extracellular phytase 2009

8/7/2019 extracellular phytase 2009

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R E S E A R C H A R T I C L E

Extracellular release of a heterologous phytase from roots of

transgenic plants: does manipulation of rhizosphere biochemistry

impact microbial community structure?

Timothy S. George1, Alan E. Richardson2, Sumei S. Li3, Peter J. Gregory1 & Tim J. Daniell1

1SCRI, Invergowrie, Dundee, UK; 2CSIRO Plant Industry, Canberra, ACT, Australia; and 3ISSCAS, Nanjing, China

Correspondence: Timothy S. George, SCRI,

Invergowrie, Dundee, DD2 5DA, UK. Tel.:

144 1382 562 731; fax: 144 1382 562 426;

e-mail: [email protected]

Received 12 February 2009; revised 14 July

2009; accepted 27 July 2009.

Final version published online 10 September

2009.

DOI:10.1111/j.1574-6941.2009.00762.x

Editor: Kornelia Smalla

Keywords

inositol phosphate; mycorrhizae; phytase;

phosphate; rhizosphere; transgenic plants.

Abstract

To maintain the sustainability of agriculture, it is imperative that the reliance of

crops on inorganic phosphorus (P) fertilizers is reduced. One approach is to

improve the ability of crop plants to acquire P from organic sources. Transgenic

plants that produce microbial phytases have been suggested as a possible means to

achieve this goal. However, neither the impact of heterologous expression ofphytase on the ecology of microorganisms in the rhizosphere nor the impact of

rhizosphere microorganisms on the efficacy of phytases in the rhizosphere of

transgenic plants has been tested. In this paper, we demonstrate that the presence

of rhizosphere microorganisms reduced the dependence of plants on extracellular

secretion of phytase from roots when grown in a P-deficient soil. Despite this, the

expression of phytase in transgenic plants had little or no impact on the microbial

community structure as compared with control plant lines, whereas soil treat-

ments, such as the addition of inorganic P, had large effects. The results

demonstrate that soil microorganisms are explicitly involved in the availability of

P to plants and that the microbial community in the rhizosphere appears to be

resistant to the impacts of single-gene changes in plants designed to alter

rhizosphere biochemistry and nutrient cycling.

Introduction

Organic phosphorus (P) tends to accumulate in soils

predominantly as derivatives of inositol phosphates (pri-

marily as phytate or myo-inositol hexakisphosphate) (An-

derson, 1980; Turner et al., 2002). In order to provide plants

access to P present in soil as phytate, transgenic plants

(Arabidopsis thaliana, Nicotiana tabacum L., Trifolium sub-

terraneum L. and Solanum tuberosum L.) that express

phytase genes from soil microorganisms (Aspergillus sp.,

Bacillus sp.) have been developed and characterized (Ri-

chardson et al., 2001; Mudge et al., 2003; Zimmermann

et al., 2003; George et al., 2004, 2005a; Lung et al., 2005).

These plants exude heterologous phytase into the rhizo-

sphere and, when grown in controlled environments (e.g.

sterile and nonsorbing media such as agar), can accumulate

significantly more P than control lines when supplied solely

with phytate (Richardson et al., 2001; Mudge et al., 2003;

George et al., 2004, 2005a). However, the benefit of exuding

phytase from roots has been shown to be compromised in

soil environments, where transgenic T. subterraneum

showed only small (up to 20%) and inconsistent increases

in P accumulation (George et al ., 2004). Transgenic

N. tabacum showed a more consistent improvement in

P accumulation (up to 50%), but only in soils that were

amended with phytate to increase its availability (George

et al., 2005a). Possible reasons for the relatively poor

capacity of transgenic plants to acquire P from phytate in

soil include low availability of substrate for mineralization

by phytase, inhibitory effects of the soil environment on the

activity of phytase exuded to the rhizosphere and, as

investigated in this paper, the presence of microorganisms

(phytase exuding or otherwise), which may compensate for

the lack of phytase exuded by wild-type plants.

Soil microorganisms are involved in many soil functions

(Dunfield & Germida, 2001; Devare et al., 2004) including

the cycling of nutrients, particularly N and P (Oehl et al.,

2001; George et al., 2006; Bunemann et al., 2008). Their

presence in some cases (i.e. mycorrhizal fungi) is critical in

order for plants to survive under extremely P-deficient

FEMS Microbiol Ecol 70 (2009) 433445 c 2009 Federation of European Microbiological Societies

Published by Blackwell Publishing Ltd. All rights reserved

MICROB

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mailto:[email protected]:[email protected]


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conditions (Smith & Read, 1997). More specifically, soil

microorganisms are considered to be critical to the mobili-

zation and cycling of P within the rhizosphere (Jakobsen

et al., 2005). For this and other reasons, the impact of

transgenic plants on the presence of individual species of

microorganisms, or the biodiversity and functionality of the

soil microbial biomass itself, has been of some interest(Bruinsma et al., 2003). However, the results have been

mixed, with some studies showing small effects of transgenic

plants on specific components of the microbial community

(Siciliano et al., 1998; Dunfield & Germida, 2001; Gyamfi

et al., 2002; Bruinsma et al., 2003; Sessitsch et al., 2003;

Castaldini et al., 2005; Henault et al., 2006), while others

have demonstrated no impact at all (Heuer & Smalla, 1999;

Heuer et al., 2002; Brusetti et al., 2004; Devare et al., 2004;

Fang et al., 2005; Griffiths et al., 2006, 2007a, b; Philippot

et al., 2006; Lupwayi et al., 2007). Moreover, a number of

recent studies have suggested that differences between

transgenic plants lie within the range of natural variation

that occurs either between genotypes or due to shifts in

agronomic practice (Griffiths et al., 2006, 2007a, b). Most of

these studies, however, have focused on the impact of plants

that have been genetically modified to produce herbicide

tolerance or resistance to insect pests or plant pathogens

(Kowalchuket al., 2003; Liu et al., 2005; Ikeda et al., 2006;

Griffiths et al., 2007b) and not those that directly alter

rhizosphere biochemistry, such as transgenic plants that

exude phytase.

