Plant and Soil

, Volume 359, Issue 1–2, pp 297–319 | Cite as

Microarray analysis of humic acid effects on Brassica napus growth: Involvement of N, C and S metabolisms

  • Laëtitia Jannin
  • Mustapha Arkoun
  • Alain Ourry
  • Philippe Laîné
  • Didier Goux
  • Maria Garnica
  • Marta Fuentes
  • Sara San Francisco
  • Roberto Baigorri
  • Florence Cruz
  • Fabrice Houdusse
  • José-Maria Garcia-Mina
  • Jean-Claude Yvin
  • Philippe Etienne
Regular Article


Background & aims

Winter rapeseed (Brassica napus) is characterized by a low N recovery in seeds and requires high rates of fertilization to maintain yield. Its nutrient use efficiency could be improved by addition of a biostimulant such as humic acids whose physiological effects have been described previously in some plant species. However, to our knowledge, no study has focused on transcriptomic analyses to determine metabolic targets of this extract.


A preliminary screening of ten humic acids revealed a significant effect of one of them (HA7) on rapeseed root growth. Microarray analysis was then used on HA7-treated or non-treated plants to characterize changes in gene expression that were further supported by physiological evidence.


Stimulation of nitrogen uptake (+15% in shoots and +108% in roots) and assimilation was found to be increased in a similar manner to growth while sulfate content (+76% in shoots and +137% in roots) was more strongly stimulated leading to higher sulfate accumulation. In parallel, microscopic analysis showed an enhancement of chloroplast number per cell.


It is therefore suggested that HA7, which promotes plant growth and nutrient uptake, could be used as a supplementary tool to improve rapeseed nitrogen use efficiency.


Brassica napus Humic acid Microarray Growth promotion Nutrient uptake Chloroplast 


Any improvement in agricultural practices that increases plant nutrients capture efficiency should reduce the negative environmental impacts of agriculture and increase crop production and sustainability in reduced input systems. Thus, many approaches have been studied to increase nutrients capture and yield, such as genetic selection which has included allele selection, selection of domestication genes, gene and genome duplication, new genotypes creation and QTLs. (for a review, see Vaughan et al.2007). More recently, new strategies such as the use of biological molecules that act as biostimulants have been evaluated. As defined by Zhang and Schmidt (1997), biostimulants correspond to “materials, other than fertilizers, that promote plant growth when they are applied in small quantities”. Among these biostimulants, bioactive substances extracted from humic acids are the most studied.

Humic substances are ubiquitous organic materials in terrestrial and aquatic ecosystems. They are collections of diverse low molecular mass components forming dynamic associations that are stabilized by hydrophobic interactions and hydrogen bonds (Conte and Piccolo 1999; Smejkalova and Piccolo 2008). These associations are capable of organizing into micellar structures in suitable aqueous environments (Varga et al.2000; Sutton and Sposito 2005). The predominant components of humic substances are humic acids, which are very active in interacting with organic and inorganic contaminants. Kozuch and Pempkowiak (1996), Christl and Kretzschmar (2001), Muscolo et al. (2007a) and Canellas et al. (2010) show that biological activity of the humic fraction depends on their chemical structure and molecular weight. These authors demonstrate that humic acids presenting a lower molecular weight had the greatest effects on plant growth or ion binding.

The first hypothesis on humic acid action is the improvement of physical and chemical properties of soil. Cyclic wetting and drying conditions in soil result in the breakdown of large aggregates into smaller aggregates susceptible to erosion and dispersion. Authors showed that humic acid application to soil could enhance aggregate stability (Piccolo et al.1997a; Imbufe et al.2005; Abiven et al.2009), thus decreasing soil erosion (Piccolo et al.1997b) and avoiding C and N leaching (Andersson et al.2000; Bossuyt et al.2007). The capacity of humic substances to form complexes permits sequestration of nutrients (Zn, Cu, Fe for example) by formation of metal-humic complexes, thus protecting them from leaching and making them more available for plant uptake (Esparza et al.2005; Garcia-Mina 2006). In addition, several authors have shown that a humic acid application has a beneficial effect on the soil bacterial community (Cookson et al.2005; Arancon et al.2006; Puglisi et al.2009) which could be due to the enhanced secretion of organic carbon from humi acid-treated plant roots (Puglisi et al.2009). These effects on physical, biological and chemical properties of soil could create improved conditions for plant development and explain the enhanced growth observed when plant are treated with humic acids. In contrast, other authors have proposed a direct effect of humic acids on plants.

Studies on the effects of humic acids on maize have shown accelerated development cycles, i.e. earlier germination, flowering and fructification (Eyheraguibel et al.2008). Other authors have related an increase in the total dry weight, and more specifically the proliferation of secondary root systems in response to foliar spraying with humic extract on Arabidopsis thaliana (Schmidt et al.2007), barley (Ayuso et al.1996), cucumber (Atiyeh et al.2002; Mora et al.2010), and maize (Canellas et al.2002; Zandonadi et al.2007). In parallel, enhancement of leaf chlorophyll content has also been reported in rice (Tejada and Gonzalez 2004). This better development and increase in crop growth led to an increase in yield in pepper, rice, wheat and rapeseed when treated by foliar spraying with humic acids (Keeling et al.2003; Tejada and Gonzalez 2004; Arancon et al.2005). In addition, other studies have revealed that humic acids application enhances plant nutrient uptake, especially N (Dell’Agnola and Nardi 1987; Pinton et al.1999; Nardi et al.2000b; Keeling et al.2003) associated with an increased expression of genes involved in nitrate transport (Quaggiotti et al.2004). Accumulation of iron (Cesco et al.2002) and other micronutrients (Ayuso et al.1996; Garcia-Mina et al.2004; Eyheraguibel et al.2008) in plants was also reported. All these authors suggest that humic acid could have a hormone-like action on plants, which would explain the effects on growth and yield. Indeed, Muscolo et al. (1998, 1999, 2007b) demonstrated the presence of auxins (IAA) in humic fractions presenting biological activity.

Focusing on plant metabolism, Nardi et al. (2000b) and Carletti et al. (2008) have shown a change in maize protein synthesis in response to humic acid treatment. Proteins that were impacted by humic acids were classified in: energy and metabolism, cellular transport, transport facilitation and transport routes, interaction with the environment, signaling, defense and cell rescue (Carletti et al.2008). In addition, Muscolo et al. (1996) demonstrated that a humic acid application on Daucus carota enhance enzymatic activity of glutamate dehydrogenase, glutamine synthetase and phosphoenolpyruvate carboxylase. In the same way, Nardi et al. (2002a, 2007) have shown the activation of N uptake and assimilation, the glycolytic pathway and the Krebs cycle in maize seedlings treated with humic extracts. According to these studies, modification of plant nutrition promoted by humic extract could be due to its hormone-like action. Moreover, Nardi et al. (2000a, 2002b) have shown that humic substances extracted from plant root exudates possesses a higher biological activity than water-extracted humic substances, probably because root exudate-extracted humic substances contained a higher content of phytohormones (Nardi et al.2000c) and a lower molecular size (Nardi et al.2005).

Winter oilseed rape (WOSR, Brassica napus L.) is an important agricultural crop cultivated for its oil, which can be used as an edible product or for industrial applications (animal nutrition, cosmetics, diester production…). However, oilseed rape shows a low Nitrogen Use Efficiency (NUE, defined by the ratio of seed N content to total plant N content) especially due to the default in leaf N mobilization (Etienne et al.2007; Desclos et al.2008) during the vegetative stage. As a consequence, N remaining in fallen leaves is a loss for dry matter production but also increases the risk of nitrate leaching following the mineralization of leaf organic N. Indeed, the fall of WOSR leaves with high N content (up to 2% of the dry matter, Malagoli et al.2005) leads to a return of N to the soil that can reach 100 kg N ha−1 (Dejoux et al.2000). Due to this low NUE, WOSR requires excessive N fertilization to maintain high N content of harvested tissues. Whatever the rate of N fertilization, the oilseed rape N Harvest Index (NHI) is low (approximately 70%, Dreccer et al.2000) compared with cereals. Thus, the aim of the present study was to identify new humic acids that could increase the mineral nutrition and growth of WOSR so as to reduce the dose of fertilizer currently used.

