Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths : 13 C labeling evidence

Introduction Conclusions References

nacea. The experiment was designed to prevent C leakage from the labeled nutritive solution to the chamber atmosphere. After 14 days of growth, the 13 C and 15 N enrichments ( 13 C-excess and 15 N-excess) in the roots, stems and leaves, and phytoliths, as well as the 13 C-excess in AAs extracted from roots and stems and leaves, were quantified relatively to a control experiment in which no labelled AAs were added. The net 15 uptake of 13 C derived from the labeled AAs supplied to the nutritive solution (AA-13 C) by Festuca arundinacea represented 4.5 % of the total AA-13 C supply. AA-13 C fixed in the plant represented only 0.13 % of total C. However, the experimental conditions may have underestimated the extent of the process under natural and field conditions. Previous studies showed that 15 N and 13 C can be absorbed by the roots in several 20 organic and inorganic forms. In the present experiment, the fact that phenylalanine and methionine, that were supplied in high amount to the nutritive solution, were more 13 Cenriched than other AAs in the roots and stems and leaves strongly suggested that part of AA-13 C was absorbed and translocated in its original AA form. The concentration of AA-13 C represented only 0.15 % of the phytolith C (phytC) but was still 1.5 times higher 25 in phytoliths than in leaves supporting that, ultimately, AA-13 C was preferentially fixed in organic compounds subject to occlusion in phytoliths. Plausible forms of AA-13 C and 19752

Introduction
In the rhizosphere, there are numerous known interactions between carbon (C) and nitrogen (N) processes that have yet to be accurately assessed in qualitative and quan-5 titative terms for their consideration in carbon cycle models (Heimann and Reichstein, 2008). Among those interactions the uptake of low molecular weight C and N (e.g. organic acids, sugars and amino acids (AAs)) by plant roots (both mycorrhizal and non-mycorrhizal plants) has been well documented through experiments using hydroponic solutions, artificial substrats or soils (e.g. Chapin et al., 1993;Bardgett et al., 10 2003; Kuzyakov and Jones, 2006;Persson et al., 2006;Jones et al., 2009a;Näsholm et al., 2009;Sauheitl et al., 2009;Gioseffi et al., 2012 ;Moran-Zuloaga et al., 2015). The aim of most of these studies was to investigate in which extent and under which conditions, organic N could be utilized by plants as a direct source of N (i.e., without going through a mineralization step). The answers are still debated (Jones and Darrah,15 1992; Jones et al., 2009a;Moran-Zuloaga et al., 2015) but it has been showed that the net uptake of AAs, that may reach half of the total N uptake of grasses (Sauheitl et al., 2009), occurred for a wide range of AA concentrations (Chapin et al., 1993;Jämtgård et al., 2008 ;Harrison et al., 2008;Sauheitl et al., 2009), and different levels of soil microorganism competition (e.g. Jones et al., 2005a;Sauheitl et al., 2009 Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | from old soil organic C, and possibly old AAs reached by the roots (Santos et al., 2012;Reyerson et al., 2015). Silicon (Si) is the second most abundant element of the earth surface after oxygen. Its uptake by plants is widespread and generates, at the ecosystem scale, important fluxes from the soil to plants (Conley, 2002). For instance, Si absorption represents 2 to 10 times the amount of dissolved Si exported to stream 5 water in tropical ecosystems (Alexandre et al., 2011). If part of the organic C uptake is linked to Si uptake in the rhizosphere, the involved flux may thus also be significant.
Here, we aim to investigate whether and in which extent organic C absorbed by grass roots, either under the form of intact AAs or microbial metabolites, can feed the organic C occluded in phytoliths. For this purpose we added 13 C-and 15 N-labeled 10 AAs to the Si-rich nutrient solution of the grass Festuca arundinacea. AAs are ubiquitous in soil organic matters of various residence times (Bol et al., 2009). The experiment was designed to prevent C leakage from the labeled nutritive solution to the chamber atmosphere. After two weeks of growth, the 13 C and 15 N enrichments in the roots, stems/leaves, and phytoliths of the grass (two replicates) were quantified, and 15 expressed in 13 C-excess and 15 N-excess relatively to a control experiment in which no labelled AAs were added. ering three 12 L tanks containing the nutrient solutions (24 plant pots per tank) (Fig. 1). This assembly enabled close contact between seeds or seedling roots and the nutrient solutions. In order to prevent escape of the C-and N-bearing gas from the nutrient solution to the chamber atmosphere, O-rings sealed the plant pots to the perforated platform and the perforated platform to the tank. Additionally, the seeds were covered 5 with agar-agar (polysaccharide agarose). Each tank was hermetically connected to two 20 L containers (an input container filled with the nutrient solutions and a waste container). Seeds were first germinated for 10 days in a germination solution and seedlings were then grown for 14 days in a growth solution. Both solutions were renewed continuously at a rate of 2 L/24 h using a peristaltic pump (Fig. 1). Additionally, the growth 10 solution was entirely renewed once, after 8 days of growth.
Overall, 10 L of germination solution and 64 L of growth solution were used per tank. Germination and the growth nutrient solution composition were described in detail in Guigues et al. (2014) during the day. 20 At the end of the experiment, all samples were cleaned (to remove agar-agar), rinsed in deionized water and oven-dried at 50 • C (to constant mass). When the tanks were filled (1st and 8th days of growth) and emptied (8th and 14th days of growth), the growth solution was sampled and kept frozen for future analyses.

