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Microbial nitrogen transformations in the root environment of barley
soil Bid. &chem. Vol. 19, No. 5, pp. W-558, Printed in Great Britain. All rights rcKlvcd
~38~717/87
1987
S3.00 + 0.00
Copyright Q 1987 Fcrgamon JoumaIs L&i
MICROBIAL NITROGEN TRANSFORMATIONS ROOT ENVIRONMENT OF BARLEY
IN THE
LEIF KLEMEDTSON,*PER BERG, MARIANNEGARHOLM, JOHANSCHN@RJZR Department of Microbiology, Swedish University of Agricultural Sciences, S-75007 Uppsala, Sweden THOMASROSSWALL
Department of Water in Environment and Society, University of LinkGping. S-58183 Linkiiping, Sweden (Accrpred 4 February1987) !Smnmary-To determine the influence of barley roots on microorganisms and N-transforming processes in soil, numbers of nitrificrs and potential nitrification and denitrification rates were measured every week for 5 wks. The barley plants were grown in growth chambers in which the root-containing soil layer (A) was separated from three outer soil layers (B, C, D). The numbers and biomass of bacteria, numbers of flagellates and amoebae, total and FDA-active hyphal lengths. microbial biomass carbon and respiration
were also determined. The numbers of ammonium oxidizers were positively correlated with root biomass but did not differ significantly between soil layers. Potential ammonium oxidation was stimulated in the ioot-layer, while potential nitrite oxidation was stimulated in the B- and C-layers. The denitrification activity (measured anaerobically in the presence of excess NO;) was positively con-elated with root biomass in the A-layer. Denitritication activity in the B-layer was positively correfated with the water content of the soil. When roots grew near the nets separating the root layer from the other layers, denitrification activity was stimulated in the next layer (B). We propose that nitrite oxidation in the root zone partly depends on the reduction of nitrate. This would explain why nitrite-oxidizer numbers were usually several orders of magnitude higher than ammoniumoxidizernumbers. Bacterial numbers decreased between wks 1 and. 5. Increases in bacteria, naked amoebae and flagellates in all layers between wks 2 and 3 indicated that bacteria were produced until wk 3. There were no signs of bacterial production after wk 3. The total length of hyphae and the length of FDA-active hyphae were not significantly different between layers. However. both of these parameters, as well as total microbial biomass carbon and respiration. were consistently highest in the A-layer.
Microorganisms in soil are affected by plant roots in several ways. For example, root exudates can stimulate heterotrophic growth and lead to local competition for inorganic nutrients between roots and microorganisms. Respiration by roots and Oz consumption by heterotrophic organisms decrease O2 concentrations close to actively growing roots, thereby also affecting the microenvironment. However, 0, diffusion into the soil is also stimulate by roots, since their uptake of water reduces a major barrier of gas diffusion in the soil. The decomposition of dead roots creates air channels, which also facilitate O2 diffusion. The rhizosphere is an illdefined zone a few miltimetres thick that surrounds plant roots. It is a soil volume characterized by the increased activities of fungi but especially bacteria (Rovira and Davey, 1974; Woldendo~, 1981) and protozoa (Darbysbire and Greaves 1973). Tbe limits of the rhizosphere are difficult to establish, and it is probabfe that there are two spheres of influence around roots: (i) a smaller one where enrichment of organic compounds occurs *To whom all correspondence should be addressed, 551
and (ii) a larger one where the roots affect the partial pressures of 0,. CO2 and other soil gases. This larger sphere may be particularly important in regulating the activity of nitrification and denitriiication, which is influenced by the aerobic-anaerobic state of the rhizospheric environment (Woldendorp, 1963; Klemedtsson et al., 1987). Since bacteria depend on easily-available organic compounds for energy (Stotzky and Norman, 1%3), most bacterial activity in soil should thus be linked to either roots or to fresh organic material undergoing decomposition. By far the largest proportion of bacteria in other regions of the soil is dormant (Lynch, 1984). Protozoa are likewise more numerous close to roots, and there is evidence that by feeding on bacteria they cause a local increase in nitrogen mineralization (Clarholm, 1985). In the bulk soil, the biomass of fungi is normally larger than that of bacteria, but either group can dominate in the rhizosphere (Newman, 1985). Ammonium-oxidizers compete with roots and heterotrophic microorganisms for available ammonium. Inhibiting effects of roots on nitrification have been discussed by several authors (Rice and Pancholy, 1972; Woldendorp, 1975; Wheeler and Donaldson, 1983) and reviewed by Rice (1984). Favourable con-
LEIFKLEKEDTSSONet
552
(II
ditions for nitrification do not seem to occur in the rhizosphere (Woldendorp, 1981). since higher concentrations of energy-rich substances will favour hetammonium-immobilizing organisms. erotrophic, Stimulating effects of roots on nitrification have, however, been reported for arable soils (Molina and Rovira, 1963; Berg and Rosswall, 1987). Denitrification activity may be stimulated (Woldendotp, 1963; Stefanson, 1972a-c; Void et al., 1976; Klemedtsson et al., 1987) or inhibited (Smith and Tiedje, 1979) by roots. Inhibition is probably due to competition between denitrifiers and plants for NO; (Smith and Tiedje, 1979). Our aim was to study the influence of roots on the amounts and activities of soil microorganisms at delined distances from plant roots. Of particular interest was whether nitrification and denitrification were stimulated or inhibited by the presence of roots. MATERIALS AND METHODS
Soil characteristics
Topsoil (0- 10 cm) was taken at Kjettslinge, 40 km north of Uppsala in central Sweden, at the site of the Ecology of Arable Lund project. The plow layer (O-27 cm) of the soil was a loam (clay content, 19%; pH 6.3; organic C content, 2.2%; total N content 0.23%) cultivated with unfertilized barley (Ho&urn dktichum L.) (Steen ef al., 1984). The soil was dried, sieved (c 2 mm), mixed and rewetted to a water potential of - 165 kPa (60% of the water holding capacity, WHC). The soil was held at 60% WHC for I wk before starting the experiment to eliminate the easily-available organic carbon produced during the drying and rewetting of the soil. Experimental
system and sampling plan
The soil was packed in growth chambers (Fig. 1) constructed from sheets and edgings of Plexiglas (Helal and Sauerbeck, 1981). Nylon mesh nets were used to establish four compartments at various distances from the roots with four soil layers on each side of the roots of the barley seedlings (Fig. 1). The thickness of the soil layers were: A 20 mm (300 g soil wet wt at 60% WHC); B 4mm (55 g); C 6mm (100 g); D IO mm (150 g). The nets delimiting the soil closest to the roots (A-layer) were of 30 pm mesh to confine all roots to this layer. The other nets had a mesh-size of 100 pm. The pots were kept moist for 4 days, after which two barley seedlings (Ho&urn distichum L. cv. Gunilla) were planted in the A-layer of each pot. The pots (4 replicates x 5 sampling occasions = 20 pots) were kept in a greenhouse at a temperature of 25°C and at a light intensity of 100 W m-*. Four replicates were sampled every week for 5 wk. The soil was kept at 90% WHC during the first 3 wk by placing the pots on a watered sand bed. In addition, the pots were watered from the top every second day. Thus water potentials varied depending on evapotranspiration and watering. At the third sampling the barley plants showed signs of nutrient deficiency, hence a nutrient solution, containing mineral salts but no nitrogen, was added (Jensen, 1942). For 3 days between the third and fourth samplings the temperature control of the greenhouse failed, causing a rise in temperature from 25 to 37°C. As a
2cm I
4
Fig. I. The growth chamber. Thickness of soil layers: A 20 mm, B 4 mm, C 6 mm, D 10nun. The outer dimensions were: 200 x 120 x 65 mm.
consequence, the soil dried out and remained dry (especially in the A-layer) for several days. Sampling
One day before each of the first three samplings the soil was moistened to near water-holding capacity. However, the soil proved to be too wet for efficient use of the fumigation method for microbial biomass determination; thus the water content was reduced after 3 wk to 70-80% WHC. To sample soil, the sheets of Plexiglass were carefully removed and the soil from each layer was collected. Soil from one half of the growth chambers was used to measure denitrification activities. The other half was used for estimating the abundance of bacteria, protozoa and fungi as well as the number of nitrifying bacteria and their potential activity. The A-layer was divided down the middle to form two subsamples, each containing one barley plant with intact roots. Root biomass,
soil moisture
and inorganic nitrogen
Roots were inspected visually at each sampling. Root biomass was determined by washing the roots with tap water over a sieve (0.5 mm mesh) to remove soil particles. The weight of the roots was determined after drying at 85°C overnight. The soil water con-
Microbial N transformations
tents were determined gravimetrically after drying at 10X overnight. Inorganic nitrogen was extracted with 2 M KC1 (10 g wet soil in 50 ml 2 M KCI) on a rotary shaker for 2 h. NO; and NO;- were analyzed by flow injection analysis (FIA; Bifok FIA 05 and Bifok FIA 06 Flow-Photometer, Tecator, Hogan&, Sweden; Ruxicka and Hansen, 1975). Ammonium was assayed by the indophenol-blue method (Keeney and Nelson, 1982). Nitrifer
numbers
The method of soil extraction and microtiter plate
technique used for most-probable-number (MPN) enumeration of ammonium oxidizers have been described by Berg and Rosswall (1985); but the nitrite oxidizer medium was prepared according to Schmidt and Belser (1982). The plates were incubated at 24°C for 8 wks, after which the presence of ammonium oxidizers was determined using a pH-indicator (Sarathchandra, 1979). The rate of nitrite disappearance was used to estimate the abundance of nitrite oxidizers using Griess-Illosvays’ reagents. Potential ammonium- and nitrite-oxidation
rates
in the root environment
553
Bacteria, protozoa, fungi and microbial biomass carbon
Bacteria were counted in a&dine orange-stained soil smears under fluorescent light at 1000x magnification. To estimate biomass, the bacteria were grouped into six size classes with nominal volume values (Clarholm and Rosswall, 1980). A density of l.Igcm-‘andadrywtof30%wereassumedforthe biomass calculations (Bakken and Olsen, 1983). Protozoa were enumerated by a modified MPN technique (Clarholm, 1981). Total hyphal length was estimated by the Jones and Mollison (1948) agar 8lm technique as modified by Schniirer et al. (1985). FDA-active hyphal lengths were determined according to Soderstriim (1977) as modified by Schnilrer et al. (1985). The carbon content of the microbial biomass was determined by the chloroform fumigation technique (Jenkinson and Powlson, 1976) with modifications (Schniirer et al., 1985). Statistical analyses. All statistical analyses conceming the rates of denitrification potential were performed using the Statistical Analysis System (SAS, SAS Institute, Gary, NC U.S.A.) version 4.10. All regressions and analyses of variance were performed using the General Linear Models (GLM) procedure within SAS. All other analyses were made using standard textbook procedures.