In a first step, we have screened ten humic acids resulting from different extraction processes for their effects on rapeseed. For this, easily and rapidly measurable parameters such as improvement of dry weight production (roots and/or shoots), chlorophyll content or improvement of Nitrogen Use Efficiency have been used. From this basic screening, only one extract, named HA7, led to significantly higher dry weight and chlorophyll content of rapeseed after 30 days of supply to the roots. In order to better understand the HA7 effects on WOSR growth, a detailed characterization (elementary and hormonal composition) of this humic acid has been performed and coupled with transcriptomic (microarray), physiological, biochemical and light and electron microscopic analyses. This approach allowed the identification of specific genes or groups of genes that were up- or down-regulated when WOSR was treated with HA7. Furthermore, and to the best of our knowledge, this study is one of the pioneer works (Trevisan et al.2011, on Arabidopsis thaliana) that has used a specific Brassica napus microarray associated with a physiological approach to identify the main the metabolic pathways induced by humic acid.

Materials and methods

Growth conditions and experimental design

Seeds of Brassica napus var. Capitol were surface-sterilized by exposure to 80% ethanol for 30s followed by 20% sodium hypochlorite for 10 min. After 10 washes with demineralized water, seeds were germinated on perlite over demineralized water for 2 d in the dark and 1 week in the light in a greenhouse. Just after first leaf emergence, seedlings were transferred to a 20 L plastic tank containing Hoagland nutrient solution with the following composition: KNO3 1 mM, CaCl2 1.25 mM, KCl 250, KH2PO4 0.25 mM, MgSO4 0.5 mM, EDTA-2NaFe 0.2 mM, H3BO3 14 μM, MnSO4 5 μM, ZnSO4 3 μM, CuSO4 0.7 μM, (NH4)6Mo7024 0.7 μM and CoCl2 0.1 μM. This nutrient solution was renewed every two days. Plants were grown under greenhouse conditions with a thermoperiod of 20°C/17°C day/night and a photoperiod of 16 h. Natural light was supplemented with high pressure sodium lamps (Philips, MASTER Green Power T400W) supplying an average photosynthetically active radiation of 280 μmol photons.m-2.s−1 at canopy height. Plants were grown for one week before treatment with HA7 humic extract.

Humic acid (HA7) sample preparation and characterization

Method of preparation of the humic acid (HA7) sample

The humic sample (HA7) was principally obtained from black peat (Galicia, Spain). The methodology for the extraction and purification used was as indicated by the International Humic Substances Society (IHSS) (Swift 1996), and described in Aguirre et al. (2009).

Briefly, 100 g of potassium humate were mixed with 1500 ml of 0.1 M NaOH in a 2 L flask. After 48 h of stirring at 25°C in darkness, the supernatant containing the unfractionated humic extract was separated from the solid fraction by centrifugation (15 min, 11100 g). The alkaline extract sample, containing humic acids and fulvic substances was acidified with 6 M hydrochloric acid (HCl) to a pH of 1.5. After 12 h, the acidified sample was centrifuged (30 min, 7650 g) in order to separate the precipitated HA7 from the supernatant containing fulvic acids and other acid-soluble organic compounds. After washing with water to eliminate Cl contamination, HA7 was freeze-dried. Dry HA7 contains 39.65% of organic carbon and before use, it was dissolved in water in order to reach a concentration of 2.6% of organic carbon.

13C RMN characterization of HA7

Solid-state 13C RMN spectra were obtained on a Bruker Avance AV-400WB (9.4T) spectrometer at 100.47 MHz using the cross-polarization magic angle spinning technique (CPMAS), with a spinning speed of 12 kHz, 90° pulse width, 30 ms acquisition time and 4.0 s delay.

HPSEC characterization of HA7

The molecular size distribution was evaluated by high performance size exclusion chromatography. The chromatographic system consisted of a Waters 600 Controller pump followed by two detectors in series: a Waters 996 Photodiode Array Detector set at 400 nm, and a Waters 2424 Refractive Index Detector (RI). Size exclusion separation occurred through a PL Aquagel-OH 30 column (Polymer Laboratories), followed by a guard column with the same stationary phase. The overall molecular weight range of separation for this column was 100–300,000 Da.

For each sample, solutions of 800 ppm of carbon were prepared in 0.05 M NaNO3. The injection volume of all samples was 100 μl, the eluent used was 0.05 M NaNO3 (pH 7), and the flow rate was 1 ml/min. Void volume (V0 = 6.65 ml) and permeation volume (Vp = 11.82 ml) were determined with polyethylene oxide of MW of 43.250 Da and methanol, respectively.

In order to evaluate an approximate MW distribution from HPSEc chromatograms for humic samples, a universal calibration was carried out. Curves of log J vs. elution volume were obtained using polyethylene glycol and polyethylene oxide standards of known MW. The parameter J is defined as the product of the intrinsic viscosity [η] and the molecular weight (Ji = [η]IMi), and it is proportional to the hydrodynamic volume. This means that two macromolecules with the same hydrodynamic volume will have equal J values.

The Mark-Houwin-Sakurada equation relates [η] to MW as follows:
$$ \left[ \eta \right] = K \cdot {{M}^a} $$
where K and a are proper constants for each macromolecule, solvent and temperature. In this study we have used the values of K and a reported by Visser (1985) for a soil humic acid (K = 2.724 × 10−2 mL g−1 and a = 0.45).

Analysis of the concentration of IAA, ABA and Cytokinins in HA7

The general method is extensively described in Aguirre et al. (2009) and Mora et al. (2010). As an example we describe the method used for cytokinins (CKs) extraction and quantification.

CKs Extraction, purification and determination procedures cytokinins (CKs) by HPLC MS/MS: Endogenous CKs were extracted and purified following the procedure described by Dobrev and Kamiek (2002) with slight modifications. Deuterated CKs (2H5-trans-zeatin (d-tZ), 2H5-trans-zeatin riboside (d-tZR), 2H6-isopentenyladenine (d-iP), 2H6-N6-isopentenyladenosine (d-iPR) (OlChemin Ltd, Olomouc, Czech Republic)) were added as internal standard. About 0.5 g fresh weight of plant tissue was homogenized in liquid N with 5 mL of cold (−20°C) extraction mixture of methanol/water/formic acid (15/4/1, v/v/v) containing 40 μL of a mixture of deuterated internal standards (50 ng.mL−1). After overnight extraction at −20°C, solids were separated by centrifugation (10 min, 12000 g). The pellet was re-extracted for 60 min with 4 mL of extraction mixture at −20°C. Pooled supernatants were purified using Strata C18-E cartridge, pre-conditioned with 4 mL of methanol and 2 mL of methanol/water/formic acid (15/4/1, v/v/v). The eluted fraction was evaporated until methanol is removed (Vortex evaporator mod. 432–2100 from Labconco Corporation (Kansas City, MO, USA)). The residue was re-dissolved in 2 mL of 1 M formic acid and applied to an Oasis MCX column pre-conditioned with 4 mL of methanol followed by 2 mL of 1 M formic acid. The column was washed with 2 mL of 1 M formic acid followed by 2 mL of methanol and 2 mL of NH4OH (0.35 M in water). CKs bases and ribosides were eluted with 2 mL of NH4OH 0.35 M in 60% (v/v) of methanol before evaporation to dryness at 40°C. The residue was dissolved with 250 μL of methanol/formic acid at 0.05% in water (40/60, v/v) and centrifuged (5 min, 10000 g).

