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Assessing Denitrifiers Density, Diversity, and Activity

Assessing Denitrifiers Density, Diversity, and Activity

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Soil Denitrifiers


A widely used ex situ assay based on C2H2 inhibition has been developed

to measure the N2O production rate from the pool of active or activateddenitrification enzymes in a sample at the time of sample collection (Smith

and Tiedje, 1979b; Tiedje, 1982). This assay, termed the denitrifying

enzyme assay, is performed by incubating soil slurries under nonlimiting

denitrifying condition (i.e., no oxygen, saturating NOÀ

3 concentration, and

addition of a surplus of electron donors). To avoid de novo enzyme synthesis,

samples are either incubated during a short period of time or in presence of

chloramphenicol, which blocks protein synthesis. The rate of N2O production, which is positively correlated to the amount of denitrification enzymes

in the samples, is then measured. As an alternative, the assay can be used

without addition of chloramphenicol and the denitrification rate can be

estimated by nonlinear regression (Pell et al., 1996). These assays can be used

to compare the effect of agronomical treatments on denitrification.

However, it does not provide information on field rates.

4.1.2. The isotope N-labeled methods

Denitrification activity can be determined using stable nitrogen isotopes in

both laboratory incubations and in field measurements. With this approach,

one or several 15N-labeled nitrogen compounds, such as NOÀ

3 , ammonium,

fertilizers, or plant litter, are added to the soil. The subsequent production

dinitrogen and N2O by denitrification is measured by quantifying the

increase of 15N-labeled gases by mass spectrometry. As with the C2H2

inhibition method, closed chambers are used to estimate denitrification

activity in the field (Nason and Myrold, 1991). This method is limited by

the high cost of 15N and the need to add nitrogen in the soil. Methods based

on the use of 13N have also been described (Smith et al., 1978; Tiedje et al.,

1979), but these cannot be applied in the field (Tiedje et al., 1989).

4.2. Resolving diversity of denitrifiers

Over several decades, diversity of denitrifiers in soil was studied by isolating

bacterial strains. Basically, dilutions of soil suspension were spread on various

agar medium supplemented with NOÀ

3 . After incubation under anaerobic

conditions, isolated colonies were characterized using phenotypic or metabolic tests, and later on by using molecular approaches (Che`neby et al., 2000,

2004; Garcia, 1977; Pichinoty et al., 1976a,b). The most complete survey

was reported by Gamble et al. (1977). From 19 soils, 3 freshwater lake

sediments, and oxidized poultry manure, around 1500 bacteria were isolated

and characterized. The dominant denitrifier populations in most samples

were related to Pseudomonas fluorescens. However, these isolation-based

techniques are limited by the fact that only a fraction of the bacterial

community is cultivable. Research on microbial diversity was completely

revolutionized 20 years ago by the application of molecular methods to


Laurent Philippot et al.

explore microorganisms in the environment without including a cultivation

step. These culture-independent molecular approaches have then been used

to assess the composition of denitrifier communities in soils.

The most frequently used approaches today to target denitrifiers in soil

start with extraction of nucleic acids (DNA or RNA) from the soil (Fig. 3).

The extracted nucleic acids are then purified and amplified by PCR using

primers targeting the denitrifier community. Since the ability to denitrify is

sporadically distributed both within and between different genera, and

cannot be associated with any specific taxonomic group, a 16S rRNAbased approach is not possible to target denitrifiers. However, in the late

1990s, the genes nirS and nirK encoding the key enzymes of the denitrification pathway were first used as molecular markers to describe the diversity

of the denitrifier community (Braker et al., 1998; Hallin and Lindgren,

1999). Since then, this approach has been extended to all the denitrification

genes (Braker and Tiedje, 2003; Flanagan et al., 1999; Philippot et al., 2002;

Scala and Kerkhof, 1999). Amplification of extracted nucleic acids using

primers targeting the denitrification genes is actually the most common way

to analyze denitrifier communities (Bothe et al., 2000; Hallin et al., 2007;

