University of Reading

School of Animal and Microbial Sciences

Regulation of Carbon Metabolism in Rhizobium leguminosarum

By

Emma Mary Lodwig

Submitted in partial fulfilment of the requirement for the degree of Doctor of Philosophy 2001

II

I declare that this is my own account of my research and that this work has not

been submitted for a degree at any other university. I would like to acknowledge

that chapter 4 (Alanine secretion by pea bacteroids) was carried out jointly with

David Allaway, and that certain vectors and strains were constructed by members

of the laboratory, as described in the text.

Emma Mary Lodwig

III

ACKNOWLEDGEMENTS

The work presented in this thesis is a component of the research carried out in

two prosperous research groups headed by my supervisors Philip Poole and

Allan Downie. I would like to thank them both for giving me the opportunity to

carry out this research and for their guidance throughout the last three years. I

wish them both continued success.

I would like to thank several people who have made significant contributions to

the work presented. Firstly, and most importantly, David Allaway for his inputs to

the alanine secretion and regulation work which has ranged considerably from

the harvesting hundreds of peas at the break of dawn through to the running of

biochemical assays and the construction of plasmids and strains. Kim Findlay for

carrying out all the microscopy work. Philip Poole, Mary Leonard, Alexandre

Bourdes, Sylvia Marroqui and David Walshaw for the construction of strains and

plasmids. Tim Wheeler for the provision of greenhouse space at PEL and advise

on experiments. Les Crompton for amino acid analysis and Richard Parsons for

15N analysis.

I would like to acknowledge the support from technical staff at both AMS central

services and at PEL, whose names are too many to mention.

I would also like to acknowledge the BBSRC for funding and my parents for their

ongoing financial support.

IV

ABSTRACT

Rhizobium leguminosarum may form determinate or indeterminate nodules with

beans and peas (biovars phaseoli and viciae respectively). During the symbiotic

interaction of both nodule type, dicarboxylates are provided by the plant to the

bacteroid to fuel nitrogen fixation. Bacteroids produce carbon storage compounds

and / or amino acids to regulate carbon metabolism under the microaerobic

conditions of the nodule.

Bacteroids of bean nodules produce significant quantities of the carbon storage

product poly-β-hydroxybutyrate (PHB), yet bacteroids of pea produce none.

Mutation of phaC (PHB synthase) in R. leguminosarum strain A5 (mutant

RU1329) prevents PHB synthesis in free-living cells and in bacteroids but has no

significant effect on the efficiency of the bean symbiosis (nitrogenase activity or

dry weight of bean plants). However, a phaC mutant (RU1328) in A34, an

isogenic strain of A5 that infects pea, which is unable to accumulate PHB in free-

living cells exhibits a variable effect on the symbiotic performance of pea plants.

Transmission electron micrographs show PHB is present in wild type bacteria

present inside the infection thread which disappears during bacteroid

differentiation. Also plant cells in interzone II-III of the indeterminate pea nodule

formed by RU1328 do not contain starch. Together these data suggest both PHB

and plant starch are used to fuel bacteroid differentiation.

Along with ammonium, considerable amounts of the amino acid alanine is

excreted from purified pea bacteroids actively fixing nitrogen. The amount of

alanine excreted can be manipulated by addition of exogenous ammonium

V

chloride or by concentrating the bacteroid suspension. Therefore, excretion of

alanine is dependant on, and proportional to, the relative ammonium

concentration of the bacteroid suspension medium. Concentrated bacteroids

incubated with 15N2 gas excrete 15N labelled alanine demonstrating that the

nitrogen incorporated in alanine is derived from nitrogen fixation.

Mutation of the gene encoding L-alanine dehydrogenase (aldA) results in

bacteroids devoid of AldA activity that are unable to excrete alanine, confirming

that alanine is synthesised by AldA. This mutant strain (RU1327) does not have

altered nitrogenase activity (acetylene reduction) but reduces the dry weight of

peas by 20% indicating a role for AldA in the pea symbiosis.

A mutant in the regulatory gene dadR (RU1275) is unable to grow on alanine

indicating the dadXA gene products are used in the catabolism of L-alanine via

alanine racemase (DadX) and D-alanine dehydrogenase (DadA). AldA is not

utilised in the degradation of L-alanine, but is able to suppress RU1275 for growth

on alanine when aldA is present in multiple copies on a cosmid. L-alanine

dehydrogenase activity was induced by growth of free living cells on

dicarboxylates and alanine. Northern analysis of aldA mRNA levels and GUS

fusions made to aldA and aldR (the regulatory gene) confirmed this induction

pattern.

VI

LIST OF ABBREVIATIONS

AAT Aspartate aminotransferase

ACN Aconitase

ALD L-Alanine dehydrogenase

aldA Gene encoding L-alanine dehydrogenase

aldR Gene encoding regulator for L-alanine dehydrogenase

ω-AM ω-Amidase

amp Ampicillin

AMS Acid minimal salts

ATP Adenosine triphosphate

BM Bacteroid membrane

bp base pair

bv. biovar

BSA Bovine serum albumin

cam Chloramphenicol

Ci Curie

CPM Counts per minute

CS Citrate synthase

cv. cultivar

Cy5 Carbocyanine dye for labelling oligonucleotides

dadA Gene encoding D-alanine dehydrogenase

dadR Gene encoding regulator of dadXA

dadX Gene encoding alanine racemase

Dct Dicarboxylate transport

DEPC diethylpyrocarbonate

VII

DME NAD+ dependent malic enzyme

DNA Dioxyribonucleic acid

[32P]dCTP 2’-deoxycytidine 5’-[α-32P] triphosphate

dNTP 2’-deoxynucleoside 5’-triphosphate

DTT Dithiothreitol

EDTA Ethylenediamine tetraacetic acid

EMBL European molecular biology laboratory

EPS Exopolysaccharide

et al. et alii

FA Formaldehyde agarose

Fig Figure

Fix+ Nitrogen fixing phenotype

Fix- Non-nitrogen fixing phenotype

FUM Fumarase

G6P-DH/HK glucose-6-phosphate dehydrogenase/hexokinase

GABA γ-Aminobutyrate

GAM Glutaminase

GAT γ-aminobuyrate aminotransferase

GC Gas chromatography

GDC Glutamate decarboxylase

GDH Glutamate dehydrogenase

GMAT Glutamine aminotransferase

glgA Gene encoding glycogen synthase

GOGAT Glutamate synthase

gen Gentamicin

GS Glutamine synthetase

VIII

GUS β-glucuronidase

HEPES N-[2-hydroxyethyl]piperazine-N’-[2-ethanesulphonic acid]

ICDH Isocitrate dehydrogenase

ICL Isocitrate lyase

IPTG Isopropyl β-D-thiogalactoside

IS50 (R & L) Insertion sequence 50 (right and left)

ITM Infection thread membrane

Kb Kilobase

kan Kanamycin

KDC α-Ketoglutarate decarboxylase

KGDH α-Ketoglutarate dehydrogenase

Km Substrate concentration at which reaction rate is half of its maximum

LB Luria-Bertani broth

MDH Malate dehydrogenase

ME Malic enzyme

MOPS 3-[N-morpholino]propanesulfonic acid

MS Malate synthase

mRNA Messanger ribonucleic acid

N2ase Nitrogenase

NAD+ Nicotinamide adenine dinucleotide (oxidised form)

NADH Nicotinamide adenine dinucleotide (reduced form)

NADP + Nicotinamide adenine dinucleotide phosphate (oxidised form)

NADPH Nicotinamide adenine dinucleotide phosphate (reduced form)

OD Optical density

PBM Peribacteroid membrane

PBS Peribacteroid space

IX

PBU Peribacteroid unit

PCR Polymerase chain reaction

PDH Pyruvate dehydrogenase

PEPCK Phosphoenolpyruvate carboxykinase

phaC Gene encoding poly-β-hydroxybutyrate synthase

PHB Poly-β-hydroxybutyrate

PNPG ρ-nitrophenyl β-D-galacto pyranoside

pSym Symbiotic plasmid

RIME Rhizobium-specific intergenic mosaic element

rif Rifampicin

RMS Rhizobium minimal salts

RNA Ribonucleic acid

RNase Ribonuclease

rpm Revolutions per minute

SCS Succinyl-CoA synthetase

SE Standard error

SDH Succinate dehydrogenase

SDS Sodium dodecyl sulphate

SS Sucrose synthase

SSC Sodium citrate solution for hybridising DNA and RNA protocols

SSDH Succinic semialdehyde dehydrogenase

SSPE Sodium phosphate EDTA solution for hybridising DNA and RNA protocols

spc Spectinomycin

str Streptomycin

TCA Tricarboxylic acid

tet Tetracycline

X

TEM Transmission electron microscopy

TME NADP+ dependent malic enzyme

Tn5 Kanamycin resistant transposon

TnB20 Tn5 based transposon carrying LacZ reporter gene

TnB60 Tn5 based transposon carrying tac promoter

TRIS 2-amino-2-(hydroxymethyl)-1,3-propanediol

TY Tryptone-yeast media

UV Ultra violet

Vmax Maximum rate

w/v Weight for volume

WT Wild type

X-Gal 5-bromo-4-chloro-3-indolyl-β-D-galactoside

XI

CONTENTS

1 INTRODUCTION..............................................................................2

1.1 Rhizobium taxonomy ........................................................................................ 3

1.2 Nodule formation, structure and environment .................................................. 5

1.2.1 Molecular basis for nodulation................................................................... 5

1.2.2 Events of nodulation.................................................................................. 6

1.2.2.1 Determinate nodules .......................................................................... 6

1.2.2.2 Indeterminate nodules........................................................................ 7

1.2.3 Formation of the symbiosome.................................................................10

1.2.4 The microaerobic nodule environment....................................................10

1.3 Exchange of metabolites in the nodule...........................................................12

1.3.1 Carbon provision to the bacteroid ...........................................................12

1.3.2 Nitrogen provision to the plant.................................................................15

1.4 Carbon metabolism in the bacteroid...............................................................20

1.4.1 The tricarboxylic acid (TCA) cycle...........................................................20

1.4.2 Mutation analysis of TCA cycle enzymes................................................21

1.4.3 Formation of acetyl-CoA: the role of malic enzymes ..............................26

1.4.4 Poly-β-hydroxybutyrate (PHB).................................................................32

1.4.4.1 Molecular basis for PHB metabolism...............................................32

1.4.4.2 PHB metabolism in bacteroids.........................................................37

1.4.4.3 Role of PHB in bacteroids................................................................40

1.4.5 Glycogen .................................................................................................51

1.5 Amino acid biosynthesis and metabolism ......................................................56

1.5.1 Bacteroid glutamate metabolism.............................................................56

1.5.2 Bacteroid alanine metabolism .................................................................61

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1.5.3 Bacteroid aspartate metabolism..............................................................67

1.6 Research objectives .......................................................................................70

2 MATERIALS AND METHODS.......................................................72

2.1 General ...........................................................................................................72

2.2 Culture conditions ...........................................................................................72

2.3 List of antibiotic concentrations ......................................................................73

2.4 List of strains...................................................................................................74

2.5 List of plasmids, cosmids and phage..............................................................76

2.6 List of primers .................................................................................................79

2.7 DNA manipulation...........................................................................................80

2.7.1 Agarose gel electrophoresis....................................................................80

2.7.2 DNA isolation...........................................................................................80

2.7.3 Restriction digests ...................................................................................80

2.7.4 Partial restriction digests .........................................................................81

2.7.5 Fill in of overhangs ..................................................................................81

2.7.6 Ligation ....................................................................................................81

2.7.7 Transformation ........................................................................................81

2.7.8 Southern blot & hybridisation ..................................................................82

2.7.9 Quantitation of DNA ................................................................................83

2.7.10 Polymerase Chain Reaction (PCR).......................................................83

2.7.11 Sequencing & Sequence Analysis ........................................................84

2.8 Transposon mutagenesis ...............................................................................85

2.8.1 Phage propagation ..................................................................................85

2.8.2 Mutagenesis ............................................................................................85

2.9 Conjugation.....................................................................................................87

2.10 RNA manipulation.........................................................................................88

XIII

2.10.1 RNA isolation.........................................................................................88

2.10.2 Quantitation of RNA ..............................................................................88

2.10.3 RNA electrophoresis .............................................................................89

2.10.4 Northern blot & hybridisation.................................................................89

2.11 Assay of β-glucuronidase (GUS) activity ......................................................91

2.12 PHB quantification ........................................................................................92

2.13 Glycogen quantification ................................................................................93

2.13.1 Extraction of glycogen from free-living cells..........................................93

2.13.2 Extraction of glycogen from bacteroids .................................................93

2.13.3 Digestion of glycogen to glucose ..........................................................93

2.13.4 Determination of glucose formed ..........................................................94

2.14 Enzyme assays.............................................................................................95

2.14.1 Preparation of protein extracts ..............................................................95

2.14.2 Assays of malate dehydrogenase, citrate synthase, isocitrate dehydrogenase and L-alanine dehydrogenase................................................95

2.15 Protein Determination ...................................................................................96

2.16 Plant Experiments.........................................................................................97

2.16.1 Seed sterilisation, sowing and inoculation ............................................97

2.16.2 Assay of nitrogenase activity (acetylene reduction)..............................98

2.16.3 Plant dry weights ...................................................................................99

2.16.4 Determination of nodulating strain ........................................................99

2.16.5 Microscopy of nodule sections ..............................................................99

2.17 Bacteroid excretion studies ........................................................................101

2.17.1 Percoll purification of bacteroids .........................................................101

2.17.2 Incubation of isolated bacteroids.........................................................101

2.17.3 Quantification of alanine......................................................................102

2.17.4 Quantification of ammonium................................................................103

XIV

2.17.5 15N2 labelling studies ...............................................................................104

2.17.6 Amino acid analysis.............................................................................105

3 IDENTIFICATION & CHARACTERISATION OF CARBON STORAGE COMPOUNDS IN RHIZOBIUM LEGUMINOSARUM SYMBIOSES ...................................................................................107

3.1 Analysis of the sequence of phaC (PHB synthase)......................................109

3.2 Construction of mutants in PHB synthase....................................................118

3.3 Growth of PHB synthase mutants ................................................................122

3.4 PHB content of free-living strains .................................................................125

3.5 PHB content of pea and bean bacteroids.....................................................126

3.6 Symbiotic performance of PHB synthase mutants on pea and bean...........145

3.7 TCA cycle enzyme activity of pea bacteroids...............................................152

3.8 Glycogen content of pea bacteroids .............................................................154

3.9 Second test of symbiotic performance of PHB mutant on pea.....................155

3.10 Construction of a mutant in glycogen synthase..........................................156

3.11 Growth of mutant in glycogen synthase .....................................................162

3.12 Glycogen content of free-living strains .......................................................164

3.13 Symbiotic phenotype of glycogen synthase mutant on pea .......................164

3.14 Discussion...................................................................................................167

4 IDENTIFICATION OF THE ROLE OF ALANINE SYNTHESIS AND EXCRETION BY BACTEROIDS OF PEA ......................................174

4.1 Isolation of pure bacteroid samples..............................................................176

4.2 Optimisation of conditions for nitrogen fixation.............................................178

4.2.1 Oxygen concentration ...........................................................................178

4.2.2 Carbon source .......................................................................................179

4.3 Ammonium and alanine are excreted from pea bacteroids as products of N2 fixation.................................................................................................................183

XV

4.3.1 Alanine is a product of de novo synthesis.............................................185

4.3.2 Alanine is the sole amino acid excreted................................................185

4.4 Alanine production is dependant on the ammonium concentration .............186

4.5 Alanine synthesis is not directly coupled to N2 fixation ................................189

4.6 Identification of the pathway of alanine synthesis in the bacteroid ..............191

4.7 Isolation of mutants in alanine metabolism ..................................................193

4.7.1 Isolation of mutant RU1275 unable to grow on alanine ........................193

4.7.2 Isolation of multicopy aldA as a suppressor of strain RU1275 .............194

4.7.3 Construction of a mutation in L-alanine dehydrogenase (aldA)............194

4.8 Effect of mutation RU1327 on plant symbiotic performance ........................195

4.9 Discussion.....................................................................................................198

5 REGULATION OF L-ALANINE DEHYDROGENASE .................203

5.1 Biochemical analysis of L-alanine dehydrogenase activity ..........................205

5.2 Analysis of the sequence of aldA (L-alanine dehydrogenase) .....................207

5.3 Construction of a mutant in the putative regulator AldR...............................212

5.4 Growth of mutant in AldR..............................................................................215

5.5 Transcriptional analysis of aldA and aldR ....................................................216

5.6 Analysis of aldA mRNA levels ......................................................................222

5.7 Discussion.....................................................................................................224

6 GENERAL DISCUSSION.............................................................229

REFERENCES................................................................................236

XVI

LIST OF FIGURES

Fig 1.1 Diagram of indeterminate (A) and determinate (B) nodules highlighting the differences in morphology....................................................................................... 9

Fig 1.2 Ammonium assimilation pathways...........................................................16

Fig 1.3 Schematic representation of nutrient exchange between a nitrogen fixing bacteroid and the plant cell...................................................................................19

Fig 1.4 The TCA cycle and possible integration of pathways used in its regulation in the bacteroid .....................................................................................................30

Fig 1.5 Pathways leading to the synthesis and degradation of poly-β-hydroxybutyrate in rhizobia...................................................................................36

Fig 1.6 Pathway leading to the synthesis and degradation of glycogen in rhizobia…..............................................................................................................52

Fig 1.7 Organisation of glycogen biosynthetic genes in Rhizobium tropici..........53

Fig 1.8 Formation of L-alanine via alanine dehydrogenase.................................66

Fig 2.1 Experimental vessel for bacteroid excretion assays ..............................102

Fig 3.1 Sequence obtained from pRU99 subclones & pRU720 .........................111

Fig 3.2 Amino acid alignment of poly-β-hydroxybutyrate synthase....................115

Fig 3.3 Results of Testcode analysis .................................................................116

Fig 3.4 Southern hybridisation to confirm phaC was not present on pSym.......117

Fig 3.5 Vectors used in the construction of mutants in PHB synthase (RU1328 and RU1329).......................................................................................................119

Fig 3.6 Southern hybridisation to confirm PHB mutant indicated by increase of 2 Kb in phaC gene size..........................................................................................121

Fig 3.7 Growth of A34 and RU1328 on malate ..................................................123

Fig 3.8 Section of bean nodule showing PHB accumulation by bacteroids of A5……….............................................................................................................130

Fig 3.9 Section of bean nodule showing PHB accumulation by bacteroids of A5……….............................................................................................................131

Fig 3.10 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329 .............................................................................................................132

XVII

Fig 3.11 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329 .............................................................................................................133

Fig 3.12 Section of bean nodule showing PHB accumulation by bacteroids of A5……….............................................................................................................134

Fig 3.13 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329 .............................................................................................................135

Fig 3.14 Section of pea nodule showing characteristic Y-shape of bacteroids..136

Fig 3.15 Section of pea nodule showing no PHB accumulation by bacteroids of A34…….. ............................................................................................................137

Fig 3.16 Section of pea nodule. Note no PHB accumulation by bacteroids of A34, but PHB in bacteria inside infection thread.........................................................138

Fig 3.17 Section of pea nodule showing no PHB accumulation by bacteroids of RU1328 .............................................................................................................139

Fig 3.18 Section of pea nodule infected by A34 showing starch (S) accumulation at periphery of infected plant cells in interzone II-III ...........................................140

Fig 3.19 Light micrograph of a transverse section of a pea nodule infected by A34. Zones I, II, II-III, III, IV are indicated ...........................................................141

Fig 3.20 Light micrograph of a transverse section of a pea nodule infected by A34 showing zones I, II, and II-III .......................................................................142

Fig 3.21 Light micrograph of a transverse section of a pea nodule infected by RU1328. Zones I, II, III, IV are indicated ............................................................143

Fig 3.22 Light micrograph of a transverse section of a pea nodule infected by RU1328. Zones I and II are indicated.................................................................144

Fig 3.23 Peas grown in the greenhouse ............................................................146

Fig 3.24 Beans grown in the greenhouse ..........................................................147

Fig 3.25 Dry weight of (A) peas) and (B) beans)...............................................150

Fig 3.26 Representative pea plants from the second harvest at 53 days .........151

Fig 3.27 Southern hybridisation to confirm glgA is carried on cosmid pIJ9019. 157

Fig 3.28 Orientation of transposon TnB60 in glgA.............................................159

Fig 3.29 Southern blot and hybridisation to confirm mutations in glgA .............161

Fig 3.30 A Growth of A34 and RU1448 on glucose; B growth of A34 and RU1448 on succinate; and C growth of A34 and RU1448 on mannitol ..........................163

XVIII

Fig 3.31 Dry weight of peas inoculated with A34 and RU1448 .........................166

Fig 4.1 Results of a typical percoll gradient centrifugation.................................176

Fig 4.2 Alanine and ammonium concentration in bacteroid supernatant spiked with or without 1mM alanine ...............................................................................177

Fig 4.3 Effect of O2 concentration on the rate of ammonium excretion by bacteroids ...........................................................................................................179

Fig 4.4 Production of ammonium by bacteroids under a range of malate concentrations.....................................................................................................180

Fig 4.5 A representative bacteroid excretion assay showing both ammonium and alanine were produced at linear rates by high density bacteroids .....................184

Fig 4.6 Production of alanine from low density bacteroids bacteroids supplemented with (A) no malate; (B) 1mM malate; and (C) 10mM malate and ammonium ..........................................................................................................188

Fig 4.7 Alanine production by bacteroids isolated in air incubated with 10mM NH4Cl or 2mM malate or both.............................................................................190

Fig 4.8 Alanine excretion from a low density preparation of bacteroids incubated for 1 hr at various ammonium concentrations with malate at 2mM....................192

Fig 4.9 Alanine produced by bacteroids of RU1327 and 3841 over 2 hours when supplemented with 10mM NH4Cl and 2mM or 10mM malate ............................195

Fig 4.10 Average dry weight (g) of peas inoculated with RU1327 and 3841 harvested at 6 weeks..........................................................................................196

Fig 5.1 Translation of aldR showing helix-turn-helix motif located in the N-terminus of the protein in red ..............................................................................207

Fig 5.2 Sequence obtained from pR3135..........................................................209

Fig 5.3 Amino acid alignment of L-alanine dehydrogenase...............................211

Fig 5.4 Vectors used in the construction of RU1422 .........................................213

Fig 5.5 Southern hybridisation to confirm AldR mutant indicated by increase of 2 Kb in aldR gene size...........................................................................................214

Fig 5.6 Plasmids pRU731 (aldA GUS fusion) and pRU730 (aldR GUS fusion. 217

Fig 5.7 Representative northern analysis of aldA mRNA levels from 3841 grown on alanine (A), alanine/glucose (A/G), succinate (S), and glucose (G) .............222

XIX

LIST OF TABLES

Table 1.1 Examples of Rhizobium-plant associations............................................ 4

Table 2.3 List of antibiotic concentrations ............................................................73

Table 2.4 List of strains.........................................................................................74

Table 2.5 List of plasmids, cosmids and phage ...................................................76

Table 2.6 List of primers .......................................................................................79

Table 3.1 Growth of PHB mutants and parental strains on different carbon sources ..............................................................................................................122

Table 3.2 PHB content of PHB mutants and parental strains grown on 10mM fructose / 5mM NH4Cl .........................................................................................125

Table 3.3 Acetylene reduction (ethylene production) by wild type & PHB mutants…………………………………….. ..........................................................149

Table 3.4 Enzyme activity of A34 and RU1328 in bacteroids and in free living (FL) cells grown on malate .........................................................................................153

Table 3.5 Glycogen content of pea bacteroids...................................................154

Table 3.6 Growth of glycogen mutant and wild type on different carbon sources…………….. ...........................................................................................162

Table 3.7 Glycogen content of 10mM sucrose/5mM NH4-grown R. leguminosarum strains RU1448 & A34 ..............................................................164

Table 3.8 Acetylene reduction (ethylene production) by A34 & RU1448...........165

Table 5.1 L-Alanine dehydrogenase activity of 3841, RU1327, RU1371 and RU1422 grown on various carbon sources ........................................................206

Table 5.2 Growth of 3841 and RU1422 on different carbon sources on agar plates…...............................................................................................................215

Table 5.3 GUS activity of RU1414, RU1415 and RU1416 grown on various C/N sources………….................................................................................................219

Table 5.4 GUS activity vs. L-alanine dehydrogenase activity in cultures grown on glucose, succinate, and alanine/glucose ............................................................221

CHAPTER 1

2

1 INTRODUCTION

Rhizobia are bacteria that form mutualistic associations with members of the

plant family Leguminosae. The bacteria reduce (fix) molecular nitrogen to

ammonia which is made available to the plant in exchange for a carbon and

energy source derived from photosynthate. The symbiotic interaction takes place

in a specialised outgrowth of the plant root (or occasionally shoot) termed the

nodule. For the formation and functioning of the nodule both organisms undergo

a complex series of developmental steps resulting in changes to fundamental

cellular processes such as cell division and metabolism. For these reasons and

because the association is both ecologically and agronomically significant,

contributing the largest single organic nitrogen input to the global nitrogen cycle,

the Rhizobium-legume symbiosis attracts a wealth of research.

The work presented in this thesis primarily considers carbon metabolism by the

bacterial partner and its impact on the efficiency of nitrogen fixation. To

understand the complexities of the nodule environment and how that might effect

the carbon and nitrogen metabolism of the microsymbiont an overview of the

rhizobia and the symbiosis is presented followed by a review of literature based

on bacteroid carbon and nitrogen metabolism that is pertinent to this thesis.

3

1.1 Rhizobium taxonomy

Traditionally, the taxonomy of rhizobia has been based on the selective

interaction with plant hosts and the rate of growth of isolates on laboratory media

(for example ‘slow growers’ versus ‘fast growers’). More recently, the taxonomic

classification has been in a state of flux due to the wealth of molecular

information such as analysis of 16S ribosomal RNA, DNA restriction fragment

length polymorphisms (RFLP), and multilocus enzyme electrophoresis (MLEE).

This modern molecular data has indicated that the rhizobia are polyphyletic and

has lead to the division of the rhizobia into the genera: Azorhizobium,

Bradyrhizobium, Mesorhizobium, Sinorhizobium and Rhizobium. These divisions

have frequently reinforced previous divisions, for example the classical ‘slow

growers’ now form the genus Bradyrhizobium. Within the genus Rhizobium

several strains nodulate a common host, but are distinct according to genetic and

phenotypic properties and are therefore classified as distinct species (eg. R.

tropici and R. etli). Strains of Rhizobium leguminosarum cannot be distinguished

other than their host range, therefore the species is further classified into three

biovars (bv.) that nodulate clover, peas and beans. The nitrogen fixation and

nodulation genes, including the host range determinants, are present on large

symbiotic plasmids (Sym plasmids), therefore the biovar classification is

essentially a description of the plasmid rather than the chromosomal background

of the strain (MartinezRomero & CaballeroMellado, 1996; Young, 1996). When a

Sym plasmid is transferred to another strain cured of its symbiotic plasmid, the

recipient strain acquires nodulation characteristics of the donor strain. In this way,

Allan Downie (pers. comm.) has produced strains of R. leguminosarum that are

4

able to nodulate both pea and bean (see chapter 3). In the ‘slow-growing‘

Bradyrhizobium strains the symbiosis genes are chromosomal. Many of the

Rhizobium – plant associations referred to throughout the thesis are given in

Table 1.1.

Table 1.1 Examples of Rhizobium-plant associations.

Bacterial species Legume host

Azorhizobium caulinodans Sesbania (Sesbania rostrata)†

Bradyrhizobium japonicum Soybean (Glycine max)

Bradyrhizobium sp. Parasponia*

Mesorhizobium ciceri Chickpea (Cicer arietinum)

Mesorhizobium loti Trefoil (Lotus sp.), Lupine (Lupinus sp.), Seradella (Ornithopus sativus)

Rhizobium sp. NGR234 Broad host range. Genera including Vigna, Glycine, Mimosa, Lotus.

Rhizobium etli Common beans (Phaseolus vulgaris)

Rhizobium leguminosarum

biovar viciae Peas (Pisum sp.), vetches (Vicia sp.)

biovar trifolii Clovers (Trifolium sp.)

biovar phaseoli

Common beans (Phaseolus vulgaris)

Rhizobium tropici Common beans (Phaseolus vulgaris)

Sinorhizobium meliloti Alfalfa (Medicago sp.)

†forms root and stem nodules * the only known non-legume nodulated

5

1.2 Nodule formation, structure and environment

1.2.1 Molecular basis for nodulation

Plant roots secrete large quantities of organic compounds into the soil. These

compounds include carbohydrates, amino acids, organic acids, vitamins and

phenolic derivatives, most of which supports growth of microorganisms in the

rhizosphere. Of these compounds, flavanoids (also betaines and erythronic and

tetronic acids) are important from the symbiotic perspective because they trigger

the induction of bacterial nodulation (nod) genes that encode proteins involved in

the production and transport of Nod factors (Perret et al., 2000). Nod factors,

which are modified lipo-chitooligosaccharides, act as bacterial signal molecules

initiating nodule organogenesis (Denarie et al., 1996; Long, 1996). Nodulation

specificity is determined by chemical modification of the nod factors. Nod factor

structure is controlled by the protein products of the nodulation genes nod, nol

and noe. The proteins NodA (acyl-transferase), NodB (deacetylase) and NodC

(N-acetylglucosaminyltransferase or chitin synthase) together function to catalyze

the synthesis of the monoacylated tetrameric or pentameric chitin core structure

(Spaink, 1996). Enzymes encoded by host specific nodulation genes modify this

core molecule and impose specificity (Denarie et al., 1996; Long, 1996; Spaink,

1996).

