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Keywords:

  • arbuscular mycorrhiza;
  • elevated carbon dioxide;
  • root length;
  • root production;
  • root loss;
  • turnover

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  • • 
    Root responses to elevated CO2 concentrations, where nutrient demand was expected to be higher than at ambient CO2, and possible interactions with mycorrhizal symbionts are reported for pea (Pisum sativum). These are important below-ground components affecting carbon flow into the soil.
  • • 
    A video-minirhizotron system was used to study root growth in pot-grown mycorrhizal (inoculated with Glomus caledonium) and nonmycorrhizal pea plants at ambient or elevated CO2 concentrations over 9 wk. Analyses were made of root length changes, cohort size and survivorship.
  • • 
    Root length production at ambient, but not at elevated CO2, was higher in nonmycorrhizal than in mycorrhizal plants from week 4–7. Root loss began at week 5, peaking 2 wk later with 40–50% loss of the root length produced by week 8. The decline in root production and increase in root loss coincided with the onset of flowering.
  • • 
    Neither mycorrhizal inoculation nor CO2 concentration has a strong effect on pea root production and root loss, although mycorrhizal infection has a greater effect than CO2.

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Plant and ecosystem responses to increasing atmospheric CO2 are directly influenced by the size and activity of below-ground sinks (Curtis et al., 1990) and the magnitude of terrestrial ecosystem carbon storage may be determined by below-ground processes. Roots and their symbionts (mycorrhizas, nodulating bacteria) are responsible for most nutrient uptake but are also major sinks for plant photosynthates during vegetative growth (Paul & Kucey, 1981; Harris & Paul, 1987). Carbon enters the soil as a consequence of growth, activity and death of roots and root symbionts. The presence of nodulating bacteria and mycorrhiza can increase C allocation below-ground and use up to 10–20% of the fixed C (Harris et al., 1985; Jakobsen & Rosendahl, 1990; Peng et al., 1993).

Mycorrhizas usually promote plant growth by improving plant uptake of P and other immobile nutrients (Smith & Read, 1997). However, mycorrhizas may also alter the morphology, size and longevity of root systems (Eissenstat et al., 2000). Mycorrhizas increased root branching in the grass species Andropogon gerardii (Daniels Hetrick et al., 1988) and in Populus (Hooker et al., 1992), which had longer and more ramified roots than nonmycorrhizal controls, but decreased root branching and specific root length of cotton, Gossypium hirsutum (Price et al., 1989). Root systems of mycorrhizal leek, Allium porrum (Berta et al., 1990) and mycorrhizal grapevine, Vitis vinifera (Schellenbaum et al., 1991), formed more, but shorter and more profusely ramified roots than nonmycorrhizal plants. The size of root systems of mycorrhizal vs nonmycorrhizal plants is variable and this is due, at least in part, to differences in the effectiveness of the colonizing arbuscular mycorrhizal fungi and in soil fertilitity (Graham et al., 1996).

It is generally accepted that ectomycorrhizas increase root longevity (Vogt & Bloomfield, 1991) since increased longevity is consistent with an increased efficiency of nutrient acquisition (Eissenstat & Yanai, 1997). The effect of arbuscular mycorrhizas on root production and mortality, however, has only been studied in poplar trees (Hooker et al., 1995), where root longevity was actually decreased by mycorrhizal colonization. Root production and root mortality have been well studied, in particular since rhizotron and video-recording technology became available (Fitter et al., 1997). There are, nevertheless, no published reports on the effect of mycorrhiza on root production and loss of annuals or on how this might be affected by global climate change.

There are conflicting results on the effect of elevated CO2 on arbuscular mycorrhizas and some variation in magnitude and direction of the response of different arbuscular mycorrhizal fungi to consider (Klironomos et al., 1998; Fitter et al., 2000). However, studies in which changes in plant size are accounted for have not shown an effect of elevated CO2 on intra- and extra-radical mycorrhizal colonization (Staddon & Fitter, 1998; Staddon et al., 1999). Results on the effect of elevated CO2 on root production and mortality are also variable, but most of them indicate increased root production and mortality under elevated CO2 (Rogers et al., 1994; Pregitzer et al., 1995; Fitter et al., 1996).