Philippot and Hallin, 2005, 2006). The sequence polymorphism of the

obtained mixed pool of PCR amplicons should reflect the composition of

the denitrifier community in the studied environment. The mixture of

PCR amplicons is analyzed by separating them based on their nucleotide

sequence polymorphism using either clone libraries combined with

sequencing or by fingerprinting techniques (Bothe et al., 2000; Hallin

et al., 2007; Philippot and Hallin, 2006). The most commonly used fingerprinting techniques to study denitrifier communities are terminal restriction

fragment length polymorphism (T-RFLP), restriction fragment length

polymorphism (RFLP), and denaturing gradient gel electrophoresis

(DGGE). These cultivation-independent approaches have limitations

related to the nucleic acids extraction, the choice of PCR primers, and

the PCR itself (Martin-Laurent et al., 2001; Philippot and Hallin, 2005).

4.3. Quantification of denitrifiers

Denitrifiers were first quantified by plating serial dilutions of soil suspension

and counting true denitrifying isolates based on their ability to reduce NOÀ


into gaseous nitrogen production. However, the most common way to

count denitrifiers using a cultivation technique is to apply the most probable

number (MPN) method (Volz, 1977). Serial dilutions of soil suspension are

inoculated into anaerobic replicates medium tubes amended with NOÀ

3 and

C2H2. Dilution tubes are then scored positive when N2O is detected, and

results are then converted into cell numbers copy using the McCrady table.

These methods refer only to microorganisms that can be cultivated and

therefore underestimate the actual number of denitrifiers in the sample.


Soil Denitrifiers

To circumvent this problem, molecular methods have also been developed

to quantify this functional community (Cho and Tiedje, 2002; Gruntzig

et al., 2001; Mergel et al., 2001; Michotey et al., 2000; TaroncherOldenburg et al., 2003; Tiquia et al., 2004; Ward et al., 1993). Two reviews

of these quantitative methods have been published (Philippot, 2006; Sharma

et al., 2007). Today, quantitative PCR is the main method used in soil

environments (Henry et al., 2004, 2006; Kandeler et al., 2006; LopezGutierrez et al., 2004; Qiu et al., 2004) (Fig. 3) with the same bias as for

the cultivation-independent approach for resolving community structure

outlined earlier.

1.000 E+1


Density analysis

1.000 E-1

1.000 E-2

1.000 E-3

1.000 E-4

1.000 E-5










Real time-PCR



Nucleic acids


Structure analysis

Soil samples

Fingerprint analysis


Clone-library analysis

50 100 150 200 250 300 350 400 450 500 550 600 650






Figure 3 Methods used to assess diversity and density of denitrifiers with a PCR-based



Laurent Philippot et al.

5. Natural Factors Causing Variations in


5.1. Temperature and water

Both the overall denitrification rates and the proportions of N2O and

dinitrogen gas produced by denitrifying microbes can vary depending on


numerous environmental factors, such as pH, carbon, NOÀ

3 , and NO2

availability, soil moisture, pore structure, aeration, temperature, freezing–

thawing, and drying–wetting events. Several of these are natural factors

influenced by climatic conditions that cannot be managed. In addition,

they are not constant, but show large variation over the vegetation period

as well as between field sites. The estimated nitrogen losses are therefore

highly variable in time and space. Emissions of N2O and dinitrogen show no

consistent seasonal pattern. In some studies, the largest N2O emissions were

recorded during spring (Kaiser and Heinemeyer, 1996; Parsons et al., 1991;

Ryden, 1985), in others during spring and autumn (Ambus and Christensen,

1995; De Klein and Van Logtestijn, 1994), or in summer (Bremner et al.,

1980; Cates and Keeney, 1987). The difference in the results could not be

related to environmental factors and management practices. A better understanding of factors contributing to variability of denitrification activity

would be helpful to improve estimations and modeling of nitrogen fluxes

by denitrification.