Nod gene expression is regulated via NodD. The nodD gene is expressed

constitutively but there is little or no expression of other nod genes unless the

bacteria are exposed to plant root exudates. The NodD proteins act as sensors to

plant signals in root exudates and bind to a highly conserved 49 bp DNA motif

(nod box) in the promoter region of nodulation loci, thus activating transcription of

6

the nod loci. Rhizobia possess one to many copies of NodD that may associate

with several flavanoids (Broughton et al., 2000).

1.2.2 Events of nodulation

Nod factors act with high specificity at very low concentrations, suggesting that

Nod factor perception is probably mediated by specific plant receptors. Nod

factors stimulate the expression of corresponding nodulation genes in the plant

(nodulins) which cause the root hairs to curl around the bacteria and stimulate

cortical cell division. Root hair curling facilitates infection by preventing the

enzymes involved in root hair penetration from diffusing away. The root hair cell

wall is dissolved away at a localised site and the bacteria enter the root hair via

this invagination. On entry a new plant cell wall is formed which encloses the

bacteria and forms an infection thread. The bacteria multiply within the infection

thread which extends to beyond the base of the root hair, then pass through the

cell wall of the adjacent cell, and down through several layers of cortical cells.

When the infection thread has penetrated the cortex the inner cortical cells are

stimulated to divide to form the nodule primordia. Bacteria are released from the

infection thread into newly divided plant cells. The nodule is formed by the

subsequent infection and expansion of plant cells. At this stage the infection

process can lead to one of two types of nodule depending on the host plant:

determinate as formed on beans or indeterminate nodules as formed on peas.

1.2.2.1 Determinate nodules

The infection thread does not continue to extend, or extends little, through the

cortex. Continued infection of cells arises from division of pre-infected plant cells,

and as a result bacteria are distributed evenly through the central mass of the

7

nodule. Increase in nodule size occurs through cell enlargement hence nodules

are more or less spherical in shape and are limited in size (Fig 1.1).

1.2.2.2 Indeterminate nodules

Infected plant cells do not divide further and the bacteria become distributed

throughout the nodule by the infection thread. The infection thread becomes

elongated and branched in order to infect cells behind the dividing cells which

form a meristematic zone at the apex of the nodule. Thus the nodule increases in

size due to the divisions of meristematic cells and the continued infection of new

plant cells by the infection thread. This results in nodules that are generally

elongated but the meristem may also divide giving rise to branched nodules.

Five distinct zones of development can be defined along a longitudinal section

through an indeterminate nodule (Fig 1.1): the bacteria free meristematic zone I;

the infection zone II, in which cell differentiation begins; the interzone II-III,

characterised by a histologically distinct band of 1 - 3 layers of host cells

containing significant deposits of starch; nitrogen fixing zone III; and the

senescent zone IV (Vasse et al., 1990). In interzone II-III bacteria undergo major

differentiation and the genes required for the production of nitrogenase are

transcribed. Recently, a sixth zone of development proximal to the senescent

zone IV (zone V) that is present in nodules of alfalfa more than 6 weeks old has

been defined (Timmers et al., 2000). This zone is characterised by the re-

invasion of plant cells that have completely senesced. The bacteria resemble

intracellular saprophytic bacteria, and do not differentiate into bacteroids and are

not bound by plant membrane (Timmers et al., 2000). It has been hypothesised

that this zone represents a pool of viable bacteria that will contribute to soil

8

bacterial populations in the vicinity of the nodulated legume after nodule

breakdown (Timmers et al., 2000).

9

Fig 1.1 Diagram of indeterminate (A) and determinate (B) nodules highlighting the differences in morphology. See text for explanation.

ZONE I

II

II – III

III

IV

V

Expansion of nodule forwards

Expansion of nodule outwards

Infected cells

Gradient of cell size

A

B

10

1.2.3 Formation of the symbiosome

Infection threads are surrounded by a membrane which is continuous with the

plasma membrane of the infected plant cell. Bacteria are taken up into the host

cell at an un-walled area of the infection thread by endocytosis of the infection

thread membrane. After endocytosis the bacteria divide. Bacterial division may

be accompanied by division of the plant membrane (the peribacteroid membrane

or PBM), resulting in bacteria that are singly enclosed by PBM, such as pea. In

other associations such as bean, the PBM does not divide or does not divide as

frequently as the bacteria, resulting in several bacteria being enclosed by the

same PBM. When the bacteria stop replicating they differentiate into their

symbiotic form known as bacteroids. Differentiation results in the formation of

cells that produce nitrogenase and a range of other accessory proteins that

provide a suitable environment for nitrogen fixation to occur. The symbiotic unit

thus formed, composed of the peribacteroid membrane, the bacteroids, and the

intervening peribacteroid space (PBS), is termed the symbiosome or

peribacteroid unit (PBU) (Whitehead & Day, 1997). The consequence of this

arrangement is that exchange of metabolites between bacteroid and host must

take occur across both peribacteroid membrane and bacteroid membrane (BM).

1.2.4 The microaerobic nodule environment

A low oxygen concentration is essential for symbiotic nitrogen fixation because

nitrogenase is irreversibly inhibited by oxygen. Both determinate and

indeterminate nodules have a layer of uninfected cells that surrounds the inner

core of infected cells. These cells serve to restrict the flux of oxygen into the

infected zone of the nodule by acting as a diffusion barrier (Minchin et al., 1985;

11

Sheehy et al., 1985; Delorenzo et al., 1993; Iannetta et al., 1993; Sheehy et al.,

1983). This, together with high rates of metabolism by bacteroids and

mitochondria inside the infected cells, imposes microaerobic conditions within the

nodule (oxygen concentrations between 3 and 22nM (Witty & Minchin, 1990;

Hunt & Layzell, 1993)). High rates of bacteroid metabolism and therefore efficient

production of ATP and reductant for nitrogenase can occur under these

conditions due to specialisations in the nodule environment. These include: (1)

Production of leghemoglobin, a plant derived oxygen binding protein, in the

cytoplasm of infected cells that facilitates oxygen transport to bacteroids

(Appleby, 1969). (2) Utilisation of a unique high oxygen affinity cytochrome

system by bacteroids that ensures ATP production can occur at oxygen

concentrations of under 10nM (Bergersen & Turner, 1975). (3) Modifications to

aerobic carbon metabolic pathways (reviewed in section 1.4 and McDermott et

al., 1989).

12

1.3 Exchange of metabolites in the nodule

1.3.1 Carbon provision to the bacteroid

Carbon from the plant is supplied to the bacteroid to fuel the process of nitrogen

fixation (see Fig 1.3 for diagram of nutrient exchange in nodule). Photosynthate is

transported to the nodule in the form of sucrose via the phloem (Streeter, 1981;

Reibach & Streeter, 1983; Gordon et al., 1985; Kouchi & Yoneyama, 1986;

Streeter, 1987). In the nodule tissue soluble sugars are abundant but do not

directly support nitrogen fixation. Several lines of evidence support this: (1)

Isolated bacteroids of Bradyrhiobium japonicum are not stimulated to fix nitrogen

by sugars (Bergersen & Turner, 1967). (2) The uptake of neutral sugars by

isolated soybean symbiosomes has been shown to occur by diffusion and the

uptake rates are not adequate to support nitrogenase activity (Day & Udvardi,

1989; Udvardi et al., 1990). (3) There are low levels of glycolytic enzymes in

bacteroids (Reibach & Streeter, 1983; Salminen & Streeter, 1987; Copeland et

al., 1989; Copeland et al., 1995). (4) With one exception (Duncan, 1981), mutants

of Rhizobium that are defective in the catabolism of sugars form effective nodules

in all symbioses studied (Arias et al., 1979; Ronson & Primrose, 1979;

Cervenansky & Arias, 1984; Glenn et al., 1984; Glenn et al., 1984; Arwas et al.,

1986; Dilworth et al., 1986; El-Guezzar et al., 1988; Lafontaine et al., 1989).

In the nodule tissue sucrose is cleaved by sucrose synthase (to UDP-glucose

and fructose) or alkaline invertase (to glucose and fructose). Sucrose synthase

(SS) activity is nodule enhanced (Thummler & Verma, 1987) and it is required for

nitrogen fixation (Craig et al., 1999; Gordon et al., 1999) suggesting that it is the

main enzyme of sucrose cleavage in the nodule. Following cleavage by SS, the

13

hydrolysed products are used as substrates for cellulose or starch biosynthesis

and/or are metabolised by glycolytic enzymes to produce phosphoenolpyruvate

(PEP). Enzymes of the glycolytic pathway have been shown to be high in plant

cytosol (Reibach & Streeter, 1983; Salminen & Streeter, 1987; Copeland et al.,

1989; Copeland et al., 1995; Kaur & Singh, 1999). PEP is further metabolised to

produce malate via phosphoenolpyruvate carboxylase (PEPC) and malate

dehydrogenase (MDH) (Day & Copeland, 1991). The concentration of malate is

high in nodules (estimated to be 3.4mM (DeVries et al., 1980)) as is the

concentration of other dicarboxylates (DeVries et al., 1980; Kouchi & Yoneyama,

1986; Streeter, 1987; Rosendahl et al., 1990).

Dicarboxylates have been shown to stimulate bacteroid nitrogen fixation in vitro

(Bergersen & Turner, 1967) and labelling data on whole nodule tissue are

consistent with dicarboxylates being the carbon source for the bacteroid in planta.

Malate, fumarate and succinate are rapidly labelled in plant cytosol when nodules

are exposed to 14CO2 (Kouchi & Nakaji, 1985; Vance et al., 1985; Kouchi &

Yoneyama, 1986; Rosendahl et al., 1990) and this label is rapidly incorporated

into bacteroids primarily as malate (Kouchi & Yoneyama, 1986; Rosendahl et al.,

1990; Salminen & Streeter, 1992). Furthermore, isolated bacteroids transport

dicarboxylates (Glenn et al., 1980) and mutants in C4-dicarboxylate transport

(Dct) form ineffective nodules on pea, clover and alfalfa (Glenn & Brewin, 1981;

Ronson et al., 1981; Finan et al., 1983; Glenn et al., 1984; Arwas et al., 1985;

Arwas et al., 1986; Engelke et al., 1987; Watson et al., 1988; Engelke et al.,

1989). Bacteroids formed by Dct mutants tend to develop as the wild type strain

but senesce shortly after release from the infection thread. Therefore in

indeterminate nodules there is a large senescent zone, and infected plant cells

14

contain higher levels of starch (Ronson et al., 1981; Finan et al., 1983; Arwas et

al., 1985; Watson et al., 1988). In determinate nodules, such as soybean, mutant

bacteroids contain fewer granules of poly-β-hydroxybutyrate (PHB) and the

periphery of infected cells and the adjacent uninfected cells contain more starch

(Humbeck & Werner, 1989). This suggests that rhizobia use carbon sources

other than dicarboxylates during invasion of the nodule, but have an absolute

requirement for organic acids to fix nitrogen.

Since transport and catabolism of dicarboxylic acids are required to fuel nitrogen

fixation by rhizobia in legume nodules a great deal of attention has been focused

on the dicarboxylic acid transport system. The Dct system consists of three

genes: dctA, which codes for the transport protein and two divergently

transcribed genes, dctB and dctD, which activate transcription of dctA in

response to the presence of dicarboxylates (Ronson et al., 1984; Watson et al.,

1988; Engelke et al., 1989; Yarosh et al., 1989; Watson, 1990; Jording et al.,

1992; Jording et al., 1994). DctA has 12 membrane spanning helices typical of

membrane transport proteins, with the N-terminus and C-terminus located in the

cytoplasm (Jording & Pühler, 1993). DctB and DctD are a two component sensor-

regulator pair (Ronson et al., 1987; Watson, 1990). DctB is a sensor protein that

responds to the presence of dicarboxylates and phosphorylates the

transcriptional activator DctD (Giblin et al., 1995). Upon phosphorylation DctD

binds to the dctA promoter and initiates transcription (Giblin et al., 1995).

The nature of the dicarboxylate transporter on the peribacteroid membrane is

unknown.

15

1.3.2 Nitrogen provision to the plant

Bacteroids use the enzyme nitrogenase to catalyse the reduction of dinitrogen to

ammonium in a reaction that requires a significant amount of reductant and

energy:

That ammonium is the primary stable product of symbiotic nitrogen fixation was

first concluded in the 1960’s. Bergersen (1965) showed that more than 90% of

the soluble fixed nitrogen in nodule tissue was ammonia in detached soybean

nodules incubated for 1 minute with 15N2 gas (Bergersen, 1965). Bergersen

further showed that the vast majority of label recovered from isolated soybean

bacteroids exposed to 15N2 gas was found as ammonium in the medium in which

bacteroids were suspended (Bergersen & Turner, 1967), suggesting that all fixed

nitrogen was transferred from the bacteroid as ammonia. However, recent papers

by Waters et al., (1998) and Allaway et al., (2000) have challenged the sole

excretion of ammonium from the bacteroid. This is addressed in section 1.5.2.

Kennedy showed that glutamate and glutamine were the primary amino

compounds synthesised from the ammonia in the plant cytosol (Kennedy, 1966a;

Kennedy, 1966b). Ammonia is converted to glutamate directly via glutamate

dehydrogenase (GDH), or by the coupled activities of glutamine synthetase

(which synthesises glutamine) and glutamate synthase (GS-GOGAT) (Fig 1.2

and Fig 1.3). In the plant cytosol there are high levels of GS-GOGAT. Over 90%

of the total nodule glutamine synthetase (GS) is in the plant tissue, where it can

account for as much as 2% of total soluble protein and its activity is induced over

N2ase N2 + 8H+ + 8e- + 16ATP 2NH4

+ + H2 + 16ADP + 16Pi

16

that in uninnoculated roots (Cullimore & Bennett, 1988). Nodule glutamate

synthase (GOGAT) activity is increased over that in other plant organs (Chen &

Cullimore, 1988). The high levels of nodule GS and GOGAT are achieved by the

nodule specific induction of a single enzyme or by the induction of nodule specific

iso-enzymes (Chen & Cullimore, 1988; Cullimore & Bennett, 1988; Bennett &

Cullimore, 1989; Bennett et al., 1989; Gregerson et al., 1993; Roche et al., 1993).

Fig 1.2 Ammonium assimilation pathways. Glutamine synthetase (GS) and glutamate synthase (GOGAT); glutamate dehydrogenase (GDH).

As ammonium assimilation primarily occurs in the plant cytosol, ammonium must

cross the symbiotic membranes (Fig 1.3). The extent to which ammonium

traverses the bacteroid membrane and peribacteroid membrane by diffusion or

active transport has not been clearly established. A significant proportion of fixed

nitrogen is believed to be transferred from the bacteroid cytoplasm to the plant

cytoplasm by diffusion of ammonia/ammonium across the membranes. There is a

50-fold lower concentration of ammonium in the plant cytoplasm compared to the

NH4+ + L-glutamate + ATP L-glutamine + ADP + Pi

L-glutamine + α-ketoglutarate + NADPH 2 L-glutamate + NADP+

NH4+ + α-ketoglutarate + NADPH L-glutamate + NADP+

GS

GOGAT

GDH

17

bacteroid due to the high activity of ammonia assimilating enzymes in the host

(Dilworth & Glenn, 1982; Streeter, 1989). This suggests that rapid movement to

host cytosol could be accomplished by diffusion and might be adequate to

support measured rates of ammonium assimilation.

The possible transport of ammonium across both the bacteroid and peribacteroid

membranes has also been investigated. An active transport system for

ammonium out of the bacteroid has not been identified on the bacteroid

membrane. Free-living rhizobia have a mechanism for the active transport of

ammonium ions, but in all cases studied, the carrier has been shown to be

inoperative in the symbiotic state (Ohara et al., 1985; Howitt et al., 1986; Jin et

al., 1988). This prevents bacteroids from competing with plant cells for the

ammonium that has diffused into the peribacteroid space from the bacteroid

(Udvardi & Day, 1997; Howitt & Udvardi, 2000).

Transport systems for ammonium have, however, been identified on the PBM.

The PBS is acidic (Udvardi et al., 1991) due to the presence of an H+ATPase on

the PBM (Blumwald et al., 1985). Therefore, as the proportion of NH3/NH4+ is

dependent on pH, a considerable proportion of the ammonia in the PBS is

protonated (NH4+) (Whitehead & Day, 1997). As the re-uptake of ammonium by

the bacteroid is prevented there is a steep NH3 concentration gradient from the

bacteroids to the PBS thus maintaining a high concentration of NH4+ in the PBS.

Recent patch clamp studies with isolated soybean symbiosomes have identified a

monovalent cation channel on the PBM that is capable of exporting this NH4+ out

of the PBS at rates that would maintain the high rates of nitrogen assimilation in

plant cytosol (Tyerman et al., 1995). The channel has little biochemical similarity

18

to other plant transporters permeable to NH4+ suggesting it is unique to the PBM

(Tyerman et al., 1995). A similar channel has also been identified on the PBM of

pea nodules using energised bacteroid side out symbiosome membrane vesicles

(Mouritzen & Rosendahl, 1997).

Fixed nitrogen is further converted from glutamate to other nitrogenous

compounds before translocation from the nodule via the xylem. The form of

nitrogen carried by the xylem depends on the legume species. Temperate

legumes (such as pea, clover and alfalfa which mainly form indeterminate

nodules) export amides (mainly asparagine) and tropical legumes (such as

soybean and Phaseolus bean) which mainly form determinate nodules export

ureides (allantoin and allantoic acid) (Schubert, 1986).

19

Fig 1.3 Schematic representation of nutrient exchange between a nitrogen fixing bacteroid and the plant cell.

TCA

SUCROSE

UDP-GLUCOSE

PEP

OXALOACETATE

MALATE

SS

GLYCOLYSIS

PEPC

MDH

N2ASE

N2

NH3

NH4+

H+ e- ATP BM PBM

ATP

ADP + Pi

NH4+

GLU

GS

GLN

GLU

GOGAT

α KG

ATP

ADP + Pi

NADP+

NADPH

AMINO ACIDS ? ?

ATP ADP + Pi

H+

20

1.4 Carbon metabolism in the bacteroid

1.4.1 The tricarboxylic acid (TCA) cycle

Our current knowledge of bacteroid metabolism is based on the presence or

absence of enzyme steps and the response of nitrogen fixation to mutations in

enzyme steps. We do not know the actual metabolite fluxes within wild type

bacteroids and therefore ideas are often speculative based on whether a

metabolite is available and how the bacteroid might respond. The rhizobia are

obligate aerobes, therefore dicarboxylates must be metabolised via the

tricarboxylic acid (TCA) cycle in the bacteroid. The operation of the TCA cycle

was confirmed in soybean bacteroids in the 70’s. Stovall and Cole (1978)

reported the activity of a full TCA cycle in B. japonicum bacteroids based on the

14CO2 evolution profiles from bacteroids incubated with 14C labelled succinate

and pyruvate (Stovall & Cole, 1978). Since then the activity of all TCA cycle

enzymes has been confirmed in Bradyrhizobium japonicum (Karr et al., 1984;

Waters et al., 1985) and demonstrated in bacteroids of Rhizobium

leguminosarum (McKay et al., 1989), Sinorhizobium meliloti (Miller et al., 1991),

Rhizobium tropici (Romanov et al., 1994) and Mesorhizobium ciceri CC 1192

(Kim & Copeland, 1996). Although bacteroids have all the enzymes of the TCA

cycle it has been suggested the cycle does not operate to its full aerobic capacity

in bacteroids due to inhibition of enzyme activity by reduced nucleotides

(McDermott et al., 1989; Day & Copeland, 1991). High levels of NADH

accumulate in the bacteroid because the microaerobic conditions in nodules limit

the ability of the respiratory chain to oxidise reduced nucleotides. Citrate

synthase activity in M. ciceri CC 1192 and both malate dehydrogenase and

21

isocitrate dehydrogenase activity in B. japonicum are strongly inhibited by NADH

in vitro, indicating that during symbiotic nitrogen fixation the enzyme activities

would be markedly reduced in planta (Karr & Emerich, 2000; Tabrett & Copeland,

2000). α-Ketoglutarate dehydrogenase (KGDH) activity is strongly inhibited by

reduced nucleotides, with the degree of inhibition dependent on the ratio of

NADH to NAD+. Salminen and Streeter (1990) estimated the B. japonicum

bacteroid NADH:NAD+ ratio to be 0.83, which results in more than 50% inhibition

of KGDH in vitro (Salminen & Streeter, 1990). The impact of this enzyme

inhibition on carbon metabolism in vivo is that the bacteroid must employ specific

pathways to (1) by-pass certain steps in the TCA cycle; (2) maintain a balanced

input and removal of carbon from the TCA cycle; and (3) balance the redox

potential of the cell. It has been proposed that these pathways include the

formation of carbon storage compounds and of amino acids (Fig 1.4) (McDermott

et al., 1989).

1.4.2 Mutation analysis of TCA cycle enzymes

Mutational analysis has provided information on the steps in the TCA cycle that

are essential in bacteroids and have implied the existence of alternative

pathways which by-pass enzyme steps that are presumed to be blocked in vivo

by high levels of reductant inhibiting enzyme activity (eg. KGDH). However, even

though studies of this type have proven useful in certain instances, more often

than not they highlight the complexity of the symbiotic interaction and how limited

our knowledge of bacteroid metabolism actually is.

Succinate dehydrogenase (SDH) catalyses the dehydrogenation of succinate to

fumarate (Fig 1.4). S. meliloti and R. leguminosarum mutants deficient in SDH

22

are not able to grow on succinate, but can on malate and fumarate (Finan et al.,

1981; Gardiol et al., 1982; Gardiol et al., 1987). The SDH mutants of both R.

leguminosarum and S. meliloti form ineffective nodules on their respective plant

hosts. Ineffective SDH- bacteroids isolated from peas are able to transport

succinate (Finan et al., 1981) indicating that the metabolism of succinate via

succinate dehydrogenase is required for an effective pea symbiosis. Given this

data one would expect that other enzymes (fumarase and malate

dehydrogenase) that catalyse key reactions in the initial breakdown

dicarboxylates coming from the plant might have similar symbiotic phenotypes.

However the case for fumarase and malate dehydrogenase is not as simple.

Fumarase (FUM) cataylses the hydration of fumarate to malate (Fig 1.4). A

deletion mutant in fumC of B. japonicum retains a high level of fumarase activity

(60% of the wild type) and is able to fix nitrogen in symbiosis with soybean,

strongly suggesting the presence of a second gene encoding fumarase in B.

japonicum (Acuna et al., 1991). There are two genes encoding fumarase in the

genome sequence of M. loti (Kaneko et al., 2000) but there is only one present in

S. meliloti (http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001).

Therefore few conclusions on the bacteroids requirement for fumarase can be

drawn from this study.

Malate dehydrogenase (MDH) catalyses the conversion of malate to oxaloacetate

(Fig 1.4). Mutants in malate dehydrogenase have not been isolated as yet. The

gene for malate dehydrogenase is part of an operon in R. leguminosarum, mdh-

sucCDAB, which includes the α-ketoglutarate dehydrogenase complex and

succinyl-CoA synthetase (Walshaw et al., 1997). Mdh codes for malate

dehydrogenase, sucCD code respectively for the β and α subunits of succinyl-

23

CoA synthetase, while sucAB code for the α-ketoglutarate dehydrogenase

component (E1) and the dihydrolipoamide succinyltransferase component (E2) of

the α-ketoglutarate dehydrogenase complex. Mutations in any upstream gene of

the mdh-suc operon prevents expression of downstream genes, while mutation of

a downstream gene increases transcription and thus total enzyme activity of the

products of upstream genes (Walshaw et al., 1997). Mutation of sucD increases

the transcription of the upstream mdh seven-fold (Poole et al., 1999). The mdh

promoter is the principal one for the whole operon. One consequence of this

transcriptional coupling is that it is not possible to transcriptionally split the two

sides of the TCA cycle as happens in E. coli : Mdh and the α-ketoglutarate

dehydrogenase complex will always be expressed together. Attempts to isolate a

malate dehydrogenase mutant in R. leguminosarum bv. viciae by transposon

mutagenesis have not been successful either because this enzyme is essential or

is highly polar on downstream genes of the mdh-suc operon (Poole et al., 1999).

α-Ketoglutarate dehydrogenase (KGDH) cataylses the oxidative decarboxylation

of α-ketoglutarate to succinyl-CoA, which is further hydrolysed to succinate by

succinyl-CoA synthetase (SCS) (Fig 1.4). Mutants have been isolated in α-

ketoglutarate dehydrogenase and succinyl-CoA synthetase that form ineffective

nodules on alfalfa (Duncan & Fraenkel, 1979) and pea (Walshaw et al., 1997)

suggesting that these steps of the TCA cycle are required for bacteroid

metabolism in these symbioses. However, an α-ketoglutarate dehydrogenase

(sucA) mutant of B. japonicum formed effective nodules on soybean (Green &

Emerich, 1997). Both nodulation and nitrogen fixation were severely delayed in

the B. japonicum mutant, but isolated effective bacteroids recovered from

24

nodules behaved as wild type with respect to nitrogen fixation (Green et al.,

2000). The mutant in sucA allowed growth on malate and succinate (Green &

Emerich, 1997) suggesting that B. japonicum does not require the normal

complete TCA cycle to generate sufficient energy from these substrates and

indicating the functioning of an alternative pathway. Potential mechanisms for by-

passing α-ketoglutarate dehydrogenase include metabolism via the glyoxylate

cycle or α-ketoglutarate decarboxylase. The glyoxylate cycle consisting of

isocitrate lyase (ICL) and malate synthase (MS) (Fig 1.4), is an inducible pathway

required for the growth of many bacteria including free-living rhizobia on two-

carbon substrates such as acetate (Mandal & Chakrabartty, 1992; Cronan &

LaPorte, 1996; Mandal & Chakrabartty, 1999). The cycle bypasses the

decarboxylating reactions of the TCA cycle, providing a net production of

succinate from two molecules of acetyl-CoA. B. japonicum bacteroids showed

very little isocitrate lyase activity, implying that a complete glyoxylate cycle is not

functional during symbiosis (Green et al., 1998). However exogenously supplied

glutamate and α-ketoglutarate are converted to succinic semialdehyde in the B.

japonicum sucA mutant (Green et al., 2000). The activity of α-ketoglutarate

decarboxylase is linear in both free-living cells and bacteroids, indicating the

alternative pathway by-passing α-ketoglutarate dehydrogenase in B. japonicum is

via α-ketoglutarate decarboxylase (KDC) and succinic semialdehyde

dehydrogenase (SSDH) to form succinate (Fig 1.4) (Green et al., 2000).

The first step in the TCA cycle is the condensation of the acetyl group of acetyl-

CoA with oxaloacetate to form citrate. This step is catalyses by citrate synthase

(CS) (Fig 1.4). R. tropici has two citrate synthase genes, one on the symbiotic

25

plasmid (pcsA) and one on the chromosome (ccsA). A mutant in ccsA has a

diminished citrate synthase activity and nodulation capacity, but fixes nitrogen. A

citrate synthase double mutant (ccsA-pcsA-) forms ineffective nodules (Pardo et

al., 1994). A clone carrying the pcsA gene confers a higher nodulation capacity in

correlation with a higher citrate synthase activity in R. leguminosarum bv.

phaseoli (which naturally lacks a plasmid citrate synthase gene) (Pardo et al.,

1994). Furthermore, mutation of citrate synthase (gltA) in S. meliloti alters the cell

surface polysaccharides and produces nodules on alfalfa that are devoid of

bacteroids (Mortimer et al., 1999). Therefore the symbiotic defects shown by

citrate synthase mutants appear to result from disruption of the infection process

rather than bacteroid carbon metabolism. It is not known why mutations in CS are

detrimental to nodulation, but perhaps this highlights the complexity of the

symbiotic interaction and / or our limited knowledge of the microsymbionts

metabolism.

Isocitrate dehydrogenase (ICDH) catalyses the oxidation of isocitrate to α-

ketoglutarate (Fig 1.4). Mutation of icd in S. meliloti results in no ICDH activity

suggesting just one gene encodes ICDH. The icd mutant forms nodules which

are ineffective (McDermott & Kahn, 1992) suggesting this enzyme is required for

bacteroid metabolism. Pseudorevertants, that grow faster than the mutant, have

been isolated which have no CS activity due to a spontaneous secondary

mutation. Nodule formation by the ICDH/CS double mutant (A39L) is severely

affected and plants only form small callus structures devoid of bacteroids

(McDermott & Kahn, 1992) again indicating a role for citrate synthase during

nodulation. However, the secondary mutation has not been mapped to a

particular gene and attempts to reintroduce the wild type gltA gene encoding

26

citrate synthase into A39L were unsuccessful suggesting the unmapped mutation

is incompatible with a functional citrate synthase (Mortimer et al., 1999).

Therefore the secondary mutation in A39L is likely to be more complex than a

single mutation in the gene encoding for citrate synthase.

The symbiotic role of aconitase (ACN), which cataylses the reversible

isomerisation of citrate and isocitrate (Fig 1.4), has not been clearly established.

A mutation in B. japonicum acnA encoding aconitase decreases activity of

aconitase in free-living cells by more than 70%. The ability of the mutant to

establish an effective root nodule symbiosis with soybean plants is not affected.

This may be due to the presence of a second aconitase but unfortunately this

was not assayed in bacteroid extracts (ThonyMeyer & Kunzler, 1996). There is

only one aconitase in both the S. meliloti

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001) and M. loti

(Kaneko et al., 2000) genomes.

