Experimental setup

This experiment included seven C₃ herbaceous species covering three different functional groups, based on a priori plant biological characteristics related to C and nitrogen allocation strategies: Arrhenatherum elatius L., Dactylis glomerata L., Lolium perenne L. (forage grasses), Hordeum vulgare L., Triticum aestivum L. (crop grasses bred for grain production, expected to favour aboveground productivity and allocation to seeds), Medicago sativa L., Trifolium pratense L. (legumes, less prone to nitrogen limitation, but likely allocating significant amounts of C to their microbial partner in the symbiosis). Square pots (18x18x17 cm height) were filled with sieved (1 cm mesh) clay loam soil (30% clay, 41.8% silt, 28.1% sand; pH 6.8, organic matter content 28.5%). One week after seed germination on a thin layer of soil, the plantlets were transferred to the pots, following an even pattern (16 individuals in monoculture per pot, i.e., 658 plants m^-2) on three quarters of the pot area. Despite the high plant density, we did not find any indication that root growth was impaired at any stage of the experiment. A PVC collar (7 cm diameter, 5 cm high) for soil CO₂ efflux measurements was inserted 2.5 cm deep in the remaining quarter of the pot surface.

Plants were grown under similar and optimal conditions in six growth chambers (PGV36, Conviron, Winnipeg, Canada) set to a 14 hour light period (photosynthetically active radiation of ca. 400 μmol m^-2 s^-1) at 20°C and an 8 hour dark period at 15°C, with two gradual 1h transitions between light and dark periods. To avoid position effects, pots were randomly distributed within each chamber, and their position within the chambers was changed weekly. Additionally, enough space was left between the pots to avoid shading from taller species over smaller ones. The spacing was increased at each stage and facilitated by the removal of one third of the pots at each stage for measurements and sampling (see below). The chambers were not airtight: outside air was continuously pumped in from above the roof of the building to the room with the growth chambers–which acted as a large buffer from which the growth chamber pumped their air from–, resulting in chamber CO₂ concentration that was monitored to be at approximately 400 μmol mol^-1 (±10%), close to equilibrium with atmospheric CO₂ concentration. Air humidity was maintained between 60 and 70%.

Three vegetative ontogenetic stages were defined for each species prior to the experiment: 1) young foliage, where plants had autotrophic leaves and were just large enough (10 cm high) to be used for our measurements (1 to 4 weeks after germination, depending on the species; termed “young”), 2) maximum growth rate, based on weekly measurements of plant height increase (5 to 8 weeks; termed “mature”), and 3) beginning of senescence (8 to 10 weeks; termed “old”), dominated by maintenance respiration. All measurements were performed on non-senescent stems and leaves. These three growth stages were expected to be associated with different contributions of growth and maintenance respiration to total respiration [8], ranging from mostly growth respiration at the young stage due to the limited maintenance requirements of the small existing biomass, to both growth and maintenance respiration contributing to total respiration at the mature stage, and finally to mostly maintenance respiration at the old stage, during which biomass is no longer produced. Due to growth conditions of constant photoperiod and temperature, flowering and seed production were not initiated during this experiment.

Leaf-gas exchange measurements

The following ecophysiological variables were measured on fully expanded leaves of three plant replicates (i.e., growing in three different climate chambers) per ontogenetic stage and species, between the 6^th and the 8^th hour of the light period: transpiration rate (E), stomatal conductance of leaves to H₂O in the light (g_s) as well as net CO₂ assimilation rate (A). Additionally, leaf respiration rate (R_l) was measured at the end of the dark period (between 5^th and 7^th dark hour) to avoid light-enhanced dark-respiration [20]. Five measurements were averaged per replicate. Measurements were carried out under standardised conditions with a portable photosynthesis system (Li-6400, Li-Cor Inc., Lincoln, NE, USA). A dewpoint generator (Li-610, Li-Cor Inc.) and a CO₂ source ensured constant relative humidity (60%) and CO₂ concentration (400 μmol mol^-1) in the incoming flow of the Li-6400 leaf chamber. Temperature and light conditions were set to growth chamber conditions: 20°C, 400 μmol m^-2 s^-1 (light source: 6400-02B, Li-Cor Inc.). Leaf gas exchange was measured using a leaf chamber covering 6 cm² of leaf area. When the leaf was too small to cover the whole chamber, leaf area was determined after leaf gas exchange, using a portable area meter (Li-3000C, Li-Cor Inc.), and the gas exchange values were recomputed for the correct leaf area. Total leaf area (cut 1 cm above the ground) was measured (portable area meter LI-3000C) before drying (48h at 60°C) and weighing.