Liquid Chromatography-Mass Spectrometry quantification of cytokinins: CKs were quantified by HPLC linked to a 3200 Q TRAP LC/MS/MS system (Applied Biosystems/MDS Sciex, Ontario, Canada), equipped with an electrospray interface, using a reverse-phase column (Synergi 4 μ Hydro-RP 80A, 150 × 2 mm, Phenomenex, Torrance, CA). A linear gradient of methanol (A) and 0.5% acetic acid in water (B) was used: 35% A for 1 min, 35% to 95% A in 9 min, 95% A for 4 min and 95% to 35% A for 1 min, followed by a stabilization time of 5 min. The flow rate was 0.20 mL.min−1, the injection volume was 50 μL and the column and sample temperatures were 30°C and 20°C, respectively.

The detection and quantification were performed by multiple-reaction monitoring (MRM) in the negative-ion mode, employing a multilevel calibration graph with deuterated hormone as internal standard. The monitored fragmentation processes are described in Aguirre et al. (2009).

The source parameters were curtain gas: 25.0 psi, GS1: 50.0 psi, GS2: 60.0 psi, ion spray voltage: −4000 V, CAD gas: medium, and temperature: 600°C.

Plant treatment

After one week of growth, plants were separated in two sets: (i) control plants (control) were grown in the Hoagland solution described above with 15 N labeling (3% atom excess) and (ii) treated plants (HA7) grown in the same labeled solution supplied with HA7 humic acid at 100 mg of organic carbon L-1. Nutrient solutions were renewed every two days. Each set of plants (control and HA7) was grown for 30 days and time courses in the experiment were expressed in days after treatment (Day 0, addition of HA7 to the nutrient solution).

For each treatment (Control and HA7) and each time of harvest (1, 3 and 30 days), ten plants (i.e. ten plants pooled in 3 replicates) were harvested and separated from shoot and root samples. Thereafter, roots and shoots were frozen in liquid nitrogen and stored at −80°C for further analysis. An aliquot of each tissue was weighed and dried (60°C) in a drying oven for dry weight (DW) determination and ground to fine powder for mineral and ion analysis. Likewise, at each time of harvest, a fresh aliquot of shoots and roots was used for real-time in vivo nitrate reductase activity and RNA extraction.

Determinations of chlorophyll concentration and net photosynthetic rates

During the experiment, determinations of chlorophyll concentration and net photosynthetic rates were monitored at 1 d and 3 d after HA7 application and then weekly until the end of the treatment (i.e. 30 days). Determination of relative chlorophyll concentration was performed using a non-destructive method: a SPAD (Soil Plant Analysis Development) chlorophyll meter (Minolta, SPAD-502 model, Tokyo, Japan). The determination was carried out on 10 measures per leaf and on 5 leaves per replicate. Moreover, the net photosynthetic rate was measured using a LI-6400 photosynthesis system (LICOR, Lincoln, Nebraska, USA) at 23°C on leaves under ambient CO2 atmospheric concentration (~ 400 ppm) and at 1000 μmol.m−2.s−1 photosynthetic photon flux (PPF) provided by a LED light. Temperature and PPF parameters were previously validated as the optimal values for photosynthesis analysis in leaves of oilseed rape.

Total N and 15N analysis

An aliquot of each plant organ (shoots or roots) was placed in thin capsules for isotopic analysis in order to analyze between 60 and 80 μg of N. The total N amount and 15 N excess in plant samples were determined with a continuous flow isotope mass spectrometer (Isoprime, GV Instrument, Manchester, UK) linked to a C/N/S analyzer (EA3000, Euro Vector, Milan, Italy):
  • Total N (Ntot) content in a tissue ‘i’ at a given time was calculated as:
    $$ {\text{Nto}}{{\text{t}}_{\text{i}}} = \left( {\% {{\text{N}}_{\text{i}}} \cdot \times \cdot {\text{D}}{{\text{W}}_{\text{i}}}} \right)/100 $$
  • The natural 15N abundance (0.36636 ± 0.0004%) of atmospheric N2 was used as a reference for 15N analysis. Nitrogen derived from current N uptake (Nupti) in a given organ, at a given time, was calculated as:
    $$ {\text{Nup}}{{\text{t}}_{\text{i}}} = \left( {{\text{Nto}}{{\text{t}}_{\text{i}}} \cdot \times \cdot {{\text{E}}_{\text{i}}}} \right)/{{\text{E}}_{\text{s}}} $$
    with Ei (%) is the atom 15N excess in shoots or roots, Es is the nutrient solution atom 15N excess (3%).

Nutrient composition analysis

Except for N whose determination method was described in previous paragraph, the total nutrient concentration in rapeseed shoots and roots, in HA7 extract or in Hoagland nutrient solution was measured by ICP-OES (Thermo Elemental Co. Iris Intrepid II XDL) with prior microwave acid sample digestion (8 mL of concentrated HNO3 and 2 mL of H2O2 for 0.5 g DW) using protocol previously described in Mora et al. (2010).

Nitrate and sulfate analysis

Nitrate and sulfate were extracted and analyzed as previously described (Abdallah et al.2011) from 30 mg of DW (shoots or roots) in 1.5 ml of 50% ethanol solution at 40°C for 1 h. After centrifugation (20 min, 10 000 g) the supernatant (called S1) was recovered and 1.5 ml of 50% ethanol was added to the pellet. After a second incubation (1 h, 40°C) and centrifugation (20 min, 10 000 g), the remaining supernatant was taken up and added to the previous supernatant (S1). All these operations (i.e. incubation and centrifugation) were repeated twice but now with 1.5 ml of ultra-pure water and incubation at 95°C. All supernatants were finally pooled then dried under vacuum for 16 h without heating (Concentrator Evaporator RC 10.22 Jouan, Saint Herblain, France). The dry residues containing both nitrate and sulfate were solubilized in 1 ml of ultra-pure water. Thereafter, nitrate and sulfate concentrations in each tissue were determined by using ion chromatography (HPLC, ACS3000, Dionex corp. Sunnyvale, CA, USA), with a conductivity detector. The eluent solution consisted of 1.8 mM Na2CO3 and 1.7 mM Na2HCO3, and was pumped isocratically over an AS17 guard column.

In vivo nitrate reductase activity in plants

The nitrate-reductase (NR) activity was determined in each replicate using an in vivo assay adapted from Bungard et al. (1999). Shoot or root tissue (0.1 g FW) was vacuum infiltrated for 3x 30 s with 10 mL of phosphate buffer (pH 7.5) containing 1% (v/v) propanol and 1 M KNO3. After vacuum infiltration, buffer solutions containing plant material were separated in two sets. One part was incubated in a shaking water bath at 30°C for 15 min in darkness and then boiled to stop any enzymatic activity. The other part was boiled immediately after vacuum infiltration for initial nitrite concentration determination. The colorimetric reaction was performed with 1 mL of plant extract, 1 mL of 0.3% (w/v) sulfanilic acid in 30% acetic acid and 1 mL of 0.05% (w/v) ά-naphtalamin in 30% acetic acid. The amount of nitrite (NO2) formed in each buffer solution was measured spectrophotometrically (UV-1601, UV-visible spectrophotometer, Shimadzu, Champs-sur-Marne, France) at 540 nm. Thereafter, NR activity was calculated as μmol NO3- reduced per g FW and per hour.

RNA extraction

Total RNA was extracted from 200 mg of root and shoot FW. Frozen samples were ground to a powder with a pestle in a mortar containing liquid nitrogen. The resulting powder was suspended in 750 μl of extraction buffer [0.1 M TRIS, 0.1 M LiCl, 0.01 M EDTA, 1% SDS (w/v), pH 8] and 750 μl of hot phenol (80°C, pH 4). This mixture was vortexed for 30 s and, after addition of 750 μl of chloroform/isoamylalcohol (24:1), the homogenate was centrifuged at 15 000 g (5 min, 4°C). The supernatant was transferred into 4 M LiCl solution (w/v) and incubated overnight at 4°C. After centrifugation (15 000 g, 30 min, 4°C), the pellet was suspended in 100 μl of sterile water. RNA was then purified with an RNeasy mini kit according to the manufacturer’s protocol (Qiagen, Courtaboeuf, France). Quantification of total RNA was performed by spectrophotometry at 260 nm (BioPhotometer, Eppendorf, Le Pecq, France) before Reverse Transcription (RT) and real-time PCR (q-PCR) analysis.