Soil temperature and soil water content are known factors that affect

gaseous nitrogen losses and the N2O/N2 ratio. Under constant laboratory

conditions, this ratio increased exponentially with increasing soil temperature (Maag and Vinther, 1996). However, the ratio was strongly influenced

by soil type, although these data could not be confirmed by field measurements. Whereas Bailey (1976) and McKeeney et al. (1979) found a positive

correlation between soil temperature and denitrification activity, others

observed no relationship with temperature (Focht, 1974; Lensi and

Chalamet, 1979). The reason might be the lower water content caused by

increased plant transpiration rates at higher temperatures, which leads to a

water deficiency. Under laboratory conditions, similar to the effects of

increasing temperature, the overall denitrifying activity and N2/N2O ratio

increased with increasing soil water content (Colbourne and Dowdell,

1984; Vinter, 1984). This was also confirmed in a pasture after harvest

(Rudaz et al., 1997).

Linked to soil water content is oxygen availability. Hochstein et al.

(1984) showed that soil oxygen concentrations below 5% resulted in denitrification being the main microbial respiratory process when NOÀ

3 was

available. In addition, at 10% oxygen concentration and moisture content

between 40% and 60%, denitrification was the main source of emitted N2O.

Soil Denitrifiers


Water content depends on the pore structure of the soil, which in turn is

affected by soil type, organic matter content, and land use. Bakken et al.

(1987) demonstrated that the pore space structure appears to be the major

factor explaining the difference in mean denitrification rates by comparing

pasture and cropped soil. In the field, Bijay-Singh et al. (1989) found higher

actual denitrification in cropped soil than in pasture, despite similar NOÀ


contents. They explained their results as the consequence of better drainage

in the pasture soil, due to the higher porosity of this soil. Complementary

measurements after the application of various amounts of water showed

denitrification activity in pasture soil was higher than denitrification in

cropped soil only at water suctions greater than À5.5 kPa (Bijay-Singh

et al., 1989). In contrast, potential denitrification has often been reported

to be higher in pasture than in cropped soil (Bijay-Singh et al., 1989; Lensi

et al., 1995; Sotomayor and Rice, 1996).

5.2. Freeze–thaw cycles

5.2.1. Freeze–thaw effects on nitrous oxide emissions

Christensen and Tiedje (1990) were the first to report peak N2O emissions

from arable soils in spring during thaw periods. Emissions of carbon dioxide

and N2O and uptake of methane throughout the snow-covered period even

at temperatures near 0  C were later reported (Sommerfeld et al., 1993).

In order to decide whether N2O production can be attributed also to

nonmicrobial processes in soil, emissions from a g-ray sterilized and a

nonsterilized soil were compared in a laboratory experiment, where the

freezing and thawing cycles were simulated. The results clearly indicated

that microbial processes were responsible for N2O production in thawing

and even frozen soils (Roăver et al., 1998). Therefore, efforts have been done

to investigate the effects of freezing and thawing cycles on microbial

denitrification, and to understand the mechanisms behind. Sehy et al.

(2003) first demonstrated the importance of denitrification for nitrogen

losses during winter in arable soil. They separated the 12 months of investigation into the growing season (March to November) and the winter period

(December to February). Independent of the amount of applied fertilizer,

about 70% of the annual N2O amounts was emitted during the winter

period. The temporal changes of the N2O emission rates were correlated to

changes in soil temperature. Similarly, Doărsch et al. (2004) found persistently high N2O emissions in arable soil with peak emissions during midwinter thawing, diurnal freezing–thawing, and spring thaw. Low and stable

temperatures below the insulating snow or ice cover, in contrast, decreased

N2O emissions. Several other field studies in the temperate regions also

reported high N2O emissions from agricultural soils during freeze–thaw

periods reaching 20–70% of the annual budget (Flessa et al., 1995; Nyborg

et al., 1997; van Bochove et al., 1996, 2000; Wagner-Riddle et al., 1997).


Laurent Philippot et al.

Nevertheless, a few studies have also reported that moderate freeze–thaw

fluctuations had little impact on nitrogen dynamics and N2O emissions in

soils (Grogan et al., 2004; Neilsen et al., 2001).

There is considerable debate on which factors could be critical controllers of winter N2O emissions from arable soils. However, most authors state

that emissions during winter are related to the release of nutrients.