Ammonium oxidation was estimated as described by Berg and Rosswall (1985), and was a modification of the method used by Belser and Mays (1980). Samples of soil (25 g) were mixed with medium (4 mM (NH&SO,, pH 7.2) according to Sarathchandra (1979) in capped bottles (300 ml). Chlorate was added to give a final concentration of 15 mM in the soil slurry. The bottles were held on a rotary shaker (200 rev min-‘) at 24°C. Samples (2 ml) were removed from the soil slurry 0, 3 and 6 h after chlorate addition and put in test tubes with 2 ml of 4 M KC1 to inhibit ammonium oxidizer activity. The samples were then centrifuged, filtered and analyzed for accumulated nitrite. Nitrite oxidation was determined by measuring nitrite disappearance at 0, 3 and 6 h after incubating soil in a nitrite oxidizer medium (Schmidt and Belser, 1982) containing 60~~ nitrite (pH 7.2). No chlorate was added; otherwise, procedures were as described above for ammoniumoxidation.
After 1 wk. mainly fine roots were found and they had accumulated close to the nets that confined the A-layers. After 2-3 wk. suberised roots dominated. The fine roots were less abundant near the nets; instead, a concentration of fine roots occurred towards the center of the A-layers. Root biomass decreased between wks 3 and 4 (Fig. 2) due to the unintentional desiccation. At 4 wk the amount of fine roots had decreased further and some dead roots were observed. The root biomass, mainly the fine root fraction, increased again between the fourth and fifth week, after the soil moisture contents had been re-adjusted.
Potential denitrification
Inorganic nitrogen
The soil layers were carefully transferred
to aluminium containers, and KNO,-solution was added to a final concentration of 40 fig NO;-N g-’ dry wt. To determine gaseous nitrogen losses, the containers were placed in gas-tight plastic bags (Cryovac Bags, W. R. Grace Ltd, St Neots, Cambs., England) with butyl-rubber stoppers. The acetylene inhibition technique at 10kPa C,H, (Klemedtsson et al., 1977; Yoshinari et al., 1977) was used to determine denitrification rates. The plastic bags were rapidly evacuated and refilled with N, (< 5~1 Ozl-‘) five times, after which 50 ml neon and 100 ml C2H, were added. The neon was added to determine the gas volume of the bags. The amount of gas dissolved in soil water was calculated using the Bunsen coefficients given by Tiedje (1982). Gases were sample at 1,2.3,4,6 and 8 h with evaculated Venoject tubes (Terumo Europe, N.V. Leuven, Belgium) and analyxed by gas chromatography (Klemedtsson et al., 1986).
RESULTS Root growth
Nitrate-N varied between 0.5 and 2 pg N g-l dry soil. Nitrite concentrations were always below 10 ng N g-’ dry soil. Ammonium concentrations were highest in the A-layer from samplings 1 and 2 (5.2 f 0.8 pg N g-i dry soil; mean + standard error; N = a), compared with the B, C and D layers (1 .l + 0.2; N = 24). It was not possible to analyze ammonium from samplings 3, 4 and 5 due to a technical error. Numbers of nitrlpers
Numbers of ammonium-oxidizers in individual samples varied greatly with time (from 2 x 10’ to 102 x 10’ cells g-i dry soil, Fig. 2). No significant differences were found between layers. The mean numbers of ammonium-oxidizers in all four layers were correlated with the amounts of root biomass in the A-layer (r = 0.91, N = 5, P < 0.05). Numbers of nitrite-oxidizers in individual samples varied less with time (21 x 10’ to 98 x 10’ cells g-i dry soil, Fig. 2)
0.4
!
b1
.-s--
Time
twrrkr
)
Potential ammonium oxidation (a) and nitrite oxidation (b) in the four soil layers (C(g NOT-N g-’ dry soil Fig. 3.
Fig. 2. hot biomass, 11~~bers of NH;- and NO,‘-oxidizers
and potential denitrification during 5 wks with barley growing in a compartmentalized growth chamber. Bars show SE. than did the ammonium-oxidizers, and reached a maximum I wk later than the ammonium-oxidizers (Fig. 2). Potential nitri@ation
rates
Potential ammonium-oxidation rates were consistently highest in the root-layer (Fig. 3a). The rates increased between samplings 2 and 3 (Fig. 3a) 1 wk after the increase in ammonium oxidizer numbers (Fig. 2). No correlation was found between the numbers and activities of ammonium-oxidizers. Potential nitrite-oxidation rates were consistently highest in the B- and C-layers (Fig. 3b). The highest potential rates were found after 2 wk, after which they declined. The numbers and potential rates of nitrite-oxidizers were correlated in the A-layer (r = 0.44; N = 20; P < 0.05), but not in the root-free layers. DenitriJScation activities
Except for the first sampling, the highest potential deniMcation rates (PDRs) were found in the Alayer (Fig. 2). At samplings 1-3 PDRs in the A-layer were correlated with root biomass (r = 0.53; P < 0.02; n = 12). However, the A-layer PDRs were negativeiy correlated with root biomass at the fourth sampling, by which time the root biomass had decreased (r = -0.96; P -C0.04; n = 4). When the activity in the B-layer of all samples was considered, no
h-‘; A’ = 4). Bars show SE.
correIation with root biomass was found, but there was correlation with soil water content (r = 0.45; P < 0.05; n = 20). The highest PDRs were found in the B-layer after 1 wk of plant growth (Fig. 2). These PDRs were correlated with root biomass (r = 0.87; P < 0.05; n = 4). The PDRs in the C- and D-layers were not correlated either with water content or root biomass. At the first sampling, the activity of the B-layer was significantly higher than that of the other layers (P < 0.05). Except for the C-layer at sampling 4 and D-layer at sampling 5, the activities in the A-layer of samples 2-5 were all significantly higher than those in the other layers (P <0.05), Bacteria, protozoa, fmgi
and total microbial biomass
The numbers and biomass of bacteria at sampling 5 were significantly lower (P c 0.05) than those at the first sampling (Table 1). Bacterial numbers had decreased by 43-45%, resulting in 24-33% lower biomass. The bacterial and flagellate development with time was similar in all four layers. The highest values were recorded at the first sampling and the second largest values were recorded at the third (data not shown). The naked amoebae were initially present in low numbers, increased until the third samphng, and then decreased, remaining low until the end of the experiment (data not shown). At the last sampling, mean values of total and FDA-active hyphae as well as biomass-C and U&-production were highest in the A-layer (Table 2). The differences were, however, not statistically significant.
Microbial
in the root environment
N transformations
555
Table I. Numbers and biomass of bacteria and numbers of flagellates and naked amccbac at sampling wks I and 5. All values are expressed g-’ dry soil, mean f SE; n - 4. A = the middle layer. co&sing the root; B, C. D layers at dcfmed distances from the root (set Fin. II
No. of No. of bacteria Layer
Sampling week
A : D
Bacterial biomass
wg-‘)
naked amoebae (lo’g_‘)
wg-‘)
I
5
6.SkO.S 6.3 f 0.4 6.2kO.5 6.1 f 0.2
3.5kO.5 3.5 iO.5 3.5kO.3 3.3 f 0.7
(:gdwg-‘) 520*35 500 f 45 490245 520 k 25
5
I
5
,
5
3SOk40 350 f IS 37Ok40 380 f 25
35 f 6 47&17 3629 32 i 5
21*5 32 f 16 g7*22 48 * 34
66 f 22 &0*4 77* I3 a6* I3
52* I4 51 f I2 4Sf:8 3a*9
DIBCUBBION
Root growth and water conditions
Early in the experiment, most roots were found close to the nets, where physical resistance of the soil to root growth was likely to have been the lowest. After about 2 wks roots began penetrating deeper into the center of the A-layer and there was a corresponding decrease in 8ne roots near the nets. These changes probably resulted from an increasing nutrient deficiency in the soil close to the nets. Water uptake by the plants probably resulted in a net flow of water from the D-, C- and B-layers towards the A-layer. It is therefore unlikely that non-volatile root exudates diffused from the A-layer into the B-layer, because exudates would have had to diffuse against the direction of water flow. Water transport between the A- and B-layers may have been slightly restricted by the fine mesh net. Bacteria, protozoa
No. of flagellates
and figi
The numbers and biomass of bacteria at the first sampling were in the upper range of that found for the same soil in the field (Schniirer et al., 1986). This was probably an effect of the treatment of the soil before the experiment. The increase in bacteria between wks 2 and 3 coincided with an increase in the values for potential ammonium oxidation. Since the increase occurred in all layers, it was probably unrelated to root activity. The dynamics of the flagellate populations were similar to those observed for bacteria. Flagellates require less dense bacterial populations to multiply, compared with naked amoebae. Thus, the flagellate numbers might have only lagged l-2 days behind the bacteria, but this would not have been detected with the sampling interval used. The mean production of amoebae between samplings 1 and 3 was 5 x 10’ amoebae g-i dry wt soil. This is equivalent to a consumption of 2 x 10’ bacteria or 80-1OOpg C, since each new amoeba represents 3000-4000 con-
sumed bacteria (Clarholm, 1981). The carbon for bacterial growth must have been derived mainly from organic matter in the soil, since bacterial and protozoan population dynamics were similar in all four layers. The constant size of the amoeba1 population during the remainder of the experiment (samplings 4 and 5) indicated no further production of bacteria during this period. At the first sampling, hyphal lengths were within the range-although at the lower end-found in field samples of the same soil (Schniirer et al., 1986). Both hyphal lengths and total microbial biomass C at the last sampling were very close to published values for this soil (Schniirer, 1985; Schniirer et al., 1986). Although total and FDA-active hyphae. respiration and microbial biomass C were not significantly higher (P > 0.05) in the A-layer than in the outer layers, all values were numerically highest in the A-layer. The differences in total microbial biomass C between the A-layer and the other layers ranged between 68 and 78~g C g-’ dry wt. Based on the numerical relationships provided by Newman (1985). a standing crop of 175 mg of roots at sampling 5 (Fig. 2) would be equivalent to a total input of from 90 to 210 pg C g-’ soil. This carbon seemed to have been in a form more accessible to fungi than to bacteria, since bacterial biomass decreased during the experiment and FDA-active hyphae was the only biological component that increased in the A-layer between the 1st and 5th samplings. An additional C-source for this increase could have been the roots killed between the 3rd and 4th samplings. The microbial biomass in the same soil grown with barley increased at most 100 pg C g-’ dry wt over a growing season, both in field and greenhouse experiments (Schniirer, 1985; Schniirer and Rosswall, 1987). The initial microbial biomass was about 4OOpg C g-’ dry wt in both cases. Thus, the total amount of microbial biomass in this soil cannot apparently increase by more than 25% in response to carbon additions from the growing barley. This is
Table 2. Total and FDA-active hyphal lengths, microbial biomau carbon and respiration on the tint and the last sampling data and at the last C sampling date (mean values f SE. n = 4). A = the middle layer, enclosing the root: B. C. D lovers at dcfmcd distances from the root (see Fia. I)