The soil CO₂ efflux rate was measured on three replicates (i.e., growing in three different climate chambers) per ontogenetic stage and plant species. A custom-made PVC chamber (27 cm high, 14.5 cm diameter) was fitted on the collar installed in the pots, and connected to a CO₂/H₂O gas analyser (Li-840, Li-Cor Inc.). The soil CO₂ efflux rate was calculated over 1 minute of linear CO₂ concentration increase in the chamber.

Sample collection

Leaves, roots, soil, and phloem sap organic matter were collected (n = 6 replicates per ontogenetic stage and species combination, i.e. one per growth chamber) between the 3^rd and 5^th hour of the dark period. Note that these measurements preceded R_l measurements by 2 hours, during which some C export to phloem may have occurred. However, the potential offset between measured respiration rate and isotopic signature of phloem sap organic matter should be small under our experimental conditions [7]. The individuals sampled for organic matter were different from those used for leaf gas exchange measurements to avoid any confounding effects of the δ¹³C value of the CO₂ source used during leaf gas exchange measurements. Aboveground biomass was cut 1 cm above the root crown. A first soil core (1.5 cm diameter core over the whole pot depth) was taken for bulk soil δ¹³C measurements, after manually removing roots. A second soil core (5 cm diameter core over the whole pot depth) was taken for microbial biomass δ¹³C measurements (see below). Roots were separated from the soil remaining in the pot by wet sieving. Leaf, root and soil samples for isotope composition analysis were dried (48h at 60°C) and finely ground (see below). Gravimetric soil water content was calculated after drying approximately 10 g soil at 105°C. Bulk phloem sap organic matter was collected by using an exudation method [21] adapted for herbaceous plants [5]. Briefly, for each species and ontogenetic stage replicate, one to three stems (or pseudostems) were cut at 1 cm height, rinsed with ultrapure water and carefully dabbed. Then, stems were inserted in a tube filled with 2 mL of 0.15M polyphosphate buffer at pH 7.5, sealed with parafilm® and placed in the dark (100% humidity, 4°C). After five hours, 1.5 mL of solution were collected, lyophilized, and used for C isotope composition analysis (see below).

For collection of soil CO₂, a soil chamber (Li-6400-09, Li-Cor Inc.) was adapted to fit the collars installed in all pots. For collection of leaf respired CO₂, a custom-made PVC chamber (15x7x4 cm) was used. The chamber was connected to the closed path infrared gas analyser of the Li-6400 portable photosynthesis to monitor chamber [CO₂]. Both chambers were equipped with a septum to allow sampling CO₂ respired by soil and leaves for isotope analysis.

C isotopic signatures of CO₂ respired by soil and leaves (individual leaves, not total aboveground biomass) were calculated using a Keeling plot approach. Briefly, this is a two end-member mixing model between the CO₂ emitted by a source and the background atmospheric CO₂, which allows to determine the isotopic signature of the source [22]. If more than two components mix, it is however possible to integrate several of them as one, provided they are well mixed and their relative contribution to the overall flux remains constant over the sampling period [23]. For example, belowground autotrophic respiration and heterotrophic respiration can be considered as an overall soil respiration component when using soil chambers (see below) The mixing model was based on five samples of chamber air, collected at regular intervals over a [CO₂] increase of at least 100 μmol mol^-1, and injected in vials (Exetainer®, Labco Ltd, High Wycombe, UK) that had been previously evacuated (<4.10³ Pa) and filled with N₂. Sample volume and vial volume for each measured compartment were chosen to represent less than 1% of the chamber volume and be within a practical [CO₂] range for isotope composition measurements (soil: 5 mL sample, 2.7 mL vial; leaf: 3 mL sample, 2 mL vial). All samples were collected between the 3^rd and 5^th hour of the dark period, thus effectively avoiding light-enhanced dark-respiration [20]. Vials were stored in a CO₂-free environment and measured for δ¹³C within 48h.

To allow comparison between samples taken at different times, all δ¹³C measurements were corrected for changes in background δ¹³C, weighed by the duration of each stage. Background δ¹³C was measured in a companion study [5] and the values are provided in S1 Table (δ¹³C_air).

Soil microbial biomass was extracted by fumigation-extraction [24] after soil sieving (2 mm mesh [24, 25]). For each soil sample, one 10 g subsample was fumigated 24 h with chloroform vapour before extracting microbial biomass, while a second 10 g subsample was extracted without prior fumigation. Microbial C was extracted by vigorous shaking for 30 minutes in 30 mM K₂SO₄ solution, then the extracts were filtered, frozen (-18°C) and lyophilised before isotope composition and C concentration analyses.

