influence of an indigenous european alder (alnus glutinosa (l.) gaertn) rhizobacterium (bacillus...

New Forests 25: 149–159, 2003. 2003 Kluwer Academic Publishers. Printed in the Netherlands.

Influence of an indigenous European alder (Alnusglutinosa (L.) Gaertn) rhizobacterium (Bacilluspumilus) on the growth of alder and its rhizospheremicrobial community structure in two soils

´ ´*BEATRIZ RAMOS , JOSE A. LUCAS GARCIA, AGUSTIN PROBANZA,˜JEZABEL DOMENECH and F. JAVIER GUTIERREZ MANERO

Facultad CC. Experimentales y de la Salud, Universidad San Pablo CEU, PO Box 67, 28660 Boadilla*del Monte, Madrid, Spain; Author for correspondence (e-mail: [email protected]; phone: 34 91

3724785/33; fax: 34 91 3510496)

Received 5 October 2001; accepted in revised form 5 September 2002

Key words: Biostimulant, Inoculation, PGPR, PLFAs, Rhizosphere structure

Abstract. European alder seedlings were inoculated with a suspension of the putative plant growthpromoting rhizobacterium (PGPR) Bacillus pumilus (CECT 5105), or left non-inoculated (controls) intwo different soils, and grown under controlled conditions. Soil A showed a thick texture, slightly acidicwith a high mineral nitrogen content, while soil B showed a thin texture, basic and with a lower nitrogencontent. At each sampling time, over an 8-week period, shoot and root systems of the plants weremeasured, nodules counted, and shoot and root length and surface were determined. In addition, changesin the microbial rhizosphere structure were evaluated by the phospholipid fatty acid (PLFA) profileextracted directly from the rhizosphere soil. The increases detected in shoot surface were significant onlyin soil A, while the root system was affected in both soils. In soil A, inoculation with B. pumilus caused aperturbation that subsequently disappeared, while the rhizosphere community structure was seriouslyaltered in soil B. All biometric parameters were enhanced to a greater extent in soil A, in which theinoculum did not alter the existing rhizosphere communities and nutrient availability was better.

Abbreviations: PGPR – plant growth promoting rhizobacteria, PLFAs – phospholipid fatty acids,˜´CECT – Coleccion Espanola Cultivos Tipo (Spanish type culture collection)

Introduction

Plant growth promoting rhizobacteria (PGPR) are naturally occurring soil bacteriathat colonise plant roots and stimulate plant growth when applied to roots, tubers orseeds (Kloepper et al. 1980). Various mechanisms may be involved, such as therelease of metabolites (auxins, gibberellins, cytokinins or ethylene) that directly

˜stimulate growth (Frankenberger and Arshad 1995; Glick 1995; Gutierrez Maneroet al. 2001). Indirect mechanisms involve production of metabolites that affect otherfactors in the rhizosphere, resulting in enhanced plant growth (Kloepper 1993; DeFreitas et al. 1997). The best known mechanisms in this group are inhibition ofdeleterious rhizobacteria and plant pathogens, and the release of either siderophoresand/or antibiotics or lytic enzymes or HCN (Scher and Baker 1982; Shanahan et al.1992; Frilender et al. 1993).

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Inoculation of agricultural crops with PGPRs has been done for many years withencouraging results. However, except for possible disease control in forest nurseries(Chanway 1997), research on trees has just begun (Enebak et al. 1997). Most studieson growth promotion of trees have been made on gymnosperm species (O’Neill etal. 1992; Chanway and Holl 1993; Bent et al. 2000; Enebak et al. 2000; Probanza etal. 2000; Shishido and Chanway 2000). The only study concerning angiosperm treesis that of Mohammad and Prassad (1988), who coinoculated Eucalyptus globuluswith two bacteria, a Bacillus strain and an Azotobacter.

In addition to their PGPR trait(s), the inoculated microorganisms must berhizosphere competent and able to survive in soil (Cattelan et al. 1999). One of themain limitations in the application of PGPRs as biostimulants for the improvementof plant productivity is that a certain degree of specificity may exist between theplant species and the bacterial strain. Nevertheless, there are many studies thatsupport the nonspecificity of many PGPRs with regard to the plant species on whichtheir stimulating effect is exerted (Enebak et al. 1997; Bent et al. 2000).

