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Ecology |
4 Department of Land, Air, and Water Resources, University of California, Davis, California 95616-8627 USA
Received for publication 25 September 2007. Accepted for publication 3 January 2008.
ABSTRACT
Rising CO2 levels in the atmosphere have drawn attention to the important role of soil in sequestering carbon. This project goal was to quantify soil carbon deposition associated with border cell release and exudation from root growth zones. Carbon was measured with a Carlo Erba C/N analyzer in soil from the rhizosphere of mature grasses and, in separate experiments, in soil collected around root growth zones. Root border cells in "rhizosphere soil" (silica sand) were counted using a compound microscope after soil sonication and extraction with surfactant. For sand-grown Bromus carinatus, Zea mays, and Cucumis sativus, young seedlings (with roots shorter than 2 cm) released thousands of border cells, while older root tips released only hundreds. For a variety of native annual and perennial grasses and invasive annual grasses (Nassella pulchra, B. carinatus, B. diandrus, B. hordeaceus, Vulpia microstachys, Aegilops triuncialis, Lolium multiflorum, Zea mays), the rhizosphere of mature root systems contained between 18 and 32 µg C g–1 sand more than that of the unplanted controls. Spatial analysis of the rhizosphere around the cucumber growth zone confirmed C enrichment there. The root tip provided C to the rhizosphere: 4.6 µg C in front of the growing tip, with the largest deposition, 20.4 µg C, to the rhizosphere surrounding the apical 3 mm (root cap/meristem). These numbers from laboratory studies represent the maximum C that might be released during flooding in soils. Scaling up from the organ scale to the field requires a growth analysis to quantify root tip distributions in space and time.
Key Words: border cells • carbon • carbon deposition • growth • rhizosphere • root • soil carbon deposition
Rising levels of CO2 and associated global warming trends are leading to intense interest in carbon cycles and the role of soil in sequestering carbon. It has long been known that a large fraction of carbon recently fixed in leaves is rapidly (on the time scale of minutes to hours) released to the rhizosphere (e.g., Horwath et al., 1994
). Carbon released from the root to the rhizosphere, also termed rhizodeposition, may account for 40% of the carbon allocated to roots of annual species and as much as 70% in Pseudotsuga menzeseii (Douglas-fir; Lynch and Whipps, 1990). Carbon-containing exudates are particularly copious at root tips and at points of initiation of branch roots (e.g., McDougall and Rovira, 1970). The root tips, including root caps, meristems, and elongation zones, secrete sugars, organic acids, and amino acids as well as more complicated compounds such as phytosiderophores. Mucilage that contains polysaccharides with a diverse array of glycosyl residues, including arabinogalactan proteins, is produced copiously by root caps and cortical cells (Bacic et al., 1986; Guinel and McCully, 1986; Knee et al., 2001). These carbon sources sustain the rich microflora of the rhizosphere. In a groundbreaking study, Jaeger et al. (1999) used microbial biosensors to show that roots of Avena barbata have a pattern of exudation that varies with developmental stage of the root. Sugars and organic acids are released to the rhizosphere by apical, young tissue, while amino acids, rich in nitrogen, are exuded by cells in older regions near the base of the root.
Another source of carbon is provided by border cells (Fig. 1), formed as part of the root cap and released from the exterior of the cap to live freely in the soil for a time (Hawes and Pueppke, 1986; Guinel and McCully, 1987). Border cells have been recognized as serving many important functions, including protection of the root tip from pathogenic fungi (Hawes, 1990; Wen et al., 2007). Recent quantitative studies have shown that roots of maize seedlings have hundreds of border cells at any given time (Iijima et al., 2000, 2003a, b, 2004). These published works suggest that, when the root tip grows from an unsaturated into a saturated soil stratum, it may provide a pulse of 3 µg of fixed C to the rhizosphere of the growth zone, representing perhaps 1 mg C per g soil in the boundary layer of soil next to the meristem. This rough calculation is the motivation for our study of C exudation from roots to their rhizospheres.
