Production of phytohormones, siderophores and population fluctuation of two root-promoting rhizobacteria in Eucalyptus globulus cuttings

ORIGINAL PAPER

Production of phytohormones, siderophores and populationfluctuation of two root-promoting rhizobacteria in Eucalyptusglobulus cuttings

Katy Dıaz Peralta • Tamara Araya • Sofıa Valenzuela •

Katherine Sossa • Miguel Martınez • Hugo Pena-Cortes •

Eugenio Sanfuentes

Received: 18 October 2011 / Accepted: 6 January 2012 / Published online: 15 January 2012

� Springer Science+Business Media B.V. 2012

Abstract Vegetative propagation by stem cuttings and

mini-cuttings has been used worldwide for growing

Eucalyptus plants. However, clones and hybrids of this

plant present a great variability in their rooting capacity,

apart from a gradual decrease in the rooting potential due

to the ontogenetic age of the mother plant. Several studies

have demonstrated that some bacteria promote plant

growth and rooting through the action of direct and indirect

mechanisms that are not still completely clear. Considering

this, the objective of this study was to assess the production

of auxins, abscisic acid and siderophores in Bacillus

subtilis and Stenotrophomona maltophilia, which in pre-

vious studies increased rooting of E. globulus cuttings.

Additionally, the population of these bacteria in the

rhizosphere, superficial tissues of the stem-base and callus

of the mini-cuttings was identified, and quantified by

real-time PCR. Only S. maltophilia produced IAA in the

presence of tryptophan; none of the bacterial strains pro-

duced ABA, but both produced siderophores. A compara-

tive analysis of the separation profiles showed that there is

a diverse microbial community in the rhizosphere, and only

S. maltophilia was capable of keeping its population at a

density of 2.03 9 107 cells/mg in different tissues of the

mini-cuttings. The results would indicate that the rooting

stimulus in E. globulus could be related to the action of one

or several mechanisms such as the production of auxins

and siderophores, and it could also be associated with the

ability of bacteria to stay in the rhizosphere or in plant

callus tissues.

Keywords Stenotrophomona � Bacillus �Phytohormones � Siderophores � Rhizosphere � DGGE

Introduction

Currently, Eucalyptus is the second exotic genus impor-

tance in Chile with a total planted area of 661,394 ha,

Eucalyptus globulus being the most important with

471,743 ha. This specie has a valuable economic role in the

forestry industry in the country because of their wide dis-

tribution (including a range of climate and soil), their

ability for the manufacture of quality paper, and their rapid

growth (INFOR 2008).

The worldwide forest breeding programs are based on

the clonal plantation of species from the genus Eucalyptus,

employing vegetative propagation using macro and micro-

propagation techniques. However, vegetative propagation

in some species does not provide the best reproduction

system due to particular effects of endogenous and exog-

enous factors on the mother plant, alterations of root

K. D. Peralta � E. Sanfuentes

Forest Pathology Laboratory, Biotechnology Center, University

of Concepcion, Concepcion, Chile

K. D. Peralta (&) � H. Pena-Cortes

Biotechnology Center ‘‘D. Alkalay L’’, Universidad Tecnica

Federico Santa Marıa, Valparaiso, Chile

e-mail: [email protected]

T. Araya � K. Sossa

Biofilms and Environmental Microbiology Laboratory,

Biotechnology Center, University of Concepcion, Concepcion,

Chile

S. Valenzuela

Molecular Biology and Genomics Laboratory, Biotechnology

Center, University of Concepcion, Concepcion, Chile

M. Martınez

Microbiology Laboratory, Faculty of Biological Sciences,

University of Concepcion, Concepcion, Chile

123

World J Microbiol Biotechnol (2012) 28:2003–2014

DOI 10.1007/s11274-012-1003-8

system architecture and manifestation of topophysis effects

that determine differences in rooting potential. Thus,

E. globulus, E. nitens and hybrids of these species present a

great variability in their rooting capacity, even at the

clone level, and are considered to be recalcitrant species

(Martellet and Fett-Neto 2005).

Over the last decades, various studies using symbiotic

bacteria and plant growth-promoting rhizobacteria (PGPR)

in agriculture, horticulture and forestry have been con-

ducted in order to control pathogen attack and to generate

biological products that improve crop yield (Vessey 2003;

Lucy et al. 2004). Among the rhizobacteria identified to

promote growth are species of Pseudomonas, Bacillus,

Acetobacter, Agrobacterium, Azospirillum, Stenotropho-

mona, Enterobacter, Serratia, Erwinia, Klebsiella, Strep-

tomyces and Rhizobium (Arshad and Frankerberger 1998;

Kumar et al. 2006; Dimkpa et al. 2008).

Bacillus subtilis and S. maltophilia showed high fre-

quency of occurrence in the rhizosphere (Kim et al. 1997;

Berg et al. 2005) and have been described as plant growth

promoters (Suckstorff and Berg 2003; Ryu et al. 2005),

biological control agents of fungal diseases (Nakayama

et al. 1999; Vespermann et al. 2007) and are being involved

in the synthesis of phytohormones such as indole-3-acetic

acid (IAA), gibberellic acid (Gibberellin), zeatin (Cytoki-

nin), abscisic acid (ABA), and ethylene (Martellet and

Fett-Neto 2005; Karadeniz et al. 2006). Abscisic acid

controls plant growth and inhibits root elongation (Pilet

and Chanson 1981), which means that there is a negative

correlation between growth and the endogenous ABA

content of plants (Pilet and Saugy 1987). It has been

reported that some bacterial species that interact with

plants or live in the soil, synthesize abscisic acid (Tuomi

and Rosenquist 1995; Karadeniz et al. 2006). Using IAA

deficient mutants of Azospirillum brasilense, the increase

in wheat roots was associated with the IAA synthesis and

suggested that this hormone could act as a signaling mol-

ecule in the Azospirillum-plant interaction (Dobbelaere

et al. 1999; Spaepen et al. 2007). In several PGPR, it has

been demonstrated that the increased proliferation of roots

might be related to bacterial IAA biosynthesis (Spaepen

et al. 2007; Teixeira et al. 2007), thus suggesting that 80%

of the bacteria isolated from the rhizosphere produce IAA

(Patten and Glick 2002). It is also likely that the combined

action of both IAA and ABA in wheat roots increases plant

growth Muller et al. (1989).