We conducted an experiment with pea plants to study root responses to elevated CO2 and their possible interactions with the presence of mycorrhizal symbionts in the roots. We hypothesized that nonmycorrhizal controls would produce more roots of smaller diameter and shorter life compared with roots of mycorrhizal plants thereby improving soil exploration for nutrients. We expected the differences between nonmycorrhizal and mycorrhizal plants to be larger at elevated than at ambient CO2 because mycorrhizal plants would overcome an increased nutrient demand at elevated CO2 by enhancing nutrient uptake by mycorrhizas rather than by altering their root system.

We present here the effects of atmospheric CO2 and mycorrhizal inoculation on weekly root production and root loss followed with a video-minirhizotron system. Results from the same experiment concerning the evaluation of mycorrhizal colonization, plant biomass production and allocation, and nutrient uptake are published in a separate paper (Gavito et al., 2000).

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Soil

The soil we used was collected from the arable layer of an organic cropping site in Denmark. This soil (49.9% sand, 31.8% silt, 16% clay, 1.36% O. M.) had 0.27 meq Mg l−1, 0.06 meq Na l−1, 10.98 meq Ca l−1, 0.27 meq K l−1, 11.58 meq CEC l−1, 0.14% total N and 27 mg kg−1 NaHCO3-extractable P. Soil was air-dried, sieved through an 8 mm mesh and mixed with quartz sand 1 : 1 w/w. This resulted in a reduction of plant available P to 18 mg kg−1. The soil was then irradiated to eliminate mycorrhizal propagules (10 kGy, 10 MeV electron beam), and mixed with the following nutrients as powder (mg kg−1): KCl (100), MgSO4·7H2O (100), MnSO4·H2O (10), CuSO4·5H2O (5), ZnSO4·7H2O (5), CoSO4·7H2O (1), Na MoO4·2H2O (0.5) and KH2PO4 (131.7). An initial application of 20 mg N kg−1 was also added as NH4NO3. The pots were PVC cylinders (20 cm diameter and 50 cm depth), sealed at the bottom. A minirhizotron butyrate-plastic tube was placed diagonally in the pot and the pots filled with 14 kg of the soil : sand mixture with or without mycorrhizal inoculum. The tube was scribed with 1 cm2 quadrats numbered from soil surface to pot bottom.

Biological materials

Mycorrhizal inoculum consisting of soil with mycelium, spores and root pieces colonized with Glomus caledonium (Nicol. and Gerd.) Trappe and Gerdemann BEG 15 isolate was incorporated to the mycorrhizal treatments, 1 : 9 w/w, mixing thoroughly with the soil : sand mixture. One d after planting we added a filtrate obtained from the mycorrhizal inoculum to all pots in an attempt to restore some of the soil free-living microorganisms. The filtrate was cleansed of mycorrhizal propagules by passing a 1 : 9 v/v inoculum : water suspension through three overlapped layers of 25 µm nylon mesh. Pea (Pisum sativum cv. Solara) seeds weighing between 0.27 and 0.32 g were germinated between wet tissue paper and planted 2 d later. Five ml of a yeast–mannitol broth culture of Rhizobium leguminosarum (Risø 18a isolate) were placed with each of the seeds planted to promote the formation of nodules. Plants were thinned to five plants per pot after seedling emergence. The inoculation treatment was effective in establishing differences in mycorrhizal colonization. Roots inoculated with G. caledonium were more than 50% colonized 5 wk after planting at both CO2 concentrations (Gavito et al., 2000). Uninoculated plants had no mycorrhizas until 5 wk after planting. Some pots became contaminated with another mycorrhizal fungus (‘fine endophyte’) towards the end of the experiment, but colonization in contaminated pots was very low, and the only control pot where contamination exceeded 5% colonized root length was eliminated from the data analysis. Roots of all plants were extensively nodulated.

Experimental design

A complete randomized block factorial experiment was set up allocating half of the pots to a growth room with ambient atmospheric CO2 and the other half to elevated CO2. Half of the plants in each room were then inoculated with a mycorrhizal fungus (M) and half were left uninoculated (NM). Each treatment combination had six replicates and is referred to as follows: AM (ambient CO2, mycorrhizal), ANM (ambient CO2, nonmycorrhizal), EM (elevated CO2, mycorrhizal) and ENM (elevated CO2, nonmycorrhizal).