1.4.3 Formation of acetyl-CoA: the role of malic enzymes

Metabolism of C4-dicarboxylates as sole carbon source via the TCA cycle is

dependent on the availability of acetyl-CoA. There must be a molecule of acetyl-

CoA for each molecule of oxaloacetate made available for condensation by

citrate synthase to allow the formation of citrate and hence proper functioning of

the TCA cycle.

McKay (1988) first reported a high level of malic enzyme activity (210-240 nmol

min-1mg protein-1) in bacteroids of R. leguminosarum suggesting this enzyme is

significant in bacteroids (McKay et al., 1988). Malic enzyme (ME) decarboxylates

27

malate to pyruvate which can be further decarboxylated to acetyl-CoA via

pyruvate dehydrogenase (Fig 1.4). The activity of two malic enzymes, a NAD+

dependent malic enzyme (DME) and a NADP+ dependent malic enzyme (TME),

has been demonstrated in B. japonicum (Kouchi et al., 1988; Copeland et al.,

1989; Tomaszewska & Werner, 1995), R. leguminosarum (McKay et al., 1988),

and R. meliloti (Driscoll & Finan, 1993) bacteroids. The activity and regulation of

the two malic enzymes has been most extensively studied in S. meliloti and B.

japonicum.

It has been suggested that the NAD+ linked ME has a special role in symbiotic

nitrogen fixation. In B. japonicum bacteroids the affinity of the NAD+ dependant

ME has a lower affinity for malate than the NADP+ enzyme (Km of 1.9 mM malate

compared to 0.1 mM in crude extracts of B. japonicum bacteroids (Copeland et

al., 1989)). This suggests that DME functions in the decarboxylation of malate

only when there was ample substrate from the plant (ie. during periods of active

nitrogen fixation). Furthermore expression of DME is stimulated by C4-

dicarboxylates and by ammonium ions (Copeland et al., 1989) which would

further ensure maximum activity during nitrogen fixation. Affirmation of the

importance of DME comes from studies using mutants in the malic enzymes of S.

meliloti. A mutant in DME (dme-) that retains TME activity is unable to fix nitrogen

on alfalfa plants whereas a mutation in TME (tme-) has no adverse effect on the

symbiosis (Driscoll & Finan, 1996). Furthermore tme placed under control of the

dme promoter in a dme- background does not restore the symbiotic phenotype of

the dme- strain. Therefore, DME is required for symbiosis and TME is not able to

functionally replace DME during symbiosis. Acetyl-CoA inhibits DME activity in

vitro suggesting that this metabolite is the end product of a DME/PDH pathway in

28

vivo (Driscoll & Finan, 1997). The gene (pdhA) encoding PDH has been reported

to be induced during the symbiotic stage of S. meliloti (Cabanes et al., 2000 a &

b). Mutation in a gene coding for a putative arylesterase (ada) which is clustered

with the genes for pyruvate dehydrogenase (pdhAβ), dihydrolipoamide

acetyltransferase (pdhB) and lipoamide dehydrogenase (lpd) results in a Fix-

phenotype in S. meliloti (Soto et al., 2001). This mutation results in a 16-fold

reduction of pyruvate dehydrogenase activity, consistent with an essential role for

pyruvate dehydrogenase in nitrogen fixation. Together these results suggest the

NAD+ dependent ME (DME), together with pyruvate dehydrogenase (PDH), form

the pathway for conversion of malate to acetyl-CoA in bacteroids.

Purified DME has a Km of 9.4mM for malate, and Km‘s of 89µM and 1.56mM for

the NAD+ and NADP+ cofactors respectively. Whereas TME has a Km of 2.6mM

for malate and 33µM for the NADP+ cofactor (Voegele et al., 1999). DME affinity

for malate is also stimulated by both succinate and fumarate and inhibited by

acetyl-CoA and oxaloacetate (Voegele et al., 1999). TME shows no such

regulation (Mitsch et al., 1999). It has been suggested from this data that DME

detects increases in carbon flow to the TCA cycle via succinate and fumarate and

increases activity to produce more pyruvate and therefore acetyl-CoA for citrate

production. This results in an increase in flux through the TCA cycle (Mitsch et

al., 1999). Also, increases in acetyl-CoA concentration down regulate DME

activity to ensure malate is available for malate dehydrogenase (Mitsch et al.,

1999). Thus the NAD+ dependent malic enzyme might be important in

maintaining the correct balance of TCA cycle substrates (malate and acetyl-CoA)

and hence regulate carbon flux into the TCA cycle.

29

The role of the second malic enzyme (TME) in bacteroids is uncertain. In

bacteroids of S. meliloti TME has much lower activity in compared to DME

(Driscoll & Finan, 1993). This lower activity arises from a fall in TME transcription

to 20% relative to free-living cells, whereas DME is unaltered in the bacteroid and

when grown on different carbon sources (Driscoll & Finan, 1997). The down

regulation of TME in association with the plant host is highlighted when dme is

placed under control of the tme promoter in the dme- strain. Plants are decreased

in both dry weight and acetylene reduction to 20% compared to plants inoculated

with the wild type (Mitsch et al., 1999). On the other hand TME has substantial

activity in B. japonicum (Kouchi et al., 1988; Copeland et al., 1989; Kimura &

Tajima, 1989). The relative activity of the two malic enzymes in B. japonicum has

been suggested to contribute to the accumulation the carbon storage compound

poly-β-hydroxybutyrate (PHB) (see section 1.4.4.2).

30

Fig 1.4 The TCA cycle and possible integration of pathways used in its regulation in the bacteroid.

TCA cycle diag2.ppt

31

Addendum to Fig 1.4. List of Enzymes. AAT Aspartate aminotransferase ACN Aconitase ALD L-Alanine dehydrogenase ω-AM ω-Amidase ASP Aspartase CS Citrate synthase FUM Fumarase GAM Glutaminase GMAT Glutamine aminotransferase GAT γ-aminobutyrate aminotransferase GDC Glutamate decarboxylase GDH Glutamate dehydrogenase GOGAT Glutamate synthase GS Glutamine synthetase ICDH Isocitrate dehydrogenase ICL Isocitrate lyase KDC α-Ketoglutarate decarboxylase KGDH α-Ketoglutarate dehydrogenase MDH Malate dehydrogenase ME Malic enzyme MS Malate synthase PDH Pyruvate dehydrogenase PEPCK Phosphoenolpyruvate carboxykinase SCS Succinyl-CoA synthetase SDH Succinate dehydrogenase SSDH Succinic semialdehyde dehydrogenase

32

1.4.4 Poly-β-hydroxybutyrate (PHB)

Forsyth et al. (1958) first noted the presence of poly-β-hydroxybutyrate (PHB) in

the root nodules of legumes (Forsyth et al., 1958). PHB is a polyester composed

of the D-(⎯) stereoisomer of β-hydroxybutyrate. A co-polymer has also been

described in S. meliloti containing β-hydroxybutyrate and β-hydroxyvalerate

(Tombolini & Nuti, 1989). PHB and its co-polymers are classified as poly-β-

hydroxyalkanoates. On purely chemical grounds, PHB provides an excellent

carbon storage reserve due to its highly reduced state and because its virtually

insoluble crystalline structure exerts negligible osmotic pressure on the cell

(Dawes & Senior, 1973). Where PHB accumulates in the bacterial cell it forms

large spherical granules that can be stained with sudan black or can be

visualised on electron micrographs as electron transparent globules in the cell.

1.4.4.1 Molecular basis for PHB metabolism

PHB synthesis and degradation are collectively referred to as the PHB cycle. The

extent of PHB accumulation is dependant on the relative rates of synthesis and

degradation which in turn are controlled by growth conditions in free-living cells.

The factors governing the rates of PHB synthesis and breakdown are not fully

understood in bacteroids.

The most common pathway for PHB production in microorganisms, and the one

used by rhizobia, begins with the condensation of 2 molecules of acetyl-CoA to

form acetoacetyl-CoA. This step is catalysed by the enzyme β-ketothiolase.

Acetoacetyl-CoA is then reduced by an NADPH dependent acetoacetyl-CoA

33

reductase to β-hydroxybutyryl-CoA and incorporated into PHB by PHB synthase

(Fig 1.5).

The genetics of rhizobial PHB synthesis was first elucidated in S. meliloti. β-

ketothiolase (phaA) and acetoacetyl-CoA reductase (phaB) form an operon and

there is another open reading frame immediately upstream of phaAB, but coded

in the opposite direction (Tombolini et al., 1995). This ORF, designated aniA, is a

putative transcriptional regulator but its role has not been clearly defined (Povolo

& Casella, 2000). The aniA gene is expressed only when cells are incubated

under low oxygen concentrations. Mutation of aniA does not prevent PHB

production but it dramatically alters carbon partitioning under microaerobic

conditions resulting in an overproduction of exopolysaccharide (EPS). In

symbiosis the mutant produces small colorless nodules dramatically reduced in

nitrogen fixation (Povolo & Casella, 2000). The gene encoding PHB synthase

(designated phaC for poly-β-hydroxyalkanoate synthase or sometimes referred to

as phbC for poly-β-hydroxybutyrate synthase) is not structurally linked to phaA

and phaB (Tombolini et al., 1995; Willis & Walker, 1998; Cai et al., 2000). The

gene has been mapped to the chromosome (Cai et al., 2000). This genetic

organisation has been confirmed by the more recent genome project of S. meliloti

strain 1021. The phaA and phaB genes lie on the chromosome from positions

3547725 to 3546547 and 3546426 to 3545704 respectively

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). The PHB

synthase gene lies on the chromosome from position 1868133 to 1869965. There

are no other genes associated with PHB metabolism in the vicinity of phaC. A

gene involved in sugar fermentation stimulation is located downstream and a

Comment: Page: 1

34

Rhizobium-specific intergenic mosaic element (RIME) is located downstream of

PHB synthase (Willis & Walker, 1998;

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). RIMEs

are approximately 100 bp repeat elements, characterised by two large

palindromes, located outside of the coding regions and are thought to be involved

in regulation (Osteras et al., 1995). As there are no PHB synthesis genes

structurally associated with the gene encoding PHB synthase, it is assumed to be

monocistronic (Tombolini et al., 1995; Willis & Walker, 1998).

The degradation of PHB is initiated by the depolymerisation of the polymer by

PHB depolymerase to form β-hydroxybutyrate. This is oxidised to acetoacetate

by β-hydroxybutyrate dehydrogenase. Acetoacetate is converted to acetyl-CoA in

two steps involving the enzymes acetoacetyl-CoA synthetase and β-ketothiolase

(Fig 1.5). Acetoacetyl-CoA synthetase is required for the activation of

acetoacetate to acetoacetyl-CoA and β-ketothiolase degrades acetoacetyl-CoA to

two molecules of acetyl-CoA. The only detailed analysis of the degradation

pathway in rhizobia has been provided by Charles and co-workers in S. meliloti.

They have isolated 4 loci that are unable to utilise β-hydroxybutyrate and / or

acetoacetate as sole carbon sources (Charles & Finan, 1991; Charles et al.,

1997). Further characterisation of the genes has shown they encode

methylmalonyl-CoA mutase (bhbA) (Charles & Aneja, 1999), β-hydroxybutyrate

dehydrogenase (bdhA) (Aneja & Charles, 1999), acetoacetyl-CoA synthetase

(acsA) and poly-β-hydroxybutyrate synthase (phaC) (Cai et al., 2000). The

requirement for the enzymes methylmalonyl-CoA mutase, which catalyses the

interconversion of succinyl-CoA and methylmalonyl-CoA, and poly-β-

35

hydroxybutyrate synthase in the degradation of PHB are not yet known. It has

been speculated that methylmalonyl-CoA or succinyl-CoA might be involved in

the donation of CoA to acetoacetate during the formation of acetoacetyl-CoA on

breakdown of PHB (Charles & Aneja, 1999). The authors also suggest that the

PHB synthase mutation might increase in vivo levels of acetoacetyl-CoA (and β-

hydroxybutyryl-CoA) such that activity of acetoacetyl-CoA synthetase is inhibited.

Consistent with this the acsA gene in multicopy supresses the growth of the PHB

synthase mutant on acetoacetate (Cai et al., 2000).

Unlike the PHB synthesis genes some of these degradative genes are encoded

on Sym plasmids in S. meliloti. S. meliloti has one chromosome of ca 3.7 Mb and

two large plasmids of ca. 1.4 and 1.7 Mb that are commonly described as pSymA

and pSymB, respectively, which carry genes essential for symbiosis. PHB

depolymerase is located on pSymA from position 1116492 to 1115368

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). β-

hydroxybutyrate dehydrogenase (bdhA) is located on pSymB (Aneja & Charles,

1999) from position 1244108 to 1244881

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). This

implies a role for PHB metabolism during symbiosis even though the polymer

does not accumulate in bacteroids.

Comment: Page: 1

36Fig 1.5 Pathways leading to the synthesis and degradation of poly-β-hydroxybutyrate in rhizobia.

TCA

Pyruvate

Malate

CARBON FROM PLANT

PHB β-Hydroxybutyryl-CoA

Acetoacetate β- Hydroxybutyrate

Acetoacetyl-CoA

Acetyl-CoA

CH3 O

O.CH.CH2.C n

PHB depolymerase

PHB synthase

β-Hydroxybutyrate dehydrogenase

CoA transferase

β-Ketothiolase

Acetoacetyl-CoA reductase

NADP

NADPH

37

1.4.4.2 PHB metabolism in bacteroids

The function of PHB in bacteroid metabolism has been the subject of debate

since the 1970’s. Bacteroids have a high demand for carbon to meet their

respiratory requirements yet PHB forms considerable reserves in bacteroids (up

to 70% dry weight) of several rhizobia that form determinate nodules on legumes,

such as B. japonicum, R. etli and R. leguminosarum bv. phaseoli. The polymer

does not accumulate in bacteroids of indeterminate nodules, such as S. meliloti

and R. leguminosarum bv. viciae. In S. meliloti, which forms indeterminate

nodules on alfalfa, PHB may accumulate up to 50% dry weight of free-living cells

(Tombolini & Nuti, 1989). Ultrastructural studies have indicated PHB granules

may be present in cells inside the infection thread (Paau & Cowles, 1978) but

they disappear when, or shortly after, cells are released from the infection thread.

Mature nitrogen fixing bacteroids contain no visable granules of PHB (Hirsch et

al., 1983; Vasse et al., 1990). PHB can accumulate in R. etli, which forms

determinate nodules on bean, in both free-living cells and bacteroids.

Ultrastructural analysis has shown that PHB is not present in the Rhizobium etli

cells present in the infection thread. It starts to accumulate in young bacteroids

and forms a considerable reserve in mature bacteroids (Cermola et al., 2000).

Bacteroids of a R. etli mutant in nifA produces little or no PHB (Cermola et al.,

2000). NifA is the positive transcriptional activator for various nitrogenase

structural genes. Therefore a mutation in nifA abolishes the ability to synthesise

nitrogenase and hence fix nitrogen. This result strongly suggests that in

determinate nodules the accumulation of PHB in bacteroids is correlated with

nitrogen fixation. However the precise initiating factor(s) for accumulation has yet

to be clarified. Conversely factors that obviate the accumulation of PHB in

38

bacteroids of indeterminate symbioses are also not known. Hypotheses based on

the division of carbon between malate utilising enzymes have been suggested to

explain the accumulation or not of PHB in bacteroids. However, there is little

experimental evidence to back these hypotheses so they remain circumstantial.

It has been suggested by Day et al. (1994) that the relative affinities of the two

malic enzymes (see section 1.4.3) for malate contributes to the accumulation of

PHB in B. japonicum bacteroids. Under circumstances of low malate provision,

the higher affinity TME would be active favoring supply of NADPH for

nitrogenase. Because DME would be virtually inactive under these

circumstances, proportionally more malate would flow into the TCA cycle via

MDH and acetyl-CoA would be consumed via citrate synthase (CS) thus

preventing PHB biosynthesis. Under circumstances of high malate provision the

lower affinity DME would also be active. This would lead to inhibition of TCA

cycle enzymes by NADH and the concomitant utilisation of a portion of the

NADPH and acetyl-CoA in PHB biosynthesis, thus making less reductant

available for nitrogenase. Indeed it has been demonstrated that high levels of

malate supply to B. japonicum bacteroids incubated ex planta primarily increases

PHB reserves and there is inhibition of nitrogen fixation under these

circumstances (Bergersen & Turner, 1990; Bergersen & Turner, 1992). Levels of

TME differ between soybean and alfalfa bacteroids. TME has substantial activity

in B. japonicum (Kouchi et al., 1988; Copeland et al., 1989; Kimura & Tajima,

1989), whereas in bacteroids of S. meliloti TME activity is repressed (Driscoll &

Finan, 1993; Driscoll & Finan, 1996; Driscoll & Finan, 1997). Therefore the type

of regulation of carbon flux described above may not be important in S. meliloti

bacteroids.

39

Factors that prevent accumulation of PHB in bacteroids of indeterminate nodules

have been studied by Copeland and co-workers. Mesorhizobium ciceri CC1192,

which forms nodules on chickpea, can produce PHB in free-living cells but

bacteroids do not accumulate the reserve (Lee & Copeland, 1994). The activities

of β-ketothiolase, acetoacetyl-CoA reductase, PHB depolymerase, and β-

hydroxybutyrate dehydrogenase are similar in free-living cells and bacteroids of

CC1192 and GC analysis has shown that small amounts of PHB are present in

bacteroids (Kim & Copeland, 1996). Label derived from 14C-malate is

incorporated into β-hydroxybutyrate by isolated CC1192 bacteroids providing

evidence that these cells retain the enzymatic capacity to make PHB (Chohan &

Copeland, 1998). This data strongly suggests that the potential to metabolise

PHB is not lost upon differentiation and that biochemical controls are likely to

operate that direct acetyl-CoA into the TCA cycle rather than towards PHB

synthesis in CC1192 bacteroids. β-Ketothiolase from CC1192 bacteroids and

free-living cells show no significant differences in kinetic properties (Kim &

Copeland, 1997) and the enzyme has similar kinetic properties in B. japonicum

(Suzuki et al., 1987). The condensation of acetyl-CoA by β-ketothiolase is

strongly inhibited by free CoA (Ki value for CoA of less than 10µM). The Km for

acetyl-CoA of β-ketothiolase is 1mM but is reduced to 0.23mM on addition of N-

ethylmaleimide (NEM). NEM reacts with free CoA therefore presumably relieves

the inhibition by CoA thus lowering the Km for acetyl-CoA (Kim & Copeland,

1997). It has been hypothesised that in free-living CC 1192 cells, the acetyl-

CoA/CoA ratio reaches a value that allows condensation activity of β-

ketothiolase, but that in CC 1192 bacteroids, the ratio is poised so that the

formation of acetoacetyl-CoA is not favored (Kim & Copeland, 1997). A second

40

possible control mechanism might be the greater potential for oxidizing malate to

oxaloacetate in CC1192 bacteroids than in free-living cells (Kim & Copeland,

1996). This would lead to the faster incorporation of acetyl-CoA into the

tricarboxylic acid cycle in bacteroids compared to free-living cells. It is perhaps

worth noting that in other indeterminate symbioses (R. leguminosarum bv. viciae)

a similar increase in MDH has been reported (McKay et al., 1989) whereas in

determinate symbioses (R. tropici and B. japonicum) there is little or no change in

MDH activity (Romanov et al., 1994; Kim & Copeland, 1996).

In considering bacteroid PHB accumulation much emphasis has been placed the

on factors that may or may not initiate its synthesis. The extent of PHB

accumulation is dependant on the relative rates of synthesis and degradation

however little attention has been placed on the rate and regulation of degradation

in either the indeterminate or determinate bacteroid. In R. etli free-living cells,

PHB is continually synthesised and degraded even under conditions in which the

polymer does not build up (Encarnacion et al., 1995). Bergersen (1992) has

presented evidence to show PHB is also continually cycled in B. japonicum

bacteroids (Bergersen & Turner, 1992). Therefore the formation of large reserves

in determinate nodules might merely result from a slower breakdown than

synthesis rate. Conversely in indeterminate nodules PHB might be rapidly cycled

through such that reserves do not accumulate.

1.4.4.3 Role of PHB in bacteroids

As well as questioning why PHB accumulates in certain bacteroids much

emphasis has been placed on the function of PHB in bacteroids, particularly in

determinate nodules. The main theories that occur on the role of PHB in

41

determinate symbioses are: (1) PHB sequesters carbon and reduced nucleotides

to maintain TCA cycle activity under microaerobic conditions. (2) PHB

metabolism may support nitrogenase activity by providing carbon skeletons, ATP,

and/or reducing equivalents via hydroxybutyrate dehydrogenase when alternative

(plant) carbon sources are scarce. (3) Conversely, PHB metabolism may be

independent of nitrogenase fixation and compete with or operate at the expense

of Nitrogenase.

During symbiosis, PHB accumulation begins at the onset of microaerobic

conditions in B. japonicum and its content increases during nodule development

when there is a decrease in the oxygen uptake activity (Karr et al., 1984; Karr &

Emerich, 1988; Karr et al., 1990). It has been suggested that bacteroid PHB

synthesis serves to regulate the TCA cycle under microaerobic conditions by

drawing reductant and carbon away from the cycle (McDermott et al., 1989). The

removal of reduced nucleotides would alleviate TCA cycle enzyme inhibition and

the removal of carbon would balance the amount of carbon metabolised by the

cycle.

PHB metabolism has been extensively studied in Azotobacter. In Azotobacter

factors that limit growth, such as ammonia or phosphate limitation, lead to the

accumulation of PHB although the amounts that accumulate are smaller than

those found under oxygen limiting conditions (Dalton & Postgate, 1969; Senior &

Dawes, 1971; Senior et al., 1972; Senior & Dawes, 1973; Jackson & Dawes,

1976). Batch cultures of Azotobacter beijerinckii start to accumulate PHB towards

the end of exponential growth and during stationary phase when oxygen

becomes limiting (Senior et al., 1972). Under relaxed oxygen concentrations

42

acetyl-CoA is fed into the tricarboxylic acid cycle, and the resultant CoA inhibits

the β-ketothiolase activity. Under oxygen limitation and carbon excess, NADPH

increases and inhibits citrate synthase and isocitrate dehydrogenase, raising

levels of acetyl-CoA and lowering CoA levels hence alleviating inhibition of β-

ketothiolase by CoA and allowing synthesis of PHB to proceed (Senior & Dawes,

1973). Therefore accumulation of PHB in Azotobacter is controlled by the redox

state of the cell.

The extent of PHB accumulation in rhizobia in free-living culture is dependant on

both the culture condition and the strain (Zevenhuizen, 1981; Tombolini & Nuti,

1989). But it is apparent that initiation of PHB synthesis is essentially the same to

that described in Azotobacter. Any growth condition that does not interrupt

carbon supply but reduces the flow of carbon into biosynthetic reactions or

respiration increases PHB content in free-living cells. Azorhizobium caulinodans

ORS571 accumulates PHB up to 37% in O2 limited conditions; 11% under

ammonium limitation; 10% under nitrate limitation; and 26% under Mg2+ limitation

(De Vries et al., 1984; De Vries et al., 1986). In A. caulinodans ORS571 PHB is

degraded only under conditions of high oxygen (Stam et al., 1986).

Studies to assess the accumulation of PHB in response to impairment of the TCA

cycle have been carried out in R. etli. Upon serial sub-culture on media

(succinate) without vitamins (biotin) a fermentative like response is shown by R.

etli. This response is characterised by a reduction or loss of TCA and

anapleurotic enzyme activity (citrate synthase, isocitrate dehydrogenase,

oxoglutarate dehydrogenase and pyruvate dehydrogenase), the excretion of

amino and organic acids into the medium (alanine, glutamate, pyruvate, malate,

43

fumarate) and by PHB accumulation (Encarnacion et al., 1995). Mutation in PHB

synthase (phaC) in R. etli results in the fermentative response being more

extreme with respect to an increase in excretion of organic acids and decrease in

enzyme activity indicating a lowered ability of the PHB mutant to metabolise

carbon under these conditions of stress (Cevallos et al., 1996). The restriction of

growth and metabolism in these free-living cells might mimic the bacteroid

metabolic state and hence indicate the importance of certain metabolic pathways

in the bacteroid. Therefore this data suggests that PHB synthesis is required by

R. etli to help regulate carbon metabolism in the bacteroid. However, the

presence of large reserves of PHB under biotin limiting conditions might also be

influenced by a biotin regulation of the PHB degradation pathway. Cells of S.

meliloti accumulate significantly more PHB granules under biotin limitation

suggesting that either PHB synthesis or degradation are influenced by biotin

(Hofmann et al., 2000). Mobilisation of a β-hydroxybutyrate dehydrogenase-lacZ

(bdhA-lacZ) fusion into S. meliloti and subsequent β-galactosidase assays have

shown the addition of biotin increases the transcription of bdhA by up to 5-fold

(Hofmann et al., 2000).

Restriction of growth due to oxygen limitation also increases the PHB content in

R. etli (Encarnacion et al., 1995). Decreasing the oxygen available to cultures

from 100% saturation of growth medium with oxygen to 5% saturation increases

PHB content from 0.15 to 0.75mg/mg of protein respectively (Encarnacion et al.,

1995). This strongly suggests that in R. etli PHB would accumulate in the

bacteroid in response to low oxygen conditions. If this pathway is important in

regulating carbon metabolism under microaerobic conditions then mutation of the

metabolic pathway might have an adverse effect on carbon metabolism by the

44

bacteroid. This is apparently not the case as mutation of phaC (PHB synthase)

does not adversely effect nitrogen fixation in the bean nodule (Cevallos et al.,

1996). However preventing PHB synthesis resulted in the re-direction of carbon

flux to other pathways. Blocking TCA enzyme activity in the mutant by serial sub-

culture on media without vitamins resulted in an increase in glycogen

accumulation of up to 50-fold (Cevallos et al., 1996). This redirection of carbon

flux indicates the flexibility of carbon metabolism in the rhizobia and the potential

use of alternative pathways to PHB synthesis in the regulation of bacteroid

carbon metabolism.

Nitrogen fixation by a phbC (PHB synthase) mutant of Azorhizobium caulinodans

is defective. The strain forms Fix- nodules and is Nif- in free-living state (Mandon

et al., 1998). The free nucleotide content of the mutant is higher, reflecting a

lowered NAD/NADH ratio, and the AMP level is higher, reflecting decreased ATP

availability, indicating the ability of the mutant to metabolise carbon is severely

impaired. However, the defect in nitrogen fixation is not due to impaired carbon

metabolism but is due to a lack of nitrogenase. A NifH-lacZ fusion is not

expressed in the mutant background indicating nitrogenase is not made.

Expression of nitrogen fixation (nif) genes depends on the transcriptional

activator NifA which is in turn under the control of various genes regulated by the

oxygen and ammonium status of the cell. Fusions under these controls (ie.

oxygen and ammonium status) are active in the mutant (Mandon et al., 1998).

Therefore the reason for the lack of nitrogenase in the mutant seems complex

and further study is required in this mutant to elucidate the PHB/nitrogenase

interaction.

45

If PHB exists inside bacteroids purely as a carbon storage compound PHB

degradation may support nitrogenase activity by providing carbon skeletons,

ATP, and/or reducing equivalents via β-hydroxybutyrate dehydrogenase when

plant carbon sources are scarce. However, the initial experiments to ascertain

this by Wong & Evans (1971) did not show PHB supported nitrogen fixation in

soybean. The authors had previously shown that β-hydroxybutyrate is able to

support nitrogenase activity of bacteroids in vitro (Klucas & Evans, 1968). The

1971 paper investigated the ability of PHB to support nitrogenase activity in

plants incubated in the dark. Soybeans incubated in the dark show a reduction in

nitrogenase activity (down to 37% of the initial rate). This lowered level of

nitrogen fixation is not apparently supported by PHB as there is little change in

the reserve size. A second experiment using detached nodules showed there is a

decrease in nitrogenase activity to 35% but a decrease in PHB content by only

1% 10 hours after nodule excision (Wong & Evans, 1971). The authors

concluded that PHB does not support nitrogen fixation even though β-

hydroxybutyrate depolymerase is present in high levels suggesting bacteroids

have the capacity to break down PHB (Wong & Evans, 1971).

However, the majority of reports since then have concluded PHB can support

nitrogenase activity (Klucas, 1974; Kretovich et al., 1977; Romanov et al., 1980;

Bergersen & Turner, 1990; Bergersen et al., 1991). Within determinate

symbioses where PHB accumulates in the bacteroid, the % dry weight formed by

PHB shows diurnal fluctuations and developmental changes in response to

nitrogenase activity. Both bacteroids of R. leguminosarum bv. phaseoli and R.

lupini show daily fluctuations in PHB content which correlate to the daily

fluctuations in nitrogenase activity which is attributed to differences in carbon

46

supply from the plant between night and day. PHB content is higher at night when

nitrogenase activity is at its lowest and vice versa (Kretovich et al., 1977).

In the majority of cases, nitrogen fixation ceases when seeds start to form as a

consequence of competition between developing seeds and nodules for

photosynthate (Phillips, 1980). In this situation PHB content tends to increase

inside bacteroids throughout seed formation (Klucas, 1974; Romanov et al.,

1980). For example, the PHB content of lupin bacteroids is reportedly high at the

beginning of the symbiosis (13-14% dry weight), declines as nitrogenase activity

increases (to 3-4% dry weight at the period of maximum nitrogen fixation), and

accumulates again (to 17% dry weight) when nitrogen fixation and bacteroid

respiration decrease at the time of seed ripening (Romanov et al., 1980).