Laver A B
Sampling week
Total hyphae (“;” g-’ dry WC) 5 o.s*o.1 0.4*0.1 0.6 f 0.2 0.7 f 0.2
0.8 * 0.2 0.7*0.1 0.4 f 0.0 0.8 f 0.1
FDA-active hThae (yedry wt) 5 Is*2 14*4 l4*6 IO f 2
2125 l7*2 l3*2 Is* I
Rapiration Biomass-C (PgCg-’ (pgCgldry wt) dry wt liday-’ 445*20 367 f 21 370 * I3 377 f I9
163 f I7 105*7 112+5 103+4
)
556
LEIF
KLEMED~SSON
consistent with current theories on a protection capacity for a given size of the microbial biomass, specific for each soil (van Veen et of., 1984). Inorganic nitrogen No inorganic N was added since we wanted to examine the effects of roots on the N-mineralization from soil organic matter. The increased amounts of ammonium in the root-containing A-layer of the two first samplings support the idea of root-mediated N-mineralization (Clarhoim, 1985), especially since potential ~onium-oxidation rates were generally highest in the root layer. A diffusion of N from the other layers could have been another possible cause, but the absence of a gradient of NH: or NO, in B-C layers makes this less probable. Since we were unable to obtain data on the NH: for the three last samplings, it was impossible to determine to what extent local mine~lization had occurred during the rest of the experiment. Numbers and activities of nitrifiers
Numbers of ammonium- and nitrite-oxidizers in the A-layer were higher than was found earlier in soil from Kjettslinge (Berg and Rosswall, 1987). This was probably a result of high ammonium concentrations produced by the rapid mineralization occurring in the dried, sieved and rewetted soil. In the field the ammonium concentrations were about l-2 pg N g-’ dry soil (P. Berg, unpublished). Since potential ammonium-oxidation rates were highest in the Alayer, the corresponding numbers of ammonium oxidizers were also expected to be highest in this layer. This may have been true but was impossible to confirm due to the large variations in the results from the MPN method. Considering the heterogeneity of the soil, the small size (1 g) of the soil sample used for MPN determinations in this study-compared with the log normally used (Berg and Rosswall, 1987)-may be why no differences were detected. Both the potential rates of ammonium-oxidation and nitrite-oxidation varied less than the MPN counts and were of the same order of magnitude found in the Kjettslinge soil (Berg and Rosswall, 1987). Since ammonium-oxidation was not inhibited by the experimental procedure, potential nitriteoxidation rates may have been underestimated, because production and oxidation of nitrite would have proceeded simultaneously. The observed decrease in nitrite must therefore represent a net disappearance. The increase in potential ammonium-oxidation rates between the 2nd and 3rd samplings occurred 1 wk after a corresponding increase in MPN numbers. This is consistent with the results of Berg and Rosswall (1985), which showed that potential ammonium-oxidation rates respond more slowly and proportionally less to short-term changes in soil than MPN numbers. These observations were also supported by the adverse effects on the number and activity of ammonium oxidizers observed after the drying-out incident. Earlier investigations found no relationship between nitrifier numbers and nitrification rates (Sarathchandra, 1978; Belser and Mays, 1980; Steele er al., 1980). We also found that there was no relationship between ammoniumoxidizer numbers and nitrification rates, although
et uf.
such a relationship has sometimes been recorded (Berg and Rosswall, 1985; Berg and Rosswall, 1987). The apparent positive effect of roots on the growth of ammonium-oxidizers, as reflected in the positive correlation between root biomass and ammoniumoxidizer numbers, may actually be coupled to the uptake of water by the roots and thereby to the oxygen status of the soil, since the correlation was found for all layers and not solely in the root layer. Due to the watering of the chambers before sampling, it was not meaningful to calculate correlations between nit~~cation rates and water contents. Several authors have proposed that plant roots could inhibit nitrification (Leather, 1911; Theron, 1951; Woldendorp, 1981). Since the water-flow was assumed to be mostly towards the A-layer, possible soluble inhibitory substances produced by the root could not have played a major role in orbiting nit~fi~tion. The present data show that ~~~~tion can be stimulated in soil within a few millimetres of roots, at least in the early stages of barley growth in arable soil. This is consistent with the results of Molina and Rovira (1963), who showed that growth of Nitrosomonas and Nitrabacter was stimulated by corn and luceme roots within 2 wk after planting. However, this growth promoting effect disappeared after S wks. Potential
denirrification
The denitrification activity in the A-layer positively associated with root biomass, was also recorded by Bailey (1976), Smith and Tiedje (1979) and Klemedtsson et al. (1987). The increase in PDRs in the A-layer, which was associated with increasing root biomass during the first 3 wk, supported the results of Bailey (1976), who studied the effects of roots on denitrification and measured rates in a similar way. Although the denitrification activity in the B-layer was higher than in the A-Iayer at the first sampling, it should be noted that only a fraction of the soil in the A-layer was affected by roots, since these were concentrated in the soil near the smaller B-layer. As the fine roots grew towards the center of the soil in the A-layer, denitrifier activity was even more stimulated in the A-layer (Fig. 2). The stimulating effect was probably restricted to the inner part of the A-layer and therefore the B-layer was less affected during the remainder of the experiment. The stimulating effect on PDRs was largest when the roots grew near the nets (Fig. 2d, sampling I), at which time denitrification activity was correlated with root biomass. Anaerobiosis in soil develops when the rate of OZ consumption is higher than the rate of OZ diffusion into the soil. which is mainly regulated by soil moisture. Denitrification rates in the B-layer were significantly correlated with water content (at the sampling after watering the previous day) throughout the experiment. This was not true for the C- and D-layers. The higher sensitivity of B-layer denitrification to water was probably related to the high consumption of O2 in the A-layer, creating anaerobic conditions, but root exudates cannot be ruled out as a cause. As the root biomass decreased between the 3rd and 4th week, denitrification activity also decreased. This
Microbial N transformations in the root environment
557
nitrification and denitrification measurements, and Inger decrease was more pronounced where larger amounts Ohlsson for assisting in protozoa and bac&a ermmerof root biomass were initially present. Water conations. We also thank Dr Stephen Si for statistical sumption probably increased with increasing biomass advice and Dr David Tilles for revising the English. This and as the soil moisture decreased Oz diffusion should work was supported by grants to the project Ecology o/ have improved, inhibiting denitritication. In addition, Arable Lund. The role of organisttu in nutrient cycling from the decomposition of roots leaves channels which the Swedish Council for the Planning and Coordination of should help oxygenate the soil. Research, the Swedish Council for Forestry and AgriculA sharp decrease in denitrification activity just tural Research, the Swediih Natural Science Research outside the rhizosphere as reported by Smith and Council and the Swedish Environment Protection Board. Tiedje (1979) was not detected at the first sampling of REFERENCES our study. We were not able to determine how far the stimulating root effect reached into the B-layer, but Bailey L. D. (1976) Effects of temperature and root on denitrification in a soil. Canadian Journal of Soil Science it did not reach past the B-layer, which was 4mm 56, 79-87. thick. The PDRs seemed to be energy-limited in all four soil layers, since denitrification activity was less Bakken L. R. and Olsen R. A. (1983) Buoyant densities and dry matter contents of microorganisms: conversion of a than 10% of that found in the same soil during measured biovolume into biomass. AppBed and Ennironincubations with continuous soil stirring (Klemental Microbio/ogy 4!i, 1188-l 195. medtsson ef al., 1987). Myrold and Tiedje (1985) Belser L. W. (1977) Nitrate reduction to nitrite, a possible found lower denitrification rates in anaerobic soil source of nitrite for growth of nitrite-oxidizing bacteria. cores rich in nitrate than in stirred soil slurries of the Applied and Environmentat Microbiology 34, 403-410. same soil. The differences in activity were assumed to Belscr L. W. and Mays E. L. (1980) Specific inhibition of nitrite oxidation by chlorate and its use. in assessing be due to a limitation in the amount of energy nitrification in soils and sediments. Applied and Enuironavailable to denitrifiers. Roots did not appear to mental Microbiology 39, 505-510. increase denitrification rates in the A-layer at samplings 4 and 5. Haider et al. (1985) did not observe Berg P. and Rosswall T. (1985) Ammonium oxidiir numbers, potential and actual oxidation rates in two Swedish any stimulating effects by roots on the denitrification arable soils. Biology and Perfitify of SoiLc 1, 131-140. process. They attributed this lack of stimulation to Berg P. and Rosswall T. (1987) Seasonal variations in the very low concentration of water-soluble root abundance and activity of nittifiers in four arable cropexudates in their soil. Although the denitrification ping systems. Microbial Ecology 13, 75-87. process in our soil was probably energy-limited, it Clarholm M. (198 I) Protozoan grazing of bacteria in soilimpact and importance. Microbial Ecology 7, 343-350. may also have been nitrate limited and inhibited by Ciarhoim M. (1985) Possible roles for roots, bacteria, oxygen. Cycling of denitrification
nitrogen
between
nitrification
and
On a theoretical basis, nitrite produced during ammonium oxidation can maintain a nitrite-oxidizer population one-third the size of the ammoniumoxidizer population (Nicholas, 1978). In our investigation, the numbers of nitrite-oxidizers were usually much higher than those of the ammonium oxidizers and the ratios between the two groups varied considerably, implying the existence of an alternate nitrite source. In a field study, numbers of nitrite-oxidizers were more numerous than ammonium oxidizers (Berg and Rosswall, 1987). Furthermore, the same study concluded that nitrite oxidizers were more numerous close to barley roots, where denitrification is most active (Svensson et al.. 1985). In the present investigation a significant correlation was found between numbers of nitrite-oxidizers and PDRs in the A-layer (r = 0.64; P < 0.01; n = I6 values from the first sampling were omitted, since only a small part of the soil was affected by roots at that time). The above-mentioned results support the theory of a nitrogen cycling between nitrate-reduction to nitrite and nitrite-oxidation back to nitrate (Belser, 1977; Berg and Rosswall. 1987). The nitrate-reductive step may have been performed both by denitrifiers and by nitrate-respirers, since both groups are more numerous in the rhizosphere compared with the bulk soil (Jagnow.