On the other hand, the production of siderophores is an

indirect mechanism associated with the increase in plant

growth by PGPR. The production of bacterial siderophores

stimulates plant growth by increasing iron availability in

the rhizosphere and inhibiting pathogen growth in the

rhizosphere. Thus, siderophores become a feasible option

for biocontrol (Kloepper et al. 1980; Hernandez et al.

2004). Among these siderophores we can find catecholate-

type, hydroxamate-type and a-carboxylate-type sidero-

phores, depending on their chemical nature and iron

coordination sites (Winkelmann 2002).

In recent years, it has been determined that those

bacteria able to produce both siderophores and auxins

could be potential candidates to be used in phytoremedia-

tion of metal contamination. Nonetheless, there are metals

that inhibit auxin production by bacteria and, therefore,

make bacterial involvement in plant growth promotion less

efficient (Dimkpa et al. 2008).

Recently, it has been demonstrated that some rhizo-

bacteria have the ability to increase proliferation of roots in

micropropagated plants of the genus Eucalyptus (Teixeira

et al. 2007; Dıaz et al. 2009). Thus, B. subtilis and

S. maltophilia increased rooting of E. globulus cuttings

over 50% (Dıaz et al. 2009). Several mechanisms have

been postulated to explain how rhizobacteria promote plant

growth due to either a direct or indirect effect (Persello-

Cartieaux et al. 2003; Ryu et al. 2005); however, these

mechanisms are not fully understood, and the effects they

may have on rooting are even less studied. Rhizosphere

bacteria can directly or indirectly promote plant growth and

rooting; however, their effect depends on the capacity of

settling or increasing their population in roots. Several

studies have revealed the structural and functional diversity

of the bacterial population in the rhizosphere, as well as the

variables that determine this diversity, such as the plant

species, the differences in the amount, location and com-

position of the radicular exudates, and the effect of the soil

type, cultivation practices and other environmental factors

(Grayston et al. 1998; Yang and Crowley 2000; Duineveld

et al. 2001). Nevertheless, diversity and population

dynamics of bacteria in the rhizosphere have probably been

underestimated since most studies have addressed the

culturable bacteria, which would only represent a small

proportion (0.1–10%) of the total bacteria present in the

rhizosphere. Recently, molecular techniques based on the

extraction of 16S ribosomal DNA from the bacteria present

in the soil or the rhizosphere have been used in studies of

microbial ecology for a wide range of habitats (Muyzer and

Smalla 1998; Smalla et al. 2001). The use of DGGE offers

a culture-independent method for tracking rhizosphere

dominant bacterial populations in space and time (Muyzer

et al. 1993; Heuer and Smalla 1997). Another method used

for studying the population dynamics of microorganisms,

both in medical and environmental research, is the quan-

titative real-time PCR technique, which offers the oppor-

tunity to detect and quantify specific genes in order to

determine the bacterial population dynamics. Some studies

have quantified specific genes from Desulfotomaculum

cells in environmental samples of soil (Stubner 2002).

Using a set of probes and primers and with the help of PCR

2004 World J Microbiol Biotechnol (2012) 28:2003–2014

123

and probe hybridization techniques, it has been possible to

quantify and localize in situ the total amount of Entero-

bacter radicincitans cells in roots of Brassica oleracea

cuttings (Ruppel et al. 2006).

The objectives of this study were to analyze the ability

of the strains of B. subtilis and S. maltophilia, that have

shown potential as bioproducts for rooting promotion in

E. globulus, to synthesize phytohormones such as IAA and

ABA and siderophores and to determine the population

dynamics of these bacteria present in the rhizosphere and

superficial tissues of the stem-base and callus of the

E. globulus mini-cuttings.

Materials and methods

Bacterial strains

The strains used for the study were B. subtilis (C1K30) (448/

451 bp, access number EU826029; Genbank) and S. malto-

philia (C3P7) (501/501 bp, access number CP001111.1;

GenBank). These bacterial strains were isolated from the

rhizosphere of E. globulus and E. nitens plants and increased

the rooting of E. globulus cuttings (Dıaz et al. 2009). The

strains were preserved in Dimethyl sulfoxide (DMSO) (5%

v/v) and glycerol (87%) and stored at -20�C.

Detection of auxins (IAA) in bacterial cultures

The production of auxins by both strains was measured

using the Salkowski colorimetric technique (2 mL 0.5 M

FeCl3 ? 98 mL 35% HClO4) (Asghar et al. 2002). After

growing for 24 h at 30�C in Petri dishes containing R2 A

solid medium, the bacterial inoculum was transferred to

flasks with 25 mL of R2 A culture broth with and without

5 mL of tryptophan (0.5%), and then incubated at 25�C on

a horizontal shaker (120 rpm). After 48 h, 1.5 mL were

removed from each flask and these were centrifuged for

15 min at 1,000 rpm to collect the supernatant (Torres

et al. 2000), which was then mixed with the Salkowski

reagent (ratio 2:3). After 30 min, a pink color was observed

and was measured using a spectrophotometer (Cole-Parmer

1100 RS, Spectrophotometer) at 535 nm. The auxin con-

centrations were calculated with a calibration curve as

standard ranging from 0.2 to 45 lg/mL.