Enzymes involved in PHB metabolism in R. lupini have been shown to be

especially active during periods of intensive nitrogen fixation (Romanov et al.,

1980).

In certain symbioses nitrogen fixation continues into seed production (Bergersen

et al., 1989; Bergersen et al., 1992). Prolonged nitrogen fixation may benefit the

plant by increasing seed size and nitrogen content (Ismande, 1989). In these

plants, bacteroid PHB content declines as nitrogen fixation continues into seed

development (Bergersen et al., 1991) further suggesting PHB is used to support

nitrogenase activity in the bacteroids. Evidence to support this was obtained from

experiments where plants were stem girdled to prevent the flow of substances

through the phloem to the root. In flowering soybean plants in which the phloem

was not disrupted the previous evening, there was no overnight degradation of

PHB (Bergersen et al., 1991). However, when the plants are stem girdled there is

47

a decrease in PHB content of bacteroids. The same experiments at seed

formation result in an overnight decrease in the PHB content and increase in β-

hydroxybutyrate levels of bacteroids whether plants are girdled or not, suggesting

PHB is degraded to meet metabolic requirements of the bacteroid when plant

carbon supply is sub-optimal (Bergersen et al., 1991). It has been calculated that

PHB may sustain nitrogen fixation of soybeans for up to two days when there is

no other source of substrate available (Bergersen & Turner, 1990).

Bergersen showed that PHB accumulates in bacteroids of B. japonicum and is

used to support nitrogen fixation using a series of flow chamber experiments

(Bergersen & Turner, 1990 a & b; Bergersen & Turner, 1992; Bergersen &

Turner, 1993). The flow chamber set up was composed of bacteroids incubated

in a stirred liquid medium, with controlled variable substrate and oxygen

concentrations, and with continuous removal of substrate. Initial experiments

show that bacteroids are able to fix nitrogen using endogenous reserves (no

substrate supplied) (Bergersen & Turner, 1990). Further experiments show that

when supplied with 10mM 14C-malate bacteroids do not fully oxidise the substrate

and accumulate 14C in PHB (90% total bacteroid label as PHB after 5 hours)

(Bergersen & Turner, 1990). In association with the accumulation of PHB under

these conditions there is an increase in bacteroid demand for oxygen and CO2

production, presumably due to an increase in action of malic enzyme and

pyruvate dehydrogenase to supply acetyl-CoA (see section 1.4.3). Under these

conditions there is inhibition of nitrogen fixation until substrate levels are lowered

through bacteroid metabolism or through washing away from the medium

(Bergersen & Turner, 1990; Bergersen & Turner, 1992). This suggests that

exogenous reserves primarily serve to build up endogenous reserves, which are

48

in turn used to support nitrogen fixation. When supplied with no substrate PHB

reserves are depleted by 9.2% over a 5 hour period, and the vast majority of

carbon in CO2 is derived from PHB (Bergersen & Turner, 1992).

That dramatic changes in bacteroid PHB content (increases and decreases) can

be determined within hours suggests that there is a rapid flux of carbon through

PHB. Furthermore, exposure of whole plants to 14CO2 has been shown to result

in the accumulation of 14C in PHB, but in no change in the overall percentage

content PHB of bacteroids, suggesting the constant cycling of bacteroid carbon

through PHB (Romanov et al., 1980). This implies an important role for the PHB

degradation pathway in bacteroids where PHB accumulates.

However PHB metabolism might be independent of nitrogen fixation and actually

compete with or operate at the expense of nitrogenase. PHB accumulates up to

70% the dry weight of soybean bacteroids (Bergersen & Turner, 1990) and this

sequestration of photosynthate has been suggested to be wasteful (Wong &

Evans, 1971). If this is the case then eliminating the capacity to make PHB might

enhance the ability to fix nitrogen (Emerich & Evans, 1984). In agreement with

this, a mutation in phaC in R. etli results in an improvement in the efficiency of

symbiosis. The nitrogenase activity of the mutant is higher than the wild type and

the mutation prolongs the capacity of the bacteroids to fix nitrogen. Plants are

increased in dry weight, have more seeds, and the seeds have a higher N

content compared to plants inoculated with the wild type (Cevallos et al., 1996).

This suggests that in determinate symbioses, PHB synthesis may compete with

or operate at the expense of nitrogenase activity for substrates (reductant and

ATP).

49

As suggested previously the absence of PHB reserves in indeterminate

bacteroids might be due to the rate of synthesis versus breakdown. Therefore the

formation of large deposits of PHB is not in itself an indication of the importance

of PHB metabolism. One of the simplest ways to evaluate the importance of PHB

metabolism in symbiosis is to mutate PHB metabolic enzymes. Whereas in the

determinate R. etli – bean symbiosis there was an increased capacity to fix

nitrogen in a PHB synthase mutant, mutation of PHB synthase in S. meliloti has

no effect on nitrogen fixation (Povolo et al., 1994; Willis & Walker, 1998; Cai et

al., 2000). Mutation of the genes methylmalonyl-CoA mutase (bhbA), β-

hydroxybutyrate dehydrogenase (bdhA) and acetoacetyl-CoA synthetase (acsA),

that are involved in the degradation of PHB in S. meliloti, does not effect nitrogen

fixation (Aneja & Charles, 1999; Charles & Aneja, 1999; Cai et al., 2000). There

are no significant differences in shoot dry weight between plants infected with

wild type versus mutant in each case. These data suggest neither PHB synthesis

nor degradation is important for an effective symbiosis between S. meliloti and

alfalfa.

However, mutation of PHB synthase affects the competitive ability of the mutant

to form bacteroids. Alfalfa plants co-inoculated with the wild type strain and a

mutant in phbC are nodulated more frequently by the wild strain (Willis & Walker,

1998). The stage at which this competitive advantage occurs during the process

of nodule development is not known. Circumstantial evidence hints at a role for

PHB metabolism during bacteroid development. Ultrastructural studies have

indicated PHB granules are present in cells inside the indeterminate infection

thread (Paau & Cowles, 1978) but they disappear when, or shortly after, cells are

released from the infection thread. Mature nitrogen fixing bacteroids contain no

50

visable granules of PHB (Hirsch et al., 1983; Vasse et al., 1990). Furthermore

bacA mutant cells, that abort bacteroid development, are not depleted of PHB

reserves upon release from the infection thread whereas wild type cells are

(Glazebrook et al., 1993). This potential role of PHB metabolism has not been

studied in detail.

51

1.4.5 Glycogen

Unlike PHB, the occurrence of large reserves of bacteroid glycogen has received

very little attention and there is scant Rhizobium – glycogen related literature.

The presence of glycogen in bacteroids of R. leguminosarum bv. trifolii was noted

by Dixon (1967). Upon emergence from the infection thread bacteroids increase

the content of a diffuse electron transparent material that becomes a predominant

feature of the mature bacteroid cytoplasm. The material was confirmed as being

glycogen by staining bacteroids for glycogen before and after digestion with

amylase (Dixon, 1967). The presence of glycogen has also been noted in

Rhizobium strains infecting lotus (Craig & Williamson, 1972).

For the formation of glycogen a source of the precursor molecule glucose must

be available in bacteroids. The peribacteroid membrane has very limited

permeability to sugars (Udvardi et al., 1990) suggesting the glucose is derived via

gluconegenesis. Phosphoenolpyruvate carboxykinase (PEPCK) has been shown

to be active in pea bacteroids (McKay et al., 1985). The pathway for glycogen

production in E.coli (Preiss & Romeo, 1994), and the one used by rhizobia,

begins with the isomerisation of glucose-6-P to glucose-1-P by

phosphoglucomutase. This is converted to ADP-glucose by ADP-glucose

phosphorylase. This undergoes polymerisation via α-1,4 linkages by glycogen

synthase. Subsequently the branching enzyme forms α-1,6-glycosidic linkages.

Glycogen is degraded by removal of the α-1,6-glycosidic linkages by the

debranching enzyme, then by removal of the α-1,4 linkages by glycogen

phosphorylase to give glucose-1-P (Fig 1.6).

52

Fig 1.6 Pathway leading to the synthesis and degradation of glycogen in Rhizobium

UDP Glucose Exopolysaccharide (EPS)

SYNTHESIS

DEGRADATION

ADP glucose phosphorylase

Phosphogluco-mutase

Glycogen phosphorylase

Glycogen debranching Glucose-6-P Glucose-1-P

Glycogen synthase

Glycogen branching

ADP Glucose 1,4-Glucosen Glycogen

53

Genetic organisation of glycogen synthesis genes has been studied in

Agrobacterium tumefaciens (Uttaro & Ugalde, 1994; Ugalde et al., 1998; Uttaro et

al., 1998) and has recently been shown to be the same in R. tropici (Marroqui et

al., 2001) and S. meliloti (http://sequence.toulouse.inra.fr/meliloti.html,

unpublished data, 2001). The nucleotide sequence of the entire rhizobia glg

cluster reveals that a continuous DNA fragment of over 7 Kb contains six genes

arranged similarly to E. coli glg genes (Preiss & Romeo, 1994) (Fig 1.7). All six

genes are transcribed in the same direction and analysis with lacZ gene fusions

suggest the first five genes are organised in one operon, although pgm also has

its own promoter (Marroqui et al., 2001). glgX is transcribed independently

(Marroqui et al., 2001). The operon is located on the chromosome of S. meliloti at

position 3065247 to 3076592. There are second copies of glgA, glgX and glgB on

plasmid pSymB but these do not form an operon.

Fig 1.7 Organisation of glycogen biosynthetic genes in Rhizobium tropici from Marroqui et al. (2001)

Little is known about the factors that lead to the accumulation of glycogen in free-

living cells or in bacteroids. As with PHB synthesis in free-living cells, glycogen

accumulates under growth limiting conditions such as nitrogen limitation

(Zevenhuizen, 1981). This suggests that glycogen metabolism may fulfill a similar

glgP glgB glgC glgA glgX pgm

Glycogen branching

ADP glucose phosphorylase

Glycogen synthase

Phosphogluco-mutase

Glycogen debranching

Glycogen phosphorylase

54

role as PHB metabolism. Free-living B. japonicum, R. leguminosarum and S.

meliloti can produce glycogen at the same time as PHB (Tsien & Schmidt, 1977;

Zevenhuizen, 1981; Povolo et al., 1994; Povolo & Casella, 2000). Preventing

PHB synthesis by mutation of phaC results in a greater accumulation of glycogen

(50-fold) when the TCA cycle is impaired by serial sub-culture on media without

vitamins (Cevallos et al., 1996). This suggests that the control of carbon flux into

either of these two pools is closely regulated.

The role of glycogen metabolism during symbiosis has only been studied in R.

tropici. A mutant in glycogen synthase (glgA) was isolated by screening Tn5

induced mutants for enhanced respiration via cytochrome oxidase activity. The

glgA mutant has increased levels of the cytochromes c1 and CycM and a small

increase in the amount of cytochrome aa3 but the reason for these increases is

not clear. The mutation results in an increase in the symbiotic efficiency of bean

plants as measured by an increase in dry weight of up to 38%. This might be a

similar response to the phaC mutant of R. etli that also enhanced nitrogen fixation

(see section 1.4.4.3). In this case glycogen synthesis could then be considered

as competing with or operating at the expense of nitrogenase activity for

substrates (reductant and ATP). However, the reason for the increased symbiotic

efficiency of the glgA mutant is uncertain. It is most likely to be due to increased

nodulation rather than increased nitrogenase activity. The reason for increased

nodulation is not understood however the mutant had lowered levels of EPS

which might have effected nodulation. Or the mutant might have impaired

nitrogen fixation (delayed onset of fixation or regulatory problem due to reduced

efficiency of carbon metabolism) thus inducing the plant to initiate the formation

55

of additional nodules, actually resulting in a net increase in symbiotic

performance (Marroqui et al., 2001).

56

1.5 Amino acid biosynthesis and metabolism

There are a substantial number of studies that show amino acids, particularly

glutamate, aspartate and alanine, are synthesised intracellularly by bacteroids. It

has been proposed that amino acid synthesis is involved in the regulation of the

TCA cycle under microaerobic conditions (McDermott et al., 1989).

1.5.1 Bacteroid glutamate metabolism

Isolated B. japonicum bacteroids incubated anaerobically with 14C-malate or

succinate accumulate 20-40% of the label in glutamate (Salminen & Streeter,

1987) and bacteroid extracts analysed for carbohydrate, organic acid and amino

acid content show that glutamate is a major component of the total metabolite

content (Streeter, 1987 a & b). Furthermore, detached nodules of both soybean

and pea rapidly label glutamate following short term exposure to 14CO2 (Salminen

& Streeter, 1992). Malate is rapidly labelled in both bacteroid and plant cytosol in

nodules of incubated in the presence of 14CO2, consistent with the prior evidence

of it being the primary carbon source for the bacteroid. Glutamate is labelled

slowly in the cytosol but rapidly labelled in the bacteroid, suggesting that the

glutamate label is derived from the metabolism of malate by the bacteroid

(Salminen & Streeter, 1992). This high rate of glutamate synthesis may be due to

inhibition of α-ketoglutarate dehydrogenase activity under microaerobic

conditions. As stated previously the low NADH:NAD+ ratio in B. japonicum

bacteroids probably results in more than 50% inhibition of KGDH in vivo

(Salminen & Streeter, 1990). This significant diversion of labelled carbon from the

TCA cycle to glutamate is consistent with the inhibition of α-ketoglutarate

57

dehydrogenase. The genes for the α-ketoglutarate dehydrogenase complex in R.

leguminosarum are part of the mdh-sucCDAB which includes malate

dehydrogenase and succinyl-CoA synthetase (see section 1.4.2) (Walshaw et al.,

1997). The sucDAB have the same order in B. japonicum and it seems likely that

mdh and sucC will also have the same order in B. japonicum since they are

reported to be clustered with sucA (Green & Emerich, 1997). Mutation of sucD or

sucA results in the synthesis and secretion of large quantities of glutamate by R.

leguminosarum bv. viciae (Walshaw et al., 1997). Therefore glutamate synthesis

appears to be a sink for carbon when α-ketoglutarate dehydrogenase is blocked

(Walshaw et al., 1997).

Glutamate can be synthesised from α-ketoglutarate either by glutamate

dehydrogenase (GDH), or by GS-GOGAT, or by various aminotransferases. The

GS-GOGAT pathway is the main ammonium assimilatory pathway in free-living

rhizobia (Brown & Dilworth, 1975; Kondorosi et al., 1977; Osburne & Signer,

1980; Ali et al., 1981; Donald & Ludwig, 1984; Howitt & Gresshoff, 1985; Bravo &

Mora, 1988). Traditionally it was thought that bacteroid ammonium assimilation

via GS-GOGAT did not occur during symbiosis as the enzyme activity had been

reported to be low (Brown & Dilworth, 1975; Werner et al., 1980). However, GS

and GOGAT activities have been detected (Duran et al., 1995) and the

ammonium assimilation pathway is active in R. etli bacteroids (Mendoza et al.,

1995; Mendoza et al., 1998).

The fate of accumulated glutamate in the bacteroid is not known. In certain

symbioses the GABA (γ-aminobuyrate) shunt, which metabolises α-ketoglutarate

to succinate via glutamate, GABA and succinic semialdehyde (Fig 1.4), may

58

operate. This pathway has been studied as a potential pathway by-passing α-

KGDH via glutamate because γ-aminobuyrate levels in bacteroids are often high (

Streeter, 1987; Miller et al., 1991). It has been suggested that alfalfa bacteroids

may possess the required enzymes for the GABA shunt, although the activities

vary remarkably between studies (Fitzmaurice & O'Gara, 1991; Miller et al.,

1991). Consistent with a role of GABA shunt in the alfalfa symbiosis, a mutant

that lacks a functional GABA by-pass, and is therefore unable to grow on

glutamate, forms bacteroids with a reduced ability to fix nitrogen (Fitzmaurice &

O'Gara, 1993). However, in other symbioses the pathway does not appear to

operate. Bacteroids of strain NGR234 isolated from snakebean nodules have

extremely low levels of GABA catabolic enzymes indicating the pathway does not

operate (Jin et al., 1990). It has also been concluded that the GABA shunt does

not operate in soybean bacteroids, as six different strains have been shown to

have no glutamate decarboxylase (GDC) activity (Salminen & Streeter, 1990) and

bacteroids of a sucA mutant have very low activity of glutamate decarboxylase

(Green & Emerich, 1997; Green et al., 2000).

It has been proposed that a glutamine cycle operates in Rhizobium that serves to

regulate carbon and amino pools (Duran & Calderon, 1995; Duran et al., 1995;

Encarnacion et al., 1998; Castillo et al., 2000). In this cycle glutamine

transaminase transfers the α-amino group of glutamine to a variety of 2-oxo acid

acceptors such as pyruvate and glyoxylate. The 2-oxo acid of glutamine, 2-

oxolutaramate, is released by glutamine transaminase and it is further degraded

to 2-oxoglutarate (α-ketoglutarate) and ammonium by an ω-amidase. Ammonium

is then reassimilated into glutamine/glutamate by the GS-GOGAT pathway, whilst

59

α-ketoglutarate can be fed back into the TCA cycle or re-cycled (Fig 1.4). When

the glutamine cycle is perturbed by mutation in GSI and GSII, or in GOGAT,

glutamine is preferentially used over succinate as carbon source by free-living

bacteria suggesting the GS-GOGAT pathway is necessary for efficient succinate

consumption in R. etli and S. meliloti (Encarnacion et al., 1998; Castillo et al.,

2000). Concomitantly, there is an increase in PHB accumulated from glutamine

carbon suggesting a pathway for PHB synthesis from glutamine carbon. The

amino acid is converted to β-hydroxybutyrate, via glutamate, GABA, succinic

semialdehyde and γ-hydroxybutyrate (Fig 1.4). β-hydroxybutyrate is then

polymerised to PHB (Encarnacion et al., 1998). The role of this pathway in

bacteroids has yet to be determined but could be a potential method of utilising

accumulated glutamate.

The activity of glutaminase, which catalyses the hydrolytic deamination of

glutamine to glutamate and NH3 (Fig 1.4), has been shown to be high in

bacteroids of R. lupini (Kretovich et al., 1981). Two glutaminases (A and B) have

been identified in R. etli. Glutaminase A is required for growth on glutamine as

sole carbon and nitrogen source. Glutaminase B plays a minor role in the

utilization of glutamine as a carbon source (Duran et al., 1996). Mutation of

glutaminase A results in higher levels of glutamine but lower levels of glutamate

in the bacteroid compared to the wild type. However the mutation has no effect

on nitrogen fixation suggesting the enzyme is active in maintaining the balance

between glutamate and glutamine but that this pathway is not critical to symbiosis

(Duran et al., 1995).

60

Mutational analysis of GS and GOGAT show the importance of glutamine /

glutamate metabolism during symbiosis depends on the particular association. B.

japonicum single mutants in glnA (GSI) and glnII (GSII) have been shown to

increase nitrogen fixation, but double mutants are not able to nodulate soybean

(Carlson et al., 1987). The double mutants of B. japonicum are glutamine

auxotrophs indicating this is a general affect on bacterial growth and is not

directly related to carbon and nitrogen metabolism in the bacteroid (Carlson et al.,

1987). In S. meliloti single mutation and double mutation of glnA and glnII

(Debruijn et al., 1989) and mutation of GOGAT (Ali et al., 1981; Lewis et al.,

1990) has no effect on nitrogen fixation. Mutation of GOGAT in R. etli leads to an

increase in efficiency of nitrogen fixation (Castillo et al., 2000). In free-living cells

the mutant excretes of large amounts of ammonium and has significantly

decreased amino acid pools (Castillo et al., 2000). In symbiosis the mutant

results in an increase in ureides in xylem sap suggesting more ammonium is

excreted to the plant cell if the bacteroid is unable to assimilate ammonium via

GS-GOGAT. Concomitantly there is an increase in total nitrogen content of both

leaves and seeds (Castillo et al., 2000).

Conversely, increasing bacteroid ammonium assimilation has been shown to

have detrimental effects on symbiosis. In S. meliloti mutation of glutamate

synthase (GOGAT) abolishes the ability to grow on ammonium but does not alter

levels of GDH which remain insufficient to allow growth on ammonium. The

mutation does not alter the ability to form nodules or fix nitrogen. A revertant, with

sufficient (x30 increase) GDH activity to allow NH3 assimilation, has been isolated

which has reduced nitrogenase activity in the nodule (Osburne & Signer, 1980).

Bacteroids recovered from nodules formed by the revertant have the same levels

61

of GDH and GOGAT as the original mutant. This indicates that increased GDH

activity in S. meliloti is incompatible with symbiotic nitrogen fixation (Osburne &

Signer, 1980). Furthermore constitutively expressing a foreign glutamate

dehydrogenase gene (gdhA from E. coli) from a plasmid in R. phaseoli is also

incompatible with symbiotic nitrogen fixation. Plants inoculated with the GDH+

strain were only nodulated by bacteria that had lost the plasmid (Bravo et al.,

1988). The presence of gdhA reduces the induction of nod genes, which inhibits

synthesis of nod factors and hence nodulation (Mendoza et al., 1995). This

inhibition of nodulation is overcome when gdhA expression is controlled by NifA

thus delaying expression of GDH activity until after establishment of the nodule

(Mendoza et al., 1998). Therefore it appears that when a high intracellular amino

acid pool accumulates prior to or during nodule formation, due to the high

constitutive activity of GDH, nod factor production is downregulated and

consequently nodulation is inhibited (Mendoza et al., 1995). In bacteroids where

GDH activity is overexpressed, the partitioning of ammonium is modified such

that there is an increase in the bacteroid glutamate and amino acid pools, and a

decrease in ammonium exported to the plant. As a consequence the ureide

content of xylem sap is dramatically reduced (Mendoza et al., 1995).

1.5.2 Bacteroid alanine metabolism

Salminen & Streeter (1992) also showed that aspartate and alanine are the next

most rapidly labelled compounds after glutamate in bacteroids isolated from

intact whole nodules of both pea and soybean (Salminen & Streeter, 1992). In

pea nodules alanine was the most highly labelled amino acid in the first three

minutes but glutamate was the most highly labelled after 6 min suggesting rapid

62

turnover of the alanine pool (Salminen & Streeter, 1992). Similarly when isolated

symbiosomes of pea are incubated with 14C malate for 30 minutes, 12% of the

label recovered in the incubation medium occurs in the amino acid fraction.

Approximately 60% of this total amino acid is alanine, and 40% is aspartate

(Rosendahl et al., 1992). The authors suggested that the labelled malate is

converted to aspartate and alanine from transamination of amino donors such as

glutamate in the bacteroid. Alanine and aspartate excretion has also been

detected from isolated pea bacteroids after the addition of malate or α-

ketoglutarate and glutamate (Appels & Haaker, 1991). These studies were

considered evidence that some sort of amino acid shuttle mechanism operated

across the symbiotic membranes. Kahn et al. (1985) proposed the operation of a

bacteroid malate-aspartate shuttle similar to the shuttle that operates to transfer

reducing equivalents into mitochondria. In the model, malate and glutamate are

imported into the bacteroid in exchange for α-ketoglutarate and aspartate

respectively. Malate is taken up by the bacteroid and oxidised to oxaloacetate

and then transaminated to aspartate by a glutamate dependant aspartate

aminotransferase. Aspartate is exported to the plant cytosol where it

transaminates α-ketoglutarate to yield glutamate, which is taken up by the

bacteroid. The shuttle would result in a net transfer of NADH into the bacteroid

without a net transfer of carbon so could not support growth, but could support N2

fixation (Kahn et al., 1985). However, closer scrutiny of the labelling data of

Rosendahl (1992) shows the operation of the shuttle cannot occur in the pea

symbiosis. If the PBM posses a malate/aspartate shuttle the addition of

unlabelled glutamate to incubated symbiosomes would be expected to enhance

the transfer of 14C from the organic acid pool to the amino pool of the incubation

63

medium. Symbiosomes of pea incubated with 14C-malate accumulate 14C-

aspartate in the medium, and this accumulation does increase three-fold on

addition of unlabelled glutamate. The shuttle would also lead to the uptake of 14C-

glutamate into the symbiosome and its transamination to, and export of, 14C-α-

ketoglutarate in exchange for malate. However, symbiosomes incubated with 14C-

glutamate do not excrete 14C-α-ketoglutarate with or without the addition of

unlabelled malate (Rosendahl et al., 1992).

Furthermore even though bacteroids are able to transport glutamate (Salminen &

Streeter, 1987; Bergersen & Turner, 1988; Udvardi et al., 1988) and glutamate

has been shown to support nitrogen fixation in isolated bacteroids (Bergersen &

Turner, 1988; Kouchi & Fukai, 1988; Kouchi et al., 1991), the PBM of soybean,

bean and pea is impermeable to glutamate (Price et al., 1987; Udvardi et al.,

1988; Herrada et al., 1989; Ouyang & Day, 1992; Hernandez et al., 1996). Also

mutation of the general amino acid permease (Aap) of R. leguminosarum

prevents uptake of glutamate but there is no detrimental effect to the symbiosis

(Poole et al., 1985; Walshaw, 1995; Walshaw & Poole, 1996). These data are not

consistent with the operation of a shuttle system involving glutamate uptake.

Work using purified bacteroids of Rhizobium lupini suggests that aspartate and

alanine formed by bacteroids have a direct role in the assimilation of fixed

nitrogen. Bacteroids excrete aspartate and alanine when incubated with

dicarboxylates and when actively fixing nitrogen. The amount excreted correlated

with the rate of nitrogen fixation (acetylene reduction) and when nitrogen fixation

was prevented no aspartate or alanine excretion occured (Kretovich et al., 1986).

64

The possible significance of amino acid excretion as a mechanism for

incorporation of fixed nitrogen in the bacteroid and its subsequent transfer to the

plant was not fully appreciated until a 15N labeling study with purified B.

japonicum bacteroids showed that alanine was the sole excretion product of

nitrogen fixing bacteroids incubated in a sealed vessel with no removal of

products (Waters et al., 1998). This contradicts the reports of Bergersen who

showed amino acids are not excreted, and that ammonium is the sole nitrogen

excretion product, from B. japonicum bacteroids in flow chamber experiments

where there is constant removal of products (Bergersen & Turner, 1990; Li et al.,

2001). Waters et al. (1998) suggested that ammonium might have been detected

in other studies as the sole nitrogen secretion product because bacteroids were

contaminated with plant extract, which has alanine deaminase activity (Waters et

al., 1998). To examine these contradictions we have isolated bacteroids using

Percoll gradients, that produce clean bacteroid preparations with no associated

alanine deaminase activity, and looked at the excretion from pea bacteroids

(chapter 4). In data presented in chapter 4 and in Allaway et al. (2000) we have

shown that the disparity between these previous studies can be explained by the

relative concentration of ammonium experienced by the bacteroid. In summary,

purified pea bacteroids isolated anaerobically produce ammonium as the sole

excretion product when bacteroids are kept at a low to moderate density.

However as bacteroid density increases alanine synthesis is switched on due to

an accumulation of ammonium from nitrogen fixation. Therefore fundamental

differences in assay conditions between the studies of Bergersen (1990) and

Waters et al. (1998) are likely to have produced the differing results.

65

L-Alanine dehydrogenase catalyses the reversible NADH dependent reductive

amination of pyruvate to L-alanine (Fig 1.8). It has been suggested that it is

responsible for biosynthesis of alanine in bacteroids. In intact soybean nodules

labelled alanine showed a steady linear accumulation over 10 minutes in the

cytosol and bacteroid following exposure to 15N2 gas. The 15N labelling pattern

was independent of that of glutamine (which followed a hyperbolic curve)

indicating that alanine was produced in bacteroids by the direct incorporation of

fixed nitrogen into alanine by L-alanine dehydrogenase, and not via formation of

glutamine or glutamate then transamination to alanine (Ohyama & Kumazawa,

1980). The activity of L-alanine dehydrogenase in nodule tissue is exclusive to

the bacteroid cytoplasm (Dunn & Klucas, 1973). Its activity has been reported to

be high in soybean bacteroids (10-fold to100-fold higher than most other

bacteroid enzymes) (Stripf & Werner, 1978; Smith & Emerich, 1993). In B.

japonicum the specific activity in free-living nitrogenase active cells is higher than

in nitrogenase repressed cells and is similar to that in bacteroids (Werner &

Stripf, 1978). Activity from effective bacteroids is six-fold higher than that of

ineffective bacteroids and its activity increases during the period of nodule

development when nitrogenase also increases (Stripf & Werner, 1978; Werner et

al., 1980). These data suggest a role for L-alanine dehydrogenase during

nitrogen fixation.

66

Fig 1.8 Formation of L-alanine via L-alanine dehydrogenase

The apparent Km of the soybean L-alanine dehydrogenase for NADH, NH4+ and

pyruvate are 86 mM, 8.9 mM and 0.49 mM respectively (Smith & Emerich, 1993).

The Km for ammonium is much lower than that seen for other bacterial L-alanine

dehydrogenases, that have Km's in the range of 20-300 mM, and is within the

estimated concentration of ammonium (12mM) in soybean bacteroids (Streeter,

1989). We also show (chapter 4; Allaway, 2000) the Km for ammonium of L-

alanine dehydrogenase from R. leguminosarum is low (5.1 mM) consistent with

an aminating role for the enzyme in bacteroids.

