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a Ecole Superieure d'Ingenieurs et de Techniciens Pour l'Agriculture, 13, rue du Nord, 76000 Rouen, France
b Department of Horticulture, Plant Science Building, Cornell University, Ithaca, NY 14853
Corresponding author (led24{at}cornell.edu)
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: ANOVA, analysis of variance • NPP, net primary productivity • POM, particulate organic matter • REF, reference • RL, root-labeled plot • SL, shoot-labeled plot • SOC, soil organic C • SOM, soil organic matter • 
13C, 13C natural abundance
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Aboveground and belowground NPP are fundamentally distinct in terms of interactions with soil microbial systems because C inputs from roots include root production, turnover, and exudation. The influence of roots on SOC pools could be relatively greater than the influence of aboveground C inputs because of the continuous release of C from roots and the complex nature of the rhizosphere–soil interface (Michulnas et al., 1985; Boone et al., 1994; Norby and Cotrufo, 1998). Roots influence aggregate stability directly by physically enmeshing soil particles and indirectly by stimulating microbial biomass that in turn synthesizes polymers that act as binding agents (Tisdall and Oades, 1979; Jastrow et al., 1998). The formation of aggregates protects SOC from biodegradation by reducing the access of decomposers to these encapsulated substrates (Elliott, 1986; Oades, 1988).
In SOM dynamics studies, particular attention is given to the POM fraction. This fraction, composed mainly of plant residues in different stages of decomposition, is regarded as a labile pool of SOM and is very sensitive to changes in C input and loss with time (Christensen, 1992). This fraction obtained by density or size separation has been used extensively as an early indicator of SOM dynamics (e.g., Janzen et al., 1992; Cambardella and Elliott, 1992; Gregorich and Ellert, 1993; Biederbeck et al., 1994). Because of its labile nature, the decomposition of POM can be strongly influenced by its location inside the soil structure (Golchin et al., 1994; Gregorich et al., 1997; Six et al., 1998).
The long-term goal of this experiment, which was designed to run for 3 to 5 yr, was to test the hypothesis that root-derived C persists in SOC pools to a greater extent than shoot-derived C. Additionally, we sought to identify factors controlling C dynamics and susceptibility to decomposition. Our investigation partitioned shoot-derived C from root-derived C during decomposition of hairy vetch, a leguminous plant that commonly serves as a green manure in annual cropping systems in the temperate zone. Our previous work comparing fertilizer-driven and legume-based cropping systems indicated that C derived from the green manure in the legume-based rotation may be retained in the soil longer than C derived from corn (Zea mays L.) residues (Drinkwater et al., 1998). We labeled hairy vetch in situ with 13CO2 and followed both root- and shoot-derived C in total SOC, CO2 released through soil respiration, microbial biomass C, and C in POM fractions during the growing season following hairy vetch incorporation. Here we report the results of the first 6 mo following the incorporation of the green manure.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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6.5 and cation-exchange capacity is 9.8 to 11.2 cmol kg-1 soil, mainly saturated by Ca (63–66%), Mg (10–12%), and K (<2%) (Penn State Agricultural Analytical Lab, State College, PA). In the plow layer (top 20 cm) soil C content is 3088 ± 343 g C m-2 and N content is 470 ± 36 g N m-2.
Labeling Procedure and Vetch Incorporation
We selected four sites within the organic legume-based treatment in which hairy vetch had been planted in September 1996 and allowed to overwinter. In early April 1997, four replicates consisting of three microplots (1.30 by 0.94 m) each were established. A 30-cm-deep metal frame was driven into the soil at each location to enclose the plow layer and support the labeling chamber (Berg et al., 1991). In one of the three microplots in each replicate, hairy vetch was labeled in situ with 99% enriched 13CO2 (ISOTEC Inc., Miamisburg, OH). At the initiation of labeling, hairy vetch aboveground biomass averaged 194 g m-2.