Isotope ratio mass spectrometry measurements

The measurement of δ¹³C values and C concentrations of plant biomass, soil and phloem sap organic matter was performed using a Flash EA 1112 Series elemental analyzer (Thermo Italy, former CE Instruments, Rhodano, Italy) coupled to a Finnigan MAT Delta^plusXP isotope ratio mass spectrometer (Finnigan MAT, Bremen, Germany) via a 6-port valve [26] and a ConFlo III [27]. The positioning of samples, blanks and laboratory standards in a measurement series followed the principle described by Werner and Brand [28]. Post-run off-line calculations like blank-, offset- and possibly drift-corrections for assigning the final δ¹³C values on the V-PDB scale were performed according to Werner and Brand [28]. Calibration of laboratory standards (acetanilide, caffeine, tyrosine) was periodically performed by comparing them to the corresponding international reference materials (NBS 22, USGS24) provided by the IAEA (Vienna, Austria). The long-term precision (~1.5 years) of our quality control standard (caffeine) was 0.09 ‰ or better for δ¹³C.

The δ¹³C values of CO₂ derived from gaseous samples were measured with a modified Gasbench II periphery (Finnigan MAT, Bremen, D) coupled to an isotope ratio mass spectrometer (Delta^plusXP; Finnigan MAT). The modification of the Gasbench comprises the replacement of the GC-type split by a ConFloIII-like split and the addition of a home-built cold trap (filled with Ni-wire) instead of the loop of the 8-port valve inside the Gasbench (modification as described by Zeeman et al. [29]).

Post-run off-line calculation like offset- and drift correction for assigning the final δ¹³C values on the V-PDB scale were done as described above. The δ¹³C values of the laboratory CO₂-in-air standards were determined at the Max-Plack-Institut für Biogeochemie (Jena, Germany) as described by Werner et al. [28]. The measurement of the aliquots of the laboratory standards is routinely better than 0.15 ‰.

C isotopic composition is expressed as the relative difference of the isotope abundance ratio of a sample relative to that of the VPDB international standard. This difference is expressed in per mil and defined as: (1) δ¹³C value of atmospheric CO₂ over each ontogenetic stage was calculated using phytometer leaves (see [5]).

For clarity, we use the following notation depending on whether the product is 1) the bulk δ¹³C value of a given plant part (δ¹³C_p: δ¹³C_leaf, δ¹³C_root, δ¹³C_phloem) or of soil microbial biomass (δ¹³C_mic) or 2) the δ¹³C value of respired CO₂ by either leaf or soil CO₂ efflux (δ¹³C_CO2: δ¹³C_CO2-leaf, δ¹³C_CO2-soil).

δ¹³C_mic was calculated using the following mass balance equation: (2) where δ¹³C_F and δ¹³C_NF are the δ¹³C values measured in fumigated and non-fumigated subsamples, respectively, and C_F and C_NF are C concentrations in fumigated and non-fumigated subsamples, respectively.

Carbon isotope discrimination

C isotope discrimination during a biochemical reaction (i.e., from a source to a product) can be calculated according to Farquhar et al. [30]: (3) where Δ is the discrimination between δ_source and δ_product, the δ¹³C values of the source and the product of the reaction, respectively. Unlike δ values, discrimination is independent of the isotopic composition of the source.

Respiratory fractionation (Δ_{Rsubstrate-product}) is estimated as discrimination between the δ¹³C value of CO₂ respired (either leaf or soil) and its putative substrate: (4) where δ_s is the δ¹³C value of the putative substrate (i.e. phloem sap organic matter in the main part of the study) and δ_CO2-X is the δ¹³C value of the CO₂ respired by X. We use the explicit term Δ_{Rsubstrate-product}, based on “Δ_R” used in previous studies [31].

Since phloem is the main source of C for non-autotrophic tissues and its isotope signature a good proxy for total canopy C discrimination [32], we chose to present and discuss respiratory C isotope fractionation between leaf-respired CO₂ and phloem sap organic matter (Δ_Rphloem-leaf). Nevertheless, respiratory fractionation was also calculated based on other C pools (i.e., leaf biomass) that may represent proxies for the actual leaf respiration substrate. Since CO₂ efflux from the soil has multiple sources and quantifying their relative contributions was beyond our objectives, we chose to calculate belowground respiratory C isotope fractionation between soil CO₂ efflux and bulk phloem sap organic matter (Δ_Rphloem-soil). However, as we did for leaf respiration, we also calculated soil respiratory fractionation based on other C pools that may represent proxies to soil respiration (i.e., leaf biomass and root biomass).

C isotope discrimination during photosynthesis is predicted from non-isotopic gas exchange measurements based on the widely accepted simplified model developed by Farquhar et al. [30], that assumes infinite internal conductance and neglects the effect of (photo)respiration, and further referred to as predicted photosynthetic C isotope discrimination (Δ_i): (5) where a is the fractionation occurring during CO₂ diffusion in air through the stomatal pore (a = 4.4‰, [33]), b is the net fractionation caused by carboxylation, c_a and c_i are ambient and substomatal partial pressures of CO₂, respectively. For higher C₃ plants, b mostly results from the fixation of CO₂ by RuBisCO, the carboxylation enzyme, estimated at 29‰ in spinach [34]. Additional fractionation effects resulting from PEP-carboxylase fixation, CO₂ transfer through the leaf boundary layer, dissolution of CO_2, liquid-phase diffusion (in the mesophyll cells), photorespiration and dark respiration during photosynthesis are gathered under the term d [35], which–when neglected–typically leads to the use of b = 27‰ ([16] see also [36]), as commonly used in ecological studies [37, 38] and also used in the present study. Note that d was discussed in detail in our companion study [5].

