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# Control of soil N cycle processes by Pteridium aquilinum and Erica cinerea in heathlands along a pH gradient

Clément Bardon, Boris Misery, Florence Piola, Franck Poly, Xavier Le Roux

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Clément Bardon, Boris Misery, Florence Piola, Franck Poly, Xavier Le Roux. Control of soil N cycle processes by Pteridium aquilinum and Erica cinerea in heathlands along a pH gradient. Ecosphere, 2018, 9 (9:e02426), 10.1002/ecs2.2426. [https://hal.science/hal-01878045](https://hal.science/hal-01878045)

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# Control of soil N cycle processes by *Pteridium aquilinum* and *Erica cinerea* in heathlands along a pH gradient

CLÉMENT BARDON<sup>1,2</sup> BORIS MISERY,<sup>1</sup> FLORENCE PIOLA,<sup>2</sup> FRANCK POLY,<sup>1</sup> AND XAVIER LE ROUX<sup>1,†</sup>

<sup>1</sup>Microbial Ecology Centre, INRA, CNRS, UMR INRA 1418, UMR CNRS 5557, Université Lyon 1, Université de Lyon, Villeurbanne Cedex F-69622 France

<sup>2</sup>LEHNA, UMR CNRS 5023, ENTPE, Université Lyon 1, Université de Lyon, Villeurbanne Cedex F-69622 France

**Citation:** Bardon, C., B. Misery, F. Piola, F. Poly, and X. Le Roux. 2018. Control of soil N cycle processes by *Pteridium aquilinum* and *Erica cinerea* in heathlands along a pH gradient. Ecosphere 9(9):e02426. 10.1002/ecs2.2426

**Abstract.** Nitrate is a limiting resource in heathland acid soils. Nitrate levels increase in heathland soils after *Pteridium aquilinum* invasions, and this species is assumed to biologically control nitrogen cycle processes, thus increasing nitrate availability. We compared how *P. aquilinum* (bracken) and *Erica cinerea* (bell heather) modify processes driving nitrate availability along a soil pH gradient in a Natura 2000 reserve facing bracken invasion. Soil nitrate and ammonium concentrations, substrate-induced respiration (SIR), denitrification and nitrification enzyme activities (DEA and NEA, respectively), root procyanidin concentrations, and denitrification inhibition by procyanidins were measured on five sites under *P. aquilinum* and *E. cinerea* stands. NEA and nitrate levels were higher, and ammonium levels and SIR lower, for *P. aquilinum* in the most acid soils. Procyanidins from both species induced the same level of denitrification inhibition, soil nitrate being correlated with root procyanidin concentration for both species. Soil nitrate correlated with NEA only for *P. aquilinum*. Our results show that both species increased procyanidin production in the most acid soils, thereby reducing denitrification and decreasing nitrate loss, this process being more efficient for *E. cinerea*. However, *P. aquilinum* additionally increased nitrification, and this double control on nitrification and denitrification was very efficient to increase soil nitrate availability in the most acid soils. This may participate to the success of *P. aquilinum* invasions in heathlands. This shows that approaches for bracken control in heathlands should better account for belowground processes and, more generally, that biological denitrification inhibition by plants may be a widespread phenomenon influencing soil N dynamics in N-poor environments.

**Key words:** biological denitrification inhibition; biological invasion; bracken; heather; heathland; invasive species; Natura 2000; nitrate; nitrification; procyanidins; soil nitrogen cycling; soil pH.

**Received** 22 February 2018; revised 24 July 2018; accepted 31 July 2018. Corresponding Editor: Kristofer D. Johnson.

**Copyright:** © 2018 The Authors. This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

† **E-mail:** xavier.le-roux@univ-lyon1.fr

## INTRODUCTION

Nitrogen (N) is a major factor that limits plant growth in many terrestrial ecosystems (Vitousek and Howarth 1991, LeBauer and Treseder 2008). Plants depend heavily on the N availability in soils (Bothe et al. 2006). The N used by plants is mainly in the form of amino acids, $NO_3^-$, and $NH_4^+$ (Boudsocq et al. 2012), and the competition

between roots and microorganisms for these N forms is intense (Hodge et al. 2000, Kuzyakov and Xu 2013). Microorganisms are often considered as better competitors than plant roots for this resource (Hodge et al. 2000, Kuzyakov and Xu 2013). However, during the last twenty years, it has been increasingly reported that plants can modify soil $NO_3^-$ and $NH_4^+$ availability by modulating nitrification and denitrification (Patra
et al. 2006, Le Roux et al. 2013), sometimes through the release of specific compounds by roots (Subbarao et al. 2009, DeLuca et al. 2013, Dietz et al. 2013, Bardon et al. 2014, Srikanthasamy et al. 2018). These strategies to control N cycling processes greatly contribute to plant survival or capacity to colonize adverse environments, particularly in ecosystems with low soil N availability (Ishikawa et al. 2003, Bardon et al. 2014).

