and if so, (2) to determine if enzymatic degradation of border cell exDNA, like NET exDNA, interferes with resistance to infection (Gunawardena and Hawes 2002). The results of these experiments revealed that, like the plant proteins exported from the root cap and border cells (Brigham et al. 1995), plant DNA is synthesized and exported into the root cap extracellular matrix during a 1-h period when no cell death occurs (Wen et al. 2009). Initial sequence analysis revealed that the exDNA structure is related to nuclear DNA, but is enriched in repetitive sequences. When this exDNA was degraded by addition of DNase I concomitant with the inoculation by a pathogenic fungus, the frequency of root cap infection increased from a mild local necrosis in <5 % of inoculated roots to 100 % infection, with rotting of each root tip and proliferation of fungal hyphae (Wen et al. 2009). As in exDNA-based extracellular trapping in mammals, the root tip resistance to fungal infection is associated with aggregation of the fungus and inhibition of its growth (Gunawardena et al. 2005; Medina 2009). The extracellular trapping phenomenon is host-microbe specific, with no aggregation or growth inhibition of nonpathogenic fungi (Gunawardena and Hawes 2002; Jaroszuk-Scisel et al. 2009).
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Fig. 2

Host specific binding of bacteria within an inducible extracellular ‘trap’ produced by border cells. a Within seconds of adding the pathogen Agrobacterium tumefaciens to border cells of a host species (P. sativum L.), chemotaxis toward the cells is evident. The large arrow denotes the leading edge of the bacterial swarm; the small arrow denotes border cell sample. b Within 1–2 h, interconnecting strands of bacteria (white arrow) develop between border cells (black arrow). c Trapping of bacteria is evident within the surrounding mucilage of individual border cells of the host species, pea (arrow); d No chemotaxis occurs in response to cells from a nonhost species (Avena sativa L.), and bacteria are excluded from rather than trapped within the surrounding mucilage layer (visualized using India ink, which does not penetrate the mucilage) (arrow). No mucilage layer is induced when a nonpathogenic strain of E. coli is added to border cells of pea (e) or oats (f), and no trapping occurs at the cell surface. Scale bar: 15 μm

Host specific chemotaxis and extracellular trapping of pathogens by border cells was described previously, but was presumed to involve aspects of pathogenesis, not defense (Goldberg et al. 1989; Hawes and Pueppke 1987; Hawes and Smith 1989). Agrobacterium tumefaciens chemotaxis toward border cells of a host species was measured using swarm agar assays (Fig. 2a) (Hawes et al. 1988) or direct microscopic observation (Fig. 2b,c). Within hours, strings and strands of immobilized bacteria develop (Fig. 2b). Bacteria adhere to the surface in a layer that is impervious to removal by washing in water (Fig. 2c). Adding the plant pathogen to border cells of a nonhost species triggers no chemotaxis or attachment within the surrounding mucilage (Fig. 2d). The human pathogen E. coli added to border cells (Fig. 2e, f) was not associated with chemotaxis, attachment, or production of a mucilage layer in either plant species. Minimal growth can be measured in remaining unattached bacteria or in bacteria growing on mucilage as a sole carbon source, but whether the trapped pathogenic bacteria are viable is unclear (Knee et al. 2001; Zhu et al. 1997). Similar patterns of specificity were reported in association between maize border cells and bacterial species including Rhizobium, E. coli, Pseudomonas, Bacillus, Streptomyces and Cytophaga (Gochnauer et al. 1990). It will be of interest to examine the role of exDNA in this phenomenon, and to explore the possibility that clusters and strings of viable but not culturable (VBNC) colonies found in the rhizosphere might be related to exDNA based trapping (Gamalero et al. 2004).