Another limiting factor is the influence that soil may have on the process, eitherbecause of soil properties, or because of the existing bacterial community, whichalready has a sophisticated network of established interactions that may or may notbe altered (Pierson and Pierson 2000).

The purpose of our study was to determine, under laboratory conditions, theability of a Bacillus strain (B. pumilus CECT 5105), indigenous to the rhizosphereof European alder (Alnus glutinosa (L.) Gaertn), to promote growth and to inducechanges in the rhizosphere structure after inoculation. The effects were assesed intwo soils i) the indigenous soil from which the bacterium was isolated and ii)another soil, in which the inoculated strain would interact with a new rhizospherecommunity in a substrate with different physicochemical characteristics.

Materials and methods

Bacterial strain

B. pumilus (CECT 5105) was isolated from the rhizosphere of European alder(Alnus glutinosa (L.) Gaertn) (Probanza et al. 1996). The bacterial culture medium,free of bacteria, showed a strong growth promoting activity on alder seedlings

˜(Probanza et al. 1996). Auxin-like compounds (Gutierrez Manero et al. 1996) and˜gibberellins (Gutierrez Manero et al. 2001) were characterised in culture medium.

Soils

Two soils were used. Soil A was collected from the same alder stand from which thebacteria was isolated (Probanza et al. 1996), located in San Martin de Valdeiglesias(Avila, Spain). Its texture was 53% sand, 24% clay, 23% silt, with pH 6.36, 0.5 mg

1 2NH /g soil, 9.4 mg NO /g soil, 2.1% organic carbon and a maximum water4 3

holding capacity (WHC) of 0.5 mL/g. Soil B was collected from experimental plots

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at the University of San Pablo CEU (Madrid, Spain); the texture was 29% sand, 41%1 2clay, 30% silt, with pH 8.01, 0.24 mg NH /g soil, 4.05 mg NO /g soil, 0.2%4 3

organic carbon and a maximum WHC of 0.36 mL/g. The experimental plots hadonly been used to grow a legume (Vicia villosa) for one year and lay fallow the nextyear, after which sampling was done.

Plants

Alder seeds were collected from the alder trees in the stand from which bacteriawere isolated. After 15 d at 80% RH and 4 8C, the seeds were surface-sterilised in asolution of sodium hypochlorite:sterile distilled water (4:1, v /v) for 1 min, followedby three rinses with sterile distilled water. Then the seeds were spread on sterilevermiculite (termite no.3) at maximum WHC, in trays covered with vented film andkept in a culture chamber (16 h light /8 h dark; photon flux density (400–700 nm)

22 21150 mmol m s at pot surface; temperature, 22 8C/18 8C; relative humidity 70%)until the first two true leaves developed (approximately after one month). Sub-sequently, two hundred seedlings were transplanted to nursery pots (14.5 3 4.5 3

4.5 cm, 200 g capacity), filled with vermiculite and grown until four mature leavesdeveloped (approximately after another two months). During this period, wateringwas done alternately with Hoagland solution (Hoagland and Arnold 1950) anddistilled water (DW). At this time, pots were filled with Soil A or B, and a total of100 seedlings were transplanted, one per pot. Fifty were selected for inoculation andfrom then on, plants were watered with DW. After 10 days in the new substrate,seedlings were ready to be inoculated.

Inoculum

Bacteria were grown in nutrient broth (Pronadisa, Spain), incubated in an orbitalshaker at 300 rpm for 24 h at 28 8C. The culture medium was centrifuged for 30 min,and resuspended in sterile DW. Bacterial growth was determined by turbidimetry at

8660 nm to achieve 10 cfu /g soil occupied by the root system at inoculation.Fifty three-month-old seedlings were transplanted to pots filled with Soil A and

the other fifty to Soil B. Half of the plants in each soil were inoculated at the stage of4 /5 mature leaves. Watering was suspended two days before and two days afterinoculation.