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Spatially averaged rhizosphere C of mature grasses grown in sand
Plastic pots approximately 900 cm3 were filled with 30-mesh silica sand. The pots were then densely seeded with a single species of the following California grassland native and perennial grass species (N = 3 replicates per species): Nassella pulchra (Hitchc.) Barkworth (purple needle grass; native perennial), Bromus carinatus Hook. and Arn. (California brome; native perennial), Vulpia microstachys (Nutt.) Munroe (small fescue; native annual), Aegilops triuncialis L. (barbed goatgrass; invasive annual), Lolium multiflorum L. (Italian ryegrass; invasive annual), B. diandrus Roth (ripgut brome; invasive annual), or B. hordeaceus L. (soft chess; invasive annual). Native grass seed was supplied by Hedgerow Farms, Winters, California; seeds of invasive grass species were collected locally. An agricultural crop grown throughout California, Zea mays L. (corn, supplied by Davis Lumber, Davis, California), was also examined. Control pots containing only silica sand were also established to monitor microbial C contributions to the sand. The pots were completely randomized in a Conviron CMP4030 growth chamber (Controlled Environments, Winnipeg, Manitoba, Canada). The growth chamber conditions were 80% humidity, oscillating 12 h temperature cycle from 21°C daytime maximum to 10°C nighttime minimum, and an illumination of 700 µmol•m–2•s–1 at bench height for 10 h a day (day) followed by 14 h of darkness (night). The pots were maintained near field capacity with full-strength Hoaglands solution (Taiz and Zeiger, 1998).
Thirty-seven days after planting, the plants were harvested. Aboveground biomass was clipped at the sand surface and discarded. The root–sand mass was then removed. It was noted that the roots of every species had thoroughly explored the entire pot volume and become root bound. Therefore, the sand within each pot was regarded as rhizosphere substrate. The substrate was thoroughly washed from the root mass into a metal tray and dried at 100°C. Rapid substrate drying (high temperature) was essential to prevent microbial proliferation in the sample. The cleaned root mass was retained for root length and biomass measurements. Root length was measured using a Comair rotating table root length scanner (Hawker De Havilland Victoria Ltd., Melbourne, Australia) and then dried in a drying oven at 60°C (adequate to measure root dry mass for calculations and for root C determinations). The dried rhizosphere substrate and root mass were weighed and then ground for C analysis. Ground rhizosphere substrate and root samples were analyzed with a Carlo ERBA C/N analyzer (Fusions Instruments, Milan, Italy).
Root border cell counting
Seedlings of Bromus carinatus, Zea mays, and Cucumis sativus L. ‘Straight Eight' (cucumber, an agricultural crop grown in California, seeds supplied by Davis Lumber, Davis, CA), were examined for root border cell sloughing. For this study, rhizotrons were constructed from 24.5 x 24.5 x 2.0 cm clear polystyrene bioassay plates (Corning Inc., Corning, New York, USA) by cutting away one of the 2.0 cm sidewalls to serve as the open top of the rhizotron. The former bottom half of the bioassay plates were bound together with large binder clips and oriented vertically to provide a narrow container with wide horizontal dimensions suitable for root growth observations. Rhizotrons were filled with silica sand, watered to field capacity with full strength Hoagland's solution, and planted with several seeds of a single species including B. carinatus, Z. mays, or C. sativus. Rhizotrons were inclined 30° from vertical in a Conviron growth chamber set to the environmental conditions described previously. Seedlings were grown for 3–5 d following germination.
Modified methods of Iijima et al. (2004) were used for border cell extraction and counts. At harvest, the rhizotrons were positioned horizontally, and the upper rhizotron wall was removed. Individual seedlings were gently removed, root length was measured, and the apical centimeter of root was excised and placed in a 1.5 ml centrifuge tube. Tween 80 solution (0.5 ml) was added to each centrifuge tube. Tubes were sonicated for 1 min, 1–2 drops of aqueous toluidine blue stain (3 x 10–5 g•ml–1) were added to each tube, and the tubes were placed on a shaker for 30 min to complete the cell staining process. Border cells had characteristic shapes for each species. Border cells were counted using a compound microscope and a Sedgewick-Rafter slide.