Quantification of auxins and abscisic acid using high

performance liquid chromatography (HPLC)

Sample preparation

Bacterial colonies of both strains were cultured in flasks

containing 100 mL of R2 A culture medium, with and

without 5 mL of tryptophan (0.5%), and then they were

incubated on a shaker at 120 rpm and 25�C for 48 h. The

bacterial cells were separated from the supernatant by

centrifugation at 8000 rpm and 4�C for 25 min. A 2 mL

aliquot of the supernatant was adjusted to pH 2.8 using 1 M

HCl and it was extracted three consecutive times with

2 mL of ethyl acetate. Subsequently, the extract was

evaporated to dryness under a nitrogen stream and dis-

solved in 600 lL of mobile phase, to be later stored at

-20�C. All the samples were filtered with 0.22 lm

hydrophobic Millipore filters before being injected (Ahmad

et al. 2005).

Indoleacetic acid (IAA) and abscisic acid (ABA)

calibration curve

A standard IAA Merck solution of 75 lg/mL (99% purity)

and a standard SIGMA ABA solution (99% purity) were

prepared by weighing 7.5 mg of IAA and ABA suspending

in 100 mL of absolute methanol. To determine the cali-

bration curve for IAA, successive dilutions of 0.03, 0.2,

0.4, 0.5, 3, 5 and 9 lg/mL were performed. In the case of

B. subtilis, the curve was adjusted using concentrations of

0.03–0.5 lg/mL; for S. maltophilia, using concentrations

of 0.5–9 lg/mL. To determine the ABA calibration curve,

successive dilutions of 40, 50, 60 and 75 lg/mL were

performed for both strains.

HPLC operating conditions and parameters

The chromatographic separation was performed isocrati-

cally in a C18 reversed-phase column (LiChroCART�

250-4 LiChospher� 100 RP-18 (5um) Merck KGaA

HPLC). The mobile phase was consisted of a mixture of

45% methanol and 0.1 M acetic acid in a proportion of

60:40 (v/v), filtered and degassed at a 0.8 mL/min flow rate

and at room temperature. A volume of 20 lL was injected

to each sample. IAA was detected using a fluorescence

detector with excitation at k 280 nm and emission at k340 nm; ABA amountwas detected by using a UV detector

262 nm. The HPLC system consisted of a Merck-Hitachi

D-7000 HPLC system composed of a D-7000 autosampler,

a D-7200 pump and an L-7400 UV Detector (Merck-

Hitachi).

Experimental design and evaluation

To evaluate the production of IAA and ABA, a chro-

matogram peak area ratio was determined by using a cal-

ibration straight line of IAA and ABA.

A completely randomized design was used, and three

replicates per treatment were performed. The control

treatment was composed of only R2A culture medium for

World J Microbiol Biotechnol (2012) 28:2003–2014 2005

123

the samples with and without tryptophan. The data were

analyzed by linear regression analysis performed with

Origin 6.1 software.

Detection of siderophores

Siderophores were detected qualitatively by CAS assay in

solid medium (Schwyn and Neilands 1987) and a few

modifications. To make 1 L of CAS agar medium, the

following were added to 750 mL of water and 1 g/L Pipes

(SIGMA): NaOH (1 N) to adjust pH to 6.8, 109 MM9

(100 mL) and agar (15 g). After autoclaving and cooling

the medium to 50�C 30 mL deferrated casamino acids

(10%), 10 mL of glucose (20%), 1 mL 1 M MgCl2, 1 mL

100 mM CaCl2 and 4 mL 500 lg/mL thiamine were

added. Finally, 100 mL Chrome Azurol S-hexadecyl-

trimethylammonium bromide (CAS-HDTMA) solution

were added.

Once prepared, the medium was placed in 10 cm Petri

dishes. 24 h later, the bacterial cultures that were growing

in R2A culture medium were grown at 30�C for 24 h. Agar

plates containing concentrations of both strains, a positive

control (B. steratermophillus) and a negative control (agar

without bacteria) were cast upon the CAS medium and then

incubated at 30�C for 48 h. The plates showing yellow

halos indicated the production of siderophores. The assays

were performed in triplicate.

Determination of the type of siderophores

with the O-CAS method

Siderophores were determined using the overlay CAS

culture medium (Perez-Miranda et al. 2007), which was

prepared as described by Schwyn and Neilands (1987) with

some modifications. The medium contained the following:

CAS 60.5 mg/L, HDTMA 72.9 mg/L, Piperazine-1,4-bis

(2-ethanesulfonic acid) (PIPES) 30.24 g/L, and 1 mM

FeCl3 9 6H2O in 10 mM HCl 10 mL/L. Agarose (0.9%

w/v) was used as gelling agent.

Siderophore detection was achieved after 15 mL over-

lays of this CAS medium were applied over those agar

rhizosphere siderophore medium (RSM) (Buyer et al.

1989) plates containing cultivated microorganisms to be

tested for siderophore production. After 30 min, a change

in color was observed in the medium; for catecholate-type

siderophores, the medium turned purple; for hydroxamate-

type siderophores, it turned orange; and for carboxylate-

type siderophores, it turned yellow. These experiments

were performed twice, with three replicates for each strain.

The strain used as a positive control was B. stearother-

mophilus and a plate lacking bacteria was used as a neg-

ative control.

Determination of microbial diversity in the rhizosphere,

superficial tissues of the stem-base and callus of the

minicuttings of E. globulus inoculated with B. subtilis

and S. maltophilia

Production of inoculum and application of bacteria

To produce the inoculum, plates containing PDA (potato

dextrose agar) culture medium were inoculated with the

strains and incubated for 24 h at 25�C. A loopful of the

bacterial inoculum was obtained from the medium to be

cultured in flasks containing 60 mL of R2A medium at

25�C, shaken at 120 rpm for 36 h. Then, 500 lL extracted

from each flask were spread on 10 plates with a Drigalsky

spatula. After incubating for 24 h, the bacterial biomass

was removed from each plate and placed in a 100 mL flask

containing a saline solution. Before the application, the

concentrated suspension was diluted in 1.5 L of sterile

distilled water to obtain a cell density of approximately

4 9 108 colony forming unit (UFC) 9 mL-1.