Growing conditions

Pots were watered, by weight to measured field capacity, for the first 2 wk and then the water supply was increased by 1% of the gravimetric water content every subsequent week to ensure adequate moisture in the soil column as plants increased water up take. The pots were placed inside air-cooled chambers where soil temperature in the pots could be controlled separately from air temperature. We sought to maintain soil temperature in the middle of the pots at approx. 15°C, which is an average soil temperature at 25 cm depth for the corresponding growth period in Denmark (Jakobsen & Nielsen, 1983; B. Jensen, pers. comm.). Actual soil temperatures were 14°C constant at the bottom of the pots, 15–17°C at 20 cm, and 15–20°C towards the soil surface.

Pots were not rotated within the soil-cooling chambers but each chamber, holding 12 pots, was divided into six blocks to account for position effects. The soil-cooling chambers were placed in two growth rooms at the Risø Experimental Risk Assessment Facility (RERAF, Risø National Laboratory, Denmark) set at ambient (368 ppm mean concentration across the experiment) and elevated (688 ppm mean concentration across the experiment) atmospheric CO2. Each growth room was maintained at either ambient or elevated atmospheric CO2 for 1 wk period and then the CO2 concentration was switched every wk. The plants within the soil-cooling chambers were moved to the appropriate room to maintain the CO2 treatments and rotated within the space available in the rooms to minimize differences due to position within a growth room.

Plants were grown for 9 wk with gradual increases in air temperature typical of field conditions; 14 : 8°C day : night for the first 3 wk and 16 : 10°C day : night during the last 6 wk. Day length was also increased from 14 h during the first 3 wk to 16 h during the rest of the experiment. The selected air temperature and photoperiod regimes reflect average air temperatures and day lengths for a typical pea growth period in Denmark. Photosynthetic photon flux density in the rooms increased gradually to, and decreased gradually from, a midday maximum of 750 ± 50 µmol m−2 s−1, with a 6 h period at maximum light intensity during the first 3 wk which was increased to 8 h during the rest of the experiment.

Video recording and image processing

Root video images from the marked quadrats were taken every week using a mini-rhizotron video camera (BTC-1.125, Bartz Technology Co., Santa Barbara, CA, USA) inserted into the rhizotron tubes. Root images were digitized and analysed using the software program RooTracker (The Duke University Phytotron, Durham NC, USA). Measurements of root length and root diameter were obtained from the images in each quadrat, on each sampling date. These were used to calculate cumulative and weekly root production, cumulative and weekly root loss, standing root length (root production minus root loss), mean root diameter of standing root length at each week, and the number of root segments born in three selected cohorts and the survivorship of one selected cohort. Root production was defined as the appearance of new root segments or any increase in root length of previously measured segments. Root loss was defined as the entire or partial disappearance of root segments from the quadrat. Disappearance was used as a criterion for root loss instead of changes in appearance because both dark and light root segments were observed in newly produced roots. A cohort was defined as the number of root segments that appeared for the first time on a selected wk (therefore not including growth of segments previously recorded). The cohort of root segments born on wk 3, the most homogeneous of early cohorts, was followed from birth to the end of the experiment in order to estimate survivorship (fraction of the cohort still alive at each subsequent week). For cohort analysis, the number of discrete root segments born and lost at each date was used rather than changes in absolute root length.

Statistical analysis

Effects of atmospheric CO2 and mycorrhizal inoculation on root length variables and the number of segments born in a cohort were tested by analysis of variance (ANOVA). Repeated measures ANOVA and ANOVAs for each measuring date were carried out to explore the significance of main factor effects and factor interactions. When the interaction was significant and we could identify a trend in the data, treatment means were compared with paired t-tests. If an isolated interaction became significant but without any obvious trend or meaning in the results, it was not pursued further. Data sets not meeting assumptions for ANOVA were transformed as required, but the results are presented in their original scale of measurement. Unless otherwise stated, we considered marginally significant differences at P < 0.1 and significant differences at P < 0.05. Survivorship of the cohort born on wk 3 was analysed with Savage and Wilcoxon rank tests and likelihood ratio tests for right-censored data (Pyke & Thompson, 1986) using the LIFETEST procedure in SAS 6.12, alpha = 0.05.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Standing root length

