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Reciprocal transfer of carbon and nitrogen by decomposer fungi at the soil–litter interface
Soil Biology & Biochemistry 35 (2003) 1001–1004 www.elsevier.com/locate/soilbio
Short communication
Reciprocal transfer of carbon and nitrogen by decomposer fungi at the soil –litter interface S.D. Freya,*, J. Sixb, E.T. Elliottc a Department of Natural Resources, University of New Hampshire, Durham, NH 03824, USA Department of Agronomy and Range Science, University of California, Davis, CA 95616, USA c School of Natural Resource Sciences, University of Nebraska, Lincoln, NE 68588, USA
b
Received 29 April 2002; received in revised form 11 March 2003; accepted 28 March 2003
Abstract We have investigated whether decomposer fungi translocate litter-derived C into the underlying soil while simultaneously translocating soil-derived inorganic N up into the litter layer. We also located and quantified where the translocated C is deposited within the soil aggregate structure. When 13C-labeled wheat straw was decomposed on the surface of soil amended with 15N-labeled inorganic N, we found that C and N were reciprocally transferred by fungi, with a significant quantity (121 –151 mg C g21 whole soil) of litter-derived C being deposited into newly formed macroaggregates (. 250 mm sized aggregates). Fungal inhibition reduced fungal biomass and the bidirectional C and N flux by approximately 50%. The amount of litter-derived C found in macroaggregates was positively correlated with litter-associated fungal biomass. This fungal-mediated litter-to-soil C transfer, which to our knowledge has not been demonstrated before for saprophytic fungi, may represent an important mechanism by which litter C enters the soil and becomes stabilized as soil organic matter within the macroaggregate structure. q 2003 Elsevier Science Ltd. All rights reserved. Keywords: Carbon; Decomposition; Fungi; Litter; Nitrogen; Translocation
Filamentous fungi are unique amongst the soil biota in their ability to translocate nutrients, a trait enabling them to establish extensive hyphal networks in a heterogeneous environment and to utilize spatially separated nutrient resources that may differ significantly in nutrient quality. Wood decay fungi, in particular, are well known to translocate significant quantities of C, nutrients (especially N, P, and S), and base cations through their mycelial cord and rhizomorph systems (Wells and Boddy, 1995a,b; Connolly and Jellison, 1997; Lindahl et al., 1999). Translocation is often viewed as a unidirectional process, with movement from a nutrient source (e.g. soil) to a sink (e.g. wood). However, bidirectional translocation of nutrients by wood decay fungi has also been observed (Connolly and Jellison, 1997; Lindahl et al., 2001). While extensive research has been carried out with cordforming, wood decay fungi, much less is known about fungal translocation associated with litter decomposition. * Corresponding author. Tel.: þ1-44-603-8623880; fax: þ 1-44-6038624976. E-mail address: [email protected] (S.D. Frey).