1983).
Acknowledgements-We thank Gunbrit Hansson and Annica Norberg for technical assistance in handling the
protozoa and fungi in supplying nitrogen to plants. In Ecological Interactions in Soil (A. H. Fitter Ed.), pp. 355-365. Special publication No. 4 of the British Ecological Society. Blackwell, Oxford. Clarholm M. and Rosswall T. (1980) Biomass and turnover of bacteria in a forest soil and a peat. Soil Biofogy & Biochemistry 12, 49-57. Darbyshire J. F. and Greavcs M. P. (1973) Bacteria and protozoa in the rhizosphere. Pesticide Science 4.349-360. Haider K., Mosier A. and Heinemcyer 0. (1985) Phytotron experiments to evaluate the effect of growing plants on denitrification. Soil Science Society of America Journal 49, 636-641. Helal H. M. and Sauerbeck D. (1981) Ein Verfahren zur trennung von Bodenzonen underschiedlicher WurzelnPhe. Zeitschrift fiir Pflanzenemaehrung und Bodenkunde 144, 524-527. Jagnow G. (1983) Relations between aerobic heterotrophic, nitrite reducing, dinittifying and nitrogen tixing bact&ia, nitrogenase activity, soil moisture and nitrogen fertilization in soil and rhizosphere of field-grown wheat and barley. In New Tretub in Soil Biology (H. M. Lebrun, A. Andre, A. DeMedts. C. Gregoire-Wibo and G. Wauthy, Eds), pp. 225-235. Proceedings of VIII International Colloquium of Soil Zoology. Louvaine-la-Neuve (Belgium), Dieu-Brichart, Ottignies-Louvain-la Neuve. Jenkinson D. S. and Powlson D. S. (1976) The effects of biocidal treatments on metabolism in soil-V. A method for measuring soil biomass. Soil Biology & Biochemisrry 8, 209-213. Jensen H. L. (1942) Nitrogen Exation in kguminous plants. 1. General characteristics of root-nodule bacteria isolated from ~pc-cics of Medicago and Triforiwn in Australia. Proceedings of rhe Linnean Society of New South Wales 66, 98-108. Jones P. C. T. and Mollison J. E. (1948) A new technique for the quantitative estimation of soil microorganisms. Journal of General Microbiology 2, 54-69.
558
LElF KLEMEDTSSON er al.
Keeney D. R. and Nelson D. W. (1982) Nitrogen-inorganic forms. In Merhodr o/Soil Analysis, Part 2 (A. L. page, Ed.), pp. 643-698. American Society of Agronomy, Madison. Klemedtsson L., Simkins S. and Svensson B. H. (1986) Tandem thermal-conductivity, electron-capture detectors, and non-linear calibration curves in quantitative N,O analysis. Journal of Chromarogruphy 361. 107-I 16. Klemedtsson L., Svensson B. H.,L&lberg T. and Rosswall T. (1977) The use of acetylene inhibition of nitrous oxide reductase in quantifying denitrification in soil. Swedish Journal of Agricultural
Research 7, 179-185.
Klemedtsson L., Svensson B. H. and Rosswall T. (1987) Dinitrogen and nitrous oxide production by denitrification and nitrification in soil with and without roots. Plant and Soil 99, 303-319. Leather J. W. (191 I) Records of drainage in India. Memoirs of the Deparrmenr of Agriculture in India, Chemistry Series 31, 63-140.
Lynch J. M. (1984) Interactions between biological processes, cultivation and soil structure. Planr and Soil 76, 307-3
18.
Molina J. A. E. and Rovira A. D. (1963) The influence of plant roots on autotrophic nitrifying bacteria. Canadian Journal of Microbiology
10, 249-257.
Myrold D. D. and Tiedje J. M. (1985) Diffusion constraints on denitritication in soil. Soil Science Society of America Journal 49, 65 I-657.
Newman E. I. (1985) The rhizosnhere: carbon sources and microbial populations. In Ecological lnteracrions in Soil (A. H. Fitter. Ed.). DD. 107-121. Soecial publication No. 4 of the British &&ical Society: Blackwell. Oxford. Nicholas D. J. D. (1978) Intermediary metabolism of nitrifying bacteria, with particular reference to nitrogen, carbon and sulfur compounds. In Microbiology (D. Schlessinger, Ed.), pp. 305-309. American Society of Microbiology, Washington, D. C. Rice E. L. (1984) Allelopurhy. 2nd edn. Academic Press, Orlando. Rice E. L. and Pancholy S. K. (1972) Inhibition of nitrification by climax vegetation. American Journal of Bofuny 59, 1033-1040. Rovira A. D. and Davey C. B. (1974) Biology of the rhizosphere. In The plam root and its environment (E. W. Carson, Ed.), pp. 153-194. University Press of Virginia, Charlottesville, U.S.A. Ruxicka J. and Hansen E. H. (1975) Flow injection analysis. Part I. A new concept of fast continuous flow analysis. Analytica
Chimica Acta 78, 145-157.
Sarathchandra S. U. (1978) Nitriflcation activities of some New Zealand soils and the effect of some clay types on nitriflcation. New Zealand Journal of Agricultural Research 21, 615-621.
Sarathchandra S. U. (1979) A simplified method for estimating ammonium oxidising bacteria. Plant and Soil 52, 305-309.
Schmidt E. L. and Belser L. W. (1982) Nitrifying bacteria. In Methods of Soil Analysis, Part 2 (A. L. Page, Ed.), pp. 1027-1041. American Society of Agronomy, Madison. Schniirer J. (1985) Fungi in arable soil-Their role in carbon and nitrogen cycling. Dissertation Report 29. Department of Microbiology, Swedish University of Ag ricultural Sciences, Uppsala. Schnilrer J. and Rosswall T. (1987) Mineralization of nitrogen from i5N-labelled fungi, soil microbial biomass and roots and its uptake by barley plants. Plant and Soil (in press). Schniirer J.. Clarholm M. and Rosswall T. (1985) Microbial
biomass and activity in an agricuIturaI soil with diBerent organic matter contents. Soif Biology & BiochemLWy 17, 61 l-618. Schniirer J., Clarhohn M. and Rosswall T. (1986) Fungi, bacteria and protozoa in soil from four arable cropping systems. Biology and Fertility of Soils 2. 119-126. f&h S. M. and-Tiedje J. M. (19?9) The effect of roots on soil denitrification. Soil Science Society of America Journal 43, 951-955.
Steele K. W.. Wilson A. T. and Saunders W. M. H. (1980) Nitrification activity in New Zealand grassland soils. New Zealand Journal of Agricultural
Research 23, 249-256.
Steen E., Jansson P.-E. and Persson J. (1984) Experimental site of the “Ecology of Arable Land” project. Acra Agricullurae
Scandikvica
3, 153- 166.
_
Stefanson R. C. (1972a) Soil denitrification in sealed soilplant systems. I. EtTect of plant, soil water content and soil organic matter content. Plant and Soil 33, 113-127. Stefanson R. C. (1972b) Soil denitriflcation in sealed soilplant systems. II. Effect of soil water content and applied nitrogen. Plum and Soil 33, 129-140. Stefanson R. C. (1972~) Soil denitrification in sealed soilplant systems. III. Effect of disturbed and undisturbed soil samples. P/am and Soil 33. 141-147. Stotxky G. and Norman A. G. (1963) Factors limiting microbial growth activities in soil. III. Supplementary substrate additions. Cunudiun Journal of Microbiology 10, 143-147.
Svensson B. H., Klemedtsson L. and Rosswall T. (1985) Preliminary field denitrification studies of NO;-fertilized and nitrogen-fixing crops. In Denifrifieafion in the Nirrogen cycle (E. L. Golterman Ed.), pp. 157-169. Pro&dings NATO Workshop. Plenum press, New York. SGderstriim B. E. 11977) Vital staining of fungi in pure cultures and in soil with fluorescein d&&ate. Soil Eioiogy & Biochemistry
Tiedje
9, 59-63.
J.
M. (1982) Denitritication. In Methods of Soil Analysis, Part 2 (A. L. Page, Ed.), pp. 1011-1026. American Society of Agronomy, Madison. Theron J. J. (1951) The influence of plants on the mineralization of nitrogen and the maintenance of organic matter in the soil. Journal o/Agriculrural Sciences, Cambridge 41, 289-296. van Veen J. A., Ladd J. N. and Frissel M. J. (1984) Modelling C and N turnover through the microbial biomass in soil. Plant and Soil 76, 257-274. Volz M. G., Ardakani M. S.. Schultz R. K., Stolxy L. H. and McLaren A. D. (1976) Soil nitrate loss during irrigation: Enhancement by plant roots. Agronomy Journal 68, 621-627.
Wheeler G. L. and Donaldson J. M. (1983) Nitrification in an upland forest sere. Soil Biology t Biochemistry IS, 119-121.
Woldendorp J. W. (1963) The influence of living plants on denitrification. Mededelingen van de Lanabouwhogeschool re Wu eningen 63, I-100. Wolden d at-n J. W. f1975) Nitrification and denitritication in the rhixo’sphere. *In L Rhizosphere (F. Mangenot. Ed.), pp. 89-107. Societi Botanique Francaise Colloquium Rhizosphbre. Woldendorp J. W. (1981) Nutrients in the rhizosphere. In Proceedings from I&h Colloquium lntemarional Institure. pp. 99-125. Bern. Switzerland.
Potash
Yoshinari T., Hynes R. and Knowles R. (1977) Acetylene inhibition of nitrous oxide reduction and measurement of denitrification and nitrogen fixation in soil. Soil Biology & Biochemistry
9, 177-183.
