N 775777775711

Soluble i 1 Solubilized i 1 ^ï™^1 i 1 Solubilized i_

extractives 1 ! acid solubles 1 'acid insolubles1

acid solubles . >

Microbial biomass

Passive (acid insolubles)

J"

Slow

The thickness of the arrow relates to the size ol the C flux

Model Flow and Organization

Years n Extractives □ Acid solubles □ Protected acid solubles * Acid insolubles o* microbial origin ■ Acid insolubles of lignin origin.

Figure 8.6 Proposed model for describing carbon isotope variation during litter decomposition and humus formation due solely to compound-specific decomposition effects ('microbes are what they eat, but they eat different things') and the hypothesis that DOC derives primarily from the by-products of 13C-depleted lignin ('microbes degrade but don't eat lignin'). In actuality, a combination of compound-specific decomposition, anaplerotic CO2 fixation, and the Suess effect may explain much of the enrichment in <513C values with depth in aerobic soils. For simplicity, all microbially active substrates (extractives and cellulose) are given the same ¿¡13C value (-25%o), which contrasts with the <513C value of the microbially undesirable lignin (-29%o). Initially (year = 0), litter has a carbon isotope ratio of -26.5%o (top inset graph) due to the abundance of the 13C-depleted acid insolubles (e.g., lignin) relative to the 13C-enriched compounds such as extractives (e.g., sugars, amino acids), acid solubles (cellulose), and protected acid solubles (lignin-bound cellulose). As decomposition progresses, microbes preferentially incorporate the 13C-enriched cellulose and extractives into their biomass. A fraction of this assimilated C is used to synthesize secondary compounds, which are ultimately converted to resistant acid insoluble substances after they are exuded or released upon death and degradation of the cells (the rest is recycled back to the extractive, acid soluble, and protected acid-soluble pools). These lignin-like acid-insoluble substances of microbial origin carry the isotopic signature of the enriched microbial biomass (e.g., -25%o), which reflects their enriched substrate (extractives and cellulose). As lignin is progressively degraded, solubilized, and lost as DOC, these resistant microbial compounds increasingly dominate the mass balance of remaining organic matter and therefore its overall isotopic composition. The DOC flux required to sustain the lignin losses (bottom inset graph) and, in this case, the eventual 1.5%o enrichment of the remaining organic matter (top inset graph) is within the range cited in the literature (l-84gCm-2 yr~'; Neff and Asner, 2001).

should be small (<20%). Moreover, field measurements indicate that soil surface respiration is typically enriched in 13C relative to plant litter inputs (Fig. 8.2), contrary to the depleted-respiration hypothesis.

If both SOM and respired CO2 are enriched in 13C in aerobic soils, a mechanism such as 13C-depleted DOC leaching is required to close the carbon isotope mass balance. Measurements of carbon isotope ratios in DOC (Schiff et al, 1990; Trumbore et al, 1992; Ludwig et al, 2000) suggest that DOC is indeed more depleted in 13C than both surface litter inputs and SOC with depth in soils where SOC is also enriched (Fig. 8.7). Further, radiocarbon dating (Trumbore et al., 1992; Schiff et al, 1997) suggests that the age of the DOC increases with depth consistent with its autochthonous production from in-situ microbial activity rather than by transfer from overlying layers. Thus, with increasing depth in the soil profile, each soil layer is progressively more isolated from fresh litter inputs at the surface (aside from C inputs from deep roots and some leaching). At more mature stages of decomposition deeper in the soil profile, the significance of 13C-enriched polysaccharide-derived microbial products that are effectively recycled by the microbial community increases relative to 13C-depleted lignin-derived products that are continuously degraded and lost as DOC (Fig. 8.6). Although 13C-C02 losses through respiration greatly exceed 12C-DOC losses, the longer effective retention time of microbial resynthe-sized 13C allows the cumulative effect of small DOC fluxes (see bottom inset graph in Fig. 8.6) to eventually cause significant shifts in soil isotope ratios (see top inset graph in Fig. 8.6). The cumulative nature of the DOC effect

Figure 8.7 Variation in carbon isotope ratios of dissolved organic carbon (DOC) and soil organic carbon (SOC) with depth in the soil profile. From Ludwig et al. (2000).