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In the two first tanks (two replicates), a mixture containing four 13 C-and 15 N-labeled AAs was added to the growth solution containing only inorganic C and N (as described above). Those AAs were selected for their following characteristics: L-Alanine (l-ALA) 19755 Introduction

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Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | (C/N = 2.6) is ubiquist and occurs in high proportions in soils and plants proteins. The D-enantiomer of Alanine (D-ALA) which is present in natural soils (Hill et al., 2011), was expected to be more resistant to degradation and, if absorbed, less subject to metabolization. Consequently, we speculated that D-ALA may accumulate as a waste product first in cell vacuoles and later in phytoliths. L-Phenylalanine (L-PHE) 5 (C/N = 7.7) comprises a phenolic ring resistant to decomposition by microorganisms in soils, solutions or plants. L-Methionine (L-METH) (C/N = 4.3) is a sulfur amino acid expected to be recovered at low abundance in plants, but is easily identifiable in Gas Chromatography-Isotope Ratio Mass Spectrometry (GC-IRMS All AAs were uniformly labeled except (i) D-ALA, which was not 15 N-labeled but was 13 C-labeled on one atom (C-2) and, (ii) an equivalent fraction of L-ALA because the labeled D-ALA was provided as a racemic mixture (DL-ALA). In each tank, the follow- The third tank was only filled with the growth solution, without labeled AAs. It served as a control experiment to calculate the 13 C and 15 N enrichments of the plants from the labeled tanks, and verify that 13 C and 15 N derived from AAs (AA-13 C and AA-15 N), that may have contaminated the chamber CO 2 , were not photosynthesized by the plants.