In acid soils, selection pressure for $NO_3^-$ is intense because of its low levels, and $NH_4^+$ and amino acids are considered as the predominant N forms (Havill et al. 1974, Troelstra et al. 1995). Plants growing on acid soils often develop a preference for $NH_4^+$ and high abilities to assimilate N from amino acids, but some also assimilate $NO_3^-$ (Rorison 1986, Raven et al. 1992, Troelstra et al. 1995, Jones et al. 2005). In these soils, selection pressure could also have promoted strategies for biological control of key N cycle processes such as mineralization, nitrification, and denitrification to increase soil $NO_3^-$ availability and allow a significant uptake of $NO_3^-$ by plants, but this largely remains to be explored. Nitrification is an aerobic process, which consists in the oxidation of $NH_4^+$ to $NO_3^-$ (Bothe et al. 2006), and is principally driven by abiotic factors such as $NH_4^+$ availability, pH, and temperature, along with the abundance of nitrifiers (Bothe et al. 2006, Le Roux et al. 2008). Nitrification is highly sensitive to soil acidity. Below pH 5, nitrification, and hence $NO_3^-$ production, often remains very low (De Boer and Kowalchuk 2001, Yao et al. 2011). On the other hand, denitrification is a reductive process, which transforms water-soluble $NO_3^-$ into the gaseous forms NO, $N_2O$, and $N_2$ (Zumft 1997), and is a major pathway for the loss of $NO_3^-$ (Bothe et al. 2006). Denitrification is mainly driven by abiotic factors such as the availability of $NO_3^-$, carbon substrate, $O_2$, pH, and temperature, along with the abundance of denitrifiers (Zumft 1997, Wallenstein et al. 2006). Denitrification is less affected than nitrification by low pH (Bothe et al. 2006), which also contributes to the decrease in $NO_3^-$ availability in acid soils (ŠImek and Cooper 2002).

Developed on acid soils, heathland is an ecosystem of great conservation value because of the important cultural and ecological services

it provides, such as water purification, carbon sequestration, and biodiversity conservation (Gimingham 1972, Fagundez 2013). French Atlantic heathlands are dominated by ericaceous dwarf shrubs (heathers) such as *Erica cinerea* (bell heather; Gallet and Roze 2001). They are characterized by $NO_3^-$-poor soils (De Boer et al. 1990, Troelstra et al. 1990). In many heathlands, the heather cover is threatened by the invasion of *Pteridium aquilinum* (Mitchell et al. 1999, Pakeman et al. 2002, DeLuca et al. 2013). Native in temperate ecosystems, *P. aquilinum* is one of the most widespread plant species around the world and is highly tolerant to marked soil acidity (Marrs et al. 2000). *Pteridium aquilinum* invasion of heathlands has been attributed to its fast growth, high biomass production, and ability to outgrow ericaceous shrubs despite the low soil pH (Marrs et al. 2000, DeLuca et al. 2013). Different strategies have been proposed for bracken control in heathlands (Milligan et al. 2016). However, a major limitation for vegetation management and restoration ecology is the ability to predict the outcome of proposed treatments across a range of sites, which is essential to meet the objectives of both policy-makers and practitioners implementing management schemes. In the case of bracken control in heathlands, meta-analyses have demonstrated that the effectiveness of control strategies varies between sites (Stewart et al. 2008) and there is a need to ascertain why the outcomes of different strategies are so variable (Pakeman et al. 2002).

Actually, bracken control strategies often rely on aerial spraying of herbicide and/or cutting (Pakeman et al. 2002, Stewart et al. 2008) and they have largely disregarded the specificities of belowground processes associated with this plant species (but see Milligan et al. 2018). It has been reported that $NO_3^-$ concentration is up to 10 times higher in soil under *P. aquilinum* than in surrounding soil in heathland (Mitchell et al. 1999, DeLuca et al. 2013). Moreover, Stams and Lutke Schipholt (1990) reported that in a woodland acid soil under $NH_4^+$ fertilization, *P. aquilinum* concentrates $NO_3^-$ in its leaves up to 100 times more than other plant species. These results suggest that *P. aquilinum* has developed an efficient strategy to increase $NO_3^-$ availability in acid soils.
Recently, Bardon et al. (2016) demonstrated that soil denitrification is reduced in response to the release of dimeric to tetrameric B-type procyanidins by roots of some plant species. This phenomenon is called biological denitrification inhibition (BDI). Norris et al. (2011) and Bardon et al. (2017) reported that, in contrast to different condensed tannin mixtures, purified B-type procyanidins with a chain length below 4 do not inhibit soil nitrification and N mineralization. The addition of these compounds to a soil can increase soil $NO_3^-$ content, likely through denitrification inhibition (Norris et al. 2011, Bardon et al. 2017), which can limit $NO_3^-$ losses from soils and increase N availability to plants (Bardon et al. 2014). However, both *P. aquilinum* and *E. cinerea* are known to produce B-type procyanidins (Wang et al. 1976, Hofland-Zijlstra and Berendse 2009) and thus may inhibit denitrification and limit $NO_3^-$ losses. Differences in procyanidin production (in terms of type and quantity) may explain differences in $NO_3^-$ availability in soils under *P. aquilinum* and *E. cinerea*. In addition, some plant species stimulate nitrification (Lata et al. 2000, Hawkes et al. 2005), which increases soil $NO_3^-$ availability. This is another explanation (non-exclusive to BDI) for the different $NO_3^-$ levels reported for soils under the two species.