Microplots were enclosed with portable chambers and were pulse-labeled on 16, 21, 25, and 30 April, and 8 May with 1.5, 1.6, 1.6, 1.6, and 2.0 liters of 13CO2 for a cumulative total of 8.3 L per labeled microplot. The portable chamber consisted of a Tedlar plastic bag (Plastic Film Enterprise, Royal Oak, MI) supported by polyvinyl chloride tubing. During the labeling procedure, total CO2 concentration inside the labeling chamber was monitored using a portable Infra-Red Gas Analyzer (EGM1, PP system, Haverhill, MA). The chamber was removed when net CO2 uptake by the hairy vetch had stopped, usually after 45 to 65 min. Labeling was usually conducted between 1000 and 1130 h and 1300 and 1600 h. We followed a rotation among the field replications for the time of day when pulse-labeling occurred (Swinnen et al., 1996). Air temperatures in chambers during labeling were comparable between the control and the chamber receiving 13CO2, averaged 21 to 25°C, and normally did not exceed 
5°C above ambient air temperatures. There were two occasions when temperatures reached 31°C, which was 7°C above ambient.
On 12 May 1997 all microplots were subjected to simulated tillage and corn (Pioneer 3527) was planted. To establish three treatments per replication, shoot biomass from the labeled microplot was exchanged with shoot biomass from one of the unlabeled microplots in the same field replication, and then incorporated. This resulted in one microplot with labeled roots (RL) and unlabeled shoots, and one with labeled shoots (SL) and unlabeled roots. The third microplot was used as a reference (REF) for the 13C calculations and received only hairy vetch residues (both roots and shoots) with naturally occurring levels of 13C. Prior to incorporation, shoot biomass was weighed and subsampled to determine dry matter and to analyze for C, N, 13C, and litter chemistry. For vetch incorporation and corn planting, all microplots were treated in the same manner. Soil from each microplot was removed to a depth of 20 cm and weighed. As soil was returned to the microplot, coarsely chopped vetch shoots (
8 cm) were incorporated. Since this was an in situ experiment, roots were already in the soil and were coarsely chopped with shovels during the mixing process. Corn was planted in two rows and then thinned to a density of six corn plants per microplot, similar to the normal plant density.
Soil Sampling
Soil samples were collected on 16 April before pulse labeling started, on 12 May just before hairy vetch incorporation, and on 7 October at corn senescence (physiological stage of black layer). A composite sample consisting of 20 to 30 soil cores (1.5–2 kg total moist weight) was taken from each microplot to a depth of 0.2 m. Fresh soil was immediately processed for microbial biomass, standing root biomass determination, and soil moisture, whereas the remaining soil was air dried for additional analysis of organic matter fractions.
Estimation of Belowground Carbon Inputs
Both standing root biomass and total root-derived C inputs were determined. Standing root biomass was determined on 16 April and 12 May by wet sieving soil and sorting out living roots. The slurries resulting from these root determinations were retained and used for microbial biomass extraction as described below. To quantitatively extract roots, field moist soil (3 replicates per microplot, 125 g) was suspended in 300 mL of 0.05 M K2SO4. After shaking (1 h at 200 rev. min-1) the sample was wet sieved at 2 and 0.5 mm. Living roots were separated from sand and POM by hand, with the aid of tweezers. Visual criteria, such as color and elasticity, were used to distinguish living roots from dead roots and other organic residues. After removal of the roots, sand and POM were returned to the soil suspension for microbial biomass extraction. This standing root biomass assessment excluded very fine roots and dynamic C contributions from fine root turnover and root exudates and thus did not represent total root-derived inputs. To estimate total root-derived C inputs at vetch incorporation (12 May) we used the 13C enrichment of the whole soil and large POM in root-labeled plots relative to the 13C natural abundance (13C) values of soil from unlabeled reference plots. Enrichment values for root-derived C inputs used in the calculations was based on 13C of the hand picked living roots.