Confirmation of the role of L-alanine dehydrogenase in alanine synthesis by

bacteroids requires the gene to be identified and mutated. In data presented in

chapter 4 and in Allaway et al. (2000) we have shown that mutation in L-alanine

dehydrogenase (aldA) of R. leguminosarum bv. viciae, which abolishes AldA

activity, prevents alanine synthesis and excretion by isolated bacteroids of pea.

Peas inoculated with the aldA mutant are Fix+ as measured by acetylene

reduction, although six week old plants show a small but significant (20%)

decrease in dry weight. This shows that alanine cannot be the sole nitrogen

COOH

H C NH2

CH3

L-ALANINE

COOH

C O

CH3

PYRUVATE

ALANINE DEHYDROGENASE NADH + H+ NAD+ NH4

+ H2O

67

excretion product from pea bacteroids, although the data are consistent with the

excretion of both alanine and ammonium. However, while a 20% decrease in dry

weight is consistent with labelling studies showing alanine excretion by isolated

bacteroids it can be argued that such a small drop in plant biomass may be

explained by an indirect effect on dicarboxylate metabolism (chapter 4).

The gene encoding L-alanine dehydrogenase (aldA) has been found to be

transcribed divergently from a transcriptional regulator (aldR) in R.

leguminosarum (chapter 5; Allaway, 2000). Mutation of either aldA or aldR

abolishes L-alanine dehydrogenase activity indicating that only one copy of each

gene is present in R. leguminosarum. This genetic organisation is the same in S.

meliloti. The aldA and aldR genes lie on the chromosome from positions 1760861

to 1759746 and 1761019 to 1761477 respectively

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). However,

in M. loti there are two copies of L-alanine dehydrogenase: one on the

chromosome which is associated with a regulator gene and one on a Sym

plasmid (Kaneko et al., 2000)

1.5.3 Bacteroid aspartate metabolism

Although the principal excreted amino acid detected from pea and soybean

bacteroids is alanine, aspartate has also been detected from pea peribacteroid

units (Rosendahl et al., 1992). The reason for this discrepancy is not clear

although it would be interesting if the presence of the peribacteroid membrane

were to alter the amino acids secreted, perhaps by allowing transamination to

occur in the peribacteroid space. Aspartate aminotransferase (AAT) catalyses the

reversible transamination of aspartate and glutamate to yield oxaloacetate (Fig

68

1.4). The activity of aspartate aminotransferase is more than 4 times higher in the

pea PBS (400 nmol min-1 mg protein-1) compared to the bacteroid cytosol (90

nmol min-1 mg protein-1) (Rosendahl et al., 2001). Moreover aspartate has been

shown to be taken up by inverted symbiosome vesicles (bacteroid side facing

out) against a concentration gradient, indicating a transport system capable of

transporting aspartate from the bacteroid side of the PBM to the plant cytosol is

present in pea (Rudbeck et al., 1999).

However, as stated previously, ‘naked’ bacteroids of R. lupini excrete both

aspartate and alanine suggesting both are synthesised in the cytoplasm of these

bacteroids (Kretovich et al., 1986). The importance of aspartate metabolism in

Rhizobium is highlighted by a mutant in aspartate aminotransferase (aatA) of S.

meliloti that forms ineffective nodules on alfalfa (Rastogi & Watson, 1991). This

symbiotic phenotype suggests that aspartate catabolism is essential for bacteroid

nitrogen fixation. The mutation in aatA reduces the aspartate aminotransferase

activity to 40% of the wild type. Other genes that encode aminotransferase

enzymes capable of aspartate utilisation have been identified such as tatA

(tyrosine (aromatic) aminotransferase) (Watson & Rastogi, 1993) and batA

(branched chain aminotransferase) (Alfano & Kahn, 1993). Aminotransferases

tend to have overlapping substrate specificity (Jensen & Calhoun, 1981) and the

mutation in aatA can be complemented by the aatA gene or by tatA, which shows

substantial aspartate aminotransferase activity (Rastogi & Watson, 1991; Watson

& Rastogi, 1993). A gene designated aatB encodes a second aspartate

aminotransferase in S. meliloti which shares sequence homology to AatA

(Watson & Rastogi, 1993). Mutants in the aatB and tatA genes form effective

nodules on alfalfa (Alfano & Kahn, 1993) implying that aatA serves a special role

69

or is the most active aspartate aminotransferase during symbiosis. A mutant in

aspartate aminotransferase of B. japonicum formed nodules on soybean that did

fix nitrogen, but plants went through a phase of yellowing, (Zlotnikov et al., 1984;

Streeter & Salminen, 1990) suggesting activity of this enzyme is important to the

soybean symbiosis but that alternative pathways can replace its activity.

Aspartate aminotransferase is important for aspartate and glutamate metabolism,

and is also important in converting TCA cycle intermediates to amino acids

therefore significantly linking carbon and nitrogen metabolism. Therefore it might

have a role in the regulation of amino acid and carbon pools in the bacteroid and

in some instances perturbation of these has a detrimental effect on nitrogen

fixation.

70

1.6 Research objectives

At the beginning of this research we were interested in carbon metabolism in R.

leguminosarum in general, particularly pathways in Rhizobium that might regulate

carbon metabolism within the nodule and subsequently affect the efficiency of

nitrogen fixation. Both carbon strorage compounds poly-β-hydroxybutyrate (PHB)

and glycogen may be formed by R. leguminosarum bacteroids. As there is no

defined role for PHB metabolism described in the literature and as such a role

might differ depending on the symbiosis (indeterminate or determinate) we have

looked at PHB metabolism. R. leguminosarum may form indeterminate nodules

on peas or determinate nodules on beans depending on the symbiotic plasmid

present (see section 1.1). Therefore we have made advantage of this and have

looked at PHB metabolism in a determinate (bean) versus indeterminate (pea)

symbiosis using isogenic strains of R. leguminosarum that differ only in symbiotic

plasmid (chapter 3). As the role of bacteroid glycogen metabolism is little studied

and is therefore not clear, we have made initial studies into its role in the R.

leguminosarum - pea symbiosis (chapter 3). In chapter 4 we have looked at

alanine metabolism in pea bacteroids. As the role of alanine synthesis and

excretion was unclear, we initially aimed to identify factors that led to the

synthesis and excretion of alanine from pea bacteroids. Subsequently we

questioned whether alanine was produced via the direct incorporation of fixed

nitrogen with pyruvate by L-alanine dehydrogenase (AldA). On finding that

alanine synthesis via AldA has a significant role in bacteroid metabolism we were

prompted to look at the regulation of L-alanine dehydrogenase (chapter 5).

71

CHAPTER 2

72

2 MATERIALS AND METHODS

2.1 General

All chemicals and reagents were from Sigma or BDH unless otherwise stated.

2.2 Culture conditions

R. leguminosarum strains were grown at 28°C on either Tryptone-Yeast media

(TY) (Beringer, 1974) supplemented with 6mM CaCl2, or on acid minimal salts

(AMS) which is derived from Rhizobium minimal salts (RMS) medium (Brown &

Dilworth, 1975) with the following adaptions: 0.5mM potassium phosphate, 2mM

MgSO4, 0.17mM CaCl2 and 20mM MOPS buffer (pH 7.0). R. tropici was grown

on TY or Y minimal medium (Sherwood, 1970). Minimal medium was

supplemented with carbon and nitrogen sources at 10mM unless otherwise

stated.

E.coli strains were grown at 37°C in Luria-Bertani broth (LB) consisting of

tryptone (10 g.l-1), yeast extract (5 g.l-1), NaCl (5 g.l-1). Agar (Difco, Bitek) was

added to the media as necessary (1.5%). Strains were routinely stored at -20°C

and -80°C in 15% glycerol after snap freezing in liquid nitrogen.

73

2.3 List of antibiotic concentrations

Media were supplemented with antibiotics or stains where appropriate at the

following concentrations:

E. coli Rhizobium

Antibiotics Concentration (µg/ml)

Ampicillin 50 -

Chloramphenicol 10 -

Gentamicin 25 20

Kanamycin 20 40

Streptomycin - 500

Spectinomycin 50 100

Tetracycline 10 2 (Minimal Media) 5 (TY)

Rifampicin - 20

Fungicide

Nyastatin - 50

Stains

IPTG 40 -

X-GAL 40 -

74

2.4 List of strains

Strain Description or Genotype Source or Reference

C600 E. coli supE44 (Sambrook et al., 1989)

DH5α E. coli supE44, recA1, endA1 (Sambrook et al., 1989)

MC1061 E. coli non suppressor strain for phage library (Sambrook et al., 1989)

TOP10 E. coli recA1, endA1 Invitrogen

CIAT899 Rhizobium tropici WT, rifr (Martinez et al., 1985)

A656 Rhizobium tropici ∆glgA, rifr (Marroqui et al., 2001)

3841 Rhizobium leguminosarum bv. viciae, strr (Glenn et al., 1980)

A5 Rhizobium leguminosarum (Sym plasmid pRL2JI – infects bean), rifr

Downie, pers. comm.

A31 Sym plasmid cured derivative of 8002, strr Downie, pers. comm.

A34 Sym plasmid pRL1JI derivative of A31 (infects pea), strr

Downie, pers. comm.

RU1275 3841 derived dadR::Tn5, strr, kanr This work

RU1327 3841 derived aldA::TnB20, strr, kanr This work

RU1328 A34 derived phaC::Ω, strr, spcr This work

RU1329 A5 derived phaC::Ω, rifr, spcr This work

RU1371 pRU708 (full length aldA) in RU1327, strr, kanr, tetr, ampr

This work

RU1414 pRU730 (AldR GUS fusion) in 3841, strr, tetr This work

RU1415 pRU731 (AldA GUS fusion) in 3841, strr, tetr This work

RU1416 pJP2 (GUS fusion vector) in 3841, strr, tetr This work

75

RU1422 3841 derived aldR:: Ω, strr, spcr This work

RU1448 A34 derived glgA::TnB60, strr, kanr This work

76

2.5 List of plasmids, cosmids and phage

Plasmid, cosmid or phage

Description or Genotype Source or Reference

λTn5 λ carrying Tn5 transposon (Simon et al., 1989)

λTnB20 λ carrying transposable lacZ promoter probe (Tn5 derivative)

(Simon et al., 1989)

λTnB60 λ carrying transposable tac-promoter (Tn5 derivative)

(Simon et al., 1989)

pBluescript® II SK- pUC19 derivative, f1 origin, ColE1 replicon, ampr, lacZ

Stratagene

pBC SK+ pUC19 derivative, f1 origin, ColE1 replicon, camr, lacZ

Stratagene

pCR®2.1-TOPO TA PCR cloning vector, f1 origin, ColE1 replicon, ampr, kanr, lacZ

Invitrogen

pJQ200SK P15A origin from pACYC184, genr, lacZ, sacB, traJ

(Quandt & Hynes, 1993)

pLAFR1 Wide host range P-group cloning vector, RK2 derivative, tetr

(Freidman et al., 1982)

pTR101 Wide host range P-group cloning vector, RK2 derivative, tetr

(Weinstein et al., 1992)

pJP2 pTR101 derivative, uidA, tetr Boesten, pers. comm.

pRK2013 RK2 transfer genes for mobilisation of P-group plasmids, kanr

(Figurski & Helinski, 1979)

pPHJI1 P-group chaser plasmid, genr (Ruvkun & Ausubel, 1981)

pHP45Ω pBR322 derivative carrying Ω, pHP45 replicon, ampr, spcr

(Prentki & Krisch, 1984)

pIJ7843 0.7 Kb EcoRI/XhoI fragment of glgA from R. tropici in pKS, ampr

Marroqui, pers. comm.

pRU99 8.9 Kb Tn5 bearing EcoRI fragment of RU137 in pSK-, kanr, ampr

(Walshaw, 1995)

77

pRU100 HindIII digestion/re-ligation of pRU99 carrying IS50R insertion sequence of Tn5, ampr

(Walshaw, 1995)

pRU101 3.2 Kb BamHI fragment from pRU99 in pBCSK+ , kanr, camr

(Walshaw, 1995)

pRU475 BamHI digestion/re-ligation of pRU99 carrying IS50L insertion sequence of Tn5, ampr

This work

pRU493 3 Kb HindIII fragment from pRU99 in pBCSK+, camr

This work

pRU549 Tn5 bearing EcoRI fragment of RU1275 in pBCSK+, kanr, camr

This work

pRU566 P138/P166 1.6 Kb PCR fragment of phaC in pCR®2.1-TOPO, ampr, kanr

This work

pRU575 Ω cloned in pRU566 (phaC), ampr, kanr, spcr

This work

pRU577 phaC::Ω in pJQ200SK, genr, spcr This work

pRU593 6 Kb EcoRI transposon subclone of pRU3138 in pSK-, kanr, ampr

This work

pRU626 EcoRI transposon subclone of pRU3146 in pSK-, kanr, ampr

This work

pRU636 EcoRI digestion/re-ligation of pRU626, kanr, ampr

This work

pRU638 1.5 Kb HindIII fragment from pRU636 carrying IS50R insertion sequence from TnB60 in pSK-, ampr

This work

pRU640 P199/P200 1.45 Kb PCR product of aldA in pCR®2.1-TOPO, ampr, kanr

This work

pRU679 P219/P220 700 bp PCR product of aldA/aldR intergenic region in pCR®2.1-TOPO, kanr, ampr

This work

pRU693 3 Kb aldA/aldR SalI fragment from pRU3135 in pSK-, ampr

This work

78

pRU701 P219/P220 700 bp PCR product of aldA/aldR intergenic region in pSK-, ampr

This work

pRU708 1.45 Kb SacI/KpnI fragment from pRU640 in pTR101, tetr, ampr

This work

pRU720 P139/P234 1.8 Kb PCR product of phaC in pCR®2.1-TOPO, ampr, kanr

This work

pRU730 AldR GUS fusion (pJP2), tetr This work

pRU731 AldA GUS fusion (pJP2), tetr This work

pRU734 Ω cloned in pRU693 (aldR), ampr, spcr

This work

pRU735 aldR:: Ω cloned in pJQ200SK, genr, spcr

This work

pRU758 glgA::TnB60 SacI fragment in pJQ200SK, genr, kanr

This work

pIJ9019 cosmid from gene bank of 8002 which complements A656 (glgA-), tetr

Downie, pers. comm.

pRU3131 Dad cosmid from 3841 gene bank that complements RU1275 (dadR-), tetr

This work

pRU3135 Ald cosmid from 3841 gene bank that complements (suppresses) RU1275 (dadR-), tetr

This work

pRU3138 aldA::TnB20 cosmid (parent pRU3135), tetr, kanr

This work

pRU3146 glgA::TnB60 cosmid (parent pIJ9019), tetr, kanr

This work

79

2.6 List of primers

Name Priming site Sequence 5’ – 3’ Cy5 labelled

M13 Forward

universal vector CGACGTTGTAAAACGACGGCCAGT yes

M13 Reverse

universal vector CAGGAAACAGCTATGAC yes

P113 IS50R AGGTCACATGGAAGTCAGATC yes

P120 IS50R TTGATTTACCAGAATATTTTGCC

P132 extends phaC sequence

GTGAAGAGTGTCGCGGTTTT yes

P137 extends phaC sequence

CTCGGCCAGCATCTTCAT yes

P138 phaC C-terminus CTGCCGAAGAAGACGGAC

P139 phaC N-terminus CGCTCGAAAATCTCGGAC

P151 extends phaC sequence

GAGGCTGCTTGTCCGAGAT yes

P152 extends phaC sequence

TTCGAGGGATAGTTGAGGAT yes

P166 phaC N-terminus GGATAAAAAGAGTGGTGCTG

P199 aldA C-terminus ATACAAAGAAGGCGGCATCC

P200 aldA N-terminus AGCTCGGCGTTGGTGATGC

P219 aldR N-terminus GAGCGCCTTGTGTGAAAGCC

P220 aldA N-terminus TCCGGCGCCAGATGCAGATAG

P234 C terminus of phaC: from R.etli sequence

TCGCGAGGATGACGAATTAT

80

2.7 DNA manipulation

2.7.1 Agarose gel electrophoresis

DNA was separated on 0.8% agarose (Gibco BRL) gels in x1 Tris Acetate

(40mM) EDTA (1mM) (TAE) buffer at 80 – 100volts. A solution of 30% glycerol,

0.25% bromophenol blue was used as DNA loading buffer (x6). A 1 Kb ladder

(Gibco BRL) was used as a size marker and a high DNA mass ladder (Gibco

BRL) was used if the DNA was to be quantified. Gels were stained in ethidium

bromide solution (0.5µg/ml) and the DNA was visualised under UV. Gel images

were scanned into Grab-IT 2.0 image capture programme and stored as bitmap

files (*.bmp).

2.7.2 DNA isolation

Plasmid and cosmid DNA was isolated using the Pharmacia Flexiprep Kit

following the manufacturers protocols. Chromosomal DNA was isolated using the

BioLine DNAce Clini Pure kit following the protocol for isolation of DNA from

whole blood.

2.7.3 Restriction digests

DNA was digested using restriction enzymes from Gibco BRL or New England

Biolabs following the manufacturers specifications. Restriction enzyme activity

was removed from the DNA sample prior to ligation by heat inactivation or by

using the Qaigen Qaiex II Gel Extraction Kit (following manufacturers protocol for

desalting DNA solutions).

81

2.7.4 Partial restriction digests

Partial digests were carried out in 150µl volumes using 1/10th normal enzyme

concentration, incubated for 5, 10 and 20 minutes at 37°C and immediately heat

inactivated at 65°C for 25 minutes. All 3 digests were separated on an agarose

gel and the appropriate linear bands were excised with a clean scalpel blade and

DNA extracted using the Qaiex II Gel Extraction Kit (Qaigen) following

manufacturers protocol.

2.7.5 Fill in of overhangs

5’ overhangs were filled in using 0.5U Klenow (Gibco BRL), 20µM dNTPs, 1x

React2 buffer at room temperature for 15 minutes. The enzyme was heat

inactivated at 75°C for 20 minutes.

2.7.6 Ligation

DNA was ligated using T4 DNA Ligase (Promega) following the manufacturers

specifications. Reactions were carried out overnight at 16°C.

2.7.7 Transformation

Plasmid and cosmid DNA was routinely transformed into competant cells of E.

coli strain DH5α. Cells were grown in LB to OD600nm 0.3 - 0.4, chilled on ice for 20

minutes, washed in cold 0.1M CaCl2 then resuspended and concentrated (x20) in

cold 0.1M CaCl2. Competant DH5α cells were routinely made in batches and

stored at -80°C in 15% glycerol after snap freezing in liquid nitrogen.

82

DNA was incubated with the competant cells on ice for 30 – 60 minutes, heat

shocked at 42°C for 90 seconds, incubated on ice for a further 2 minutes then

heat shocked a second time by the addition of LB at 42°C to a final volume of 1

ml. Cells were incubated for 1hr at 37°C with shaking at 250rpm then plated onto

selective media.

For higher efficiency transformation, DNA was transformed into XL2-blue

ultracompetant cells (Stratagene) following the manufacturers protocol.

2.7.8 Southern blot & hybridisation

DNA was transferred from agarose gels to positively charged nylon membranes

(Hybond-N+) as recommended by the manufacturers (Amersham Pharmacia

Biotech). The gels were soaked in 250mM HCl for 15 minutes to depurinate the

DNA and rinsed in water. DNA was denatured by soaking the gel in a solution of

1.5M NaCl, 0.5M NaOH for 30 minutes. The gel was neutralised in 1.5M NaCl,

0.5M Tris HCl (pH7.5) then blotted overnight using x20 SSC (0.3M sodium

citrate, 3M NaCl, pH7-8). The membrane was rinsed in x2 SSC, air dried, and the

DNA was fixed onto the membrane by a 3 minute exposure to UV light.

The membrane was pre-hybridised for 1 hr at 65°C in a solution of x5 SSPE

(0.9M NaCl, 0.05M Sodium phosphate, 0.005M EDTA pH 7.7), x5 Denhardt’s

(0.1% Bovine serum albumin, 0.1% Ficoll, 0.1% Polyvinylpyrrolidine) and 0.5%

SDS, containing non-homologous denatured salmon sperm (1mg/ml). The Gene

Images™ Random Prime Labelling Module (Amersham Pharmacia Biotech) was

used to label the probe DNA following the manufacturers protocol. The probe was

added to the pre-hydridisation solution and incubated at 65°C for 12 hrs. Both

83

pre-hybridisation and hybridisation steps were carried out in glass cylinders on

rollers inside a hybridisation oven (Techne Hybridiser HB-1).

Following hybridisation, the membrane was washed as follows:

2x SSPE, 0.1% SDS at room temperature for 10 minutes

1x SSPE, 0.1% SDS at 65°C for 15 minutes

0.1x SSPE, 0.1% SDS at 65°C for 10 minutes (repeated)

After the washes the probe was detected using the CDP-star™ Detection Module

(Amersham Pharmacia Biotech) following the manufacturers protocol. The blot

was exposed to Hyperfilm™ MP (Amersham Pharmacia Biotech), developed

using an automated developer (Agfa), then the image was scanned and saved as

a bitmap file (*.bmp).

2.7.9 Quantitation of DNA

The purity and concentration of DNA was determined by measuring the

absorbance at 260nm and 280nm. The ratio of OD260nm / OD280nm was used as an

indication of quality (samples with values of between 1.8 and 2.0 were

considered pure). The OD260nm was used to estimate DNA concentration (OD260nm

of 1.0 was equivalent to 50µg/ml DNA).

2.7.10 Polymerase Chain Reaction (PCR)

PCR primers (see 2.6) were designed using the Wisconsin GCG Package

(Version 8) or in Vector NTI 5.5 and obtained from Genosys. DNA was amplified

from Rhizobium by PCR using 2U Bio-X-Act enzyme in x1 Optibuffer (Bioline);

84

1.5mM MgCl2; 0.2mM of each dNTP and 50pmol of each primer in a final reaction

volume of 50µl. DNA template was provided at 1ng µl-1. Thermal cycler (Hybaid

Omnigene) conditions were 95°C for 5 minutes, followed by 30 cycles of 95°C for

1.5 minutes, 57°C - 59°C for 1.5 minute, 72°C for 1 minutes (per Kb of DNA),

followed by a final 10 minutes extension at 72°C. PCR products were visualised

on an agarose gel and cloned using the vector pCR®2.1-TOPO (Invitrogen)

following the manufacturers protocol.

2.7.11 Sequencing & Sequence Analysis

Sequencing of plasmids was carried out in a Amersham Pharmacia Biotech. ALF

Express automated DNA sequencer using a cycle sequencing programme at the

AMSEQ Sequencing Unit, University of Reading. 1µg of DNA was provided for

sequencing reactions. Primers were designed using Oligo 4.0 or Vector NTI 5.5

and were obtained Cy5 labelled from Genosys or MWG-Biotech AG (see 2.6).

Sequences were processed and analysed by ALFwin™ Software (Version 2.0,

Amersham Pharmacia Biotech). Cosmids were sequenced by MWG-Biotech AG.

The sequences of primers used by MWG-Biotech AG were not disclosed.

Homology searches were made using the BLAST program from the ExPASy

Molecular Biology web site (http://expasy.hcuge.ch/) or from the National Center

for Biotechnology Information web site (http://www.ncbi.nlm.nih.gov/). DNA

sequences were analysed for restriction sites and putative coding regions using

the GCG package at http://www.hgmp.mrc.ac.uk or Vector NTI 5.5. Comparisons

of DNA and amino acid sequences were made using GCG, BLAST or Vector NTI

5.5.

85

2.8 Transposon mutagenesis

2.8.1 Phage propagation

Lambda (λ) phage containing the transposon was propagated in E. coli strain

C600 to obtain a titre of 1010 or above before carrying out mutagenesis. E. coli

was grown in 10 ml LB containing 10mM MgSO4 overnight at 37°C with shaking

at 225rpm. 100µl of this culture was sub cultured into 10 ml LB 10 mM MgSO4

and grown for 3 - 4 hrs. Phage was serially diluted from 10-2 to 10-7 in SM buffer

(0.1M NaCl, 0.008M MgSO4.7H2O, 0.001% gelatin, 0.05M Tris HCl, pH 7.5). 0.9

ml of bacterial culture was mixed with 0.1 ml of each phage dilution in 4 ml of

0.75% LA containing 10mM MgSO4 (top agar) and spread on 1.5% LA containing

10mM MgSO4 (bottom agar) in a petri dish. Minus phage and minus bacteria

controls were made up in top agar and spread onto bottom agar. The plates were

incubated at 37°C overnight. Confluent plates were flooded with 10 ml SM and

rocked for 1 hour. The supernatant was pipetted from the plate, filtered through a

sterile millipore membrane and stored containing 2 drops of chloroform at 5°C.

This phage solution was titred by carrying out the above procedure to overnight

incubation. The number of plaques per plate was used to determine the number

of plaque forming units per ml.

2.8.2 Mutagenesis

Cosmids were transferred into a non supressor strain of E. coli (MC1061) to carry

out mutagenesis. Cosmids were mutagenised essentially as described by Simon

et al. (1989). Cultures of MC1061 containing pIJ9019 was grown in 10 ml LB

containing 10mM MgSO4 to mid log stage. The phage was diluted to 10-2, 10-3,

86

10-4, 10-5, 10-6. 0.9 ml culture was mixed with 0.1 ml of each phage dilution and

incubated at 37°C for 30 minutes without agitation then for a further 90 minutes

with gentle shaking at 125rpm. Cultures were diluted to 10-1, 10-2, 10-4. Bacteria-

phage suspensions were plated onto LA agar containing kanamycin and

tetracycline and incubated overnight at 37°C. 10,000 - 20,000 colonies were

bulked together in 2 ml LB and stored in 15% glycerol at -20°C prior to purifying

and screening mutagenised cosmids.

87

2.9 Conjugation

Conjugations to transfer DNA into Rhizobium were done using the helper plasmid

pRK2013 (kanr) (Figurski & Helinski, 1979). Cultures of donor plasmid and helper

plasmid were grown overnight in 10 ml LB. 0.2 ml of the overnight culture was

sub-cultured into 10 ml fresh LB and incubated with shaking at 100rpm for a

further 2 – 3 hrs until the OD600nm was 0.2 – 0.6. 1 ml of each donor and helper

plasmid was harvested and washed twice in TY to remove any traces of

antibiotics then resuspended in 1 ml TY. Rhizobium was washed off fresh TY

slopes with 3 ml TY. 400µl of donor plasmid, 400µl Rhizobium and 200µl

pRK2013 were mixed, pelleted and resuspended in 30µl TY. The resuspended

culture was transferred to a sterile filter placed on a TY plate and incubated

overnight at 25°C. Bacteria were washed off the filter with TY and streaked, or a

serial dilution was plated, onto selective media.

88

2.10 RNA manipulation

To minimise the contamination with RNAses, all RNA work was carried out using

γ-irradiated disposable plasticware and glassware that had been thoroughly

cleaned and oven-baked at 200°C overnight. Distilled water and solutions (except

those containing Tris) were treated with DEPC to inactivate RNAses. DEPC was

added to solutions at 0.1%, allowed to stand for 12hrs at 37°C then autoclaved

twice to remove all trace of DEPC. Tris-buffered solutions were made up with

DEPC treated water. Electrophoresis tanks and any other equipment required

was cleaned with RNaseZAP™ (Invitrogen).

2.10.1 RNA isolation

RNA was isolated from Rhizobium grown in 50 ml AMS to OD600nm 0.2 – 0.8

using the Qaigen RNeasy® Midi Kit following the protocol for bacteria. The lysis

step was modified by incubating cells in 10mM Tris (pH 8) with 20% sucrose and

1mg/ml lysozyme at room temperature for 15 minutes followed by a further 20

minute incubation with EDTA at 1mM.

2.10.2 Quantitation of RNA

The purity and concentration of RNA was determined by measuring the

absorbance at 260nm and 280nm. The ratio of OD260nm / OD280nm was used as an

indication of quality (samples with values of between 1.8 and 2.0 were

considered pure). The OD260nm was used to estimate RNA concentration (OD260nm

of 1.0 was equivalent to 40µg/ml RNA).

89

2.10.3 RNA electrophoresis

RNA was separated on a formaldehyde agarose (FA) gel (1.2% agarose, 0.65%

formaldehyde, 0.1µg/ml ethidium bromide) in x1 FA gel running buffer (20mM

MOPS, 5mM sodium acetate, 1mM EDTA, 0.74% formaldehyde, pH 7.0). The FA

gel was equilibriated in running buffer for 30 minutes prior to electrophoresis. 1

volume of x5 RNA loading buffer (see below) was added to 4 volumes of RNA

sample, incubated at 65°C for 3 minutes then chilled on ice before loading onto

the gel. Gels were run at 70 volts.

x5 RNA loading buffer

16 µl saturated bromophenol blue solution

80 µl 500mM EDTA, pH 8.0

720 µl 12.3M formaldehyde

2 ml 100% glycerol

3084 µl formamide

4 ml x10 FA gel buffer

RNase-free water to 10 ml

2.10.4 Northern blot & hybridisation

RNA was transferred from FA gels to positively charged nylon membranes

(Hybond-N+) as described by the manufacturers (Amersham Pharmacia Biotech).

The membrane was air dried and the RNA was fixed onto the membrane by a 3

minute exposure to UV light.

