Enrichmentandgenotypicdiversityof phlD-containing£uorescentPseudomonasspp. in twosoils aftera centuryofwheatand£axmonocultureBlanca B. Landa1,2, Olga V. Mavrodi1, Kurtis L. Schroeder3, Raul Allende-Molar1 & David M. Weller3

1Department of Plant Pathology, Washington State University, Pullman, WA, USA; 2Departamento de Agronomıa, Escuela Tecnica Superior de

Ingenieros Agronomos y de Montes, Universidad de Cordoba, Cordoba, Spain; and 3US Department of Agriculture, Agricultural Research Service,

Root Disease and Biological Control Research Unit, Washington State University, Pullman, WA, USA

Correspondence: David M. Weller, US

Department of Agriculture, Agricultural

Research Service, Root Disease and Biological

Control Research Unit, PO Box 646430,

Washington State University, Pullman, WA

99164-6430, USA. Tel.: 11 509 335 6210;

fax: 11 509 335 7674;

e-mail: wellerd@mail.wsu.edu

Received 25 April 2005; revised 6 September

2005; accepted 8 September 2005.

First published online 19 December 2005.

doi:10.1111/j.1574-6941.2005.00038.x

Editor: Gary King

Keywords

antibiotic-producing bacteria; host preference;

2,4-diacetylphloroglucinol (DAPG);

Pseudomonas fluorescens.

Abstract

Fluorescent Pseudomonas spp. producing the antibiotic 2,4-diacetylphloroglucinol

(2,4-DAPG) play a key role in the suppressiveness of some soils to take-all of wheat

and other diseases caused by soilborne pathogens. Soils from side-by-side fields on

the campus of North Dakota State University, Fargo, USA, which have undergone

continuous wheat, continuous flax or crop rotation for over 100 years, were

assayed for the presence of 2,4-DAPG producers. Flax and wheat monoculture, but

not crop rotation, enriched for 2,4-DAPG producers, and population sizes of log

5.0 CFU g root� 1 or higher were detected in the rhizospheres of wheat and flax

grown in the two monoculture soils. The composition of the genotypes enriched

by the two crops differed. Four BOX-PCR genotypes (D, F, G, and J) and a new

genotype (T) were detected among the 2,4-DAPG producers in the continuous flax

soil, with F- and J-genotype isolates dominating (41 and 39% of the total,

respectively). In contrast, two genotypes (D and I) were detected in the soil with

continuous wheat, with D-genotype isolates comprising 77% of the total. In the

crop-rotation soil, populations of 2,4-DAPG producers generally were below the

detection limit, and only one genotype (J) was detected. Under growth-chamber

and field conditions, D and I genotypes (enriched by wheat monoculture)

colonized the wheat rhizosphere significantly better than isolates of other

genotypes, while a J-genotype isolate colonized wheat and flax rhizospheres to

the same extent. This study suggests that, over many years of monoculture, the

crop species grown in a field enriches for genotypes of 2,4-DAPG producers from

the reservoir of genotypes naturally present in the soil that are especially adapted to

colonizing the rhizosphere of the crop grown.

Introduction

Genes encoding resistance to foliar plant pathogens are

abundant in crop species, but resistance to some of the most

widespread soilborne pathogens (e.g. Gaeumannomyces gra-

minis, Pythium spp., Rhizoctonia spp. and many Fusarium

spp.) (Weller et al., 2005) that cause root rots, crown rots,

damping-off and wilts is often lacking. As an alternative,

crops have evolved a strategy of stimulating and supporting

specific groups of indigenous antagonistic microorganisms

as the first line of defence against soilborne pathogens (Cook

et al., 1995; Weller et al., 2005). Isolates of fluorescent

Pseudomonas spp. producing the antibiotic 2,4-diacetyl-

phloroglucinol (2,4-DAPG) are responsible for some of the

best examples of natural microbial defence of plant roots

(Weller et al., 2002). For example, in both Washington State

and Dutch fields, 2,4-DAPG producers play a key role in

take-all decline (TAD), a natural suppression of take-all

developing when a field is continuously cropped to wheat or

barley following a severe outbreak of the disease (Raaij-

makers & Weller, 1998; Weller et al., 2002; de Souza et al.,

2003). The population of 2,4-DAPG producers must reach a

threshold density of 105 CFU g root� 1 to be effective against

take-all (Raaijmakers & Weller, 1998; Weller et al., 2002).

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

2,4-DAPG also contributes to the suppressiveness of certain

soils in Switzerland to black root rot of tobacco caused by

Thielaviopsis basicola (Stutz et al., 1986; Weller et al., 2002;

Ramette et al., 2003). In addition, evidence is mounting that

monoculture of other crops results in an enrichment of 2,4-

DAPG producers (Weller et al., 2002). Landa et al. (2002a)

reported that 2,4-DAPG producers were abundant

(4105 CFU g root� 1) on pea grown in a soil in Mount

Vernon, Washington that had undergone over 30 years of

continuous pea monoculture and is suppressive to Fusarium

wilt caused by Fusarium oxysporum f. sp. pisi. The key role

that 2,4-DAPG plays in disease suppression has been repeat-

edly demonstrated both by studies using genetic mutational

analysis with Pseudomonas fluorescens strains CHA0, F113,

Q8r1-96, Q2-87 and SSB17 (Vincent et al., 1991; Keel et al.,

1992; Shanahan et al., 1992; Harrison et al., 1993; de Souza

et al., 2003) and by direct isolation of the antibiotic from

rhizospheres colonized by these bacteria (Keel et al., 1992;

Bonsall et al., 1997; Raaijmakers et al., 1999).

The 2,4-DAPG biosynthetic locus includes the five-gene

operon phlACBDE (Bangera & Thomashow, 1999). Particu-

larly noteworthy is phlD, which functions in the synthesis of

the 2,4-DAPG precursor monoacetylphloroglucinol

(MAPG). The phl biosynthetic genes are conserved among

all known 2,4-DAPG-producing fluorescent Pseudomonas

spp. (Keel et al., 1996; McSpadden-Gardener et al., 2000; Lee

& Kim, 2001; Mavrodi et al., 2001; de la Fuente et al.,

2004b). Thus, the terms phlD1 fluorescent Pseudomonas

spp. and 2,4-DAPG producer are used synonymously be-

cause, among the strains tested, detection of the gene

correlates with the capacity to produce 2,4-DAPG.

Fluorescent Pseudomonas spp. strains producing 2,4-

DAPG exhibit considerable genetic and phenotypic diver-

sity. Keel et al. (1996) and McSpadden-Gardener et al.

(2000) reported three distinct phylogenetic groups using

amplified ribosomal DNA restriction analysis (ARDRA),

designated ARDRA Groups 1, 2 and 3, or A, B and C,

respectively. The well-described ARDRA Group 1 strains P.

fluorescens CHA0 and Pf-5 are known for producing a much

broader range of biologically active metabolites than Group

2 and 3 strains. Strain F113 from Ireland (Fenton et al.,

1992) is one of the few known representatives of ARDRA

Group 3 (Keel et al., 1996; McSpadden-Gardener et al.,

2000). Genotype analysis of phlD1 fluorescent Pseudomonas

spp. based on genomic fingerprinting by random amplified

polymorphic DNA (RAPD) analysis (Keel et al., 1996; Lee &

Kim, 2001; Raaijmakers & Weller, 2001; Mavrodi et al., 2001;

Bergsma-Vlami et al., 2005b), whole-cell repetitive se-

quence-based (rep)-PCR analysis (McSpadden-Gardener

et al., 2000; Landa et al., 2002a), restriction fragment length

polymorphism (RFLP) analysis of phlD (RFLP-phlD) (Mav-

rodi et al., 2001; McSpadden-Gardener et al., 2001; Ramette

et al., 2001; Wang et al., 2001; Landa et al., 2002a; Mazzola

et al., 2004), phylogenetic analysis of phlD (Mazzola et al.,

2004; Bergsma-Vlami et al., 2005b) and denaturing gradient

gel electrophoresis (DGGE) analysis of phlD (Bergsma-

Vlami et al., 2005b) revealed a considerable amount of

genetic variation among phlD1 Pseudomonas spp. (Keel

et al., 1996; McSpadden-Gardener et al., 2000; Picard et al.,

2000; Weller et al., 2002; Picard & Bosco, 2003). Seventeen

different phlD1 genotypes (designated A to Q) were recog-

nized by rep-PCR, with the BOXA1R primer (BOX-PCR) or

RFLP-phlD analysis (McSpadden-Gardener et al., 2000,

2001; Mavrodi et al., 2001; Landa et al., 2002a), and the

clusters defined by the different genomic fingerprinting

methods or by phylogenetic analysis of phlD correlate well

(de la Fuente et al., 2005). New genotypes have been

described, however, some of them using different ap-

proaches (Mazzola et al., 2004; Bergsma-Vlami et al.,

2005b; McSpadden-Gardener et al., 2005).