Reverse transcription (RT) and q-PCR analysis

For RT, 1 μg of total RNA was converted to cDNA with an ‘iScript cDNA synthesis kit’ according to the manufacturer’s protocol (Bio-Rad, Marne-la-Coquette, France).

Q-PCR amplifications were performed using specific primers for each housekeeping gene (EF1-ά [forward 5′-gcctggtatggttgtgacct-3′ and reverse 5′-gaagttagcagcacccttgg-3′] and 18S rRNA [forward 5′-cggataaccgtagtaattctag-3′ and reverse 5′-gtactcattccaattaccagac-3′]) and target gene: BnNRT1.1 forward 5′-tggtggaataggcggctcgagttg-3′ and reverse 5′- gtatacgttttgggtcattgccat-3′, BnNRT2.1 forward 5′-atggtaacggaagtgccttg-3′ and reverse 5′- tgattcgagctgtgtgaagc-3′, BnSultr1.1 forward 5′-agatattgcgatcggaccag-3′ and reverse 5′- gaaaacgccagcaaagaaag-3′, BnSultr1.2 forward 5′-ggtgtagttcgctggaatggt-3′ and reverse 5′-aacggagtgaggaagagcaa-3′, BnSultr4.1 forward 5′-gaccagacccgttaaggtca-3′ and reverse 5′- ttggaatccatgtgaagcaa-3′, BnSultr4.2 forward 5′-agcaagatcagggattgtgg-3′ and reverse 5′- tgcaacatttgtgggtgtct-3′. Q-PCRs were performed with 4 μl of 100× diluted cDNA, 500 nM of primers, and 1x SYBR Green PCR Master Mix (Bio-Rad) in a ChromoFour System (Bio-Rad). For each pair of primers, a threshold value and PCR efficiency have been determined using a cDNA preparation diluted >10-fold. For all pairs of primers, PCR efficiency was c.a. 100%. The specificity of PCR amplification was examined by monitoring the presence of the single peak in the melting curves after q-PCRs and by sequencing the q-PCR product to confirm that the correct amplicons were produced from each pair of primers (Biofidal). For each sample, the subsequent q-PCRs were performed in triplicate. The relative expression of the genes in each sample was compared with the control sample (corresponding to untreated plants at the same time of harvest) and was determined with the delta delta Ct (ΔΔCt) method using the following equation (Livak and Schmittgen 2001):
$$ \matrix{{*{20}{c}} {{\text{Relative}}\,{\text{expression}} = {{{2}}^{{ - \left[ {\Delta {\text{Ct}}\,{\text{treated}} - \Delta {\text{Ct}}\,{\text{control}}} \right]}}},} \hfill \\ {{\text{with}}\,\Delta {\text{Ct}} = {\text{C}}{{\text{t}}_{{{\text{target}}\,{\text{gene}}}}}--\left[ {\surd \left( {{\text{C}}{{\text{t}}_{{{\text{EF}}1 - \mathop{a}\limits^{\prime } }}} \times {\text{C}}{{\text{t}}_{\text{18S}}}} \right)} \right]} \hfill \\ } $$
Where Ct refers to the threshold cycle determined for each gene in the exponential phase of PCR amplification and [√(CtEF1-ά x Ct18S)] corresponds to the geometric average of Ct of reference genes. Using this analysis method, relative expression of the different genes in the control sample of the experiment was equal to 1 (Livak and Schmittgen 2001), and the relative expression of other treatments was then compared with the control.

Microarray analysis

Briefly, each test sample was hybridized on a Brassica napus Gene Expression Microarray 4 X 44 K (Agilent Technologies®) using a two-color microarray-based gene expression protocol. In this procedure, controls and treated samples (HA7) were compared and respectively labeled with cyanine 3 and cyanine 5. For each plant tissue (shoots and roots) and each time of harvest (1, 3 and 30 days), the same control was used for the different hybridizations. After the labeling step, cRNA sample sizes ranged from 50 to 3000 nucleotides. Thus, fragmentation was required to take away secondary structures (specific buffer at 60°C for 30 min) enabling cRNA lengths of between 50 to 200 nucleotides to be obtained and then optimal hybridization with an Agilent 60-mer oligonucleotide microarray to be carried out. Thereafter, hybridizations were performed at 65°C for 17 hours.

Scanning of microarrays was performed with the Agilent scanner using default parameters for 4 × 44 K formats. Data were extracted with Feature Extraction 10.1 software (Agilent Technologies®).

Fluorescence confocal microscopy and transmission electron microscopy

After 1 and 3 days of contact with HA7, microscopy observations were made on young plants with four leaves. We then chose to make observation on the last fully expended leaf: leaf #3. After, 30 days of treatment, we chose to make the observations on a leaf in the center of the plant. The first leaves that had emerged were nearly senescent and young leaves just emerged were not representative of the whole plant. Thus, among the eight leaves emerged at the end of culture (day 30), we have chosen to focus our observations on leaf #5.

A square of rapeseed leaf from each replicate of each treatment (Control and HA7-treated plant) and each time (1 d, 3 d and 30 d) was embedded in LMPT agar (low melting point) 5% in phosphate buffer at 40°C. After cooling, slices of 50 μm thickness were cut with a vibratome (Microm HM650V). Half of these slices were directly observed with confocal microscopy (Olympus FV1000) with laser excitation of autofluorescence at 458 nm and emission at 650–700 nm. The remaining slices were fixed with 2.5% glutaraldehyde in phosphate buffer 0.1 M pH 7.4 from 1 hour to several days at 4°C. The sections were rinsed in phosphate buffer 0.1 M pH 7.4 three times, post-fixed for 1.5 hours with 1% osmium tetroxide in 0.1 M phosphate buffer pH 7.4, and then rinsed in phosphate buffer three times. The cells were then dehydrated in progressive ethanol dilutions (70–100%) and 100% propylene oxide, embedded in Aradite 502 resin and polymerised for 48 h at 60°C. Ultrathin sections of 80 nm thickness were cut and contrasted with uranyle acetate and lead citrate. The sections were observed with a JEOL 1011transmission electron microscope and images were taken with a Mega View III Camera and Analysis FIVE software.

Data and statistical analysis

Regarding growth, nitrate uptake, SPAD and IRMS analysis, experiments were conducted with 10 replicates. In the case of ICP and DIONEX analysis, experiments were conducted with 6 replicates. For the microscopy, q-PCR and microarray experiments, 3 replicates were used for each sample.

All data were analyzed for significant differences by Student’s test and marked by asterisks (*: p < 0.05, **: p < 0.01 and ***: p < 0.001).


For each experiment (each time and each tissue), probes with [Marginal] flag and at least one channel above the background for the three biological replicates were retained. A t-test was applied on each filtered gene list with the following parameters: (i) t-test against zero, (ii) Benjamini-Hochberg correction and (iii) p-value < 0.05. Moreover, only genes whose expression was modified at least by a fold change of 5 (chosen as a threshold) were included in the list of differentially expressed genes.