Statistical analysis

Data were analysed using R 3.0.2 [39]. The effects of ontogeny and species identity were tested by analysis of variance, nesting species identity within the functional group, and including growth chambers as blocks to remove any chamber-related effects from the analyses. Differences of means among groups were tested with a Tukey honest significant difference test (HSD). Linear regression models were used to fit regression coefficients.

Leaf gas-exchange

Species, functional group and ontogeny had significant effects on all leaf gas exchange variables (S2 Table). Furthermore, the effect of ontogeny differed among species (significant ontogeny × species interaction): in particular, leaf respiration (R_l) varied by an order of magnitude among different plant species and ontogenetic stages growing under controlled conditions (Table 1). Leaf respiration rates were the highest in mature Medicago foliage (1.68 μmol m^-2 s^-1), and the lowest in old Hordeum leaves (0.16 μmol m^-2 s^-1). A and g_s data were previously published in Salmon et al. [5]. Assimilation was highest in mature Triticum leaves (14.45 μmol m^-2 s^-1) and lowest in old Hordeum leaves (5.63 μmol m^-2 s^-1). Stomatal conductance values ranged from 43.8 mmol H₂O m^-2 s^-1 (old Hordeum) to 708.1 mmol H₂O m^-2 s^-1 (young Triticum). Resulting Δ_i ranged from 14.8‰ to 26.3‰, with an overall average of 23.4±0.6‰ [5]. Aboveground biomass and leaf area at the whole pot level both increased with age for all species (p<0.001), except for Trifolium that was more advanced in senescence and had already lost leaves between mature and old stages (Table 1). Soil CO₂ efflux rate (R_s) also varied among species and ontogenetic stages (0.6 to 44.3 μmol m^-2 s^-1, p<0.001). Arrhenatherum, Dactylis and Trifolium exhibited high soil CO₂ efflux rates at mature and old stages, like Medicago and Lolium at the old stage.

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Table 1. Leaf respiration rate (R_l), net assimilation rate (A), leaf stomatal conductance to H₂O (g_s), instantaneous predicted photosynthetic discrimination (Δ_i), aboveground biomass at the whole pot level (B), leaf area at the whole pot level (LA) and soil respiration rate (R_s) for different plant species at three ontogenetic stages (young foliage, maximum growth rate and beginning of senescence, indicated by young, mature and old, respectively).

A, g_s and Δ_i (recalculated with b = 27‰) values are from Salmon et al. (2011). Values are mean±1SE (n = 3).

https://doi.org/10.1371/journal.pone.0151583.t001

Isotopic signature of plant tissues and leaf-respired CO₂

δ¹³C_leaf, δ¹³C_phloem and δ¹³C_root values were previously published [5]. The isotopic signature of leaf biomass (δ¹³C_leaf) was on average -31.1±0.3‰ (mean±SE, S1 Table), significantly lower than phloem sap organic matter (δ¹³C_phloem, -30.3±0.3‰, 0.001<p≤0.01) and root biomass (δ¹³C_root, -29.4±0.4‰, p<0.001). Ontogeny and species significantly affected δ¹³C_leaf, δ¹³C_phloem and δ¹³C_root (p<0.001 in all three cases). The effect of ontogeny differed among species (significant ontogeny × species interaction) for all δ¹³C_p (p<0.01), except for δ¹³C_phloem.

Across all ontogenetic stages and species, δ¹³C_CO2-leaf ranged from about -30‰ to -22‰, showing larger variation than δ¹³C_CO2-soil (Table 2, S1 Table). ¹³C of leaf-respired CO₂ was enriched compared to leaf biomass and phloem sap organic matter (Fig 1). However, as mentioned above, we only calculated Δ_R based on phloem sap organic matter (see below). Over all ontogenetic stages and species, δ¹³C_CO2-leaf (-26.0±0.3‰ overall mean±SE) was significantly lower (p<0.001) than δ¹³C_CO2-soil (-24.4±0.3‰ overall mean). δ¹³C_CO2-leaf was significantly affected by ontogenetic stage and species, but not by functional group (Table 3, S1 Fig). Furthermore, over all ontogenetic stages and species, δ¹³C_CO2-leaf was positively related with leaf biomass (R² = 0.23, p = 0.028, Fig 2A) and negatively related with Δ_i (R² = 0.25, p = 0.021). However, the latter relationship was driven only by mature stage plants (R² = 0.83, p = 0.0045, Fig 2B).