We demonstrated that the genetic profiles generated by

BOX-PCR and RFLP-phlD analysis are predictive of the

rhizosphere competence of 2,4-DAPG-producing fluores-

cent Pseudomonas spp. in the wheat and pea rhizospheres,

and now think that isolates of some genotypes preferentially

colonize the roots of certain crop species (Raaijmakers &

Weller, 2001; Weller et al., 2002; Landa et al., 2002a, 2003).

This idea is supported by findings that D-genotype isolates

are dominant in the rhizosphere of wheat grown in Wa-

shington State TAD soils, even though three other genotypes

(B, L and E) also are found in these soils (McSpadden-

Gardener et al., 2000; Raaijmakers & Weller, 2001). Simi-

larly, in the rhizosphere of pea grown in Mt. Vernon pea

monoculture soil, D- and P-genotype isolates are dominant,

even though A, L, O and Q isolates also are found in the soil

(Landa et al., 2002a). In addition, introduced isolates of

genotype D, and genotypes D and P colonized the rhizo-

spheres of wheat (Raaijmakers & Weller, 2001; Landa et al.,

2003) and pea (Landa et al., 2002a), respectively, signifi-

cantly better than the other genotypes that were not

dominant in these monoculture soils.

In this study, the hypotheses that crop monoculture

enriches for phlD1 isolates and that the crop species grown

modulates the genotypes enriched from those present in the

soil were further tested. The study was conducted using soil

from a unique site on the campus of North Dakota State

University, Fargo, North Dakota where side-by-side fields

have undergone continuous wheat, continuous flax or crop

rotation for over 100 years. We show that flax monoculture,

like wheat monoculture, enriches for 2,4-DAPG producers

in the rhizosphere, but the composition of the genotypes

enriched by the two crops is quite different. BOX-PCR-

defined D-genotype isolates were dominant in the wheat

monoculture soil, but flax monoculture enriched for F- and

J-genotype isolates. In addition, a new T genotype was

described from the Fargo flax monoculture soil.

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

352 B.B. Landa et al.

Materials andmethods

Soils andplants

Three soils (fine, smectitic, frigid Typic Epiaquerts)

were collected from field plots located on the campus of

North Dakota State University (NDSU), Fargo, ND, in 1997.

One soil, designated Fargo continuous wheat (FCW),

was from a plot (NDSU wheat plot no. 2) established by

W. M. Hays in 1882 to study soilborne diseases of wheat,

and every year since then had been cropped to several

different varieties of hard red spring wheat, making it the

oldest continuous spring wheat plot in the world. A second

soil, designated Fargo continuous flax (FCF), was from a

plot (NDSU flax plot no. 30) established by H. L. Bolley in

1894 to evaluate Fusarium wilt of flax (FWF) caused by

Fusarium oxysporum f. sp. Lini, and by 1997 the field had

been cropped for 103 continuous years to several different

varieties of flax. The third plot (NDSU plot R5) designated

Fargo crop rotation (FCR) had been rotated to a variety

of crops such as bean, corn, oat, soybean, sugar beet

and sunflower, or left fallow, and in 1997, when the soil

was collected, was cropped to sugar beet. The three fields

are located side by side and share similar physical and

chemical properties (data not shown). The wheat and flax

plots gained entry into the National Register of Historic

Places in 1992.

The soils were collected from the upper 30 cm of the soil

profile, shipped to Pullman, WA in plastic buckets, air-dried

for 1 week, passed through a 0.5-cm-mesh screen, and a

portion of the soils were planted to wheat within 2 months

from the date they were collected (experiment 1). The

remainder of the soils was stored in a covered shed exposed

to outdoor temperatures ranging from a low of about

� 7 1C to a high of about 38 1C. After three (experiment 2)

or six (experiment 3) years of storage, portions of the

original soils were again assayed.

A Shano sandy loam soil (Weller et al., 1997) (Quincy

virgin) from a noncropped site near Quincy, WA that

supports native vegetation was used in studies of the root-

colonizing ability of 2,4-DAPG producers as previously

described (Landa et al., 2002a, 2003). The soil was collected

from the upper 30 cm of the soil profile, air-dried for 1 week,

and passed through a 0.5-cm-mesh screen. The soil was

stored inside a headhouse at 24 1C.

A field plot (Thatuna silt loam) located on the Wash-

ington State University Plant Pathology Research Farm,

Pullman, WA, was used for studying the rhizosphere

competence of selected phlD1 genotypes under field con-

ditions.

Flax (Linum usitatissimum L.) cv. Norlin and the spring

wheat (Triticum aestivum L.) cv. Penawawa were used

throughout the study.

Bacterial strains,growthmediaand inoculumproduction

Twenty-nine reference strains of 2,4-DAPG-producing

(phlD1) Pseudomonas fluorescens representing 17 BOX-

PCR genotypes were used in this study (Table 1) (McSpad-

den-Gardener et al., 2000; Mavrodi et al., 2001; Landa et al.,

2002a). P. fluorescens strain 2-79, which does not contain

phlD, was used as a negative control for PCR (Landa et al.,

2002a). Chemicals were obtained from Sigma Chemical Co.,

St Louis, MO, unless noted otherwise. All strains were

routinely cultured on a modified semiselective medium for

fluorescent Pseudomonas spp. consisting of one-third-

strength King’s B medium (1/3�KMB) supplemented with

antibiotics (1/3�KMB111) (McSpadden-Gardener et al.,

2000) at 27 1C. Indigenous phlD1 fluorescent pseudomo-

nads and total pseudomonads were isolated from the rhizo-

sphere on 1/3�KMB111 agar or 1/3�KMB111 broth as

described below. Introduced Pseudomonas strains used in

colonization studies were spontaneous rifampicin-resistant

mutants and were recovered and quantified in 1/3�KMB111 broth supplemented with rifampicin (100

mg mL� 1) (1/3�KMB111rif). Rifampicin-resistant and

their respective wild-type strains were similar phenotypi-

cally. Frozen stock cultures of all strains were stored in

1/3�KMB111 plus 35% glycerol at � 80 1C.

Determinationofpopulationdensitiesoftotalpseudomonadsand indigenous2,4-DAPGproducers in theFargosoils

Three cycling experiments were conducted with wheat and/

or flax as described previously (Raaijmakers & Weller, 1998;

Landa et al., 2002b). The purpose of the cycling was to

quantify the population sizes of phlD1 isolates that devel-

oped in the rhizosphere and to collect isolates for studies of

genetic diversity within the populations of these soils. Wheat

(cv. Penawawa) was selected in the three experiments to

assay the soils because it is supportive of phlD1 isolates

belonging to the known BOX-PCR genotypes (McSpadden-

Gardener et al., 2000). In cycling experiment 1 initiated in

1997, wheat was cycled in the three soils beginning about 2

months after the soils were collected. In cycling experiment

2 conducted in 2000, either wheat or flax (cv. Norlin) was

grown in the FCW and FCF soils after 3 years of storage. In

cycling experiment 3 conducted in 2003, wheat was grown

in all three Fargo soils after the soil had been stored for

6 years.

Twelve wheat seeds (experiment 1), 10 wheat or flax seeds

(experiment 2) or 20 wheat seeds (experiment 3) were sown

in PVC pots (8 cm high, 7.5 cm wide) containing approxi-

mately 200 g of sieved Fargo soils (experiments 1 and 2) or

in PVC pots (17.5 cm high, 15.5 cm wide) containing

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

353Pseudomonas spp. in soils after wheat and flax monoculture

Table 1. Reference strains and isolates of 2,4-diacetylphloroglucinol-producing Pseudomonas spp. from the rhizospheres of flax and wheat grown in

soils collected from three field plots at the North Dakota State University, Fargo, ND

Prefix or strain�Isolate

number GenotypewPlant

source

Soil

samplez Ref.‰

Reference strains

Pf-5, CHA0 A Cotton,

tobacco

Howell & Stipanovic (1979);

Keel et al. (1996)

Q2-87, Q2-1 B Wheat Bangera & Thomashow (1999);

McSpadden-Gardener et al. (2000)

STAD384-97 C Wheat Mavrodi et al. (2001)

OC4-1, Q8r1-96,

QT1-5, W2-6

D Wheat McSpadden-Gardener et al. (2000);

Raaijmakers & Weller (2001)

Q2-2, QT1-6 E Wheat McSpadden-Gardener et al. (2000)

JMP6 F Wheat McSpadden-Gardener et al. (2000)

FFL1R18 G Wheat McSpadden-Gardener et al. (2000)

CV1-1 H Wheat McSpadden-Gardener et al. (2000)

FTAD1R36 I Wheat McSpadden-Gardener et al. (2000)