Humic acid (HA7) characterization

As can be observed in the Fig. 1a, the HPSEC chromatogram shows four peaks for HA7 that correspond to the four groups of different molecular weight presented in Fig. 1b. The HA7 humic acid was composed of a mix of molecules ranging between 9.65·102 and 6.88·104 Da with a high representation (65%) of molecules between 1.41·103 and 5.07·103 Da. HA7 was therefore an extract with a high molecular weight. To have a better view of HA7 composition, 13 C RMN analysis was used (Fig. 2a). This characterization showed a high representation of aromatic C (42%) following by alkyl C (27.7%). To complete this analysis, biochemical characterization of HA7 was also undertaken.
Fig. 1

High Pressure Size Exclusion Chromatography analysis of humic acid HA7 a Size-exclusion HPSEC chromatogram for HA7. b Molecular weight distribution of HA7, obtain from A, expressed as the molecular weight (MW) corresponding to the maximum of the main peaks or as the MW corresponding to the interval of the whole peak

Fig. 2

13C RMN analysis of humic acid HA7 a 13C RMN spectra of HA7. b relative abundance of different carbon types (in%) determined by 13C RMN (A) for HA7

Except for C, H and O, which are the main components (2.20, 102 and 53.90 mmol.L−1 respectively), HA7 extract principally contains K, Na and S. Surprisingly, this humic acid did not contain significant N (Table 1). Concerning phytohormones, as can be observed in Table 2, no auxin (IAA) or abscissic acid (ABA) was detected in the HA7 extract. There were only very small amounts of certain cytokinins, such as Z (Zeatine), tZR (trans Zeatine), iP (isopentyladenine) and iPR (isopentyladenosine). However, these concentrations are too low to have physiological significance.
Table 1

Elemental nutrient composition of humic acid HA7 (†: determination by difference).


Concentration (mmol.L−1)


Concentration (mmol.L−1)
















53.90 †


















Table 2

Hormonal composition of humic acid extract HA7. Value are expressed in pmol.g−1. Phytohormones measured were auxin (IAA), abscisic acid (ABA) and cytokinins (Z: Zeatin, DHZ: dihydrozeatin, tZR: trans-zeatin, cZR: cis-zeatin, DHZR: dihydrozeatin riboside, iP: isopentenyladenine, iPR: isopentenyladenosine, BAR: benzyladenine riboside, mT: meta-topolin, mTR: meta-topolin riboside, oT: ortho topolin, oTR: ortho-topolin riboside). ND : Not detected

































Growth analysis

In order to determine the impact of HA7 on plant growth, the relative total dry weight (DW, defined as a percentage of control plant dry weight) was monitored after 1, 3 and 30 days of treatment (Fig. 3a). Figure 3a shows that the addition of HA7 humic acid (100 mg.L−1) in the root growth medium did not significantly affect the total DW of rapeseed after 1 or 3 days of treatment. Indeed, the total DW of control plants was 0.30 ± 0.03 g and 0.57 ± 0.02 g.plant−1 after 1 and 3 days, respectively vs 0.37 ± 0.02 g and 0.37 ± 0.04 g.plant−1 for treated plants at the same time of harvest. However after 30 days, plants treated with HA7 showed a significant increase of total DW (+29 ± 11.50%) compared to control plants (3.60 ± 0.41 g.plant−1 for control vs 4.66 ± 0.41 g.plant−1). As shown in Fig. 3b and c, this increase in total DW could be explained by a significantly increased root DW (+88 ± 19.00% compared to control, Fig. 3c) after 30 days of treatment by HA7, the shoot DW (Fig. 3b) not being significantly affected by HA7 application.At 30 days, this higher root DW than shoot DW resulted in a lower shoot/root ratio for treated plants compared to control plants (8.36 ± 1.89 and 2.84 ± 0.05 in control and treated plants respectively (Fig. 3d).
Fig. 3

Effect of humic acid on rapeseed dry weight (DW) after 1, 3 or 30 days of treatment: a Relative comparison of total dry weight (shoots + roots) of treated plants (squares and dotted line) with control (circles and continuous line), in the percentage of control plants. Values near the points are the total DW expressed in g. b Shoot DW of control (white histogram) and treated plants (hatched histogram),expressed in g.plant−1. c Root DW of control (white histogram) and treated plants (hatched histogram), expressed in g.plant−1. d Shoot/root ratio of control (circles and continuous line) and treated plants (square and dotted line). For all data, indicated values are means and vertical bars indicate ± standard deviation for n = 10 when exceeding the symbol. Significant differences at p < 0.05 and p < 0.01 are indicated by one and two asterisks, respectively

In order to check that supplemental mineral nutrients provided by the addition of HA7 was not responsible for the enhanced dry weight of treated rapeseed, the nutrient solution with or without HA7 was analyzed (Table 3). It appeared that HA7’s contribution to the mineral supply of the Hoagland solution was negligible (from 0% for Mg to +14.4% for Cu, with notable contributions of Na: +27% and Si: +57%). Furthermore, regarding plant nutrient uptake by both control and treated plants (Table 3), the fraction taken up by the plants was always lower than 15% of the total nutrient supply, whichever the nutrient was considered. Thus, even in the control solution (without HA7), plants were not exposed to any limiting conditions and the increase in dry weight of treated rapeseeds did not result from the amelioration of any kind of starvation. However, it is evident that although the percentage of nutrients taken up by plants was low, it was always higher for treated plants than for control plants.
Table 3

Cumulated nutrient supply to the plant during 30 days by Hoagland solution with or without HA7 contribution supply. The plant uptake for each nutrient was calculated from the different content between day 30 and day 0 and is expressed as percentage of cumulated nutrient available in the nutrient solution with or without HA7 during 30 days


Content in Hoagland solution

Control plant uptake (% of Control Hoagland solution)

HA7 plant uptake (% of Hoagland solution + HA7)

Control (mmol)

With HA7 (mmol)
























































Microarray data

Microarray analysis was performed with 3 replicates for each plant tissue (roots or shoots) and for each harvest time (1, 3 and 30 days). A total of 31 561 genes were analyzed on the microarray. The analysis of significantly and differentially expressed genes between control and treated plants was undertaken using a minimal fold change of expression of 5 (p-value <0.05). Using this threshold, no differentially expressed genes were found in shoots and roots after 1 day of treatment. More than 300 genes were differentially expressed after 3 days of HA7 treatment (720 genes in shoots and 366 genes in roots, Fig. 4a). 30 days of treatment with HA7 drastically lowered the number of differentially expressed genes (102 genes in shoots and no differentially expressed genes in roots, Fig. 4b).
Fig. 4

Distribution among metabolic pathways (according to DFCI annotation) of genes differentially expressed in Brassica napus roots and shoots after 3 days (a) or in shoots after 30 days (b) of treatment with HA7. Numbers in parenthesis indicated the total number of genes differentially expressed in each conditions (each time of treatment and each part of the plant, p < 0.05). Numbers near pie charts indicated the percentage of differentially expressed genes implicated in each metabolic pathways

All differentially expressed genes have been classified in metabolic pathways according to DFCI annotations (, Fig. 4). From this global classification, at first sight it appears that about 60% of the differentially expressed genes did not have a known function (supplemental data, Tables S1 and S2). However, the DFCI classification revealed that several metabolic pathways were affected by treatment (Fig. 4a and b). Among these, some metabolic pathways (such as fatty acids, phytohormones, senescence, plant development and ion transport) were represented in low numbers among the differentially expressed genes. In contrast, four metabolic pathways were more specifically affected in shoots and roots by the treatment (Fig. 4a and b): general cell metabolism (10.6% of the total differentially expressed genes on average), nitrogen and sulfur metabolism (6.6% on average), carbon metabolism and photosynthesis (6.1% on average) and stress responses (6.1% on average). Thus, according to our initial aim which was to target an improvement in nutrient use efficiencies (such as N and S) of WOSR following humic acid treatment, this study focused on N, S and C (photosynthesis) metabolisms. Accordingly, non-exhaustive lists of genes involved in these metabolisms and differentially expressed after 3 days of contact with HA7 are detailed in Table 4.
Table 4

Partial list of differentially expressed shoots and roots genes involved in photosynthesis,and nitrogen and sulfur metabolisms after 3 days of treatment with HA7 extract. The first two letters of the gene description indicates the species. (At: Arabidopsis thaliana, Bc: Brassica campestris, Bj: Brassica juncea, Bn: Brassica napus, Bo: Brassica oleracea, Br: Brassica rapa, Ca: Capsicum annuum, Gm: Glycine max, Mc: Mesembryanthemum crystallinum, Me: Manihot esculenta, Th: Thellungiella halophila, Zm: Zea mays). Positive fold change indicates that the gene is specifically over-expressed in response to humic acid (orange boxes); negative fold change indicates that the gene is specifically repressed in response to humic acid (green boxes). Boxes marked with “−” (gray boxes) indicate genes with expression levels that are not significantly different from control. P-values are Bonferroni-corrected. Genes were considered as differentially expressed at a p-value <0.05

Reflecting the results showed in Fig. 4, in Table 4 it appears that the number of genes differentially expressed in shoots was higher than in roots. Moreover, most of the differentially expressed genes were root-specific or shoot-specific, with only a few commons to the entire plant, whichever metabolic pathway was considered.

