Transport of root-respired CO2 via the transpiration stream affects aboveground carbon assimilation and CO2 efflux in trees

Authors


Author for correspondence:

Jasper Bloemen

Tel: +32 9 264 6115

Email: jasper.bloemen@ugent.be

Summary

  • Upward transport of CO2 via the transpiration stream from belowground to aboveground tissues occurs in tree stems. Despite potentially important implications for our understanding of plant physiology, the fate of internally transported CO2 derived from autotrophic respiratory processes remains unclear.
  • We infused a 13CO2-labeled aqueous solution into the base of 7-yr-old field-grown eastern cottonwood (Populus deltoides) trees to investigate the effect of xylem-transported CO2 derived from the root system on aboveground carbon assimilation and CO2 efflux.
  • The 13C label was transported internally and detected throughout the tree. Up to 17% of the infused label was assimilated, while the remainder diffused to the atmosphere via stem and branch efflux. The largest amount of assimilated 13C was found in branch woody tissues, while only a small quantity was assimilated in the foliage. Petioles were more highly enriched in 13C than other leaf tissues.
  • Our results confirm a recycling pathway for respired CO2 and indicate that internal transport of CO2 from the root system may confound the interpretation of efflux-based estimates of woody tissue respiration and patterns of carbohydrate allocation.

Introduction

The exchange of carbon between terrestrial ecosystems and the atmosphere has been the subject of many studies (e.g. Baldocchi et al., 2005; Luyssaert et al., 2007, 2010). Carbon dioxide (CO2) is assimilated in ecosystems by photosynthesis, expressed as gross primary productivity, and is returned to the atmosphere by a variety of metabolic processes both aboveground and belowground, which comprise total ecosystem respiration (Trumbore, 2006). Belowground, the respiratory processes that contribute to soil CO2 efflux are functionally divided into autotrophic (CO2 released by roots and associated rhizosphere organisms) and heterotrophic (CO2 released during decomposition of nonliving organic matter) components (Hanson et al., 2000). Different techniques are applied to quantify soil CO2 efflux, but all conventional methodology is based on the assumption that root-respired CO2 diffuses through the soil and upward to the atmosphere (Kuzyakov, 2006).

A number of studies have investigated the relationship between tree canopy photosynthesis and belowground autotrophic respiration, which are connected through the transport of photosynthates from the leaves to the roots via the phloem (e.g. Hogberg et al., 2001; Subke et al., 2009; Kuzyakov & Gavrichkova, 2010). In contrast, few studies have focused on the reciprocal relationship: the coupling of autotrophic respiration and carbon assimilation resulting from the internal transport of dissolved CO2 from belowground. Ford et al. (2007) demonstrated in Pinus taeda seedlings that small amounts of soil-dissolved inorganic carbon can be taken up by roots, transported upward in the xylem stream, and assimilated by foliage. Aubrey & Teskey (2009) observed that a large amount of root-respired CO2 was transported upward in the transpiration stream to aboveground tissues of Populus deltoides trees. Therefore, the transport of root-respired CO2 in the transpiration stream has potentially important implications for measuring belowground respiration and assessing the role of transpiration in forest carbon cycling (Hanson & Gunderson, 2009).

The concentration of CO2 in the xylem (range < 1 to over 26%) is usually many times greater than that of the atmosphere (c. 0.04%) (e.g. MacDougal & Working, 1933; McGuire & Teskey, 2002; Teskey et al., 2008). This internal CO2 can be transported upward through the plant via the transpiration stream (Teskey & McGuire, 2002; McGuire & Teskey, 2004), where it is released radially into the atmosphere (Teskey & McGuire, 2005; Steppe et al., 2007) or assimilated within the plant. Small-scale experiments that introduced carbon isotope-labeled solutions into detached leaves or branches revealed that CO2 transported in the transpiration stream can be assimilated in leaf veins, petioles (Stringer & Kimmerer, 1993), and woody branch tissue (McGuire et al., 2009). Consequently, internally transported root-respired carbon in trees could provide substrate for carbon assimilation which has previously been overlooked. In addition, when large quantities of root-respired CO2 are transported internally and diffuse to the atmosphere remote from the site of production, efflux-based approaches fail to accurately estimate respiration of both roots and aboveground woody tissues.

Few studies have examined the fate of CO2 transported internally from belowground. Ford et al. (2007) and Ubierna et al. (2009) applied 13C-labeled solution to soil around seedlings in pots and around large field-grown trees, respectively, and they found that labeled carbon contributed only 0.8% to whole seedling carbon gain (Ford et al., 2007) and 1–3% to stem CO2 efflux (Ubierna et al., 2009). However, these studies did not address the transport and fate of CO2 derived internally from root respiration. Our objective was to label the xylem sap at the base of large trees in situ with 13CO2 to determine the fate of internally transported carbon. We hypothesized that, as the label moved upward in the transpiration stream, a portion would be assimilated by chlorophyll-containing woody and leaf tissues and a portion would diffuse to the atmosphere from stems and branches.

Materials and Methods

Overview

To investigate the role of internally transported root-respired CO2 as a potential carbon substrate for photosynthesis, we infused 13C-labeled solution into the xylem at the base of four intact eastern cottonwood (Populus deltoides Bartr. ex Marsh) trees and subsequently determined its presence in various tissues throughout the trees. The 13C label served as a proxy for dissolved CO2 entering the stem from the roots. The experimental trees were part of a 7-yr-old plantation in Whitehall Forest, a research facility of the University of Georgia near Athens, GA, USA. Details of the experimental plant material are given in Table 1.