In this paper, we investigate the impact of rhizosphere

microorganisms on the P nutrition of plants and whether

direct manipulation of rhizosphere biochemistry through

genetic engineering had any impact on microbial commu-nity structure. Transgenic tobacco plants that express and

release extracellular microbial phytases from their roots were

used and compared with both a transgenic control line and

wild-type plants. The efficacy of the expression of phytase in

relation to the P nutrition of plants was examined in relation

to the structure of the bacterial community both within the

rhizosphere and associated with the root (surface and

endophytic bacteria), and on arbuscular mycorrhizal (AM)

fungi associated with the roots.

Materials and methods

Transformation of N. tabacum

Nicotiana tabacum plants (var. W38) were independently

transformed with phytase genes (phyA) from Aspergillus

niger (An) and Peniophora lycii (Pl) using Agrobacterium-

mediated transformation. The fungal phytase genes (phyA)

were expressed independently under the control of either

the CaMV 35S promoter or the A. thaliana phosphate

transporter (AtPt) promoter, with all constructs being

modified for extracellular secretion by inclusion of an

extracellular targeting sequence from the carrot extensin

(ex) gene (Richardson et al., 2001; Mudge et al., 2003;

George et al., 2005a). The P. lycii phytase was synthesized

according to the amino acid sequence reported by Lassen

et al. (2001) with codon usage optimized for expression in

tobacco (Geneart, Regensburg, Germany; A.E. Richardson,unpublished data). Primary transformant calli containing

ex


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of soil wash solution per kg soil or 5% (w/w) live soil,

respectively, with sterilized soil in a cement mixer for

30 min.

Additions of phosphate were based on the amount of

orthophosphate required to increase plant available P by

10-fold (75 mg P kg1 soil) and was incubated for 28 days

before analysis and plant growth. Incubated soils wereanalysed for pH (1 : 5 w/v deionized H2O), anion exchange

resin-extractable P (resin-P or plant available P) (Saggar

et al., 1990), water-extractable Pi and Po (H2O Pi and Po)

and 0.5 M NaHCO3-extractable inorganic P (Olsen-P) (Olsen

& Sommers, 1982; George et al., 2007). The anion exchange

resin method of Saggar et al. (1990) was modified in that

resins were charged with Na2HCO3 and eluted with HCl.

Growth and P uptake by transgenic plants in soil

Five replicate pots containing 450 g (weight at 80% field

capacity) of each soil treatment were sown with five tobacco

seedlings of each plant line separately, which had beengerminated and grown for 7 days on agar containing

100mg kanamycinL1. Wild-type plants were grown to the

same stage on nutrient agar in the absence of kanamycin.

Because of the impact of these differences in the nursery

conditions on the initial vigour of the transformed plants

and the wild type, it was only possible to compare the plant

growth parameters for transgenic plants against the trans-

genic vector control. Pots were thinned to three plants per

pot after establishment and maintained at $80% field

capacity during growth, by weight. All nutrients except P

were supplied weekly by addition of 5 mL of nutrient

solution [3 mM (NH4)2SO4, 2mM KNO3, 1mM MgSO4,

10 mM Ca(NO3)2, 80mM FeEDTA and micronutrients (B,

Cu, Mn, Zn, Mo and Co)]. Plants were grown in a

randomized design in a glasshouse at 561270N and 031040W

between 14 and 22 1C with an approximate daylight length

of 16 h. Before the plants became pot bound, shoots were

harvested after 28 days of growth and biomass was deter-

mined after oven drying at 65 1C. Shoot materials were

milled and analysed for the total P content after digestion

with a sulphuric acidhydrogen peroxide mix (Heffernan,

1985).

DNA extraction from roots and soil

For each of the five replicate pots, rhizosphere soil that was

closely adhering to roots after shaking of all plants in each

pot was brushed from plants and collected into a microcen-

trifuge tube before freezing in liquid N2. A composite

subsample of the roots from each plant in each pot was also

collected into a microcentrifuge tube and also frozen in

liquid N2; this sample included microorganisms associated

with the root surface but also any endophytic microorgan-

isms such as AM fungi. Roots were freeze-dried and

pulverized by bead beating with 1-mm stainless-steel beads

(Atlas ball, UK) using a Mixer Mill 301 (Retsch GmbH,

Haan, Germany). Total DNA was then extracted from roots

(root bacteria and AM fungi) using a Nucleospins 96 Plant

DNA extraction kit (Macherey-Nagel, Germany) following

the manufacturers procedures. Total DNA was extracted

from the soil (rhizosphere bacteria) according to Deng et al.(2009). Briefly, soil was extracted with 1 : 2 w/v 0.12M

Na2HPO4 in a 1% SDS solution on a bead beater with

DEPC-treated glass beads. Following centrifugation (4960 g

for 5 min), the supernatant was added to an equal volume of

phenol : chloroform : IAA (25 : 24 : 1) and mixed. Following

a further centrifugation (4960 g for 1 min), the supernatant

was added to an equal volume of 0.3 M Na acetate in

isopropanol in clean tubes and frozen ( 20 1C) overnight.