Particulate Organic Matter Fractionation
We performed two types of SOM separation. The first was based on the size of organic material, and it involved a complete dispersion of the soil followed by wet sieving (Christensen, 1992). Air-dried soil (50 g) was placed in a 250-mL Nalgene bottle with 125 mL of 0.5% (w/w) sodium hexametaphosphate and shaken for 16 h on a rotary shaker at 150 rev. min-1. The soil suspension was then sequentially wet sieved through sieves of 2000, 250, and 50 µm (stainless sieves, 10-cm diam., Newark Company, Newark, NJ). Organic material >50 µm in each of these resulting fractions was separated from the mineral particles (Feller, 1979). Briefly, minerals (sand) were separated from POM by densimetric separation in water after hand agitation, swirling, and decantation of the organic floating material. This fractionation yielded three POM fractions: >2000, 250 to 2000, and 50 to 250 µm, as well as a clay and silt fraction consisting of organo-mineral particles finer than 50 µm and soluble C. During protocol development, complete dispersion was confirmed by observation of POM under a dissecting microscope.
A second procedure was used to distinguish POM in terms of location inside (occluded POM) or outside (free POM) of stable aggregates. We adapted a procedure developed by Golchin et al. (1994). Air-dried soil (30 g) was gently shaken for 1 h at 100 rev. min-1 with 75 mL of sodium polytungstate (SOMETU-US, Van Nuys, CA) adjusted to a density of 1.7 g cm-3. The resulting soil suspension was allowed to settle for 16 h. The free POM on top of the solution was aspirated as described by Strickland and Sollins (1987). The recovered light fraction was washed with 90 mL of diH2O. The heavy fraction, consisting of mineral soil and POM trapped inside aggregates, was returned to the shaker for 16 h at 150 rev. min-1. After shaking, POM was recovered by wet sieving at 50 µm and separated from sand as described previously.
Measurement of Microbial Biomass
Soil microbial biomass was determined using the chloroform fumigation–extraction method (Brookes et al., 1985). However, because chloroform destroys cell membranes of both microorganisms and living roots, this procedure results in overestimation of microbial C and N in the presence of living roots. On the basis of the work of Mueller et al. (1992), we used a root-free slurry for the fumigation–extraction procedure for two early soil sampling dates when significant amounts of living hairy vetch roots were present. By October, there were essentially no living roots present, so soil samples taken then were treated using the conventional microbial biomass method (Voroney et al., 1993). In order to directly analyze the samples for C content and 13C, all samples were extracted with 0.05 M K2SO4 following Bruulsema and Duxbury (1996).
For all microbial biomass determinations, triplicate soil samples were processed within 1 wk from soil that had been stored at 4°C. For the root-free extractions, after living roots were separated from field moist soil as described above, the slurry was split into two subsamples for the control and fumigated extraction. For the October sampling date, field moist soil samples were fumigated directly.
All samples were fumigated in the presence of 10 to 20 mL of chloroform under a vacuum for 24 h in glass desiccators. For the root-free slurries, two or three drops of chloroform were also added directly to the slurry prior to fumigation (Mueller et al., 1992). Fumigated and control samples were extracted in 0.05 M K2SO4, (Bruulsema and Duxbury, 1996) and then filtered through a Buechner funnel fitted with a filter paper (Whatmann 41, Whatman, Clifton, NJ). For the slurry extractions, soil retained on the filter paper was dried at 60°C and the weight was recorded for future calculations. All extracts were stored in the freezer until they were lyophilized and ground for analysis. We present data as chloroform-extractable C since we did not use a Kc correction factor.
Soil Respiration
Soil respiration measurements began on 14 May (2 d after hairy vetch incorporation). Soil respiration was monitored in closed chambers (an inverted white bucket with an area of 0.06 m2) using 40 g of soda lime as a CO2 trap. This method is described in detail elsewhere (Edwards, 1982; Zibilske, 1994). Soda lime traps were left in the field for 24 h in the early part of the season (14 May–1 July) and for 48 h from July until freezing (24 Nov.). To collect continuous respiration data and allow time for drying and weighing of the soda lime, sets of two or three soda lime traps were alternated in the same microplot until their weight gain approached 7%. Soda lime from each set was then combined to give a composite sample representing 2 to 3 wk. A subsample was ground under N2 atmosphere for 13C analyses. To avoid modifying soil profiles, chambers were shifted between two sites within microplots every 24 to 48 h and chamber rings were moved every 8 to 10 d.
