soil Bid. &chem. Vol. 19, No. 5, pp. W-558, 1987 Printed in Great Britain. All rights rcKlvcd

~38~717/87 S3.00 + 0.00 Copyright Q 1987 Fcrgamon JoumaIs L&i

MICROBIAL NITROGEN TRANSFORMATIONS IN THE ROOT ENVIRONMENT OF BARLEY

LEIF KLEMEDTSON,* PER BERG, MARIANNE GARHOLM, JOHAN SCHN@RJZR Department of Microbiology, Swedish University of Agricultural Sciences, S-75007 Uppsala, Sweden

THOMAS ROSSWALL

Department of Water in Environment and Society, University of LinkGping. S-581 83 Linkiiping, Sweden

(Accrpred 4 February 1987)

!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 ammonium- oxidizer numbers.

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 stimu- late heterotrophic growth and lead to local com- petition for inorganic nutrients between roots and microorganisms. Respiration by roots and Oz con- sumption by heterotrophic organisms decrease O2 concentrations close to actively growing roots, thereby also affecting the microenvironment. How- ever, 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 facili- tate O2 diffusion.

The rhizosphere is an illdefined zone a few milti- metres 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,

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; Kle- medtsson 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 bac- teria 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 miner- alization (Clarholm, 1985). In the bulk soil, the biomass of fungi is normally larger than that of bacteria, but either group can dominate in the rhi- zosphere (Newman, 1985).

Ammonium-oxidizers compete with roots and het- erotrophic 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-

551

552 LEIF KLEKEDTSSON et (II

ditions for nitrification do not seem to occur in the rhizosphere (Woldendorp, 1981). since higher con- centrations of energy-rich substances will favour het- erotrophic, ammonium-immobilizing organisms. 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 (Wol- dendotp, 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 dis- tances 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 min- eral 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 10 nun. 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 in the root environment 553

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 de- scribed 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 (Sar- athchandra, 1979). The rate of nitrite disappearance was used to estimate the abundance of nitrite oxi- dizers using Griess-Illosvays’ reagents.

Potential ammonium- and nitrite-oxidation rates

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 re- moved 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 ac- cumulated 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, pro- cedures were as described above for ammonium- oxidation.

Potential denitrification

The soil layers were carefully transferred to alu- minium 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 tech- nique 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 C2 H, 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 ana- lyxed by gas chromatography (Klemedtsson et al., 1986).

Bacteria, protozoa, fungi and microbial biomass car- bon

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). Pro- tozoa were enumerated by a modified MPN tech- nique (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 accord- ing to Soderstriim (1977) as modified by Schnilrer et al. (1985). The carbon content of the microbial biomass was determined by the chloroform fumi- gation technique (Jenkinson and Powlson, 1976) with modifications (Schniirer et al., 1985).

Statistical analyses. All statistical analyses concem- ing the rates of denitrification potential were per- formed 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.

Root growth RESULTS

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 to- wards 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.

Inorganic nitrogen

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)

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 consist- ently 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. Po- tential nitrite-oxidation rates were consistently high- est 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 A- layer (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 de- creased (r = -0.96; P -C 0.04; n = 4). When the ac- tivity in the B-layer of all samples was considered, no

0.4 ! b 1

s- .- -

Time twrrkr )

Fig. 3. Potential ammonium oxidation (a) and nitrite ox- idation (b) in the four soil layers (C(g NOT-N g-’ dry soil

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 de- creased by 43-45%, resulting in 24-33% lower bio- mass. 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 N transformations in the root environment 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 Bacterial No. of naked bacteria biomass amoebae

Sampling wg-‘) (:gdwg-‘) 5

flagellates

wg-‘) (lo’g_‘) Layer week I 5 I 5 , 5

A 6.SkO.S 3.5kO.5 520*35 3SOk40 35 6 21*5 f 66 22 52* I4 f : 6.3 f 0.4 3.5 iO.5 500 f 45 350 f IS 47&17 32 f 16 &0*4 51 f I2

6.2kO.5 3.5kO.3 490245 37Ok40 3629 g7*22 77* I3 4Sf:8 D 6.1 f 0.2 3.3 0.7 f 520 25 380 f 25 k 32 5 48 34 i * a6* I3 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 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 be- tween 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 un- related 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 proto- zoan 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 be- tween the A-layer and the other layers ranged be- tween 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 experi- ment 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 experi- ments (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)