Carbon in the rhizosphere of the growth zone
Because of ease of cultivation and reproducible C exudation, Cucumis sativus was chosen for closer examination of spatial patterns of C secretion into the rhizosphere. Rhizotrons were slightly modified to facilitate harvesting of rhizosphere substrate. The former bottom half of the bioassay plates was filled with the moist silica sand, scraped to an even surface, and wrapped with a commercial plastic wrap stretched to prevent wrinkling on the sand surface. The former top of the bioassay plate was placed onto the sand-filled chamber, and the two halves were bound together with large spring clamps. Cucumis sativus seeds were planted in the rhizotrons and grown inclined in a growth chamber at conditions described previously. Root elongation rate was approximately 1 mm/h. Seedlings were grown for 3–5 d until the radicle had grown to a length of 1 or 4.5 cm. At harvest, the rhizotrons were positioned horizontally, the upper rhizotron wall was removed, and the plastic wrap was peeled away with care not to disturb roots or sand. A rhizosphere-harvesting device was constructed by cutting a drinking straw into a scoop with a half-cylindrical top 3 mm long and 4.8 mm inner diameter. The diameter of the rhizosphere-harvesting scoop includes the spatial extent over which C is expected to be exuded from the root (0.5 mm from the root; Kim et al., 1999; Nichol and Silk, 2001). A clean razor blade was placed in front of the root tip, and the half-cylindrical volume in front of the root was removed from the rhizotron and placed in a glass vial. Then a razor blade was used to slice the root and neighboring soil 3 mm behind the tip, and the half-cylindrical volume containing the root segment and its rhizosphere was harvested in a similar manner. Rhizosphere sand was collected separately by position from 10 roots to make a composite sample for each of the different positions. The rhizosphere samples were saturated with 2.5 ml deionized water and then sonicated for 1 min to free any loose border cells from the root segments. Rhizosphere samples were saturated with 2.5 ml deionized water and then sonicated for 1 min to free border cells from the root segments. The root segments were then removed and dried at 60°C. Rhizosphere substrate (extractant) was dried at 100°C to prevent microbial proliferation. Dried root segments and rhizosphere substrate were weighed. Rhizosphere substrate was ground for C assay as described previously. Five replicate samples were assayed per location for each of the two root lengths. The assays were repeated on three sets of roots, and the 15 replicates were used to obtain mean C and standard deviations.
RESULTS
Roots of all grass species deposited C into their rhizospheres
After 37 d of growth, all planted species had rhizosphere sand that was enriched with carbon relative to control sand.
As shown in Table 1, in units of µg C•g–1 sand, the enrichment ranged from 18 (for Nassella pulchra, a native bunchgrass with fine, long roots) to 32 (for Bromus diandrus and B. hordaceus) more C than in the control sand. The soil C deposition represented as much as 58 mg C•g–1 root (for the native perennial B. carinatus, a C3 plant) and as little as 13 mg C•g–1 root for Zea mays (a C4 plant that has a particularly wide primary root). The percentage of belowground C provided by exudates and sloughed cells ranged from 3% for Z. mays (where the large roots dominate the belowground C stores) to 13% for B. carinatus. These results are within the range reported for root exudation in other species, reviewed in Liebersbach et al. (2004) who also emphasized that environmental conditions, particularly soil moisture content, have a large effect on mucilage production, so large natural variation in C deposition is expected.
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; Nichol and Silk, 2001). The assay results displayed in Fig. 3 are useful for comparing spatial and developmental trends. Other C cycle relations are of interest and are displayed in Table 2 showing total C deposited around the harvested zone, C per unit root length, C per unit root biomass, and the estimated rhizosphere C concentration if the measured C has been deposited in a boundary layer of soil within 0.5 mm of the root.
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Stage-dependent border cell release
Border cells play important roles in rhizosphere ecology. Released into the soil by the higher plant, they have been shown to aid penetration of the soil (Iijima et al., 2003), promote growth of certain microorganisms, and also to exert ecologically important antibiotic activity against some common pathogenic fungi (Gochnauer et al., 1990; Hawes, 1990; Wen et al., 2007). It has long been recognized that the number of border cells released depends on species (Hawes and Pueppke, 1986; Hamamoto et al., 2006), and recent studies show variation also with soil strength (Iijima et al., 2003). Previous work has also shown that a minimum developmental stage must be achieved, corresponding to a root length 5 mm in pea, before border cells are released (Hawes and Lin, 1990). The strong developmental trend reported here was observable in all species examined and suggests an important functional role. The large number of border cells produced during the young stages of root development may provide protection during the vulnerable period of radicle penetration of the soil crust.