Table 1. Diameter at breast height (DBH), tree height, total amount of 13CO2-labeled solution infused, amount of 13C uptake and cumulative sap flow during the infusion period for four Populus deltoides trees
TreeDBH (cm)Height (m)Label solution infused (l)13C infused (g)Cumulative sap flow (l)
  1. Trees 1 and 2 were infused with low-label solution; trees 3 and 4 were infused with high-label solution.

112.79.841.00.71152.6
211.97.240.00.70157.9
313.911.145.06.76189.5
49.78.645.06.76139.1

Baseline tissue sampling for isotopic analysis

Before infusion, on 23 June 2010, samples of woody tissue and leaves were taken from all experimental trees to determine the natural abundance carbon isotopic composition (δ13C) to which δ13C of labeled tissues would be compared. For this sampling, a mid-canopy branch of each tree was detached and a stem core was taken with an increment borer at approximately the same canopy position. Samples were immediately frozen in liquid nitrogen and then moved to a freezer at −9°C for storage before processing for carbon isotope analysis.

13C label infusion

13CO2-labeled solutions were prepared as described in McGuire et al. (2009) at low and high 13CO2 concentrations. To prepare each labeled solution, a 20-l polycarbonate container was completely filled with deionized water, which was amended with KCl to 40 mM concentration to facilitate solution uptake by the trees (Zwieniecki et al., 2001). Depending on the desired concentration, 1–3 l of the solution was displaced with 13CO2 gas from a cylinder of compressed 100% CO2 at 99 atom% 13C (ICON Services, Summit, NJ, USA). The gas was then circulated with a pump through the water in a closed loop for at least 3 h. The solutions were amended with sodium bicarbonate to adjust pH and achieve the target dissolved 13CO2 concentrations. Mean pH, gaseous CO2 concentration ([CO2], %), and dissolved inorganic carbon concentration ([13CO2*], mol l−1) were 5.30 ± 0.05, 3.77 ± 0.20%, and 0.0014 ± 0.0001 mol l−1 for the solutions at low 13CO2 concentration (low-label treatment) and 6.90 ± 0.02, 6.97 ± 0.05%, and 0.0120 ± 0.0005 mol l−1 for the solutions at high 13CO2 concentration (high-label treatment), respectively. The pH and [CO2] of the enriched solutions were within the range measured previously in Populus deltoides (Aubrey et al., 2011).

The trees were infused with the labeled solutions in the field. Preliminary experiments showed that infusing solution directly into a lateral root resulted in limited solution uptake, as it was only a small part of the root system, so infusion was performed at the base of the stem instead. A 19-mm-diameter hole was drilled 4.5 cm deep into the xylem on two sides of each tree c. 10 and 15 cm above ground level. A threaded brass hose-barb fitting was inserted 1.5 cm into each hole. The holes were drilled deeper than needed for installation of the fittings to increase the contact surface area of the solution with the sapwood. Both fittings were connected to the 20-l reservoir of labeled solution with 12.8-mm inner-diameter CO2-impermeable tubing (Bev-a-Line IV; Thermoplastic Processes, Georgetown, DE, USA). The solution reservoirs were placed 1.5 m above the holes to provide a small pressure head. Two trees were labeled simultaneously for 2 d starting at noon (trees 1 and 2 on 25–27 June 2010; trees 3 and 4 on 3–5 July 2010). Trees 1 and 2 were infused with the low-label solution and trees 3 and 4 were infused with the high-label solution. Based on previous measurements of sap flow, we estimated that it would take c. 2 d for the solution to reach the top of the canopy. Therefore, we waited two additional days after infusion ceased before harvesting the trees for tissue sampling to allow adequate time for assimilation of the 13C label by woody and leaf tissue.

Environmental conditions

Weather during infusion and measurement periods was hot and mostly sunny without precipitation. Mean maximum air temperature and minimum relative humidity were 30.7°C and 42% and 35.4°C and 45% during the June and July measurement periods, respectively.

Biomass determination and tissue sampling for isotopic analysis

Early on the third morning after the infusion ended, the trees were felled and tissues of the tree organs (stem, branch, and leaf) were sampled for carbon isotope analysis according to Fig. 1. Each tree was divided vertically into four strata: one below the canopy and three of equal length within the canopy (lower-, mid-, and upper-canopy). From each stratum, three 20-cm segments of the main stem were collected, one each from the bottom, middle, and top portions of the stratum (at 10%, 50%, and 90% of the total stratum length, respectively). Two branches from each canopy stratum were divided into three equal-length sections (A, B, and C; Fig. 1). A 20-cm segment of woody tissue and all leaf tissue were collected from each branch section. All samples were placed in plastic bags and immediately transferred to an ultralow freezer at −25°C to stop metabolic activity. Samples were later moved to a walk-in freezer and stored at −9°C. All remaining tissue not sampled for isotopic analysis was collected, separated according to Fig. 1, and dried for biomass determination.

Figure 1.

Scheme for biomass harvest and tissue sampling for isotopic analysis of four Populus deltoides trees infused with 13CO2-labeled solutions. Trees were subdivided into four strata: one below-canopy stratum and three canopy strata of equal length (lower-, mid-, and upper-canopy). Two branches from each canopy stratum were sampled. Branches were divided into three sections (A, B, and C) of equal length according to distance from the stem. The arrow indicates the location of infusion of the labeled solution.