FDA-active Rapiration Total hyphae hThae Biomass-C (PgCg-’

Sampling Laver week

(“;” g-’ dry WC) 5

(ye- dry wt) 5

(pg Cgldry wt) dry wt liday-’ )

A o.s*o.1 0.8 * 0.2 Is*2 2125 445*20 163 f I7 B 0.4*0.1 0.7*0.1 14*4 l7*2 367 f 21 105*7

0.6 f 0.2 0.4 f 0.0 l4*6 l3*2 370 * I3 112+5 0.7 f 0.2 0.8 f 0.1 IO 2 f Is* I 377 f I9 103+4

556 LEIF KLEMED~SSON et uf.

consistent with current theories on a protection ca- pacity for a given size of the microbial biomass,

such a relationship has sometimes been recorded

specific for each soil (van Veen et of., 1984). (Berg and Rosswall, 1985; Berg and Rosswall, 1987). The apparent positive effect of roots on the growth

Inorganic nitrogen of ammonium-oxidizers, as reflected in the positive correlation between root biomass and ammonium-

No inorganic N was added since we wanted to examine the effects of roots on the N-mineralization

oxidizer numbers, may actually be coupled to the

from soil organic matter. The increased amounts of uptake of water by the roots and thereby to the oxygen status of the soil, since the correlation was

ammonium in the root-containing A-layer of the two first samplings support the idea of root-mediated

found for all layers and not solely in the root layer.

N-mineralization (Clarhoim, 1985), especially since Due to the watering of the chambers before sampling, it was not meaningful to calculate correlations be-

potential ~onium-oxidation rates were generally tween nit~~cation rates and water contents. highest in the root layer. A diffusion of N from the other layers could have been another possible cause,

Several authors have proposed that plant roots could inhibit nitrification (Leather, 1911; Theron,

but the absence of a gradient of NH: or NO, in B-C 1951; Woldendorp, 1981). Since the water-flow was layers makes this less probable. Since we were unable to obtain data on the NH: for the three last sam-

assumed to be mostly towards the A-layer, possible soluble inhibitory substances produced by the root

plings, it was impossible to determine to what extent could not have played a major role in orbiting local mine~lization had occurred during the rest of nit~fi~tion. The present data show that ~~~~tion the experiment. can be stimulated in soil within a few millimetres of

Numbers and activities of nitrifiers roots, at least in the early stages of barley growth in arable soil. This is consistent with the results of

Numbers of ammonium- and nitrite-oxidizers in the A-layer were higher than was found earlier in soil

Molina and Rovira (1963), who showed that growth of Nitrosomonas and Nitrabacter was stimulated by

from Kjettslinge (Berg and Rosswall, 1987). This was probably a result of high ammonium concentrations

corn and luceme roots within 2 wk after planting.

produced by the rapid mineralization occurring in the However, this growth promoting effect disappeared after S wks.

dried, sieved and rewetted soil. In the field the ammonium concentrations were about l-2 pg N g-’ Potential denirrification

dry soil (P. Berg, unpublished). Since potential The denitrification activity in the A-layer positively ammonium-oxidation rates were highest in the A- associated with root biomass, was also recorded by layer, the corresponding numbers of ammonium oxi- Bailey (1976), Smith and Tiedje (1979) and Kle- dizers were also expected to be highest in this layer. medtsson et al. (1987). The increase in PDRs in the This may have been true but was impossible to A-layer, which was associated with increasing root confirm due to the large variations in the results from biomass during the first 3 wk, supported the results the MPN method. Considering the heterogeneity of of Bailey (1976), who studied the effects of roots on the soil, the small size (1 g) of the soil sample used for denitrification and measured rates in a similar way. MPN determinations in this study-compared with Although the denitrification activity in the B-layer the log normally used (Berg and Rosswall, was higher than in the A-Iayer at the first sampling, 1987)-may be why no differences were detected. it should be noted that only a fraction of the soil in