~38~717/87
1987
S3.00 + 0.00
Copyright Q 1987 Fcrgamon JoumaIs L&i
MICROBIAL NITROGEN TRANSFORMATIONS ROOT ENVIRONMENT OF BARLEY
IN THE
LEIF KLEMEDTSON,*PER BERG, MARIANNEGARHOLM, JOHANSCHN@RJZR Department of Microbiology, Swedish University of Agricultural Sciences, S-75007 Uppsala, Sweden THOMASROSSWALL
Department of Water in Environment and Society, University of LinkGping. S-58183 Linkiiping, Sweden (Accrpred 4 February1987) !Smnmary-To determine the influence of barley roots on microorganisms and N-transforming processes in soil, numbers of nitrificrs and potential nitrification and denitrification rates were measured every week for 5 wks. The barley plants were grown in growth chambers in which the root-containing soil layer (A) was separated from three outer soil layers (B, C, D). The numbers and biomass of bacteria, numbers of flagellates and amoebae, total and FDA-active hyphal lengths. microbial biomass carbon and respiration
were also determined. The numbers of ammonium oxidizers were positively correlated with root biomass but did not differ significantly between soil layers. Potential ammonium oxidation was stimulated in the ioot-layer, while potential nitrite oxidation was stimulated in the B- and C-layers. The denitrification activity (measured anaerobically in the presence of excess NO;) was positively con-elated with root biomass in the A-layer. Denitritication activity in the B-layer was positively correfated with the water content of the soil. When roots grew near the nets separating the root layer from the other layers, denitrification activity was stimulated in the next layer (B). We propose that nitrite oxidation in the root zone partly depends on the reduction of nitrate. This would explain why nitrite-oxidizer numbers were usually several orders of magnitude higher than ammoniumoxidizernumbers. Bacterial numbers decreased between wks 1 and. 5. Increases in bacteria, naked amoebae and flagellates in all layers between wks 2 and 3 indicated that bacteria were produced until wk 3. There were no signs of bacterial production after wk 3. The total length of hyphae and the length of FDA-active hyphae were not significantly different between layers. However. both of these parameters, as well as total microbial biomass carbon and respiration. were consistently highest in the A-layer.
Microorganisms in soil are affected by plant roots in several ways. For example, root exudates can stimulate heterotrophic growth and lead to local competition for inorganic nutrients between roots and microorganisms. Respiration by roots and Oz consumption by heterotrophic organisms decrease O2 concentrations close to actively growing roots, thereby also affecting the microenvironment. However, 0, diffusion into the soil is also stimulate by roots, since their uptake of water reduces a major barrier of gas diffusion in the soil. The decomposition of dead roots creates air channels, which also facilitate O2 diffusion. The rhizosphere is an illdefined zone a few miltimetres thick that surrounds plant roots. It is a soil volume characterized by the increased activities of fungi but especially bacteria (Rovira and Davey, 1974; Woldendo~, 1981) and protozoa (Darbysbire and Greaves 1973). Tbe limits of the rhizosphere are difficult to establish, and it is probabfe that there are two spheres of influence around roots: (i) a smaller one where enrichment of organic compounds occurs *To whom all correspondence should be addressed, 551
and (ii) a larger one where the roots affect the partial pressures of 0,. CO2 and other soil gases. This larger sphere may be particularly important in regulating the activity of nitrification and denitriiication, which is influenced by the aerobic-anaerobic state of the rhizospheric environment (Woldendorp, 1963; Klemedtsson et al., 1987). Since bacteria depend on easily-available organic compounds for energy (Stotzky and Norman, 1%3), most bacterial activity in soil should thus be linked to either roots or to fresh organic material undergoing decomposition. By far the largest proportion of bacteria in other regions of the soil is dormant (Lynch, 1984). Protozoa are likewise more numerous close to roots, and there is evidence that by feeding on bacteria they cause a local increase in nitrogen mineralization (Clarholm, 1985). In the bulk soil, the biomass of fungi is normally larger than that of bacteria, but either group can dominate in the rhizosphere (Newman, 1985). Ammonium-oxidizers compete with roots and heterotrophic microorganisms for available ammonium. Inhibiting effects of roots on nitrification have been discussed by several authors (Rice and Pancholy, 1972; Woldendorp, 1975; Wheeler and Donaldson, 1983) and reviewed by Rice (1984). Favourable con-
LEIFKLEKEDTSSONet
552
(II
ditions for nitrification do not seem to occur in the rhizosphere (Woldendorp, 1981). since higher concentrations of energy-rich substances will favour hetammonium-immobilizing organisms. erotrophic, Stimulating effects of roots on nitrification have, however, been reported for arable soils (Molina and Rovira, 1963; Berg and Rosswall, 1987). Denitrification activity may be stimulated (Woldendotp, 1963; Stefanson, 1972a-c; Void et al., 1976; Klemedtsson et al., 1987) or inhibited (Smith and Tiedje, 1979) by roots. Inhibition is probably due to competition between denitrifiers and plants for NO; (Smith and Tiedje, 1979). Our aim was to study the influence of roots on the amounts and activities of soil microorganisms at delined distances from plant roots. Of particular interest was whether nitrification and denitrification were stimulated or inhibited by the presence of roots. MATERIALS AND METHODS
Soil characteristics
Topsoil (0- 10 cm) was taken at Kjettslinge, 40 km north of Uppsala in central Sweden, at the site of the Ecology of Arable Lund project. The plow layer (O-27 cm) of the soil was a loam (clay content, 19%; pH 6.3; organic C content, 2.2%; total N content 0.23%) cultivated with unfertilized barley (Ho&urn dktichum L.) (Steen ef al., 1984). The soil was dried, sieved (c 2 mm), mixed and rewetted to a water potential of - 165 kPa (60% of the water holding capacity, WHC). The soil was held at 60% WHC for I wk before starting the experiment to eliminate the easily-available organic carbon produced during the drying and rewetting of the soil. Experimental
system and sampling plan
The soil was packed in growth chambers (Fig. 1) constructed from sheets and edgings of Plexiglas (Helal and Sauerbeck, 1981). Nylon mesh nets were used to establish four compartments at various distances from the roots with four soil layers on each side of the roots of the barley seedlings (Fig. 1). The thickness of the soil layers were: A 20 mm (300 g soil wet wt at 60% WHC); B 4mm (55 g); C 6mm (100 g); D IO mm (150 g). The nets delimiting the soil closest to the roots (A-layer) were of 30 pm mesh to confine all roots to this layer. The other nets had a mesh-size of 100 pm. The pots were kept moist for 4 days, after which two barley seedlings (Ho&urn distichum L. cv. Gunilla) were planted in the A-layer of each pot. The pots (4 replicates x 5 sampling occasions = 20 pots) were kept in a greenhouse at a temperature of 25°C and at a light intensity of 100 W m-*. Four replicates were sampled every week for 5 wk. The soil was kept at 90% WHC during the first 3 wk by placing the pots on a watered sand bed. In addition, the pots were watered from the top every second day. Thus water potentials varied depending on evapotranspiration and watering. At the third sampling the barley plants showed signs of nutrient deficiency, hence a nutrient solution, containing mineral salts but no nitrogen, was added (Jensen, 1942). For 3 days between the third and fourth samplings the temperature control of the greenhouse failed, causing a rise in temperature from 25 to 37°C. As a
2cm I
4
Fig. I. The growth chamber. Thickness of soil layers: A 20 mm, B 4 mm, C 6 mm, D 10nun. The outer dimensions were: 200 x 120 x 65 mm.
consequence, the soil dried out and remained dry (especially in the A-layer) for several days. Sampling
One day before each of the first three samplings the soil was moistened to near water-holding capacity. However, the soil proved to be too wet for efficient use of the fumigation method for microbial biomass determination; thus the water content was reduced after 3 wk to 70-80% WHC. To sample soil, the sheets of Plexiglass were carefully removed and the soil from each layer was collected. Soil from one half of the growth chambers was used to measure denitrification activities. The other half was used for estimating the abundance of bacteria, protozoa and fungi as well as the number of nitrifying bacteria and their potential activity. The A-layer was divided down the middle to form two subsamples, each containing one barley plant with intact roots. Root biomass,
soil moisture
and inorganic nitrogen
Roots were inspected visually at each sampling. Root biomass was determined by washing the roots with tap water over a sieve (0.5 mm mesh) to remove soil particles. The weight of the roots was determined after drying at 85°C overnight. The soil water con-
Microbial N transformations
tents were determined gravimetrically after drying at 10X overnight. Inorganic nitrogen was extracted with 2 M KC1 (10 g wet soil in 50 ml 2 M KCI) on a rotary shaker for 2 h. NO; and NO;- were analyzed by flow injection analysis (FIA; Bifok FIA 05 and Bifok FIA 06 Flow-Photometer, Tecator, Hogan&, Sweden; Ruxicka and Hansen, 1975). Ammonium was assayed by the indophenol-blue method (Keeney and Nelson, 1982). Nitrifer
numbers
The method of soil extraction and microtiter plate
technique used for most-probable-number (MPN) enumeration of ammonium oxidizers have been described by Berg and Rosswall (1985); but the nitrite oxidizer medium was prepared according to Schmidt and Belser (1982). The plates were incubated at 24°C for 8 wks, after which the presence of ammonium oxidizers was determined using a pH-indicator (Sarathchandra, 1979). The rate of nitrite disappearance was used to estimate the abundance of nitrite oxidizers using Griess-Illosvays’ reagents. Potential ammonium- and nitrite-oxidation
rates
in the root environment
553
Bacteria, protozoa, fungi and microbial biomass carbon
Bacteria were counted in a&dine orange-stained soil smears under fluorescent light at 1000x magnification. To estimate biomass, the bacteria were grouped into six size classes with nominal volume values (Clarholm and Rosswall, 1980). A density of l.Igcm-‘andadrywtof30%wereassumedforthe biomass calculations (Bakken and Olsen, 1983). Protozoa were enumerated by a modified MPN technique (Clarholm, 1981). Total hyphal length was estimated by the Jones and Mollison (1948) agar 8lm technique as modified by Schniirer et al. (1985). FDA-active hyphal lengths were determined according to Soderstriim (1977) as modified by Schnilrer et al. (1985). The carbon content of the microbial biomass was determined by the chloroform fumigation technique (Jenkinson and Powlson, 1976) with modifications (Schniirer et al., 1985). Statistical analyses. All statistical analyses conceming the rates of denitrification potential were performed using the Statistical Analysis System (SAS, SAS Institute, Gary, NC U.S.A.) version 4.10. All regressions and analyses of variance were performed using the General Linear Models (GLM) procedure within SAS. All other analyses were made using standard textbook procedures.