We have demonstrated (Frey et al., 2000) that litter decomposers can bridge the soil – litter interface and transport a significant quantity of soil inorganic N up into the plant litter layer, stimulating fungal biomass production. Here we tested whether decomposer fungi simultaneously translocate litter-derived C into the underlying soil. To do this, we carried out a two-way factorial experiment with two litter treatments (fungicide and no fungicide), two soil inorganic N concentrations (10 and 100 mg N g21 soil) and three replicates per treatment. The soil used was a Weld silt loam classified as a fine, montmorillonitic, mesic aridic Paleustoll (41% sand, 36% silt, and 23% clay) collected from the Central Great Plains Research Station at Akron, CO, USA (Halvorson et al., 1997). This soil was selected to be representative of a cultivated soil and because it has a low N mineralization potential, minimizing dilution of added inorganic 15N with native unlabeled N during the incubation period. The soil was air dried and sieved (250 mm) to remove large macroaggregates. The sand remaining on the sieve was then added back to the sieved soil so as not to alter the soil
0038-0717/03/$ - see front matter q 2003 Elsevier Science Ltd. All rights reserved. doi:10.1016/S0038-0717(03)00155-X
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S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
texture. Subsamples of 150 g of air-dried soil were added to incubation units consisting of 9 cm dia. plastic containers. This amount of soil was used because it gave a 2.5 cm layer of soil in each unit, mimicking the narrow interface zone between litter and surface soil in a no-tillage system. The soil was amended with 10 or 100 mg N g21 soil as (15NH4)2SO4 (5.0 at.%) in sufficient deionized water to bring the soil to field capacity (0.19 g g21). The total organic C, organic N, and inorganic N concentrations of unamended soil were 6820 ^ 948, 820 ^ 105, and 10.94 ^ 0.38 mg g21 soil, respectively. Following soil N amendment, 1.3 g (dry weight basis) of untreated or fungicide treated, 13C-labeled wheat straw (40.48% C, 0.659% N, d13C: 1803 ^ 2.44‰) was placed on the soil surface. This C input amount is equivalent to approximately twice the annual C input rate in a no-tillage wheat-fallow cropping system. Fungicide-treated litter material was prepared by soaking each sample (1.3 g) overnight in 20 ml deionized water containing the fungicide Captan (50 W, wettable powder, 48.7% active ingredient, 2.1% N) in the amount of 4.4 mg ml21, equivalent to 68 mg Captan g21 dry litter. This application rate was used successfully to significantly inhibit fungal populations in a field study (Frey et al., 2000). Untreated litter (no fungicide) was soaked in deionized water only. At experiment initiation, a circular piece of nylon mesh (3 £ 5 mm2 openings) was placed between the soil and surface litter to reduce soil contamination of the litter and facilitate its removal at the end of the incubation. The containers were sealed inside 1.9 l food jars and incubated at 25 8C. A small amount of water was placed in the bottom of each jar to maintain a humid atmosphere and reduce soil moisture loss. Every 2 – 3 d, the jars were opened and flushed with compressed air to maintain aerobic conditions. After 35 d, surface litter was removed and analyzed for gravimetric moisture content, fungal biomass, total C and N concentrations, and 15N enrichment. The soil was separated into aggregate size classes and analyzed for total organic C and 13 C isotopic composition. Total physiologically active fungal biomass associated with the wheat litter was measured using a substrateinduced respiration (SIR) method developed for plant residues (Beare et al., 1991). Total fungal biomass C was estimated using the equation: 231.5 þ 17.3 (fungal SIR), where fungal biomass C has units of mg C g21 residue dry weight and fungal SIR has units of mg CO2 – C g21 residue dry weight h21 (Beare et al., 1991). The size distribution of water-stable aggregates was measured by the method described by Elliott (1986). Briefly, a 100 g subsample of air-dried soil was wet sieved through a series of three sieves to obtain the following aggregate size fractions: . 2000, 250– 2000, 53– 250 and , 53 mm. Soil remaining on each sieve was backwashed into an aluminum pan, dried overnight at 50 8C, and weighed. Total organic C and N were determined on finely ground litter and soil subsamples by dry combustion using a NC2100 soil
analyzer (CE Elantech, Lakewood, NJ). These data are reported on an oven dry (105 8C) basis. The amount of litter-associated N derived from the soil inorganic N pool and the amount of wheat-derived C that entered the soil during the incubation was determined by measuring the isotopic composition (d15N and d13C) of finely ground litter, whole soil and soil aggregate fractions using a Carlo Erba NA 1500 CN analyzer (Carlo Erba, Milan, Italy) coupled to a Micromass VG isochrom-EA mass spectrometer (Micromass UK Ltd. Manchester, UK). Results were expressed as
d15 N‰ ¼ ½ð15 Rsample =15 Rstandard Þ 2 1 £ 1000 d13 C‰ ¼ ½ð13 Rsample =13 Rstandard Þ 2 1 £ 1000 where 15R ¼ 15N/14N and 13R ¼ 13C/12C. The proportion of litter-derived C(f) in the soil was calculated using the equation f ¼ ðdt 2 ds Þ=ðdw 2 ds Þ where dt is the isotopic composition of the whole soil or aggregate fraction at the end of the incubation, ds the isotopic composition of the original soil (2 17.86 ^ 0.46‰), and dw the isotopic composition of the wheat material (1803 ^ 2.44‰). We found that significant quantities of C and N were reciprocally transferred (Figs. 1 and 2). When fungal growth was not inhibited (no fungicide), 2.1– 4.7% of the total soil C was litter-derived after just 5 weeks. Fungal biomass C on untreated wheat straw was 4.42 ^ 0.72 mg g21 litter. Treating litter with a fungicide reduced fungal biomass by 47% on average (to 2.34 ^ 0.43 mg fungal C g21 litter) and both C and N transfer by nearly 50%. Nitrogen transfer in sterile controls accounted for less than 8% of total upward N transport (Frey et al., 2000), indicating that abiotic processes did not contribute substantially to nutrient flux under our experimental conditions. When we increased soil
Fig. 1. Soil-derived 15N found in the litter layer. Error bars represent 1 standard error of the mean.