Processing of tissue samples

Each tissue sample was thawed individually before subsamples were taken for carbon isotope analysis. Woody tissue was subsampled as follows: for stem sections where the diameter was > 5 cm, a radial xylem core was taken with an increment borer at the middle of each section (three per canopy stratum) for analysis. At the same position on the section, a 2-cm ring of outer (nonliving) bark was removed. The cortex (defined as live inner bark) of this entire ring was then sampled for analysis. For stem sections where the diameter was < 5 cm, a 2–10-cm-long (depending on diameter) subsample was taken from the middle of the section. Outer bark was removed and the entire subsample was separated into cortex and xylem for analysis. Branch samples (two branches per canopy stratum; three sections per branch) were subsampled in the same manner as the stems < 5 cm diameter. Leaves were sampled as follows: a subsample of 20 mature leaves was removed from each sample branch section and dissected into petiole, primary (central) vein, secondary veins, and remaining leaf mesophyll. A second subsample of 10 mature leaves was taken and processed as whole leaves. Subsamples of tissues harvested before labeling were processed in a similar manner to provide baseline carbon isotope values. All subsamples were dried to constant weight in an oven at 65°C and then ground to powder in a ball mill (8000-D Mixer Mill; SPEX SamplePrep, Metuchen, NJ, USA) for 13C analysis.

Isotopic analysis of tissue samples

A small portion of each ground tissue subsample was weighed to microgram precision in a tin capsule, flash combusted, and analyzed by isotope-ratio mass spectrometry at the Stable Isotope and Soil Biology Laboratory (SISBL), Odum School of Ecology, University of Georgia, Athens, GA, USA.

Enrichment of the labeled tissues (δ13Ct , ‰) was calculated as the difference between the δ13C value of the labeled sample (δ13Cs–t, ‰) and the δ13C value of the baseline sample (δ13Cb–t , ‰) of similar tissue:

display math(Eqn 1)

The ratio of 13C to 12C related to tissue enrichment (Rt) in each sample was calculated as:

display math(Eqn 2)

where 0.0112372 is the ratio of 13C : 12C of the PeeDee Belemnite standard.

Scaling isotope measurements of tissue component samples to organ and whole-tree levels

Biomass measurements of the subsamples were used to determine the mass proportions of the woody (cortex and xylem) and leaf (petiole, primary vein, secondary veins, and mesophyll) tissue components for each organ (stem, branch, and leaf) at every canopy stratum and branch section of each tree. These proportions were used to calculate the total biomass of the woody and leaf organs by tissue component for every canopy stratum and branch section combination (Fig. 1) for each tree. The amount of 13C assimilated in each tissue component (13Ct, mg) was calculated as:

display math(Eqn 3)

(DM, the dry mass of the tissue component per stratum and section (mg); C, the carbon content of the tissue component (%).)

To determine the amount of 13C assimilated by the three organs (stem, branch, and leaf) and the whole tree, the values of 13Ct for the individual tissue components of each stratum and section were summed to organ and tree level. The sum of the amount of 13C assimilated by the whole tree (13Cassim) and the amount of 13C lost to the atmosphere (13Cefflux) were assumed to be equal to the amount of 13C taken up by the tree (13Cuptake), based on the following mass balance equation:

display math(Eqn 4)

Thus, for each tree we estimated 13Cefflux by subtracting 13Cassim from 13Cuptake. 13Cuptake was calculated by multiplying the dissolved 13CO2 concentration of the labeled solution by the amount of solution taken up by the tree as described in McGuire et al. (2009).

Sap flow

Sap flow was determined by scaling sap flux density measured with homemade thermal dissipation probes (TDPs) (Granier, 1985) with sapwood area. Two TDP sensors were installed in the stem on opposite sides of each tree at a height of 0.45 m. Thermocouple depth was 10 mm and vertical separation of the sensor needles was 50 mm. Zero sap flow conditions were assumed between 03:00 and 05:00 h. Sap flow calibration parameters developed for Populus deltoides at this site (Sun et al., 2011) were applied to the sap flow measurements as recommended by Steppe et al. (2010). Sap flow was recorded with a datalogger (23X; Campbell Scientific, Logan, UT, USA) at 5-min intervals.

Gas sampling for isotopic analysis

Gas samples were collected from a cylindrical cuvette installed around a section of the stem and a section of a branch of each tree to confirm that 13C label diffused from woody tissues after uptake of the 13C enriched solution. These cuvettes were originally constructed to measure stem and branch CO2 efflux, but efflux data will not be presented in this report. Stem cuvettes were installed just below the base of the canopy at an average height of 1.05 m. Branch cuvettes were installed close to the stem in the lower third of the canopy at an average height of 1.70 m. Both stem and branch cuvettes were 15 cm long, constructed of 0.18-mm-thick Mylar film (Ridout Plastics, San Diego, CA, USA) and sealed to the stem with adhesive closed-cell foam gasket material and noncaustic silicone sealer (RTV162; MG Chemicals, Surrey, British Columbia, Canada). Compressed air at atmospheric composition was supplied from a cylinder to each cuvette at 500 ml min−1 with a mass flow controller (FMA5514; Omega Engineering Inc., Kingston, Ontario, Canada). For gas sampling, a needle attached to a 35-ml syringe was inserted through the foam gasket of the cuvette and a sample was withdrawn and injected directly into a 10-ml evacuated vial (Vacutainer; BD, Franklin Lakes, NJ, USA) that had been pre-filled with 1 ml of distilled deionized water. The septa of the vials were sealed with parafilm and samples were stored upside down to minimize gas diffusion through the septa seal. Starting 24 h after the beginning of label infusion, gas samples were collected every 2 h over a 24-h period and an additional sample was taken at 72 h. This timing was selected to detect changes in the enrichment of the air inside the cuvettes that might be related to the timing of label infusion and/or sap flow rate. Baseline samples of the air inside the cuvettes were taken before the start of infusion to determine natural abundance isotope concentrations.