Ammonium oxidation was estimated as described by Berg and Rosswall (1985), and was a modification of the method used by Belser and Mays (1980). Samples of soil (25 g) were mixed with medium (4 mM (NH&SO,, pH 7.2) according to Sarathchandra (1979) in capped bottles (300 ml). Chlorate was added to give a final concentration of 15 mM in the soil slurry. The bottles were held on a rotary shaker (200 rev min-‘) at 24°C. Samples (2 ml) were removed from the soil slurry 0, 3 and 6 h after chlorate addition and put in test tubes with 2 ml of 4 M KC1 to inhibit ammonium oxidizer activity. The samples were then centrifuged, filtered and analyzed for accumulated nitrite. Nitrite oxidation was determined by measuring nitrite disappearance at 0, 3 and 6 h after incubating soil in a nitrite oxidizer medium (Schmidt and Belser, 1982) containing 60~~ nitrite (pH 7.2). No chlorate was added; otherwise, procedures were as described above for ammoniumoxidation.
After 1 wk. mainly fine roots were found and they had accumulated close to the nets that confined the A-layers. After 2-3 wk. suberised roots dominated. The fine roots were less abundant near the nets; instead, a concentration of fine roots occurred towards the center of the A-layers. Root biomass decreased between wks 3 and 4 (Fig. 2) due to the unintentional desiccation. At 4 wk the amount of fine roots had decreased further and some dead roots were observed. The root biomass, mainly the fine root fraction, increased again between the fourth and fifth week, after the soil moisture contents had been re-adjusted.
Potential denitrification
Inorganic nitrogen
The soil layers were carefully transferred
to aluminium containers, and KNO,-solution was added to a final concentration of 40 fig NO;-N g-’ dry wt. To determine gaseous nitrogen losses, the containers were placed in gas-tight plastic bags (Cryovac Bags, W. R. Grace Ltd, St Neots, Cambs., England) with butyl-rubber stoppers. The acetylene inhibition technique at 10kPa C,H, (Klemedtsson et al., 1977; Yoshinari et al., 1977) was used to determine denitrification rates. The plastic bags were rapidly evacuated and refilled with N, (< 5~1 Ozl-‘) five times, after which 50 ml neon and 100 ml C2H, were added. The neon was added to determine the gas volume of the bags. The amount of gas dissolved in soil water was calculated using the Bunsen coefficients given by Tiedje (1982). Gases were sample at 1,2.3,4,6 and 8 h with evaculated Venoject tubes (Terumo Europe, N.V. Leuven, Belgium) and analyxed by gas chromatography (Klemedtsson et al., 1986).
RESULTS Root growth
Nitrate-N varied between 0.5 and 2 pg N g-l dry soil. Nitrite concentrations were always below 10 ng N g-’ dry soil. Ammonium concentrations were highest in the A-layer from samplings 1 and 2 (5.2 f 0.8 pg N g-i dry soil; mean + standard error; N = a), compared with the B, C and D layers (1 .l + 0.2; N = 24). It was not possible to analyze ammonium from samplings 3, 4 and 5 due to a technical error. Numbers of nitrlpers
Numbers of ammonium-oxidizers in individual samples varied greatly with time (from 2 x 10’ to 102 x 10’ cells g-i dry soil, Fig. 2). No significant differences were found between layers. The mean numbers of ammonium-oxidizers in all four layers were correlated with the amounts of root biomass in the A-layer (r = 0.91, N = 5, P < 0.05). Numbers of nitrite-oxidizers in individual samples varied less with time (21 x 10’ to 98 x 10’ cells g-i dry soil, Fig. 2)
0.4
!
b1
.-s--
Time
twrrkr
)
Potential ammonium oxidation (a) and nitrite oxidation (b) in the four soil layers (C(g NOT-N g-’ dry soil Fig. 3.
Fig. 2. hot biomass, 11~~bers of NH;- and NO,‘-oxidizers
and potential denitrification during 5 wks with barley growing in a compartmentalized growth chamber. Bars show SE. than did the ammonium-oxidizers, and reached a maximum I wk later than the ammonium-oxidizers (Fig. 2). Potential nitri@ation
rates
Potential ammonium-oxidation rates were consistently highest in the root-layer (Fig. 3a). The rates increased between samplings 2 and 3 (Fig. 3a) 1 wk after the increase in ammonium oxidizer numbers (Fig. 2). No correlation was found between the numbers and activities of ammonium-oxidizers. Potential nitrite-oxidation rates were consistently highest in the B- and C-layers (Fig. 3b). The highest potential rates were found after 2 wk, after which they declined. The numbers and potential rates of nitrite-oxidizers were correlated in the A-layer (r = 0.44; N = 20; P < 0.05), but not in the root-free layers. DenitriJScation activities
Except for the first sampling, the highest potential deniMcation rates (PDRs) were found in the Alayer (Fig. 2). At samplings 1-3 PDRs in the A-layer were correlated with root biomass (r = 0.53; P < 0.02; n = 12). However, the A-layer PDRs were negativeiy correlated with root biomass at the fourth sampling, by which time the root biomass had decreased (r = -0.96; P -C0.04; n = 4). When the activity in the B-layer of all samples was considered, no
h-‘; A’ = 4). Bars show SE.
correIation with root biomass was found, but there was correlation with soil water content (r = 0.45; P < 0.05; n = 20). The highest PDRs were found in the B-layer after 1 wk of plant growth (Fig. 2). These PDRs were correlated with root biomass (r = 0.87; P < 0.05; n = 4). The PDRs in the C- and D-layers were not correlated either with water content or root biomass. At the first sampling, the activity of the B-layer was significantly higher than that of the other layers (P < 0.05). Except for the C-layer at sampling 4 and D-layer at sampling 5, the activities in the A-layer of samples 2-5 were all significantly higher than those in the other layers (P <0.05), Bacteria, protozoa, fmgi
and total microbial biomass
The numbers and biomass of bacteria at sampling 5 were significantly lower (P c 0.05) than those at the first sampling (Table 1). Bacterial numbers had decreased by 43-45%, resulting in 24-33% lower biomass. The bacterial and flagellate development with time was similar in all four layers. The highest values were recorded at the first sampling and the second largest values were recorded at the third (data not shown). The naked amoebae were initially present in low numbers, increased until the third samphng, and then decreased, remaining low until the end of the experiment (data not shown). At the last sampling, mean values of total and FDA-active hyphae as well as biomass-C and U&-production were highest in the A-layer (Table 2). The differences were, however, not statistically significant.
Microbial
in the root environment
N transformations
555
Table I. Numbers and biomass of bacteria and numbers of flagellates and naked amccbac at sampling wks I and 5. All values are expressed g-’ dry soil, mean f SE; n - 4. A = the middle layer. co&sing the root; B, C. D layers at dcfmed distances from the root (set Fin. II
No. of No. of bacteria Layer
Sampling week
A : D
Bacterial biomass
wg-‘)
naked amoebae (lo’g_‘)
wg-‘)
I
5
6.SkO.S 6.3 f 0.4 6.2kO.5 6.1 f 0.2
3.5kO.5 3.5 iO.5 3.5kO.3 3.3 f 0.7
(:gdwg-‘) 520*35 500 f 45 490245 520 k 25
5
I
5
,
5
3SOk40 350 f IS 37Ok40 380 f 25
35 f 6 47&17 3629 32 i 5
21*5 32 f 16 g7*22 48 * 34
66 f 22 &0*4 77* I3 a6* I3
52* I4 51 f I2 4Sf:8 3a*9
DIBCUBBION
Root growth and water conditions
Early in the experiment, most roots were found close to the nets, where physical resistance of the soil to root growth was likely to have been the lowest. After about 2 wks roots began penetrating deeper into the center of the A-layer and there was a corresponding decrease in 8ne roots near the nets. These changes probably resulted from an increasing nutrient deficiency in the soil close to the nets. Water uptake by the plants probably resulted in a net flow of water from the D-, C- and B-layers towards the A-layer. It is therefore unlikely that non-volatile root exudates diffused from the A-layer into the B-layer, because exudates would have had to diffuse against the direction of water flow. Water transport between the A- and B-layers may have been slightly restricted by the fine mesh net. Bacteria, protozoa
No. of flagellates
and figi
The numbers and biomass of bacteria at the first sampling were in the upper range of that found for the same soil in the field (Schniirer et al., 1986). This was probably an effect of the treatment of the soil before the experiment. The increase in bacteria between wks 2 and 3 coincided with an increase in the values for potential ammonium oxidation. Since the increase occurred in all layers, it was probably unrelated to root activity. The dynamics of the flagellate populations were similar to those observed for bacteria. Flagellates require less dense bacterial populations to multiply, compared with naked amoebae. Thus, the flagellate numbers might have only lagged l-2 days behind the bacteria, but this would not have been detected with the sampling interval used. The mean production of amoebae between samplings 1 and 3 was 5 x 10’ amoebae g-i dry wt soil. This is equivalent to a consumption of 2 x 10’ bacteria or 80-1OOpg C, since each new amoeba represents 3000-4000 con-
sumed bacteria (Clarholm, 1981). The carbon for bacterial growth must have been derived mainly from organic matter in the soil, since bacterial and protozoan population dynamics were similar in all four layers. The constant size of the amoeba1 population during the remainder of the experiment (samplings 4 and 5) indicated no further production of bacteria during this period. At the first sampling, hyphal lengths were within the range-although at the lower end-found in field samples of the same soil (Schniirer et al., 1986). Both hyphal lengths and total microbial biomass C at the last sampling were very close to published values for this soil (Schniirer, 1985; Schniirer et al., 1986). Although total and FDA-active hyphae. respiration and microbial biomass C were not significantly higher (P > 0.05) in the A-layer than in the outer layers, all values were numerically highest in the A-layer. The differences in total microbial biomass C between the A-layer and the other layers ranged between 68 and 78~g C g-’ dry wt. Based on the numerical relationships provided by Newman (1985). a standing crop of 175 mg of roots at sampling 5 (Fig. 2) would be equivalent to a total input of from 90 to 210 pg C g-’ soil. This carbon seemed to have been in a form more accessible to fungi than to bacteria, since bacterial biomass decreased during the experiment and FDA-active hyphae was the only biological component that increased in the A-layer between the 1st and 5th samplings. An additional C-source for this increase could have been the roots killed between the 3rd and 4th samplings. The microbial biomass in the same soil grown with barley increased at most 100 pg C g-’ dry wt over a growing season, both in field and greenhouse experiments (Schniirer, 1985; Schniirer and Rosswall, 1987). The initial microbial biomass was about 4OOpg C g-’ dry wt in both cases. Thus, the total amount of microbial biomass in this soil cannot apparently increase by more than 25% in response to carbon additions from the growing barley. This is
Table 2. Total and FDA-active hyphal lengths, microbial biomau carbon and respiration on the tint and the last sampling data and at the last C sampling date (mean values f SE. n = 4). A = the middle layer, enclosing the root: B. C. D lovers at dcfmcd distances from the root (see Fia. I)