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Fig 1.

δ¹³C of leaf-respired CO₂ compared to δ¹³C of aboveground C pools, i.e., leaf biomass (a) and phloem sap organic matter (b), and δ¹³C of soil-respired CO₂ compared to above and belowground C pools, i.e., leaf biomass (c), phloem sap organic matter (d), root biomass (e), bulk soil (f) and microbial biomass (g) at three ontogenetic stages (young foliage, “young”, white circles; maximum growth rate, “mature”, grey squares; beginning of senescence, “old”, black diamonds). Each point represents the average value (n = 6) for a given plant species at a given ontogenetic stage. Errors bars indicate ±1SE, the dashed line represents y = x.

https://doi.org/10.1371/journal.pone.0151583.g001

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Fig 2.

Response of δ¹³C values of leaf-respired CO₂ (δ¹³C_CO2-leaf) to leaf biomass (a) and to Δ_i (b) at three ontogenetic stages: young foliage (young, white circles), maximum growth rate (mature, grey squares) and beginning of senescence (old, black diamonds). Each point represents the average value (n = 6) for a given plant species at a given ontogenetic stage. Regressions are based on average values and given across all species and ontogenetic stages in (a) (y = 0.19x-26.82, R² = 0.23, p = 0.028) and for all ontogenetic stages (dashed line, y = -0.31x-19.26, R² = 0.25, p = 0.021) as well as mature stage only in (b) (y = -0.21x-21.31, R² = 0.83, p = 0.0045). Error bars indicate ±1SE.

https://doi.org/10.1371/journal.pone.0151583.g002

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Table 2. δ¹³C values of leaf-respired CO₂ (δ¹³C_CO2-leaf), soil-respired CO₂ (δ¹³C_CO2-soil), microbial biomass (δ¹³C_mic) and bulk soil δ¹³C (δ¹³C_soil) at three ontogenetic stages (young foliage, “young”; maximum growth rate, “mature”; beginning of senescence, “old”) in seven species (Arrhenatherum elatius, Dactylis glomerata, Hordeum vulgare, Lolium perenne, Medicago sativa, Trifolium pratense, Triticum aestivum).

Values are given as mean±1SE (n = 6). Within one column, different letters indicate significant differences between ontogenetic stages within a species (p≤0.05). Note that only significant differences are shown.

https://doi.org/10.1371/journal.pone.0151583.t002

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Table 3. ANOVA results for δ¹³C values, and respiratory C isotope fractionation between phloem sap organic matter and leaf- and soil-respired CO₂ (Δ_Rphloem-leaf and Δ_Rphloem-soil, respectively).

The variables are growth chamber, plant ontogenetic stage, functional group identity, and species identity (block, onto, group and species, respectively). F_x,y, x refers to the degrees of freedom of the tested variable and y to the d.f. of the residuals.

https://doi.org/10.1371/journal.pone.0151583.t003

Plant respiratory fractionation

Δ_Rphloem-leaf values were generally negative (Fig 3 and S2 Fig; except in young Medicago). Δ_Rphloem-leaf was significantly affected by ontogeny (Table 2, Table 4, overall mean±SE: -3.6±0.3‰), and differed significantly among functional groups (Table 3, S2 Fig). Δ_Rphloem-leaf tended to be positively related to assimilation rate (R² = 0.19, p = 0.051; Fig 3).

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Fig 3. Response of respiratory C isotope fractionation (Δ_R) between phloem sap organic matter and leaf-respired CO₂ (Δ_Rphloem-leaf) to leaf assimilation rate at three ontogenetic stages: young foliage (young, white circles), maximum growth rate (mature, grey squares) and beginning of senescence (old, black diamonds).

Each point represents the average value (n = 6) for a given plant species at a given ontogenetic stage. The regression is given across all species and ontogenetic stages and based on the average values (y = 0.28x-6.51, R² = 0.19, p = 0.051). Error bars indicate ±1SE.

https://doi.org/10.1371/journal.pone.0151583.g003

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Table 4. Respiratory C isotope fractionation (Δ_R) at three ontogenetic stages (young foliage, “young”; maximum growth rate, “mature”; beginning of senescence, “old”) in seven species (Arrhenatherum elatius; Dactylis glomerata; Hordeum vulgare; Lolium perenne; Medicago sativa; Trifolium pratense; Triticum aestivum).