The static chamber technique usually overestimates CO2 fluxes when respiration rates are low, and underestimates CO2 flux when respiration rates exceed 0.24 g m-2 h-1 (Nay et al., 1994; Kaye and Hart, 1998). We used this data to determine the relative contribution of various sources and did not attempt to construct a C budget based on respiration data.
We used a mass balance approach to calculate the contribution of SOM pools to soil respiration. Four sources of CO2 were used in our calculations: (i) decomposition of native organic matter, (ii) decomposition of hairy vetch roots, (iii) decomposition of vetch shoots, and (iv) corn root respiration. We assumed that the values of 13CO2 reflected 13C of the source. This is probably a reasonable assumption for newly added C sources such as the vetch residues; however, we expect that the estimate for native SOM is approximate because of isotopic fractionation and also because of the alternation of C3 and C4 plants in the rotation. We also assumed that the decomposition of root and shoot residues was identical in both microplots regardless of the 13C signature. Thus, we were able to solve for three sources of 13C simultaneously because we had separate microplots for tracing labeled roots vs. labeled shoots. Because there were actually four sources of CO2 during most of the sampling period, we had to conduct our calculations in two steps.
First, the contribution of the native SOM decomposition was estimated as follows. During the first few weeks and after about the first week of October, corn root respiration was either absent or extremely low. Early in the season, plants were germinating and just beginning to develop roots. At the end of the season, corn plants reached physiological senescence in early October. We calculated the contribution of native SOM during these two periods using Eq. [1] and [2]
 | (1) |
 | (2) |
13CRL-CO2 represents 13C of CO2 respired from RL microplots, 13CRL represents 13C of labeled vetch roots, 13CSU represents 13C of unlabeled vetch shoots, 13CSL-CO2 represents 13C of CO2 respired from SL microplots, 13CSL represents 13C of labeled vetch shoots, 13CRU represents 13C of unlabeled vetch roots, and 13CN-OM represents 13C of total SOM in reference plots. Using these equations, we solved for x, which is the proportion of CO2 respired from root residues, for y, the proportion of CO2 respired from shoot residues, and for (1 - x - y), the proportion of CO2 respired from native SOM. Equation [1] used data from the root-labeled microplot, while data from the shoot-labeled plot was used in Eq. [2]. First the equations were algebraically rearranged to solve for x (Eq. [1]) and y (Eq. [2]). Then, to solve Eq. [1] for x, we replaced the unknown y with the equation for y (modified Eq. [2]) so that Eq. [1] contained only one unknown (x).
We calculated CO2 respired from native SOM when corn root respiration was absent to be 0.59 + 0.01 g C m-2 d-1. This value was then extrapolated over the growing season and subtracted from total CO2 respired at each sampling date. The 13C signature of CO2 respired was adjusted to reflect the absence of CO2 derived from native SOM. We did not make adjustments for soil temperature because, although there was a twofold difference in soil temperature among the four time points used in this calculation, CO2 evolved from native SOM was not significantly correlated with soil temperature (Pearson correlation, r = 0.15, P > 0.05).
After CO2 derived from native SOM had been removed from total CO2 respired, we could replace the native SOM term with an unknown for corn root respiration. We used Eq. [3] and [4] to solve for the proportion of CO2 respired from root residues (x), shoot residues (y), and corn root respiration (1 - x - y) for the remaining sampling dates when corn root respiration was present. Parameters defined above remain the same, and one additional known value was added: 13CCorn = 13C of corn roots. We used the value of -12
for the signature of CO2 respired from corn roots (Rochette and Flanagan, 1997).
 | (3) |
 | (4) |
Analytical Methods
Plant residue samples and POM fractions were dried at 60°C for 48 h. Carbon, N, and 13C signature were determined after dry combustion on a Europa CN auto-analyzer coupled to a Europa Tracermass mass spectrometer (Europa Scientific Ldt., Crewe, UK). Litter biochemistry was determined for hairy vetch shoot residues from all four replicate microplots. The hand picked root samples from each replicate were needed for 13C determinations, so two additional composite samples were collected from locations other than the microplots and roots were sorted using the same protocol described above. A total of eight shoot samples and two root samples were analyzed gravimetrically for cellulose, hemicellulose, cell walls, and lignin contents after digestion in neutral detergent and acid solutions according to the Van Soest method (Van Soest and Wine, 1968). Nonstructural carbohydrates were obtained after hydrolysis in a 0.1 M H2SO4 solution and quantified by titration with sodium thiosulfate (Analysis Lab., Colorado State University).