FFL1R22, CC3-1 J Wheat McSpadden-Gardener et al. (2000)

F113 K Sugarbeet Fenton et al. (1992)

W4-4, 1M1-96 L Wheat McSpadden-Gardener et al. (2000);

Raaijmakers & Weller (2001)

PILH1, D27B1 M Tomato, wheat Keel et al. (1996); McSpadden-

Gardener et al. (2000)

HT5-1 N Wheat McSpadden-Gardener et al. (2000)

7MA12, 7MA20 O Pea Landa et al. (2002a)

7MA15, MVP1-4 P Pea Landa et al. (2002a)

MVW4-2, MVW4-3 Q Wheat Landa et al. (2002a)

Experiment 1

FFL1-R 9 D Wheat FCF soil McSpadden-Gardener et al. (2000);

Mavrodi et al. (2001), ts

16, 23, 24, 25, 26 G Wheat FCF soil McSpadden-Gardener et al. (2000);

Mavrodi et al. (2001), ts

8, 10, 11, 13, 14, 15, 17 J Wheat FCF soil McSpadden-Gardener et al. (2000);

Mavrodi et al. (2001), ts

FTAD1R 5, 25, 26, 27, 33, 34, 35, 37, 38 D Wheat FCW soil McSpadden-Gardener et al. (2000);

Mavrodi et al. (2001), ts

FFL2R 65 D Wheat FCF soil ts

3, 7, 11, 13, 17, 18, 19, 20, 32

34, 40, 44, 49, 50, 51, 52, 53, 57

F Wheat FCF soil ts

60 ts

9, 33, 62, 71 G Wheat FCF soil ts

6, 8, 10, 12, 14, 15, 21, 22, 24

26, 27, 28, 29, 35, 36, 38, 42, 43

J Wheat FCF soil ts

45, 46, 56, 63, 66, 68, 69, 70, 72

4, 5, 54, 55, 61 T Wheat FCF soil ts

FTAD2R 3, 11,16, 18, 19, 23, 43, 47, 48,

49, 52, 53, 55, 56, 57

D Wheat FCW soil ts

21, 22 I Wheat FCW soil ts

FFL3R 7, 29, 30, 47, 54, 59, 60 F Wheat FCF soil ts

39, 48, 55 G Wheat FCF soil ts

41, 58, 61, 63, 72 J Wheat FCF soil ts

49 T Wheat FCF soil ts

FTAD3R 49, 51, 52 I Wheat FCW soil ts

FSB3R 24, 28, 29, 32 J Wheat FCR soil ts

FFL4R 18, 19, 21, 23, 24, 25, 26, 28, 29,

30, 31, 32, 33, 34, 36, 37, 38

F Wheat FCF soil ts

35 J Wheat FCF soil ts

FTAD4R 10, 30 D Wheat FCW soil ts

9, 11 I FCW soil ts

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

354 B.B. Landa et al.

approximately 1200 g of sieved Fargo soils (experiment 3),

and seeds were covered with a 1.0-cm layer of soil. Each pot

received 50–200 m L� 1 of water (experiment 3) or water

supplemented with metalaxyl (once per cycle) (Syngenta,

Greensboro, NC) at 2.5 mg mL� 1 active ingredient (experi-

ments 1 and 2) to control Pythium root rot. Each treatment

was replicated six to eight times depending on the experi-

ment, with each pot serving as a replicate. Plants were grown

in a growth chamber at 15–18 1C (experiments 1 and 3) and

at 21–23 1C (experiment 2), 40–60% relative humidity

(RH), and with a 12-h photoperiod. Pots were watered as

needed, and twice a week received 50–100 mL of a Miracle-

Gro solution 15–30–15 (1 g L� 1; Scotts, Port Washington,

NY). After three weeks, one or two plants were harvested

from each pot, and the roots with tightly adhering soil were

prepared as described below to determine the population

size of indigenous phlD1 isolates. The soil and remaining

root systems from pots of each treatment were decanted

into a plastic bag, mixed by shaking, returned to the same

pots immediately (experiments 2 and 3) or after 1 week of

storage at 15 1C (experiment 1), and planted again with new

seeds.

To determine the population densities of total and phlD1

Pseudomonas spp. in the rhizospheres of wheat and flax, the

entire root system was excised from the seedling, and roots

with associated rhizosphere soil were placed in 50-mL

screw-cap centrifuge tubes containing 10 mL of sterile water.

The tube was agitated vigorously for 1 min on a Vortex

mixer and then sonicated in an ultrasonic bath for 1 min. In

experiment 1, the wash solution was serially diluted (1 : 10)

and 100-mL aliquots were plated in duplicate onto 1/3�KMB111 agar plates. Plates were incubated at room

temperature (22–25 1C) in the dark, and total colonies were

counted after 48 h. Colonies of phlD1 fluorescent

Table 1. Continued.

Prefix or strain�Isolate

number GenotypewPlant

source

Soil

samplez Ref.‰

FSB4R 2, 27, 28, 29, 36, 37, 38, 64, 65,

66

J Wheat FCR soil ts

Experiment 2

P-C1Ffz-y 2-2 J Flax FCF soil ts

P-C2Ffz-y 3-1, 3-2, 4-2, 5-1, 5-2, 7-1, 7-3,

8-1, 8-2, 8-3

J Flax FCF soil ts

P-C3Ffz-y 2-1 J Flax FCF soil ts

P-C4Ffz-y 5-1, 5-2 J Flax FCF soil ts

4-2, 8-4 T FCF soil ts

P-C1Fwz-y 2-1, 2-2 J Wheat FCF soil ts

P-C3Fwz-y 1-1, 5-1 J Wheat FCF soil ts

7-1 T FCF soil ts

P-C4Fwz-y 5-1 J Wheat FCF soil ts

2-1, 3-1 T ts

P-C1Wfz-y 5-1 J Flax FCW soil ts

P-C2Wfz-y 1-1, 3-1, 3-3, 6-3 J Flax FCW soil ts

P-C3Wfz-y 1-1, 3-1, 5-1, 6-1, 8-1 J Flax FCW soil ts

P-C4Wfz-y 1-1 J Flax FCW soil ts

P-C2Wwz-y 4-2 J Wheat FCW soil ts

P-C3Wwz-y 4-1 J Wheat FCW soil ts

P-C4Wwz-y 4-1, 4-2 J Wheat FCW soil ts

�In experiment 1, phlD1 fluorescent Pseudomonas spp. were isolated from the rhizosphere of wheat grown in Fargo continuous flax (FCF), Fargo

continuous wheat (FCW) and Fargo crop rotation (FCR) soils and isolates obtained were designated as FFLxRy, FTADxRy and FSBxRy, respectively, where

x represents the cycle number, and y the isolate number. In experiment 2, phlD1 fluorescent Pseudomonas spp. were isolated from the rhizospheres of

wheat and flax grown in FCW or FCF soils. Isolates obtained from wheat in FCW and FCF soils were designated as P-CxWwz-y and P-CxFwz-y,

respectively; isolates obtained from flax in FCW or FCF soils were designated as P-CxWfz-y and P-CxFfz-y, respectively; x represents the cycle number, y

the isolate number, and z the replicate.wGenotype was determined by whole-cell repetitive sequence-based (rep)-PCR using the BOX-PCR and restriction fragment length polymorphism (RFLP)

analysis of phlD (RFLP-phlD) with HaeIII, MspI and TaqI restriction enzymes [McSpadden-Gardener et al. (2001), Landa et al., (2002a)] in experiment 1

and by RFLP- phlD in experiment 2.zThe soil designated as FCW had been cropped to hard red spring wheat since 1882, and the soil designated as FCF had been cropped to flax since 1894.

The soil designated as FCR had been under crop rotation, and at the time of sampling was cropped to sugarbeet. All soils were collected in 1997 and

planted in the same year (experiment 1) or were stored at outdoor temperature before planting in 2000 (experiment 2) or 2003 (experiment 3).‰Source of reference; ts, this study.

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

355Pseudomonas spp. in soils after wheat and flax monoculture

Pseudomonas spp. were then determined by colony hybridi-

zation as described previously (Raaijmakers et al., 1997;

Landa et al., 2002b). Representative phlD1 fluorescent

Pseudomonas isolates (about 2–4 colonies per plate from

each soil-crop-cycle-replication combination, to avoid sam-

pling clones of the same isolate) were subcultured on

1/3�KMB111 plates, and then stored in 35% glycerol at

–80 1C. In experiment 1, isolates from wheat grown in FCF,

FCR and FCW soils were designated as FFLxRy, FSBxRy and

FTADxRy, respectively, where x represents the cycle number

and y the isolate number. The genotype of the phlD1 isolates

was determined by BOX-PCR and RFLP-phlD analysis as

described below.