Analyses
For each tank (one control tank and two labeled tanks) stems/leaves were separated from the roots into two samples. 200 µm. After alkaline fusion, they were analyzed in SiO 2 , using Inductively Coupled Plasma Optical Emission Spectrometry (ICP-OES). For each sample, total C, 13 C/ 12 C, total N and 15 N/ 14 N were determined after dry combustion by IRMS using a Carlo Erba NA 1500 elemental analyzer (EA) coupled to a Thermo-Finnigan Delta-plus mass-spectrometer. Solutions were also analyzed for 5 total C and 13 C/ 12 C by IRMS, after evaporation by dry combustion in tin capsules. Proteic carbon was analyzed as the sum of 19 individual AAs representing ca. 95 % of all AAs. Quantification and 13 C/ 12 C determination of individual AAs were performed using a GC-IRMS (Thermo Fisher Scientific). The extraction and purification procedure was a slightly modified version (Rubino et al., 2014) of the protocol devel- with helium as a carrier gas. AA identification and quantification were performed using commercial mixtures of 20 proteinogenic AAs (Sigma Aldrich). Norvaline (Sigma-Aldrich Co. Ltd.) was added to plant samples before hydrolysis as an internal standard for quantification. Due to the addition of non-labeled carbon by TBDMS, AA 13 C enrichment was subsequently calibrated for each AA (Shinebarger et al., 2002). Briefly, 25 this calibration was based on the independent measurement of 13 C of the TBDMSderivatives of the commercial AA and an additional set of four 13 C-labelled AA (Rubino et al., 2014). The calibration equation involves the number of carbon atoms added as TBDM and the isotopic composition of the latter. This isotopic composition term BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths disappears in the calculation of the isotope excess (cf Eq. 1 below). When multiple peaks were encountered for a single AA, the main or the best individualized peak was chosen for both quantification and isotope ratio determination. The isotope ratios were calculated using ammonium sulfate IAEA-N1 (δ 15 N = 0.43 ± 0.07 ‰) and IAEA-N2 (δ 15 N = 20.41±0.07 ‰), polyethylene IAEA CH7 (δ 13 C = −32.15±0.05 ‰) as sec-5 ondary standards. The sucrose standard IAEA CH6 used as a control yielded a mean value of 10.43 ‰. Phytoliths were extracted from plants using a high purity protocol based on acid digestion and alkali immersion steps previously described in detail by Corbineau et al. (2013). Phytolith samples were observed in natural light microscopy to determine 10 their morphological assemblage and check for the absence of residual organic matter particles. An additional purity check was done via Scanning Electron Microscopy (SEM) (Corbineau et al., 2013;Reyerson et al., 2015). Then, phytolith samples were analyzed for their C and N contents, as well as their 13 C and 15 N abundances by EA (Fisons NA 1500NC) coupled to a continuous flow IRMS (Finnigan Delta-Plus). About 15 6-10 mg of phytolith concentrates were weighed using a pre-calibrated microbalance (Sartorius AG, Göttingen, Germany) into tin capsules (5 × 9 mm capsules, Costech Analytical Technologies Inc., Valencia, CA, USA) and pre-baked at 100 • C for 2 h to remove extraneous contaminants. To assure accurate integration and linearization of the raw analytical data obtained from the lower C and N peaks, we decreased the helium 20 carrier flow rate and measured several aliquots of in-house collagen L-Cystine 99 % (δ 13 C = −28.74 ± 0.13 ‰ and δ 15 N = −6.14 ± 0.07 ‰; from Sigma Aldrich Co. Ltd) and Atropina (δ 13 C = −21.30 ± 0.06 ‰ and δ 15 N = −2.90 ± 0.10 ‰; from Costech 031042) as well as the internationally certified reference materials (e.g. Graphite USGS24 δ 13 C = −16.05 ± 0.07 ‰ and, Amonium sulfate − IAEA-N1 δ 15 N = +0.43 ± 0.07 ‰).

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Aliquots of baked-clean silicon dioxide (SiO 2 ; mesh# −325, Sigma Aldrich, St. Louis, MO, USA) and fossil phytoliths (MSG70; Crespin et al., 2008) were also analyzed to provide independent blank data (Santos et al., 2010) and to check accuracy. To serve as quality assurance, note that the reproducibility obtained on the phytolith laboratory BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | standard MSG70 was SD ± 0.2 ‰ for δ 13 C and δ 15 N, SD ± 0.01 % for C and N, and SD ± 0.09 for C/N.

Calculations
For the control tank (no labeling), results are reported as δ values in ‰ relative to the Vienna Pee Dee Belemnite (V-PDB) for δ 13 C, and in ‰ relative of the atmospheric N 2 where 13 C atom sample is the 13 C atom abundance of a sample (stems and leaves, roots 10 or phytoliths) from a labeled tank and 13 C atom control is the 13 C atom abundance of the same sample from the control tank. The concentration of AA-13 C in a sample, expressed in % of total C in the sample, was calculated using Eq. (2): where 13 C atom solution is the 13 C atom abundance in the labeled solution at the beguinning of the growth phase. Recovery of AA-13 C in a sample, expressed in µg g −1 of the dry matter weight (d.wt), was calculated using Eq. (3): BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths Net uptake of C derived from the supplied AAs (AA-13 C uptake), expressed in % of AA-13 C initially supplied to the solution, was calculated using Eq. (4): AA-13 C uptake (%) = Recovery AA-13 C (mg g −1 ) × d.wt sample (g) AA-13 C max, supplied (mg) where d.wt sample is the dry weight of sample and AA-13 C max, supplied is the maximum AA-13 C supplied to the solution (322 mg of AA-13 C).