The membrane was pre-hybridised in x5 SSPE, x5 Denhardt’s solution, 0.5%

SDS, with 1mg/ml non-homologous denatured calf thymus DNA (Sigma) at 62°C

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for 4 hrs. The DNA probe was labelled with [32P]dCTP using Ready To Go™ DNA

labelling beads and purified using ProbeQuant™ G-50 Micro Columns following

the manufacturers protocols (both Amersham Pharmacia Biotech). Probe was

added to the pre-hybridizing solution and the membrane was incubated at 62°C

for 12 hrs. Both pre-hybridisation and hybridisation steps were carried out in

glass cylinders on rollers inside a hybridisation oven (Techne Hybridiser HB-1).

Following hybridisation, the membrane was washed as for DNA blots (see

section 2.7.8) then exposed to a Phosphor Screen for approximately 48hrs. The

screen was scanned using a phosphor imager (Molecular Dynamics Phosphor

Imager TMSI) and analysed by Image Quant (version 5.1) software. The relative

intensities of hybridisation was measured using densitometer software

(Amersham Pharmacia Biotech, Total Lab version 1.00).

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2.11 Assay of β-glucuronidase (GUS) activity

Cells were grown overnight in 10 ml AMS containing the appropriate carbon and

nitrogen sources and antibiotics in universals. 1.5 ml cells were harvested at

OD600nm 0.2 – 0.8 and spun down at 6,500 rpm for 5mins. Cells were

resuspended in 1.5 ml Z buffer (0.06M Na2HPO4, 0.04M NaH2PO4, 0.01M KCl,

0.001M MgSO4.7H2O, pH 7.0). In duplicate eppendorfs, 330µl cells were

combined with 300µl Z buffer and 70µl lysozyme solution (0.05g lysozyme, 350µl

mercaptoethanol in 10 ml of 10mM phosphate buffer (pH 7.8) giving a final

reaction concentration of 0.5mg/ml lysozyme). Samples were incubated at room

temperature for 5 minutes. The remainder of the resuspended cells were used to

read their OD600nm. 15µl of 0.5M EDTA (pH8.0) was added and incubated for 15

minutes. 7µl of 1% SDS (final concentration of 0.01%) was added and samples

were equilibrated at 30°C for 5 minutes. 140µl of ρ-nitrophenyl β-D-galacto

pyranoside (PNPG) solution (0.08g PNPG, 70µl mercaptoethanol in 20 ml of Z

buffer) was added and samples were incubated for 10min or until a yellow colour

developed. The reaction was stopped by adding 350µl of 1M Na2CO3. Samples

were centrifuged at 13,000rpm for 30 minutes to pellet fine cell debris. The

OD420nm of the supernatant was read.

The extinction co-efficient of 4.012 x 103 l mol-1 cm-1 was used to calculate the

rate at which ρ-nitrophenol was hydrolysed from PNPG by β-glucuronidase

(GUS).

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2.12 PHB quantification

PHB content was determined in free-living cultures of R. leguminosarum grown in

50 ml AMS (10mM fructose/10mM NH4Cl) to an OD600nm of approximately 1.0.

Cells were washed in RMS (section 2.2) and resuspended in 5 ml of technical

grade sodium hypochlorite. Suspensions were incubated at 37°C for 1 hour. The

solid fraction from 1 ml of sample was pelleted by centrifugation at 13,000rpm for

30 minutes at 4°C. The pellet was washed with H2O, acetone and ethanol. The

polymer was extracted from the pellet by dissolving in three changes of 100µl

boiling chloroform (in a water bath at 61°C). The chloroform containing the

dissolved PHB was boiled to evaporate off the chloroform. Polymer was heated

to 100°C for 10 minutes in 3 ml of concentrated H2SO4. This produced crotonic

acid which was quantified by measuring its absorbance at 235nm, using

concentrated H2SO4 as blank. The extinction coefficient of crotonic acid (1.56 x

104 l mol-1 cm-1) was used to determine the quantity of PHB (Law & Slepecky,

1961).

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2.13 Glycogen quantification

2.13.1 Extraction of glycogen from free-living cells

Glycogen content was determined in free-living cultures of R. leguminosarum

grown overnight in AMS (10mM glucose/ 10 mM NH4Cl) to OD to an OD600nm of

0.2 – 0.8. Cells were washed and resuspended in water to OD600nm1.0. Cells

were hydrolysed by combining 50µl cells with 200µl KOH (30% w/v) and heating

in a water bath at 100°C for 90 minutes. Glycogen was precipitated by adding

600µl ice cold absolute ethanol to the cooled samples. Samples were incubated

on ice for 1 – 2 hrs prior to collecting the glycogen by centrifugation (5 minutes at

13,000rpm). The pellet was washed twice in ethanol then dried for 10 minutes at

60°C. The glycogen pellet was resuspended in 300µl 100mM acetate buffer (pH

4.75).

2.13.2 Extraction of glycogen from bacteroids

Bacteroids were crudely purified from plant tissue by macerating 2g (wet weight)

nodules in 5 ml 100mM phosphate buffer (pH 7). The breis was strained through

muslin then centrifuged at 250 x G for 5 minutes to pellet the plant material. The

supernatant was removed and bacteroids were pelleted by a second spin at 3000

x G for 10 minutes. The pellet was resuspended in 1 ml buffer and 50µl of this

was used to extract glycogen as section 2.13.

2.13.3 Digestion of glycogen to glucose

Glycogen was digested in duplicate (plus undigested control) with

amyloglucosidase (4units/assay) and amylase (8units/assay) for 1 hr at room

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temperature or 25 minutes at 55°C. Commercially bought glycogen (Sigma) was

digested for a standard curve (0-200µg). Insoluble membrane fragments were

removed by centrifugation and the supernatant was used in to determine the

glucose content.

2.13.4 Determination of glucose formed

100µl supernatant was assayed by a standard hexokinase/glucose-6-phosphate

dehydrogenase assay coupled to NADP+ reduction in 1 ml reaction volumes in

duplicate together with a control containing no G6P-DH/HK. Final concentrations

of buffer, cofactor, substrates, and enzymes were: 256mM Triethanolamine buffer

(pH 7.6), 3.3mM MgCl2, 1mM ATP, 1mM NADP+, 0.5U G6P-DH/2.8U HK

(Boerhinger Mannheim). Samples were incubated for 90 minutes at 37°C.

Glycogen content was assessed by comparing OD340nm values for the sample

with those for the standard.

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2.14 Enzyme assays

2.14.1 Preparation of protein extracts

Rhizobium was grown in 400 ml AMS with the appropriate carbon and nitrogen

source to OD600nm 0.2 – 0.8. Cells were pelleted at 4°C and washed twice in cold

10mM Hepes (pH 7.4) then resuspended in 12 ml of cold 40mM Hepes (pH 7.4)

with 2mM DTT and 20% glycerol. Cells were broken open using a French®

pressure cell press (SLM Instruments Inc.) at 1000 psi. The lysed cells were spun

at 15,000rpm (Sorvall SS-34) for 30 minutes to pellet cell membranes and debris.

The cytosolic fraction was kept on ice prior to enzyme assays.

2.14.2 Assays of malate dehydrogenase, citrate synthase, isocitrate dehydrogenase and L-alanine dehydrogenase

L-Alanine dehydrogenase, malate dehydrogenase and isocitrate dehydrogenase

activities were assayed at 28°C by measuring the change in absorbance at

340nm due to oxidation of NADH or reduction of NAD. The enzyme rate was

monitored in reaction volumes of 1 ml inside glass cuvettes. Reactions were

initiated upon addition of enzyme substrate. Final concentrations of substrates,

cofactors and buffers were as follows:

Malate dehydrogenase 50mM sodium phosphate buffer (pH 7.8); 0.2mM NADH &

0.5mM oxaloacetic acid.

Isocitrate dehydrogenase 20mM Tris-HCl buffer (pH 7.5); 2mM MnCl2; 0.5mM

NADP & 0.5mM isocitrate.

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L-Alanine dehydrogenase 50mM Tris-HCl buffer (pH8.5); 0.2mM NADH; 5mM

pyruvate & 100mM NH4Cl.

Citrate synthase activity was assayed at 28°C by measuring the change in

absorbance at 412nm due production of mercaptide. Final concentrations of

reactants were:

Citrate synthase 200mM Tris-HCl buffer (pH 8.1); 0.2mM DTNB; 0.1mM Acetyl-

CoA; 0.5mM oxaloacetic acid.

Activities were calculated from the initial linear rates (NADH extinction coefficient

= 6.22x103 l mol-1 cm-1 and mercaptide ion extinction coefficient = 1.36x104 l mol-1

cm-1).

2.15 Protein Determination

The total protein concentration for enzyme extracts, free-living cells and

bacteroids was determined by the Lowry-Folin method (1951) using BSA as

standard (0 - 100µg range) (Lowry et al., 1951).

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2.16 Plant Experiments

2.16.1 Seed sterilisation, sowing and inoculation

To prevent nodulation by other Rhizobia, peas seeds (cv. Avola) and bean seeds

(cv. Tendergreen) were surface sterilised before sowing. Seeds were washed in

70% ethanol for 30 seconds, rinsed twice in sterile water, washed in 2% Na-

hypochlorite for 8 minutes (peas) or 4 minutes (beans), then rinsed in 5 changes

of sterile water to remove the hypochlorite.

Seeds were sown in sterile vermiculite contained in 2L beakers for studies in the

plant growth room (3 seeds were sown and thinned to 1 seedlings after

germination) or in a sterile mixture of sand, gravel and vermiculite (4:2:2) in 15L

pots for studies in the greenhouse (12 seeds were sown and thinned to 7

seedlings after germination). In each case the growth medium was wetted and

plants were subsequently watered with nitrogen free rooting solution (see over).

Plants in the growth room were grown at a constant temperature of 22°C with a

16 hour light cycle.

To inoculate seeds, cultures of Rhizobium were grown over night in TY broth.

Culture was harvested at an OD600nm of 0.5-0.7 and the cells were washed with

sterile water to remove any remaining nitrogen compounds. Cells were

resuspended to approximately OD600nm 0.5 and seeds were inoculated with 1 ml

of this cell suspension at the time of sowing.

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N-Free Rooting Solution

100µM KH2PO4

100µM Na2HPO4

1mM CaCl2.2H20

100µM KCl

800µM MgSO4.7H2O

10µM Fe EDTA

35µM H3BO3

9µM MnCl2.4H2O

0.8µM ZnCl2

0.5µM Na2MoO4.2H2O

0.3µM CuSO4.5H2O

2.16.2 Assay of nitrogenase activity (acetylene reduction)

For acetylene reduction assays, plants were harvested at flowering during peak

nitrogen fixation. Plants were placed in sealed glass vessels lined with moist

tissue to prevent the plant from drying during the course of the experiment (3

plants per 1 litre kilner jar or 1 plant per 250 ml schott bottle sealed with

neoprene). 5% of the jars volume of air was removed with a syringe and replaced

with the same volume of acetylene. Samples of gas were taken from the vessels

at 60 minutes using a 1 ml syringe and injected onto a gas chromatogram

(Column temperature 30°C; injector temperature 45oC; FID detector temperature

120oC) (Varian STAR 3400cx). The acetylene reduction rate was calculated using

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the ratio of ethylene to acetylene (and assuming 1 mole acetylene occupies

22.41 litres at STP).

2.16.3 Plant dry weights

Aerial parts of plants were dried in an oven at 75°C for 48 hrs before weighing.

2.16.4 Determination of nodulating strain

Nodules were picked from roots then surface sterilised with 2% Na-hypochlorite

for 5 minutes and rinsed in 5 changes of sterile water. Nodules were transferred

into the wells of microtitre plates (1 nodule per well) and crushed with a sterile

metal rod to release the bacteria. These were then patch plated onto selective

media.

2.16.5 Microscopy of nodule sections

Nodules were cut in half longitudinally and placed in a solution of 2.5% (v/v)

glutaraldehyde in 0.05M sodium cacodylate (pH 7.3) and briefly vacuum

infiltrated until they sank. The fixative was replaced with fresh and left overnight

to adequately fix all the cells. The fixative was washed out by three successive

10 minute washes in 0.05M sodium cacodylate and the nodules were post-fixed

in 1% (w/v) OsO4 in 0.05M sodium cacodylate for one hour at room temperature.

The osmium fixation was followed by three, 10 minute washes in distilled water

before beginning the ethanol dehydration series (30%, 50%, 70%, and 95% each

for about 20 minutes, then 100% ethanol for an hour). Once dehydrated, the

samples were gradually infiltrated with LR White resin (London Resin Company)

by successive changes of resin:ethanol mixes over approximately 24 hours at

room temperature (1:1 for 1hr, 2:1 for 1hr, 3:1 for 1hr, 100% resin for 1 hr then

100

100% resin for 16 hrs and a fresh change again for a further 6 hrs) then the

samples were transferred into Beem capsules full of fresh LR White and placed

at 60oC for 16 hrs to polymerize. The material was sectioned with a glass knife

using a Reichert ultramicrotome (Leica).

For light microscopy, 0.5µm thick sections were dried onto glass slides and

stained with 0.5% (w/v) Toluidine blue “O” in 0.5% (w/v) borax. Photographs

were taken on a Nikon E800 light microscope with Kodak technical Pan film.

For electron microscopy, ultrathin sections of approximately 90nm were picked

up on 200 mesh copper grids which had been pyroxylin and carbon coated. The

sections were stained with 2% (w/v) uranyl acetate for 1hr and 1% (w/v) lead

citrate for 1 minute, washed in distilled water and air dried. The grids were

viewed in a Jeol 1200 EX transmission electron microscope at 80kV and

photographs were taken on Kodak electron image film.

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2.17 Bacteroid excretion studies

2.17.1 Percoll purification of bacteroids

Nodules were picked off the plant roots onto ice. The total yield of nodules was

weighed. Anaerobic isolation of bacteroids was carried out within an AtmosBag

(Aldrich) filled with argon using buffers that had been purged with argon. Nodules

were crushed in a mortar and pestle in isolation buffer (100mM potassium

phosphate, 0.3M sucrose, 2mM MgCl2, pH 7.4). The resulting breis was strained

through 4 layers of muslin to remove large debris. Percoll gradients were pre-

formed by spinning a solution of 57% Percoll (Pharmacia) 43% isolation buffer for

45 minutes at 20,000rpm (Sorvall SS-34). The filtrate was spun through the

gradient (20,000 rpm for 15 minutes) to separate the plant fraction from the

bacteroids (Reibach et al., 1981). The bacteroid fraction, which formed a band

approximately half way down the gradient, was collected using a 3.5 inch 17

gauge needle and syringe. Bacteroids were washed twice in isolation buffer to

remove the Percoll and resuspended in 50mM phosphate buffer with 0.3M

Sucrose and 2mM MgCl2 (pH 7.4) (2 ml of buffer per gram of wet weight

nodules).

2.17.2 Incubation of isolated bacteroids

The bacteroid suspension was injected into a 100 ml Schott bottle sealed with

neoprene that had been pre-sparged with 100% N2 (Fig 2.1). Experimental

conditions were varied for O2 concentration and substrate (malate and/or

ammonium) depending on experiment. Bacteroids were incubated at 26°C with

shaking at 100rpm. Samples were taken from the bottle using a syringe and

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needle at different time points. Samples were centrifuged to pellet the bacteroids

and the supernatant was assayed for alanine and ammonium concentration. A

sample of bacteroid suspension was used to determine the protein level.

Fig 2.1 Experimental vessel for bacteroid excretion assays. 100 ml Schott bottle in which holes were drilled through the lid and bottle was made air-tight with a layer of neoprene placed inside the lid.

2.17.3 Quantification of alanine

The concentration of alanine in the bacteroid supernatant was determined by

measuring the absorbance of the NADH produced in an L-alanine dehydrogenase

linked enzyme assay using alanine as standard (0 – 120nmole range). Reactants

were 50µl sample (or standard), 60µmol hydrazine, 75µmol, glycine, 4µmol NAD+

and 0.5U of L-alanine dehydrogenase (Sigma) made up to a final reaction volume

of 1.5 ml with water. Samples were assayed in duplicate with a minus enzyme

Neoprene seal

Bacteroids resuspended in 50mM PO4 buffer

Nitrogen atmosphere with variable [O2]

Variable mM malate / NH4Cl

Samples injected in & taken out through lid

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control for every sample. Reactions were incubated at 37oC for 90 min. NADH

absorbance was read at 340nm on a spectophotometer.

2.17.4 Quantification of ammonium

Ammonium was distilled from the bacteroid supernatant in order to determine its

concentration. All distillation was carried out in glassware that had previously

been acid washed in 10% HCl for at least ½ hour to remove any contaminating

ammonia. Sample (0.2 ml) and H2O (0.8 ml) was added to 1 ml of saturated

K2CO3 in a universal. An etched glass rod dipped in 1M H2SO4 was used to

collect the ammonia during the distillation process. The universal was sealed

immediately by a bung (through which the etched glass rod had been inserted).

Universals were incubated in an orbital shaker at 25°C overnight with shaking at

80rpm. The end of the etched glass rod was washed in 5 ml water contained in a

7 ml sterilin bijou.

The ammonium concentration of the distillate was determined by Phenol

Hypochlorite assay (Dilworth & Glenn, 1982) using NH4Cl as standard (0-50

nmole range). Each sample was quantified in triplicate. A solution of NaPhenate

was freshly prepared by mixing 0.5 ml 6.25% phenol with 7.5 ml 12.50% NaOH.

To 0.9 ml of sample (or standard) 0.15 ml of the NaPhenate was added and

mixed. 225µl each of 0.02% Sodium Nitro Prusside (Na2Fe(CN)5NO) and 0.04M

Na hypochlorite solutions were added simultaneously. Samples were mixed

thoroughly and incubated for 30 minutes at room temperature. The absorbance at

650nm was read on a spectrophotometer.

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2.17.5 15N2 labelling studies

Labelling with 15N2 was done with high-density bacteroids as described in 2.18.2

except that samples were incubated in 1% O2, 20% 15N2 (99 atom percent, Isotec

Inc), 79% argon. The ammonium released was distilled, acidified and dried

slowly over a period of 36 h at 25oC, and analysed for total nitrogen and 15NH4+

using a Europa Scientific Roboprep combustion analyser interfaced to a VG 622

isotope ratio mass spectrometer referenced against IAEA quality standard 305.

To analyse alanine for 15N, samples were purified using a SP-25 column as for

total amino acid analysis and freeze dried. Dried samples were redissolved in 20

ml pyridine and derivitized using an equivalent volume of N-(tert-

butyldimethylsilyl)-N-methyltrifluoroacetamide (MTBSTFA), mixed and heated in

locking eppendorf tubes for 30 minutes at 70°C. Samples (1.5 ml) were injected

into a VG Masslab GC-MS MD800 using an AS800 autosampler, separated on a

BPX5 column and analysed, initially in scan mode, using the mass spectrometer

(Mawhinney et al., 1986). Subsequently, the mass spectrometer was

programmed in single ion monitoring (SIM) mode and a second injection used to

monitor the ion count for the largest significant fragment (for alanine; mass 260),

mass + 1 (for alanine; mass 261), and mass + 2 (for alanine; mass 262). The

amount of 15

N incorporated into alanine was assessed by measuring the ratio of

the M peak to the M+1 peak and the atom percent excess (APE) calculated as:

APE = (Re – Rc)/(1+(Re – Rc)) × 100

Where Re is M+1/ M for enriched sample and Rc is M+1/M for control samples

(Robinson et al., 1991). Analysis of norleucine as an internal standard revealed

no significant enrichment. Alanine synthesis by low-density bacteroids was also

105

measured using the GC-mass spectrometer, where either 10 or 20 mM NH4Cl

(Sigma 10% atom percent excess) was included.

2.17.6 Amino acid analysis

Bacteroid supernatant was run through a Sephadex SP-25 (Pharmacia) column

and the amino acid eluate freeze-dried (Redgwell, 1980). Reconstituted samples

were analysed for individual free amino acids using a Biochrom 20 amino acid

analyser (Biochrom Ltd.).

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CHAPTER 3

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3 IDENTIFICATION & CHARACTERISATION OF CARBON STORAGE COMPOUNDS IN RHIZOBIUM LEGUMINOSARUM SYMBIOSES

To gain some understanding of the role of PHB metabolism in determinate and

indeterminate symbioses, Kim and Copeland (1996) looked at the enzymatic

capacity for metabolism of PHB in both soybean and chickpea bacteroids. They

suggested a greater potential for oxidising malate to oxaloacetate in chickpea

bacteroids might favour the utilisation of acetyl-CoA in the TCA cycle over PHB

synthesis. Therefore PHB metabolism is lower in chickpea bacteroids and hence

PHB does not form large granules as typically seen in soybean bacteroids.

However, it is difficult to draw conclusions from the study due to the inherent

differences between the bacterial species. Mesorhizobium ciceri (the chickpea

microsymbiont) and Bradyrhizobium japonicum (the soybean microsymbiont)

differ dramatically in growth rate, carbon metabolism and molecular

characteristics (MartinezRomero & CaballeroMellado, 1996; Young, 1996).

We have looked at PHB metabolism in a determinate versus indeterminate

symbiosis using isogenic strains of R. leguminosarum. Strains A5 and A34 are

derived from the same field isolate (strain 8002) of R. leguminosarum bv.

phaseoli which infects bean (Phaseolus vulgaris). This field isolate was selected

for resistance to rifampicin (giving strain A5) and for resistance to streptomycin.

The streptomycin resistant strain was cured of its bean (pRL2JI) Sym plasmid

(giving strain A31) and subsequently the pea (pRL1JI) Sym plasmid was inserted

enabling the infection of pea (Pisum sativum) by strain A34 (Downie, pers.

comm.).

108

By mutating phaC in both these strains of R. leguminosarum we have

investigated and compared the relevance of PHB metabolism to each pea and

bean symbiosis. Bacteroids in symbiosis with bean would be expected to produce

considerable reserves of PHB and therefore the effect of blocking PHB synthesis

on nitrogen fixation might be similar to the effect seen in the R. etli – bean

symbiosis where the efficiency of nitrogen fixation increased (Cevallos et al.,

1996). If the absence of PHB in pea bacteroids is an indication that PHB

metabolism is not relevant to bacteroid metabolism one might expect a mutation

in PHB synthase to produce a null effect on the overall efficiency of nitrogen

fixation as seen in previous studies using S. meliloti (Povolo et al., 1994; Willis &

Walker, 1998; Cai et al., 2000). Here we show different symbiotic phenotypes to

those reported previously in both the determinate and indeterminate symbiosis.

There is no effect of the mutation in the bean symbiosis whilst in the pea

symbiosis the mutation has a variable effect on symbiotic performance.

We aim to investigate the role of glycogen metabolism in the same two R.

leguminosarum backgrounds. To this end we have made preliminary studies into

the role of glycogen metabolism during symbiosis by mutating glgA (glycogen

synthase) in strain A34 and show that the mutation has no effect in the pea

symbiosis.

Some of the data presented here was presented at the 12th International

Congress on Nitrogen Fixation, Brazil in September 1999.

109

3.1 Analysis of the sequence of phaC (PHB synthase)

Rhizobium leguminosarum bv. viciae strain 3841 mutated in phaC (PHB

synthase) by Tn5 (RU137) was previously isolated in this laboratory (Walshaw,

1995). The transposon and adjacent DNA was cloned as a 9.2 Kb EcoRI

fragment in pSK- (pRU99). The transposon insertion site was confirmed by

sequencing the DNA adjacent to the transposon using a primer that annealed to

the IS50 insertion sequence of Tn5 (P113). The IS50 sequence is duplicated at

both ends of the transposon and therefore sub-clones of pRU99 (plasmids

pRU100 and pRU101) were constructed such to contain only one end of Tn5

(hence only one priming site) (Walshaw, 1995). For the purpose of this study the

remaining phaC sequence was obtained in order to design PCR primers and to

map restriction sites to enable the construction of PHB- mutants in the isogenic

strains. Sub-clones of pRU99 (pRU475 and pRU493) were made to sequence

DNA using M13 forward and reverse primers and primers were designed from

data to extend sequences until the entire sequence carried by pRU99 was

obtained.

The 3365 nucleotide sequence from pRU99 contained two truncated open

reading frames. The largest open reading frame showed a high level of amino

acid identity to other rhizobia PHB synthase genes. The phaC open reading

frame was truncated at the C-terminus therefore to complete the sequence DNA

was amplified from 3841 using a primer specific to R. leguminosarum sequence

that read out of the sequence (P139) and a primer (P234) that bound 69 bp

outside the stop codon of R. etli phaC sequence (GenBank database accession

110

no. U30612). The 1.8 Kb fragment obtained was cloned into pCR®2.1-TOPO

(pRU720) and the C-terminus was sequenced using the M13 reverse primer.

The entire sequence obtained is shown in Fig 3.1. As it is mainly single strand

sequence it has not been submitted onto a public database.

111

Fig 3.1 Sequence obtained from pRU99 subclones & pRU720. Restriction sites shown in pink (EcoRI); grey (HindIII); green (BamHI); yellow (EcoRV); blue (PshAI).

phacseq1.doc

112

113

The entire phaC sequence was used in an amino acid alignment with the PHB

synthase genes isolated from R. etli (Cevallos et al., 1996) and from two strains

of S. meliloti (Tombolini & Nuti, 1989; Willis & Walker, 1998). The gene from R.

leguminosarum showed 92%, 83% and 82% amino acid identity to phaC genes

from R. etli (EMBL accession number U30612) and S. meliloti strains Rm1021

(EMBL accession number U17227) and Rm41 (EMBL accession number

AF031938) respectively (Fig 3.2).

The second divergently transcribed open reading frame identified in this study

encoded a peptide with greatest amino acid identity to a hypothetical

aminotransferase (82% identity) which was located upstream of PHB synthase in

S. meliloti strain Rm1021 (Willis & Walker, 1998). The peptide showed weaker

homology (25% identity) to S. meliloti aspartate aminotransferase (aatA) (Rastogi

& Watson, 1991). A mutation in AatA isolated in S. meliloti reduced aspartate

amino transferase activity to 40% of the wild type. The mutant formed nodules

that were unable to fix nitrogen, suggesting that aspartate metabolism via

aspartate aminotransferase is essential for bacteroid nitrogen fixation. However,

multiple forms of the enzyme are present in S. meliloti and not all of these are

required for nitrogen fixation (see section 1.5.3). There is at least one other

putative aspartate aminotransferase in R. leguminosarum 3841 whose sequence

shows strong homology to AatA of S. meliloti (Allaway, pers. comm.). Therefore

as the role of aminotransferases in Rhizobium is complex it was decided that

further investigation of the gene adjacent to phaC was beyond the scope of this

study. However, the original mutant in PHB synthase mutant (RU137) isolated in

the laboratory (Walshaw, 1995) was able to grow on toxic levels of aspartate. The

114

location of the putative aminotransferase adjacent to phaC might suggest that in

RU137 aminotransferase transcription is up-regulated resulting in an improved

ability by the mutant to detoxify aspartate.

As the DNA sequence between phaC and the putative aminotransferase was

approximately 700bp we analysed the DNA using Testcode to search for putative

coding sequence. The results (Fig 3.3) suggested the DNA was coding

sequence, but no matches were found when this section of DNA was subject to a

BLAST search. In S. meliloti this region contains a Rhizobium-specific intergenic

mosaic element (RIME) (Willis & Walker, 1998). RIMEs are approximately 100 bp

repeat elements, characterised by two large palindromes, located outside of the

coding regions (Osteras et al., 1995). To ascertain whether the sequence in R.

leguminosarum contained such an element we searched for repetitive sequences

and compared the nucleotide sequence against that of S. meliloti. We were

unable to identify a RIME in the sequence.