In shoots (Table 4), c.a. 50% of genes involved in nitrogen and photosynthetic pathways were up-regulated. Among the genes for nitrogen metabolism nitrate reductase (+24.507 fold), nitrite reductase (+9.552 fold) and genes involved in amino acid metabolism were found. Genes that were up-regulated in photosynthetic pathways were ferredoxin and ferritin, i.e. components of the electron transport chain. Among the genes that were down-regulated in nitrogen metabolism and photosynthetic pathways, proteinases (−86.773 and −5.381 fold), chlorophyllases (−9.972 and −7.938 fold) and stay green protein (−10.253 fold) were found, i.e. genes implicated in chlorophyll catabolism during senescence. In contrast, 80% of genes involved in sulfate metabolism were up-regulated with a high representation of genes involved in sulfate uptake and assimilation (sulfate transporter, ATP sulfurylase and serine acetyltransferase).

Microarray analysis of roots (Table 4) revealed that, like shoots (Table 4), 50% of the differentially expressed genes were up-regulated for the three metabolisms considered. Similar to nitrogen metabolism in shoots, genes implicated in amino acid metabolism were up-regulated and senescence proteinases were down-regulated. In photosynthetic pathways, also in common with shoots, an up-regulation of ferredoxin and carbonic anhydrase was retained. However, a large down-regulation of the plastid division regulator, MinE, was observed (26.014 fold in shoots and 30.623 fold in roots after 3 days). Focusing on S metabolism in roots, expression of a sulfate transporter gene was up-regulated but it was a different one than in the shoots, while glutathione S transferase was down-regulated.

After 30 days of treatment with HA7, only one gene was differentially expressed in shoots in the chosen metabolic pathways (supplemental data, Table S2). It was an S-adenosylmethionine decarboxylase of the sulfur metabolic pathway, which was down-regulated 11.597 fold.

N metabolism

Microarray analysis highlights the up-regulation of genes encoding proteins involved in N metabolism (especially nitrate and amino acids). From these data, physiological and biochemical approaches have been developed to confirm the beneficial effect of humic acid extract on N metabolism of WOSR. Thus, in the first instance, the total amount of N in plants treated by humic acid extract has been compared to control plants (Fig. 5a). Secondly, the effects of humic acid extract on nitrate uptake (using 15NO3, Fig. 5b), expression of two major nitrate transporters (BnNRT1.1 and BnNRT2.1, Fig. 5c) and nitrate reductase activity in shoots and roots (Fig. 5d) were quantified. The total N amount in treated plants (Fig. 5a) showed that after 30 days, treatment with HA7 significantly affected the N content of plant roots. Thus, at days 30, the N content in roots of treated plants was significantly higher than in control plants (+108 ± 21.9%). Indeed, compared to control, plants treated by HA7 showed an increase of nitrate uptake (Fig. 5b) at day 1 and day 30 (+32 ± 15.3 and +31 ± 13.7%, respectively). Because these last two data points were of the same order of magnitude as the increase in DW (Fig. 3), this show that N uptake followed the stimulation of growth. In the meantime (Fig. 5c), the BnNRT2.1 expression level was increased by 10.7 ± 5.4 fold in 1-day treated plants and remained over-expressed at day 3 (22.1 ± 18.7 fold) but it was not detectable after 30 days of treatment while BnNRT1.1 expression was more strongly induced later on (40.7 ± 17.3 and 630.0 ± 180.7 fold at 3 and 30 days respectively, Fig. 5c). Additionally, compared to control plants, nitrate reductase activity (Fig. 5d) increased significantly in roots after 1 day (+74 ± 18.0%) and in shoots after 30 d (+59 ± 15.2%) of treatment. However, no nitrate accumulation in shoot or root tissues was detected in plants (data not shown).
Fig. 5

Effects of HA7 extract on N metabolism: a Total N amount in shoots and roots of control plants (white histogram) or treated plants with HA7 (hatched histogram). b Net nitrate uptake. c q-PCR analysis of the expression level of BnNRT1.1 (black histogram) and BnNRT2.1 (gray histogram), two genes encoding root transporters implicated in N uptake. For q-PCR analysis, control (white histogram, value 1) corresponds to the expression level of each gene in control plants at each time of harvest. d In vivo nitrate reductase activity in shoots and roots of control (white histogram) and treated plants (hatched histogram). For all data, indicated values are means and vertical bars indicate ± standard deviation for n = 10 when exceeding the symbol. Significant differences at p < 0.05 and p < 0.01 are indicated by one and two asterisks, respectively

S metabolism

Microarray analysis highlighting the up-regulation of genes involved in sulfur uptake and assimilation, the total amount of S (Fig. 6a) and the sulfate content (Fig. 6b) have been quantified in roots and shoots. Moreover the expression levels of sulfate transporters, such as BnSultr1.1 and BnSultr1.2 genes (plasmalemmic transporters, Fig. 6c) and BnSultr4.1 and BnSultr4.2 genes (tonoplastic transporters, Fig. 6d) were quantified in roots and shoots, respectively. The total S amount in treated plants (Fig. 6a) shows that treatment with HA7 resulted in significantly higher shoot (+76 ± 30.8%) and root (+137 ± 13.7%) S contents after 30 days of treatment. Unlike N, these data were higher than the magnitude of the DW increase (Fig. 3), reflecting a stimulation of sulfate uptake per s.e. (Fig. 6b). Indeed, the total sulfate amount in the plants (Fig. 6b) shows that treatment with HA7 resulted in accumulation of sulfate in shoots (+21 ± 2.3% and +46 ± 16.0% at 1 and 30 days respectively) and in roots (+103 ± 25.8% at 30 days) after 1 and 30 days of treatment but with a decrease at 3 days (−35 ± 1.6% and −46 ± 2.7% in shoots and roots respectively). In the meantime, compared to control plants, BnSultr1.1 and BnSultr1.2 (Fig. 6c) were induced after 1 and 3 days of treatment with HA7 (2.8 ± 1.7 fold and 14.1 ± 1.6 fold at 1 and 3 days respectively for BnSultr1.1; 14.8 ± 4.8 fold and 18.3 ± 3.1 fold at 1 and 3 days respectively for BnSultr1.2). Then after 30 days, the expression levels of both sulfate transporters were not detectable in plants. Moreover, BnSultr4.1 and BnSultr4.2 genes (Fig. 6d), encoding tonoplastic transporters involved in vacuolar fluxes of sulfate, were induced only after 3 days of treatment (1.8 ± 0.2 and 7.3 ± 0.4 fold respectively).
Fig. 6

Effects of HA7 on S metabolism: a Total S amount in the shoots and roots of control plants (white histogram) or plants treated (hatched histogram) with HA7. b Total S-sulfate content in the shoots and roots of plant control or treated with HA7. c q-PCR analysis of the expression level of BnSultr1.1 (black histogram) and BnSultr1.2 (gray histogram), two genes encoding transporters involved in S uptake in roots. d q-PCR analysis of the expression level of BnSultr4.1 (black histogram) and BnSultr4.2 (gray histogram), two genes encoding sulfate transporters involved in sulfate sequestration in shoot vacuoles. For q-PCR analysis, control (white histogram, value 1) corresponds to the expression level of each gene in control plants at each time of harvest. For all data, indicated values are means and vertical bars indicate ± standard deviation for n = 10 when exceeding the symbol. Significant differences at p < 0.05 and p < 0.01 are indicated by one and two asterisks, respectively. N/D: not detected