Figure 8.7 Variation in carbon isotope ratios of dissolved organic carbon (DOC) and soil organic carbon (SOC) with depth in the soil profile. From Ludwig et al. (2000).

is evidenced by the greater 13C-enrichment observed with depth in the soil profile where enough time has elapsed to degrade lignin and its derivatives.

Resolving the complexities of root-microbe-soil stable isotope interactions and their application to partitioning ecosystem respiration will undoubtedly benefit from studies aimed at closing the ecosystem carbon isotope budget. Such studies should include compound-specific 13C measurements of the major carbon fractions (e.g., soluble carbohydrates, amino acids, alkanes, cellulose, lipids, and lignin) and 14C analyses to better understand the fate and transformations of stable carbon isotopes as they move from leaves to roots to litter and among various soil microbial, DOC, and SOC pools.

Determining Source Signatures

What to Measure

Partitioning CO2 fluxes using carbon isotope ratios requires direct measurement of carbon isotope ratios in CO2. That is, temporal, metabolic and kinetic fractionation effects can cause unpredictable isotopic differences between bulk organic matter and respired CO2 such that the adequacy of using only organic matter signatures and assuming they can act as surrogates for the respired CO2 signatures will depend on site conditions and in many cases will not work (Fig. 8.1). For example, Cheng (1996) found minimal difference between root respiration and ¿13C values of bulk root C under controlled growth conditions. However, in a naturally dynamic growth environment, it is not uncommon to find isotopic disequilibrium between bulk tissues and respired CO2. For example, during drought, significant differences can arise between 513C values of leaf organic matter and leaf respired CO2 (Duranceau et al., 1999; Ghashghaie et al., 2001), presumably as a result of transient reductions in discrimination caused by reduced stomatal conductance. Similar disequilibrium can occur with respect to whole plant respiration as the 513C values of metabolically active C in phloem can differ from the 513C values of leaf dry matter (Fig. 8.8; Pate and Arthur, 1998). This may in part stem from the fact that organic matter reflects C fixed at an earlier date (Smedley et al., 1991; Terwilliger, 2001) whereas respired CO2 reflects recent photosynthates (Pate, 2001). For example, leaf-to-phloem isotope differences reported by Pate and Arthur (1998) apparently arose because leaf dry matter was synthesized during relatively unstressed wet season conditions whereas leaf respiration during peak summer conditions (Fig. 8.8) reflected recent water stress.

Owing to the dynamic nature of carbon allocation among plant sources and sinks, the CO2 respired by the leaves may not reflect the CO2 respired by the roots. Aside from the frequently cited isotopic differences between

Figure 8.8 Differences between carbon isotope composition of leaf dry matter and phloem carbon during peak summer. Leaf dry matter was presumably synthesized during less-stressful wet season conditions. From Pate and Arthur (1998).

leaf and root organic matter (Gleixner et al, 1993; Schweizer et al, 1999; Arndt and Wanek, 2002), Terwilliger and Huang (1996) found a 1-3%o enrichment of heterotrophic plant parts adjacent to photoauto-trophic leaves. Similar results from other studies (Gleixner et al, 1998; Terwilliger, 2001) suggest that isotope effects related to C translocation (e.g., de-novo sucrose synthesis) and anaplerotic CO2 fixation can contribute to isotopic differences among organs and their metabolically active C pools. Anaplerotic CO2 fixation by roots or mycorrhizae may be particularly important during nitrogen assimilation (Wingler et al, 1996; Dunn, 1998).