Christensen and Christensen (1991) could show that soluble carbon, applied

as plant extract, was necessary to induce N2O production during freezing

and thawing events. Therefore, plant residues from catch crops and green

manure may play an important role in the regulation of N2O emissions in

winter, since frost enhances the release of organic compounds from plant

residues. Additionally, freeze–thaw events may result in transient pulses of

carbon and nitrogen due to disruption of soil aggregates (Christensen and

Christensen, 1991; Muăller et al., 2002) and lysis of microorganisms (Schimel

and Clein, 1996; Skogland et al., 1988). Muăller et al. (2002) showed that the

increased ammonium and NOÀ

3 concentrations during freezing were associated to peak N2O emissions during the following thawing period.

Enhanced oxygen consumption during degradation of plant residues combined with a high water content of the thawing soil increases the anaerobic

volume, thus enhancing denitrification. The freeze–thaw-induced emission

of N2O could thus be a straightforward result of enhanced denitrification.

N2O may also be produced by microorganisms in unfrozen water films on

the soil matrix during freezing. Several authors showed that an ice layer

covering the unfrozen water film could be a diffusion barrier, which reduces

oxygen supply to the microorganisms and partly prevents the release of

N2O to the air (Burton and Beauchamp, 1994; Goodroad and Keeney,

1984; Teepe et al., 2001).

Nitrification could also be of significance for N2O emissions during

winter. It has been demonstrated that freeze–thaw cycles enhances nitrogen

mineralization, which results in the release of substrate for ammoniaoxidizing bacteria (Deluca et al., 1992). Lowered oxygen availability during

freeze–thaw-induced respiration could also induce higher N2O emissions


from nitrifiers, since the N2O/(NO

3 ỵ NO2 ) ratio of nitrification increases

sharply in response to oxygen limitation (Davidson, 1991; Dundee and

Hopkins, 2001; Goreau, 1980). However, it has been demonstrated that

only a few percent of the measured N2O originate from nitrification.

Denitrification was the main N2O source at various oxygen concentrations

investigated in freeze–thaw-affected soil (Ludwig et al., 2004; Mrkved et al.,


5.2.2. Freeze–thaw effects on denitrifier communities

Although microbial denitrification is believed to be the major source of

N2O during freeze–thaw events, few have analyzed the denitrifier communities involved. Actually, little is known about the significance of the

Soil Denitrifiers


denitrifier community composition for N2O emissions in general, since

most of the work conducted has focused on gas and soil analysis. Freeze–

thawing effects on total bacterial community structure are contradictory.

Eriksson et al. (2001) observed a change in ribosomal internal spacer analysis

patterns during freeze–thaw events, whereas Koponen et al. (2006) concluded that neither microbial biomass nor community structure was affected

in boreal soils.

It has been postulated that the relative activity of N2O reductase can be

lowered at near-freezing temperatures (Holtan-Hartwig et al., 2002b; Melin

and Noămmik, 1983), possibly resulting in high N2O/(N2 ỵ N2O) ratios in

soil during thawing. A high N2O/(N2 ỵ N2O) ratio could also be a

‘‘postfreezing trauma’’; the N2O reductase appears to be more vulnerable

to perturbations than the other denitrification enzymes, and if this holds

for frost damages, it would result in a higher proportion of produced N2O

to total denitrification after freezing (Doărsch and Bakken, 2004; HoltanHartwig et al., 2002; Melin and Noămmik, 1983). Nevertheless, how specific

enzymes involved in denitrification are influenced by freezing and thawing

is still not answered.

Sharma et al. (2006) investigated the mRNA levels of genes encoding


the periplasmic NOÀ

3 reductase gene (napA) and cytochrome cd1 NO2

reductase (nirS) in the upper horizon of a grassland soil during thawing in

a laboratory experiment. By using a MPN-based reverse transcriptase PCR

approach they could show that high transcript levels occurred for both

genes 2 days after thawing had begun, followed by a decrease. The peak

of N2O production coincided with the peak for napA and nirS transcripts,

and it timely shifted after 2 days. In the same study, the napA and nirS

genotype diversity was analyzed. Interestingly, DNA-based profiles showed

no change in banding patterns, whereas those derived from cDNA showed

a clear succession of the genotypes, with the most diverse community

structure at the time point of the highest gene expression.