S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
Fig. 2. Litter C deposited into three soil aggregate fractions. Error bars represent 1 standard error of the mean for the total amount of litter-derived soil C across all three aggregate size classes within a given litter treatment and soil N level ðn ¼ 3Þ:
inorganic N from 10 to 100 mg N g21 soil, we found a 7-fold increase in N flow into the litter (Fig. 1), indicating that fungal N translocation is stimulated by increased soil N availability. While there was a trend toward increased litterderived soil C when soil N concentrations were increased (Fig. 2), the effect of soil N availability on C flux was not significant. Since filamentous fungi influence soil structure by binding soil particles together into aggregates (Tisdall and Oades, 1982), and aggregates stabilize organic matter by physically protecting labile substrates from microbial attack (Elliott, 1986; Beare et al., 1994), we were interested in determining, where the litter-derived C was located within the soil matrix. A large proportion (68%) of this C in the control treatment was found in soil macroaggregates (. 250 mm; Fig. 2)—an aggregate fraction known to play a significant role in the stabilization of soil organic matter
Fig. 3. Relationships between litter-associated fungal biomass and the amount of litter-derived C found in soil aggregate fractions (n ¼ 12 for each aggregate fraction).
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(Six et al., 2000). Litter-derived C concentrations in macroaggregates were significantly correlated with the amount of fungal biomass observed in the litter layer (Fig. 3; r 2 ¼ 0:68; P , 0:05), providing further evidence that the downward flow of C can be attributed to fungal activity. There was not a strong relationship between fungal biomass and the labeled C found in microaggregates (53 – 250 mm) and non-aggregated soil (, 53 mm fraction). We conclude that litter-associated fungi carry out simultaneous, bidirectional translocation of soil-derived N and litter-derived C, and in so doing, transport a significant quantity of C belowground. Most of this C is preferentially located in soil macroaggregates, where we hypothesize, it is physically protected from decay and subsequently stabilized as soil organic matter. Fungal-mediated litterto-soil C transfer has not previously been quantified in terrestrial ecosystems. If this process is widespread, it could represent a potentially important mechanism of soil C sequestration.
Acknowledgements This work was supported by grants from the Ohio Agricultural Research and Development Center of the Ohio State University and the US Department of Agriculture. Sylvie Recous provided the 13C-labeled wheat. We thank Tanna Holtrey and Rodney Simpson for technical assistance. Comments by two anonymous reviewers significantly improved the quality of the manuscript.