As soil pH is a major factor that drives $NO_3^-$ production in heathland (Troelstra et al. 1990), we explored the control strategy of N cycle processes that drive $NO_3^-$ availability in *P. aquilinum* and *E. cinerea* stands in Atlantic heathland sites along a soil pH gradient. We hypothesized that both species could influence nitrification and denitrification, but that *P. aquilinum* would have a more efficient strategy than *E. cinerea* to increase soil $NO_3^-$ availability particularly at lower pH. Our objectives were (1) to test whether both species can induce BDI—in particular, we studied how each plant species influences denitrification according to the quantity and type of procyanidins produced; (2) to assess whether the influence of *P. aquilinum* and/or *E. cinerea* on nitrification can also contribute to increase soil $NO_3^-$ availability; and (3) to discuss the possible implications of the control of denitrification and nitrification by *P. aquilinum* and *E. cinerea* for the performance of these species and the invasive capacity of *P. aquilinum* in Atlantic heathland.

We also discuss our results in the context of bracken control strategies in heathlands.

## METHODS

### Site selection and biological material sampling

This study was conducted in *Erica cinerea*-dominated heathlands of the Natura 2000 site Récifs et landes de la Hague (Normandy, France) in March 2015. Bracken invasion is recognized as an important threat for heathlands at this Natura 2000 site (INPN 2017). Soils are poorly developed acidic podsolic soils. Five sites where bracken invasion is observed and with contrasting pH were selected, each with *E. cinerea* and *Pteridium aquilinum* stands. The sites were organized as a gradient of decreasing pH (see Appendix S1: Table S1 for soil characteristics): site 1 (pH = 4.75; 49°39′,715″ N 001°52′,325″ W), site 2 (pH = 4.69; 49°41′, 908″ N 001°49′748″ W), site 3 (pH = 4.67; 49°41′, 840″ N 001°49′784″ W), site 4 (pH = 4.59; 49°41′ 343″ N 001°55′ 975″ W), and site 5 (pH = 4.50; 49°41′, 352″ N 001°56′019″ W). Sixteen soil samples per site (eight from *P. aquilinum* and eight from *E. cinerea* stands) were collected from the 0–15 cm soil layer after removing the plant litter. Large organic fragments were removed. Soil samples were homogenized by sieving (2 mm) and stored at 4°C before laboratory measurements. Eight root samples per site (four under *P. aquilinum* cover and four under *E. cinerea* cover) were randomly collected from the 0–10 cm soil layer using an auger (10 cm diameter) and cleaned with water. Soils aliquots and roots were lyophilized, crushed in a TissueLyser II bead mill (Qiagen, Venlo, The Netherlands), and stored at –20°C.

### Measurements of soil moisture, $NO_3^-$ and $NH_4^+$, and soil and plant N concentrations

The moisture content and water-holding capacity of the sieved soils were determined by gravimetric and pressure plate techniques, respectively. $NO_3^-$ and $NH_4^+$ were extracted from freshly sampled soils (3 g eq. of dried soil) supplemented with 30 mL of a 0.01 M solution of $CaCl_2$. Soil suspensions were shaken at 140 rpm for 2 h at 10°C. The suspension was filtered (0.2 µm), and the $NO_3^-$ and $NH_4^+$ were quantified using an ICS-900 ion chromatography system (Thermo Scientific Dionex, Sunnyvale,
California, USA). Total N content was measured on 10 mg of lyophilized and crushed soil and 2 mg of root material enclosed in tin capsules, using a Flash EA 1112 NC elemental analyzer (Thermo Electron, Milan, Italy).

## Measurement of soil microbial activities

Substrate-induced respiration (SIR) was measured according to Anderson and Domsch (1978). Soil samples (5 g eq. of dried soil) were placed in an airtight flask in 21% O₂ atmosphere, supplemented with a nutrient solution, including glucose (1.2 mg of C/g of dried soil) to reach 70% of the water-holding capacity. Soil was incubated at 28°C, and CO₂ levels were measured each hour for 8 h. The slope of the linear CO₂-time regression was used to estimate aerobic respiration (µg C-CO₂·[g of dried soil]⁻¹·h⁻¹).

Denitrifying enzyme activity (DEA) was measured according to Patra et al. (2005). Soil samples (10 g eq. of dried soil) were placed in a 150-mL airtight plasma flask sealed with a rubber stopper. In each flask, the air was removed and replaced with a He/C₂H₂ mixture (90/10; v/v) to create anoxic conditions and inhibit N₂O-reductase activity. A nutrient solution containing glucose (0.5 mg of C/g of dried soil), glutamic acid (0.5 mg of C/g of dried soil), and potassium nitrate (50 µg of N/g of dried soil) was added to the soil to reach 100% of the water-holding capacity. During the incubation at 28°C, the amount of N₂O in the headspace was measured each hour for 5 h. The slope of the linear N₂O-time regression was used to compute DEA (µg of N-N₂O·[g of dried soil]⁻¹·h⁻¹). Gases (CO₂ and N₂O) were measured using gas chromatography coupled to a microkatharometer detector (µGC-R3000; SRA Instruments, Marcy l'Etoile, France).