The high molecular weight polysaccharide-based mucilage exported from root caps has been studied in various species but DNA has not been included in analyses to date (Bacic et al. 1986; Chaboud and Rougier 1990; Foster 1981a, b, 1982; Jones and Morre 1973; Knee et al. 2001; Lynch and Staehelin 1995; Miki et al. 1980; Newcomb 1967; Oades 1978; Read et al. 1999; Sealey et al. 1995; Watt et al. 1993). Therefore details of how exDNA is synthesized, exported, and integrated into the extracellular matrix remain to be established (Hawes et al. 2011). However, the presence of nucleic acids among exudates of healthy roots was reported (Curl and Truelove 1986; Fries and Forsman 1951; Lundegarth and Stenlid 1944; Stenlid 1944), and its active synthesis and export into the extracellular matrix of the root cap periphery also were documented (Phillips and Torrey 1971). Using the fluorescent stain DAPI, which binds to A-T rich strands of DNA and can pass through intact cell membranes to reveal DNA within cells or outside the cell boundaries (Kubista et al. 1987), exDNA is readily detected within border cell mucilage (Wen et al. 2009). In the presence of stimulating bacteria, DAPI staining occurs within border cells, throughout the surrounding expanded mucilage layer (Fig. 3a) and within trapped bacteria (Fig. 3a, arrow). Staining border cell populations and associated mucilage with SYTOX green, a high-affinity nucleic acid stain which is not taken into living cells, reveals extracellular material ranging from strands (Fig. 3b) to distinctive structures (Fig. 3c). These structures are similar in appearance to those produced by neutrophils (Patel et al. 2010; Pilszik et al. 2010).
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Fig. 3

exDNA from the root tip of pea. a DAPI staining of a pea border cell with bacteria trapped within the surrounding mucilage layer. Scale bar: 15 μm. b SYTOX green staining of border cell exDNA strands and c other structures. Scale bar: 10 μm. 3A, 3 C, photos by Fushi Wen. 3B, photo by Sarah O’Connor

Questions of particular interest are the nature of the exDNA structure(s) involved in trapping and how they might interface with other polymers within the root cap mucilage (Bacic et al. 1986; Knee et al. 2001). One possibility is that the ‘stickiness’ of DNA alone might be sufficient to trap added microorganisms. If so, then addition of DNA alone would be predicted to result in trapping. No such result occurred upon addition of salmon sperm DNA or pea genomic DNA to microbes (Wen et al. 2009). An alternative hypothesis is that distinct sequences organized in specific structures are required. In support of this model are observations by Van’t Hof and colleagues (Van’t Hopf and Bjerknes 1982; Kraszewska et al. 1985), who described a distinct class of DNA produced by P. sativum root caps during the G2-M transition, the point in the root cap meristem cell cycle when border cell separation occurs (Brigham et al. 1998). Like root cap exDNA (Wen et al. 2009), this ‘extrachromosomal DNA’ is related to nuclear DNA but is distinguishable based on the prevalence of repetitive sequences (Kraszewska et al. 1985). The programmed delivery of characteristic exDNA patterns as an integral component of the matrix could provide a tool to examine underlying patterns of rhizosphere carbon deposition and microbial colonization and allow progress toward exploiting the system for crop improvement. Factors known to influence border cell delivery are summarized below.

Factors controlling delivery of exDNA-based traps from root caps

Border cell populations

The presence of DNA from plants and other organisms in the soil is well established (Izano et al. 2008; Vlassov et al. 2007; Whitchurch et al. 2002). Plant exDNA has been presumed to be derived by leakage from dead cells (Levy-Booth et al. 2007). The discovery that secretion of exDNA from root caps instead is a component of a complex, inducible, and carbon-expensive defense mechanism may be useful in tracking as well as modelling rhizosphere community structure. The programmed separation of cells from the root cap was long presumed to be a product of continuous cell cycle activity within the root cap meristem in parallel with such activity in the apical meristem (e.g. Clowes 1971; Whipps and Lynch 1983). If correct, then a continuous detection of exudates at the tip would be a predicted result. Direct observations of rhizosphere structure even under controlled conditions do not support this paradigm (Iijima et al. 2003). The viability and number of border cells that a root cap can release daily are conserved within families and can range from 0 to 10,000 cells a day (Hawes et al. 2003; Hawes and Pueppke 1986). For a given root, the process of root cap turnover is not continuous but instead is induced or repressed in a species- and genotype- specific manner in response to endogenous and environmental signals (Brigham et al. 1998; Ponce et al. 2005). Therefore, when seedlings are grown under identical conditions side by side in petri dishes, the delivery of mucilage and border cells can vary from nothing to intermittent clumps to a continuous sheath surrounding the root from base to tip (Fig. 4). The variation is illustrated schematically because even with direct microscopic observation on sterile plates the differences can be difficult to detect (Fig. 4, inset photos). Some species exhibit border cell specific expression of pigmented metabolites which provide a convenient marker for cell dispersal (Brigham et al. 1999). Thus, Saccharum officinarum, Sorghum vulgare and Lithospermum erythrorhizon have pink, purple, and red border cells, respectively. This pigmentation facilitates recognition of rhizosphere distribution patterns that otherwise would be obscure (Fig. 4, inset center). Variation in root exudation and rhizosphere colonization has been proposed to be a major obstacle to agronomic application of promising discoveries like biological control (Cooper and Rao 2006; Sylvia et al. 1998). Understanding factors controlling carbon delivery via border cells may be key to monitoring and controlling rhizosphere community structure (Lee and Hirsch 2006; Smucker and Erickson 1987).
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Fig. 4