There were no significant differences between treatments in standing root length (net accumulation of roots at a given time as a result of root production minus root loss) from weeks 1 to 3 (Fig. 1). There was a CO2-inoculation interaction at week 4 (P = 0.04), 5 (P = 0.07), 6 (P = 0.05) and 7 (P = 0.09) showing higher accumulation of roots without mycorrhiza than roots with mycorrhiza at ambient but no differences at elevated CO2. The nonmycorrhizal–ambient CO2 treatment had, therefore, higher standing root length than the other three treatment combinations, which had similar standing root length, across the measuring period (P < 0.0001). Standing root length increased steadily in all treatments during the first wk of the experiment, reached its maximum between wk 5 and 6, and declined at wk 7. There were no significant differences between treatments in mean root diameter at any sampling time (data not shown).

image

Figure 1. Standing root length of Pisum sativum (root production minus root loss = total root length measured at a given time) per minirhizotron tube at weekly intervals. Closed columns, mycorrhizal treatments; open columns, nonmycorrhizal treatments. A, ambient CO2, E, elevated CO2. Error bars represent 1 SE of the mean (n = 5–6). Asterisks indicate that the treatment combination ambient CO2–nonmycorrhizal differs significantly (P < 0.05) from the other treatment combinations.

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Root production

Root production increased rapidly from planting to wk 2 and remained fairly constant in all treatments until plants flowered, in wk 6 (Fig. 2). Root production dropped markedly after flowering. There was a trend towards higher production by roots without mycorrhiza than by roots with mycorrhiza at both CO2 concentrations (Fig. 2), but variation was very high and the comparative increase in root length of nonmycorrhizal plants reached significance only at wk 5 (P = 0.07). Atmospheric CO2 had no consistent effect on root production. However, since root production accounted for almost all of the observed standing root length during the first 5 wk, standing root length was equivalent to cumulative root production during that period. Therefore, the same interaction observed in standing root length was significant for root production per wk (P = 0.056) and cumulative root production (P < 0.0001) across the experiment.

image

Figure 2. Pisum sativum root length produced per minirhizotron tube at weekly intervals. Closed columns, mycorrhizal treatments; open columns, nonmycorrhizal treatments. A, ambient CO2, E, elevated CO2. Error bars represent 1 SE of the mean (n = 5–6). Asterisks indicate that the treatment combination ambient CO2–nonmycorrhizal differs significantly (P < 0.05) from the other treatment combinations.

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Root loss

There was no root loss observed during the first 4 wk (Fig. 3). Root loss started at wk 5, increased markedly at flowering and decreased at wk 8, indicating a very well defined period of root shedding. There were no differences among treatments in root loss per wk until wk 6. We observed higher loss of nonmycorrhizal roots than mycorrhizal roots at wk 7 (P = 0.07) and wk 8 (P = 0.04), but at wk 8 an interaction (P = 0.07) revealed that the effect of mycorrhiza occurred only at ambient CO2. These results indicated that root length loss was becoming proportional to root length production. Neither mycorrhizal inoculation nor elevated atmospheric CO2 concentration had a strong effect on root loss at the time we finished our measurements. A ratio of cumulative root loss to cumulative root production showed that by the end of the expt 64 ± 5% (mean ± SE) of the entire root length production was still alive in treatment AM, 49 ± 4% was alive in treatment ANM, 58 ± 6% was alive in treatment EM and 57 ± 6% was alive in treatment ENM (Fig. 4). There were no significant main effects or interaction in root loss ratio but the means for inoculation treatments were significantly different (P = 0.049) at ambient CO2.

image

Figure 3. Root length loss of Pisum sativum per minirhizotron tube at weekly intervals. Closed columns, mycorrhizal treatments; open columns, nonmycorrhizal treatments. A, ambient CO2, E, elevated CO2. Error bars represent 1 SE of the mean (n = 5–6).