In order to measure nitrifying enzyme activity (NEA), soil samples (3 g eq. of dried soil) were placed in an airtight flask with 21% O₂ atmosphere and supplemented with 30 mL of a water solution containing (NH₄)₂SO₄ at 5 µg of N/mL. The flasks were incubated at 28°C and shaken at 140 rpm. The soil suspension was sampled and filtered every two hours for 10 h. The amount of NO₃⁻ produced during incubation was measured in samples using an ICS-900 ion chromatography system. The slope of the linear NO₃⁻-time regression was used to estimate NEA (µg N-NO₃⁻·[g of dried soil]⁻¹·h⁻¹).

## Root metabolite extraction, procyanidin quantification, and extract analysis

Root metabolite extraction was carried out with 150 mg of dried and crushed roots suspended in 10 mL water:methanol mixture (50/50; v/v) and shaken at 140 rpm for 1 h in 15-mL Falcon centrifuge tubes. The tubes were centrifuged, and the supernatants were used as crude extracts to quantify root procyanidins. These were quantified using the acid butanol method with crude extracts (Dalzell and Kerven 1998) as in Bardon et al. (2016). Procyanidins contained in 1 mL of crude extracts were oxidatively cleaved to anthocyanidins (i.e., cyanidins and delphinidins) at 95°C for 50 min by adding 6 mL of acid butanol reagent (butanol/12 N HCl; 95/5; v/v) and 200 µL of ferric ammonium sulfate (20 mg/mL) with 2N HCl. The reaction mixture was cooled, and anthocyanidins were quantified spectrometrically at 550 nm using different commercial proanthocyanidin solutions (Laffort, Bordeaux, France) as standards (0.25, 0.5, 1, 2.5, 5, 7.5, and 10 mg/mL).

After procyanidin quantification, crude extracts were dried and re-suspended in water: methanol (50/50; v/v) at a concentration of 10 mg/mL for HPLC MS/MS analysis. Root extract analyses were performed as in Bardon et al. (2014), using an UHPLC 1290 infinity series (Agilent Technologies, Santa Clara, California, USA) coupled to a G4212A DAD and a Q-TOF 6530 series detector with jet stream electrospray ionization. MassHunter Acquisition software (Agilent Technologies) was used to run the system. A Poroshell 120 EC-C18 column (2.7 lm, 3.0 9 100 mm; Agilent Technologies) carried out separation at a temperature of 60°C. The mobile phase was a linear gradient of 0.4% of acetic acid in water (solvent A) and acetonitrile (solvent B). Using a flow rate of 0.6 mL/min, the linear gradient was as follows: 0–1.5 min 2% solvent B; 1.5–22 min 2% to 17% solvent B; 22–41 min 17% to 100% solvent B; 41–41.5 min 100% solvent B. The system was equilibrated before each sample. The injection volume was 1 µL. N₂ was used as the nebulizing gas, with a flow rate of 12 L/min and a temperature of 310°C at 40 psi. Procyanidins were identified by analyzing their UV and MS/MS spectra, using MassHunter Qualitative Analysis (Agilent Technologies). Spectra were recorded between 190 and 600 nm, and the
response at 280 nm was used for qualitative
interpretations.

Effect of root extracts on denitrification of
Pseudomonas brassicacearum

The effect of *P. aquilinum* and *E. cinerea* root extracts on the denitrification activity of *P. brassicacearum* NFM421, a denitrifying bacterium frequently encountered in soils, was tested as described by Bardon et al. (2014, 2016). Briefly, 5 mL of nutrient broth liquid culture medium with 20 mmol/L KNO₃ was supplemented with extracts of known procyanidin concentration to reach a final proanthocyanidin concentration of 0.05 mg/mL, with water:methanol mixture (50/50; v/v; 2.5% methanol final) as a control, and inoculated with a *P. brassicacearum* culture in order to reach an initial optical density of 0.1. Denitrification activity was measured by surveying N₂O production under anaerobic conditions every two hours for 4 d with a gas chromatograph. Four replicates were used for each type of root extract and for the control.

Data and statistical analyses

Two-way ANCOVAs were performed to test
the global effects of the site gradient—repre-
sented by site number, species, and their interac-
tion on soil NH₄⁺, soil NO₃⁻, total soil N, total
root N, root procyanidin concentration, SIR,
NEA, and DEA. Differences between species
among sites were tested using a Tukey HSD test
after two-way ANCOVAs. The differences in val-
ues observed for soil characteristics, microbial
activities, and procyanidin concentration between
*P. aquilinum* and *E. cinerea* along the gradient were
expressed as a function of soil pH and tested
through linear or non-linear regression. In com-
plement, mixed models were performed with site
as a random factor to analyze the effects of plant
species and possible explanatory variables on
selected soil variables: (1) soil NO₃⁻ with NEA as
explanatory variable, nitrification being the pro-
cess producing NO₃⁻, (2) NEA with soil pH,
given the possible sensitivity of nitrification to
pH, (3) NEA with its substrate soil NH₄⁺, (4)
NEA with root procyanidin concentration, (5)
DEA with root procyanidin concentration, and
(6) soil NO₃⁻ with root procyanidin concentra-
tion. When the interaction between species and

co-factors was significant, individual linear
regressions were performed for the two species.
To test the effect of *E. cinerea* and *P. aquilinum*
extracts on denitrification of *P. brassicacearum*, a
one-way analysis of variance was performed
with the treatment (i.e., control, *E. cinerea*, and
*P. aquilinum*) as factor.