This was then centrifuged (4960 g for 5 min) to pellet the

DNA, which was then washed with ice-cold 70% ethanol

and centrifuged (4960 g for 5 min). Ethanol was removed

and the pellet was allowed to dry before being resuspended

in 50 mL TE; this solution was cleaned by passing

over polyvinylpolypyrrolidone (Sigma) using Multiscreen

HTS HV plates (Millipore) after the polyvinylpolypyrroli-

done was equilibrated by repeated water addition (100mL).

DNA extracted from both roots and rhizosphere soil was

used for PCR amplification and community structure

analysis using terminal-restriction fragment length poly-

morphism (T-RFLP). It should be noted that the different

extraction procedures for the rhizosphere and root DNA

could impact the T-RFLP profile observed and comparison

between these two compartments should be made with

caution.

PCR amplification, T-RFLP and sequencing

The small subunit rRNA gene was used to assess the

community structure of AM fungi and root and rhizosphere

bacteria. Following optimization for template quantity, AM

fungal PCR was performed using 2 mL of the DNA extraction

in a 25-mL volume PCR reaction containing Expand High

Fidelity Buffer with 15 mM MgCl2, 100 nM of each of the

dNTPs, 200 nM of each of the primers NS31 and AM1

(Simon et al., 1992; Helgason et al., 1998), 20mg mL1

bovine serum albumin (BSA) and 0.7 U Expand High

Fidelity enzyme mix (Roche Applied Science, Mannheim,

Germany). NS31 was 5 0 labelled with the fluorophore FAM

and AM1 with VIC (Applied BioSystems, UK). Thermo-

cycling conditions were as follows: 94 1C for 2 min; 10 cycles

of 94 1C for 15s, 58 1C for 30s, 72 1C for 45 s; 20 cycles of

94 1C for 15s, 58 1C for 30s, 72 1C for 4515 s per cycle; and

72 1C for 7 min.

Following optimization for template quantity, bacterial

PCR was performed using 2mL of DNA extracted either

from soil or root. This was amplified in a 25-mL volume PCR



435Impact of phytase genes on rhizosphere microorganisms


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reaction containing Platinum Taq buffer with 15mM

MgCl2, 100 nM of each of the dNTPs, 200 nM of each of

the primers 16f27 (AGAGTTTGATCCTGGCTCAG; Amann

et al., 1995) and 1392R (ACGGGCGRTGTGTACA; Black-

wood et al., 2003 modified to include a variable base),

20mgmL1 BSA and 0.725 U of Platinum Taq DNA poly-

merase (Promega UK), 16f27 was 50

labelled with thefluorophore FAM and 1392R with VIC. Before amplifica-

tion, the bacterial PCR mastermix was digested with HhaI

(40min at 37 1C) to remove contaminating bacterial

template, with subsequent heat inactivation (10min at

65 1C) of the endonuclease activity. Thermocycling condi-

tions were as follows: 94 1C for 2 min; 10 cycles of 94 1C for

15 s, 58 1C for 30s, 72 1C for 45 s; 20 cycles of 94 1C for 15 s,

58 1C for 30s, 72 1C for 45s15 s per cycle; and 72 1C for

7min.

All PCR was performed using a DNA Engine PTC Dyad

thermocycler (MJ Research, Reno). The success of PCR

amplification was assessed by agarose gel electrophoresis.

The PCR product was subjected to restriction enzyme

digestion using AluI for bacterial and independent diges-

tions with HinfI and Hsp92I for AM fungal products. In the

restriction enzyme digestion step, 5 mL of PCR product was

digested with 1 mL of restriction enzyme (0.5 U) at 37 1C for

2 h, followed by a 10-min enzyme denaturation step at

65 1C. The digested PCR product was further diluted 1 : 10

with molecular-grade ultrapure water and 1mL mixed with

8.95 mL of formamide and 0.05mL of an internal length

standard (LIZ, Applied Biosystems Inc., Freemont, CA). The

terminal restriction fragments marked with fluorophore

were analysed by electrophoresis using an automated DNA

sequencer (ABI PRISMTM

3730). Blank samples (negativePCR controls from the second-round PCR and water con-

trols) were also digested and analysed. A postrun analysis

was performed using GENEMAPPER (Applied BioSystems, UK)

to allow peak sizing and generation of a peak area for each

identified peak. Peaks that were attributed to plant DNA

(130 bp peak related to chloroplast DNA) were removed

before analysis. A fixed bin width of 5 bp was used as in the

preliminary analysis as this produced uniform and stable

peak identification. Data were then processed in EXCEL

(Microsoft Corporation) to yield peak relative abundance,

with subsequent removal of peaks representing o 1% of the

total fluorescence in each sample. Hellinger transformation

was performed to reduce the effect of dominant peaks

(Blackwood, 2006).

Data presentation and statistical analyses

All data are presented as the mean of five replicates and error

bars represent one SE of the mean. Significant differences

were established using general ANOVA and treatment means

compared by LSD (P= 0.05) (GENSTAT v5; Rothamsted Ex-

periment Station, UK). All data were tested for normality

before analysis and, where required, skewed data were

transformed to natural log values before analysis. The

various microbial community assemblages in the various

compartments of the root soil interface were subjected to

principal component analysis (PCA; GENSTAT v9; Rothamsted

Experiment Station), general ANOVA was used to identify

components that were significantly affected by the experi-mental treatments and relationships between PCs and plant

performance were established using linear regression. The

ShannonWiener index was applied to T-RFLP profiles to

establish diversity that incorporates measures of evenness

and richness.