5
The same calculations were applied to 15 N.

Total C, N and AAs concentrations in the plants
After 14 days in the growth solution, the above-ground part of the plants of Festuca arundinacea were 30 cm high but had not reached maturity or flower development 10 (Fig. 1). The C/N ratios of the stems and leaves were similar in the labeled and control plants (8.4 and 9.0), whereas the C/N ratios of roots were higher in labeled (11.0) than in control (4.2) plants (Table 1; Fig. 2). Recovered AAs accounted for 52 mg C g −1 of the dry matter in both roots and leaves. This is a high AAs content, also attested by the low C/N ratio, in agreement with the high N level requirement of young plants suf-15 ficiently fertilized to support rapid protein synthesis (Mattson, 1980). However AA-13 C and AA-15 N only accounted for ca. 13-14 % of total C, and 40 % of total N. This N contribution was lower than what might be expected. Indeed, at any stage of growth, AAs (in the form of protein or free molecules) should account for more than 50 % of grass N. This discrepancy can be attributed to an underestimation of AAs by the extraction- 20 purification process (i.e., incomplete hydrolysis recovery due to recombination into strong acids, and incomplete silylation). An underestimation of the AAs is consistent with the fact that amino-acid TBDMS-derivatives, such as tryptophane or cystine ( cystein-dimer), could not be recovered. However such an underestimation should not bias the measured relative proportions of METH, PHE, and ALA.

Excess, uptake and recovery of AA-13 C and AA-15 N in the plants
The δ values of the roots and aerial parts of the control plants were respectively −31.0 and −31.7 ‰ for δ 13 C and 13.8 and 14.5 ‰ for δ 15 N. These values were in the range 5 of the ones measured for C 3 grasses in natural conditions (δ 13 C from −22 to −34 ‰; e.g. O 'Leary, 1988), ensuring that CO 2 potentially produced by decomposition of the 13 C-labeled molecules inside the tanks did not contaminate the growing chamber atmosphere. The amount of labeled C recovered in plants and phytoliths was thus considered as exclusively resulting from root uptake.

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Relative to plants from the control tank, plants from the labeled tanks were enriched by 0.05 % in 13 C and 0.5 % in 15 N in the roots and by 0.02 % in 13 C and 0.3 % in 15 N in the stems and leaves (Table 1).
Overall, the net uptake of AA-13 C and AA-15 N by the plant represented respectively 4.5 and 46.9 % of the AA-13 C and AA-15 N added to the solution (Table 1; Fig. 2).

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Whereas 35 % of absorbed AA-13 C and 19.2 % of absorbed AA-15 N were stored in the roots, 64.4 and 81.1 % of the absorbed AA-13 C and AA-15 N, respectively, were allocated to the stems and leaves (after Table 1). The associated AA-13 C/AA-15 N ratios were 0.8 in the roots and 0.3 in the stems and leaves (  Fig. 3). This contribution was higher in roots (0.28 % of C and 4.1 % of N) than in stems and leaves (0.10 % of C and 2.6 % of N). In roots, AA-13 C was more concentrated in AAs than in total plant matter (0.70 vs. 0.28 % of C) (Table 1). When translocated to leaves, AA-13 C concentration in AAs decreased to reach that of the bulk leaf matter (0.10 % of C) (Table 1).

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Among the measured AAs in plant, ALA and PHE were more abundant by more than a factor 10 relative to MET (Table 1). However, ALA was not more enriched in 13 C than BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths most of the AAs (Fig. 4). Instead, PHE and MET were significantly more enriched in 13 C than other AAs in roots, stems and leaves (Fig. 4).