5.3. Dry–wet cycles

Similar to freeze–thaw cycles in soil, dry–wet cycles can enhance N2O

emissions. Prieme´ and Christensen (2001) compared the effects of drying–

wetting and freezing–thawing cycles on the emission of N2O, carbon

dioxide, and methane from intact soil cores from farmed organic soils.

During the first week, following wetting or thawing, up to a 1000-fold

increase in N2O emission rates were recorded from the cores. The total N2O

emission ranged between 3 and 140 mg N–N2O mÀ2, and between 13 and

340 mg N–N2O mÀ2 due to the first wetting and thawing event, respectively. Nevertheless, the emission rates declined after two successive freeze–

thaw events. Many other studies have also documented differences in the

rate of denitrification following wetting (Ambus and Lowrance, 1991;


Laurent Philippot et al.

Gilliam et al., 1978; Groffman and Tiedje, 1989; Rice and Tiedje, 1982;

Robertson and Tiedje, 1985, 1988; Sexstone et al., 1986). Some studies have

also noted denitrification differences between the wet-up and dry-down

phases of soil moisture following rainfall events (Gilliam et al., 1978).

Bergsma et al. (2002) showed that a short wet-treatment significantly

decreased the relative amount of N2O emitted from cropped soil compared

with a long wet-treatment, while no effect of moisture history was seen in a

successional agrosystem. The authors hypothesized that these differences in

N2O production were due to selection of denitrifiers with enhanced capacity for enzyme maintenance at lower levels of NOÀ

3 , such as found in the

successional soil. Others later confirmed differences in denitrifier community composition in the successional and cropped soil at this site (Stres et al.,

2004). Denitrification enzymes were also more sensitive to oxygen in the

cropped soil and N2O activity was higher in the successional soil (Cavigelli

and Robertson, 2000). Soil moisture history seems to be important for

denitrification. If denitrification enzymes are induced differentially in

response to wetting, then both the overall rate of denitrification as well as

the relative amount of N2O will differ substantially among ecosystems.

6. Denitrification in the Rhizosphere of Crops

6.1. Crops as a factor influencing denitrifiers

The rhizosphere is the volume of soil influenced by plant roots (Hiltner, 1904).

The growth and activity of the root system induce significant modifications in the physicochemical and biological properties of the soil surrounding

the roots, which correspond to the so-called rhizosphere effect. It is well

known that the major factors regulating denitrification: carbon, oxygen, and


3 can be modified in the rhizosphere of plants. Thus, carbon compounds,

which can be used as electron donor by denitrifiers, are released by plants roots

in the surrounding soil through rhizodeposition. The effect of plants on

oxygen and NOÀ

3 concentration is more complex. Oxygen concentration

can be lowered in the rhizosphere by respiration of the roots and microorganisms. On the other hand, consumption of water by plant roots increases soil

gas exchange and oxygen concentration. Some plants, such as rice, also

transport oxygen from the air down to the soil in water-saturated soil. Finally,

when roots grow and penetrate the soil, they can modify soil compaction,

which affects oxygen diffusion. Nitrate is used by both plants and microorganisms and the competition for NOÀ

3 is therefore high in the rhizosphere during

the growing season. However, plants can also potentially provide NOÀ

3 for

denitrification when organic matter present in root exudates is mineralized.

Moreover, during plant senescence and litter decomposition in fall and

winter, nitrogen becomes bioavailable and can be denitrified. Overall, factors

Soil Denitrifiers


regulating denitrification in the rhizosphere are strongly interwoven and the

stimulating effect of root-derived carbon is only observed under nonlimiting

concentrations of NOÀ

3 and oxygen. It is therefore not possible to state that

plant roots always stimulate denitrification.