In experiments 2 and 3, population densities of total and

phlD1 fluorescent Pseudomonas spp. in root washings were

determined by the PCR-based dilution-endpoint method

(McSpadden Gardener et al., 2001; Landa et al., 2002b).

Briefly, 100mL of the wash solution obtained as described

above was serially diluted (1 : 3) in a 96-well microtiter plate

pre-filled with 200mL of sterile distilled water per well, and

then 50mL of each dilution was transferred to a well of a 96-

well plate containing 200mL of 1/3�KMB111. These micro-

titer plates were incubated at room temperature, and bacterial

growth in each well was assayed spectrophotometrically

(Dynatech MR5000, Dynatech Laboratories, Burlington,

MA), scoring an optical density at 600 nm (OD600nm) of

Z0.05 as positive. Population sizes of total culturable pseu-

domonads were determined on the basis of the last well

showing growth in 1/3�KMB111. Aliquots from wells

showing growth were tested for the presence of phlD1

Pseudomonas spp. by PCR analysis as described below, and

population sizes of phlD1 isolates were determined on the

basis of the last well where the phlD signal was detected. Landa

et al. (2002b) previously demonstrated that colony hybridiza-

tion and the phlD-specific PCR-based dilution-endpoint assay

used to quantify phlD isolates provide similar results.

In addition, in experiment 2, replica plates were made by

transferring a 100-mL aliquot of the culture from each well

into an equal volume of 35% glycerol, and storing at

� 80 1C. phlD1 isolates (1 isolate per well showing the last

positive signal for phlD) were later isolated by thawing the

plates and streaking a loop of broth from wells of glycerol

replica plates onto 1/3�KMB111 agar. Cells from colonies

showing a diffusible reddish-brown pigment typical of a 2,4-

DAPG producer (Bangera & Thomashow, 1996) were con-

firmed by PCR to contain phlD, stored at � 80 1C, and then

genotypes were determined by RFLP-phlD analysis as de-

scribed below. Isolates recovered from wheat grown in FCW

or FCF soils were designated as P-CxWwz-y or P-CxFwz-y,

respectively, and those from flax grown in FCW or FCF soils

were designated as P-CxWfz-y or P-CxFfz-y, respectively; x

represents the cycle number, y the isolate number and z the

replicate. Also in Experiments 2 and 3, the dominant phlD1

genotype present in a sample was determined directly by

RFLP analysis of amplified phlD fragments present in the last

dilution showing a positive signal for phlD after PCR. No

isolates from Experiment 3 were selected for storage.

Genotypic fingerprintingof indigenousphlD1

fluorescentPseudomonas spp.

A whole-cell rep-PCR fingerprinting method with the BOX-

PCR was used to determine the genotype of over 150 phlD1

fluorescent Pseudomonas isolates from experiment 1. BOX-

PCR and gel electrophoresis were performed as described

previously (McSpadden-Gardener et al., 2000; Landa et al.,

2002a). RFLP analyses of the 629-bp phlD fragment were

performed using HaeIII, MspI and TaqI restriction enzymes

(New England Biolabs, Beverly, MA) as described before

(Mavrodi et al., 2001; McSpadden-Gardener et al., 2001;

Landa et al., 2002a). RFLP-phlD analysis was used to

determine the genotype of over 190 phlD1 fluorescent

Pseudomonas isolates (experiments 1 and 2), and also from

terminal dilution cultures showing a positive signal for phlD

(experiments 2 and 3). The genotypes determined by

BOX-PCR and RFLP-phlD analysis are highly correlated

(McSpadden-Gardener et al., 2001; Landa et al., 2002a).

BOX-PCR and RFLP-phlD fingerprints were converted,

normalized, combined, and analyzed with GelCompar 4.0

software (Applied Maths, Kortrijk, Belgium). For BOX-PCR

analysis, a similarity matrix of the whole densitometric

curves of the gel tracks was calculated using the pairwise

Pearson product–moment correlation coefficient (r value).

Cluster analysis was performed by the unweighted pair group

method using arithmetic averages (UPGMA) and clusters of

genotypes were defined by using the 95th percentile simi-

larity coefficient of replicate assays for identical strains

(McSpadden-Gardener et al., 2000; Landa et al., 2002a). For

RFLP-phlD analysis, a band-matching algorithm was selected

to calculate pairwise similarity matrices with the Dice

coefficient, and cluster analysis was performed with the

UPGMA method (Landa et al., 2002a). BOX-PCR and

RFLP-phlD genotypes of phlD1 fluorescent Pseudomonas

isolates indigenous to Fargo soils were determined by com-

parison of their fingerprints to the database described by

Landa et al. (2002a) and McSpadden-Gardener et al. (2000).

Bacterial seedandsoil treatments

For studies of rhizosphere colonization by introduced

rifampicin-resistant strains, bacteria were cultured on

1/3�KMB111rif agar for 48 h at 27 1C, harvested, washed

twice by centrifugation (8,000 g for 10 min) and resus-

pended in sterile distilled water. Cell densities were deter-

mined by turbidity at OD600nm with a microplate

spectrophotometer and diluted in sterile distilled water to

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

356 B.B. Landa et al.

adjust inoculum densities as needed. In the growth-chamber

experiment, bacteria were introduced into sieved Quincy

virgin soil in a 1% methylcellulose suspension to give

approximately 104 CFU g� 1 fresh weight of soil as pre-

viously described (Landa et al., 2002b). In the field experi-

ment, bacteria were inoculated onto the seed in a 1%

methylcellulose suspension to obtain approximately

104 CFU seed� 1 as previously described (Raaijmakers &

Weller, 2001; Okubara et al., 2004). Methylcellulose was

used to enhance the survival of the bacteria and adherence of

the cells to the seed. The actual density of each strain was

determined by assaying portions of inoculated soil or

inoculated seeds as described below.

Determinationof rhizosphere colonization byselectedgenotypesof2,4-DAPGproducers ingrowth-chamberand field experiments

Seven phlD1 P. fluorescens strains representing six genotypes,

including reference strains for which we have previous

information of their rhizosphere competence (Landa et al.,

2002a, b, 2003) and isolates recovered from wheat roots

grown in FCF and FCW soils, were compared for their ability

to colonize the rhizosphere of wheat and flax under con-

trolled conditions and in the field. In the growth-chamber

experiment, five strains from four genotypes were used:

Q8r1-96 and FTAD1R33 (genotype D), FTAD1R36 (geno-

type I), FFL1R8 (genotype J) and MVP1–4 (genotype P). In

the field study, five strains were used: Q2-87 (genotype B),

Q8r1-96, FFL1R8, 1M1-96 (genotype L) and MVP1-4 (Table

1). Strains representatives of genotypes B and L were included

based on their known rhizosphere competence on wheat in

growth-chamber experiments (Landa et al., 2003, 2002b),

and a P-genotype strain was included because of its aggressive

colonization of pea roots (Landa et al., 2002a).

In the growth-chamber experiment, ten wheat or flax seeds

were sown in pots containing soil inoculated with one of the

bacterial strains and grown as described in experiment 2. The

control treatment consisted of soil amended with a 1%

methylcellulose suspension. Each treatment was replicated

three times with one pot serving as a replicate. Bacteria were

introduced into the soil only at the beginning of the first

cycle. Two plants were selected randomly from each pot after

each cycle to determine the population size of the introduced

bacteria (six plants in total). The entire root system from each

seedling was excised and processed individually as described

above. Bacterial populations of the introduced bacteria were

determined by the PCR-based endpoint-dilution assay as

described above using 1/3�KMB111rif.

In the field study, for each crop–bacterial-strain combi-

nation, two rows (2 m long and spaced 50 cm apart) were

hand-sown 3–5 cm deep with treated seeds of flax (approxi-

mately 500 seeds per row) or wheat (approximately 200

seeds per row) on 25 May 2001. Each crop–bacterial-strain

treatment was replicated six times. Weeds were removed by

hand, and insecticides and fertilizers were applied in accor-

dance with local recommendations. Plants were sampled at

25, 48, 68 and 88 days after sowing to estimate bacterial

rhizosphere colonization. At each sampling date, three

plants were dug at random from each treatment replicate,

and excess soil was removed from the roots by gentle

shaking. Plants were placed in a plastic bag, transferred to

the laboratory, and stored at 4 1C until processing the next

day. Roots (the entire root system obtained from the upper

10–15 cm layer of soil) from the three plants from each

replicate were pooled for processing.