Laver A B
Sampling week
Total hyphae (“;” g-’ dry WC) 5 o.s*o.1 0.4*0.1 0.6 f 0.2 0.7 f 0.2
0.8 * 0.2 0.7*0.1 0.4 f 0.0 0.8 f 0.1
FDA-active hThae (yedry wt) 5 Is*2 14*4 l4*6 IO f 2
2125 l7*2 l3*2 Is* I
Rapiration Biomass-C (PgCg-’ (pgCgldry wt) dry wt liday-’ 445*20 367 f 21 370 * I3 377 f I9
163 f I7 105*7 112+5 103+4
)
556
LEIF
KLEMED~SSON
consistent with current theories on a protection capacity for a given size of the microbial biomass, specific for each soil (van Veen et of., 1984). Inorganic nitrogen No inorganic N was added since we wanted to examine the effects of roots on the N-mineralization from soil organic matter. The increased amounts of ammonium in the root-containing A-layer of the two first samplings support the idea of root-mediated N-mineralization (Clarhoim, 1985), especially since potential ~onium-oxidation rates were generally highest in the root layer. A diffusion of N from the other layers could have been another possible cause, but the absence of a gradient of NH: or NO, in B-C layers makes this less probable. Since we were unable to obtain data on the NH: for the three last samplings, it was impossible to determine to what extent local mine~lization had occurred during the rest of the experiment. Numbers and activities of nitrifiers
Numbers of ammonium- and nitrite-oxidizers in the A-layer were higher than was found earlier in soil from Kjettslinge (Berg and Rosswall, 1987). This was probably a result of high ammonium concentrations produced by the rapid mineralization occurring in the dried, sieved and rewetted soil. In the field the ammonium concentrations were about l-2 pg N g-’ dry soil (P. Berg, unpublished). Since potential ammonium-oxidation rates were highest in the Alayer, the corresponding numbers of ammonium oxidizers were also expected to be highest in this layer. This may have been true but was impossible to confirm due to the large variations in the results from the MPN method. Considering the heterogeneity of the soil, the small size (1 g) of the soil sample used for MPN determinations in this study-compared with the log normally used (Berg and Rosswall, 1987)-may be why no differences were detected. Both the potential rates of ammonium-oxidation and nitrite-oxidation varied less than the MPN counts and were of the same order of magnitude found in the Kjettslinge soil (Berg and Rosswall, 1987). Since ammonium-oxidation was not inhibited by the experimental procedure, potential nitriteoxidation rates may have been underestimated, because production and oxidation of nitrite would have proceeded simultaneously. The observed decrease in nitrite must therefore represent a net disappearance. The increase in potential ammonium-oxidation rates between the 2nd and 3rd samplings occurred 1 wk after a corresponding increase in MPN numbers. This is consistent with the results of Berg and Rosswall (1985), which showed that potential ammonium-oxidation rates respond more slowly and proportionally less to short-term changes in soil than MPN numbers. These observations were also supported by the adverse effects on the number and activity of ammonium oxidizers observed after the drying-out incident. Earlier investigations found no relationship between nitrifier numbers and nitrification rates (Sarathchandra, 1978; Belser and Mays, 1980; Steele er al., 1980). We also found that there was no relationship between ammoniumoxidizer numbers and nitrification rates, although
et uf.
such a relationship has sometimes been recorded (Berg and Rosswall, 1985; Berg and Rosswall, 1987). The apparent positive effect of roots on the growth of ammonium-oxidizers, as reflected in the positive correlation between root biomass and ammoniumoxidizer numbers, may actually be coupled to the uptake of water by the roots and thereby to the oxygen status of the soil, since the correlation was found for all layers and not solely in the root layer. Due to the watering of the chambers before sampling, it was not meaningful to calculate correlations between nit~~cation rates and water contents. Several authors have proposed that plant roots could inhibit nitrification (Leather, 1911; Theron, 1951; Woldendorp, 1981). Since the water-flow was assumed to be mostly towards the A-layer, possible soluble inhibitory substances produced by the root could not have played a major role in orbiting nit~fi~tion. The present data show that ~~~~tion can be stimulated in soil within a few millimetres of roots, at least in the early stages of barley growth in arable soil. This is consistent with the results of Molina and Rovira (1963), who showed that growth of Nitrosomonas and Nitrabacter was stimulated by corn and luceme roots within 2 wk after planting. However, this growth promoting effect disappeared after S wks. Potential
denirrification
The denitrification activity in the A-layer positively associated with root biomass, was also recorded by Bailey (1976), Smith and Tiedje (1979) and Klemedtsson et al. (1987). The increase in PDRs in the A-layer, which was associated with increasing root biomass during the first 3 wk, supported the results of Bailey (1976), who studied the effects of roots on denitrification and measured rates in a similar way. Although the denitrification activity in the B-layer was higher than in the A-Iayer at the first sampling, it should be noted that only a fraction of the soil in the A-layer was affected by roots, since these were concentrated in the soil near the smaller B-layer. As the fine roots grew towards the center of the soil in the A-layer, denitrifier activity was even more stimulated in the A-layer (Fig. 2). The stimulating effect was probably restricted to the inner part of the A-layer and therefore the B-layer was less affected during the remainder of the experiment. The stimulating effect on PDRs was largest when the roots grew near the nets (Fig. 2d, sampling I), at which time denitrification activity was correlated with root biomass. Anaerobiosis in soil develops when the rate of OZ consumption is higher than the rate of OZ diffusion into the soil. which is mainly regulated by soil moisture. Denitrification rates in the B-layer were significantly correlated with water content (at the sampling after watering the previous day) throughout the experiment. This was not true for the C- and D-layers. The higher sensitivity of B-layer denitrification to water was probably related to the high consumption of O2 in the A-layer, creating anaerobic conditions, but root exudates cannot be ruled out as a cause. As the root biomass decreased between the 3rd and 4th week, denitrification activity also decreased. This
Microbial N transformations in the root environment
557
nitrification and denitrification measurements, and Inger decrease was more pronounced where larger amounts Ohlsson for assisting in protozoa and bac&a ermmerof root biomass were initially present. Water conations. We also thank Dr Stephen Si for statistical sumption probably increased with increasing biomass advice and Dr David Tilles for revising the English. This and as the soil moisture decreased Oz diffusion should work was supported by grants to the project Ecology o/ have improved, inhibiting denitritication. In addition, Arable Lund. The role of organisttu in nutrient cycling from the decomposition of roots leaves channels which the Swedish Council for the Planning and Coordination of should help oxygenate the soil. Research, the Swedish Council for Forestry and AgriculA sharp decrease in denitrification activity just tural Research, the Swediih Natural Science Research outside the rhizosphere as reported by Smith and Council and the Swedish Environment Protection Board. Tiedje (1979) was not detected at the first sampling of REFERENCES our study. We were not able to determine how far the stimulating root effect reached into the B-layer, but Bailey L. D. (1976) Effects of temperature and root on denitrification in a soil. Canadian Journal of Soil Science it did not reach past the B-layer, which was 4mm 56, 79-87. thick. The PDRs seemed to be energy-limited in all four soil layers, since denitrification activity was less Bakken L. R. and Olsen R. A. (1983) Buoyant densities and dry matter contents of microorganisms: conversion of a than 10% of that found in the same soil during measured biovolume into biomass. AppBed and Ennironincubations with continuous soil stirring (Klemental Microbio/ogy 4!i, 1188-l 195. medtsson ef al., 1987). Myrold and Tiedje (1985) Belser L. W. (1977) Nitrate reduction to nitrite, a possible found lower denitrification rates in anaerobic soil source of nitrite for growth of nitrite-oxidizing bacteria. cores rich in nitrate than in stirred soil slurries of the Applied and Environmentat Microbiology 34, 403-410. same soil. The differences in activity were assumed to Belscr L. W. and Mays E. L. (1980) Specific inhibition of nitrite oxidation by chlorate and its use. in assessing be due to a limitation in the amount of energy nitrification in soils and sediments. Applied and Enuironavailable to denitrifiers. Roots did not appear to mental Microbiology 39, 505-510. increase denitrification rates in the A-layer at samplings 4 and 5. Haider et al. (1985) did not observe Berg P. and Rosswall T. (1985) Ammonium oxidiir numbers, potential and actual oxidation rates in two Swedish any stimulating effects by roots on the denitrification arable soils. Biology and Perfitify of SoiLc 1, 131-140. process. They attributed this lack of stimulation to Berg P. and Rosswall T. (1987) Seasonal variations in the very low concentration of water-soluble root abundance and activity of nittifiers in four arable cropexudates in their soil. Although the denitrification ping systems. Microbial Ecology 13, 75-87. process in our soil was probably energy-limited, it Clarholm M. (198 I) Protozoan grazing of bacteria in soilimpact and importance. Microbial Ecology 7, 343-350. may also have been nitrate limited and inhibited by Ciarhoim M. (1985) Possible roles for roots, bacteria, oxygen. Cycling of denitrification
nitrogen
between
nitrification
and
On a theoretical basis, nitrite produced during ammonium oxidation can maintain a nitrite-oxidizer population one-third the size of the ammoniumoxidizer population (Nicholas, 1978). In our investigation, the numbers of nitrite-oxidizers were usually much higher than those of the ammonium oxidizers and the ratios between the two groups varied considerably, implying the existence of an alternate nitrite source. In a field study, numbers of nitrite-oxidizers were more numerous than ammonium oxidizers (Berg and Rosswall, 1987). Furthermore, the same study concluded that nitrite oxidizers were more numerous close to barley roots, where denitrification is most active (Svensson et al.. 1985). In the present investigation a significant correlation was found between numbers of nitrite-oxidizers and PDRs in the A-layer (r = 0.64; P < 0.01; n = I6 values from the first sampling were omitted, since only a small part of the soil was affected by roots at that time). The above-mentioned results support the theory of a nitrogen cycling between nitrate-reduction to nitrite and nitrite-oxidation back to nitrate (Belser, 1977; Berg and Rosswall. 1987). The nitrate-reductive step may have been performed both by denitrifiers and by nitrate-respirers, since both groups are more numerous in the rhizosphere compared with the bulk soil (Jagnow.