6.1.1. Effect of crops on the denitrification activity

Comparison of denitrification rates between planted and nonplanted soil in

the field or in incubation experiment has been the most common approach

to investigate the influence of crops on this process. Early reports showed

enhanced denitrification rates in the rhizosphere compared with bulk soil

(Smith and Tiedje, 1979a; Stefanson, 1972; Woldendorp, 1962). The key

role of plant on denitrification has later been confirmed in several studies,

although the mechanisms responsible for the higher denitrification rates are

still not clear. Among the agricultural plants studied, barley (Hordum vulgare)

has received the greatest attention so far. Klemedtsson et al. (1987) observed

that denitrification rates in pots planted with barley increased with time

along with increased root biomass. Stimulation of the denitrification rates in

planted pots was 2–22 times compared with the unplanted pots. Similar

results were reported by Hjberg et al. (1996) who observed an average


3 reduction and denitrification rates in the rhizosphere of barley

1.8 times higher than in the bulk soil, with the most pronounced increase

of 7 times. By using monoclonal antibodies against the copper nitrite

reductase, Metz et al. (2003) clearly showed the presence of active enzymes

in the rhizosphere of wheat.

Vinter et al. (1984) demonstrated that this increase of denitrification in

the barley rhizosphere was positively correlated with soil NOÀ

3 concentration.

Their results showed that for fertilizer applied to barley at 30 kg N ha–1, the

denitrification rate increased 2.5 times while a fivefold increase was observed

in field plots receiving 150 kg N ha–1. These results were consistent with those

of Mahmood et al. (1997), who carried out a field experiment to examine the


effect of maize plants on denitrification. At low soil NOÀ

3 levels (1–4 mg N g

dry soil), the presence of maize plants resulted in a nearly 50% increase in

–1 dry soil) the

denitrification, whereas at higher NOÀ

3 levels (7–19 mg N g

observed increase due to plants was 2.5 times. The combined effect of

plant roots and NOÀ

3 concentration on denitrification was first pointed out

by Smith and Tiedje (1979a). They found that denitrification was lower in

planted than in unplanted soil when NOÀ

3 concentration was low (0.002 g

À1 dry soil), while at higher NOÀ concentration (0.1 g NOÀ –N







kgÀ1 dry soil) the presence of plants increased denitrification. Qian et al. (1997)

also reported higher denitrification rates in the unplanted soil compared

with planted soil at late maize growth stages when the amount of NOÀ

3 was

limiting in the planted soil. These neutral or negative effects of plant roots on

denitrification were attributed to NOÀ

3 depletion around the roots.


Laurent Philippot et al.

It has also been reported that the rhizosphere effect on denitrification

was associated with air-filled porosity (Wollersheim et al., 1987). At a low

moisture tension, Bakken (1988) observed a tenfold increase in the denitrification rate in the planted soil compared with the unplanted soil. At

medium or high moisture tension, the plants had no or even a negative

effect on denitrification. Similarly, Prade and Trolldenier (1988) reported

that the rhizosphere effect on denitrification was confined to air-filled

porosity lower than 10–12% (v/v). Thus, the lack of stimulation on denitrification in the rhizosphere at nonlimiting NOÀ

3 concentrations reported

by Haider et al. (1985) was attributed to a high air-filled porosity in both

planted and unplanted pots.

Carbon, the third factor regulating denitrification, is probably responsible for the stimulating effect of plants on denitrification activity. Several

investigators have demonstrated the influence of different organic substrates

on denitrification. Denitrification was correlated with soluble organic matter (Bijay-Singh et al., 1988; Burford and Bremner, 1975; Cantazaro and

Beauchamp, 1985; McCarty and Bremner, 1993) and easily mineralizable

carbon (Bijay-Singh et al., 1988). The release of organic compounds by

living roots can directly affect denitrification rates by providing an additional source of electron donor, but also indirectly by increasing microbial

activity, which lowers the oxygen concentration. This amount of carbon

released by roots into the soil can be up to 20% of photosynthetically

fixed carbon during the vegetation period (Huătsch et al., 2002; Nguyen,

2003). The nature of the root-derived carbon is highly variable (mucilage,

exudates, root cap cells, and so on). The mucilage is composed of highmolecular-weight polysaccharides, mainly arabinose, galactose, fucose,