It is not known if isotopic differences exist between fungal hyphae and sporocarps yet fruiting bodies are widely used to assess hyphal isotopic signatures (Hobbie et al, 1999; Hogberg et al, 1999; Kohzu et al, 1999). Thus, although there does not appear to be fractionation during fungal respiration (Henn and Chapela, 2000), further studies are needed to determine if sporocarp isotope signatures can be used to provide an accurate assessment of fungal <513C-CC>2 values. It should be noted that Chapela et al (2001) reported no isotopic difference between mycorrhizal root tips and the host plant, whereas fruit bodies of the same fungal species exhibited a 3%o difference relative to the host plant. In this specific case, unusually high rates of fungal productivity suggested that this isotopic gradient between fungal tissues adjacent to roots and their more distant fruit bodies resulted from the incorporation of 13C-enriched SOC as a result of significant saprotrophic activity. This notion was supported by radiocarbon dating that indicated the presence of older carbon in the fruit bodies, consistent with different C sources between the fungal tissues adjacent to the roots and their associated fruit bodies. Saprotrophic activity of ectomycorrhizal fungi (Gadgil and Gadgil, 1987; Durall et al., 1994) in addition to long-distance transport of C from interconnected mycelia (Simard et al., 1997) may be the cause of isotopic differences between ectomycorrhizal fungi and their apparent plant host (Fig. 8.3).

Clearly, the potential for isotopic disequilibrium between plant and fungal parts necessitates careful consideration as to what to measure in order to achieve a specific partitioning goal (e.g., root vs microbial, aboveground vs belowground).

When to Measure Source Signatures

Determination of what to measure also requires a consideration of when to measure, as carbon isotope signatures of respired C02 can exhibit significant variation on daily (Duranceau et al, 1999; Ghashghaie et al., 2001; Tcherkez et al., 2003) to seasonal timescales (Pate and Arthur, 1998). Seasonal variations in the isotopic composition of plant respiration are primarily driven by changes in photosynthetic discrimination, which are in turn driven by moisture limitations to stomatal conductance (Pate and Arthur, 1988) in addition to other factors such as air temperature, genotype, leaf age, etc. (see Dawson et al, 2002). On diel timescales, leaf respiration can be enriched in 13C during the day relative to its value near the end of the night (Duranceau et al, 1999; Tcherkez et al, 2003). This pattern is most pronounced in variable environments where daytime photosynthetic discrimination is limited due to water or temperature stress, and the resulting isotopic signature of the labile C pool (e.g., sucrose) is continuously changing relative to the more stable pool of C reserves which are respired near the end of the night after the labile pool is exhausted. The turnover time of this labile pool appears to be on the order of 4 hours (Fig. 8.9), implying that the best time to sample plant respiration for the purpose of partitioning nighttime respiration is at least 4 hours after sunset. Sooner than this, significant isotopic differences between plant parts may exist (Fig. 8.10) as a result of differential allocation and thus different amounts of labile C

Figure 8,9 Change in carbon isotope composition of leaf respiration with respect to duration in darkness. Initially (i = Ohr), the S^C value of respired C02 presumably reflects light-dependent and photosynthate-driven reactions involved in growth and biosynthesis of secondary compounds such as lipids (Park and Epstein, 1961). By mass balance, synthesis of such ^C-depleted compounds will result in the *®C-enriched C02 signals shown at t = 0. As the labile photosynthate pool becomes exhausted, the biosynthetic reactions will decrease, allowing the respired C02 signal to reflect that of the more stable pool of C reserves. The turnover times (r = 1/k) were determined by fitting first order decay functions (Y = Ae~lct) to the ¿13C values that were first adjusted so that values at hour 24 equaled 0%o (r2 ranged from 0.92 to 0.99). The values for Lycopersicon esculentum are from Park and Epstein (1961) whereas those for Arctostaphylos pajaroensis and Artemisia tridentata are unpublished results. All C02 samples were collected in chambers filled with C02-free air (see also Tcherkez et al., 2003).