Isotopic analysis of gas samples

Gas samples were analyzed by isotope-ratio mass spectrometry at SISBL. Similar to Eqn 1, the enrichment of the air inside the cuvettes attributable to 13C label diffusion from the stem and branch tissues (δ13Ca, ‰,) was calculated by subtracting the δ13C value of the baseline sample before label infusion (δ13Cb–a, ‰) from the δ13C value of the samples during label infusion (δ13Cs–a, ‰).

Data processing and statistical analysis

Enrichment of tissue components (δ13Ct) of each organ (stem, branch, and leaf) as well as the total quantity of 13C assimilated in each tissue component (13Ct) were analyzed using multifactorial analysis of variance (ANOVA). At the stem level, 13C label concentration (= 2), canopy stratum (= 4), and tissue component (xylem and cortex) were treated as fixed factors, and individual tree (= 4) was treated as a random factor. A similar ANOVA model was used at the branch and leaf levels with a few differences. At the branch level, the number of canopy strata was reduced by 1 (= 3, eliminating the below-canopy stratum), an additional tissue component (leaf) was included, and branch section (= 3) was included as a fixed factor. At the leaf level, tissue components consisted only of leaf parts (petiole, primary vein, secondary veins, and mesophyll). A similar ANOVA model was used to compare enrichment of different organs (stem, branch, and leaf). In this case, we confined our measurements to those taken within the canopy and calculated the weighted average of enrichment across tissue components as a function of biomass per canopy stratum. Treatment means were compared using Fisher's least significant difference (LSD) test. Enrichment of the air inside the stem and branch cuvette (δ13Ca) was analyzed using a repeated measures analysis of variance (ANOVA) with 13C label concentration (= 2), tissue (= 2), and time (= 14) treated as fixed factors and individual tree (= 4) treated as the random subject factor. Corrected Akaike information criterion (AICC) was used to determine the covariance structure that best estimated the correlation among individual trees over time. All analyses were performed using the mixed model procedure (PROC MIXED) of sas (Version 9.1.3; SAS Inc., Cary, NC, USA) with α = 0.05.

Results

13C label uptake

Each of the four trees took up between 40 and 45 l of 13CO2-labeled solution during the 2-d infusion period, which accounted for 23.8–32.4% of total sap flow. From visual observation it was clear that most of the solution was taken up during periods of high sap flow during the day, while little was taken up at night. Soil moisture was high during the experiment, precluding downward flow into the roots at night. Although solution uptake for all trees was similar, the average amount of dissolved 13C label taken up was nearly 10-fold greater under the high-label treatment (6.76 ± 0.00 g) compared with the low-label treatment (0.71 ± 0.01 g) (Table 1).

Carbon isotope composition of woody tissue and leaves

The baseline δ13C of tissues sampled before labeling ranged from −27.74‰ to −29.71‰. Tissue enrichment results (δ13Ct) showed that the label was transported from the base of the stem and assimilated in all tissues of the tree organs. The δ13Ct of the organs (calculated by weighted average of the tissue components) was influenced by label concentration, canopy stratum, and organ; however, these factors were not independent of each other (i.e. label concentration × organ interaction, P < 0.0001 and organ × canopy stratum interaction, P < 0.0001). The δ13Ct of the stem and leaves was higher under the high-label treatment than under the low-label treatment at all canopy strata. In the branches, a significant difference in δ13Ct between label treatments was observed only at mid- and upper-canopy strata. Under the high-label treatment, δ13Ct of the branches was higher than that of the stem and leaves at all canopy strata (Fig. 2). Mean δ13Ct of branches was higher at mid-canopy than at upper- and lower-canopy strata, whereas stem and leaf δ13Ct was similar among canopy strata under the high-label treatment (Fig. 2).

Figure 2.

Mean 13C enrichment (δ13Ct, ‰) of stem (black), branch (gray), and leaf (dark gray) organs of two Populus deltoides trees infused with high-label 13CO2 solution showing interaction between organ and canopy stratum. Tissue components of the organs were sampled according to the scheme depicted in Fig. 1. δ13Ct was calculated by subtracting δ13C of baseline (natural abundance) tissue samples from δ13C of labeled tissue samples and was averaged per organ and canopy stratum. Different lowercase letters indicate significant differences (Fisher's LSD; < 0.05) in δ13Ct among organs within a canopy stratum. Different uppercase letters indicate significant differences in δ13Ct of the same organ among different canopy strata. Bars indicate standard error of the mean.

In the stems, δ13Ct was influenced by label concentration, tissue component, and canopy stratum. For example, δ13Ct of the cortex was higher under the high-label treatment (3.86 ± 0.57‰) than under the low-label treatment (0.58 ± 0.21‰) at the upper- and mid-canopy strata, but cortex δ13Ct was not influenced by label treatment at the lower- or below-canopy strata. Under the high-label treatment, δ13Ct of the cortex was higher than that of the xylem at the upper- and mid-canopy strata, but not at the lower- or below-canopy strata (i.e. label concentration × tissue component × canopy stratum interaction, = 0.0108) (Fig. 3). The δ13Ct of both the cortex and xylem under the high-label treatment was higher at the upper- and mid-canopy strata than at the lower- and below-canopy strata. (Fig. 3).