Microarray analysis shows that, compared to control, some genes involved in carbon and photosynthesis pathways were differentially expressed in plants treated by humic acid extract. For example, in shoots, genes encoding the plastid division regulator, MinE, or stay green protein (Srg) were highly down-regulated by 26.014 and 10.253 fold respectively after 3 days (Table 4). In contrast, genesencoding ferredoxin NADP reductase or carbonic anhydrase were up-regulated by HA7 (Table 4). To complete this analysis, relative chlorophyll content (Fig. 7a) and net photosynthetic rate (Fig. 7b) were measured and no significant difference was observed between control and HA7-treated plants for this parameter. However, HA7-treated plants presented significantly higher net photosynthetic rates between 3 and 21 days of treatment (+41 ± 1.7%, +42 ± 2.8%, +23 ± 1.4% and +20 ± 1.6% at 3, 7, 15 and 21 days of treatment respectively). Therefore microarray analysis, chlorophyll content and the net photosynthetic rate seem to give contradictory results. Thus, confocal microscopy was used to observe chloroplast numbers in mature leaf cells from treated or control plants (Fig. 7c). After 1 and 30 days of treatment, HA7 application significantly increased the number of chloroplasts per cell (+43 ± 10.7% and +130 ± 15.8% at 1 d and 30 d respectively). Indeed, in cells from control plants, the number of chloroplasts was relatively constant during the experiment (39.6 ± 2.1, 47.4 ± 3.7 and 46.6 ± 1.3 at 1 d, 3 d and 30 d, respectively). In leaf cells from treated plants, whatever the duration of treatment, the number of chloroplasts was always higher than controls (56.7 ± 4.2, 51.9 ± 1.8 and 107.3 ± 7.4). In order to estimate the potential humic acid effect on chloroplastic ultrastructure, transmission electronic microscopy (TEM) was performed after 30 days of treatment (Fig. 7d). Compared to the ultrastructure of chloroplasts from control plants, TEM showed no effect of HA7 application on thylakoids organization, but an enhancement of the number and the size of starch granules suggesting an accumulation of carbohydrate compounds in chloroplasts in response to treatment.
Fig. 7

Effects of HA7 on chloroplasts and photosynthesis: a Chlorophyll content in leaves from control plants (continuous line) or treated plants (dotted line) measured by SPAD. b Kinetics of the net photosynthetic rate from control (continuous line) or treated leaves (dotted line). Value c Fluorescence confocal microscopy of chloroplasts of controls or plants treated with HA7 after 1, 3 and 30 days of treatment. Bars represent a scale of 50 μm on the images. Numbers of chloroplasts per cell are indicated in the white square and are the means ± standard deviation for n = 10 cells. d Transmission Electron Microscope (TEM) observations of chloroplasts from control (top) or from leaves treated by humic acid (bottom) over 30 days. Black bars represent a scale of 2 μm. SG: starch granules. For A and B, results are means ± standard deviation for n = 5 leaves with 10 measurements per leaf. Asterisks represent the result of the Student’s test at p < 0.05 for *, p < 0.01 for ** and p < 0.001 for ***


The aim of this study was to identify a new and finely characterized humic acid that could increase growth and mineral nutrition of WOSR in order to reduce fertilizer use.From a previous set of screening experiments (data not shown), HA7 extract was selected from ten humic acids extracts. Only HA7 promoted a significantly higher growth rate and leaf chlorophyll content compared to control non-treated rapeseed. Its effects were confirmed in the present experiment on total DW (+29 ± 11.5%), and more specifically root DW (+88 ± 19.0%) (Fig. 3). This growth promotion by humic acids has been already reported by several authors and for different species, such as Arabidopsis thaliana (Schmidt et al.2007), barley (Ayuso et al.1996), cucumber (Mora et al.2010), maize (Canellas et al.2002; Zandonadi et al.2007) and tomato (Atiyeh et al.2002). Most studies have hypothesized that the observed effects in treated plants were the result of hormone-like activity by humic acids (Quaggiotti et al.2004; Muscolo et al.1998, 2007b). However, phytohormone analysis (Table 2) did not reveal any significant level of hormones (IAA, ABA, cytokinins…) in HA7 extract. Moreover, its mineral contribution to the nutrient solution was negligible compared with Hoagland solution alone (Table 3). Thus from our results, it cannot be postulated that known endogenous hormones and nutrients supplied from HA7 triggered the promotion of growth. However, these results cannot exclude de novo phytohormone synthesis in plants.

Therefore, to identify HA7 metabolic targets in plants, we used a microarray analysis specific for Brassica napus that allowed the analysis of 31 561 genes. Nevertheless, most of these genes were not previously identified (Fig. 4) due to the lack of complete sequencing of the Brassica genome. However, from the identified genes, which accounted for about 40% of gene expression analyzed after 3 days of contact with HA7, the expression of about one thousand known genes was significantly affected. Yet, after 30 days of contact with HA7, only one hundred of these genes remained differentially expressed (Fig. 4). Thus, this high number of differentially expressed genes reflects an early and substantial effect of HA7 at the molecular level that influenced almost all plant metabolic pathways. These results are in accordance with Tresivan’s study (2011) on Arabidopsis thaliana, where, after 120 min of contact with an earthworm-originating humic acid, the expression of 133 genes was changed in comparison with untreated control plants. Using DCFI annotation, HA7 affected genes could be classified in nine clusters covering the major metabolic functions of plants: carbon and photosynthesis, general cell metabolism, fatty acids, nitrogen/sulfur, phytohormones, plant development, senescence, responses to stress and transport of ions and water. Among these pathways, the most affected after 3 days of contact were carbon and photosynthesis, cell metabolism, nitrogen/sulfur metabolisms and responses to stress (Fig. 4).

Overall, microarray analysis highlighted the enhancement of gene expression related to N and S metabolism and more particularly on identified genes encoding proteins involved in uptake and assimilation (Table 4). However, physiological analyses also gave access to more precise interpretation of data. For N metabolism (Fig. 5), increases in NO3- uptake (+31 ± 13.7%) and N content (+108 ± 21.9% in roots) were of the same order of magnitude as the DW increase (+88 ± 19.0% in roots). Moreover, q-PCR analysis of the expression of the BnNRT1.1 and BnNRT2.1 genes encoding nitrate transporters showed an induction of these genes in roots from treated plants. Surprisingly, these changes were not revealed by microarray analysis. However, nitrate transporters are a very large family of genes (Daniel-Vedele et al.1998) and a BLAST analysis showed that the nitrate transporter probes used on microarrays are weakly specific for the BnNRT1.1 and BnNRT2.1 isoforms. All these data suggest that supplemental N taken up by the roots of treated plants was directly assimilated in relation to the growth rate without being stored (enhancement of nitrate reductase enzymatic activity, Fig. 5d and no N or nitrate content enhancement, data not shown). Recent studies (Castaings et al. 2010; Krouk et al. 2010) pointed out that NRT1.1 (whose gene expression was strongly induced by HA7) may have a role in N sensing as well as in IAA transport and ultimately on root growth, a process that was also largely stimulated by HA7 (Fig. 3c).

In contrast, for S metabolism (Fig. 6), the S content after 30 days (+76 ± 30.8 and 137 ± 13.7% in shoots and roots respectively) increased more than dry matter and resulted in enhanced sulfate content (+46 ± 16.0 and +103 ± 25.8% in shoots and roots respectively). This has also been related to an increased expression of the main root and tonoplastic sulfate transporter genes (Fig. 6) alongside an enhancement of assimilatory gene expression (Table 4). These results are in agreement with previous works that showed an enhanced nitrate uptake in response to humic acid (Dell’Agnola and Nardi 1987; Pinton et al.1999; Nardi et al.2000b; Keeling et al.2003; Eyheraguibel et al.2008) in Bermuda grass (Cynodon dactylon), maize, wheat and oilseed rape. In addition, the enhanced expression of genes involved in nitrate transport has also been reported in maize (Quaggiotti et al.2004). As N is not increased by the addition of HA7 to the nutrient solution (Table 3), enhancement of N uptake in response to treatment is possibly only a consequence of growth promotion and more specifically of lateral root development. Focusing on S metabolism, Eyheraguibel et al. (2008) demonstrated that maize presented an increase in S accumulation in roots in response to humic acid treatment.