The bacterial suspension was inoculated directly onto

the substrate, which was composed of a mixture of peat-

perlite-vermiculite (40:40:20) at the moment of installing

the cuttings. The control treatment consisted of the appli-

cation of sterile distilled water. The assay lasted for

30 days, and it was conducted in the summer (February–

March). During that period, the cuttings were under a

periodic misting system and were applied fungicides and

fertilizers according to the specifications of the Bioforest,

Arauco company’s. The assay was randomly conducted; it

was made up of three treatments, two bacterial strains and

one control treatment with four replicates. The experi-

mental unit was composed of three mini-cuttings. To study

the population dynamics, three samples of each treatment

were collected weekly in order to extract the bacterial

DNA; each compound sample was composed of three mini-

cuttings.

Total bacterial DNA extraction

Three samples were carried out at weekly intervals. Due to

the evolution in the callus and root formation in the cut-

tings, the material collected in each sample collection

varied. Thus, we collected: material from the superficial

tissues of the mini-cutting base and the associated rhizo-

sphere after 7 days, material from the callus tissues after

14 days and material from roots after 21 days.

For all the types of tissue collected, the treatment was as

follows: the tissue was carefully cut, washed with sterile

distilled water and dried with sterile absorbent paper tow-

els. Then, 0.05 g of the tissues was ground in a mortar with

liquid nitrogen and then put in a 1.5 mL Eppendorf tube.


123

The total bacterial DNA was extracted according to the

modified protocol by Dong et al. (2006).

The samples were suspended in 300 lL of phosphate

buffer (0.1 M Na2HPO4–NaHPO4, pH 8) and gently vor-

texed. 50 lL of Lysozyme (10 mg/mL) were added to the

samples, and then these were incubated at 37�C for 30 min.

Subsequently, 200 lL of aluminium sulfate were added to

the solution, which was vortexed for 2 min. Later, 0.35 g

of crystal spheres (2 mm) and 250 lL of lysis buffer

[100 mM NaCl, 500 mM Tris [pH 8.0]; 10% (wt./vol.)

SDS] were added to the samples and these were taken to

the disruptor for 5 min at maximum speed. The samples

were centrifuged at 10.000 g for 30 s, the proteins were

removed from the supernatant by the addition of 250 lL of

phenol/chloroform/isoamyl alcohol (25:24:1). The samples

were once again centrifuged at 10,000g for 1 min. The new

supernatant was transferred to another tube and 250 lL of

chloroform/isoamyl alcohol (24:1) were added; the super-

natant was then incubated at 4�C for 5 min and centrifuged

at 10,000g for 1 min. The supernatant was transferred to

another tube in order to precipitate the DNA; 0.5 vol of

7.5 M ammonium acetate and 1.0 vol of isopropanol were

added. After incubation at -20�C for 15 min, the DNA

was centrifuged at 12,000g for 10 min, washed twice with

70% Ethanol (1 mL) and air dried. The pellet obtained was

dissolved in 50 lL of MiliQ water. The yield and purity of

the obtained DNA were determined by spectrophotometry

(NanoDropTM 1000) using the ratio of absorbance at

260/280 nm. As additional controls we used DNA from

pure strains of B. subtilis and S. maltophilia, which was

extracted as previously described.

Fragments of the gene coding for 16S rDNA were

amplified from the bacterial DNA samples by a PCR. The

reaction mixture contained 2 lL of DNA (5–40 ng):

5 9 buffer at a concentration of 19, 25 mM MgCl2,

10 mM DNTP; 10 lM concentration of primers 341F

[50-CCTACGGGAGGCAGCAG-30] and 907R [50-CCCTCA

ATTCMTTTGAGTTT-30], and 5 U/lL GoTaq polymerase.

The DNA amplification was carried out with an

Eppendorf MasterCycler gradient, using an amplification

program at 94�C for 5 min, followed by a cycle at 94�C for

30 s, 65�C for 45 s and 72�C for 1 min 30 s, 20 touchdown

cycles of 0.5�C; 10 cycles of 5 min at 94�C, 45 s at 55�C

and 1 min 30 s at 72�C, and a final extension cycle of

5 min at 94�C, 45 s at 55�C and 5 min at 72�C and then

cooling at 10�C. The amplified product of the first PCR was

once again amplified in a nested PCR, this time using

10 lM of primers 341F with GC-clamp [50-CCTACGG

GAGGCAGCG-30] and 534R [50ATTACCGCGGCTGC

TGG-30]; the bacterial 16S rDNA targeted primer pair has

been widely used for DGGE analysis of bacterial com-

munities (Muyzer et al. 1993); the mixture was concen-

trated to 50 lL; the temperature and other conditions were

similar to the first run. The resulting PCR product was

approximately 450 bp long. Gel electrophoresis was used

to analyze this product; it was run on a 1.2% (wt/vol)

agarose-gel prepared in TAE 19 and stained with ethidium

bromide.

For the DGGE analysis, fragments of the 341F ? GC-

clamp and 534R regions of the 16S rDNA were amplified.

To separate these fragments, a denaturing gradient (7 M

urea-40% formamide) was used, considering a denaturation

between 0 and 100% (0–45–75–100%), and using poly-

acrylamide gel (acrylamide/bis-acrylamide 37,5:1) in 19

TAE buffer (Muyzer et al. 1993). Approximately 25 lL of

the PCR product were applied to the individual lanes of the

gel and the run was performed at 60�C and 110 V for 14 h,

using a D-gene system (Bio-Rad Laboratories). The gels

were stained with ethidium bromide for 20 min and

washed twice with Milli-Q water before UV transillumi-

nation. The gels were photographed and processed using

Quantity One software, Version 4.2.1 Bio-Rad Laborato-

ries. The results were analyzed with the multivariate sta-

tistics software Primer V.5 (Primer-E Ltd, Plymouth, UK).