References Beare, M.H., Neely, C.L., Coleman, D.C., Hargrove, W.L., 1991. Characterization of a substrate-induced respiration method for measuring fungal, bacterial and total microbial biomass on plant residues. Agriculture, Ecosystems and Environment 34, 65– 73. Beare, M.H., Cabrera, M.L., Hendrix, P.F., Coleman, D.C., 1994. Aggregate-protected and unprotected organic matter pools in conventional tillage and no-tillage soils. Soil Science Society of America Journal 58, 787–795. Connolly, J.H., Jellison, J., 1997. Two-way translocation of cations by the brown rot fungus Gloeophyllum trabeum. International Biodeterioration and Biodegradation 39, 181 –188. Elliott, E.T., 1986. Aggregate structure and C, N and P in native and cultivated soils. Soil Science Society of America Journal 50, 627– 633. Frey, S.D., Elliott, E.T., Paustian, K., Peterson, G.A., 2000. Fungal translocation as a mechanism of exogenous nitrogen inputs to decomposing surface residues in a no-tillage agroecosystem. Soil Biology & Biochemistry 32, 689– 698. Halvorson, A.D., Vigil, M.F., Peterson, G.A., Elliott, E.T., 1997. Longterm tillage and crop residue management study at Akron, Colorado. In: Paul, E.A., Elliott, E.T., Paustian, K., Cole, C.V. (Eds.), Soil Organic Matter in Temperate Agroecosystems: Long-Term Experiments in North America, CRC Press, Boca Raton, FL, pp. 361–370.
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S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
Lindahl, B., Finlay, Rl., Olsson, S., 2001. Simultaneous, bidirectional translocation of 32P and 33P between wood blocks connected by mycelial cords of Hypholoma fasciculare. New Phytologist 150, 189– 194. Six, J., Elliott, E.T., Paustian, K., 2000. Soil macroaggregate turnover and microaggregate formation: a mechanism for carbon sequestration under no-tillage agriculture. Soil Biology & Biochemistry 32, 2099–2103.
Tisdall, J.M., Oades, J.M., 1982. Organic matter and water-stable aggregates in soils. Journal of Soil Science 33, 141–163. Wells, J.M., Boddy, L., 1995a. Phosphorus translocation by saprotrophic basidiomycete mycelial cord systems on the floor of a mixed deciduous woodland. Mycological Research 99, 977 –980. Wells, J.M., Boddy, L., 1995b. Translocation of soil-derived phosphorus in mycelial cord systems in relation to inoculum resource size. FEMS Microbiology Ecology 17, 67 –75.
Short communication
Reciprocal transfer of carbon and nitrogen by decomposer fungi at the soil –litter interface S.D. Freya,*, J. Sixb, E.T. Elliottc a Department of Natural Resources, University of New Hampshire, Durham, NH 03824, USA Department of Agronomy and Range Science, University of California, Davis, CA 95616, USA c School of Natural Resource Sciences, University of Nebraska, Lincoln, NE 68588, USA
b
Received 29 April 2002; received in revised form 11 March 2003; accepted 28 March 2003
Abstract We have investigated whether decomposer fungi translocate litter-derived C into the underlying soil while simultaneously translocating soil-derived inorganic N up into the litter layer. We also located and quantified where the translocated C is deposited within the soil aggregate structure. When 13C-labeled wheat straw was decomposed on the surface of soil amended with 15N-labeled inorganic N, we found that C and N were reciprocally transferred by fungi, with a significant quantity (121 –151 mg C g21 whole soil) of litter-derived C being deposited into newly formed macroaggregates (. 250 mm sized aggregates). Fungal inhibition reduced fungal biomass and the bidirectional C and N flux by approximately 50%. The amount of litter-derived C found in macroaggregates was positively correlated with litter-associated fungal biomass. This fungal-mediated litter-to-soil C transfer, which to our knowledge has not been demonstrated before for saprophytic fungi, may represent an important mechanism by which litter C enters the soil and becomes stabilized as soil organic matter within the macroaggregate structure. q 2003 Elsevier Science Ltd. All rights reserved. Keywords: Carbon; Decomposition; Fungi; Litter; Nitrogen; Translocation
Filamentous fungi are unique amongst the soil biota in their ability to translocate nutrients, a trait enabling them to establish extensive hyphal networks in a heterogeneous environment and to utilize spatially separated nutrient resources that may differ significantly in nutrient quality. Wood decay fungi, in particular, are well known to translocate significant quantities of C, nutrients (especially N, P, and S), and base cations through their mycelial cord and rhizomorph systems (Wells and Boddy, 1995a,b; Connolly and Jellison, 1997; Lindahl et al., 1999). Translocation is often viewed as a unidirectional process, with movement from a nutrient source (e.g. soil) to a sink (e.g. wood). However, bidirectional translocation of nutrients by wood decay fungi has also been observed (Connolly and Jellison, 1997; Lindahl et al., 2001). While extensive research has been carried out with cordforming, wood decay fungi, much less is known about fungal translocation associated with litter decomposition. * Corresponding author. Tel.: þ1-44-603-8623880; fax: þ 1-44-6038624976. E-mail address: [email protected] (S.D. Frey).