Δ_R is calculated between phloem sap organic matter and leaf-respired CO₂ (Δ_Rphloem-leaf) as well as between phloem sap organic matter and soil-respired CO₂ (Δ_Rphloem-soil). Within one column, different letters indicate significant difference between ontogenetic stages within a species (p≤0.05). Note that only significant differences are shown.

https://doi.org/10.1371/journal.pone.0151583.t004

Calculating plant respiratory fractionation based on other potential substrates, i.e., leaf biomass and photoassimilates, showed patterns that were generally similar to that of Δ_Rphloem-leaf for the seven studied species, despite a few exceptions particularly with photoassimilates as a source for respiration (S3 Table).

Isotopic signature of microbial biomass and soil-respired CO₂

δ¹³C values of microbial biomass ranged from about -25‰ to -21‰ (-23.9±0.2‰ overall mean, Fig 3), and were overall significantly more enriched (p<0.001) than bulk soil δ¹³C (-25.5±0.0‰ overall mean). δ¹³C_mic was significantly affected by ontogeny. No significant correlations were observed between any measured ecophysiological parameter and δ¹³C_mic. Bulk soil δ¹³C was -25.48±0.01‰ (overall mean±se, Table 2) and was not affected by either ontogeny or the species growing in.

The ¹³C signal of soil CO₂ efflux was enriched compared to all plant components and generally also compared to bulk soil, but not compared to microbial biomass (Fig 1). The effect of ontogeny on δ¹³C_CO2-soil differed among functional groups (Table 3, S1 Fig), driven by significant differences among ontogenetic stages that were found only in the crops Triticum and Hordeum (Table 2). No significant correlations were observed between measured leaf-gas exchange variables and δ¹³C_CO2-soil.

Belowground respiratory fractionation

Δ_Rphloem-soil values (overall mean±SE: -6.5±0.4‰,) were negative (Table 4, S2 and S3 Figs), indicating enrichment in ¹³C of respired CO₂ compared to the putative respiration substrates. Δ_Rphloem-soil was significantly affected by ontogeny (Table 3, Table 4) and the interaction between functional group and ontogeny. Furthermore, Δ_Rphloem-soil tended to be negatively related to Δ_i (R² = 0.19, p = 0.051) and positively to WUE (R² = 0.16, p = 0.069). In contrast, Δ_Rphloem-soil was positively related to A/R_l for plants at the old stage (R² = 0.68, p = 0.021).

Considering other potential substrates for belowground respiratory fractionation (i.e., leaf and root biomass, photoassimilates,) showed similar patterns of the ontogeny response as for Δ_Rphloem-soil for the seven species studied, despite a few exceptions, particularly when choosing photoassimilates as a source for respiration (S3 Table).

Isotopic signature of plant-respired CO₂

We found large, significant effects of ontogeny and species identity on δ¹³C_CO2-leaf. The magnitude of change due to both ontogeny and species identity effects on the δ¹³C signal of respired CO₂ was up to 7‰, and the associated respiratory fractionation reached up to 7‰ in Δ_Rphloem-leaf. This is in agreement with the 6‰ increase in δ¹³C of leaf-respired CO₂ during the first 22 days of growth of French beans and peanuts, documented in the–to our knowledge–only other studies about the impact of ontogeny on δ¹³C_CO2 [3, 17, 18]. These values thus span a similar range as changes in the δ¹³C of leaf-respired CO₂ resulting from changes in environmental conditions (e.g., about 8‰ in [40]). Thus, our results emphasise the relevance of accounting for ontogeny and species identity when using respiratory δ¹³C values to infer plant responses to changing environments. Furthermore, δ¹³C_CO2-leaf was more enriched than its potential substrate in all species and stages considered, indicating that respiratory fractionation in leaves should be accounted for when using δ¹³C to study respiration in C₃ herbaceous species. The large ontogenetic and species effects that we found on δ¹³C_CO2-leaf were associated with significant changes in Δ_Rphloem-leaf driven by ontogeny and the identity of species or functional groups.

Short-term processes imprint the isotopic signature of photoassimilates that will be allocated to fast- or slow-turnover C pools. Long-term processes control the isotopic signature of slow-turnover C pools. Our results illustrate how both short- and long-term processes are involved in the changes in δ¹³C of respired CO₂ with species and ontogenetic stages. In the short-term, δ¹³C_CO2-leaf was significantly negatively related to Δ_i, especially at the mature stage, which is directly related to the isotopic signature of fast-turnover C pools [41–43]. The coupling between δ¹³C_CO2-leaf and Δ_i is in agreement with general predictions for annuals and fast-growing species with small C reserves [44], and suggests that a fraction of the newly-synthesized assimilates was likely directly respired by the leaf. Furthermore, Δ_i is mostly driven by A and g_s: it is therefore highly dependent on plant regulation of gas exchange. Under controlled conditions, in the absence of day-to-day environmental variation, changes in leaf gas exchange rates are not driven by changes in climatic conditions, but are responding to changes in the ratio of plant C source to sink strengths [45–47], either directly, as decreasing C source:sink ratios will stimulate leaf gas exchange to meet the increasing C demand, or indirectly, as increasing C sinks will translate to increasing leaf and root biomass. Increasing leaf biomass in turn results in higher photosynthesis and transpiration, while increasing root biomass allows better access to water resource, which in turn can translate in higher transpiration, A and g_s.