Plant harvest

Sampling times were 24 h, 2 wk, 6 wk and 8 wk after inoculation (wk.a.i.). At eachsampling time, three seedlings from each treatment were selected at random and thefollowing analyses were made.

Seedling growth

Seedlings were carefully removed from pots and plants were gently flattened and

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pressed for biometric analysis. Leaf surface area (LSA), shoot length (SL), rootsurface area (RS), root length (RL) and nodules of all plants, at all samplingmoments were measured with a Delta T image analysis system, with DIAS software.

Soil sampling and phospholipid fatty acid (PLFA) extraction

Before flattening, 1 g of the soil adhering to the roots was resuspended in 1.5 mL ofsterile DW and kept at 220 8C for PLFA analysis (total PLFAs). Extraction was

˚done according to Frostegard et al. (1993) based on the methodology proposed byBligh and Dyer (1959). The extracted lipids were fractionated on silicic acidcolumns (60–200 pore size, Sigma, St. Louis, MO) by eluting with chloroform,acetone and methanol. The phospholipid fraction was subjected to a mild alkalinemethanolysis (Dowling et al. 1986) which transformed the fatty acids of thephospholipids into free fatty acid methyl esters. Fatty acids were named andidentified using gas chromatography, according to the methods described by

˚Frostegard et al. (1993). All solvents were HRGC-grade.Fatty acids were designated as the total number of carbon atoms: the number of

double bonds, followed by the position of the double bond from the methyl end (v)of the molecule. C and t indicate Cis and trans-configurations, respectively. Theprefixes a and i indicate anteiso and iso branching, respectively; br indicates an

thunknown methyl branching position, 10Me indicates a methyl group on the 10carbon atom from the carboxyl end of the molecule and cy refers to cyclopropanefatty acids.

Statistics

A two-way analysis of variance (ANOVA) with replicates and two variation factors(sampling moments and inoculum) was made for each biometric variable evaluatedin each soil. When differences were significant, averages were compared using theLSD test (Sokal and Rohlf 1979).

Two Principal Component Analyses (PCA) were made (Harman 1967) with datafrom PLFAs extracted directly from the soil (total PLFAs) from the rhizosphere ofinoculated and non-inoculated plants, grown in soils A (Figure 1a) and B (Figure1b). For each PCA, data were organised in a matrix of 8 columns (4 sampling times3 2 treatments (B. pumilus and control) and 28 lines, corresponding to the fattyacids analysed.

Results

Seedling growth

Data from shoot and root systems as well as the number of nodules of seedlingsgrown in soils A and B are shown in (Table 1).

In soil A, the leaf surface area of inoculated seedlings was double that of controls

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Figure 1. (a) Principal component analysis of total PLFA-profiles from the rhizosphere of alder seedlings24 h, 2, 6 and 8 wk after inoculation with Bacillus pumilus (BP, solid dots) or non-inoculated controls (C,empty dots), grown in soil A. (b) Principal component analysis of total PLFA-profiles from therhizosphere of alder seedlings 24 h, 2, 6 and 8 wk after inoculation with Bacillus pumilus (BP, solid dots)or non-inoculated controls (C, empty dots), grown in soil B.

at the end of the trial (8 wk). In the root system, differences were non-significant atthe end of the trial, although 6 wk after inoculation, root length and surface ofinoculated seedlings were twice that of controls, and differences were significant.Nodules appeared in the root system of inoculated seedlings 2 wk after inoculation,

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Table 1. Shoot and root growth of alder seedlings grown in Soil A and Soil B, 24 h, 2, 6 and 8 weeks (wk)after inoculation with Bacillus pumilus (BP) or non-inoculated (C). Data are the average of threereplicates 6 standard error. NS: non significant according to ANOVA.