In both growth-chamber and field experiments, popula-

tion densities of introduced bacteria were enumerated using

the phlD-specific PCR-based dilution- endpoint method

described above. Soil (0.5 g) or seeds (10 seeds per treat-

ment) were placed in centrifuge tubes with 10 mL of sterile

distilled water, and roots with tightly adhering soil were

placed in tubes with 10–30 mL of sterile distilled water

(depending on the amount of roots). Samples were then

vortexed and sonicated, each for 1 min. Fifty-microliter

aliquots of each of the serial dilutions (1 : 3) were transferred

to wells of microtiter plates containing 1/3� KMB111rif

broth. Microtiter plates were incubated at room tempera-

ture, and bacterial growth was measured after 72� 4 h

(growth chamber experiment) or 96� 4 h (field experi-

ment) (McSpadden-Gardener et al., 2001; Landa et al.,

2002b). Aliquots from the terminal dilution showing growth

were confirmed to contain phlD1 cells by PCR analysis, and

the genotype of the phlD1 strain was determined by RFLP-

phlD analysis with HaeIII and MspI (growth-chamber

experiment) or HaeIII, TaqI and MspI (field experiment)

restriction enzymes. The RFLP patterns generated by diges-

tion with the respective enzymes were sufficient to distin-

guish the genotypes B, D, I, J, L and P used in the

experiments (Landa et al., 2002a).

Statistical analysis

Treatments in the growth-chamber and field experiments

were arranged in a completely randomized block design.

Population data of bacteria were log10-transformed prior to

statistical analysis. Data were analyzed using STATISTIX 7.0

(Analytical Software, St. Paul, MN). Differences in popula-

tion dynamics and densities of 2,4-DAPG producers among

soils were determined by standard analysis of variance using

the mean population density for all cycles (growth-chamber

experiments) or sampling dates (field experiment) and the

area under the rhizosphere colonization progress curve

(AUCPC) (Landa et al., 2002a). Mean comparisons among

treatments were performed using Fisher’s protected least

significant difference test at P = 0.05.

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

357Pseudomonas spp. in soils after wheat and flax monoculture

Nucleotide sequenceaccession numbers

The phlD sequences for T-genotype strains FFL2R4, FFL2R5,

FFL2R54, FFL2R55, FFL2R61, and FFL3R49 are deposited in

the GenBank nucleotide sequence database with accession

numbers AY928626, AY928622, AY928624, AY928623,

AY928625, and AY928627, respectively (de la Fuente et al.,

2005).

Results

Populationdensitiesof indigenousphlD1

fluorescentPseudomonasspp. inFargosoils

Population densities of phlD1 fluorescent Pseudomonas spp.

isolates generally remain low in bulk soil and increase in the

rhizosphere when a crop is sown. Raaijmakers et al. (1997)

showed that the cycling process ‘activates’ dormant phlD1

isolates in the bulk soil. Therefore, to assess the population

sizes of phlD1 isolates in the three Fargo soils, we cycled

wheat or flax in the soils to activate the indigenous phlD1

isolates.

In experiment 1, the three soils were sown to wheat and

cycled soon after being collected. The population dynamics

of indigenous phlD1 isolates in the wheat and flax soils

during cycling was similar to that observed in previous

studies of cycled wheat and pea monoculture soils from

Washington State (Raaijmakers et al., 1997; Raaijmakers &

Weller, 1998; Landa et al., 2002a); during cycling, phlD1

isolates reach and sustain threshold population sizes (log

5.0 CFU g root� 1) known to be required for disease suppres-

sion. At the end of cycle 1, rhizosphere population sizes of

phlD1 isolates were log 5.3, 5.1 and o log 4.0 CFU g root� 1

(detection limit of colony hybridization) on roots of wheat

grown in the FCF, FCW and FCR soils, respectively (Table

2). At the end of cycle 2, population sizes were 5.7, 5.2, and

o log 4.0 CFU g root� 1 for the FCF, FCW, and FCR soils,

respectively (Table 2). Continued cycling yielded slightly

smaller mean populations in FCF and FCW soils. Mean

population sizes of phlD1 isolates in FCF soil were higher

than in FCW soil. In cycles 3 and 4 in FCR soil, a few phlD1

isolates were detected; however, the population sizes were

close to the detection limit (Table 2).

In experiment 2, population sizes of phlD1 isolates were

determined on both wheat and flax grown in FCF and FCW

soils after 3 years of storage to determine the ability of phlD1

isolates to persist in dry soil. In general, phlD1 isolates

showed the same population dynamics as those detected

three years earlier during experiment 1 (Table 2). Popula-

tion densities of 2,4-DAPG producers on wheat grown in

FCW and FCF soils were greater than log 5.0 CFU g root� 1,

and populations were significantly greater (Po 0.05) on

wheat in the FCF soil than in the FCW soil after cycle 2.

Densities of 2,4-DAPG producers on flax grown in the FCW

and FCF soils were log 5.2 and log 6.0 CFU g root� 1,

respectively, in the first cycle. Subsequently, population

densities increased and were maintained up to the fourth

cycle, with mean values higher than log 5.5 and log

7.0 CFU g root� 1 in the FCW and FCF soils, respectively

(Table 2).

In experiment 3, the persistence of phlD1 isolates was

determined by growing wheat in the FCF, FCW and FCR

soils after 6 years of storage. In cycle 1, rhizosphere popula-

tion densities of phlD1 isolates were below the detection

limit of log 3.26 CFU g root� 1 in all three soils, but, by cycle

2, phlD1 isolates reached population densities of log 5.8 and

log 5.1 CFU g root� 1 in the FCF soil and FCW soil, respec-

tively (Table 2). Population sizes of phlD1 isolates continued

increasing in cycles 3 and 4 in the FCF soil to values close to

log 8.3 CFU g root� 1, but maintained stable population

levels in the FCW soil. phlD1 isolates were not detected in

FCR soil in any of the four cycles analyzed.

Population densities of total culturable Pseudomonas spp.

ranged from log 6.3 to log 8.7 CFU g root� 1 depending of

the growth cycle and experiment, and, with the exception of

experiment 3, did not differ significantly among soils within

an experiment (Table 2). In general, however, phlD1 isolates

comprised a larger percentage of the total culturable Pseu-

domonas spp. on roots from the FCF soil compared with

roots from the FCW soil (data not shown).

Geneticdiversitywithin phlD1 isolates collectedfromFargosoils

We assumed that the process of collecting and shipping the

soils from Fargo, ND and the length of time until the soils

were replanted in Pullman, WA would not significantly alter

the composition of the phlD1 genotypes in the soils. During

experiment 1, single phlD1 isolates from roots grown in the

three Fargo soils were randomly picked from plates of

KMB111 after colony hybridization and the isolates were

stored at � 80 1C. The genetic diversity present within a

collection of one hundred and fifty three phlD1 isolates (105

from FCF soil, 34 from FCW soil, and 14 from FCR soil) was

analyzed by BOX-PCR and RFLP-phlD analysis (Table 1).

From the new phlD1 Fargo isolates and the 29 phlD1

reference strains analyzed, 19 distinct genomic clusters were

defined based on a 95th percentile similarity coefficient of

independent replicates (McSpadden-Gardener et al., 2000),

which was calculated to be 68% for this data set. Seventeen

of these clusters correlated with BOX genotypes A–Q

described previously (McSpadden-Gardener et al., 2000;

Landa et al., 2002a), and a new BOX genotype, T, also was

identified (Fig. 1) among isolates from roots grown in the

FCF soil. We named this genotype as T because new

genotypes (R and S) have been recently described by

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

358 B.B. Landa et al.

McSpadden-Gardener et al. (2005). Furthermore, the addi-

tion of the new Fargo strains to the cluster analysis caused

subtle changes in the topology of the dendrogram shown in

Fig. 1 as compared with dendrograms previously reported

(McSpadden-Gardener et al., 2000; Landa et al., 2002a). For

example, it should be noted that isolates belonging to

genotypes G, J and the new genotype T have the highest

level of similarity of c. 62%. Furthermore, within the cluster

J, two subgroups emerged, with one of them, J1, more closely

related to cluster T, and the second one, J2, more closely

related to cluster G. Cluster J1 includes the type strains

originally identified as J genotype (Fig. 1).