We have demonstrated (Frey et al., 2000) that litter decomposers can bridge the soil – litter interface and transport a significant quantity of soil inorganic N up into the plant litter layer, stimulating fungal biomass production. Here we tested whether decomposer fungi simultaneously translocate litter-derived C into the underlying soil. To do this, we carried out a two-way factorial experiment with two litter treatments (fungicide and no fungicide), two soil inorganic N concentrations (10 and 100 mg N g21 soil) and three replicates per treatment. The soil used was a Weld silt loam classified as a fine, montmorillonitic, mesic aridic Paleustoll (41% sand, 36% silt, and 23% clay) collected from the Central Great Plains Research Station at Akron, CO, USA (Halvorson et al., 1997). This soil was selected to be representative of a cultivated soil and because it has a low N mineralization potential, minimizing dilution of added inorganic 15N with native unlabeled N during the incubation period. The soil was air dried and sieved (250 mm) to remove large macroaggregates. The sand remaining on the sieve was then added back to the sieved soil so as not to alter the soil
0038-0717/03/$ - see front matter q 2003 Elsevier Science Ltd. All rights reserved. doi:10.1016/S0038-0717(03)00155-X
1002
S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
texture. Subsamples of 150 g of air-dried soil were added to incubation units consisting of 9 cm dia. plastic containers. This amount of soil was used because it gave a 2.5 cm layer of soil in each unit, mimicking the narrow interface zone between litter and surface soil in a no-tillage system. The soil was amended with 10 or 100 mg N g21 soil as (15NH4)2SO4 (5.0 at.%) in sufficient deionized water to bring the soil to field capacity (0.19 g g21). The total organic C, organic N, and inorganic N concentrations of unamended soil were 6820 ^ 948, 820 ^ 105, and 10.94 ^ 0.38 mg g21 soil, respectively. Following soil N amendment, 1.3 g (dry weight basis) of untreated or fungicide treated, 13C-labeled wheat straw (40.48% C, 0.659% N, d13C: 1803 ^ 2.44‰) was placed on the soil surface. This C input amount is equivalent to approximately twice the annual C input rate in a no-tillage wheat-fallow cropping system. Fungicide-treated litter material was prepared by soaking each sample (1.3 g) overnight in 20 ml deionized water containing the fungicide Captan (50 W, wettable powder, 48.7% active ingredient, 2.1% N) in the amount of 4.4 mg ml21, equivalent to 68 mg Captan g21 dry litter. This application rate was used successfully to significantly inhibit fungal populations in a field study (Frey et al., 2000). Untreated litter (no fungicide) was soaked in deionized water only. At experiment initiation, a circular piece of nylon mesh (3 £ 5 mm2 openings) was placed between the soil and surface litter to reduce soil contamination of the litter and facilitate its removal at the end of the incubation. The containers were sealed inside 1.9 l food jars and incubated at 25 8C. A small amount of water was placed in the bottom of each jar to maintain a humid atmosphere and reduce soil moisture loss. Every 2 – 3 d, the jars were opened and flushed with compressed air to maintain aerobic conditions. After 35 d, surface litter was removed and analyzed for gravimetric moisture content, fungal biomass, total C and N concentrations, and 15N enrichment. The soil was separated into aggregate size classes and analyzed for total organic C and 13 C isotopic composition. Total physiologically active fungal biomass associated with the wheat litter was measured using a substrateinduced respiration (SIR) method developed for plant residues (Beare et al., 1991). Total fungal biomass C was estimated using the equation: 231.5 þ 17.3 (fungal SIR), where fungal biomass C has units of mg C g21 residue dry weight and fungal SIR has units of mg CO2 – C g21 residue dry weight h21 (Beare et al., 1991). The size distribution of water-stable aggregates was measured by the method described by Elliott (1986). Briefly, a 100 g subsample of air-dried soil was wet sieved through a series of three sieves to obtain the following aggregate size fractions: . 