Results

Soil treatments

The soil treatments affected some soil properties important

to the interpretation of subsequent data: pH varied signifi-

cantly (Po 0.001) with soil treatment, being reduced by0.2U with P addition and increased by 0.3 U with

g-irradiation and 0.4 U when soils were g-irradiated and

reinoculated with either inoculum (Table 1).

Pools of soil P were also changed by the soil treatments

(Table 1). Resin-P was increased (Po 0.001) by both P

addition (25-fold) and g-irradiation (eightfold), although

reinoculation of g-irradiated soil with either inoculum

reduced resin-P to a level not significantly different from

the live soil. Soil treatments also induced similar changes in

Table 1. Characteristics of soil used in experimental treatments

Treatment Live P-fertilized g-Irradiated Reinoculated Bacterial wash LSD (P= 0.05)

pH (H2O) 5.5 (0.0) 5.3 (0.1) 5.8 (0.0) 5.9 (0.0) 5.9 (0.0) 0.2

Resin-Pi (mg g1) 4.3 (0.5) 106.3 (0.9) 33.6 (3.6) 6.7 (0.2) 9.5 (0.8) 5.4

Olsen-Pi (mg g1) 10.8 (2.8) 16.6 (0.1) 20.9 (1.4) 14.9 (1.3) 9.2 (0.5) 4.8

H2O Pi (mg g1) 0.7 (0.3) 46.5 (7.8) 20.4 (2.0) 2.7 (0.2) 10.6 (3.9) 12.7

H2O Po (mg g1) 8.1 (1.2) 43.3 (12.9) 14.5 (4.0) 3.6 (1.2) 7.0 (1.3) 20.6

Soil was a spodosol (010 cm) collected from Tentsmuir Forest, Fife, UK, which was either left live, fertilized with KH 2PO4 (P-fertilized), sterilized by

g-irradiation or sterilized and reinoculated with live soil (5% w/w) (reinoculated) or a bacterial wash. Data represent the mean of five replicates with SEs

in parentheses.

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436 T.S. George et al.


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Olsen-P, with significantly greater P concentrations in

fertilized and sterilized soil, but not in sterilized soils that

were reinoculated. The addition of P to soils led to a large

increase (Po 0.001) in both the water-extractable Pi (inor-

ganic phosphate) and Po (organic phosphate), and

g-irradiation also increased the water-extractable Pi (29-

fold), but not the organic portion of this pool. In reinocu-lated soils, water extractable Pi and Po were again not

significantly different from live soil.

Plant growth and P accumulation

Shoot growth was significantly (Po 0.001) affected by soil

treatment, whereby plants grown in both the P-fertilized and

the g-irradiated soils were larger than those in the other soil

treatments, which were not different from each other (data

not shown). Plants grown in live soil were the smallest, being

2.8-fold smaller than in the P-fertilized and the g-irradiated

treatments. Plants grown in reinoculated soils were inter-

mediate, with the bacterial wash and live soil inoculumtreatments being 1.8-fold and 1.4-fold larger than plants

grown in live soil, respectively. There was also a significant

main effect of plant line (Po 0.001) on shoot biomass, with

plants that expressed the A. niger phytase generally being

larger than either the P. lycii phytase or the control plants.

There was no significant interaction between plant lines and

soil treatments.

Unlike shoot biomass, there was no significant main effect

due to plant line on P accumulation. However, there was a

significant main effect (Po 0.001) caused by the soil treat-

ments so that plants accumulated more P in all treatments

compared with the live soil, and with all treatments being

significantly different from one another (Fig. 1). The effect

of the soil treatments on plant P-accumulation was best

demonstrated by vector control plants, which, when com-

pared with P accumulation on live soil, accumulated 5.3-

fold more P on g-irradiated soils, 4.1-fold more on P-

fertilized soils, 3.2-fold more on bacterial wash and 1.4-fold

more on live soil inoculum treatments. There was also a

significant interaction (Po 0.001) between plant and soil

treatments so that in g-irradiated soil both of the 35S-

promoted phytase constructs (lines 35SAn and 35SPl)

accumulated more P than either the control plants or those

promoted with the AtPt promoter (Fig. 1). In all other soil

treatments, there were no significant differences betweenplant lines.

Impact on rhizosphere and root-associated

microorganisms

The bacterial community structure of the rhizosphere and

the plant root were significantly (Po 0.001) different when

analysed by PCA, with PC1 accounting for 66.4% of the

variability when the two datasets were pooled (data not

shown). However, it should be noted that this difference

may be an artefact of the different DNA extraction protocols

used for rhizosphere and root-associated communities,

although it has been demonstrated, at least for nematode

community structure, that different extraction methods

yield similar community structure analysis using T-RFLP

(Donn et al., 2008). When considered separately, PC1accounted for 35.5% and 25.2% of the variability for the

rhizosphere and root-associated communities, respectively

(Fig. 2).