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image

Figure 4. Number of root segments born in each cohort of Pisum sativum and the mean of the root segments born in all three cohorts. Closed columns, mycorrhizal treatments; open columns, nonmycorrhizal treatments. A, ambient CO2, E, elevated CO2. Error bars represent 1 SE of the mean (n = 5–6 for single cohort, n = 15–18 for mean of three cohorts).

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Cohort analysis

There were no significant differences between treatments in the number of root segments born on wk 1 and wk 3, but there was a nonsignificant trend for an increased number of root births in nonmycorrhizal treatments at both CO2 levels at wk 3 (Fig. 5). This trend became marginally significant (P = 0.059) at wk 5. We selected the cohort of roots born on wk 3, which was the most homogeneous of early cohorts, and followed its survival through wk 8. Loss of root segments from the cohort was highly variable and there were no clear treatment effects. Differences in survivorship did not reach statistical significance in any of the tests performed.

image

Figure 5. Pisum sativum root loss ratio (cumulative root loss/cumulative root production) at harvest, is used as an indicator of root longevity in our treatments. Closed columns, mycorrhizal treatments; open columns, nonmycorrhizal treatments. Error bars represent 1 SE of the mean. (n = 5–6).

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Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

A limited number of the previous studies on CO2 effects on root production and root loss have dealt with annuals, and the present work is the first to our knowledge to include mycorrhizas as a treatment factor in such a study. Our prediction of higher root production and faster root loss in nonmycorrhizal than in mycorrhizal plants was in general supported by the results. Contrary to our expectations, these differences were significant and clear only at ambient CO2. Our prediction of a CO2–mycorrhiza interaction was based, however, on the premise of increased nutrient demand at elevated CO2 presumably leading to nutrient limitation.

It is important to consider that our experiment was carried out with a nodulated legume that was given additional N and P and grown under favourable conditions to minimize nutritional or other effects on plant and mycorrhiza development. Mycorrhizal colonization was similar at both CO2 levels (Gavito et al., 2000) and there were no differences in root mass, root length density, intraradical or extra-radical mycorrhizal colonization of ambient and elevated CO2 plants that may have indicated confounding changes in total mycorrhizal colonization or fungal biomass. Our observations of biomass production and nutrient uptake in this experiment (Gavito et al., 2000) suggested that there was little persistent nutrient limitation. Assuming that there was no nutrient limitation, it is difficult to explain why mycorrhizal inoculation affected root production and root loss only at ambient CO2, a result that has been confirmed in later experiments (M. E. Gavito & I. Jakobsen, unpublished).

Our only possible explanation is based on the use of plant C by mycorrhizas. Mycorrhizal fungi are obligate symbionts and depend on plant C but would not respond to an increase in C supply unless this resource was limiting. Photosynthate use by mycorrhizal symbionts does not seem to be controlled by the plants (Fitter et al., 2000) and can be high enough to become detrimental as illustrated from reports of growth depressions caused by inoculation with mycorrhizal fungi (Eissenstat et al., 1993; Peng et al., 1993). We believe that mycorrhizal fungi are not C limited and, with adequate nutrients and at elevated atmospheric CO2, they might at some point respond to an increased availability of root space to colonize, not to an increased C supply to the roots. If so, mycorrhizal fungi would use the same amount of C at both atmospheric CO2 levels, at least during the vegetative stage where fungal growth is usually not ‘root space limited’. Without nutrient limitation but limited C supply for the plant (ambient CO2), shoot C in mycorrhizal plants would be partitioned to mycorrhizal fungi at the expense of root development whereas in nonmycorrhizal plants it would be used entirely for root development. At elevated CO2, if a larger amount of photosynthate was sent below-ground, the additional C could be used for root growth in mycorrhizal plants thereby resulting in a more similar root production in nonmycorrhizal and mycorrhizal plants. This is, at present, still highly speculative because most work on the regulation of the mycorrhizal symbiosis has been centred on the plant side and plant-limiting resources such as P (Fitter et al., 2000).