Concentrations of phytoliths, phytC, phytN, and AA-13 C and AA-15 N in phytC and phytN
Silica content measured by ICP-AES accounted for 0.08 and 0.26 % of the dry weight 5 (d.w.) of roots and stems/leaves respectively, which is lower than the > 1 % d.w. concentration previously measured for this species harvested 8 weeks after sowing (Hartley et al., 2015), possibly due to the fact that the plants did not reach maturity or/and that the volume of the roots in contact with the Si-enriched solution was small, limited by the RHYZOtest configuration. Most of the stems and leaves silica was in the form 10 of phytoliths (0.19 % d.w.) ( Table 2)  Interestingly, phytoliths were 1.5 times more enriched in 13 C ( 13 C-excess of 0.03 %) than the leaves ( 13 C-excess of 0.02 %), or leaves AAs ( 13 C-excess of 0.02 %) ( Table 1).
The AA-13 C concentration in phytoliths represented 0.15 % of phytC which is low but still 1.5 times higher than the concentrations in the bulk matter and AAs of stems and 20 leaves ( Table 1). The AA-13 C/AA-15 N ratio in phytoliths was low (0.8) but higher than in bulk stems and leaves (0.3) (Table 1; Fig. 3).
BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths

Plausible forms of AA-13 C and AA-15 N absorbed and translocated
Festuca arundinacea may have absorbed AA-13 C and AA-15 N from the labeled solution in several organic and inorganic forms described below. The AA-13 C/AA-15 N ratios of roots and leaves (0.8 and 0.3, respectively) that were significantly lower than C/N ratios  Jones and Darrah, 1993). At least, rhizospheric and endophytic microorganisms, that acquired their labeled signature from the labeled solution, may also account for the AA-13 C and AA' 15 N recovery in the plant. All these processes may have accounted for the 13 C and 15 N enrichments measured in the roots. 20 However, the fact that 13 C was more concentrated in the extracted AAs than in bulk root matter (Table 1; Fig. 3), and further, that 13 C-excess values of MET and PHE were significantly higher than 13 C-excess values of other AAs in the roots (Fig. 4), supported that some AA-13 C was absorbed by and fixed in the roots in their original AA form.
From roots to leaves, the concentration of AA-13 C (relative to total C) in AAs de-25 creased significantly (Fig. 3). This suggests that plants metabolized AA-13 C during the translocation to aerial parts. Deamination of labeled AAs, probably recombined into other AAs, sugars or other molecules constituting the plant matter may also be in-19763 BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths voked. At least some AA-13 C may have been lost via respiratory processes overtaking N losses, as suggested by the decrease of the AA-13 C/ 15 N ratio from roots to stems and leaves (Table 1). Those three processes probably occurred jointly. Thus, the present labeling experiment does not allow to precisely trace the molecular form under which the AA-13 C and AA-15 N were absorbed and fixed in roots, stems and 5 leaves, as recently done using a position-specific C and N labeling technique (Moran-Zuloaga et al., 2015). However the fact that PHE and METH that were supplied in high amount to the nutritive solution were significantly more 13 C-enriched than other AAs in the roots and stems and leaves (Fig. 4) strongly suggests that part of AA-13 C was absorbed and translocated in its original PHE and METH form. On the other hand, the low 13 C-enrichement in ALA suggests that this molecule was rapidly degraded or metabolized in the solution instead of being absorbed.