Both the potential rates of ammonium-oxidation the A-layer was affected by roots, since these were and nitrite-oxidation varied less than the MPN concentrated in the soil near the smaller B-layer. As counts and were of the same order of magnitude the fine roots grew towards the center of the soil in found in the Kjettslinge soil (Berg and Rosswall, the A-layer, denitrifier activity was even more stimu- 1987). Since ammonium-oxidation was not inhibited lated in the A-layer (Fig. 2). The stimulating effect by the experimental procedure, potential nitrite- was probably restricted to the inner part of the oxidation rates may have been underestimated, be- A-layer and therefore the B-layer was less affected cause production and oxidation of nitrite would have during the remainder of the experiment. The stimu- proceeded simultaneously. The observed decrease in lating effect on PDRs was largest when the roots grew nitrite must therefore represent a net disappearance. near the nets (Fig. 2d, sampling I), at which time

The increase in potential ammonium-oxidation denitrification activity was correlated with root bio- rates between the 2nd and 3rd samplings occurred mass. 1 wk after a corresponding increase in MPN num- Anaerobiosis in soil develops when the rate of OZ bers. This is consistent with the results of Berg and consumption is higher than the rate of OZ diffusion Rosswall (1985), which showed that potential into the soil. which is mainly regulated by soil ammonium-oxidation rates respond more slowly and moisture. Denitrification rates in the B-layer were proportionally less to short-term changes in soil than significantly correlated with water content (at the MPN numbers. These observations were also sup- sampling after watering the previous day) throughout ported by the adverse effects on the number and the experiment. This was not true for the C- and activity of ammonium oxidizers observed after the D-layers. The higher sensitivity of B-layer drying-out incident. Earlier investigations found denitrification to water was probably related to the no relationship between nitrifier numbers and high consumption of O2 in the A-layer, creating nitrification rates (Sarathchandra, 1978; Belser and anaerobic conditions, but root exudates cannot be Mays, 1980; Steele er al., 1980). We also found that ruled out as a cause. there was no relationship between ammonium- As the root biomass decreased between the 3rd and oxidizer numbers and nitrification rates, although 4th week, denitrification activity also decreased. This

Microbial N transformations in the root environment 557

decrease was more pronounced where larger amounts of root biomass were initially present. Water con- sumption probably increased with increasing biomass and as the soil moisture decreased Oz diffusion should have improved, inhibiting denitritication. In addition, the decomposition of roots leaves channels which should help oxygenate the soil.

A sharp decrease in denitrification activity just outside the rhizosphere as reported by Smith and Tiedje (1979) was not detected at the first sampling of our study. We were not able to determine how far the stimulating root effect reached into the B-layer, but it did not reach past the B-layer, which was 4mm thick. The PDRs seemed to be energy-limited in all four soil layers, since denitrification activity was less than 10% of that found in the same soil during incubations with continuous soil stirring (Kle- medtsson ef al., 1987). Myrold and Tiedje (1985) found lower denitrification rates in anaerobic soil cores rich in nitrate than in stirred soil slurries of the same soil. The differences in activity were assumed to be due to a limitation in the amount of energy available to denitrifiers. Roots did not appear to increase denitrification rates in the A-layer at sam- plings 4 and 5. Haider et al. (1985) did not observe any stimulating effects by roots on the denitrification process. They attributed this lack of stimulation to the very low concentration of water-soluble root exudates in their soil. Although the denitrification process in our soil was probably energy-limited, it may also have been nitrate limited and inhibited by oxygen.

Cycling of nitrogen between nitrification and denitrification

On a theoretical basis, nitrite produced during ammonium oxidation can maintain a nitrite-oxidizer population one-third the size of the ammonium- oxidizer population (Nicholas, 1978). In our in- vestigation, 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 ammo- nium oxidizers (Berg and Rosswall, 1987). Further- more, 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 numer- ous in the rhizosphere compared with the bulk soil (Jagnow. 1983).

Acknowledgements-We thank Gunbrit Hansson and Annica Norberg for technical assistance in handling the

nitrification and denitrification measurements, and Inger Ohlsson for assisting in protozoa and bac&a ermmer- ations. We also thank Dr Stephen Si for statistical advice and Dr David Tilles for revising the English. This work was supported by grants to the project Ecology o/ Arable Lund. The role of organisttu in nutrient cycling from the Swedish Council for the Planning and Coordination of Research, the Swedish Council for Forestry and Agricul- tural Research, the Swediih Natural Science Research Council and the Swedish Environment Protection Board.

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