As previously described, genotypes defined by DNA

polymorphisms within phlD correlated almost perfectly

with groupings based on BOX-PCR (McSpadden-Gardener

et al., 2001; Landa et al., 2002a). In the same way, cluster

analysis by UPGMA of phlD restriction patterns among

isolates in this study revealed the same eighteen groups (A

to Q and T) as identified by analysis of BOX-PCR patterns

(Figs 1 and 2). All the strains belonging to J1 and J2 BOX

genotypes shared the same HaeIII, MspI and TaqI restriction

profiles (Fig. 2). RFLP analyses of members of the newly

defined BOX-PCR genotype T yielded patterns very close to

those observed for J genotype isolates (differing slightly in

Table 2. Population densities of indigenous phlD1 fluorescent Pseudomonas spp. and total culturable Pseudomonas spp. in the rhizospheres of wheat

cv. Penawawa and flax cv. Norlin grown in three soils collected from field plots on the campus of North Dakota State University, Fargo, ND

Soil� Crop

log (CFU g root�1 fresh weight)

Cycle 1 Cycle 2 Cycle 3 Cycle 4

Indigenous total culturable Pseudomonas spp.w

Experiment 1

FCF Wheat 7.11‰ 7.12 7.18 7.48

FCW Wheat 7.18 6.82 6.88 7.12

FCR Wheat 6.96 6.92 6.99 7.33

Experiment 2

FCF Flax 6.81 7.84 7.89 7.88

FCW Flax 6.65 6.47 7.03 7.26

FCF Wheat 6.52 7.28 7.66 7.57

FCW Wheat 6.70 7.21 7.22 7.29

Experiment 3

FCF Wheat 6.72 b 6.91 b 8.70 8.70 a

FCW Wheat 7.92 a 7.87 a 8.53 7.61 b

FCR Wheat 6.28 b 7.14 b 8.38 7.27 b

Indigenous phlD1 fluorescent Pseudomonas spp.z

Experiment 1

FCF Wheat 5.30‰ 5.70 a 5.18 a 5.26 a

FCW Wheat 5.07 5.18 b 4.49 b 4.62 b

FCR Wheat oDL oDL o 4.20 c o 4.21 c

Experiment 2

FCF Flax 5.96 a 7.54 a 7.34 a 7.41 a

FCW Flax 5.20 b 5.58 b 5.52 b 5.98 b

FCF Wheat 5.22 5.81 a 6.96 a 6.66 a

FCW Wheat 5.08 5.14 b 5.37 b 5.19 b

Experiment 3

FCF Wheat oDL 5.81 a 8.28 a 8.29 a

FCW Wheat oDL 5.12 b 5.12 b 5.84 b

FCR Wheat oDL oDL oDL oDL

�FCF, Fargo soil under flax monoculture; FCW, Fargo soil under wheat monoculture; FCR, Fargo soil under crop rotation. Soils were planted shortly after

being collected (experiment 1) or stored at outdoor temperature for three (experiment 2) or six (experiment 3) years before planting.wIn experiment 1, population sizes of total Pseudomonas spp. were determined by dilution plating on agar plates of 1/3�KMB111, which is selective

for these bacteria. In experiments 2 and 3 population sizes of total Pseudomonas spp. were determined by the dilution end-point method with

1/3�KMB111.zPopulation sizes of phlD1 Pseudomonas spp. were determined by colony hybridization followed by PCR (experiment 1) (Raaijmakers et al. 1997), or by

a phlD-specific PCR-based dilution end-point method (experiments 2 and 3) (McSpadden-Gardener et al. 2001; Landa et al. 2002b).‰Means followed by the same letter in a column within an experiment or no letter are not significantly different according to Fisher’s protected least

significant difference test (P = 0.05). DL: Population levels below the detection limit. Colonization data were converted to log10 CFU g�1 fresh root

weight before data analysis.

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

359Pseudomonas spp. in soils after wheat and flax monoculture

Fig. 1. Cluster analysis of genomic fingerprint patterns generated by BOX-PCR amplification of whole-cell genomic DNA from representative phlD1

fluorescent Pseudomonas spp. isolates indigenous to Fargo continuous flax (FCF) and Fargo continuous wheat (FCW) soils (see Table 1) and phlD1

reference strains (BOX-PCR genotypes A to Q). Two independent amplifications were used for each isolate, with the exception of FFL3R54. The

unweighted pair-group method with arithmetic averaging (UPGMA) algorithm was applied to the similarity matrix generated from the tracks of the

whole patterns by using the pairwise Pearson product–moment correlation coefficient (r value). The similarity coefficient used to define distinct groups

(see Materials and methods) is noted (m). phlD1 reference strains are noted (�). The newly defined genotype T is marked in gray shade.

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

360 B.B. Landa et al.

Fig. 2. Cluster analysis of combined HaeIII, TaqI and MspI restriction fragment length polymorphism patterns of the 629-bp phlD fragment amplified

from isolates of fluorescent Pseudomonas spp. indigenous to Fargo continuous flax (FCF), Fargo continuous wheat (FCW) and Fargo crop rotation (FCR)

soils. Patterns include those of reference strains (genotypes A to Q). Two independent amplifications were used for each reference strain. The

unweighted pair-group method with arithmetic averaging (UPGMA) algorithm was applied to the similarity matrix generated with the Dice coefficient.

The similarity score (SD) defining distinct genotypes is noted (m). The newly defined genotype T is marked in gray shade.

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

361Pseudomonas spp. in soils after wheat and flax monoculture

the HaeIII restriction pattern). This HaeIII restriction

pattern of BOX–BOX genotype T was unique when

compared with previously described HaeIII patterns

(McSpadden-Gardener et al., 2001; Landa et al., 2002a)

(Fig. 2).

DifferentBOX-PCRphlD1genotypesare enrichedby long-termwheatand flaxmonoculture

PhlD1 isolates were enriched in both FCF and FCW soils, but

the BOX-PCR genotypes found in the rhizosphere of wheat

during cycling differed greatly between the soils. Five geno-

types (D, F, G, J and T) were detected among the phlD1

isolates collected in Experiment 1 from the FCF soil (Fig. 3;

Table 1). Genotypes F and J occurred at the highest frequency

and represented 41% and 39%, respectively, of the total

phlD1 isolates collected from the FCF soil. Genotypes D, G

and T were recovered at lower frequencies (2%, 12% and 6%

of the total, respectively). In contrast, only genotypes D and I

were detected on wheat from the FCW soil, and comprised

77% and 23%, respectively, of the 34 strains analyzed (Fig. 3).

Only J-genotype isolates were detected among those recov-

ered from the FCR soil (Fig. 3, Table 1).

After 3 years of storage, only J- and T-genotype isolates were

recovered from the rhizospheres of wheat and flax grown in the

FCW and FCF soils (Table 1). The results were similar whether

RFLP analysis was performed directly on fragments amplified

in the terminal dilution culture showing a positive signal for

phlD, or whether RFLP-phlD analysis was performed on

isolates recovered from the same positive dilutions by streaking

aliquots onto 1/3� KMB111 agar (Table 1). After the soil

had been stored for 6 years (experiment 3), RFLP-phlD analysis

detected only the J-genotype pattern in terminal dilution

cultures showing a signal for phlD from wheat grown in the

FCF soil (data not shown). A freezer problem prevented

subsequent RFLP-phlD analysis of terminal dilution

cultures showing a signal for phlD from wheat grown in the

FCW soil.

Genotypesof phlD1 fluorescentPseudomonasspp.differ in theirability to colonize therhizospheresofwheatand flax

In growth-chamber experiments, one hour after the bacteria

were introduced into the soil (cycle 0), population sizes

among the strains ranged from log 4.4 to 4.8 CFU g� 1 of soil

and did not differ significantly (PZ0.05). In the wheat

rhizosphere, the population dynamics of D-genotype strains

Q8r1-96 and FTAD1R33 was typical of that shown by D-

genotype strains in previous cycling experiments with wheat

(Landa et al., 2003); population densities increased above

log 6.0 CFU g root� 1 and remained above this level for the

duration of the experiment (12 weeks) (Fig. 4). In the fourth

cycle, the density of Q8r1-96 and FTAD1R33 was near

107 CFU g root� 1. The population dynamics of the I-geno-

type strain FTAD1R36 was similar to that of the two

D-genotype strains. Strains FFL1R8 (J genotype) and

MVP1-4 (P genotype) colonized the wheat rhizosphere

significantly less than the D- and I-genotype strains. By the

fourth cycle, the density of the J-genotype strain was 10-fold

less than those of the D- and I-genotype strains (Fig. 4).

These differences in colonization were also reflected in the

values for AUCPC and mean colonization (Table 3). As in

the wheat rhizosphere, strains FFL1R8 and MVP1-4 colo-

nized the flax rhizosphere to a lesser extent than the D- and

I-genotype strains. All the strains colonized the flax rhizo-

sphere significantly less (Po 0.05) than they did the wheat

rhizosphere, with the exception of FFL1R8 (J genotype),

which colonized both crops to the same extent. The AUCPC

values for FFL1R8 on wheat and flax did not differ signifi-

cantly (PZ0.05) (Table 3).

In the field experiment, at the time of sowing, population

sizes of the introduced strains on flax and wheat seeds

ranged from log 4.3 to 4.6 CFU per seed and did not differ

significantly (PZ0.05). Strain Q8r1-96 colonized the wheat

rhizosphere better than strains Q2-87 (B genotype), FFL1R8

(J genotype), 1M1-96 (L genotype) and MVP1-4 (P geno-

type), as indicated by significantly (Po 0.05) greater

AUCPC and mean colonization values (Fig. 5, Table 3). By

the fourth sampling date (90 days after sowing), the

densities of the B-, L-, J- and P-genotype strains were about

10- to 100-fold less than those of the D-genotype strain (Fig.