The second axis of our study focused on carbon assimilation by photosynthesis. In our work, no significant effect was found on chlorophyll content (Fig. 7a). However, net photosynthetic rate measurements (Fig. 7b) showed an increased activity for HA7-treated plants between 3 and 21 days of contact with the extract. Likewise, the microscopic observations (Fig. 7c) showed that, in response to HA7, chloroplasts contain higher quantities of starch compared with control non treated plants after 30 days of contact. These results suggest that HA7 has an effect not only on the clear phase of photosynthesis (net photosynthesis measurement at 1000 μmol PAR) but it also enhanced the dark phase of photosynthesis (carbon fixation and starch synthesis). To support this hypothesis, the microarray results showed that genes involved in carbon fixation (Table 4 and Supplemental Table S1 and S2) were mostly up-regulated after 3 days of treatment with HA7. For example, genes encoding carbonic anhydrase, a zinc-requiring enzyme, catalyzing the reversible hydration of carbon dioxide and thus facilitating its transfer and fixation (Ramanan et al.2009), is up-regulated 5.469 fold in shoots and 6.674 fold in roots after 3 days of treatment. Carbonic anhydrase is important in photosynthesis and respiration as it participates in the transport of inorganic carbon to actively photosynthesizing cells and away from actively respiring cells (Moroney et al.2001). Therefore, the enhanced expression of gene encoding this protein could explain the increased carbon fixation suggested by the increased number and size of starch granules observed in chloroplasts of HA7-treated plants (Fig. 7d).

Furthermore, confocal microscopy observations showed a clear effect on the number of chloroplasts per cell from 1 d of contact with HA7 extract (Fig. 7). Okazaki et al. (2009) obtained similar results with Arabidopsis thaliana in response to exogenous application of cytokinins. According to these authors, plastid division components of chloroplasts were under control of a cytokinin-responsive transcription factor (CRF2). In our study, only one “plastid division component” was found on the microarray, the “plastid division regulator MinE”, but it was clearly down regulated in response to HA7 treatment after 3 days of contact. However, the literature on this gene provides contradictory results. Itoh et al. (2001) showed that over-expression of MinE in Arabidopsis thaliana resulted in cells containing only one giant chloroplast. Kojo et al. (2009) obtained the same phenotype with an AtMinE deficient mutant (2,000-fold reduced MinE expression compared with wild type). The second aspect of the Okazaki study (2009) was the role of exogenous cytokinin in chloroplast division. However, no significant levels of phytohormone have been detected in HA7. Thus, this suggests that either HA7 acts on chloroplasts in a different way than phytohormones or that HA7 can induce early phytohormone biosynthesis in plants. To support this second hypothesis, the microarray results showed that genes involved in cytokinin and gibberellin metabolisms were up regulated in shoots after 3 days of contact with HA7 (supplemental data, Table S1).

These results suggest that the first events to occur after HA7 application to plants was an increase in chloroplast number per cell, in parallel with the enhancement of root nitrate uptake (+32 ± 15.3%) and increases in expression of the root nitrate transporter BnNRT2.1 (10.7 ± 5.4 fold after 1 d, Fig. 5) as well as the sulfur transporters BnSultr1.1 (2.8 ± 1.7 fold after 1 d, Fig. 6) and BnSultr1.2 (14.8 ± 4.8 fold after 1 d, Fig. 6). These first responses were followed by sulfate accumulation in shoots and even a greater number of chloroplast over the time of treatment. Because chloroplasts were the first organelles affected during senescence, it could be assumed that the increase in the number of chloroplasts in HA7-treated plants would promote a delay of senescence and extend the life span of leaves. This hypothesis was also supported by the microarray analysis that revealed up-regulation of some genes encoding protease inhibitors and down regulation of stay green protein (Srg) that are able to limit leaf protein degradation (Etienne et al.2007; Desclos et al.2008) and protect chlorophyll degradation (Park et al.2007) during leaf senescence, respectively. According to this work, it could be assumed that chloroplast modifications could act on the duration of leaf senescence and the leaf life span and allow an improvement in the NUE of WOSR. Thus, the HA7 extract could be especially relevant to augment or substitute the fertilizers actually used and improve the agro-environmental balance of WOSR.

This study combining microarray and physiological analyses to explain effects on WOSR growth gives clues about the metabolic targets of humic acid. Enhancement of N, C and S assimilation could explain increased growth of plants treated with HA7 extract. Furthermore, in addition to measurements of an enhanced net photosynthetic rate, this study shows an early effect of HA7 on the chloroplast numbers per cell and the number of starch granules. However, more analyses should be carried out for further determinations of the HA7 metabolic target. For example, in planta phytohormonal measurement could help to better understand HA7 effects on chloroplasts and/or the regulation of metabolic pathways. Furthermore, some genes involved in different metabolic pathways (such as responses to stress and senescence) were up- or down-regulated in response to HA7 treatment (supplemental data, Tables S1 and S2) and remain to be studied to improve our knowledge of the effects of humic acids on plant physiological processes.



This study was a part of AZOSTIMER project selected and supported by the Pôle de compétitivité Mer-Bretagne and funded by the French FUI (Fond Unique Interministériel), Brittany Region and Saint-Malo Agglomeration. We thank Marie-Paule Bataillé and Raphaël Ségura for IRMS analyses. We acknowledge Patrick Beauclair for LICOR measurements, Julie Levallois for technical assistance in RNA extractions and q-PCR analyses, Xavier Sarda and Anne-Françoise Ameline for helping with plant culture and harvest and finally Nicolas Elie from GRECAN (Groupe Régional d’Etude sur le CANcer, Histo-imagerie quantitative, Caen, France) for microscopic analysis. We thank Laurence Cantrill for improving the English of the manuscript.

Supplementary material

11104_2012_1191_MOESM1_ESM.xls (296 kb)
Supplemental Table S1List of the differentially expressed genes in shoots and roots of rapeseed after 3 days of HA7 supply to the roots. (XLS 295 kb)
11104_2012_1191_MOESM2_ESM.xls (35 kb)
Supplemental Table S2List of the differentially expressed genes in shoots and roots of rapeseed after 30 days of HA7 supply to the roots. (XLS 35 kb)


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Copyright information

© Springer Science+Business Media B.V. 2012

Authors and Affiliations

  • Laëtitia Jannin
    • 1
    • 2
  • Mustapha Arkoun
    • 1
    • 2
  • Alain Ourry
    • 1
    • 2
  • Philippe Laîné
    • 1
    • 2
  • Didier Goux
    • 3
  • Maria Garnica
    • 4
  • Marta Fuentes
    • 4
  • Sara San Francisco
    • 4
  • Roberto Baigorri
    • 4
  • Florence Cruz
    • 5
  • Fabrice Houdusse
    • 5
  • José-Maria Garcia-Mina
    • 4
  • Jean-Claude Yvin
    • 5
  • Philippe Etienne
    • 1
    • 2
  1. 1.Université de Caen Basse-Normandie, UMR 950 Ecophysiologie VégétaleCaen CedexFrance
  2. 2.INRA, UMR 950 Ecophysiologie VégétaleCaen CedexFrance
  3. 3.Université de Caen Basse-Normandie, Centre de Microscopie Appliquée à la Biologie (CMABio)Caen CedexFrance
  4. 4.TIMAC Agro SpainOrcoyenSpain
  5. 5.Centre de Recherche International en Agroscience, CRIAS-TAI, Groupe RoullierDinardFrance

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