We found that vegetative growth was accompanied by increasing root production, and the onset of flowering marked a drastic reduction in root production and a period of high root loss. Such a clear period of root shedding implies that large amounts of C and nutrients enter the soil in the form of dead roots at flowering, when between 40 and 50% of the root system dies. At ambient CO2, root length just before flowering was approx. 30% lower in mycorrhizal than in nonmycorrhizal plants. Two wk after flowering root length was reduced by 24% in mycorrhizal plants and by 43% in nonmycorrhizal plants compared to root length measured before flowering. Jakobsen (1986) and Jakobsen & Nielsen (1983) used a soil-core method to measure changes in root length of ambient CO2 field-grown peas in similar soil and temperature conditions as our study. They found that nonmycorrhizal plants produced more roots of smaller diameter and shorter life than mycorrhizal plants and that there were two clear periods of root production and loss. One period occurred before flowering and one occurred after flowering, indicating some replacement or turnover of roots at pod filling. In their study, root length before flowering was 30% lower in untreated P-fertilized soil than in fumigated soil (to eliminate mycorrhizas) and after flowering root length was reduced by 23% in untreated soil and by 33% in fumigated soil (Jakobsen, 1986). This indicates higher loss of roots without mycorrhiza. Our results at ambient CO2 are, therefore, in general agreement with previous observations of field-grown peas but differ from those of Hooker et al. (1995) and Hooker & Atkinson (1996) where arbuscular mycorrhizas decreased root longevity in the woody perennial Populus generosa inter americana. Cohort survivorship showed that between 40 and 50% of the root segments born had disappeared by week 8, but with no obvious treatment differences. Roots of annual crops seem to live for 1 or 2 months and to have a shorter life than perennials, based on few studies conducted to date (Eissenstat & Yanai, 1997). Since our criterion for root death was disappearance, and not change of colour or appearance, as in most root mortality studies, it is possible that we underestimated root loss (Comas et al., 2000).

The root systems of mycorrhizal and nonmycorrhizal plants seem to be similar under nonlimiting nutrient supply and to differ progressively as nutrient availability decreases or increases from the optimum (Daniels Hetrick et al., 1988; Amijee et al., 1989). If we assume that there was no nutrient limitation in our experiment, the differences in root length should be attributed to size or morphology changes in the root system induced by the presence of mycorrhiza independently from its effects on nutrient uptake. Hooker et al., 1992) reported such mycorrhiza-induced changes unrelated to plant nutrition in root morphology and architecture of Populus. We found no mycorrhiza effects on mean root diameter suggesting changes in root morphology or architecture. We are aware of no other published reports of mycorrhiza effects on root diameter, production and loss in annual plants with which to compare our findings. Altogether, our results indicated that under controlled growth conditions of nonlimiting nutrient supply, root production and root loss of pea plants was affected only slightly by the presence of mycorrhiza but mycorrhizal inoculation effects on root architecture could not be ruled out.

Elevated CO2 had almost no effect on standing root length or root production in our experiment, which is contrary to our expectations but in agreement with the results of van Vuuren et al. (1997) from a study with spring wheat and other results discussed by Arnone et al. (2000). The lack of response to atmospheric CO2 is consistent with the results discussed above if plants were not nutrient-limited. Root loss was also unaffected by elevated CO2, in accordance with other studies showing that elevated CO2 does not affect the relationship between root production and root loss (Pregitzer et al., 1995; Berntson & Bazzaz, 1996). That is, root production, root loss and root longevity (turnover) are affected equally by elevated CO2, although CO2 effects are likely to be different in annual and perennial plants (Pritchard & Rogers, 2000).

Overall, our results indicated modest effects of atmospheric CO2 and mycorrhizal inoculation on root production or root loss in pea. However, it is clearly premature to draw general conclusions from a single study. Further experimentation will be needed if we are to make accurate predictions regarding the effects of atmospheric and climatic change on root production and root loss, two crucially important processes affecting the amount of C entering the soil.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The authors wish to thank Anne Olsen and Anette Olsen for skilful technical assistance, Henrik Jørgensen and Erling Johannsen for assistance on video and computing equipment. This work was supported by grant no. 9501743 of the Danish Agricultural and Veterinary Research Council. Partial funding was provided by the National Institute for Global Environmental Change through the US Department of Energy (Co-operative Agreement No. DE-FC03–90ER61010). Any opinions, findings, and conclusions or recommendations expressed in this publication are those of the authors and do not necessarily reflect the views of the DOE.

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  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
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