5). Strain Q2-87 showed significantly less colonization than

FFL1R8 and 1M1-96. On flax, strain Q8r1-96 showed great-

er rhizosphere colonization than Q2-87, 1M1-96 and

MVP1-4, but did not differ significantly from those of

FFL1R8. Colonization of flax by strain FFL1R8 was signifi-

cantly greater than strains Q2-87 and MVP1-4. With the

exception of FFL1R8, all of the strains colonized wheat

Fig. 3. Percentage of genotypes of fluorescent Pseudomonas spp.

isolates (n = 153) indigenous to three natural soils [Fargo continuous flax

(FCF) soil, n = 105; Fargo continuous wheat (FCW) soil, n = 34; Fargo

crop rotation (FCR) soil, n = 14; see Table 1] collected from field plots at

the North Dakota State University (NDSU), Fargo, ND. Isolates were

characterized by BOX-PCR and RFLP-phlD analysis as members of D, F, G,

I, J or T genotypes. Values are total numbers of phlD1 isolates obtained in

four cycles of wheat growth in FCF, FCW, and FCR soils.

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

362 B.B. Landa et al.

significantly better than flax, as occurred in growth-chamber

studies (Fig. 5, Table 3).

Discussion

The rhizosphere environment consists of three habitats: the

rhizosphere, the rhizoplane and the root interior or endo-

sphere (Lynch, 1990). Within these habitats, microorgan-

isms proliferate and reach population densities 10–100-fold

higher (Andrews & Harris, 2000) than microorganisms in

the bulk soil owing to the loss of organic materials from the

roots (rhizodeposition). This highly diverse microbial com-

munity provides a basal level of buffering, known as general

suppression, against root attack by soilborne pathogens

(Weller et al., 2002). Plant species also initiate and maintain

much more sophisticated mutualistic relationships with

individual or select groups of microbial species or subspe-

cies, resulting in a very high level of pathogen suppression

known as specific suppression (Weller et al., 2002). One

example of specific suppression is TAD, described in

Fig. 4. Population dynamics of phlD1 Pseudo-

monas fluorescens strains Q8r1-96 and

FTAD1R33 (D genotype), FTAD1R36 (I geno-

type), FFL1R8 (J genotype), and MVP1-4

(P genotype) in the rhizospheres of wheat cv.

Penawawa and flax cv. Norlin grown in Quincy

virgin soil for four successive cycles of three

weeks each. Each strain was introduced into the

soil to give a final density of approximately

104 CFU g� 1 of soil (cycle 0). Mean values and

standard deviations are presented.

Table 3. Population dynamics of introduced 2,4-diacetylphloroglucinol-producing Pseudomonas spp. in the rhizospheres of wheat cv. Penawawa and

flax cv. Norlin grown in natural soils

Bacterial strain� BOX genotypew

Flax Wheat

Mean colonization

log (CFU g root�1)z AUCPC‰

Mean colonization

log (CFU g root�1)w AUCPC‰

Growth-chamber experiment

Q8r1-96 D 5.92 23.26 az 7.02 28.51 a�

FTAD1R33 D 5.87 22.82 ab 6.92 27.23 b�

FTAD1R36 I 5.74 22.30 b 6.42 25.68 c�

FFL1R8 J 5.23 20.33 c 5.65 21.85 e

MVP1-4 P o 4.89 o 18.73 d 6.17 23.31 d�

Field experiment

Q2-87 B o 3.95 o 337.59 c 4.60 411.60 c�

Q8r1-96 D 5.88 435.47 a 6.57 529.71 a�

FFL1R8 J 5.18 424.67 ab 5.63 461.85 b

1M1-96 L 4.94 384.96 bc 5.66 472.30 b�

MVP1-4 P o 4.41 o 358.20 c 5.61 431.60 bc�

�In the growth chamber, bacterial isolates were introduced into a natural virgin soil from Quincy, WA, to give a final density of approximately

104 CFU g� 1 of soil. Each treatment consisted of three pots with ten plants per pot; two root systems per pot were sampled to estimate the bacterial

population on roots. Plants were grown for four cycles of three weeks each in a growth chamber at 22� 1 1C (See Fig. 4). In the field experiment,

bacterial isolates were introduced on the seeds to give a final density of approximately 104 CFU per seed. Treated seeds of flax or wheat were hand-sown

on 25 May 2001, and plants were sampled at 25, 48, 68 and 88 days after sowing to estimate bacterial rhizosphere colonization (See Fig. 5). There were

six replications for each crop-bacterial isolate treatment, and treatments were arranged in a completely randomized block design.wGenotypes were defined previously by BOX-PCR genomic fingerprinting.zMean population density across all cycles or sampling dates except for cycle 0 or date of sowing. Population data were converted to log10 CFU g root�1.‰AUCPC: area under the rhizosphere colonization progress curve calculated using the trapezoidal integration method (Landa et al. 2002a).zMeans in a column followed by the same letter are not significantly different (P = 0.05) according to Fisher’s protected least significant difference test.

Asterisk indicates a significantly higher (Po 0.05) rhizosphere population for a bacterial strain between crops.

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

363Pseudomonas spp. in soils after wheat and flax monoculture

Washington State and the Netherlands, resulting from the

buildup of isolates of 2,4-DAPG-producing fluorescent

Pseudomonas spp. to a threshold density of

105 CFU g root� 1. In some soils, phlD1 strains may border

on being mutualistic symbionts (Andrews & Harris, 2000),

because the plant benefits from a highly effective line of

defence against pathogens for which it has no resistance, and

the 2,4-DAPG producers benefit from a substantial increase

in population size in a highly competitive environment

(Weller et al., 2002).

Our studies with the Fargo soils showed that over 100

years of flax or wheat monoculture, but not crop rotation,

led to an enrichment of 2,4-DAPG producers. In addition,

phlD1 populations showed considerable persistence in the

bulk soil once enriched by monoculture, and, although a

decrease in density occurs in the soil in the absence of the

plant, populations increase dramatically in the rhizosphere

when another crop is sown. These findings, coupled with

our previous findings of the enrichment of 2,4-DAPG

producers in the Washington State monoculture pea field

(Landa et al., 2002a), lead us to conclude that monoculture

is an important stimulus for enrichment of phlD1 isolates in

the rhizosphere environment of other plants species besides

wheat and barley. These current findings are especially

convincing because the fields from which the FCW, FCF

and FCR soils were collected are located next to each other

and the unique histories of the fields are well documented.

Although we have identified monoculture as a stimulus for

enriching phlD1 isolates, we are not ruling out the ability of

other cropping practices or crop rotations to facilitate

enrichment. For example, McSpadden-Gardener et al.

(2005) reported a greater frequency of 2,4-DAPG producers

on the roots of corn than on soybean roots in Ohio fields

where crop rotations are commonly practised.

BOX-PCR and RFLP-phlD analyses showed that a new

genotype (genotype T) was present in the FCF soil. The

novelty of this genotype has been confirmed by phylogenetic

analysis of phlD using T-genotype isolates and isolates from

the 17 other genotypes (de la Fuente et al., 2005). With the

discovery of the T genotype, and the new genotypes

reported recently by McSpadden-Gardener et al. (2005) (R

and S) and by Mazzola et al. (2004) (Yand Z), the number of

BOX-PCR genotypes now stands at 22. However, the actual

number of genotypes in soils worldwide and the basis for

their biogeography is not known. Other methods of mole-

cular fingerprinting have identified greater numbers of

phlD1 genotypes (Picard et al., 2000; McSpadden-Gardener

et al., 2000; Picard & Bosco, 2003; Bergsma-Vlami et al.,

2005a,b). We prefer rep-PCR because of the good correla-

tion between groupings identified by BOX-PCR and RFLP-

phlD analysis (Mavrodi et al., 2001; McSpadden-Gardener

et al., 2001; Landa et al., 2002a), and phylogenetic analysis of

phlD (de la Fuente et al., 2005). More importantly, however,

the BOX-PCR genotype is predictive of certain in situ

biological activities such as root colonization (Raaijmakers

& Weller, 2001; Landa et al., 2002a; Weller et al., 2002).

Therefore, it is notable that the D-genotype strain

FTAD1R33 from the FCW field showed population dy-

namics on wheat in cycling experiments typical of D-

genotype strains isolated from two Washington State TAD

fields (Fig. 4) (Landa et al., 2003). Fargo, ND is approxi-

mately 1600 km from central Washington State, making it

unlikely that the similarities among D-genotype isolates

from Fargo, ND, Lind, WA and Quincy, WA resulted from

soil-contamination events.