For mineral N and microbial activities, data
were log-transformed to ensure data normality.
All analyses were carried out using R software
(R Development Core Team 2008).

RESULTS

Effect of Erica cinerea and Pteridium aquilinum on soil NO3-, NH4+, and total N

Under both *E. cinerea* and *P. aquilinum* stands, soil NO₃⁻ concentration increased along the gradient of decreasing pH (Fig. 1A). However, NO₃⁻ concentration increased much more for *P. aquilinum*, and NO₃⁻ concentration became higher in *P. aquilinum* than *E. cinerea* soils for the lowest pH sites (sites 4 and 5; Fig. 1A, C, Table 1). In contrast, NH₄⁺ concentration was lower in *P. aquilinum* than *E. cinerea* soils for the lowest pH site (Fig. 1B, D, Table 1). No differences in total soil N concentrations or total root N concentrations were observed between the two species (Table 1).

Effect of E. cinerea and P. aquilinum on
potential soil aerobic respiration, denitrification
and nitrification, and links with soil parameters

Substrate-induced respiration was higher in *E. cinerea* than *P. aquilinum* soils except for site 1 (Table 2; Appendix S1: Fig. S1). DEA was higher for *P. aquilinum* than *E. cinerea* in the sites with highest soil pH (sites 1 and 2; Fig. 2A, C, Table 2). Nitrifying enzyme activity (NEA) was higher in *P. aquilinum* than *E. cinerea* soils for the sites with lowest soil pH (Fig. 2B, D, Table 2).

Considering site as a random factor, soil NO3- concentration was positively correlated with NEA for P. aquilinum, but not for E. cinerea (Fig. 3, Table 3). NEA was negatively and weakly correlated with soil pH with no differences between E. cinerea and P. aquilinum, whereas NEA was not correlated with soil NH4+ concentration for both E. cinerea and P. aquilinum (Fig. 4, Table 3).
<figure><img src="image_3.png" /></figure>

Fig. 1. Concentrations of (A) nitrate, NO₃⁻, and (B) ammonium, NH₄⁺, in soils from *Erica cinerea* (black) and *Pteridium aquilinum* (gray) stands for the five study sites. Vertical bars indicate standard errors (n = 8). Stars indicate significant differences between species among sites (*P* < 0.05). See Table 1 for results of statistical analyses. Panels on the right present the difference in soil (C) nitrate and (D) ammonium observed for *P. aquilinum* as compared to *E. cinerea* along the pH gradient.

### E. cinerea and P. aquilinum root procyanidins: concentrations, effects on denitrification, and correlation with soil NO₃⁻

Procyanidin concentration was lower in *P. aquilinum* than *E. cinerea* roots for all sites, and the root procyanidin concentration increased with decreasing soil pH for both *P. aquilinum* and *E. cinerea* (Fig. 5A, Table 2). However, the lower the pH, the higher the difference in root procyanidin concentration between *E. cinerea* and *P. aquilinum* (Fig. 5B). NEA and DEA were not correlated to root procyanidin concentration (Table 3).

Extracts of both *E. cinerea* and *P. aquilinum* roots (using a same procyanidin concentration of 0.05 mg/mL) inhibited denitrification of *P. brassicacearum* (55% of inhibition), with no significant difference between species (Fig. 5C). Flavan-3-ol monomers encountered in *E. cinerea* and *P. aquilinum* root extracts were catechin and epicatechin

monomers of procyanidins (Appendix S1: Fig. S2). Procyanidins encountered in *E. cinerea* were B-type procyanidins, ranging from dimeric to tetrameric forms, and B-type procyanidins with one A-type bond, ranging from dimeric to trimeric forms (Appendix S1: Fig. S2). Procyanidins encountered in *P. aquilinum* were only dimeric B-type procyanidins (Appendix S1: Fig. S2).

For both species, the soil NO₃⁻ concentration observed in the field was correlated with the procyanidin concentration in the roots, with a higher slope for *P. aquilinum* (Table 3; Appendix S1: Fig. S3).

## DISCUSSION

Here, we demonstrate, for the first time, the existence of biological controls of soil N cycle processes related to NO₃⁻ availability by two
<table><caption>Table 1. Results of two-way ANCOVA on the soil NO₃⁻ concentration, soil NH₄⁺ concentration, soil pH, total soil N content, and total root N content, using the environmental gradient, species, and their interaction as main factors.</caption><thead><tr><th>Variables</th><th>df</th><th>F</th><th>P</th></tr></thead><tbody><tr><td>(a) NO<sub>3</sub><sup>-</sup></td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>34.60</td><td>&lt;0.001</td></tr><tr><td>Species</td><td>1</td><td>5.10</td><td>&lt;0.005</td></tr><tr><td>Gradient × Species</td><td>1</td><td>3.92</td><td>&lt;0.05</td></tr><tr><td>(b) NH<sub>4</sub><sup>+</sup></td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>0.16</td><td>0.68</td></tr><tr><td>Species</td><td>1</td><td>4.07</td><td>&lt;0.05</td></tr><tr><td>Gradient × Species</td><td>1</td><td>6.52</td><td>&lt;0.01</td></tr><tr><td>(c) pH</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>16.94</td><td>&lt;0.001</td></tr><tr><td>Species</td><td>1</td><td>11.56</td><td>&lt;0.005</td></tr><tr><td>Gradient × Species</td><td>1</td><td>2.69</td><td>0.11</td></tr><tr><td>(d) Total soil N</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>3.37</td><td>0.07</td></tr><tr><td>Species</td><td>1</td><td>1.80</td><td>0.18</td></tr><tr><td>Gradient × Species</td><td>1</td><td>2.10</td><td>0.15</td></tr><tr><td>(e) Total root N</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>1.32</td><td>0.25</td></tr><tr><td>Species</td><td>1</td><td>1.11</td><td>0.29</td></tr><tr><td>Gradient × Species</td><td>1</td><td>1.40</td><td>0.24</td></tr></tbody></table>

Note: See Fig. 1 for graphical representations.