Hours in darkness

Figure 8,9 Change in carbon isotope composition of leaf respiration with respect to duration in darkness. Initially (i = Ohr), the S^C value of respired C02 presumably reflects light-dependent and photosynthate-driven reactions involved in growth and biosynthesis of secondary compounds such as lipids (Park and Epstein, 1961). By mass balance, synthesis of such ^C-depleted compounds will result in the *®C-enriched C02 signals shown at t = 0. As the labile photosynthate pool becomes exhausted, the biosynthetic reactions will decrease, allowing the respired C02 signal to reflect that of the more stable pool of C reserves. The turnover times (r = 1/k) were determined by fitting first order decay functions (Y = Ae~lct) to the ¿13C values that were first adjusted so that values at hour 24 equaled 0%o (r2 ranged from 0.92 to 0.99). The values for Lycopersicon esculentum are from Park and Epstein (1961) whereas those for Arctostaphylos pajaroensis and Artemisia tridentata are unpublished results. All C02 samples were collected in chambers filled with C02-free air (see also Tcherkez et al., 2003).

and associated anabolic reactions in the various plant organs (e.g., synthesis of 13C-depleted lipids; Park and Epstein, 1961).

How to Measure Source Signatures

Arguably the most difficult aspect of determining source signatures is not what or when to sample but how to sample without introducing artefacts while isolating and collecting CO2 from the various respiration sources. Foliage respiration CO2 is relatively easy to collect using, for example, the chamber method described by Duranceau et al (1999) in which CO2 is collected in a C02-free chamber connected to an infra-red gas analyzer (IRGA) in a closed loop to monitor CO2 concentration. Whereas this approach has the possible disadvantage of introducing artefacts related to respiration into

Figure 8.10 Relationship between carbon isotope composition of leaf and whole plant respiration after 1 and 4h in darkness (unpublished results). The two sets of samples were collected on different dates (after 1 hr—January 11, 2002; after 4 hr—December 21, 2001).

<513C of whole plant respiration (%»)

Figure 8.10 Relationship between carbon isotope composition of leaf and whole plant respiration after 1 and 4h in darkness (unpublished results). The two sets of samples were collected on different dates (after 1 hr—January 11, 2002; after 4 hr—December 21, 2001).

CC>2-free air, it has the advantage of requiring only one isotope analysis per sample.

In contrast, Keeling plot approaches, such as those described by Fessenden and Ehleringer (2002) for measuring soil surface respiration (see also Hogberg and Ekblad, 1996; Ekblad and Hogberg, 2000) generally require a minimum of five samples to determine one source signature (over a substantial CO2 concentration range of >300ppm). This type of system is generally configured in a closed loop between a sample and the IRGA and can be modified for use with detached or attached leaves, or on boles, branches, and roots (with the appropriate chamber). Because Keeling plot approaches require a range of CO2 concentrations, the sample is thus typically subjected to elevated CO2 concentrations in contrast to the aforementioned CC>2-free method. Particular attention must be given to maintaining the volume and background isotopic signature. The latter is crucial so as not to introduce a third end-member into the system and confounding the interpretation. In general, care must be exercised with chamber approaches to avoid leaks and pressure-related effects on rates of CO2 efflux (Lund el al, 1999) as soil air that is 13C-enriched relative to the source (by up to ~4.4%o) can be drawn into the chamber airspace if the chamber pressure is less than ambient. In addition to chambers placed on the soil surface, soil gas probes (Rochette et al, 1999) and wells (Andrews et al, 1999) provide practical methods for characterizing the isotopic composition of soil-respired CO2 (see also Amundson et al, 1998).

A: Syringe method

A: Syringe method

B: Chamber keeling plot method

Vial

1 Place plant or soil sample in syringe

2 Pull air into syringe through CO? scrubber

3 Inject air into vial after ~S-10 minutes

To mass spec for 6i3C measurement

Allow COj to build up inside bottle and sample over time (replace volume of air removed with equal volume of background air)

Vial

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