Therefore, the relationship between leaf gas exchange and Δ_i across species and ontogenetic stage suggests that species-related differences in δ¹³C_CO2-leaf could be related to shifts in the balance between C sources and sink strengths. Considering the possible fate of C once assimilated or released from storage, it can either be used to build biomass, be respired or exported. While information about the δ¹³C of exported C–particularly out of the plant (e.g. exudates)–are lacking, in the long-term, we found that δ¹³C_CO2-leaf became more enriched as the plants grew older. Even in the absence of the isotopic signature of exported C, this relationship between δ¹³C of C incorporated in biomass and respired is likely due to changes in the balance between maintenance and growth respiration as plant grew older. Indeed, young and growing plants synthetize large amount of proteins–in particular RuBisCO [48]–and cellulose, both of which generally slightly enriched in ¹³C compared to leaf organic matter [13], thus generating CO₂ that is more ¹³C-depleted than bulk leaf material. At later growth stages, RuBisCO degradation [48] could result in the release of more enriched ¹³C-CO₂.

Plant respiratory fractionation and C balance

Over all species and ontogenetic stages, we found a positive relationship between Δ_Rphloem-leaf and A. Under our controlled experimental conditions, and within the limits set by the available light intensity and photosynthetic capacity (e.g., as a function of available RuBisCO), changes in the balance between C source and sink strengths were very likely influencing photosynthetic activity [45–47]. Our results suggest that increased C demand from respiring and growing tissues stimulate higher assimilation rates to meet the additional C costs and is associated with less fractionation. In a conifer forest, Ubierna & Marshall [14] also found that C allocation played a key role, but the relationship between A and the difference between δ¹³C_leaf and δ¹³C_CO2-leaf was negative. This relationship was attributed to changes in respiratory fractionation resulting from changes in available C that could be diverted to secondary metabolism. Although the relationship between secondary metabolism and δ¹³C respired CO₂ has not been formally proven, this hypothesis is well supported by experimental data (see for example [4] for details). These opposite results might, however, be driven by the same underlying mechanism: higher C availability for secondary metabolism increases respiratory fractionation. Under field conditions, A is mostly regulated by environmental conditions and particularly light availability, while under constant day-to-day environmental conditions such as in our experiment, A will respond positively to an increase in C sink strength relative to C source. This can be both direct, i.e. increased carbohydrate demand by respiring and growing tissues stimulating C assimilation [45–47, 49], or indirect, when growing aboveground tissue leads to an increase in photosynthetically active tissue. Thus, under field conditions, in addition to covering respiratory C costs and growth, higher photosynthesis rates allow to have carbohydrates available for secondary metabolism, resulting in more apparent fractionation between δ¹³C of respired CO₂ and δ¹³C of respiratory substrate [14]. However, when photosynthetic rate is stimulated by higher C sink strength, higher C demand for growth and maintenance respiration will lead to both higher A and less fractionation between respired CO₂ and its substrate (smaller Δ_Rphloem-leaf in absolute value), since less C is available for secondary metabolism. Therefore, while our results only describe an ontogenetic trend among herbaceous C₃ species and do not allow formal testing of the underlying mechanisms, they are consistent with the hypothesis that respiratory fractionation in leaves is likely responding to changes in the ratio between the strength of C sinks and sources.

Isotopic signature of soil-respired CO₂

At the timescale of our study, we found little overall effects of species, functional group or ontogeny on δ¹³C_CO2-soil, in line with the expected low ontogenetic variability in δ¹³C of root-respired CO₂ (see recent review by [17]). However, the ontogenetic pattern of δ¹³C_CO2-soil differed among functional groups (significant ontogeny × functional group interaction), as a significant ontogeny effect was detected in crops (namely in Triticum and Hordeum) but not in the other groups. δ¹³C_mic showed a response pattern that was similar to that of δ¹³C_CO2-soil, suggesting that the crop-specific ontogenetic effects on the isotopic signature of soil CO₂ emissions were, at least in part, shaped by the metabolic activity of the soil microbial community. Changes in microbial community structure have been associated with changes in the isotopic signature of CO₂ respired by heterotrophs [50, 51]. Thus, our results suggest that plant-microbial interaction dynamics over time may differ between crops and the other functional groups included here.