The rhizosphere populations showed a significantly dif-

ferent and distinct community structure between soil treat-

ments, with the two live soils (live and P-fertilized) being

more similar to each other than the various g-irradiated

treatments in this dimension (Fig. 2a). There were no

significant differences between plant lines or any interaction

between plant line and soil treatment. There were also

significant differences between treatments in PC2, which

accounted for a further 11.0% of the variation. Again, the

soil treatments were all significantly (Po 0.001) different

from one another, with the exception of the bacterial wash

and live soil treatments. The g-irradiated and P-fertilized

treatments were also more similar to one another in this

dimension, as were the live soil and both reinoculation

treatments. Unlike PC1, there were significant (Po 0.05)

differences between plant treatments in PC2, with the

unplanted control being different from all the other planted

treatments. However, there was no significant difference in

the structure of the rhizosphere community between the

transgenic and the control plant lines.

Root-associated bacterial community structure was also

affected by soil treatment, but to a lesser extent than thatobserved in the rhizosphere (Fig. 2b). Soil treatments caused

a significant (Po 0.001) main effect in PC1, which ac-

counted for 25.2% of the variation. In particular, the live

soil treatment was distinct from all other soil treatments and

both the P-fertilized and the g-irradiated treatments were

distinct from the two reinoculated treatments. There was no

significant main effect of the plant line in PC1, but the

interaction between the plant line and the soil treatment was

significant (Po 0.001). Despite this, there were no consis-

tent differences between the transgenic lines and the control,

suggesting that any impact of plants was not consistently

due to the expression of phytase. The next PC to be related

to differences in experimental treatments was PC5, which

explained only 5.5% of the variation. There was a significant

(Po 0.05) impact of soil treatment in this dimension, but

only the live soil and bacterial wash treatments could be

statistically differentiated. There was no significant effect of

plant line in this dimension. As with PC1, statistically

significant (Po 0.05) interactions between treatments

showed no consistent differences between controls and

transgenic plants.





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The diversity of the dominant members of the rhizo-

sphere and root-associated bacterial populations was as-

sessed by the number of T-RFLP fragments that contributed

to the community analysis (Table 2). It is important to note

that this, however, does not reflect the entire diversity in the

system as rare individual species and groups in the popula-

tion are not counted in this instance (Bent & Forney, 2008).

Across the two sample populations, the bacterial richness of

the dominant members of the population was 48% greater

in the rhizosphere compared with the community associated

with the root. Moreover, within the rhizosphere, the greatest

richness, as measured by this method, occurred in the live

soil, which was significantly richer (Po 0.05) than all other

soil treatments including the fertilized (live) soil (Table 2).

However, no differences were evident between plant lines,

nor were there any significant interactions in either of the

rhizosphere or the root-associated populations. In contrast

to the rhizosphere, the richness of the dominant bacterial

Plant treatment

0

1000

2000

3000

0

1000

2000

3000

Plant treatment

35SAn AtPTAn 35SPl AtPTPl Vt

35SAn AtPTAn 35SPl AtPTPl Vt

PAccumulation(gPperplant)

PAccumulation(gPperplant)

0

1000

2000

3000

0

1000

2000

3000

0

1000

2000

3000

LSD P


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communities associated with the root was not influenced by

soil treatment (Table 2).

The AM fungal community structure associated with

roots was also significantly affected (Po 0.001) by soil

treatment as indicated by PC1, which accounted for 57.9%

of the variation (Fig. 2c). In this dimension, the P-fertilized

treatment was distinct from both the live soil and the

g-irradiated soil that received 5% live soil as inoculum.

Neither the g-irradiated soil nor soil that was reinoculated

with a bacterial wash had amplifiable AM fungal DNA

associated with their roots. Despite the large effect due to

soil treatment, there was no impact of plant line or its

interaction with soil treatment. The next PC that was related

to either plant or soil treatment was PC5, which explained

only 1.3% of the variability. In addition to a significant effect

of soil treatment, this component showed a significant

difference (Po 0.05) between plant lines, with both wild-

type and vector control plants being different from all four

transgenic lines.

Given the large effect of soil treatment on microbial

community structure (Fig. 2), the data were reanalysed

within each soil treatment to establish whether therewas a stronger impact of plant line without the over-riding

variation caused by soil treatment. For both the rhizosphere

and the root-associated bacterial communities, however,

there were no instances where transgenic lines collectively

led to a significant and consistent change in the community

structure relative to the two control lines (e.g. Fig. 3

for the rhizosphere communities). However, there were

instances in live soils when PCs, which explained variation

ranging from 5.0% to 27.5% of the data, showed a signifi-

cant difference (Po 0.05) between the controls and the

35SAn line, which is consistent with this construct being

the most responsive in terms of plant growth in sterilized

soil (Fig. 1).

Despite the lack of any strong effect of plant genotype (i.e.

expression of extracellular phytase) on the rhizosphere

microbial community structure, there was an impact of the

presence of plants (Fig. 3). In four of the soil treatments, a

relatively large proportion (7.020.2%) of the variability

within soil treatments could be attributed to PCs that

showed a significant difference (Po 0.05) between

unplanted controls and the planted treatments.