Statistics
Statistical analyses were done using Statistica 4.5 (Statsoft). The experiment was a randomized complete block design with four replications and three treatments (REF, RL, and SL microplots). Total C pools were compared using one-way analysis of variance (ANOVA) with date as the main effect, or separately by sampling date with source as the main effect (RL vs. SL). Differences in C pools (origin or location) were also tested with two-way ANOVAs using treatment x date as the error term. Post hoc analyses were done using LSD Fisher comparison tests.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Hairy Vetch Biomass: 13C Values and C Inputs
At incorporation (May 12), labeled hairy vetch biomass was highly enriched with 13C compared with the unlabeled biomass. The 13C signature obtained was +163.81 for the shoots, and +60.05 for the roots (Table 1), compared with 13C of -26.73 and -27.98 for their unlabeled counterparts. The signature was 2.5 times greater in shoots than in roots (Table 1), and variability across the field replicates was high (coefficient of variation 39% for the signal in both aboveground and belowground biomass). Before incorporation, labeled hairy vetch belowground C inputs significantly increased the whole soil 13C by +1.86 (±0.23 SE, ANOVA, P = 0.0045). This enrichment in 13C was still significant at the end of the season, when it averaged +0.73 for the soil receiving the labeled root C (± 0.12 SE, ANOVA, P = 0.052) and +1.26 for the soil amended with labeled shoots (±0.13 SE, ANOVA, P = 0.002).
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13C values of soil from unlabeled plots, we calculated that the total root-derived C at the time of incorporation was 48.2 g C m-2 (Table 2). Thus, standing root biomass estimates accounted for at least 60% of the hairy vetch root-derived C inputs at the time of incorporation. Compared with total soil C, C inputs from hairy vetch were small, equal to 6.5% of the total soil C.
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30% of new C (15.3 g C m-2). Aboveground vetch-derived C was composed of large pieces of standing biomass and was incorporated as large POM (>2000 µm).
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50% of the total new vetch C present in the soil. Between 12 May and 7 October, the amount of root-derived C decreased significantly in all size fractions except for the intermediate POM fraction, where it increased. The most drastic decrease occurred in the macro-organic matter fraction (POM >2000 µm). Particulate organic matter from both aboveground and belowground sources was preferentially located inside soil aggregates (occluded POM) at both sampling dates (ANOVA, P = 0.0085; Fig. 2) . Furthermore, the decrease in root-derived C from free POM between 12 May and 7 October was greater than the decrease in the occluded POM fraction (80 vs. 57% decreases, respectively). At the end of the season, a greater proportion of root-derived C was present in both free and occluded POM fractions, compared to shoot-derived C. This difference was particularly pronounced in the occluded fraction, where twice as much C was derived from roots as from shoots (ANOVA, P = 0.049).
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13C signature of microbial biomass showed a significant response to the input of labeled residues. However, assuming that new C assimilated by the microbial biomass had the same 13C signature as the labeled residues, only a small proportion of root- and shoot-derived C was assimilated by the microbial biomass (Table 3). Vetch-derived microbial biomass C ranged from 10.5 to 12.7% of the total microbial biomass C. There were no significant differences in vetch-derived microbial C between the two sampling dates or between root- and shoot-derived C (ANOVA, P = 0.63 and 0.28, respectively).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Field studies focusing on the N dynamics in legume-based rotations clearly suggest that green manure residues are rapidly decomposed during the first few weeks after incorporation, resulting in significant net N mineralization (Sarrantonio and Scott, 1988; Drinkwater et al., 2000). Based on the 13C signature of the respired CO2, it is likely that biodegradation of shoot residues is probably the primary source of this rapid increase in mineral N.