The large soil organic matter pool most likely contributed as a substrate for microbial respiration and thus for soil CO₂ efflux in our experiment, especially for plants in the young stage. Based on the relative plant assimilation and soil respiration rates at different ontogenetic stages, we can at least determine stages at which soil organic C may have been contributing to soil CO₂ efflux. At the young stage, overall photosynthesis was likely not sufficient to sustain the total soil CO₂ efflux, except in crops (S4 Table). At the mature stage, soil CO₂ efflux could have been fuelled by assimilation in all species, except Dactylis. Finally, at the old stage, the overall assimilation flux was potentially sufficient to meet respiration costs, except for legumes. Thus, the discrimination associated with soil microbial respiratory processes may change with ontogeny, as a consequence of several processes, including shifts in the relative activity of different microbial populations [50, 51] or in the organic substrate preferentially metabolized [52], but our experimental setup does not allow to quantify these processes.

Additionally, the changes in δ¹³C_CO2-soil and associated changes in Δ_Rsoil-phloem with ontogeny and functional group were possibly driven by 1) changes in the relative contribution of plants to the overall soil CO₂ efflux, both through root respiration and microbial respiration fed by plant rhizodeposits, and 2) changes in the fractionation between respired CO₂ and respiratory substrate provided by the plants, i.e., phloem sap organic matter, either directly as a source for root respiration or indirectly as C exudates. Little information is available to discuss (1), especially in terms of root exudation. Nonetheless, the contribution of root-respired CO₂ (and its isotopic signature) to total soil CO₂ efflux and its signature is expected to increase with the development of the plant root system. The lack of a significant relationship in our study between Δ_Rsoil-phloem and aboveground biomass (the latter should be correlated with root biomass [53]) does not support an increased contribution of roots to total CO₂ efflux with growth. However, our results support (2) as a mechanism shaping part of the ontogeny and functional group effects on δ¹³C_CO2-soil. As expected, ontogeny strongly affected Δ_Rsoil-phloem, since it also affected its putative substrate, δ¹³C_phloem. Nevertheless, considering a different proxy for respiration substrate, such as root biomass, showed very similar results (S3 Table). Thus, the Δ_Rsoil-phloem response to ontogeny was not solely driven by change in δ¹³C_phloem, but reflected actual changes in respiratory processes. In contrast, we detected no overall ontogeny effects on δ¹³C_CO2-soil, suggesting that although ontogeny affects the ¹³C signature of plant material, these changes do not propagate to the isotopic signature of soil respiration.

Moreover, our study showed consistent relationships on different time scales between Δ_Rsoil-phloem and leaf gas exchange. Δ_Rsoil-phloem was negatively related to Δ_i (i.e., dependent on A and g_s) and positively related to the ratio of assimilation to leaf respiration (A/R_l) at the old stage. Note that similar relationships were found when considering another proxy than phloem as respiration substrate, indicating that these results were not an artefact due to the correlation between δ¹³C of phloem and leaf gas exchange. Both relations suggest that, in agreement with results for aboveground tissues, increased C assimilation (higher A, resulting in both higher A/R_l and lower Δ_i) leads to smaller respiratory fractionation in absolute values, most likely since most assimilated C is directed towards primary rather than secondary metabolism. This hypothesis is supported by several studies that documented the importance of aboveground controls of C allocation to root respiration ([54] and references therein). Additionally, C fixation by PEPc in belowground plant tissues might also play an important role in the isotopic signature of respiratory substrates in roots [17] and thus lead to an apparent fractionation between phloem sap organic matter and respired CO₂.

The ontogeny effect on Δ_Rsoil-phloem was specific to each functional group. In particular, crops showed a significant decrease in absolute values between younger and older stages, while little differences were found in forage grasses and legumes. Crops, unlike legumes and forage grasses, are selected for grain rather than vegetative biomass production. Despite grain filling did not take place in this study (see above), C allocation and nutrient uptake in crops may have been switching towards storage (for example as sucrose and fructans in the stem [55] and references therein) in preparation for grain filling, possibly leading to the measured changes in apparent respiratory C fractionation and consequently in δ¹³C_CO2-soil, efflux. Furthermore, crop is the only functional group for which total assimilation was sufficient to cover total C lost through respiration at every ontogenetic stage (S4 Table). This is consistent with a tighter coupling between respiration fractionation in soil and plant C dynamics in crops than in other functional groups.

Conclusion

Our results show that biological factors (i.e., ontogeny and species identity) can strongly affect the carbon isotopic signature of respired CO₂ and respiratory fractionation. These factors need to be accounted for in studies inferring physiological responses of plants to their biotic and abiotic environment based on δ¹³C of respired CO₂ and respiratory fractionation. Furthermore, we show that these ontogeny-related changes in δ¹³C of respired CO₂ and respiratory fractionation are associated with changes in physiological processes involved in the plant C budget and particularly with changes in photosynthesis, biomass and respiration. Therefore, our results suggest that ontogeny-related changes in the ratio of plant internal C source to sink strengths drove the measured changes in δ¹³C of respired CO₂ and respiratory fractionation.