Spatial and temporal patterns of carbon release in the soil
In the 1960s, work with 14CO2 supplied to wheat plants revealed that significant C is exuded by the apical centimeter, little exudation (or even net uptake) occurs in older regions of the primary root, and the most exudation (10 times that from the apex) occurs in the region of lateral root emergence and is associated with lateral root tips (McDougall and Rovira, 1970). Until recently, most studies of root exudation were performed in solution culture or on filter paper. Controlled conditions were necessary for the study of the nature and quantity of the exudates and to work out the physiology and biochemistry of the plant metabolism and the effects of environmental variation. However, an understanding of C cycling in soils requires attention to spatial and temporal aspects of root architecture and function relative to soil strata. Recently published methods for isolating and counting border cells in soils (Bengough et al., 2001; Iijima et al., 2000, Iijima et al., 2004) inspired our efforts to extend in vitro studies to natural environments. The sand culture reported here, while much less complex than a real soil, does offer a more natural mechanical impedance and pore structure than does culture in solution or on germination paper.
Because our assays involve extracting soil with aqueous solutions, our results represent the C that might be released as a pulse during rainfall, irrigation, or growth into a saturated soil layer. Our calibration trials with Zea mays roots (not shown) gave results for border cell and C release that were quantitatively similar to results published by Iijima et al. (2000). However, other approaches are necessary to estimate the C exuded into the soil during growth in soils at field capacity. Iijima et al. considered the timing of replacement of the root cap by cell production and displacement as might occur during quasisteady state conditions. Their estimates for C released by root tips growing in nonsaturated conditions are an order of magnitude lower than those reported here. This discrepancy makes sense, because laboratory results show that a period of several hours is required for recovery and growth to regenerate sloughable border cells after rapid shedding induced by immersion (Hawes and Lin, 1990).
Several pioneering studies have drawn attention to the importance of considering spatial–temporal interactions in the rhizosphere. Van Bruggen and colleagues (2000, 2002) have shown that waves of microbial colonization and activity develop on growing roots. The peaks of bacterial activity are maintained at fixed distances from the root tip and are translocated relative to the soil surface during the growth of the root. The authors hypothesized that exudation from the root tip leads to microbial attraction and colonization near the tip. Death of the colony provides substrate for a later burst of bacterial growth in colonies that will be localized in more basal root regions, while the original colony, attached to a material group of root cells, is left behind the growing root tip.
A physically similar model was used in work from this laboratory to analyze pH patterns in the rhizospheres of growth zones (Kim et al., 1999; Nichol and Silk, 2001). We recognized that to understand the rhizosphere of the root tip, the time-dependent, classical models of charge balance across the root surface needed to be extended to include convection that resulted from root displacement during growth. When our convection–diffusion model was used to predict pH in the rhizosphere of the growth zone, we found that steady patterns developed around the moving tip. This pattern is in contrast to the time-dependent patterns that are observed in the stationary reference frame of the classical models. Recently, Watt and colleagues are extending this approach to investigate whether growth analysis can be a useful tool to understand the importance of seedling vigor in inhibiting pathological microbial infestation (reviewed in Watt et al., 2006). On a larger scale, a few modeling efforts have attempted to integrate C fluxes across the soil continuum. Roose et al. (2001) added root growth to the models of mineral uptake and efflux at particular developmental stages in individual roots. They treated as modules the transport at known stages and assumed the modules propagated through the soil in time. For an understanding of C sequestration, this approach could be extended with close attention to the complex spatial patterns of transport within the growth zone, coupled with extensive spatial–temporal observations of the propagation of the growth zones relative to the soil structure.
This review of some recent literature suggests the utility of studying C released from the root tip on a fine spatial scale (millimeters from the root tip and in the rhizosphere), with attention to the displacement of the root system on a larger scale (meters in the soil). For ecological relevance, the systems to be studied should include adult plants in real soils. An exciting research avenue is the construction of microbial sensors to find pulses of bacterial growth to monitor in situ the spatial and temporal patterns of C release from root systems (Herron et al., 2007). Preliminary results with the luxCDABEG reporter genes in common soil bacteria show in situ C release patterns qualitatively similar to those reported here. These experimental and theoretical approaches can be combined so that we can understand the complicated belowground C fluxes that interact to affect soil sequestration and release of carbon.
FOOTNOTES
1 This work was supported by a grant from the Kearney Foundation of Soil Science (Grant #2004.200) to W.K.S. The authors thank Prof. T. Rost for assistance in imaging of root border cells. 
2 Present address: Bureau of Land Management, 20 Hamilton Court, Hollister, California 95023 USA
3 Author for correspondence (e-mail: wksilk{at}ucdavis.edu), fax 1-530-752-1552
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