Figure 3.

Mean 13C enrichment (δ13Ct, ‰) of cortex (black) and xylem (gray) of the stem at four canopy strata of two Populus deltoides trees infused with high-label 13CO2 solution showing interaction between tissue component and canopy stratum. Tissue components of the stem were sampled according to the scheme depicted in Fig. 1. δ13Ct was calculated by subtracting δ13C of baseline (natural abundance) tissue samples from δ13C of labeled tissue samples and was averaged for the cortex and xylem per canopy stratum. Different lowercase letters indicate significant differences (Fisher's LSD;< 0.05) in δ13Ct between cortex and xylem within a canopy stratum. Different uppercase letters indicate significant differences in δ13Ct of the same tissue component among different canopy strata. Bars indicate standard error of the mean.

Overall, in the branches, δ13Ct was highest in the mid-canopy stratum, intermediate in the upper-canopy stratum and lowest in the lower-canopy stratum (P = 0.0013), and was also influenced by label concentration and tissue component; However, the effects of label concentration and tissue component were not independent of each other (i.e. label concentration × tissue component interaction, < 0.0001). The δ13Ct of the cortex was higher under the high-label treatment than under the low-label treatment. It also differed among the branch tissue components (xylem, cortex, and leaves) under the high-label treatment but not under the low-label treatment. Mean δ13Ct across all canopy strata was highest in the cortex (10.81 ± 1.13‰), intermediate in the xylem (5.27 ± 0.55‰), and lowest in the leaves (1.04 ± 0.12‰) under the high-label treatment.

Mean δ13Ct of the leaves was higher under the high-label treatment than under the low-label treatment at the lower-canopy stratum (1.57 ± 0.39‰ vs −0.67 ± 0.12‰) and at the mid-canopy stratum (2.16 ± 0.34‰ vs 0.53 ± 0.09‰) (i.e. label concentration × canopy stratum interaction, = 0.0007). Under the high-label treatment, petioles were significantly more enriched than the other leaf tissue components regardless of canopy stratum or branch section. However, under the high-label treatment, δ13Ct of the petioles decreased with increasing distance from the stem (i.e. label concentration × tissue component × branch section interaction, = 0.0008) (Fig. 4).

Figure 4.

Mean 13C enrichment (δ13Ct, ‰) of petiole (black) and other tissue components (combined primary vein, secondary veins, and mesophyll) (gray) of the leaves of three equal-length branch sections in the canopy of two Populus deltoides trees infused with high-label 13CO2 solution showing the interaction between tissue component and branch section. Tissue components of the leaves were sampled according to the scheme depicted in Fig. 1. δ13Ct was calculated by subtracting δ13C of baseline (natural abundance) tissue samples from δ13C of labeled tissue samples and was averaged for the petioles and other leaf tissue components per branch section. Different lowercase letters indicate significant differences in δ13Ct (Fisher's LSD; < 0.05) between petioles and other leaf tissue components within a branch section. Different uppercase letters indicate significant differences in δ13Ct of either petioles or other leaf tissue components among different branch sections. Bars indicate standard error of the mean.

Amount of 13C assimilated

Generally, the total amount of 13C assimilated (13Ct) was higher under the high-label treatment than under the low-label treatment (Table 2), but it also depended on canopy stratum, and organ; however, these individual effects were not independent of each other (i.e. label concentration × organ interaction, = 0.0009 and organ × canopy stratum interaction, = 0.0002). The highest 13Ct was observed in the branches compared with stem and leaves under both label treatments (Table 2). Mean 13Ct of the branches was higher at the lower-canopy stratum than at the mid- and upper-canopy strata (< 0.0001) under the high-label treatment.

Table 2. Mean (SE) total biomass of tissue components (kg), amount of 13C fixed per tissue (13Ct, mg), and amount of 13C assimilated relative to 13C uptake (13Ct : 13Cuptake, %) of Populus deltoides trees infused with low-label (two trees) and high-label (two trees) 13CO2 solutions
Low-label treatmentStem xylemStem cortexBranch xylemBranch cortexLeaf petioleLeaf primary veinLeaf secondary veinLeaf mesophyllTotal
  1. All data have been scaled to the whole-tree level. Standard errors displayed as (0.00) were not zero, but have been truncated as a result of rounding.

Total biomass (kg)15.37 (1.73)2.56 (0.34)8.54 (3.37)2.75 (1.19)0.76 (0.16)0.25 (0.05)0.18 (0.01)5.87 (1.29)36.28 (2.10)
13Ct (mg)18.00 (0.01)4.20 (0.00)62.00 (0.00)21.00 (0.00)8.40 (0.00)0.24 (0.00)0.35 (0.00)10.00 (0.00)124.19 (0.01)
13Ct : 13Cuptake (%)2.52 (0.19)0.59 (0.04)8.69 (0.01)2.94 (0.06)1.18 (0.01)0.03 (0.01)0.05 (0.01)1.40 (0.06)17.40 (0.10)
High-label treatment
Total biomass (kg)15.34 (6.42)2.56 (0.85)7.82 (2.88)2.48 (0.86)0.70 (0.27)0.27 (0.11)0.16 (0.07)5.83 (2.05)35.16 (3.72)
13Ct (mg)64.00 (0.02)24.00 (0.00)140.00 (0.01)120.00 (0.02)15.00 (0.00)1.20 (0.04)0.61 (0.00)14.00 (0.00)378.81 (0.02)
13Ct : 13Cuptake (%)0.94 (0.04)0.36 (0.01)2.10 (0.01)1.72 (0.03)0.22 (0.01)0.01 (0.00)0.05 (0.00)0.20 (0.01)5.60 (0.02)

Within the stem, 13Ct was solely dependent on tissue component (P = 0.014). Averaged across canopy strata and label treatments, 13Ct of the xylem was higher than 13Ct of the cortex (Table 2).