PC1 (35.5%)

0.4 0.2 0.0 0.2 0.4

PC2(11.0%)

0.3

0.2

0.1

0.0

0.1

0.2

0.3

Bacterial washReinoculated-IrradiatedP-fertilizedLive soil35SAnAtPTAn35SPlAtPTPlNPVtWT

PC1 (25.2%)0.90 0.85 0.80 0.75 0.70 0.65 0.60 0.55

PC5(5.5%)

0.2

0.1

0.0

0.1

0.2

PC1 (57.9%)

0.5 0.4 0.3 0.2 0.1 0.0 0.1 0.2

PC5(1.3%)

0.04

0.02

0.00

0.02

0.04

0.06

Rhizosphere bacteria

Root bacteria

AM fungi

(a)

(b)

(c)

Fig. 2. Mean loadings of PCs derived from (a) rhizosphere bacterialcommunity structure, (b) root-associated bacterial community structure

and (c) the root AM fungal community. In each case, the first two PCs to

be significantly affected by the plant and soil treatments are plotted

against each other. Values in parenthesis for each component indicate

the amount of total variability represented. Filled stars represent the

mean loading for each soil treatment (n = 35); live (cyan); P-fertilized

(pink); sterilized by g-irradiation (red); and sterilized soil reinoculated with

a bacterial wash (green) or with 5% of live soil (blue). The mean effects

of Nicotiana tabacum lines (n = 5) that express phyA from either

Aspergillus niger (An) or Peniophora lycii (Pl) under the control of either

the 35S constitutive (35SAn and 35SPl, respectively) or the Arabidopsis

thaliana AtPT1 phosphate transporter (AtPTAn and AtPTPl, respectively)

promoters and vector (Vt) and wild-type (WT) control lines along with a

no plant control (NP) arealso shown by separate symbols. On each panel,the bars show the LSD (P= 0.05) for the interaction between plant and

soil treatments in both dimensions.

Table 2. Effect of soil treatment on bacterial richness, as measured by

T-RFLP in rhizosphere and root-associated populations

Treatment

Number of T-RFLP peaks scored

Rhizosphere Root associated

Live 36.4 (4.1) 21.1 (3.9)

g-Irradiated 32.0 (2.7) 22.0 (2.6)

Bacterial wash 30.1 (2.2) 21.3 (2.7)

Reinoculated 32.2 (2.6) 22.5 (1.8)

P-fertilized 30.1 (3.6) 21.9 (1.8)

LSD (P40.05) 1.7 n.s.

Data are presented as the mean number of T-RFLP peaks scored (based

on a 1% threshold) foreachsoil treatmentpooled across all sixplantlines

(n = 5 for each plant line). Means are shown with one SD in parentheses

and, where significant, the LSD is provided (P= 0.05).





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Relationship between rhizosphere community

structure and plant P accumulation

There was a significant relationship (R2= 0.75, Po 0.001)

between rhizosphere community structure (as determined

for PC1; Fig. 1) and shoot P accumulation by the plants (Fig. 4).

However, a major shift in microbial community structure

was not required in order to achieve an increase in shoot P

accumulation, as demonstrated by the P-fertilized treat-

ment. Likewise, it was also evident that a large change in

the structure of the microbial community did not necessa-

rily change the ability of the plant to accumulate P asdemonstrated by the comparison between live soil and

the various inoculum treatments of sterilized soil. While

g-irradiation caused the largest change in both the commu-

nity structure and increase in the availability of P, the soil

inoculated with a bacterial wash appeared to transform the

microbial community back towards that of the live soil, and

reduced the availability of P as measured in both soil extracts

(Table 1) and by uptake of P by plants (Fig. 4). Moreover,

the sterilized soil that was reinoculated with 5% live soil

shifted the community structure even further toward that of

the live soil and further reduced the availability of soil P for

plant uptake.

Discussion

This study demonstrates that the heterologous expression of

a microbial phytase gene in plants had no detectable impact

on the community structure of microorganisms in the

rhizosphere or those associated with the root, given the

known limits of the methodology applied. This was despite

these plant lines previously being shown to have a clear

effect on the biochemistry of the rhizosphere (George et al.,

2005b, 2007). In contrast, soil treatments aimed at perturb-

ing the soil microbiology or altering the P status of the soil

had a significant effect on microbial community structure.The effects of soil treatment were particularly evident on

mycorrhizal associations with roots and on the structure of

the bacterial community in rhizosphere soil as compared

with that associated with roots. Furthermore, and despite

the lack of impact of the expression of phytase in different

transgenic lines on soil microorganisms, it was apparent that

PC1(20.2%)

0.2

0.1

0.0

0.1

0.2

Plant treatment

35SAn

AtPTAn

35SPl

AtPTPl

NP V

t

WT

PC4

(7.0%)

0.10

0.05

0.00

0.05

0.10

PC1(16.7%)

0.1

0.0

0.1

Live

Bacterial wash

P-fertilized

PC3(10.9%

)

0.05

0.00

0.05

0.10

0.15

-Irradiated

LSD (P


9/13

the presence of microorganisms themselves had an influence

on the efficacy of the expression of phytase in plants on their

growth, and subsequently, the ability to acquire additional

phosphate from soil.

Impact of rhizosphere microorganisms on the

efficacy of heterologous expression of phytase

Plants that constitutively expressed either the A. nigeror the

P. lycii phytase accumulated more P than control plants

when grown in g-irradiated soils, but not in any of the live

soil treatments (either unamended or P-fertilized) or in

either of the reinoculation treatments. While not knowing

exactly the effectiveness of the g-irradiation treatment in

sterilizing the soil, the dose was considered to be sufficient

to eliminate most soil microorganisms (McNamara et al.,

2003), and was effective for the removal of mycorrhizal

fungi. No attempt was made, however, to maintain sterility

in the soil, which was allowed to recolonize naturally over 28

days of incubation and a subsequent period of 28 days of

plant growth. While it is tempting to suggest that the

presence of mycorrhizal fungi might be responsible for

compensating plants for the inherent lack of phytase expres-

sion in all treatments except for the sterilized soils, this was

not supported by the results (i.e. mycorrhizal fungi were also

absent from the bacterial-wash treatment and no differences

were observed between the various plant lines). This sug-

gests that the presence of some other component of the

microbial biomass, or the general presence of microorgan-

isms themselves in the rhizosphere, renders the expression

of phytase in plants ineffective relative to control lines. This

could be for a number of reasons, which include (1)

microbial decomposition or inactivation of the phytase

when exuded into the rhizosphere, although we have shown

previously that the enzyme can remain active in live soils(George et al., 2007), (2) that the microbial biomass

indirectly mediates the availability of inositol phosphates

for mineralization by phytase (LAnnunziata, 1975;