1983).
Acknowledgements-We thank Gunbrit Hansson and Annica Norberg for technical assistance in handling the
protozoa and fungi in supplying nitrogen to plants. In Ecological Interactions in Soil (A. H. Fitter Ed.), pp. 355-365. Special publication No. 4 of the British Ecological Society. Blackwell, Oxford. Clarholm M. and Rosswall T. (1980) Biomass and turnover of bacteria in a forest soil and a peat. Soil Biofogy & Biochemistry 12, 49-57. Darbyshire J. F. and Greavcs M. P. (1973) Bacteria and protozoa in the rhizosphere. Pesticide Science 4.349-360. Haider K., Mosier A. and Heinemcyer 0. (1985) Phytotron experiments to evaluate the effect of growing plants on denitrification. Soil Science Society of America Journal 49, 636-641. Helal H. M. and Sauerbeck D. (1981) Ein Verfahren zur trennung von Bodenzonen underschiedlicher WurzelnPhe. Zeitschrift fiir Pflanzenemaehrung und Bodenkunde 144, 524-527. Jagnow G. (1983) Relations between aerobic heterotrophic, nitrite reducing, dinittifying and nitrogen tixing bact&ia, nitrogenase activity, soil moisture and nitrogen fertilization in soil and rhizosphere of field-grown wheat and barley. In New Tretub in Soil Biology (H. M. Lebrun, A. Andre, A. DeMedts. C. Gregoire-Wibo and G. Wauthy, Eds), pp. 225-235. Proceedings of VIII International Colloquium of Soil Zoology. Louvaine-la-Neuve (Belgium), Dieu-Brichart, Ottignies-Louvain-la Neuve. Jenkinson D. S. and Powlson D. S. (1976) The effects of biocidal treatments on metabolism in soil-V. A method for measuring soil biomass. Soil Biology & Biochemisrry 8, 209-213. Jensen H. L. (1942) Nitrogen Exation in kguminous plants. 1. General characteristics of root-nodule bacteria isolated from ~pc-cics of Medicago and Triforiwn in Australia. Proceedings of rhe Linnean Society of New South Wales 66, 98-108. Jones P. C. T. and Mollison J. E. (1948) A new technique for the quantitative estimation of soil microorganisms. Journal of General Microbiology 2, 54-69.
558
LElF KLEMEDTSSON er al.
Keeney D. R. and Nelson D. W. (1982) Nitrogen-inorganic forms. In Merhodr o/Soil Analysis, Part 2 (A. L. page, Ed.), pp. 643-698. American Society of Agronomy, Madison. Klemedtsson L., Simkins S. and Svensson B. H. (1986) Tandem thermal-conductivity, electron-capture detectors, and non-linear calibration curves in quantitative N,O analysis. Journal of Chromarogruphy 361. 107-I 16. Klemedtsson L., Svensson B. H.,L&lberg T. and Rosswall T. (1977) The use of acetylene inhibition of nitrous oxide reductase in quantifying denitrification in soil. Swedish Journal of Agricultural
Research 7, 179-185.
Klemedtsson L., Svensson B. H. and Rosswall T. (1987) Dinitrogen and nitrous oxide production by denitrification and nitrification in soil with and without roots. Plant and Soil 99, 303-319. Leather J. W. (191 I) Records of drainage in India. Memoirs of the Deparrmenr of Agriculture in India, Chemistry Series 31, 63-140.
Lynch J. M. (1984) Interactions between biological processes, cultivation and soil structure. Planr and Soil 76, 307-3
18.
Molina J. A. E. and Rovira A. D. (1963) The influence of plant roots on autotrophic nitrifying bacteria. Canadian Journal of Microbiology
10, 249-257.
Myrold D. D. and Tiedje J. M. (1985) Diffusion constraints on denitritication in soil. Soil Science Society of America Journal 49, 65 I-657.
Newman E. I. (1985) The rhizosnhere: carbon sources and microbial populations. In Ecological lnteracrions in Soil (A. H. Fitter. Ed.). DD. 107-121. Soecial publication No. 4 of the British &&ical Society: Blackwell. Oxford. Nicholas D. J. D. (1978) Intermediary metabolism of nitrifying bacteria, with particular reference to nitrogen, carbon and sulfur compounds. In Microbiology (D. Schlessinger, Ed.), pp. 305-309. American Society of Microbiology, Washington, D. C. Rice E. L. (1984) Allelopurhy. 2nd edn. Academic Press, Orlando. Rice E. L. and Pancholy S. K. (1972) Inhibition of nitrification by climax vegetation. American Journal of Bofuny 59, 1033-1040. Rovira A. D. and Davey C. B. (1974) Biology of the rhizosphere. In The plam root and its environment (E. W. Carson, Ed.), pp. 153-194. University Press of Virginia, Charlottesville, U.S.A. Ruxicka J. and Hansen E. H. (1975) Flow injection analysis. Part I. A new concept of fast continuous flow analysis. Analytica
Chimica Acta 78, 145-157.
Sarathchandra S. U. (1978) Nitriflcation activities of some New Zealand soils and the effect of some clay types on nitriflcation. New Zealand Journal of Agricultural Research 21, 615-621.
Sarathchandra S. U. (1979) A simplified method for estimating ammonium oxidising bacteria. Plant and Soil 52, 305-309.
Schmidt E. L. and Belser L. W. (1982) Nitrifying bacteria. In Methods of Soil Analysis, Part 2 (A. L. Page, Ed.), pp. 1027-1041. American Society of Agronomy, Madison. Schniirer J. (1985) Fungi in arable soil-Their role in carbon and nitrogen cycling. Dissertation Report 29. Department of Microbiology, Swedish University of Ag ricultural Sciences, Uppsala. Schnilrer J. and Rosswall T. (1987) Mineralization of nitrogen from i5N-labelled fungi, soil microbial biomass and roots and its uptake by barley plants. Plant and Soil (in press). Schniirer J.. Clarholm M. and Rosswall T. (1985) Microbial
biomass and activity in an agricuIturaI soil with diBerent organic matter contents. Soif Biology & BiochemLWy 17, 61 l-618. Schniirer J., Clarhohn M. and Rosswall T. (1986) Fungi, bacteria and protozoa in soil from four arable cropping systems. Biology and Fertility of Soils 2. 119-126. f&h S. M. and-Tiedje J. M. (19?9) The effect of roots on soil denitrification. Soil Science Society of America Journal 43, 951-955.
Steele K. W.. Wilson A. T. and Saunders W. M. H. (1980) Nitrification activity in New Zealand grassland soils. New Zealand Journal of Agricultural
Research 23, 249-256.
Steen E., Jansson P.-E. and Persson J. (1984) Experimental site of the “Ecology of Arable Land” project. Acra Agricullurae
Scandikvica
3, 153- 166.
_
Stefanson R. C. (1972a) Soil denitrification in sealed soilplant systems. I. EtTect of plant, soil water content and soil organic matter content. Plant and Soil 33, 113-127. Stefanson R. C. (1972b) Soil denitriflcation in sealed soilplant systems. II. Effect of soil water content and applied nitrogen. Plum and Soil 33, 129-140. Stefanson R. C. (1972~) Soil denitrification in sealed soilplant systems. III. Effect of disturbed and undisturbed soil samples. P/am and Soil 33. 141-147. Stotxky G. and Norman A. G. (1963) Factors limiting microbial growth activities in soil. III. Supplementary substrate additions. Cunudiun Journal of Microbiology 10, 143-147.
Svensson B. H., Klemedtsson L. and Rosswall T. (1985) Preliminary field denitrification studies of NO;-fertilized and nitrogen-fixing crops. In Denifrifieafion in the Nirrogen cycle (E. L. Golterman Ed.), pp. 157-169. Pro&dings NATO Workshop. Plenum press, New York. SGderstriim B. E. 11977) Vital staining of fungi in pure cultures and in soil with fluorescein d&&ate. Soil Eioiogy & Biochemistry
Tiedje
9, 59-63.
J.
M. (1982) Denitritication. In Methods of Soil Analysis, Part 2 (A. L. Page, Ed.), pp. 1011-1026. American Society of Agronomy, Madison. Theron J. J. (1951) The influence of plants on the mineralization of nitrogen and the maintenance of organic matter in the soil. Journal o/Agriculrural Sciences, Cambridge 41, 289-296. van Veen J. A., Ladd J. N. and Frissel M. J. (1984) Modelling C and N turnover through the microbial biomass in soil. Plant and Soil 76, 257-274. Volz M. G., Ardakani M. S.. Schultz R. K., Stolxy L. H. and McLaren A. D. (1976) Soil nitrate loss during irrigation: Enhancement by plant roots. Agronomy Journal 68, 621-627.
Wheeler G. L. and Donaldson J. M. (1983) Nitrification in an upland forest sere. Soil Biology t Biochemistry IS, 119-121.
Woldendorp J. W. (1963) The influence of living plants on denitrification. Mededelingen van de Lanabouwhogeschool re Wu eningen 63, I-100. Wolden d at-n J. W. f1975) Nitrification and denitritication in the rhixo’sphere. *In L Rhizosphere (F. Mangenot. Ed.), pp. 89-107. Societi Botanique Francaise Colloquium Rhizosphbre. Woldendorp J. W. (1981) Nutrients in the rhizosphere. In Proceedings from I&h Colloquium lntemarional Institure. pp. 99-125. Bern. Switzerland.
Potash
Yoshinari T., Hynes R. and Knowles R. (1977) Acetylene inhibition of nitrous oxide reduction and measurement of denitrification and nitrogen fixation in soil. Soil Biology & Biochemistry
9, 177-183.