Within the branches, 13Ct values of cortex and xylem were higher under the high-label treatment than the low-label treatment, but only in the lower- and mid-canopy strata. Averaged across canopy strata and branch sections, 13Ct was higher in the xylem than in the cortex under both label treatments (Table 2). Under the high-label treatment, 13Ct of the xylem decreased with increasing distance from the stem. However, these individual effects were not independent of each other, that is, 13Ct of xylem and cortex varied with label concentration, tissue component, canopy stratum, and branch section (canopy stratum × label concentration interaction, < 0.0001, canopy stratum × tissue component interaction, = 0.0004, and branch section × tissue component interaction, = 0.0003).

In the leaves, 13Ct was affected by label concentration, and varied among tissues and branch sections. Averaged across canopy strata and branch sections, 13Ct of the petioles was higher under the high-label treatment than under the low-label treatment (tissue component × label concentration interaction, P = 0.0274) (Table 2). Under the high-label treatment, 13Ct of the petioles was higher than that of the other leaf tissue components at the more distal branch sections B and C, but not at the proximal branch section A, resulting in a branch section × tissue component interaction (P = 0.0002).

Carbon isotope composition of air inside stem and branch cuvette

Isotopic signatures of the air inside the stem and branch cuvette confirmed that the 13C label was transported via the transpiration stream and diffused to the atmosphere from aboveground woody tissue. Enrichment of the air inside the cuvettes (δ13Ca) was influenced by label concentration and organ (stem and branch) and changed temporally; however, these individual effects were not independent of each other (i.e. organ × time × label concentration interaction, = 0.0003). Significant temporal variation in δ13Ca was observed in the stem, but not in the branch under both low- (Fig. 5a) and high-label (Fig. 5b) treatments. The highest δ13Ca of the stem was observed at 36 and 24 h from the start of label infusion under low- and high-label treatments, respectively. At 72 h from the start of label infusion, δ13Ca of the branch under both label treatments and δ13Ca of the stem under the low-label treatment returned to baseline, whereas δ13Ca of the stem under the high-label treatment remained high relative to the baseline. δ13Ca of the stem, averaged across the observation period (45.40 ± 6.75‰) was significantly higher than that of the branch (3.75 ± 1.92‰) under the low-label treatment. Similarly, δ13Ca of the stem, averaged across the observation period (41.96 ± 3.75‰), was significantly higher than that of the branch (3.75 ± 1.92‰) under the high-label treatment.

Figure 5.

Enrichment of the air in cuvettes (δ13Ca, ‰) installed on the stem (black circles) and a branch (gray circles) of Populus deltoides trees infused with low-label (a; two trees) and high-label (b; two trees) 13CO2 solutions. Gas samples were taken hourly from 24 to 48 h and at 72 h after the start of label infusion. *Significant (< 0.05) differences in δ13Ca between the stem and branch at each observation. Bars indicate standard error of the mean.

Assimilation and efflux of 13C relative to the amount of label uptake

On average, 17.40 ± 0.10% and 5.60 ± 0.02% of the infused 13C label was assimilated in aboveground tree organs under low- and high-label treatments, respectively (Fig. 6). 13C assimilation occurred mainly in the branches and, to a lesser extent, in the stem and leaves (Table 2, Fig. 6). Most of the 13C label was not assimilated and was therefore assumed to have diffused to the atmosphere both via the stem and the branches (82.60 ± 0.10% and 94.40 ± 0.02% under low- and high-label treatments, respectively).

Figure 6.

Overview of the fate of a 13C label infused at the base of Populus deltoides trees calculated by mass balance. Values are mean assimilation or efflux of the label (percentage of total label infused in brackets) for two trees infused with low-label solution (left panel) and two trees infused with high-label solution (right panel). Total efflux was calculated by subtracting total label assimilated from total label infused.

Discussion

Our results provide the first experimental evidence that internally transported root-respired CO2 can be assimilated in stems, branches, and leaves of large trees in the field. We found that up to 17% of a 13C label infused at the base of the stem was assimilated in woody and leaf tissues, providing evidence of an internal recycling mechanism for respired CO2. Moreover, based on mass balance calculations, we estimated that most of the putative root-respired CO2 diffused to the atmosphere from higher in the stem and branches, suggesting that efflux-based estimates of aboveground and belowground autotrophic respiration are inaccurate. Based on the previous observation that a substantial quantity of CO2 was transported from roots into shoots in xylem sap in this species (Aubrey & Teskey, 2009), the objective of this experiment was to determine the fate of root-respired CO2 that dissolved in xylem sap. Our infusion of the 13C label at the base of the stem was designed to serve as a proxy for dissolved CO2 that was transported in xylem sap from the root system into the stem. However, the study also provides insights into the potential fate of internally transported dissolved CO2 that originates from respiring cells in stems and branches.