glucose, and xylose, and up to 6% is proteins. In contrast, exudates are

low-molecular-weight compounds released passively from roots such as

sugars, amino acids, and organic acids. As expected, daily addition of

70 mg C g–1 dry soil of maize mucilage to an agricultural soil increased

denitrification 2.8 times compared with water addition (Mounier et al.,

2004). Similarly, daily addition at a rate of 150 mg C g–1 dry soil of different

mixtures of amino acids, organic acids, and sugars mimicking maize root

exudates greatly stimulated denitrification rates (Henry et al., unpublished

data). In addition, several investigations have shown that denitrification

rates were also positively related to the distribution of fresh plant residues

in the soil profile (Aulakh et al., 1984, 1991; Cantazaro and Beauchamp,

1985; Christensen and Christensen, 1991; Parkin, 1987).

6.1.2. Effect of crop on the denitrifier community

In contrast to denitrification activity, there have been fewer studies of the

effect of plant on the denitrifier community. Vinther et al. (1982) reported

some early estimates of the diversity and the density of denitrifiers in

agricultural soils under continuous barley cultivation. Counts of denitrifiers

Soil Denitrifiers


performed using the most-probable-number method with NOÀ

3 agar broth

as growth medium revealed densities ranging between 103 and 106 bacteria

gÀ1 of dry soil, which represented less than 1% of total bacteria. In contrast,


3 reducers counts for less than 10% of total viable count. Identification

of denitrifying isolates based on selected physiological and morphological

properties showed that numerically predominant denitrifiers belonged to

Pseudomonas spp., Alcaligenes sp., and Bacillus sp. The effect of plant roots on

the taxonomic diversity of denitrifiers has further been investigated by

isolating denitrifiers from unplanted or maize planted soil in a 3-month

incubation experiment (Che`neby et al., 2004). Density of denitrifiers was

2.4 Â 106 and 1.6 Â 107 cells gÀ1 of dry soil in the unplanted and planted

soil, respectively. A total of 3240 NOÀ

3 -reducing isolates were obtained and

188 of these isolates were identified as denitrifiers based on their ability to

reduce at least 70% of the NOÀ

3 to N2O or N2. Comparison of the

distribution of the denitrifying isolates between planted and unplanted soil

showed a difference in the composition of the denitrifier community with

an enrichment of phylogenetically Agrobacterium-related denitrifiers in the

planted soil. In addition, these predominant Agrobacterium-related isolates

from the rhizosphere soil were not able to reduce N2O while dominant

isolates from the unplanted soil emit N2 as end denitrification product.

Direct molecular approaches have recently been applied to investigate

the effect of maize on NOÀ

3 reducers community performing the first step of

the denitrification pathway. The narG gene encoding the membrane-bound


3 reductase was used as molecular marker to analyze the composition of

the NOÀ

3 reducers community from planted and unplanted pots after

3 months of repeated maize culture. A shift in the community composition

between unplanted and planted soils was reported without significant modification of the diversity indices (Philippot et al., 2002b). Clone library

analysis revealed that most of the dominant sequences in the planted soil

were related to narG from the Actinomycetes suggesting a specific selection

` neby

of NOÀ

3 -reducing actinobacteria by the maize roots. In contrast, Che

et al. (2003) detected a reduction of the reciprocal Simpson’s diversity index

in the maize planted soil compared with the unplanted soil, but without any

major modification of the composition of the NOÀ

3 -reducing community

in another soil type. The results from these two studies suggest that the

rhizosphere effect on the structure of the denitrifier community is strongly

dependent on the soil type. Several studies aiming at sorting out the relative

importance of plant and soil confirmed that these two factors might act

simultaneously in determining the composition of the indigenous soil

microbial community (Clays-Josserand et al., 1999; Costa et al., 2006;

Marschner et al., 2004; Wieland et al., 2001).

In two studies, effort has been devoted to disentangle the mechanism of

the rhizosphere effect by investigating the influence of the two major

rhizodeposits, mucilage and exudates, on the genetic structure of denitrifiers

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