In studies of the genetic diversity within phlD1 isolates in

European and American collections, McSpadden-Gardener

et al. (2000) concluded that these bacteria showed a sub-

stantial degree of endemicity; however, certain genotypes

such as the A and D appear to be more widely distributed

than others (McSpadden-Gardener et al., 2001). Genotype A

has been isolated from birdsfoot trefoil, corn, cotton,

cucumber, tobacco, pea and soybeans from soils in the

Czech Republic, Ghana, Italy, Switzerland, Uruguay and

various states in the USA (McSpadden-Gardener et al.,

2000, 2005; Wang et al., 2001; Landa et al., 2002a; de la

Fig. 5. Population dynamics of phlD1 Pseudo-

monas fluorescens strains Q2-87 (B genotype),

Q8r1-96 (D genotype), FFL1R8 (J genotype),

1M1-96 (L genotype), and MVP1-4 (P genotype)

in the rhizospheres of wheat cv. Penawawa and

flax cv. Norlin grown in a field plot located at the

WSU Plant Pathology Farm in Pullman, WA.

Each strain was introduced onto the seed to give

a final density of approximately 104 CFU per

seed at sowing. Mean values and standard

deviations are presented.

FEMS Microbiol Ecol 55 (2006) 351–368c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

364 B.B. Landa et al.

Fuente et al., 2004b); and genotype D has been isolated from

a variety of crops in France, Germany, Spain, Switzerland,

and various states in the USA [McSpadden-Gardener et al.,

2000, 2005, B. B. Landa and D. M. Weller, unpublished

results]. Within a given soil, it is not yet clear how many

phlD1 genotypes are naturally present and whether all the

genotypes present at very low densities are being found by

current approaches. For example, in soil from the mono-

culture pea field at Mt. Vernon, WA, genotypes A, D, L, O, P

and Q were isolated, whereas in soils from TAD fields in

Quincy and Lind, WA, genotypes B, D and E, and genotypes

and D and L, respectively, were found. Among the three

Fargo soils sampled, six genotypes, D, F, G, I, J and T, were

found. However, in some soils collected from monoculture

wheat fields throughout the US only one genotype has so far

been isolated (McSpadden-Gardener et al., 2000).

A key unanswered question is why certain genotypes

become dominant in the rhizosphere environment of a plant

species when multiple genotypes are present in the soil. For

example, in the Washington State TAD soils, D-genotype

strains are dominant, and in the Mt. Vernon continuous pea

soil, D and P genotypes are both in abundance (Landa et al.,

2002a, 2003). In this study, we showed that in the FCW soil

D and I genotypes comprised 77% and 23% of the phlD1

isolates, whereas in the FCF soil F and J genotypes were

dominant, comprising about 41% and 39%, respectively, of

the phlD1 isolates when wheat was sown in the soils soon

after they were collected from the field.

One possible explanation for these findings is that plant

species play a central role in establishing a rhizosphere

environment that is physically, chemically or nutritionally

more ‘hospitable’ for certain genotypes already present in

the soil ‘seed bank’ of genotypes, and ultimately over several

years of monoculture the growth of one or more of the

genotypes is favoured and becomes dominant. It is well

known that the quantity and composition of the rhizodepo-

sition and root architecture is under the genetic control of

the plant, and, as a consequence, the plant genotype plays a

central role in modulating the general microbial community

structure (Atkinson et al., 1975; Grayston et al., 1998;

Miethling et al., 2000; Wieland et al., 2001; Kuske et al.,

2002) as well as populations of specific genera and species

(Lemanceau et al., 1995; Smith et al., 1999; Smith & Good-

man, 1999; Mazzola & Gu, 2000; Berg et al., 2002; Briones

et al., 2002; Da Mota et al., 2002), including 2,4-DAPG

producers (de la Fuente et al., 2004a; Picard et al., 2004;

Bergsma-Vlami et al., 2005a), in the rhizosphere environ-

ment. Recently, Picard et al. (2004) reported that the

population sizes and genetic diversity of phlD1 isolates were

significantly greater in the rhizosphere of hybrid maize than

in the rhizospheres of the two parental lines. Mazzola et al.

(2004) reported that the wheat cultivar Lewjain grown in

apple orchard soils supported higher populations of phlD1

isolates than four other cultivars grown in the same soils.

Finally, Bergsma-Vlami et al. (2005a) reported that the

population size and genetic diversity of phlD1 isolates were

significantly lower in the rhizosphere of lily grown in take-all

suppressive and conducive soils as compared with wheat,

sugarbeet or potato grown in the same soils. Although this

explanation is appealing, it should be tempered by the fact

that few differences have been found among carbon utiliza-

tion profiles of phlD1 isolates representing different geno-

types defined by BOX-PCR or RFLP-phlD analysis

(McSpadden-Gardener et al., 2000; Raaijmakers & Weller,

2001; Wang et al., 2001). However, classical carbon and

nitrogen utilization tests probably do not reflect the full

range of organic substrates and concentrations found in

rhizodeposition.

A second possibility is that certain phlD1 genotypes have

an affinity or preference for the roots of certain plant species

or even cultivars of a species. Unknown bacterial and/or

plant traits would allow certain genotypes to colonize more

aggressively the rhizosphere environment by more efficient

attachment to the root, transport with the growing tip,

colonization of and growth in favoured sites, and/or survival

in senescent root tissues. When applied as seed or soil

treatments, isolates of the different genotypes vary consider-

ably in rhizosphere competence on certain plants. For

example, D- and K-genotype strains are significantly more

rhizosphere-competent on wheat than isolates of genotypes

A, B and L (Landa et al., 2003), and D- and P-genotype

strains are significantly more competent on pea, compared

with isolates of genotypes A, E, L, O and Q (Landa et al.,

2002a). In the current study, the genotypes tested differed in

rhizosphere competence on both wheat and flax, with the

D-genotype strains again being the most aggressive colonist.

All of the genotypes tested showed greater colonization of

wheat than of flax in growth-chamber and field experi-

ments, except for the J-genotype isolate, which was equiva-

lent on both crops. At this time, we cannot explain why

indigenous phlD1 population sizes on flax grown in FCW

and FCF soils were greater than on wheat grown in the same

soils, yet, when individual strains (with the exception of the

J genotype) were introduced on wheat and flax, population

sizes were greater on wheat. However, the physicochemical

soil properties of both soils used in growth-chamber and

field experiments and the environmental conditions under

which they were conducted may play a role in the results.

A third possibility is that the niche overlap in the rhizo-

sphere environment, expected for genotypes of a single

subspecies of P. fluorescens, results in intra-genotype compe-

tition and antagonism. Studies on both wheat (Landa et al.,

2003) and pea (Landa et al., 2002a) support this possibility.

For example, Landa et al. (2003) demonstrated that

D-genotype strains displaced A-, B- and L-genotype strains

in the wheat rhizosphere in co-inoculation studies, yet the D

FEMS Microbiol Ecol 55 (2006) 351–368 c� 2005 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

365Pseudomonas spp. in soils after wheat and flax monoculture

genotype was displaced by the K genotype. Furthermore, the

use of the de Wit replacement series demonstrated a

competitive disadvantage for strain Q2-87 (B genotype) or

strong antagonism by strain Q8r1-96 against Q2-87 in the

wheat rhizosphere (Landa et al., 2003). Recently, Validov

et al. (2005) screened 47 phlD1 isolates representing 17

genotypes and showed that, upon induction, over 70% of

the isolates produced bacteriocins inhibitory to other geno-

types. Similar to results found by Landa et al. (2003), their

study showed that in paired inoculations strain Q8r1-96

(antagonistic) out-competed different strains of different

genotypes in the wheat rhizosphere (both sensitive and

neutral), and concluded that bacteriocin production may

have a role in rhizosphere interactions among 2,4-DAPG

producers.

In conclusion, we think that all three of the possibilities

described above may be involved to some extent in the

enrichment of different genotypes during wheat and flax

monoculture. The mechanisms underpinning the broader

phenomenon of enrichment of populations of 2,4-DAPG

producers by crop monoculture and the specific enrichment

of certain genotypes are not known but are topics of

considerable interest.

Acknowledgements

This project was supported by National Research Initiative

Competitive Grant 01-35107-1011 from the USDA Coop-

erative State Research, Education, and Extension Service.

Blanca B. Landa was recipient of a postdoctoral fellowship

from the Fulbright Commission and the Spanish Ministry of

Science and Technology and a grant from the University of

Cordoba, Spain while conducting the experiments, and

currently is a contract holder of the ‘Ramon y Cajal’

programme of the Ministerio de Ciencia y Tecnologıa of

Spain. We thank S. Blouin-Banhead, S. Kalloger and E. C.

Sachs for technical assistance, Suellen, Nathan and Keith

Weller for helping to collect the soils, A. Felip Edo for her

support and useful discussions, and Robert Stack, Albert

Schneiter and Burton Johnson, North Dakota State Uni-

versity for providing access to and information about the

NDSU field plots.

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