2000, 250– 2000, 53– 250 and , 53 mm. Soil remaining on each sieve was backwashed into an aluminum pan, dried overnight at 50 8C, and weighed. Total organic C and N were determined on finely ground litter and soil subsamples by dry combustion using a NC2100 soil
analyzer (CE Elantech, Lakewood, NJ). These data are reported on an oven dry (105 8C) basis. The amount of litter-associated N derived from the soil inorganic N pool and the amount of wheat-derived C that entered the soil during the incubation was determined by measuring the isotopic composition (d15N and d13C) of finely ground litter, whole soil and soil aggregate fractions using a Carlo Erba NA 1500 CN analyzer (Carlo Erba, Milan, Italy) coupled to a Micromass VG isochrom-EA mass spectrometer (Micromass UK Ltd. Manchester, UK). Results were expressed as
d15 N‰ ¼ ½ð15 Rsample =15 Rstandard Þ 2 1 £ 1000 d13 C‰ ¼ ½ð13 Rsample =13 Rstandard Þ 2 1 £ 1000 where 15R ¼ 15N/14N and 13R ¼ 13C/12C. The proportion of litter-derived C(f) in the soil was calculated using the equation f ¼ ðdt 2 ds Þ=ðdw 2 ds Þ where dt is the isotopic composition of the whole soil or aggregate fraction at the end of the incubation, ds the isotopic composition of the original soil (2 17.86 ^ 0.46‰), and dw the isotopic composition of the wheat material (1803 ^ 2.44‰). We found that significant quantities of C and N were reciprocally transferred (Figs. 1 and 2). When fungal growth was not inhibited (no fungicide), 2.1– 4.7% of the total soil C was litter-derived after just 5 weeks. Fungal biomass C on untreated wheat straw was 4.42 ^ 0.72 mg g21 litter. Treating litter with a fungicide reduced fungal biomass by 47% on average (to 2.34 ^ 0.43 mg fungal C g21 litter) and both C and N transfer by nearly 50%. Nitrogen transfer in sterile controls accounted for less than 8% of total upward N transport (Frey et al., 2000), indicating that abiotic processes did not contribute substantially to nutrient flux under our experimental conditions. When we increased soil
Fig. 1. Soil-derived 15N found in the litter layer. Error bars represent 1 standard error of the mean.
S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
Fig. 2. Litter C deposited into three soil aggregate fractions. Error bars represent 1 standard error of the mean for the total amount of litter-derived soil C across all three aggregate size classes within a given litter treatment and soil N level ðn ¼ 3Þ:
inorganic N from 10 to 100 mg N g21 soil, we found a 7-fold increase in N flow into the litter (Fig. 1), indicating that fungal N translocation is stimulated by increased soil N availability. While there was a trend toward increased litterderived soil C when soil N concentrations were increased (Fig. 2), the effect of soil N availability on C flux was not significant. Since filamentous fungi influence soil structure by binding soil particles together into aggregates (Tisdall and Oades, 1982), and aggregates stabilize organic matter by physically protecting labile substrates from microbial attack (Elliott, 1986; Beare et al., 1994), we were interested in determining, where the litter-derived C was located within the soil matrix. A large proportion (68%) of this C in the control treatment was found in soil macroaggregates (. 250 mm; Fig. 2)—an aggregate fraction known to play a significant role in the stabilization of soil organic matter
Fig. 3. Relationships between litter-associated fungal biomass and the amount of litter-derived C found in soil aggregate fractions (n ¼ 12 for each aggregate fraction).