<table><caption>Table 2. Results of two-way ANCOVA on the substrate-induced respiration, SIR, nitrifying enzyme activity, NEA, denitrifying enzyme activity, DEA, and root procyanidin concentration, using the environmental gradient, species, and their interaction as main factors.</caption><thead><tr><th>Variables</th><th>df</th><th>F</th><th>P</th></tr></thead><tbody><tr><td>(a) SIR</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>0.29</td><td>0.58</td></tr><tr><td>Species</td><td>1</td><td>50.21</td><td>&lt;0.001</td></tr><tr><td>Gradient × Species</td><td>1</td><td>12.55</td><td>&lt;0.001</td></tr><tr><td>(b) NEA</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>7.94</td><td>&lt;0.005</td></tr><tr><td>Species</td><td>1</td><td>4.53</td><td>&lt;0.05</td></tr><tr><td>Gradient × Species</td><td>1</td><td>4.16</td><td>&lt;0.05</td></tr><tr><td>(c) DEA</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>2.54</td><td>0.11</td></tr><tr><td>Species</td><td>1</td><td>4.96</td><td>&lt;0.05</td></tr><tr><td>Gradient × Species</td><td>1</td><td>0.06</td><td>0.79</td></tr><tr><td>(d) Procyanidins</td><td></td><td></td><td></td></tr><tr><td>Gradient</td><td>1</td><td>38.70</td><td>&lt;0.001</td></tr><tr><td>Species</td><td>1</td><td>126.46</td><td>&lt;0.001</td></tr><tr><td>Gradient × Species</td><td>1</td><td>15.00</td><td>&lt;0.001</td></tr></tbody></table>

Note: See Figs. 2, 5 for graphical representations.

plant species in heathlands, which may play a key role in their N acquisition and success in these acidic environments. In particular, our results show a strong BDI with high levels of root procyanidins and a low DEA/SIR ratio for *Erica cinerea*, and a biological control of both the nitrification and denitrification by *Pteridium aquilinum* that efficiently improves NO₃⁻ availability for this plant species in the most acid sites (Fig. 6). This may explain the invasion capacity of *P. aquilinum* in nutrient-poor, acid soils dominated by *E. cinerea*.

### Differences in the soil NO₃⁻ under P. aquilinum and E. cinerea are not explained by changes in NH₄⁺ and total N content

In the sites with lowest soil pH values, *P. aquilinum* was associated with a higher soil NO₃⁻ concentration than *E. cinerea* and lower NH₄⁺ concentration, which is consistent with previous observations at different heathland sites by Mitchell et al. (1999) and DeLuca et al. (2013). The variations in soil NO₃⁻ concentration observed between the two species were not explained by variations in the total soil N content, which was roughly constant across all sites. In addition, changes in NO₃⁻ concentration were not correlated with changes in NH₄⁺ concentration or changes in SIR, and an increase in mineralization cannot thus explain the higher NO₃⁻ concentration for *P. aquilinum*, as previously suggested (Marrs et al. 1992, Mitchell et al. 1997, DeLuca et al. 2013). Our results actually suggest that changes in soil NO₃⁻ and NH₄⁺ concentrations are driven by several controls acting on nitrification and denitrification depending on the plant species and that *P. aquilinum* has developed a biocontrol strategy of these N cycle processes, which is more efficient than the strategy developed by *E. cinerea* to enhance soil NO₃⁻ availability.

### Effect of P. aquilinum and E. cinerea on soil microbial processes

The DEA/SIR ratio was lower in *E. cinerea* than *P. aquilinum* soils. The DEA/SIR ratio is a useful tool to assess biological denitrification inhibition (BDI; Bardon et al. 2014). Indeed, reduction in the DEA/SIR ratio reflects a specific decrease in the activity of denitrifiers as compared to the broad activity of the whole heterotrophic
<figure><img src="image_4.png" /></figure>

Fig. 2. (A) Denitrifying enzyme activity, DEA, and (B) nitrifying enzyme activity, NEA, in soils from *Erica cinerea* (black) and *Pteridium aquilinum* (gray) stands for the five study sites. Vertical bars indicate standard errors (n = 8). Stars indicate significant differences between species among sites (P < 0.05). See Table 2 for results of statistical analyses. Panels on the right present the difference in soil (C) DEA and (D) NEA observed for *P. aquilinum* as compared to *E. cinerea* along the pH gradient.