Recycling of respired CO2 by woody tissues has been assumed to positively contribute to the overall carbon gain of plants (Aschan & Pfanz, 2003; Teskey et al., 2008). Assimilation of CO2 in woody tissues has been previously reported in the cortex of branches and stems (Aschan et al., 2001; Aschan & Pfanz, 2003; Saveyn et al., 2010) where sufficient light for CO2 assimilation is transmitted to chlorophyll-containing cells (Aschan & Pfanz, 2003). Our results showed that the cortex of the stem and branches was the most enriched tissue, confirming the potential importance of the cortex in assimilation of internally transported CO2. In branches receiving the high-label treatment, we found that the average δ13Ct of the cortex was twice as high as the xylem, which agrees with results of 13C labeling experiments on detached sycamore (Platanus occidentalis) branches (McGuire et al., 2009). Notwithstanding the fact that the upper- and mid-canopy strata were more distal from the 13C infusion, the cortex of woody tissues in these strata was more enriched than in the lower- and below-canopy strata. In the upper canopy, where young tissues are more metabolically active, corticular photosynthesis can be greater than in lower, older canopy sections (Cernusak & Marshall, 2000; Aschan et al., 2001; Pfanz et al., 2002). In addition to reduced metabolism in lower parts of the canopy, light transmission is substantially lower, which decreases corticular CO2 assimilation (Aschan & Pfanz, 2003).

The importance of woody tissue CO2 assimilation to overall carbon gain can only be evaluated when data are scaled to the whole tree. The xylem of stems and branches makes up a large proportion of total tree biomass in comparison to the cortex (in our study, on average, 65.9% vs 14.5%, respectively). Scaling δ13Ct with biomass demonstrated that, despite lower enrichment of xylem compared with cortex, the greatest quantity of 13C label was assimilated in the xylem, rather than in the cortex, in both branches and stems. The stems and branches of Populus deltoides in this study had visibly green xylem. The occurrence of chlorophyll in xylem has been reported for woody species (Rentzou & Psaras, 2008; Dima et al., 2006; Saveyn et al., 2010) and chlorophyll-containing pith cells in young poplar were found to be capable of fixing 14CO2 in the light (Van Cleve et al., 1993). Therefore, from a mechanistic perspective, assimilation of internally transported CO2 in the xylem, which is in essence a carbon recycling process, should be considered when assessing the total quantity of carbon assimilated by trees.

Few studies have described the role of assimilation of xylem-transported CO2 by leaves in the context of a recycling mechanism. Aubrey & Teskey (2009) estimated that up to 50% of root-respired CO2 could be transported internally to aboveground tree organs, and based on these results, Hanson & Gunderson (2009) argued that, if a large fraction of root-respired CO2 reached the foliage via the transpiration stream, it could have a substantial impact on the availability of CO2 for leaf photosynthesis. Powers & Marshall (2011) labeled the xylem of a Thuja occidentalis tree in situ and subsequently could not detect any label in the leaves. However, their failure to detect label in the leaves could be partly the result of the small quantity of label solution that was infused and partly a consequence of their strategy of sampling just a small amount of foliage from the top of the canopy. In our study, a portion of the infused label reached the leaves and was assimilated, with the petioles being the most enriched. Previous isotopic labeling experiments on detached branches of Platanus occidentalis (McGuire et al., 2009) and detached leaves of Populus deltoides (Stringer & Kimmerer, 1993) reported similar high values of carbon isotope enrichment in petioles compared with more distal leaf components. Apparently, the petioles scrubbed most of the label from the xylem stream before it reached the leaf mesophyll. Russin & Evert (1984) reported that, in the petioles of Populus deltoides leaves, chlorophyll-containing cells are in close contact with the vasculature, suggesting that xylem-transported CO2 may be a carbon source for photosynthetic reactions in adjacent tissues, as was reported by Stringer & Kimmerer (1993) and observed in other species (Hibberd & Quick, 2002; Berveiller et al., 2007; Leegood, 2008). However, in our study, leaf assimilation of transported 13C label was limited. It appeared that most of the infused label either diffused to the atmosphere or was assimilated by woody tissues before it could reach the foliage.

In total, we found that up to 17% of the infused label was assimilated in woody tissue and leaves. However, our results probably underestimate the actual importance of assimilation of internally transported CO2, because we could not account for any assimilation of internally sourced 12C, that is, the 13C label represented only part of the internal carbon available for assimilation. The water taken up by roots and transported upward contained dissolved CO2 (composed almost entirely of 12C) from root respiration. Based on the proportion of total sap flow that the label solution represented (23.8–32.4%), it can be expected that the amount of root-respired CO2 assimilated was three to five times more than estimated with the label. Finally, CO2 from aboveground autotrophic respiration, which is composed almost entirely of 12CO2, could be re-assimilated immediately in nearby photosynthetic cells or become part of the internal transport pool. McGuire & Teskey (2004) found that up to 55% of daily stem-respired CO2 could be transported in the xylem stream, so it is likely that this source contributed substantially to the amount of internal CO2 available as substrate for assimilation. Moreover, because the xylem was not saturated with 13CO2 label after the 2 d infusion period, photosynthetic discrimination against 13C in woody tissue (Cernusak et al., 2001; Saveyn et al., 2010) is another likely source of underestimation. Longer-term labeling would probably have resulted in even greater 13C concentrations in all tissues where the label was assimilated.