SOIL A SOIL B2Leaf Surface area (cm )

24 h 2 wk 6 wk 8 wk 24 h 2 wk 6 wk 8 wkC 3.6 6 0.3 6.6 6 2.2 30.7 6 3.2 50.7 6 7.9 5.9 6 0.6 6.2 6 0.9 37.3 6 7.4 41.4 6 1.5

BP 5.6 6 0.92 10.3 6 2.4 51.3 6 0.1 93.7 6 0.6 5.6 6 0.4 10.6 6 1.7 51.1 6 0.8 49.9 6 0.1LSD LSD 5 15.05 NS(a50.01)

Shoot length (cm)


BP 4.8 6 0.1 4.8 6 0.5 8.1 6 0.7 12.0 6 0.3 3.0 6 0.4 3.1 6 0.2 9.6 6 0.3 9.9 6 0.2LSD NS NS

2Root Surface Area (cm )


BP 4.3 6 0.97 11.7 6 5.3 27.8 6 5.6 47.5 6 5.2 2.3 6 0.2 6.1 6 0.7 21.7 6 0.1 35.4 6 2.7LSD LSD 5 12.56 LSD 5 7.93(a50.01) (a50.01)

Root Length (cm)


BP 19.9 6 1.0 45.2 6 19.6 104.4 6 19.6 151.7 6 15.5 6.18 6 0.5 24.1 6 9.8 97.2 6 6.5 150.5 6 18LSD LSD 5 43.82 LSD 5 44.22(a50.01) (a50.01)

Nodule number

24 h 2 wk 6 wk 8 wk 24 h 2 wk 6 wk 8 wkC 0 0 1 7 0 0 0 0

BP 0 1.1 6 17 0 0 0 0

while in non-inoculated samples, nodules appeared after 6 wk. At the end of thetrial, the inoculated seedlings had twice as many nodules as the controls.

In soil B, although the shoot system was not significantly affected, the root systemshowed significant differences in root surface area. It should be pointed out thatsignificant two-fold increases in root surface area and length were also found ininoculated plants as compared to controls after 6 wk of the experiment, and that theRSA was significantly higher at the end of the trial. In this soil, no nodules weredetected.

Phospholipid fatty acid analysis (PLFA)

Representation of the PCA made with data from total PLFAs from soil A (Figure 1a)shows that the inoculated sample separates from the non-inoculated control 24 hafter inoculation, along axis I, which accounts for 85.4% of the variance. Samplesfrom all other sampling times group together towards the positive ends of both axes,which together account for 90.14% of the variance. Loading factors with the highest

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weight values on both axes appear in the figure. The most abundant phospholipidswere 15:0, br16:0, br17:0, i17, Me18 and 19:1a.

In soil B, total PLFAs (Figure 1b) show how controls and inoculated samplesmerge towards the positive end of both axes in the two first wk of the experiment;both axes account for 79.42% of the variance. However, data for inoculatedseedlings at 6 and 8 wk, markedly separate from all other samples along axis I,which accounts for 63.5% of the variance. Loading factors with the highest weightvalues on both axes appear in the figures. The most abundant phospholipids were15:0, br16:0, 16:1v5, Me18 and 19:1a.

Discussion

Studies on plant growth promotion by microorganisms may be designed to studyonly the effects on growth (Enebak et al. 1997), or relating plant growth withcolonisation (Holl and Chanway 1992; Shishido et al. 1995; Schlotter et al. 1997)

˜and mechanisms of growth promotion (De Freitas et al. 1997; Gutierrez Manero etal. 2001). It is widely accepted that to affect the plant’s physiology, a closeinteraction with the plant must be established for a certain period of time (Chanwayet al. 1991).

Plant growth was better in soil A than in soil B (Table 1). The differentnodulation pattern detected in either soil (Table 2) should be considered to partiallyexplain differences in growth. The natural presence of Frankia in soil A explains thenodulation of seedlings only in this soil, contributing to the better results ascompared to soil B. In soil A, Bacillus pumilus seems to stimulate nodulation, sincenodules appear 2 wk after inoculation in inoculated seedlings, while in controls thefirst nodules show after 6 wk. Although no Bacillus strain has been reported tobenefit Frankia nodulation of alder, some Pseudomonas strains have been reportedto enhance nodulation of Frankia in this species (Knowlton et al. 1980), as well asthe Rhizobium-legume symbiosis (Frankenberger and Arshad 1995; Dashti et al.1998). This PGPR trait seems to be due to modulation of the plant hormonalbalance, (Atzorn et al. 1988) since production of plant hormones by this strain has