community (Schimann et al. 2012, Bardon et al. 2014). Our results thus indicate that *E. cinerea* reduces more efficiently denitrification than *P. aquilinum*. This is consistent with the higher concentration of root procyanidins observed for *E. cinerea* than *P. aquilinum*. Our tests in vitro show that root procyanidins produced by *P. aquilinum* allow inhibition of denitrification by this species, but our results from in situ measurements show that this inhibition may be less important than for *E. cinerea*. In addition, soils under *P. aquilinum* had a higher NEA than from under *E. cinerea* in sites with lowest soil pH values, and soil $NO_3^-$ was correlated with NEA for *P. aquilinum*, but not for *E. cinerea*. Overall, our results thus suggest that the higher soil $NO_3^-$ observed for *P. aquilinum* in particular in the most acidic soils could be explained by a

<figure><img src="image_5.png" /></figure>

Fig. 3. Relationship between soil $NO_3^-$ concentration and NEA for *Erica cinerea* (black dots) and *Pteridium aquilinum* (white triangles) stands. ***P < 0.001.
<table><caption>Table 3. Mixed model between NO₃⁻ and NEA, NEA and pH, NEA and NH₄⁺, DEA and procyanidins, and NO₃⁻ and procyanidins, with site as a random factor and species and the interaction of explanatory factors with species as co-factors.</caption><thead><tr><th>Variable</th><th>df</th><th>F</th><th>P</th></tr></thead><tbody><tr><td>(a) NO<sub>3</sub><sup>-</sup></td><td></td><td></td><td></td></tr><tr><td>NEA</td><td>1</td><td>16.97</td><td>&lt;0.001</td></tr><tr><td>Species</td><td>1</td><td>8.52</td><td>&lt;0.01</td></tr><tr><td>NEA × Species</td><td>1</td><td>11.93</td><td>&lt;0.001</td></tr><tr><td>(b) NEA</td><td></td><td></td><td></td></tr><tr><td>pH</td><td>1</td><td>21.18</td><td>&lt;0.001</td></tr><tr><td>Species</td><td>1</td><td>5.70</td><td>&lt;0.05</td></tr><tr><td>pH × Species</td><td>1</td><td>4.03</td><td>0.09</td></tr><tr><td>(c) NEA</td><td></td><td></td><td></td></tr><tr><td>NH<sub>4</sub><sup>+</sup></td><td>1</td><td>1.99</td><td>0.17</td></tr><tr><td>Species</td><td>1</td><td>5.50</td><td>&lt;0.05</td></tr><tr><td>NH<sub>4</sub><sup>+</sup> × Species</td><td>1</td><td>3.66</td><td>0.06</td></tr><tr><td>(d) DEA</td><td></td><td></td><td></td></tr><tr><td>Procyanidin</td><td>1</td><td>0.76</td><td>0.38</td></tr><tr><td>Species</td><td>1</td><td>6.30</td><td>&lt;0.05</td></tr><tr><td>Procyanidin × Species</td><td>1</td><td>1.42</td><td>0.23</td></tr><tr><td>(e) NO<sub>3</sub><sup>-</sup></td><td></td><td></td><td></td></tr><tr><td>Procyanidin</td><td>1</td><td>0.001</td><td>0.99</td></tr><tr><td>Species</td><td>1</td><td>6.59</td><td>&lt;0.05</td></tr><tr><td>Procyanidin × Species</td><td>1</td><td>11.12</td><td>&lt;0.001</td></tr></tbody></table>

synergistic effect of both nitrification stimulation and BDI.

Some plant species are known to stimulate nitrification (Lata et al. 2000, Le Roux et al. 2003,

Hawkes et al. 2005). DeLuca et al. (2013) observed that soils from *P. aquilinum* showed higher net nitrification than soils from heathland dominated by *Calluna vulgaris*. They concluded that *P. aquilinum* could stimulate nitrification through an increase in N mineralization and thus soil NH₄⁺ availability. However, in our study, the NH₄⁺ concentration was not higher in soils under *P. aquilinum* (it was actually even lower than for *E. cinerea* in site 5). Moreover, no correlation was observed between NEA and soil NH₄⁺ for both species. In addition, nitrification is sensitive to decreasing soil pH (De Boer et al. 1990, Yao et al. 2011), and the soil pH was lower in soils from *P. aquilinum* than *E. cinerea*. Thus, higher NEA and soil NO₃⁻ in *P. aquilinum* were likely explained by plant-dependent biological controls. Because it is known that nitrification contributes to decrease soil pH, particularly in heathland (De Boer et al. 1990, Yao et al. 2011), the lower pH in *P. aquilinum* stands was probably more a consequence than a driver of higher nitrification.

To sum up, the increase in soil nitrification observed under *P. aquilinum* could not be explained by the main abiotic factors (pH and NH₄⁺) known to control nitrification. Our results suggest that *E. cinerea* induced BDI more efficiently than *P. aquilinum*, while *P. aquilinum* also promoted an increase in soil nitrification.