115

1 60 R. etli phaC (1) MYNKRIKRVLPP-EEMVTDSKQESGGQKNGDKTGFDATDLKPYLLKDPETMAMNFARALE R. leg. phaC (1) MYNEWIKRVVLPPEEMVTDSKQENGG-----KAGFDATDLDPYLLKDPEAMAMNFARALE S. mel1 phaC (1) ------------------------------------------------------MARAAE S. mel2 phaC (1) ----------------VTAEKAEGATG----FAGFDPKSVEPYIVKDPESLAINMARAAE Consensus (1) MYN IKRVL P EEMVTDSKQE GG KAGFDATDLDPYLLKDPESMAMNMARALE 61 120 R. etli phaC (60) NLGQAASAWLAPRERGEITETAIDPMTDMVKTLSKISEYWISDPRRTFEAQTQLMSSFFG R. leg. phaC (56) NLGQAASALLCARERGEITESAADPMTDMVKTLSKVTEYWISDPRRTFEAQTQLMSSFFG S. mel1 phaC (7) QLGKAASAWLAPREAGEKTDSFAEPVSDMVKTLSKVSEYWLSDPRRTLEAQTHLLGSFFD S. mel2 phaC (41) QLGKAASAWLAPREAGEKTDSFAEPVSDMVKTLSKVSEYWLSDPRRTLEAQTHLLGSFFD Consensus (61) NLGQAASAWLAPRERGEKTDSFADPMSDMVKTLSKVSEYWISDPRRTLEAQTQLLSSFFG 121 180 R. etli phaC (120) IWMRSMQRMQGTRGMQGEPLPPEPDTRKDKRFSDEDWQKNPFFDFLRQVYFVTSDWVDKL R. leg. phaC (116) IWMRSMQRMQG------DPTPPEPDTRKDKRFSDEDWQKNPFFDFLRQVYFVTTDWVEKM S. mel1 phaC (67) MWSRTLQRMAA-----DAVEDPANLQHNDKRFADEDWVKNPFFDFIRQAYFVTSDWAERM S. mel2 phaC (101) MWSRTLQRMAG-----DAVEDPANLQRNDKRFADEDWVKNPFFDFIRQAYFVTSDWAERM Consensus (121) IWSRSLQRMQG DAVEPPEPLTRNDKRFADEDWVKNPFFDFIRQVYFVTSDWVEKM 181 240 R. etli phaC (180) VSETDGLDEHTKHKAGFYVKQITAALSPSNFIATNPQLYRETIASNGENLVRGMKMLAED R. leg. phaC (170) VSETEGLDEHTKHKAGFYVKQITAAFRRP--TSSHNPQSTRAIATSGANLVRGMKMLAED S. mel1 phaC (122) VKDAEGLDDHTRHKAAFYVRQIASALSPTNFITTNPQLYRETVASSGANLVKGMQMLAED S. mel2 phaC (156) VRDAEGLDDHTRHKAAFYVRQIASALSPTNFITTNPQLYRETVASSGANLVKGMQMLAED Consensus (181) VSDTEGLDDHTKHKAAFYVKQITAALSPTNFITTNPQLYRETIASSGANLVKGMQMLAED 241 300 R. etli phaC (240) IAAGKGELRLRQTDMTKFAVGRDMALTPGKVIAQNDICQIIQYEASTETVLKRPLLICPP R. leg. phaC (228) IAAGHGDLRLRQTDMTKFAVGRDMALTPGKVIAQNDICQIIQYEASTETVLKRPLLICPP S. mel1 phaC (182) IAAGRGELRLRQTDTSKFAIGENIAITPGKVIAQNDVCQVLQYEASTETVLKRPLLICPP S. mel2 phaC (216) IAAGRGELRLRQTDTSKFAIGENIAITPGKVIAQNDVCQVLQYEASTETVLKRPLLICPP Consensus (241) IAAGRGELRLRQTDTSKFAIGRNIAITPGKVIAQNDICQIIQYEASTETVLKRPLLICPP 301 360 R. etli phaC (300) WINKFYILDLNPQKSFIKWCVDQGQTVFVISWVNPDGRHAEKDWAAYAREGIDFALETIE R. leg. phaC (288) WINKFYILDLNPQKSFIKWCVDQGQTVFVISWVNPDARHADKDWAAYAREGIDFALDTIE S. mel1 phaC (242) WINKFYVLDLNPEKSFIKWAVDQGQTVFVISWVNPDERHASKDWEAYAREGIGFALDIIE S. mel2 phaC (276) WINKFYVLDLNPEKSFIKWAVDQGQTVFVISWVNPDERHASKDWEAYAREGIGFALDIIE Consensus (301) WINKFYILDLNPQKSFIKWCVDQGQTVFVISWVNPDERHASKDWEAYAREGIGFALDTIE 361 420 R. etli phaC (360) KATGEKEVNAVGYCVGGTLLAATLALHAKEKNKRIKTATLFTTQVDFTHAGDLKVFVDEE R. leg. phaC (348) KATGEKDVNTVGYCVGGTLLAATLALHAKEKNKRIKTATLFTTQVDFTHAGDLKVFVDEE S. mel1 phaC (302) QATGEREVNSIGYCVGGTLLAATLALHAAEGDERIRSATLFTTQVDFTHAGDLKVFVDDD S. mel2 phaC (336) QATGEREVNSIGYCVGGTLLAATLALHAAEGDERIRSATLFTTQVDFTHAGDLKVFVDDD Consensus (361) QATGEKEVNSIGYCVGGTLLAATLALHAKEKNKRIKSATLFTTQVDFTHAGDLKVFVDDD 421 480 R. etli phaC (420) QLAALEEHMQAAGYLDGSKMSMAFNMLRASELIWPYFVNSYLKGQEPLPFDLLFWNADST R. leg. phaC (408) QLESLEEHMQAAGYLDGTKMSMAFNMLRASELIWPYFVNNYLKGQDPLPFDLLFWNADST S. mel1 phaC (362) QIRHLEANMSATGYLEGSKMASAFNMLRASELIWPYFVNNYLKGQDPLPFDLLYWNSDST S. mel2 phaC (396) QIRHLEANMSATGYLEGSKMASAFNMLRASELIWPYFVNNYLKGQDPLPFDLLYWNSDST Consensus (421) QIRHLEENMSATGYLDGSKMASAFNMLRASELIWPYFVNNYLKGQDPLPFDLLFWNADST 481 540 R. etli phaC (480) RMAAANHAFYLRNCYLRNALTQNEMILDGKRISLKDVKIPIYNLATREDHIAPAKSVFLG R. leg. phaC (468) RMAAANHAFYLRNCYLKNALTQNEMILDGKSVSLKDVKIPIYNLATREDHIAPAKSVFFG S. mel1 phaC (422) RMPAANHSFYLRNCYLENRLSRGEMMLAGRRVSLGDVKIPIYNLATKEDHIAPAKSVFLG S. mel2 phaC (456) RMPAANHSFYLRNCYLENRLSKGEMVLAGRRVSLGDVKIPIYNLATKEDHIAPAKSVFLG Consensus (481) RMPAANHAFYLRNCYLENRLSQNEMILDGKRVSLKDVKIPIYNLATKEDHIAPAKSVFLG 541 600 R. etli phaC (540) SRFFGGKVEFVVTGSGHIAGVVNPPDKRKYQFWTGGPAKGEYETWLEQASETPGSWWPHW R. leg. phaC (528) SQFFGGKVEFVVTGSGHIAGVVNPPDRKKYQFWTGGPAKGDYETWLDQATETPGSWWPHW S. mel1 phaC (482) SSSFGGKVTFVLSGSGHIAGVVNPPARSKYQYWTGGAPKGDIETWMGKAKETAGSWWPHW S. mel2 phaC (516) SSSFGGKVTFVLSGSGHIAGVVNPPARSKYQYWTGGAPKGDIETWMGKAKETAGSWWPHW Consensus (541) SSSFGGKVTFVLSGSGHIAGVVNPPDRSKYQFWTGGPPKGDYETWLGQAKETPGSWWPHW 601 638 R. etli phaC (600) QAWIETHDGRRVAARKPGGDALNAIEEAPGSYVMERT- R. leg. phaC (588) RAWIETHDGRRVPSRKPGGDALNAIEEAPGSYVMERA- S. mel1 phaC (542) QGWVERLDKRRVPARKAG-GPLNSIEEAPGSYVRVRA- S. mel2 phaC (576) QGWVERLDKRRVPARKAG-GPLNSIEEAPGSYVRVRA- Consensus (601) QAWIETLDKRRVPARKPGGGPLNAIEEAPGSYVRVRA Fig 3.2 Amino acid alignment of poly-β-hydroxybutyrate synthase from R. etli (R. etli), R. leguminosarum bv. viciae (R. leg.), S. meliloti Rm1021 (S. mel1) and S. meliloti Rm41 (S. mel2). The amino acid identity is colour coded. black - identical, green - weakly similar, blue - conservative, grey - non homologous, pink - block of similar.

116

Fig 3.3 Results of Testcode analysis.

phacaatx testcode.ppt

117

The principle focus for this study was to ascertain the symbiotic properties of

PHB synthase mutants in two different symbioses using an isogenic strain that

differed in only its Sym plasmid. Therefore we made no further attempt to

ascertain the genetics of PHB production in R. leguminosarum. However, we

were interested to find out whether the gene was carried on the Sym plasmid or

not in R. leguminosarum. To identify whether phaC was located on the pSym in

R. leguminosarum, EcoRI digested total DNA preparations from strains A5, A34

and A31 (A5 cured of its Sym plasmid) were analysed by Southern blot and

hybridisation using a phaC PCR clone as the probe (pRU566 see section 3.2 for

details). The phaC gene was present on the same EcoRI fragment in all three

strains indicating the gene was located on chromosomal DNA or a plasmid other

than the Sym plasmid (Fig 3.4).

Fig 3.4 Southern hybridisation to confirm phaC was not present on pSym. EcoRI digests of genomic DNA. lane 1 - Kb ladder; lane 2 – A31 (no Sym plasmid); lane 3 – A34; lane 4 – A5.

1 2 3 4

3.4 kb

118

3.2 Construction of mutants in PHB synthase

A phaC PCR product was required for construction of insertion mutations in

strains A5 and A34 and for use as a probe in Southern hybridisation. The

sequence of phaC was used to design primers (P138 & P166) to amplify a 1.6 Kb

region of the gene from A5 (see Fig 3.1 for priming sites). The PCR product was

cloned into pCR®2.1-TOPO (to produce plasmid pRU566) and was sequenced

using M13 Forward and Reverse primers to the vector to confirm the correct

fragment had been cloned. The DNA sequence of phaC was analysed for

restriction sites that could be used to insert a spectinomycin resistant omega (Ω)

cassette. The cassette is flanked by BamHI, EcoRI, HindIII, SmaI and XmaI sites

which can be used to clone the DNA. PhaC was found to have a unique blunt

PshAI site (see Fig 3.1 for location). Therefore, the cassette was excised from the

vector pPH45Ω as a blunt SmaI fragment and cloned into the PshAI site within

phaC to give pRU575 (Fig 3.5).

119

Fig 3.5 Vectors used in the construction of mutants in PHB synthase (RU1328 and RU1329). Not to scale.

phaC construction1.doc

120

The disrupted gene fragment was excised from pRU575 using 2 EcoRI sites that

flank the PCR cloning site of pCR®2.1-TOPO. The 5’ overhangs of the EcoRI

ends were filled in using Klenow. The DNA was blunt cloned into a unique SmaI

site of the suicide vector pJQ200SK (genr) (Quandt & Hynes, 1993) (Fig 3.5).

This construct (pRU577) was conjugated into Rhizobium leguminosarum strains

A5 and A34. Plasmid pJQ200SK does not replicate in Rhizobium and therefore

colonies that were genr spcr resistant on primary selection plates indicated a

single recombination event into the chromosome at phaC and integration of the

vector. As pJQ200SK carries the sacB gene from Bacillus subtilis double

recombinants were selected for by sub culturing the genr spcr colonies on minimal

media containing 10% sucrose. For each A34 and A5 twelve sucrose resistant

colonies were purified and checked for gentamicin sensitivity (hence the loss of

vector). The chromosomal DNA was extracted from the strains and used in PCR

reactions using primers P138 and P166. In each case a fragment of 3.6 Kb was

amplified indicating the incorporation of the 2 Kb spectinomycin cassette into

phaC. Mutants were designated strain numbers RU1328 (A34 parent) and

RU1329 (A5 parent) and confirmed by Southern blot and hybridisation using a

phaC PCR clone (pRU566) as the probe to verify an increase in gene size by 2

Kb on an EcoRI fragment. XmnI, which cuts within the cassette, digests

confirmed the orientation of the cassette (Fig 3.6. Note - A5 wild type and mutant

(RU1329) gave the same band pattern but data is not shown due to the bad

quality of the scanned image).

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Fig 3.6 Southern hybridisation to confirm PHB mutant indicated by increase of 2 Kb in phaC gene size. lane 1 - RU1328 XmnI digest; lane 2 - A34 XmnI; lane 3 - pRU577 XmnI; lane 4 - RU1328 EcoRI; lane 5 - A34 EcoRI.

5.4kb

3.4kb

1 2 3 4 5

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3.3 Growth of PHB synthase mutants

Assessments of the growth characteristics of the PHB synthase mutants were

made to determine whether the mutation had significantly affected metabolism of

the strains in free-living conditions. Strains were streaked onto TY plates and

onto AMA plates containing a range of different carbon sources. The mutants

were severely inhibited in growth on TY (10 days to form single colonies

compared to 4 days for the wild type strains). On glucose, malate and succinate

the mutants appeared to grow slower and formed slightly smaller colonies at 4

days growth (indicated by a scoring of ++ instead of +++ in Table 3.1).

Table 3.1 Growth of PHB mutants and parental strains on different carbon sources. +++ indicates good growth; ++ growth; + poor growth; - no growth.

Carbon Source A34 RU1328 A5 RU1329

Glucose +++ ++ +++ ++

Malate +++ ++ +++ ++

Succinate +++ ++ +++ ++

Glutamate + + + +

Oxaloacetate + + + +

α-ketoglutarate + + + +

Aspartate - - - -

Pyruvate +++ +(17d) +++ +(17d)

Alanine +++ +(17d) +++ +(17d)

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To ascertain this difference in growth rate the mean doubling time of RU1328

was tested in liquid culture with malate as the carbon source (Fig 3.7). There was

no difference in the growth rate of RU1328 compared to A34 even though on

plates the PHB mutant appeared to have a slower growth rate and formed

smaller colonies than the wild type. The difference in the morphology of RU1328

on plates compared to the wild type is likely to be a result of differences in EPS.

Mutations in carbon storage compounds have been shown to affect EPS levels in

Rhizobium, for example a mutation in aniA of S. meliloti led to higher levels of

EPS and conversely the R. tropici glgA mutant had much lower levels of EPS

(Marroqui et al., 2001).

Fig 3.7 Growth of A34 and RU1328 on malate (mean doubling times 3.46 & 3.24 hrs respectively).

time (hours)

0 2 4 6 8 10

Log 10

OD

600n

m

0.1

1

A34 RU1328

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There was no difference in the growth of both the mutants and parental strains on

glutamate, oxaloacetate, α-ketoglutarate and aspartate. On alanine and pyruvate

the mutants were severely inhibited in their growth and formed small pin colonies

after 17 days (Table 3.1). Poor growth on pyruvate was also observed in the PHB

synthase mutant of R. etli (Cevallos et al., 1996). For use of both alanine and

pyruvate as carbon sources the enzyme pyruvate dehydrogenase, which forms

acetyl-CoA, is required. Acetyl-CoA can then be directed into PHB synthesis or

into the TCA cycle. For the incorporation of this carbon into the TCA cycle and

hence for the production of ATP and reduced nucleotides, oxaloacetate is

required to condense with acetyl-CoA to form citrate via citrate synthase.

Oxaloacetate is formed from pyruvate via pyruvate carboxylase. The results of

growth on alanine and pyruvate suggest that PHB metabolism is required to

remove acetyl-CoA to regulate carbon metabolism on these substrates. When

this does not occur there may be an accumulation of acetyl-CoA in the cell which

inhibits growth.

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3.4 PHB content of free-living strains

The PHB content of RU1328, RU1329, A34 & A5 was determined (Table 3.2).

Strains were grown on 10mM fructose as the carbon source with 5mM NH4Cl as

these conditions had been shown to optimise PHB production in S. meliloti

(Tavernier et al., 1997). PHB was not detected in the mutant strains indicating the

pathway for synthesis of PHB had been blocked by the insertion of the omega

cartridge into phaC.

Table 3.2 PHB content of PHB mutants and parental strains grown on 10mM fructose / 5mM NH4Cl

Strain PHB (µg mg protein-1)

A34 65.0

RU1328 ND

A5 50.4

RU1329 ND

ND not detected

The method used for determination of PHB was that of Law & Slepecky (1961).

This assay relies on measuring crotonic acid spectrophotometrically which is

formed during the depolymerisation of PHB under acidic conditions. The accurate

determination of crotonic acid can be subject to interference from common

biological compounds that also absorb in the region of 200-230nm and the

purification procedure may lose short chain length PHB polymers (Law &

Slepecky, 1961). Therefore the method is applicable to qualitative comparisons

but lacks the accurate quantitation required to determine small differences in

126

PHB content or small amounts of PHB content. Therefore no effort was made to

apply this method to look at the regulation of PHB production in free-living cells or

bacteroids.

3.5 PHB content of pea and bean bacteroids

To confirm the absence of PHB in bacteroids and to identify any ultrastructural

differences in nodules formed between the wild types and PHB mutants, sections

of pea and bean nodules were examined by light microscope and by transmission

electron microscopy (TEM). This work was kindly carried out by Kim Findlay at

the John Innes Institute.

For each pea and bean the mutant strains nodulated the plants, forming healthy

pink nodules, as the wild type. There was no discernible differences in nodule

size or number between plants inoculated with the PHB mutant and the wild type

(data not shown).

The micrographs of bean nodule sections showed characteristic features of

determinate nodules. Infected plant cells were full of symbiosomes. Multiple

bacteroids (approximately seven) were surrounded by a single PBM to form each

individual symbiosome (Figs. 3.8, 3.9, 3.10 and 3.11). There are large granules of

starch present in uninfected plant cells adjacent to the infected (Figs. 3.8, 3.9,

3.10 and 3.11). In determinate nodules these cells are involved in carbohydrate

metabolism (Day & Copeland, 1991) and ureide synthesis (Schubert, 1986)

therefore have high levels of the enzymes involved in these activities. A least

20% of an infected cells surface is in contact with an uninfected cell which

highlights the importance of the uninfected cell in exchange of nutrients between

127

plant and bacteroid (Day & Copeland, 1991). Bacteroids were elongated rod

shaped cells (Figs. 3.8, 3.9, 3.10 and 3.11), many of which had been cut

transversely during sectioning and therefore appeared spherical.

PHB formed considerable reserves in bacteroids of R. leguminosarum wild type

strain A5. Mature bacteroids were packed full of PHB granules, which can be

seen on the micrographs as spherical electron-transparent areas inside the

bacteroid (Figs. 3.8, 3.9 and 3.12). The phaC mutation in strain A5 abolishes

PHB production in free-living cells (Table 3.2) and in bacteroids. There were no

PHB granules in RU1329 bacteroids (Figs. 3.10, 3.11 and 3.13). There were no

other visible differences between bacteroids of A5 and RU1328.

The electron transparent material that is disorganised and irregularly spread

throughout the cytoplasm, seen most clearly in the high magnification TEMs

(Figs. 3.12, 3.13) is reminiscent of the material identified as glycogen in clover

bacteroids (Dixon, 1967). A mutant in glycogen synthesis is required to verify this.

If it is the case bean bacteroids accumulate both compounds during symbiosis

(Fig 3.11).

The micrographs of pea nodule sections showed characteristic features of

indeterminate nodules. Fewer uninfected cells were present in the indeterminate

nodule and starch granules were present in the periphery of some infected cells

(Fig 3.18). Pea bacteroids were packaged singly in PBM and many bacteroids

were deformed and showed Y-shape morphology (Fig 3.14). As expected for an

indeterminate nodule, no PHB accumulated in the bacteroids of A34 (Fig 3.15).

The electron transparent material putatively identified as glycogen in the bean

bacteroids was also present in the pea bacteroids (Fig 3.14). There were no

128

apparent differences between bacteroids of A34 and RU1328 as shown by the

transmission electron micrographs (Fig 3.15 and Fig 3.17).

However, sections that cut through infection threads showed that wild type

bacteria inside the infection thread contain granules of PHB (Fig 3.16). As would

be expected for a mutant in phaC, there were no granules present in RU1328

bacteria inside the infection thread (not shown). Previous ultrastructural studies

have indicated PHB granules are present in S. meliloti cells inside the infection

thread (Paau & Cowles, 1978) but they disappear when cells are released from

the infection thread. Mature nitrogen fixing bacteroids contain no visable granules

of PHB (Hirsch et al., 1983; Vasse et al., 1990). Therefore, here we present data

that confirms PHB is also present in R. leguminosarum bv. viciae cells multiplying

in the infection thread and that these reserves disappear upon differentiation into

the bacteroid form.

Transverse sections of pea nodules infected by A34 showed the characteristic

zones of indeterminate nodules (Fig 3.19). Particularly notable was a 1-3 cell

layer where cells were filled with starch granules described as interzone II-III by

Vasse et al. (1990) (Fig 3.20). This zone of cells is characterised by the presence

of the starch granules in the plant cells (Fig 3.18) and is highly significant to the

development of the symbiosis as it represents the transition of bacterial cells (in

zone II) into nitrogen fixing bacteroids (occurring in zone III). Bacterial cells in this

area undergo dramatic changes such as increase in cell volume, production of

nitrogenase and production of novel cytochromes required for the respiratory

system under microaerobic conditions (Vasse et al., 1990). Interzone II-III was

129

not apparent on the sections of nodules infected by RU1328 as there was no

distinct histological zone of cells containing starch (Fig 3.21 and Fig 3.22).

130

Fig 3.8 Section of bean nodule showing PHB accumulation by bacteroids of A5. P – PHB granules (transparent material); B – bacteroid; M – peribacteroid membrane; W – plant cell wall; S – starch; N – plant cell nucleus. Bar 2µm.

Bean wt plant cell 1 c

Bar xµm.

131

Fig 3.9 Section of bean nodule showing PHB accumulation by bacteroids of A5. P – PHB granules (transparent material); B – bacteroid; Y – symbiosome (containing up to 7 bacteroids); M – peribacteroid membrane; W – plant cell wall; S – starch. Bar 2µm.

Bean wt plant cell 2 d

Bar xµm. d

132

Fig 3.10 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329. B – bacteroid; Y – symbiosome (containing up to 7 bacteroids); M – peribacteroid membrane; W – plant cell wall; S – starch in adjacent uninfected cells. Bar 5µm.

Bean PHB- plant cell n

Bar xµm. n

133

Fig 3.11 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329. B – bacteroid; Y – symbiosome (containing up to 7 bacteroids); M – peribacteroid membrane; W – plant cell wall; S – starch in adjacent uninfected cells. Bar 2µm.

Bean PHB- plant cell o

Bar xµm. o

134

Fig 3.12 Section of bean nodule showing PHB accumulation by bacteroids of A5. P – PHB granules (transparent material); G – material that is possibly glycogen (diffuse transparent material); B – bacteroids; M – peribacteroid membrane; C – plant cytoplasm. Bar 500nm.

Bean wt close-up a

Bar xµm. a

135

Fig 3.13 Section of bean nodule showing no PHB accumulation by bacteroids of RU1329. B – bacteroids; M – peribacteroid membrane; G – material that is possibly glycogen (diffuse transparent material); C – plant cytoplasm. Bar 500nm.

Bean phb- close-up b

Bar xµm. b

136

Fig 3.14 Section of pea nodule showing characteristic Y-shape of bacteroids. B – bacteroid; M – peribacteroid membrane; G – material that is possibly glycogen (diffuse transparent material); C – plant cytoplasm. Bar 1µm.

Pea PHB- close up y-shape e

Bar xµm. e

137

Fig 3.15 Section of pea nodule showing no PHB accumulation by bacteroids of A34. B – bacteroid; Y – symbiosome (containing only 1 bacteroid); M – peribacteroid membrane; W – plant cell wall. Bar 2µm.

Pea wt plant cell f

Bar xµm. f

138

Fig 3.16 Section of pea nodule. Note no PHB accumulation by bacteroids of A34, but PHB in bacteria inside infection thread. P – PHB granules (transparent material); B – bacteroid; I – infection thread; A – bacteria. Bar 2µm.

Pea wt infection thread g

Bar xµm. g

139

Fig 3.17 Section of pea nodule showing no PHB accumulation by bacteroids of RU1328. B – bacteroid; Y – symbiosome (containing only 1 bacteroid); M – peribacteroid membrane; W – plant cell wall. Bar 2µm.

Pea PHB- plant cell h

Bar xµm. h

140

Fig 3.18 Section of pea nodule infected by A34 showing starch (S) accumulation at periphery of infected plant cells in interzone II-III. Note starch is absent from plant cells of zone III (nitrogen fixing zone). Bar 5µm.

Pea wt starch around periphery of infected cell i

Bar xµm. I

141

Fig 3.19 Light micrograph of a transverse section of a pea nodule infected by A34. Zones I, II, II-III, III, IV are indicated. Note starch (S) accumulation at periphery of infected plant cells in interzone II-III. Bar 150µm.

Light micrograph wt pea zones j

142

Fig 3.20 Light micrograph of a transverse section of a pea nodule infected by A34 showing zones I, II, and II-III. Note starch (S) accumulation at periphery of infected plant cells in interzone II-III. Bar 100µm.

Light micrograph wt pea inter zone k

143

Fig 3.21 Light micrograph of a transverse section of a pea nodule infected by RU1328. Zones I, II, III, IV are indicated. Note starch is absent from plant cells of interzone II-III. Bar 150µm.

Light micrograph PHB- pea no inter zones l

144

Fig 3.22 Light micrograph of a transverse section of a pea nodule infected by RU1328. Zones I and II are indicated. Note starch is absent from plant cells of interzone II-III. Bar 100µm.

Light micrograph PHB- pea no inter zone m

145

3.6 Symbiotic performance of PHB synthase mutants on pea and bean

Peas and beans were planted in a greenhouse in May 1999 (Fig 3.23 and Fig

3.24) in order to assess the symbiotic performance of both mutants in phaC on

their respective host plants. Plants that grew in the row of un-inoculated pots

between the pots containing wild type and mutant Rhizobium strains also became

nodulated. However the nodules were located at the crown of the root system

and were not distributed evenly throughout the root system, indicating they were

formed by contamination of the soil after the initial inoculation (probably during

watering of the plants). Nodules were harvested from the roots to determine the

source of the contamination. For peas, 100% nodules taken from plants

inoculated with A34 (strr) contained streptomycin resistant / spectinomycin

sensitive bacteria. 98% nodules taken from plants inoculated with RU1328 (strr

spcr) contained streptomycin / spectinomycin resistant bacteria. All the nodules

formed on un-inoculated plants contained strr bacteria (ie. A34 or RU1328). For

beans, 95% nodules taken from plants inoculated with A5 (rifr) contained

rifampicin resistant / spectinomycin sensitive bacteria. 90% nodules taken from

plants inoculated with RU1329 (rifr spcr) contained rifampicin / spectinomycin

resistant bacteria. Again all of the nodules formed on un-inoculated plants

contained rifampicin resistant bacteria. Therefore we were confident of the

legitimacy of the results.

146

Fig 3.23 Peas grown in the greenhouse. Note the double bucket system to allow collection of GM (genetically modified) contaminated waste. Row of plants fourth from the left are the un-inoculated.

147

Fig 3.24 Beans grown in the greenhouse. Note the double bucket system to allow collection of GM (genetically modified) contaminated waste. Row of plants fourth from the left are the un-inoculated.

148

Plants were harvested at the peak period of nitrogen fixation (during plant

flowering) (Sprent, 1982). The peas flowered during the fifth week of growth. The

beans grew more slowly and flowered later (during week 7). At the first harvest

the acetylene reduction of the mutant and wild type strain was measured. The

acetylene reduction assay does not measure the conversion of dinitrogen to

ammonium per se. Instead it gives an indication of the total amount of

nitrogenase capable of fixing nitrogen, so when used in comparative studies it

can give a good indication on any gross differences in nitrogenase activity

between strains. There was a halving of acetylene reduction in peas infected with

RU1328 compared to the wild type indicating that either less nitrogenase is

produced by RU1328 or that there is a major defect in its functioning. There was

no significant difference in bean (Table 3.3).

Dry weights of plants were taken at flowering and during the time when seeds

were maturing. There were no significant differences in dry weight of either peas

or beans inoculated with the PHB mutant compared to the parental strain at

flowering (Fig 3.25). However, by the time of seed maturation, the peas

inoculated with the PHB mutant were significantly reduced (by 37%) in dry weight

(Fig 3.25 and Fig 3.26). There was no significant difference in the beans (Fig

3.25).

149

Table 3.3 Acetylene reduction (ethylene production) by wild type & PHB mutants. Values shown are means of 40 plants plus & minus the standard error of the mean.

Strain Ethylene production (µmoles plant1hr-1)

A34 5.93 ± 0.59

RU1328 2.99 ± 0.17

P<0.001

A5 9.05 ± 0.92

RU1329 10.97 ± 1.11

P>0.05

150

Fig 3.25 Dry weight of (A) peas (A34, n=42; RU1328, n=45; P<0.001 at 53 day harvest) and (B) beans (A5, n=29; RU1329, n=32; P>0.05 at 54 day harvest).

time of harvest (days)

35 53

dry

wei

ght (

g)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

A34 RU1328

time of harvest (days)

46 54

dry

wei

ght (

g)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

A5 RU1329

A

B

151

Fig 3.26 Representative pea plants from the second harvest at 53 days.

A34 RU1328

152

3.7 TCA cycle enzyme activity of pea bacteroids

If preventing PHB synthesis disrupted the redox poise of pea bacteroids it was

possible the activity of some key enzymes in the TCA cycle would be impaired.

Citrate synthase modulates the entry of acetyl-CoA into either the TCA cycle or

PHB metabolism. The activity of citrate synthase in chickpea bacteroids is

strongly inhibited by NADH (Tabrett & Copeland, 2000). Also isocitrate

dehydrogenase activity is regulated by PHB synthesis in A. beijerinckii (Jackson

& Dawes, 1976). Therefore we measured the activity of CS, ICDH and MDH in

RU1328 bacteroids where the nitrogenase activity was reduced to determine

whether there was a correlation between TCA cycle function and the symbiotic

defect. The enzyme activity was only able to be determined once using

bacteroids purified from the entire pooled nodule harvest, hence no errors of the

mean are stated in Table 3.4. We also assessed enzyme activity in free living

cells grown on malate as this is the principle carbon source supplied to the

bacteroid. There were no gross changes to enzyme activity in bacteroids or free

living cells of RU1328 compared to A34. We found there was a slight decrease in

activity of CS and ICDH in the PHB mutant compared to the wild type however

these small differences in activity are unlikely to be physiologically relevant in

either free living cells or bacteroids. There was no difference in the malate

dehydrogenase activity of the mutant versus the wild type in bacteroids or in free

living cells.

153

Table 3.4 Enzyme activity of A34 and RU1328 in bacteroids and in free living (FL) cells grown on malate. Malate dehydrogenase MDH; citrate synthase CS; isocitrate dehydrogenase ICDH.