The similarity matrix was calculated using Bray-Curtis

similarity coefficient (Clarke and Warwick 2001). Based

on the similarity data a multidimensional scaling (MDS)

was constructed in order to compare the difference among

the treatments.

Quantification of the population of B. subtilis

and S. maltophilia in the rhizosphere, superficial tissues

of the stem-base and callus

The same DNA samples described in point 5 (b) were used.

The bacterial DNA concentration was determined by

spectrophotometer at 260 nm; the quality of the DNA

extracted was photometrically evaluated (NanoDropTM

1000) by calculating the A260/A280 ratio that is probably to

be found between 1.7 and 1.9. After being extracted from

the pure cultures, the bacterial DNA obtained was purified

using the Wizard� SV gel/PCR Clean-Up System (PRO-

MEGA) kit with incubation times from 1 to 5 min. Then,

the minicolumn was discarded. The DNA was examined

using real-time PCR; the 16S rDNA fragments were

amplified using the Bs 20–41 forward (50-CCGCGT

GAGTGATGAAGGTTT-30); Bs 99–118 reverse (50-GGT

GCCGCCCTATTTGAA-30) to B. subtilis (Goto et al.

2000) and St 164–185 forward (50-GCGTAGGTGGTCGT

TTAAGTC-30); St 246–267 reverse (50-CCACACTCTAG

TCGTCCAGTT-30) to S. maltophilia (Minkwitz and Berg

2001) primers according to the manufacturer’s instructions

(LightCycler� Faststart DNA Master SYBR Green I)

on a LightCycler� 1.0 Instrument system, software version

4.05 (ROCHE Applied Science). The DNA concentrations

were estimated by absorbance at 260 nm using a


123

spectrophotometer (NanoDropTM1000). Additionally, the

PCR products were verified by 2% agarose gel electro-

phoresis. The number of copies (copies/lL) of the PCR

products was determined using the following formula:

Molecules=lL ¼ sample concentration; ng=lLð Þ � Kð ÞPMNð Þ � amplicon length; bpð Þ

K = 6.022 9 1023 molecules/moles, PMN = 656.6 9

109 g/moles (molecular weight of nucleotide pairs).

The quantitative PCR was carried out using the Light-

Cycler� 1.0 Instrument and its corresponding software

(Version 4.05. Roche Applied Sciences). The PCR product

was measured using a fluorescence signal during the PCR

process. The number of Ct cycles at which the fluorescence

signal crosses a certain threshold is proportional to the

logarithm of the concentration of copies in the assay. The

calibration curves were constructed from a series of dilu-

tions of the standard solution with a copy concentration of

the known gene, starting from 108–102. The signal was

generated by the binding of the fluorophore SybrGreen to

double-stranded DNA. The amplification included four

stages: pre-incubation, 1 cycle at 95�C for 10 min;

amplification of 40 cycles, composed of 3 sub-stages:

denaturation at 95�C for 10 s, annealing at 55�C for 5 s and

extension at 72�C for 10 s; followed by the melting curve

performed in 1 cycle that comprised three sub-stages:

denaturation at 95�C for 10 s; annealing at 65�C for 15 s;

melting at 95�C for 10 s; and finally one cycle of cooling at

40�C for 30 s. PCR Grade water was used as a negative

control. To validate the PCR, the measurements were

performed in duplicate and the DNA extractions from

samples of the rhizosphere and superficial tissue of the

stem-base and callus of the mini-cuttings were performed

in triplicate. The concentrations of the gene copies were

calculated from the calibration curve adjusted by the

software.

Results and discussion

IAA and ABA production in both B. subtilis

and S. maltophilia

Both strains were able to synthesize IAA in vitro growth

medium with and without tryptophan. IAA levels in the

supernatants of the B. subtilis culture were low (2.61 ±

0.0180 lg/mL equivalent IAA) compared to those of

S. maltophilia (13.53 ± 0.0047 lg/mL equivalent IAA).

When tryptophan was added, IAA synthesis increased in

both strains to 13.53 ± 0.0047 lg/mL and 23.6 ± 0.026

lg/mL IAA-equivalent, respectively. These results were

expected since tryptophan is the main precursor of auxins

that favors IAA production by rhizobacteria and improves

root proliferation (Asghar et al. 2004; Khalid et al. 2004).

IAA levels obtained for B. subtilis in the absence of trypto-

phan coincides with other results obtained for other Bacillus

species. Karadeniz et al. 2006 quantified 2.15 ± 0.35 lg/

100 mL IAA-equivalent in stationary phase for B. megate-

rium. Araujo et al. (2005) indicates the potential of using

selected strains of B. subtilis due to the effect they have with

the synthesis of phytohormones and some unidentified

metabolites on growth promotion. IAA could be synthesized

by multiple pathways in a bacterial cell thanks from the

precursor tryptophan, which can be originated from

the degradation of roots and microbial cells and from root

exudates (Kravchenko et al. 2004). It should be noted that the

concentration of bacterial auxin produced depends on the

response and physiological development of the plant due to

endogenous hormone levels of its host, which may vary

according to the genotype and age of the plant (Ahmad et al.