Dynamics of Hairy Vetch Carbon in POM Fractions
Because root-derived C was present in all POM fractions in both May and October, we were able to evaluate the turnover rates in the various fractions. Carbon losses were greater in POM fractions composed of larger pieces of organic matter, confirming that larger POM fractions turn over quite rapidly. Others have reported that the largest POM fractions have very rapid turnover times, usually <30 d (Buyanovsky et al., 1994; Balesdent, 1996; Cambardella and Elliott, 1992). In our study, roots in the coarsest POM fraction (>2000 µm) had a half life of 40 d. Only the 50- to 250-µm POM fraction showed an increase in root-derived C between May and October, indicating this fraction is mainly composed of partially decomposed POM originating from larger fractions (Balesdent, 1996). Occluded POM and the C in the clay and silt fraction decomposed at a slower rate and had a half life of 122 d. Thus, occluded POM-C from hairy vetch root origin had a slower turnover rate than did free POM C. The greater retention of root-derived POM results from its greater presence as occluded POM. By October, 22% of the initial root C was present as occluded POM, while only 4% of the shoot derived C had been retained as occluded POM.
Dynamics of Labeled Carbon through the Soil Microbial Compartment
At each of the three sampling dates, a relatively small amount of vetch-derived C was present as chloroform-extractable microbial biomass. By October, 158 d after tillage and shoot incorporation, hairy vetch C accounted for 10% of the total microbial biomass C pool. Conversion rates for substrates to microbial biomass ranging from 0.1 to 15.3% have been reported previously, primarily in microcosm studies (Merckx et al., 1985; Ladd and Amato, 1988). These seemingly low conversion rates reflect shortcomings of the fumigation–extraction method, which only extracts a fraction of the cytoplasmic microbial biomass (Brookes et al., 1985; Tate et al., 1988) and also does not select for the active portion of the microbial biomass (Ladd et al, 1995; Qian et al., 1997; Hu and Van Bruggen, 1998).
The vetch contribution to microbial biomass was composed of equal parts of C from shoot and root origins. However, a greater proportion of root C was converted to microbial biomass compared with shoot-derived C (7 mg g-1 of shoot C respired compared with 24 mg g-1 of root C respired was retained as microbial biomass). The mechanisms leading to an increased retention of root C in the cytoplasmic microbial biomass during decomposition is not clear, although Jans-Hammermeister et al. (1998) found that a greater proportion of glucose C added as small daily aliquots was retained compared with glucose C added in a single large aliquot. One fundamental difference between root- and shoot-derived C is the continuous nature of root C inputs in the form of root exudates and fine-root turnover during plant growth.
Estimation of Corn Root Carbon Contributions to Soil Organic Carbon
On the basis of shifts in 13C natural abundance in our reference plots, we were able to estimate corn root contributions to different SOC compartments at corn senescence. Corn root C inputs to the top 20 cm of soil were threefold greater than C inputs from hairy vetch roots (corn: 157 g C m-2 vs. vetch: 48.22 g C m-2). The distribution of corn root-derived C among POM compartments was quite different than for the hairy vetch root-derived C. Corn root C accounted for 38% of the free POM C and only 4.3% of the C in the occluded POM, whereas hairy vetch root C was more evenly distributed between these two fractions. Hairy vetch root-derived C accounted for 10% of the occluded POM C. This suggests that hairy vetch roots were more effective at promoting aggregation than were corn roots, which could lead to an overall slower turnover rate of vetch root litter compared with corn root litter. In cropping system studies where bare fallow was replaced by a leguminous green manure, increased levels of water stable aggregation have been documented (Roberson et al., 1991). Furthermore, some plants such as corn, tomato (Lycopersicon esculentum Mill. var. esculentum), and wheat (Triticum aestivum L.) actually decreased aggregate stability while growth of perennial ryegrass (Lolium perenne L.) and lucerne (Medicago sativa L. subsp. sativa) tended to increase it (Reid and Goss, 1981). This increased aggregate stability has been attributed to polysaccarhides produced in the rhizosphere (Reid and Goss, 1981) and increased fungal populations associated with these species (Tisdall and Oades, 1979; Haynes and Beare, 1997).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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ACKNOWLEDGMENTS |
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Received for publication December 13, 1999.
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