AA-13 C fixation in phytoliths
In agreement with the radiocarbon evidence for soil organic C occlusion in phytoliths (Reyerson et al., 2015), AA-13 C accounted for a measurable part of phytC. Due to 15 the very small concentration of phytC, isotopic analysis at the molecular level was impossible. The phytolith C/N value (5.0) was close to a value previously measured in cultivated wheat phytoliths (3.7; Alexandre et al., 2015) and in the range of C/N values characteristic of AAs (4-5, Jones et al., 2009). However, the AA-13 C/AA-15 N ratio (0.8) was far from this range. The form in which AA-13 C, and most generally phytC, has been 20 occluded in the silica structure thus remains unknown. Previous Nano Secondary Ion Mass Spectrometry (NanoSIMS) investigation of phytC indicated that at least part of phytC is continuously distributed in the silica structure, at the sub-micrometric scale (Alexandre et al., 2015). This has been further supported by Raman spectroscopy mapping (Gallagher et al., 2015). The process of silica 25 precipitation has been investigated by environmental scanning electron microscope (ESEM) and TEM-EDX analyses that highlighted that silica first precipitates in the in-BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths ner cell wall, probably triggered by the presence of callose or lignin, then infills the cell lumen in a centripetal way, until most of the cell becomes silicified (e.g. Perry et al., 1987;Motomura, 2004;Laue et al., 2007;Law and Exley, 2011;Zhang et al., 2013). During this process, an organic template probably participates in the silica formation (Harrison, 1996;Laue et al., 2007). When the cell silicification is complete, residual 5 organic compounds that were not already occluded probably gather in any remaining spaces within the cell and delimitate micrometric central cavities characteristic of most phytoliths (Alexandre et al., 2015). In the present case, the higher concentration of AA-13 C (relatively to total C) in phytoliths than in leaves, supports a preferential fixation of AA-13 C in these residual organic compounds subject to occlusion in the silica structure.

10
There are two plausible hypotheses for this preferential fixation. The first hypothesis is that AA-13 C may be associated with Si when absorbed by the roots, translocated in the plant and introduced into the cells. However, 29 Si NMR spectroscopy of 29 Si-labeled exudate of wheat xylem previously indicates only the occurrence of the dissolved forms of Si (Casey et al., 2004). Although this does not preclude the subsequent formation 15 of organo-silicate complexes it weakens the hypothesis of Si and C being associated since their uptake by the roots. Additionally, in our experiment, the roots that contain the lowest amount of Si also contain the highest amount of AA-13 C which is not in agreement with AA-13 C and Si being associated when absorbed by the roots. The second hypothesis is that AA-13 C may be isolated as an unwanted substance in cell vacuoles 20 and subsequently trapped in the silica structure. In order to check this hypothesis we used D-ALA, expected to be less metabolized than L-AAs, although recent investigations suggest that plants are able to utilize D-AAs at rate comparable to those of other N forms (Hill et al., 2011). D-ALA was not specifically taken-up or retained as an intact molecule (Fig. 4) and cannot account for the AA-13 C measured in phytC. D-ALA 25 may thus not be appropriate for tracing unwanted substances. Further investigations, including the use of spectroscopies relevant for characterizing phytC at the molecular level, are necessary to support or refute one or more of these other hypotheses.