2005). It has also been proposed that the biosynthesis of

auxins in bacteria would be a strategy to detoxify tryptophan

excesses in the rhizosphere (Bar and Okon 1993). Retention

times were 10.95 and 16.98 min for IAA and ABA, respec-

tively. The production of auxin was confirmed by HPLC

analysis, and it was only detected and quantified in

S. maltophilia (Fig. 1a).

Higher rates of tryptophan assimilation increase the

production of auxins in Azospirillum brasilense and other

bacteria (Martınez-Morales et al. 2003). It could be

suggested that there exists a synthesis pathway independent

of tryptophan in S. maltophilia, which produces IAA in

the absence of tryptophan. This pathway would permit to

detoxify the excess of tryptophan in the rhizosphere. For

the species studied, there are previous references about the

IAA production and its relationship with growth promo-

tion. S. maltophilia strains isolated from different clinical

and environmental sources produced IAA levels ranging

from 5.2 to 0.7 lg/mL. When environmental sourced

strains were applied to strawberry plants, these obtained

Without Trptophan

With Tryptophan

0

2

4

6

8

10

S. maltophiliaB. subtilis

IAA

(u

g/m

l)

Fig. 1 a Detection of IAA in B. subtilis and S. maltophilia cultures

grown with and without tryptophan (0.025 M) after 48 h by using

HPLC analysis


123

longer and better quality roots (Suckstorff and Berg, 2003).

B. subtilis strains secreting 0.7 lg/mL IAA increased

rooting in 6.8% and root biomass in 106.7% in minicut-

tings of an E. grandis 9 E. urophylla hybrid (Teixeira et al.

2007).

IAA production of S. maltophilia, determined by Sal-

kowski’s method, was of 23.6 lg/mL, three times higher

than that obtained using HPLC (8.7 lg/mL). These varia-

tions have already been detected by other authors, for

example for A. brasilense, where colorimetric assay

detected 26.1 lg/mL IAA, and HPLC detected 0.5 lg/mL

(Crozier et al. 1988). No synthesis of IAA was detected in

B. subtilis, although this strain increased

Eucalyptus globulus rooting (Dıaz et al. 2009), sug-

gesting that other mechanisms of action may be involved in

the process of root formation.

Probably S. maltophilia stimulates rooting, leading to a

physiological change in the plant by synthetizing low IAA

concentrations and by protecting the plant from important

pathogenic microorganisms, producing an antagonistic

effect as mentioned by Berg et al. (1994) and Dunne et al.

(2000). Along with this, S. maltophilia would avoid the

need to use a synthetic auxin source in clonal propagation

programs of recalcitrant species. Furthermore, according to

Van de Broek et al. (1999), high levels of IAA production

in vitro could be considered as a useful parameter to select

native strains of A. brasilense with plant-growth promoting

activity.

None of the bacterial strains produced ABA, which may

have different effects on plants roots, either positive or

negative. Studies have determined that exogenous appli-

cation of ABA would promote rooting (Yasmin et al.

2003). In contrast, it has also been proven that the appli-

cation of ABA would also act by inhibiting or neutralizing

the root formation (Pilet and Saugy 1987). It has also been

demonstrated the ability of Bradyrhizobium japonicum to

synthesize ABA (0.02 lg/mL), indicating that the inocu-

lation of this bacteria in plants growing in soils under salt

stress conditions would induce some tolerance; these bac-

teria are called plant stress homo-regulating rhizobacteria

(Araujo et al. 2005).

In this work, both strains produced IAA in different

concentrations under in vitro conditions. However, the

increase in bacterial population in the rhizosphere and the

continuos release of small amounts of IAA could enhance

rhizogenesis. Other indole compounds such as indole

pyruvic acid, indole acetamide acid, and indole carboxylic

acid might be involved in root formation. The ability of

S. maltophilia and B. subtilis to increase rooting, fibrosity

and biomass of E. globulus plants may depend on the

endogenous levels of IAA in the plant and those produced

by the respective bacteria.

Detection and determination of siderophores

According to the CAS assay and in comparison with the

negative control both strains produced siderophores.

B. subtilis produced a yellow halo larger than that of the

negative control, where discoloration did not occur. In the

positive control, B. stearothermophilus showed a larger

halo after 48 h of incubation, whereas S. maltophilia dis-

played a clear halo around the sample.

Some plant-growth promoting bacteria produce sidero-

phores by sequestering the limited supply of iron in the

rhizosphere, especially in alkaline and neutral soils, and

thereby reduce and inhibit the ability of some pathogens to

grow (Sharma and Johri 2003). The role of these bacteria

has also been demonstrated in the induction of resistance in

different plant-pathogen systems (Ardon et al. 1998).

According to the coloration obtained in the medium

using the overlay technique, B. subtilis produced

hydroxamate-type siderophores, similar to what was

reported previously for B. cereus strains (Perez-Miranda

et al. 2007). In S. maltophilia, a slightly pale yellow color

was observed, which had not been reported for this test

before. According to Perez-Miranda et al. (2007) this color

may be due to carboxylate-type siderophores, which would

indicate a high sensitivity of S. maltophilia to low sidero-

phore concentrations.

Hydroxamates are produced by fungi and bacteria,

whereas catecholates are produced exclusively by bacteria.

a-Carboxylates are produced by fungi of the Zygomycetes

group and a few bacteria such as Rhizobium meliloti and

Staphylococcus hycus (Baakza et al. 2004).

Methylobacterium spp and B. subtilis strains produced

hydroxamate-type siderophores (Lacava et al. 2008). It has

been demonstrated that one strain can produce more than

one type of siderophore, thus Burkholderia cepacia strains

produce different types of siderophores, namely ornibactin

and cepaciachelin, hydroxamate- and catecholate-type

siderophores, respectively (Barelman et al. 1996). This

ability to produce siderophores has also been commonly

associated with the capacity of biocontrol shown by

Pseudomonas fluorescens strains. Inhibition in the germi-

nation of chlamydospores of Fusarium oxysporum was

correlated with the synthesis of siderophores in vitro

(Hernandez et al. 2004)

Well known is the ability of S. maltophilia to suppress

fungal diseases by the production of antibiotics (Jakobi

et al. 1996), competition for nutrients through siderophores

and extracellular enzymatic activity (proteases, glucanases

and chitinases) (Suckstorff and Berg 2003). This charac-

teristic to produce carboxylate-type siderophores could

determine a potential of S. maltophilia for the biocontrol of

diverse leaf and root pathogens for Eucalyptus spp.; this


123

characteristic could also be associated with plant rooting

and growth.