Variation in border cell delivery from root caps under controlled conditions. The length of the root at time zero is denoted with white arrows. As roots elongate side by side in the same plate of water agar, the presence of border cells detectable by direct microscopic observation ranges from none (inset photo, left) to intermittent clumps (inset photo, center) to a continuous robust sheath (center). (Plant Physiol 119:417–428, (www.plantphysiol.org) Copyright American Society of Plant Biologists). Inset, center: Border cells released from tips of elongating root of Lithospermum erythrorhizon. The border cells from this species express a red pigment, shikonin, which facilitates detection of their presence at intervals along the root surface (arrows). (Plant Physiol 119:417–428, (www.plantphysiol.org) Copyright American Society of Plant Biologists)

For roots of any given plant, one major factor controlling border cell release is the availability of free water (Fig. 5) (Odell et al. 2008). Roots of legumes, cereals, cucurbits and most other crop species are programmed to produce a species-specific number of cells (Hawes and Pueppke 1986). When that number has accumulated on the cap periphery the same set may remain on the root for an extended period without any new cells being produced. This appears to result from the accumulation of an extracellular signal within the surrounding mucilage to a level that suppresses cap turnover except when diluted in water (Brigham et al. 1998). The properties of the mucilage are such that it can hold 1000X its weight in water (Guinel and McCully 1986). Yet even at 99 % humidity, in the absence of free water, the mucilage remains ‘dry’ like a sponge without moisture (Fig. 5a) and high-resolution microscopy is necessary to detect the ensheathed border cells (Fig. 5a, inset). Upon addition of water--including, for example, a drop resulting from condensation on the inside of a petri plate falling onto the root--the mucilage immediately expands (Fig. 5b) and border cells are dispersed into suspension (Fig. 5c). It is important to note that the drop of water not only causes the dissociation of the existing group of ca 4,000 cells from the cap periphery, but also triggers renewed cell cycle instantaneously (Brigham et al. 1998). Within 5 min, mitosis increases within the root cap meristem, and dozens of new cells emerge from the periphery. Activation of the quiescent center also occurs, and cell production proceeds until a new set has accumulated within 24 h (Ponce et al. 2005). It seems obvious that within the soil environment, where a continuous film of free water at the root tip would be intermittent for most crops in most conditions, this factor alone could account for much of the variability in root tip carbon deposition that occurs.
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Fig. 5

Instantaneous swelling and dispersal of P. sativum border cells in response to immersion in free water. a At 99+% humidity, the root tip is smooth, and the presence of border cells is undetectable except with scanning electron microscopy (inset). Scale bar: 0.5 mm. b Addition of a droplet of water results in swelling of border cells away from the tip within 10–15 s. Scale bar: 0.5 mm. c Gentle agitation of the plate by tapping one side results in immediate dispersal of border cells into the suspension, in the direction of the applied force. Scale bar: 0.5 mm. Staining with the vital stain fluorescein diacetate, which only accumulates inside living cells, reveals border cell viability of 95–100 % (inset). Scale bar: 10 μm