Implication for our understanding of the C cycle at the plant-soil interface
In the experiments presented here, the net uptake of AA-13 C by Festuca arundinacea represented 4.5 % of AA-13 C supplied to the nutrient solution, part of it being absorbed as intact AA molecules (here methionine and phenylalanine). AA-13 C fixed in the plant represented only 0.13 % of total C, the root absorption of AA-13 C being clearly marginal 5 compared to photosynthesis. However, our experiment may underestimate the extent of the process under natural and field conditions. Indeed, AAs uptake was shown to be inhibited by the high concentrations of mineral N preferred by the plant (Paungfoo-Lonhienne et al., 2008;Sauheitl et al., 2009;Gioseffi et al., 2012). In the present case, mineral N may come from KNO 3 initially presents in the original nutrient solution and from the supplied AAs dissociated by microbes, mycorrhizas or root exsudates (Paungfoo-Lonhienne et al., 2008;Keiluweit et al., 2015). The maximum AA content in the growth solution (0.2 mmol L −1 ) was higher than AAs concentrations that have been measured in soil solutions (from 0 to 0.1 mmol L −1 ; Vinolas et al., 2001;Jämtgård et al., 2010). The probable high concentration of inorganic N in the solution, partly com-15 ing from the mineralization of AAs supplied in high concentration, may consequently have inhibited AAs uptake. Additionally, due the RHIZOtest configuration, roots were confined to a small volume, and their contact with the renewed labeled solutions and AA-13 C available for uptake was limited. Thus, the range of AA-13 C uptake and allocation in roots, stems, leaves and phytoliths of the grass grown in the present labeling 20 experiment may underestimate what may happen in field conditions. The comparison with soil conditions is not straightforward due to the diversity of C and N biogeochemical exchanges in soils. However, to gain an idea on the order of magnitude of the C flux that may occur from soil to plant, we used the 0.13 % AA-13 C concentration in plants obtained in the present experiment and extrapolated that value 25 to the grassland ecosystem scale. Grasslands cover a global surface of 2.4 × 10 9 ha (Scurlock and Hall, 1998) and are characterized by a Net Primary Production (NPP) ranging from 7 to 20 × 10 9 t C ha −1 yr −1 (Scurlock et al., 2002). The global grassland BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | productivity thus ranges from 16.8 to 48 10 9 t C yr −1 . The obtained flux of soil-derived C absorbed by grasses then ranges from 21.8 to 62.4 × 10 6 t C yr −1 . This is nonsignificant when compared to the 2.6 × 10 9 t C yr −1 estimate for the land C sink (IPCC Staff, 2007), or to the 0.4 × 10 9 t C yr −1 estimate for the global long term soil C accumulation rate (Schlesinger, 1990). It is however higher than a possible CO 2 phytolith bioseques-5 tration flux (e.g. Parr and Sullivan, 2005;Parr et al., 2010;Song et al., 2014). The CO 2 phytolith biosequestration concept is based on the assumptions that phytC is exclusively derived from atmospheric CO 2 , and has a long residence time in soils. A recent re-examination of the CO 2 biosequestration flux by phytoliths, in the light of the lower and more realistic estimates of phytolith residence time in soils, yielded a value of 10 4.1 × 10 4 t C yr −1 for the world grasslands (Reyerson et al., 2015). The present study further minimizes the significance of CO 2 biosequestration by phytoliths showing that it could be counteracted by the flux of C potentially mobilized from soils by grass root uptake.
Recent experiments have demonstrated that root exudates promote a net loss of soil 15 C previously assumed to be stable at the millennial scale thanks to its protection by mineral constituents (e.g. clays or amorphous minerals). Root exudates would stimulate microbial and fungi digestion (priming effect) (Fontaine et al., 2003(Fontaine et al., , 2011 and promote dissolution of the mineral phase through oxalic acid production (Keiluweit et al., 2015). From the present experiment, we suggest that direct uptake of organic C by 20 roots, in conjunction with the N uptake, should be accounted for when investigating the role of roots in soil C mobilization. Radiocarbon dating of phytC (the ultimate fixation of soil C absorbed by the roots) that produced ages of several thousand years (Reyerson et al., 2015) strongly suggests that old pools of soil organic C are being accessed by root uptake. Introduction

Conclusions
In agreement with previous studies, the present labeling experiment supports that AA-C absorbed by grass roots and allocated to stems and leaves can partly preserve its original molecular form (here methionine and phenylalanine). Moreover, the experiment shows for the first time that AA-C absorbed by grass roots and allocated to stems and 5 leaves can partly feed the C ultimately fixed in organic compounds subject to occlusion in stem and leaf silica. Further analyses are required to identify the form in which phytC has been occluded. This finding complements previous radiocarbon evidence of soil C (and more particularly old soil C) contribution to phytC (Santos et al., 2012;Reyerson et al., 2015) and raise questions about the mechanisms that drive soil C mobilization by BGD Jones, D. L. and Darrah, P. R.: Re-sorption of organic compounds by roots of Zea mays L., and its consequences in the rhizosphere, Plant Soil, 178, 153-160, 1996. Jones, D. L., Shannon, D., Junvee-Fortune, T., and Farrar, J. F Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | Persson, J., Gardeström, P., and Näsholm, T.: Uptake, metabolism and distribution of organic and inorganic nitrogen sources by Pinus sylvestris, J. Exp. Bot., 57, 2651-2659, 2006 Badeck, F., Cattivelli, L., and Santos, G. M.: Evidence of old soil carbon in grass biosilica particles, Biogeosciences Discuss., 12, 15369-15410, doi:10.5194/bgd-12-15369-2015Discuss., 12, 15369-15410, doi:10.5194/bgd-12-15369- , 2015 BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths   BGD 12,2015 Direct uptake of organic carbon by grass roots and allocation in leaves and phytoliths