Bunemann et al., 2008) and (3) that the presence of

phosphatase and phytase-exuding microorganisms in the

rhizosphere themselves compensates directly for the inabil-

ity of plants to utilize P from inositol phosphates. Indeed, a

wide range of soil fungi and bacteria that exhibit phytase

activity have been reported (Tarafdar & Claassen, 1988; Hill

& Richardson, 2007; Sakurai et al., 2008) and, in some

instances, have been shown to be more prominent in the

rhizosphere (Unno et al., 2005). Other studies have sug-

gested that shifts in rhizosphere microbial community

structure are correlated with a more effective mineralizing

environment (Acosta-Martinez et al., 2003; Marschner et al.,

2006; Renalla et al., 2006). However, this was not evident in

the present study and it is important to note that under

certain conditions of enhanced availability of inositol phos-

phates, plants exuding phytase do have an advantage even in

live soils (George et al., 2005a).

The g-irradiation treatment also had a major effect on the

growth of all plant lines, with both transgenic and control

lines producing larger biomass and accumulating more P

than when grown in the unamended soil. This response may

be due to either a relief of pathogenesis or due to theincreased availability of P. For the former, it was evident

that reduced growth and P accumulation occurred in both

the reinoculation treatments and less growth and P accu-

mulation was evident in the P-fertilized soil, despite the P-

rate being considered to be sufficient for maximum plant

growth. An alternative explanation is that the irradiation

treatment resulted in a large release of P (Table 1), which

occurred presumably with the removal of any microorgan-

isms that may exert a deleterious effect on plant growth. The

lack of growth response of the transgenic lines that had

targeted expression of the phytase (i.e. from the AtPT1

promoter) may have also been affected by the availability of

P in these treatments, as it would be expected that the

expression of the phytase from this promoter would be

suppressed under such conditions (Mudge et al., 2003). The

impact of reinoculation on reduced plant growth might also

be attributed to a rapid immobilization of available P back

into microbial biomass and thus increased competition with

plants for access to this P (Jakobsen et al., 2005). The release

of cell contents other than inorganic P with g-irradiation,

i.e. monoester forms of P such as inositol phosphates

Shoot P accumulation (g P per plant)

0 1000 2000 3000 4000 5000

PC1

(35.5%)

0.6

0.4

0.2

0.0

0.2

0.4

0.6

Bacterial washReinoculated

-IrradiatedLive soilP-fertilized

Fig. 4. Relationships between shoot P accumulation (mg P per plant) of

plants grown in different soil treatments and the PC1 derived from the

rhizosphere bacterial community structure. Two separate relationships

fitted by an exponential function (y= a1b (1 exp(cx)) are shown along

with R2 values and the significance (P-value) of the relationships as

established by regression analysis. Data points represent individual

observations for all plant lines within soil treatments and include data

from live (unamended) (cyan), P-fertilized (pink) and g-irradiated (red)

soils and sterilized soil that was reinoculated with either a bacterial wash

(green) or 5% live soil (blue).





10/13

(LAnnunziata, 1975; Bunemann et al., 2008) and easily

mineralized compounds (Oehl et al., 2001), is entirely

possible, and might also go some way towards explaining

the response of the phytase-exuding transgenic plants over

the controls.

Impact of the heterologous expression of

phytase in plants on the microbial community

structure in the rhizosphere and that associated

with roots

Despite the apparent interaction of rhizosphere microor-

ganisms and ability of the heterologous phytase to improve

the P nutrition of plants, there was little impact of the

different plant lines on the structure of the bacterial or

AM fungal communities. However, the presence of the

plants did have a major influence on selecting a community

structure that was different between the rhizosphere and

that associated with roots, with the latter showing less

richness of dominant members of the population and being

affected to a lesser extent by soil treatment. Other studies

have similarly shown differing community structure of

bacteria with increasing proximity to plant roots as opposed

to that in the rhizosphere (Brusetti et al., 2004; Crecchio

et al., 2007).

The different plant lines used in the study, though, did

not explain any significant variability in the structure of the

rhizosphere bacterial community, even when soil treatment

was removed as a variable, with the only consistent effect

being the presence or absence of a plant (Fig.3). The

influence of the presence of a plant compared with anunplanted control has commonly been observed and is

considered to be a major driver of microbial community

structure in soil and within the rhizosphere (Dunfield &

Germida, 2003; Philippot et al., 2006).

In contrast, soil treatment had large effects on the rhizo-

sphere microbial community structure, with a major com-

ponent of this variability being explained by the effect of

both g-irradiation and P availability. Irradiation of the soil

significantly reduced the richness of the bacterial population

as measured by T-RFLP and, although the change in micro-

bial biomass and the structural basis for this population shift

was not investigated, it was evident that reinoculation of the

sterilized soil shifted the bacterial community structure back

towards that of the unamended soil. This observation was

apparent in both the bacterial wash and addition of 5% fresh

soil treatments, with the latter having a more pronounced

effect (Fig. 4). While this may be a direct consequence of the

reintroduction of specific groups of microorganisms and

their interaction with plant roots, it may also be a result of

interactions of the microbial population with other aspects

of the soil environment.