˜been reported (Gutierrez Manero et al. 1996, 2001).Production of auxin-like compounds could explain the increases in root growth

(Selvadurai et al. 1991). Auxins affect both root growth and pattern and increaseroot-soil surface, improving nutrient and water absorption potential (Germida andWalley 1996). This improvement can strongly influence plant growth capacity,subject to nutrient availability, which is lower in soil B. However, the improvementof plant nutrition is not sufficient to explain the increases detected in shoot surface insoil A in such a short period, and this striking increase may be attributed to the

˜gibberellins produced by this strain (Gutierrez Manero et al. 2001).In this sense, the question of the substrate-dependence for gibberellin synthesis on

culture media has been demostrated for other microorganisms (Janzen et al. 1992;Bastian et al. 1998). This substrate-dependence could be extrapolated to the soilsassayed, to partially explain the different effects of this strain on the shoot system.

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The bacterium is effective only in the original soil where, despite the better nutrientavailability, there may be a nutritional factor, which is absent in soil B, limiting thebacterial capacity to synthetise growth regulators in the latter. This hypothesis issupported by other authors, who have reported that inoculation with bacterial strainsin different soils may result in loss of a certain PGPR or biocontrol ability, such asIAA synthesis (Frankenberger and Arshad 1995) or antibiotic synthesis (Elsherifand Grossmann 1994).

Despite any PGPR trait that the inoculated strain may exhibit in vitro, thestimulating effect on growth has to be demonstrated in soil, in which the existingrhizobacterial communities play a definitive role. The indigenous communities havealready established an equilibrium, which should be mantained after the inoculation,so the bacteria introduced can exert their beneficial effect without disrupting theequilibrium (Pierson and Pierson 2000). Therefore, the relevance of adaptational

¨and competence phenomena (Wiehe and Hoflich 1995; Chabot et al. 1996) toimprove plant production with PGPRs is a key point.

The phospholipid fatty acid analysis (PLFA) was used to detect changes in themicrobial community structure following rhizosphere inoculation, comparing thePLFA profiles of the total microbial population (viable, nonviable, culturable andnonculturable) extracted directly from soil (total PLFAs), of inoculated and non-inoculated seedlings. The arrangement of samples in (Figure 1a) (soil A) reflects themarked alteration that has taken place in the structure of the rhizosphere communityimmediately after inoculation. This perturbation disappears within a 2 to 6 wkperiod, as shown by clumping of inoculated and non-inoculated samples 6 and 8 wkafter inoculation. Conversely, introduction of inoculum in soil B (Figure 1b) washardly reflected at first, since samples 24 h and 2 wk after inoculation were together,which could suggest a failure in the inoculation. However, the marked perturbationreflected by the separation of samples after 6 and 8 wk confirms that inoculation wassuccessful and that the perturbation must be caused by the exogenous population. Insoil A, the most abundant PLFAs are characteristic of gram positive bacteria(branched fatty acids), while in soil B, monounsaturated fatty acids, characteristic ofgram negative bacteria, and branched fatty acids (Steinberger et al. 1999) showedthe same levels.

In view of the better PGPR efficiency of B. pumilus in soil A, its indigenous soil,the competitiveness of this strain in the original soil has been evidenced. This maybe because the inoculation does not affect the equilibrium of the system and allowsit a proper structure. Therefore, B. pumilus may act as a PGPR without disturbingthe existing rhizosphere microbial community. However, in soil B, the inoculumstrongly alters the equilibrium in the rhizosphere, and the original structure is notlikely to recover; hence, the structure of the total population of inoculated andnon-inoculated samples differs, following the introduction of the foreign inoculatedbacteria.

In conclusion, the plant growth promoting activity shown by B. pumilus isstrongly conditioned by the physicochemical characteristics and the native rhizo-sphere community of the substrate. Therefore, before this strain can be used as abiostimulant, further studies should be carried out to determine its good per-formance in other soils and whether it can be extended to other plant species.

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Acknowledgements

We wish to thank Brian Crilly and Linda Hamalainen for help in preparing themanuscript.

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