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(Six et al., 2000). Litter-derived C concentrations in macroaggregates were significantly correlated with the amount of fungal biomass observed in the litter layer (Fig. 3; r 2 ¼ 0:68; P , 0:05), providing further evidence that the downward flow of C can be attributed to fungal activity. There was not a strong relationship between fungal biomass and the labeled C found in microaggregates (53 – 250 mm) and non-aggregated soil (, 53 mm fraction). We conclude that litter-associated fungi carry out simultaneous, bidirectional translocation of soil-derived N and litter-derived C, and in so doing, transport a significant quantity of C belowground. Most of this C is preferentially located in soil macroaggregates, where we hypothesize, it is physically protected from decay and subsequently stabilized as soil organic matter. Fungal-mediated litterto-soil C transfer has not previously been quantified in terrestrial ecosystems. If this process is widespread, it could represent a potentially important mechanism of soil C sequestration.
Acknowledgements This work was supported by grants from the Ohio Agricultural Research and Development Center of the Ohio State University and the US Department of Agriculture. Sylvie Recous provided the 13C-labeled wheat. We thank Tanna Holtrey and Rodney Simpson for technical assistance. Comments by two anonymous reviewers significantly improved the quality of the manuscript.
References Beare, M.H., Neely, C.L., Coleman, D.C., Hargrove, W.L., 1991. Characterization of a substrate-induced respiration method for measuring fungal, bacterial and total microbial biomass on plant residues. Agriculture, Ecosystems and Environment 34, 65– 73. Beare, M.H., Cabrera, M.L., Hendrix, P.F., Coleman, D.C., 1994. Aggregate-protected and unprotected organic matter pools in conventional tillage and no-tillage soils. Soil Science Society of America Journal 58, 787–795. Connolly, J.H., Jellison, J., 1997. Two-way translocation of cations by the brown rot fungus Gloeophyllum trabeum. International Biodeterioration and Biodegradation 39, 181 –188. Elliott, E.T., 1986. Aggregate structure and C, N and P in native and cultivated soils. Soil Science Society of America Journal 50, 627– 633. Frey, S.D., Elliott, E.T., Paustian, K., Peterson, G.A., 2000. Fungal translocation as a mechanism of exogenous nitrogen inputs to decomposing surface residues in a no-tillage agroecosystem. Soil Biology & Biochemistry 32, 689– 698. Halvorson, A.D., Vigil, M.F., Peterson, G.A., Elliott, E.T., 1997. Longterm tillage and crop residue management study at Akron, Colorado. In: Paul, E.A., Elliott, E.T., Paustian, K., Cole, C.V. (Eds.), Soil Organic Matter in Temperate Agroecosystems: Long-Term Experiments in North America, CRC Press, Boca Raton, FL, pp. 361–370.
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S.D. Frey et al. / Soil Biology & Biochemistry 35 (2003) 1001–1004
Lindahl, B., Finlay, Rl., Olsson, S., 2001. Simultaneous, bidirectional translocation of 32P and 33P between wood blocks connected by mycelial cords of Hypholoma fasciculare. New Phytologist 150, 189– 194. Six, J., Elliott, E.T., Paustian, K., 2000. Soil macroaggregate turnover and microaggregate formation: a mechanism for carbon sequestration under no-tillage agriculture. Soil Biology & Biochemistry 32, 2099–2103.
Tisdall, J.M., Oades, J.M., 1982. Organic matter and water-stable aggregates in soils. Journal of Soil Science 33, 141–163. Wells, J.M., Boddy, L., 1995a. Phosphorus translocation by saprotrophic basidiomycete mycelial cord systems on the floor of a mixed deciduous woodland. Mycological Research 99, 977 –980. Wells, J.M., Boddy, L., 1995b. Translocation of soil-derived phosphorus in mycelial cord systems in relation to inoculum resource size. FEMS Microbiology Ecology 17, 67 –75.