Other factors that can vary the number of border cells and associated products released into the rhizosphere, include soil type, physical abrasion, day length, root age and growth rate (Iijima et al. 2000, 2003; Odell et al. 2008; Somasundaram et al. 2008; Wuyts et al. 2006). Sodium fluoride added to wheat roots can stimulate changes in number of border cells and in level of protein secretion (Bozhkov et al. 2007). Carbon dioxide, aluminum, boron, and plant pathogens stimulate changes in border cell production in a plant species- and genotype-specific manner with distinct responses at different developmental stages (Cannesan et al. 2011; Chen et al. 2008; Liu et al. 2007; Miyasaka and Hawes 2001; Pan et al. 2004; Tamas et al. 2005; Zhao et al. 2000; Zhu et al. 2003). For example, increased carbon dioxide inhibits border cell production in P. sativum during germination, but results in increased cell production in seedlings (Fig. 6). Border cell production in Medicago sativa seedlings, in contrast, is impervious to similar changes in carbon dioxide (Zhao et al. 2000). Continuous culture of roots in high concentrations of certain sugars and secondary metabolites results in marked increases in mucilage production by maize roots (Jones and Morre 1973; Knudson 1917). Transient exposure of roots to metabolites including rhamnose, caffeine, and flavonoids for several minutes, a condition more likely to occur under natural conditions, can specifically induce or repress border cell production without affecting rate of root growth (Curlango-Rivera et al. 2010). Altered expression of genes controlling cell cycle or cell wall solubilization at the cap periphery results in altered border cell production, and transient changes in their expression due to diverse environmental signals could influence the process as well (Wen et al. 1999; Woo et al. 2004). A new study reporting an ‘extraordinary sheath’ of material triggered on roots of Acacia magnum grown in hydroponic culture, highlights the importance of understanding factors controlling this avenue of carbon deposition and their impact on rhizosphere structure (Endo et al. 2011).
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Fig. 6

Effects of carbon dioxide on border cell separation from root tips (arrows) of P. sativum seedlings. a Cells from a single root tip observed with a dissecting microscope after 3 days in (A) ambient (0.03 % CO2 vs 21 % O2); or (b) 6 % CO2 vs 15 % O2). Increased O2 alone had no effect on cell production. From Plant Physiology 122:181–188, used with permission. (www.plantphysiol.org) Copyright American Society of Plant Biologists

Single cells

Morphology of border cell detachment from the cap periphery can range from a population of single cells in suspension to finger-like strands of cells to an entire root cap (Endo et al. 2011; Hamamoto et al. 2006; Vicre et al. 2005; Wen et al. 2008). The significance of these variations with respect to exDNA-based trapping is unknown, but the variation in amount and composition of carbon-based material can be substantial even on a single-cell basis. For many years, border cells were called ‘sloughed root cap cells’ to reflect the presumption that delivery of the cell populations must reflect a process of falling away from the root as a consequence of cell death (e.g. Uren 2001). This notion prevailed, despite repeated documentation that the cells from most species are metabolically active as they detach from the root cap and can survive for extended periods in liquid culture (Caporali 1983; Gautheret 1933; Hawes and Wheeler 1982; Stubbs et al. 2004). Knudson (1919) reported that border cells released from Zea mays or P. sativum grown in hydroponic culture, with or without glucose, remained 100 % viable for more than one month. Even more surprising was the observation that the cells export enzymes and other proteins (Rogers et al. 1942) and can remain metabolically active after detachment into the soil environment (Vermeer and McCully 1982). Continued secretion of mucilage from border cells can occur for days after detachment from roots grown in soil (Hawes and Brigham 1992; Hawes et al. 1998). Like white blood cell ‘granules’, border cells contain abundant storage particles which may provide energy for survival and response to signals in the extracellular environment (Feldman 1985; Newcomb 1967).