Based on our mass balance calculations, we found that the largest fraction of the 13C label (83% and 94% under the low- and high-label treatments, respectively) diffused to the atmosphere from the stem and branches. This observation suggests that, as root-respired CO2 is transported from belowground upward through the stem, most of it diffuses to the atmosphere before reaching the canopy. We assumed that the quantity of 13C label that remained in the xylem in dissolved and gaseous states at the end of the experiment was negligible and that diffusion of the label from the leaves was inconsequential because the direction of CO2 movement is from the atmosphere into leaves during the day and the amount of CO2 efflux from leaves at night is only a small fraction of the amount taken up during the day.

As CO2 dissolves in water, pH-dependent equilibria are established between the aqueous (CO2(aq)), bicarbonate (inline image) and carbonate (inline image) forms of CO2. In the xylem, with pH levels reported in the range of 4.7–7.4 (Teskey et al., 2008), CO2 is present only as CO2(aq) and inline image and not in its carbonate form (inline image). Measurements of the pH of the xylem sap in six trees at our site showed the average pH value to be 6.22 ± 0.07.

Similarly, the 13C solutions applied in this study, in which the average pH was 5.30 and 6.90 for the low- and high-labeled solutions, respectively, contained 13C only in its aqueous (13CO2 (aq)) and bicarbonate (inline image) forms. Other studies performed with carbon tracers have specifically supplied the bicarbonate or carbonate form of the stable or nonstable isotope to leaves (Stringer & Kimmerer, 1993), herbaceous plants (Hibberd & Quick, 2002), cuttings (Vapaavuori & Pellvonen, 1985) and small trees (Powers & Marshall, 2011) and they all concluded that there was an accumulation of the tracer in various tissues. Moreover, CO2 and inline image (12C or 13C) can be assimilated by Rubisco and phosphoenolpyruvate carboxylase (PEPc)-mediated carboxylation, respectively, and previous studies have reported that Rubisco (e.g. Pfanz et al., 2002; Saveyn et al., 2010) and PEPc (Hibberd & Quick, 2002; Berveiller et al., 2007) in woody and leaf tissues actively contribute to CO2 refixation. Thus, it is reasonable to believe that the inorganic form of the supplied 13C label was assimilated by biological processes.

Previous experiments demonstrated that CO2 dissolved in soil water contributed only a small amount to the overall carbon economy of plants (Ford et al., 2007; Jones et al., 2009). Moore et al. (2008) found that the isotopic composition of soil CO2 explained a considerable amount of the variation in the isotopic composition of CO2 that fluxed from stems, but Ubierna et al. (2009) observed that neither belowground processes nor CO2 transported in the transpiration stream had a detectable influence on stem CO2 efflux. By applying an isotope label to the soil, Ford et al. (2007) and Ubierna et al. (2009) tested the assumption that CO2 dissolved in soil water could be taken up by roots and transported in the xylem stream, while our study simulated the transport of CO2 originating from both root and soil sources by infusing a 13C label at the base of tree stems. The internal [CO2] measured at the bottom of the stem (Teskey & McGuire, 2007; Aubrey & Teskey, 2009) was substantially higher than the [CO2] in soil surrounding the roots, indicating that only a small quantity of the CO2 in the xylem at the base of the stem could have originated from the soil. Therefore, most of the internal CO2 at the base of the stem must be derived from the root system. Ford et al. (2007) and Ubierna et al. (2009) were not able to simulate the transport of CO2 derived internally from root respiration, which has been shown to comprise 92% of the CO2 transported in xylem from belowground (Aubrey & Teskey, 2009). In addition, Ford et al. (2007) and Ubierna et al. (2009) were unable to determine the total amount of label that was taken up by the roots, but by infusing the label solution directly into the xylem, we were able to quantify the uptake of 13C, which allowed us to calculate 13C efflux as the difference between uptake and assimilation.

The magnitude of root-respired CO2 transported in the xylem and subsequently lost to the atmosphere may have important implications for how we assess aboveground and belowground metabolism (Aubrey & Teskey, 2009; Grossiord et al., 2012). Given that one-half of root-respired CO2 may follow an internal flux pathway that cannot be accounted for by soil CO2 efflux measurements (Aubrey & Teskey, 2009) and that up to 94% of the 13C label infused into trees in this study was lost to the atmosphere, we suggest that up to 47% of root-respired CO2 could diffuse to the atmosphere from aboveground woody tissues. That a substantial amount of CO2 diffusing out of the stem and branches actually originates from belowground autotrophic respiration has important implications for how we interpret efflux-based measurements of soil and woody-tissue respiration and patterns of carbohydrate allocation. First, we suggest that the gas-exchange approach for estimating aboveground and belowground respiration is inaccurate and needs to be adjusted for internal transport of CO2. Secondly, our understanding of patterns of allocation of carbohydrates in trees should be reconsidered because we are routinely under- and over-estimating the carbon needed to sustain belowground and aboveground tissues, respectively, when estimates are derived from measurements of CO2 efflux. In future research, additional measurements of metabolic processes (photosynthesis, root, and stem respiration) conducted simultaneously with measurements of internal CO2 transport may reveal the implications of this recently recognized carbon flux pathway.

Acknowledgements

We thank M. Ameye, I. Bauweraerts, C. Bryars, T. M. Wertin and J. Yin for assistance with field work and A. Alred, S. Arnold, J. Audley, M. Cent, L. Coleman, A. Goijman, D. Layfield, E. Oxford and P. Tupper for sample processing. T. Maddox of Stable Isotope and Soil Biology Laboratory, University of Georgia provided advice and conducted isotope analysis. Funding was provided by the Special Research Fund (B.O.F.) of Ghent University and U.S. National Science Foundation Award No. 1021150.

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