<figure><img src="image_6.png" /><figcaption>Fig. 4. Relationships between nitrifying enzyme activity, NEA, and soil (left) pH and (right) ammonium, for *Erica cinerea* (black dots) and *Pteridium aquilinum* (white triangles) stands. The full line indicates that a significant relationship was observed.</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>Fig. 5. (A) Procyanidin concentrations in roots of *E. cinerea* (black) and *Pteridium aquilinum* (gray) for the five study sites, (B) difference in root procyanidin concentration observed for *P. aquilinum* as compared to *Erica cinerea* along the pH gradient, and (C) level of the biological denitrification inhibition, BDI, induced by *E. cinerea* and *P. aquilinum* root extracts on the denitrification activity of *Pseudomonas brassicacearum*. Vertical bars indicate standard errors (n = 4). Stars indicate significant differences between species among sites. Letters indicate significant differences between treatments (*P* < 0.05).</figcaption></figure>

E. cinerea and to a lesser extent P. aquilinum influence denitrification and soil NO₃⁻ concentrations through the production of procyanidins

Bardon et al. (2016) recently demonstrated that soil denitrification is reduced by some plant species through the release of procyanidins by their roots. In this study, we could not detect procyanidins in soil samples. Indeed, procyanidins strongly bind to soil components and are rapidly oxidized, making soil inputs particularly difficult to quantify (Kraus et al. 2004b, Norris et al. 2011). However, because of their water solubility, it is admitted that they are released by roots (Kraus et al. 2003, 2004a). Moreover, Meier et al. (2008) demonstrated that roots are the dominant source of labile phenolics to soils during the growing season. For these reasons, we used the root procyanidin concentration as a proxy of the amount released in soils. Here, we observed a lower concentration of procyanidins in *P. aquilinum* than in *E. cinerea* roots. While the procyanidins found in both species were based on the same structure (monomers catechin and epicatechin), *E. cinerea* procyanidins showed a higher degree of polymerization (dimeric to tetrameric) and A-type bonds. *P. aquilinum* showed only B-type bonds between monomers and only dimeric procyanidins (Appendix S1). In addition to the quantity of procyanidins produced by roots, the differences in procyanidin structure between the two plants could explain their different effects on denitrification. Indeed, variations in procyanidin structure greatly influence their biological activity (Norris et al. 2011). However, when applied at the same concentrations and tested on *Pseudomonas brassicacearum*, procyanidins extracted from roots of both species did not differ in BDI efficiency. These results suggest that *E. cinerea* inhibits denitrification more than *P. aquilinum*, mainly because of a higher procyanidin production, which is consistent with the lower DEA/SIR ratio observed for this species.

In contrast, soil NO₃⁻ concentration was positively correlated with the procyanidin concentration in roots for both species. This suggests that for each species, the BDI level is directly correlated with the procyanidin production level. However, the relationship between soil NO₃⁻ and procyanidins differed between the two species, *P. aquilinum* producing less procyanidins
<figure><img src="image_8.png" /><figcaption>Fig. 6. Schematic summary of the biological controls that *Erica cinerea* and *Pteridium aquilinum* exert on N cycling processes in acid soils, leading to changed availability in soil ammonium, NH₄⁺, and nitrate, NO₃⁻.</figcaption></figure>

than *E. cinerea*, but influencing NO₃⁻ concentration more efficiently in the most acid soils. This is likely due to the synergistic effect of both an increase in nitrification and BDI in *P. aquilinum* stands, while *E. cinerea* biological control strategy on N cycle processes would mostly rely on BDI.

## CONCLUSION

Our results demonstrate that both *Pteridium aquilinum* and *Erica cinerea* have a biological control strategy concerning soil N cycle processes related to NO₃⁻ availability. In particular, both *P. aquilinum* and *E. cinerea* induce a biological denitrification inhibition, but *E. cinerea* inhibits more denitrification than *P. aquilinum*, mainly due to a higher procyanidin production.

However, *P. aquilinum* also influences soil N cycling by stimulating nitrification, in particular in low pH soils, and the conjunction of nitrification stimulation and denitrification inhibition explains the higher NO₃⁻ availability observed for *P. aquilinum* compared to *E. cinerea*. This may partly explain the success of *P. aquilinum* on acid soils and its invasion capacity in Atlantic heathlands. In the context of the management of heathlands, our results may help exploring new approaches for bracken control which would better account for belowground processes, for example, exploring the possible role of nitrification inhibitors to counteract the bracken strategy for controlling soil N cycling. Further, we suggest that all evaluations of bracken control strategies should include characterization of key soil
properties, in particular nitrate/ammonium and pH. This may help understanding why the effectiveness of bracken control in heathlands varies between locations. More generally, our results highlight the role that BDI, through procyanidin production by roots, can play to help plants increasing $NO_3^-$ availability in soil, which had not been studied before. As procyanidins are produced by many plant species (Dixon et al. 2005), BDI could be a widespread phenomenon that should be taken into account when studying soil $NO_3^-$ dynamics, particularly in N-poor environments.

## ACKNOWLEDGMENTS

We would like to thank the CNRS INEE Department and the INRA EFPA Department for providing funds for this work. Denitrification and $NO_3/NO_2$ analyses were performed at the AME platform (UMR 5557, FR 41, University Lyon1). The authors would also like to thank Dr. Laurent Simon for elemental analysis performed on the "Ecologie Isotopique" platform (UMR5023, FR41, University Lyon1). The procyanidins were purified at the CESN platform (UMR5557, University Lyon1).

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## SUPPORTING INFORMATION

Additional Supporting Information may be found online at: http://onlinelibrary.wiley.com/doi/10.1002/ecs2.2426/full