Determination of microbial diversity in the rhizosphere

and tissues of E. globulus mini-cuttings inoculated

with B. subtilis and S. maltophilia

At diverse sampling times, differences were observed in

the profiles of the bacterial community associated with the

rhizosphere and superficial tissues of the stem-base and

callus of E. globulus minicuttings inoculated with both

strains. The presence of permanent, dominant populations

was observed (darkest bands) as well as the presence of

populations that appear and disappear; these populations

would indicate that a certain degree of succession occurs in

the bacterial community of the rhizosphere during the

rooting process (Fig. 4).

Patterns between the treatment inoculated with

S. maltophilia (IB lane) and the control treatment (T lane)

showed a relative abundance of species similar and higher,

respectively, than the treatment inoculated with B. subtilis.

A bacterial species present in all the profiles (Fig. 2)

could correspond to a highly competitive species with the

ability to adhere to and remain in roots. Identifying this

type of bacteria could represent a strategy to detect possible

root-growth promoting agents.

Bacillus subtilis population decreases over time

(IA lane), along with a part of the associated bacterial

community (Fig. 2). In general, some reports indicate that

Gram-positive bacteria are dominant in the rhizosphere of

plants such as chrysanthemum, barley, etc. (Smalla et al.

2001; Duineveld et al. 2001). It would suggest that this fact

may be an antagonistic effect between the indigenous

species of the rhizosphere of E. globulus and B. subtilis,

since this population is characterized by producing a higher

amount of secondary metabolites and siderophores, a fact

which affects the structure of the microbial community and

the interaction of pathogens in the plant rhizosphere. These

compounds are first secreted by binding to the root tips

along with sugars and organic acids that can quickly be

used by other microorganisms, either for their benefit or

detriment.

According to the profile, S. maltophilia (IB lane) was

able to remain in the rhizosphere of E. globulus and thus

had DNA profiles similar to those of the control treatments.

From this, it can be inferred that the associated bacterial

community is not perturbed by the introduction of this

strain. Marilley and Aragno (1999) argue that Gamma-

proteobacteria (S. maltophilia) predominates more in the

rhizosphere than Gram-positive bacteria. Soderberg et al.

(2004) reported that this group of Gram-negative bacteria

predominate in the rhizosphere due to their reproductive

rate and high activity under nutrient-lacking conditions.

However, other studies indicate that Gram-positive bacteria

would be more abundant in roots (Smalla et al. 2001).

A factor that may influence the differential detection of one

group over another is the selection of molecular methods,

especially the efficiency of DNA extraction method and

PCR experimental conditions which might explain in part

the conflicting results.

Both strains were isolated from the rhizosphere of

Eucalyptus spp. plants and inoculated in the same type of

substrate in order to promote rooting of E. globulus mini-

cuttings. However, colonization assays were performed on

other type of substrate and inoculated in another E. glob-

ulus clone. According to Ramos et al. (2003), bacteria are

more effective when inoculated into the soil or substrate

where they were isolated from. If did so, they should use

nutrients in a better way and maintain the balance within

the indigenous community after inoculation. Therefore,

adaptation and competence of bacteria in the soil envi-

ronment play an important role and thus it is advisable to

use bacteria in the same niche from which they were

isolated. The rhizosphere of plants is a dynamic environ-

ment in which many factors may affect the structure and

composition of species belonging to the microbial com-

munities that colonize the roots. Thus, for rhizobacteria to

play their role of promoting plant growth and rooting and

contributing to the biological control, they should be able

to remain in the rhizosphere of inoculated plants. Consid-

ering that 1,000 bacterial species can coexist in one gram

A B IA1 IB1 T1 IA2 IB2 T2 IA3 IB3 T3

Fig. 2 PCR-DGGE profiles of bacterial communities associated with

the rhizosphere, stem base tissues and callus tissues of E. globulusminicuttings. Control B. subtilis (a) and S. maltophilia (b) strains.

Uninoculated control treatment (T1, T2 and T3). Cuttings treated with

B. subtilis (IA) and S. maltophilia (IB). Sampling times on days 7(1),

14(2) and 21(3). The circles show the bands of pure cultures. The

arrow shows a band that may represent a highly competitive species


123

of soil (Torsvik et al. 1990) it is clear that the bacterial

community DGGE profiles represent a selective proportion

of the total community of the rhizosphere. The value of

DGGE and other detection techniques in soil microbiology

is a way to know, identify and measure the microbial

diversity in soils because most bacteria have never been

cultivated (Peixoto et al. 2002).

Based on the PCR-DGGE profiles for B. subtilis (A) and

S. maltophilia (B) inoculated in the minicuttings, there was

a similarity of 78% in the rhizosphere of plants inoculated

with B. subtilis compared with that of uninoculated plants.

As the root formation process was progressing, this rate

decreased to 51%, which indicates that the population of

B. subtilis decreased (Fig. 3). On the contrary, S. malto-

philia showed an 83% similarity between the inoculation

and the first week; this indicates its ability to adapt and

remain in roots without significantly altering the natural

microbial community of the rhizosphere (Fig. 3).