The mucilage produced by individual border cells after separation from the root cap also is a dynamic process. An increase in the diameter of the mucilage layer is induced almost instantaneously in a species- and genotype-specific manner in response to exposure to bacteria (Figs. 2, 3), fungi (Wen et al. 2009), and aluminum (Miyasaka et al. 2000). Border cells from pea, for example, can form aggregates containing hundreds of cells and associated mucilage (Fig. 7a), or exist as isolated cells with variable layers of surrounding mucilage (Fig. 7b) (Wen et al. 2007a). Given that such variation can occur in controlled environments and that each cell can trap thousands of bacterial cells, the potential for creating variable islands that confound efforts to measure carbon deposition and its impact on rhizosphere colonization in the soil, is obvious. With recognition of the ‘trapping’ function of border cells, on the other hand, these seemingly inexplicable phenomena may be easier to understand. The observation by Guinel and McCully (1987) that border cells can continue to expand after detachment from the root as single cells, also is less surprising in the context of their proposed functions in ‘border patrol.’ If border cells trap heavy metals and pathogens and control the growth of deleterious microorganisms in the vicinity of plant roots, then a capacity to achieve an increased surface area would be a predictable benefit to the plant rather than an egregious waste of fixed carbon (Fig. 8).
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Fig. 7

Variation in aggregation of detached border cell populations ranges from (a) a cohesive mass containing hundreds of border cells to (b) individual cells. Mucilage layers, detected by staining with India ink which is excluded, are present on viable cells but disintegrate rapidly after cell death (block arrow). Scale bar: 20 μm

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Fig. 8

Border cell expansion of the volume of a single cell (black arrows denote each end of the cell) by >10-fold 7 days after detachment from the root cap. The nucleus (white arrow) and cytoplasmic strands are evident within the living cell. Inset: The original size of border cells within this sample is illustrated by showing for comparison a cell within the same population which died before any growth occurred. Scale bar: 30 μm

In addition to proteins, DNA and polysaccharides, the root cap and border cell exudates include primary and secondary metabolites that function in signalling and recognition of beneficial as well as pathogenic microbes (e.g. Baluska et al. 1996; Graham 1991; Maxwell and Phillips 1990; Peters and Long 1988). The mixture also contains feedback signals that may influence rate and direction of root growth and development (Baluska et al. 1996; Caffaro et al. 2011; Moore and Fondren 1986). The potential for creating changes in the composition of border cell products has been demonstrated by studies of cotton engineered to resist insect damage by expression of crystal (CRY) proteins from Bacillus thuriengensis. BT toxin is delivered through exudates of engineered plants into the soil where it can exhibit a half-life of up to 234 days (Saxena and Stotzky 2001; Tapp and Stotzky 1997). Direct measurements of Cry proteins revealed that roots of all genetically modified lines tested synthesize and export BT toxin, and that root caps, border cells and root mucilage are sources of this material (Knox and Vadakattu 2005; Knox et al. 2007). The environmental impact is not clear at this time, but the results suggest that reproducible changes in the soil environment already have been accomplished via changes in root cap delivery systems of genetically modified crops.

Conclusions

The discovery that exDNA plays a role in plant defense raises more questions than it answers, and additional research is needed before conclusions can be drawn regarding a general role in plant immunity. The new data do reinforce the premise that a simple model of nutrient rich material leaking from roots and feeding microbial growth in general is inadequate (De-La-Pena et al. 2010). The important role of metabolites secreted into the ‘apoplast’ has long been recognized (Brisson et al. 1994; Kwon et al. 2008). Understanding the nature and function of the ‘exudates’ delivered by the root cap may offer insights into how the natural immunity of the root cap might be extended to more vulnerable sites including the region of elongation, where most soilborne pathogens initiate infections (Hawes et al. 2000). The controlled delivery of exDNA may complement new tools available to define the structural and functional dynamics of the rhizosphere and its components in the interest of fostering sustainable methods for agriculture (Ceccherini et al. 2009; Levy-Booth et al. 2007; Pietramellara et al. 2009). If used in conjunction with holistic tracking methods that combine laboratory and field assessment (e.g. Knox et al. 2009), a goal of harnessing the plant’s ability to control root exudation and rhizosphere community structure may not be unrealistic (Atkinson et al. 1975; Knox et al. 2009; Liu et al. 2005).

Acknowledgements

We gratefully acknowledge support for our research in this area from the National Science Foundation (NSF# 1032339 to MCH and ZX) and the Department of Energy (DOE DEAC02-06CH11357 to JOK). We thank Dr. Virginia Rich for critical reading of the manuscript.

We dedicate this review to the memory of W. D. ‘Dietz’ Bauer.

Copyright information

© Springer Science+Business Media B.V. 2012