Indeed, there was a relationship between changes in the

community structure of the rhizosphere bacteria and the

availability of P to plants as measured by their P accumula-

tion (Fig. 4). However, whether the community structure

was reacting to the availability of P, or whether the avail-

ability of P was a consequence of the soil treatments only,

remains uncertain. The fact that P availability could beincreased without a significant change in the community

structure, as in the P-fertilized treatment, and with large

differences in community structure occurring between cer-

tain treatments without a major change in the availability of

P, suggests that this relationship is not straightforward.

What is of interest, though, is that despite g-irradiation

releasing less inorganic P than that added in the P-fertilized

treatment (as measured by standard P-extractions; Table 1),

the P released from the biomass appeared to be more

available for plant uptake, which hints at the importance of

the microbial biomass as a source of available P to plants

(Seeling & Zasoski, 1993; Bunemann et al., 2008) and their

important role in the soil P-cycle.

Soil treatment also had a significant effect on the bacterial

community structure associated with roots, although this

was less marked than that observed in the rhizosphere. This

suggests that the plant exerts a strong influence on the

composition of the root-associated bacterial community

structure, which, to a large extent, had an over-riding

influence over the effects due to soil treatment affecting the

more distant rhizosphere. As with the rhizosphere bacterial

community structure, there were no significant main effects

due to the expression of the phytase gene in transgenic

plants. This observation is similar to that of Rasche et al.

(2006), who demonstrated that any impacts of a transgenictrait on bacterial endophytes were small in comparison with

other variables including plant genotype and soil type.

However, other studies have suggested a significant impact

of heterologous traits on the community structure of both

rhizosphere and root-associated bacteria (Siciliano et al.,

1998; Siciliano & Germida, 1999), despite these transgenic

traits not being targeted to the rhizosphere.

AM fungal community structure was also affected by the

soil treatment where g-irradiation was effective in removing

AM fungi from the soil. Consequently, no amplifiable AM

fungal DNA was present in the roots of plants grown in

either the irradiated soil or the irradiated soil that was

reinoculated with the bacterial wash. Of the other treat-

ments where AM was identified, the AM fungal community

structure in P-fertilized soil was significantly different from

the nonfertilized treatments. This suggests that P availability

not only affects the strength of mycorrhizal symbiosis but

also the community structure. Such a possibility has simi-

larly been demonstrated by others (Frey et al., 2004;

Vogelsang et al., 2006). In addition, plant line changed the

AM fungal community structure such that the control lines





11/13

had a community structure different from the transgenic

lines. However, this difference only explained 1.3% of the

variability in the data and further resolution showed that

this was only seen in the live soil inoculum treatment. Other

studies have shown a significant impact of a transgenic trait

(e.g. expression of Bt genes in maize) on the AM fungal

infection of roots (Castaldini et al., 2005; OCallaghan et al.,2005). However, our data suggest that any impact of the

heterologous expression of phytase on the AM fungal

community structure was only evident following disruption

of the microbial community structure by sterilization of the

soil.

Overall, there was little evidence for any impact of the

transgenic plant lines on the structure of microbial commu-

nities either within the rhizosphere or on the root surface.

While it is not possible to suggest whether this would be the

case in other soils with different community structures, the

weak sorption environment of this spodosol is likely to allow

the greatest impact of the heterologous protein, which has

been shown to be strongly adsorbed in other stronger

sorbing soils (George et al., 2005b, 2007). Other studies

have similarly reported little impact of transgenic plants on

rhizosphere microbial community structure (Heuer & Smal-

la, 1999; Heuer et al., 2002; Brusetti et al., 2004; Devare et al.,

2004; Fang et al., 2005; Griffiths et al., 2006, 2007a, b).

Importantly, there was no major impact on functionally

important microbial groups here, for example AM fungi,

which have been designated as indicators of environmental

perturbation (Bruinsma et al., 2003; Kowalchuket al., 2003).

This is the first study of the impacts on rhizosphere and

root-associated microbial community structure of the ex-

pression and extracellular release of a heterologous trans-gene designed to specifically alter the biochemistry and

cycling of nutrients in the rhizosphere. Our data suggest

that while soil microorganisms appear to be involved in the

availability of P to plants, the microbial community in the

rhizosphere appears to be resistant to the impact of single-

gene changes in plants designed to alter root biochemistry

and nutrient cycling in the rhizosphere. In this case, a

transgenic technology aimed at improving the sustainability

of agriculture by altering rhizosphere biochemistry appears

to have little impact with regard to the ecology of the

microbial community and thus the wider ecology of the

agricultural system.

Acknowledgements

This research was supported by the European Commission

under a Marie Curie Outgoing International Fellowship

(T.S.G.) and the contents of the paper reflect the opinion of

the authors and not that of the European Commission. The

research in the paper was also contributed to by the Royal

Society (London) through funding for a ChinaUK Science

Network between SCRI and ISSCAS (S.S.L.) and by a travel

grant from the Australian Academy of Science, International

Science Linkages Program (A.E.R.). SCRI is supported by a

grant-in-aid from the Scottish Government RERAD. The

authors thank L. Brown for technical assistance.

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