A divergence was verified between the samples grouped

in S. maltophilia (Fig. 4), which had fewer differences

from the moment of inoculation than B. subtilis. This

indicates that there are variations between the respective

microbial communities, which have positive or negative

effects on the original microbiota. However, the results of

the electrophoretic profiles of control strains were homo-

geneous and allowed us to clearly assess the ecological

disruptions that could have caused coinoculation of the

strains selected. Furthermore, other authors have demon-

strated that the genome size and the number of copies of

16S rDNA genes could affect the performance of the

partial amplification of 16S rDNA fragments leading a

bacterial community. Consequently, the current diversity

of the microbial complex could be overestimated (Vallaeys

et al. 1997).

Quantification of the population of B. subtilis and

S. maltophilia in the rhizosphere, superficial tissues

of the stem-base and callus

The standard curve constructed to determine the number of

copies of the specific gene in unknown samples was

generated from specific-DNA products amplified from

B. subtilis and S. maltophilia, and adjusted in a seven

dilution range from 108 to 102 per 2 lL of a template spe-

cific to both strains. The slopes of the curve were -5.1557

and -3.7761 and its yield was 1.5 and 1.8, respectively.

Real time PCR has been applied to different studies, e.g.

the detection of sulfate-reducing bacteria such as Desul-

fotomaculum lineage 1 in rice field soils (Stubner 2002). As

proved by Ruppel et al. (2006), we also demonstrated the

possibility of quantifying the number of specific 16S rDNA

gene copies of bacteria associated with plants in original

undiluted DNA samples.

Melting curve analyses of the selected strains show a

peak when comparing the PCR products of experimental

samples with those of standard samples. All the experi-

mental samples showed a Tm of 83 and 85�C, identical to

that of standard samples. These results confirm that

amplification of primers was specific for each strain. PCR

amplicons were analyzed in 2% agarose gel by standard

horizontal gel electrophoresis. The results show a 78 bp

fragment for B. subtilis and an 82 bp fragment for

S. maltophilia (results not shown).

Based on the specific DNA concentration of standard

samples, when transferring the results of the number of 16S

rDNA gene copies to the B. subtilis cell number (six 16S

IA1

T1

IA2

T2

T3

IA3

60 70 80 90 100

Similarity

B3

T2

B1

B2

T1

B3

60 70 80 90 100

Similarity

a

b

Fig. 3 Similarity dendrograms showing the comparisons between

profiles for: a B. subtilis (IA), b S. maltophilia (IB) and the

uninoculated control treatment (T) at different sampling times: 7(1),

14(2) and 21(3) days

IA1

IB1

T1

IA2

IB2

T2

IA3

IB3

T3

Stress: 0.11

Fig. 4 MDS based on the Bray-Curtis similarity matrix for DGGE

samples when inoculating B. subtilis and S. maltophilia in E. globulusminicuttings. B. subtilis strain (IA); S. maltophilia strain (IB);

uninoculated control treatment (T). Sampling times on days 7(1),

14(2) and 21(3). (Stress = 0.11)


123

rDNA gene copies per cell) and measuring the concentra-

tion of DNA extracted per mg of rhizosphere soil or

superficial tissues of the stem-base and callus of minicut-

tings, cell densities of 8.7 9 106, 4.3 9 106 and 1.9 9 107

cells/mg of rhizosphere soil, or stem base and/or callus

tissues were obtained, respectively. Cell densities of

4.7 9 106, 2.1 9 107 and 2.0 9 107 cells/mg of rhizo-

sphere soil or stem base and/or callus superficial tis-

sues collected after 7, 14 and 21 days were found for

S. maltophilia (Fig. 5b).

The results showed an increase in the total bacterial col-

onization of treated plants, compared with that of untreated

plants (Figs. 5a, b). This could be due to a better bacterial

adherence and a competitive ability in the root against the

native bacterial community. Populations of Pseudomonas

spp. in the rhizosphere increased from 1.3 9 106 to

5.3 9 106 at 60 and 120 days after inoculation, whereas the

control treatment increase ranged from 0.7 9 106 to

2.6 9 106 (Viswanathan and Samiyappan 2007).

In this study we demonstrated the applicability of using

new techniques such as DGGE and real-time PCR with

SybrGreenTM (SG) in a forest species, and we could

quantify the population dynamics of B. subtilis and

S. maltophilia in a wide range of native bacteria in rhizo-

sphere and superficial tissues samples of the stem base and

callus of the minicuttings.

In general, shortly after introducing B. subtilis in the

substrate, the population rapidly decreased (Fig. 5a). To

assure a beneficial effectiveness of the microorganism, this

must be established in large numbers in the rhizosphere of

the inoculated plant and it shall maintain the levels of

colonization during the phase in which microorganisms

exert the action for which they were applied (Couillerot

et al. 2010). Thus, a common feature of each inoculants is

characterized by its ability to release the appropriate

number of viable cells in good physiological conditions at

the time of inoculation (Ruppel et al. 2006) .

The bacterial population ranged between 106 and 107

cells/mg of rhizosphere, superficial tissues of the stem-base

and callus in E. globulus minicuttings, similar to what has

been obtained in other studies in which rhizospheric pop-

ulations ranged from 106 to 108 cells/mg (Gardener and

Weller 2001; Ruppel et al. 2006). Nevertheless, in this

study it was not possible to determine a relationship

between the total cells of the bacteria applied and the total

bacterial population since no previous information about

native bacterial populations inhabiting the rhizosphere of

E. globulus minicuttings was available.

These preliminary results allow us to conclude that

S. maltophilia has a high capacity to establish and prolif-

erate in the rhizosphere or in superficial tissues of the stem-

base and callus of E. globulus minicuttings. This consti-

tutes a tremendous advantage in the potential case of the

strain being used as a biofertilizer. Moreover, its ability to

produce auxins and siderophores could explain its effi-

ciency in promoting rooting.

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Production of phytohormones, siderophores and population fluctuation of two root-promoting rhizobacteria in Eucalyptus globulus cuttings

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