Strain Activity (µmol min-1mg protein-1)

MDH CS ICDH

Bacteroids

A34 8.196 0.752 0.573

RU1328 8.448 0.572 0.450

FL †

A34 3.604 ± 0.809 0.057 ± 0.001 0.645 ± 0.040

RU1328 2.780 ± 0.774 0.043 ± 0.007 0.481 ± 0.038

† values are mean of 3 experiments ± standard error

From the enzyme activities presented in Table 3.4 it can be seen that CS and

MDH activity is induced in pea bacteroids. The enzyme activities reported here

are consistent with a previous data for R. leguminosarum bacteroids (McKay et

al., 1989). Our original intention was to also measure enzyme activity in A5

bacteroids and free living cells. This would have allowed us to see whether there

were any specific differences between enzymes induced in A34 and A5

bacteroids. Experiments of this kind might give an indication of differences in

carbon metabolism that lead to the presence and absence of large deposits of

PHB in A5 and A34 bacteroids respectively. However, we were unable to obtain a

sample of bean bacteroids that was free of plant contamination using the

technique standardised in the laboratory for pea bacteroids. Therefore we were

unable to carry out a comparative study of bean and pea TCA enzyme activity.

154

3.8 Glycogen content of pea bacteroids

Cevallos (1996) reported the PHB mutant of R. etli accumulated up to 50-fold

more glycogen than the wild type strain during free-living fermentative

metabolism (Cevallos et al., 1996). This suggests that under conditions where the

TCA cycle is impaired in its function, such as microaerobic conditions, carbon flux

in Rhizobium may be easily re-directed to different pathways. We estimated the

glycogen content of purified pea bacteroids in which the nitrogenase activity was

reduced to establish whether there had been any significant effect on this reserve

in the mutant bacteroids. Bacteroids of A34 contained 4.5 times more glycogen

than RU1328 bacteroids (Table 3.5). The results indicate that under conditions

where PHB metabolism is prevented glycogen reserves are either mobilised at a

faster rate or are not built up to the same degree.

Table 3.5 Glycogen content of pea bacteroids

Strain Glycogen (µg mg protein-1)

A34 9.52

RU1328 2.08

155

3.9 Second test of symbiotic performance of PHB mutant on pea

To repeat our nitrogenase activity and growth results for pea we set up a second

greenhouse experiment in July 2000. However in this experiment we saw no

difference in acetylene reduction (A34 - 4.52 ± 0.86; RU1328 - 4.38 ± 0.61

µmoles ethylene plant-1hr-1, n=15) or dry weight (A34 - 2.55 ± 0.05 (n=40);

RU1328 - 2.47 ± 0.06 (n=48) grams) between plants inoculated with the PHB

mutant and wild type. The experiments only differed in time of planting (May

versus July) which would have affected the rate of photosynthesis of the plant

and hence the carbon supply to nodules. Therefore we suspect the relevance of

a mutant in PHB synthase, in at least indeterminate symbioses, may depend on

the status of carbon metabolism by the plant.

156

3.10 Construction of a mutant in glycogen synthase

Rhizobium tropici strain CIAT889 mutated by an in frame deletion in glgA

(glycogen synthase) was isolated (A656) by S. Marroqui (John Innes Institute).

The colony size of this mutant was smaller than that of the wild type when grown

on Y minimal media plates (10mM glucose / 10mM NH4Cl). This phenotype was

due to a reduced level of exopolysacharide (EPS) in the mutant strain. As the

deletion in glgA was in frame, the reduced levels of EPS were not a result of a

polar effect on pgm (phosphoglucomutase) located downstream of glgA (Fig 1.7)

but were a result of the glgA mutation itself (Marroqui et al., 2001). The mutant

was complemented by a cosmid (pIJ9019) from the gene bank of A5 on the basis

of restoring the ability to synthesise wild type levels of exopolysaccharide. This

cosmid was isolated by Sylvia Marroqui for the following studies. We confirmed

the cosmid carried glgA by southern blot and hybridisation (Fig 3.27) using a 0.7

Kb EcoRI/XhoI fragment of glgA from R. tropici (carried on plasmid pIJ7843 also

provided by Sylvia Marroqui). The cosmid was used to create a transposon

(TnB60) mutant of glgA in R. leguminosarum strain A34.

157

Fig 3.27 Southern hybridisation to confirm glgA is carried on cosmid pIJ9019. Lane 1 - EcoRI digest of cosmid; lane 2 - BamHI digest; lane 3 - Kb ladder.

The genes encoding enzymes in the pathway of glycogen biosynthesis in

Rhizobium are arranged as in Agrobacterium tumefaciens with glgA immediately

upstream of pgm (see Fig 1.7). Pgm encodes phosphoglucomutase, which is an

intermediary enzyme in the synthesis of both glycogen and exopolysaccharide

(see Fig 1.6). EPS is important for an effective symbiosis and in some cases

mutants in EPS have a detrimental effect on nodulation and/or nitrogen fixation.

For example, R. leguminosarum mutants not able to produce EPS fail to nodulate

peas but are able to nodulate beans (Borthakur et al., 1986; Diebold & Noel,

1989). Other studies have found no consistent correlation between rhizobial EPS

production and nodulation of beans. Some EPS mutants nodulate beans as well

as the parent strain, whereas others do not produce normal nodules (Sanders et

4.6kb

2.1kb

1 2 3

158

al., 1981; Raleigh & Signer, 1982). Thus as the primary focus of the project was

to produce glgA mutants for use in plant experiments, TnB60 was chosen for the

initial mutagenesis of glgA to prevent a polar mutation on pgm. TnB60 is a

derivative of Tn5 and carries an outwardly directed tac promoter to ensure

expression of downstream genes (Simon et al., 1989).

Cosmid pIJ9019 was mutated with λTnB60 and the mutagenised cosmids were

conjugated in bulk into R. tropici strain A656 to isolate those that no longer

complemented the glgA mutant phenotype on Y minimal media plates.

Approximately 10% of kanr tetr colonies on the initial spread plates were of the

small colony phenotype. Six of these small colonies were streaked onto Y

minimal media (10mM glucose) plates to confirm the phenotype. The colonies (4

of six isolates) that grew on these secondary streak plates were of the same

phenotype as R. tropici strain A656. Cosmids were extracted by alkaline lysis

from these 4 strains and immediately transformed into DH5α. Cosmid DNA was

extracted from DH5α and digested with EcoRI and SalI to analyse restriction

patterns. The restriction patterns of 1 TnB60 mutated cosmid differed from the

wild type and therefore was analysed by sequencing. The transposon and

flanking DNA was sub-cloned as an EcoRI fragment into pSK- (pRU626).

Although only having one IS50 insertion sequence, the primer (P113) used for

sequencing from IS50 was found to bind at both ends of TnB60 in pRU626.

Therefore a 1.5 Kb HindIII fragment, that only contained the IS50 sequence and

the DNA flanking the transposon, was sub-cloned into pSK- (pRU638). The

insertion site was then identified by sequencing using P113. The sequence data

indicated the transposon was inserted in glgA (84% amino acid identity to glgA

159

from Agrobacterium tumefaciens EMBL AF033856) and had a 9 bp repeat of

GCTCGTTAT. It was orientated such that ptac was directed to read downstream

of glgA (Fig 3.28).

Fig 3.28 Orientation of transposon TnB60 in glgA. Flags indicate direction of promoters.

The cosmid was designated number pRU3146 (tetr kanr) and was conjugated into

A34 to create a chromosomal mutation in glgA. After purification, integration of

the disrupted glgA gene was facilitated by the conjugation of a second plasmid

(pPHJI1 genr). pPHJI1 is incompatible with pLAFR1 (the parent vector of

pIJ9019) therefore conjugants that are both genr and kanr indicate at least a

single recombination event between cosmid DNA and host DNA. Genr kanr

conjugants were purified and single colonies were checked for loss of the cosmid

vector (tetracycline sensitivity), which would indicate a double recombination and

hence integration of the mutated gene (Ruvkun & Ausubel, 1981). No double

recombinants were isolated by this method therefore the transposon and flanking

DNA was directly cloned into pJQ200SK as a 12 Kb SacI fragment (pRU758)

which was conjugated into A34 and the mutation in glgA was constructed using

sucrose selection against the presence of the pJQ200SK vector (see section

IS50R

P113

glgA TnB60

ptac pgm

160

3.2). Three putative mutants (gens kanr sucroser) were streaked onto Y minimal

plates containing 10mM glucose to ascertain the size of the colonies in relation to

those of R. tropici A656 and R. leguminosarum A34. All three strains exhibited

the small colony phenotype.

The chromosomal DNA was extracted from the 3 putative mutants (glgA no’s #1,

#2 and #3) and digested with EcoRI. The mutation in glgA was confirmed by

Southern blot and hybridisation using a 0.7 Kb EcoRI/XhoI fragment of glgA from

R. tropici (carried on plasmid pIJ7843) as probe. EcoRI cuts at one extreme end

of the transposon, hence the wild type fragment of 2.1 Kb is split into two

fragments upon insertion of the transposon. One fragment includes all of the

transposon (4.8 Kb) and 370 bp of glgA sequence (fragment 5.2 Kb in size) and

the second fragment carries 1.7 Kb glgA sequence (Fig 3.29).

One glgA mutant (no.2) was designated strain number RU1448 (strr kanr) and

was used in all further analysis.

161

Fig 3.29 Southern blot and hybridisation to confirm mutations in glgA. Lane 1 - EcoRI A34; lane 2 - EcoRI glgA #1; lane 3 - EcoRI glgA #2; lane 4 - EcoRI glgA #3.

5.2kb

2.1kb 1.7kb

162

3.11 Growth of mutant in glycogen synthase

R. tropici strain A656 was affected in growth on glucose, but was unaffected by

succinate or mannitol (Marroqui et al., 2001). We grew RU1448 on minimal plates

containing different carbon sources and assessed growth in broth to ascertain its

growth phenotype. The mutation in glgA prevented growth on glucose (Table 3.6

and Fig 3.30). Growth on mannitol was slowed, but growth on sucrose, fructose

and succinate was unaffected (Table 3.6 and Fig 3.30). The normal growth on

sucrose indicates that glucose is not toxic to the cells, but must slow growth down

considerably. If glucose were toxic to the cell one would expect there to be no

growth on sucrose. The ability of the strain to grow on fructose indicates that

during growth on sucrose the cell is probably utilising fructose, derived when

sucrose is cleaved, for growth.

Table 3.6 Growth of glycogen mutant and wild type on different carbon sources. +++ indicates good growth; ++ growth; + poor growth; - no growth.

Strain Glucose Succinate Mannitol Sucrose Fructose

A34 +++ +++ +++ +++ +++

RU1448 - +++ ++ +++ +++

163

Fig 3.30 A Growth of A34 and RU1448 on glucose (mean doubling times 3.64 hrs for A34 and not determined for RU1448); B growth of A34 and RU1448 on succinate (mean doubling times 3.66 & 3.86 hrs respectively); and C growth of A34 and RU1448 on mannitol (mean doubling times 3.48 & 6.60 hrs respectively).

time (hours)

0 2 4 6 8 10 12

Log 10

OD

600n

m

0.1

13841 RU1448

time (hours)

0 2 4 6 8 10 12

Log 10

OD

600n

m

0.1

1

3841 RU1448

time (hours)

0 2 4 6 8 10 12

Log 10

OD

600n

m

0.1

13841 RU1448

A

B

C

164

3.12 Glycogen content of free-living strains

As the glgA mutation prevented growth on glucose, strains were grown on

sucrose to assess the glycogen content of RU1448 and A34. Glycogen was not

detected in the mutant strain indicating the pathway for synthesis of glycogen had

been blocked by the transposon insertion (Table 3.7).

Table 3.7 Glycogen content of 10mM sucrose/5mM NH4-grown R. leguminosarum strains RU1448 & A34

Strain Glycogen (µg mg protein-1)

A34 70.16

RU1448 ND

ND not detected

3.13 Symbiotic phenotype of glycogen synthase mutant on pea

Peas were planted in a growth room in order to assess the symbiotic

performance of RU1448. The mutant nodulated the plants, forming healthy pink

nodules, as the wild type. There were no discernible differences in nodule size or

number between peas inoculated with RU1448 and wild type (data not shown).

Uninoculated plants did not become nodulated, and were yellow and stunted in

growth due to a deficiency in nitrogen.

165

Plants were harvested at the peak period of nitrogen fixation (during plant

flowering) (Sprent, 1982) to measure the acetylene reduction of the mutant.

There was no significant difference in nitrogenase activity in RU1448 compared

to the wild type (Table 3.8). Dry weights of plants were taken at flowering and

during the time when seeds were maturing. There was no significant difference in

the dry weight of peas inoculated with RU1448 compared to the parental strain

(Fig 3.31).

Table 3.8 Acetylene reduction (ethylene production) by A34 & RU1448. Values shown are means of 8 plants plus & minus the standard error of the mean.

Strain Ethylene production (µmoles plant1hr-1)

A34 3.70 ± 0.60

RU1448 3.38 ± 0.51

P>0.05

166

harvest (days)

dry

wei

ght (

g)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4 A5 RU1448

30 50

Fig 3.31 Dry weight of peas inoculated with A34 and RU1448 (A34, n=11; RU1448, n=13; P>0.05 at 50d harvest).

A34 RU1448

167

3.14 Discussion

In this chapter we aimed to gain some understanding of the role of PHB

metabolism in determinate and indeterminate symbioses. By mutating PHB

synthase in two isogenic strains of R. leguminosarum we investigated and

compared the relevance of bacteroid PHB metabolism to both the pea and the

bean symbiosis.

Perhaps the most intriguing results to emerge from this study were those relating

to the ultrastructural differences of the PHB mutant (RU1328) in the pea nodule.

The major zone of bacteroid differentiation (interzone II-III) lacks starch granules

in pea nodules infected by RU1328. Carbon demand from the endosymbiont

versus carbon made available by the plant to the nodule would dictate the levels

of starch accumulation in plant cells. In Dct mutants, where full bacteroid

differentiation is prevented and bacteroid metabolism does not represent a sink

for plant carbon, the accumulation of starch is much greater in the plant cells

infected by the mutant compared to the wild type (Ronson et al., 1981; Finan et

al., 1983; Arwas et al., 1985; Humbeck & Werner, 1989). We propose that the

absence of starch in plant cells of interzone II-III infected by RU1328 reflects an

increase in demand for plant carbon from the bacteroid due to the absence of

bacterial endogenous reserves. As seen in alfalfa (Paau & Cowles, 1978; Hirsch

et al., 1983; Vasse et al., 1990), we have shown that in pea, wild type bacterial

cells in the infection thread contain PHB and this disappears on differentiation of

cells to the bacteroid form. Therefore, cells forming indeterminate nodules must

utilise PHB, at least in part, to meet their requirements for carbon during bacterial

growth in the infection thread and / or bacteroid differentiation. We hypothesise

168

that cells unable to utilise PHB have an increased demand for plant carbon

during differentiation. The carbon source for bacterial proliferation in the infection

thread and for bacteroid development has not yet been established although

reports have suggested the role of certain pathways. For example, in S. meliloti

mutants in phosphoenolpyruvate carboxykinase (PEPCK) form nodules on alfalfa

that are reduced in acetylene reduction activity by 60% (Finan et al., 1991). Wild

type S. meliloti bacteroids lack PEPCK activity, therefore it has been suggested

the symbiotic defect resulting from the mutation occurs before mature bacteroid

formation (Finan et al., 1991). However, wild type R. leguminosarum bacteroids

have PEPCK activity therefore this is probably not the same case in this

symbiosis (McKay et al., 1985).

Under the circumstances of no endogenous bacterial reserves (PHB) the demand

and competition for plant carbon would be greater. Therefore the hypothesis that

bacterial PHB reserves are utilised to meet requirements for carbon during

bacterial growth in the infection thread and / or bacteroid differentiation affords an

explanation to the results of Willis & Walker (1998). The authors showed a

mutant in PHB synthase was out-competed by its parental strain when equal

numbers of cells were inoculated onto alfalfa (Willis & Walker, 1998). The ability

to metabolise PHB would advantage bacterial proliferation in the infection thread

and bacteroid differentiation, and would ensure the majority of nodules were

occupied by the wild type strain. Therefore, we intend to look at the competitive

ability of RU1328 to determine whether this is the case.

From the enzyme activities presented in Table 3.4 it can be seen that the activity

of CS is increased by a factor of 13 in pea bacteroids. The enzyme activities

169

reported here are consistent with a previous data for R. leguminosarum

bacteroids (McKay et al., 1989). Of the TCA cycle enzymes assayed in the

previous paper citrate synthase and aconitase are increased in activity in the

bacteroid by at least ten-fold (McKay et al., 1989). This up regulation of citrate

synthase (and aconitase) might be a factor in determining the absence of PHB

granules in pea bacteroids. It is feasible that the high activity of citrate synthase

represents a strong sink for acetyl-CoA and prevents its accumulation and / or

utilisation by β-ketothiolase. In Rhizobium sp. (Cicer) CC1192, which forms

indeterminate nodules on chickpea, there is no such increase in citrate synthase

activity (Kim & Copeland, 1996). However chickpea bacteroids have a higher

capacity for malate oxidation via malate dehydrogenase (x3.5) than free-living

cells. It has been suggested that this increase in MDH activity is a factor in

determining the fate of malate (and hence acetyl-CoA) in the bacteroid (Kim &

Copeland, 1996). A similar increase (x2.3) in MDH activity occurred in R.

leguminosarum bv. viciae (Table 3.4). If this were the case then we would expect

to see different levels of CS and MDH in A5 bacteroids in comparison to A34

bacteroids. It would be interesting to see the citrate synthase activity of A5 in

free-living and in bacteroid form to see if the increase in rate in the bacteroid form

is the same. Unfortunately we were not able to isolate bacteroids from bean by

the method used in the laboratory for pea bacteroids. Therefore we were not able

to carry out any enzymatic analysis that might have indicated why PHB

accumulates in bean bacteroids but not in pea. We routinely purify pea bacteroids

from plant tissue using a Percoll gradient of 57%. Using this gradient with

macerated bean nodules resulted in little separation of bacteroids from cytosol.

We increased the Percoll concentration to 70%, 80% and 90% but these

170

concentrations were still not appropriate. Therefore to study biochemical

differences between bacteroids from pea and bean nodules an appropriate

purification procedure for bean must first be established.

The results presented here indicate that blocking PHB metabolism in the R.

leguminosarum bv. phaseolus symbiosis has no effect on the overall efficiency of

the symbiosis. There was no difference acetylene reduction or dry weight of

plants infected with RU1329 compared to A5. This differs to the results presented

by Cevallos et al. (1996) who showed that acetylene reduction, dry weight and

the nitrogen content of aerial parts of bean were all significantly greater after 38

days growth in the plants infected by a R. etli PHB mutant (Cevallos et al., 1996).

This indicates that preventing PHB metabolism by some means prolongs nitrogen

fixation. However, the result of Cevallos et al. (1996) contradicts that of

Bergersen et al. (1991) who showed that active PHB metabolism prolonged

nitrogen fixation in soybean (Bergersen et al., 1991). Therefore we have not

resolved any of the apparent condradictions of PHB metabolism in determinate

symbioses.

We have shown that blocking PHB metabolism in R. leguminosarum bv. viciae

can reduce the efficiency of the pea symbiosis indicating PHB metabolism is

important even though the bacteroids do not accumulate large reserves of PHB.

This result differs to previous studies using PHB synthase mutants in S. meliloti

that have shown no differences in efficiency of symbioses (Povolo et al., 1994;

Willis & Walker, 1998; Cai et al., 2000). However, it is perhaps worth noting that

in these previous studies plants were grown under artificial light and at constant

temperature (ie. in a plant growth room). The results presented here are the first

171

reported of a mutant in PHB synthase whose symbiotic efficiency has been

tested in a large scale greenhouse trial where conditions are optimised for growth

of plants. However we are unsure of the reason for the dramatic effect in the first

experiment and why we were unable to repeat the result. It is possible that PHB

metabolism is required in pea bacteroids to regulate carbon metabolism and

prevention of this regulation may significantly impair the functioning of the

microsymbiont. However, this might only occur under certain conditions of plant

carbon supply to the bacteroid. As the two experiments only differed in the time of

planting it is likely that the effect of the mutation is strongly influenced by the

status of plant carbon metabolism.

The mutant in glycogen synthase (RU1448) did not alter the symbiotic efficiency

of pea suggesting glycogen metabolism is not an important factor in determining

the efficiency of nitrogen fixation, at least under the conditions in which the plants

were grown (in the growth room). The only other study of symbiotic performance

by a Rhizobium strain unable to synthesise glycogen is that of Marroqui et al.

(2001). The authors noted an increase in the dry weight of bean plants

inoculated with the glgA mutant. The plants with increased biomass also had an

increased number of nodules and increase nodule mass. Therefore the authors

concluded the enhanced symbiotic performance was due to increased nodulation

rather than an increased specific activity of nitrogen fixation in each nodule (see

section 1.4.5).

Glycogen accumulated to a lesser extent in the PHB mutant (RU1328) that was

defective in symbiosis. This suggests the carbon flux between PHB and glycogen

pools in the bacteroid may be closely linked and that glycogen metabolism may

172

be important in certain circumstances. If it is the case that glycogen is being

metabolised as an alternative pathway to PHB then one might expect a mutant

that is unable to metabolise glycogen, in addition to PHB, might be further

impaired in symbiosis efficiency. We intend to construct a double mutant in PHB

and glycogen synthase in A34 and to investigate the symbiotic performance on

pea.

173

CHAPTER 4

174

4 IDENTIFICATION OF THE ROLE OF ALANINE SYNTHESIS AND EXCRETION BY BACTEROIDS OF PEA

The role of the excretion of alanine has been subject of much debate due to the

apparent inconsistency between studies. No amino acids were excreted by

nitrogen fixing soybean bacteroids in flow chamber experiments carried out by

Bergersen and Turner (1990). In these experiments, where the aim was to

simulate the nodule environment in vitro by the maintenance of steady state

concentrations of oxygen and dicarboxylic acids and by the constant removal of

excretion products from the bacteroids, the sole product of nitrogen fixation was

ammonia (Bergersen & Turner, 1990; Li et al., 2001). The recent paper by

Waters et al. (1998) contradicts these findings of Bergersen. In this study alanine

was the only significant nitrogen containing compound excreted by nitrogen fixing

bacteroids of soybean. Exposure of the bacteroids to 15N2 gas resulted in the

alanine being highly enriched in label, whereas the little ammonium that was

seen was not enriched. The authors argued that the presence of ammonium in

the supernatant of the bacteroids arose from the de-amination of alanine upon

addition of, or by the contamination of the sample with, plant cytosol (Waters et

al., 1998).

Symbiosomes isolated from pea have been shown to rapidly excrete alanine

(Rosendahl et al., 1992), and alanine has been detected coming from R.

leguminosarum bv. viciae bacteroids incubated aerobically (Poole, pers. comm.).

Here we have looked at the nitrogen compounds excreted by pea bacteroids that

are actively fixing nitrogen. We aimed to clarify the role of alanine excretion and

to identify the conditions under which alanine excretion may occur. By isolating

175

and mutating the gene required for growth on L-alanine dehydrogenase we hoped

to identify the pathway for alanine synthesis in the bacteroid, to characterise this

mutation in free-living cells and bacteroids, and ultimately to determine the effect

this mutation in the pea symbiosis. We have shown that both ammonium and

alanine are excreted by pea bacteroids and that alanine is synthesised via L-

alanine dehydrogenase. Peas inoculated with the L-alanine dehydrogenase

mutant are Fix+ but six week old plants show a small but significant (20%)

decrease in dry weight. The reason for this decrease in dry weight is not certain.

It suggests that either alanine excretion contributes to the total fixed nitrogen

transferred to the plant and therefore the efficiency of the symbiosis or that

alanine metabolism may help regulate dicarboxylate metabolism.

The majority of the work in this chapter was done jointly by David Allaway and

myself. However the total amino acid analysis and 15N2 analysis was done by Les

Crompton and Richard Parsons respectively, and the construction of the aldA

mutant was done by Philip Poole and David Allaway. The results were presented

at the 12th International Congress on Nitrogen Fixation, Brazil in September 1999.

176

4.1 Isolation of pure bacteroid samples

In order to assess the products of nitrogen fixation it was necessary to obtain a

pure preparation of bacteroids that were still capable of fixing dinitrogen. The

Percoll gradient purification method (Reibach et al., 1981) was adopted as it

rapidly gave pure bacteroids with very little damage due to dehydration or

repeated pelleting and resuspension. The bacteroid fraction obtained (Fig 4.1)

was observed microscopically and showed no significant contamination with plant

debris.

Fig 4.1 Results of a typical percoll gradient centrifugation.

bacteroids

high density pellet (large plant material)

plant debris

177

Waters et al., (1998) suggested that ammonium only accumulated in bacteroid

supernatants when the sample was contaminated with plant cytosol and was the

result of deamination of excreted alanine by plant enzymes. To ensure that the

current study would identify nitrogen compounds that were solely excreted by the

bacteroid and that were not a product of enzymatic reactions occurring outside of

the bacteroid, 1mM L-alanine as added to bacteroid suspensions and the

supernatant was assessed for presence of ammonium and alanine. There was no

breakdown of added alanine, or increase in the initial level of ammonium (Fig

4.2).

Fig 4.2 Alanine and ammonium concentration in bacteroid supernatant spiked with or without 1mM alanine sampled over 2 hours.

time (minutes)

0 20 40 60 80 100 120 140

amm

onia

/ al

anin

e (n

mol

mg

prot

ein-1

)

0

50

100

150

200

250

300

350

alanine - 1mM alanine addedalanine - no alanine addedammonium - 1mM alanine addedammonium - no added alanine

178

4.2 Optimisation of conditions for nitrogen fixation

4.2.1 Oxygen concentration

The isolation of bacteroids was carried out in an argon atmosphere using buffers

that had been purged with argon to prevent the inactivation of nitrogenase by

oxygen. The bacteroids were supplied with oxygen on transfer to the

experimental vessel. In the nodule leghemoglobin buffers the external oxygen

concentration and maintains the oxygen tension experienced by the bacteroid low

at 10 – 40 nM (Vance & Heichel, 1991). In this study the oxygen tension

experienced by the bacteroids was not buffered by leghemoglobin, but was

dependant on the oxygen concentration in the gas phase above the bacteroid

solution and its transfer into the liquid phase by slow shaking of the experimental

vessel. As this supply of oxygen was a critical factor in maintaining a balance

between carbon metabolism and inactivation of nitrogenase in the bacteroids the

initial experiments optimised the oxygen concentration in the gas above the

bacteroid suspension.

Bacteroids were incubated at 28°C with slow shaking under a range of O2

concentrations from 0.1% to 1% and the efficiency of nitrogen fixation under the

different O2 concentrations was measured by ammonia excretion (Fig 4.3). The

oxygen concentration was also raised to 5% oxygen which caused a total loss of

ammonium excretion (data not shown) consistent with its synthesis being

dependant on nitrogenase activity. The rate of nitrogen fixation was optimum at

1% oxygen and this rate was comparable to those seen by other studies using

soybean bacteroids (Bergersen & Turner, 1967; Bergersen & Turner, 1990).

179

Fig 4.3 Effect of O2 concentration on the rate of ammonium excretion by bacteroids (3.1mg protein ml-1).

4.2.2 Carbon source

Isolated bacteroids have been shown to most actively fix nitrogen using organic

acids as an energy source (Bergersen & Turner, 1967), and as malate is the

principal carbon source delivered to the bacteroid to fuel nitrogen fixation during

symbiosis (see section 1.3.1), bacteroids were supplied with L-malate. To find the

optimum L-malate concentration for in vitro nitrogen fixation, initial experiments

measured ammonium production under various concentrations of L-malate (Fig

4.4). Under conditions of 1% oxygen the bacteroids were capable of fixing

nitrogen with no added malate, 1mM malate or 2mM malate, at the same rate. N2

fixation was inhibited by 5mM and 10mM malate. For the purpose of subsequent

experiments to assess alanine excretion, the concentration of 2mM malate was

considered the optimum as it was the upper concentration tested that gave a

oxygen (%)

0.1 0.2 0.5 1.0

amm

onia

pro

duct

ion

(nm

ol m

g pr

otei

n-1 h

our-1

)

0

5

10

15

20

25

180

good rate of nitrogen fixation. Therefore this would maintain a sufficient carbon

supply to the bacteroid over the long time courses that the experiments were to

run.

Fig 4.4 Production of ammonium by bacteroids under a range of malate concentrations (2.32 mg protein ml-1).

The inhibition of N2 fixation at high exogenous carbon concentrations (5mM and

10mM malate) and the ability to fix nitrogen when incubated with no exogenous

substrate has been reported previously in B. japonicum. Bergersen and Turner

(1990 a & b) showed a sharp decline in N2 fixation when bacteroids were

supplied with malate after a period of N2 fixation when supplied with no carbon

source. This was accompanied by an increase in demand for O2, efflux of CO2

and a hence a higher respiratory quotient (rate of CO2 efflux/rate of O2

time (minutes)

0 20 40 60 80 100 120

amm

onia

pro

duce

d (n

mol

mg

prot

ein-1

)

0

10

20

30

40

50

60

0 mM malate1 mM malate2 mM malate 5 mM malate10 mM malate

181

consumption). The authors suggested the exogenous carbon was used to build

up endogenous reserves which then supported nitrogen fixation (Bergersen &

Turner, 1990 a & b). This was supported by the fact that after the malate was

washed away there was a more sustained demand for O2 and sometimes

increased N2 fixation when compared to the rates before the initial supply of

malate. A further study showed concentrations of succinate or malate over

0.5mM led to the build up of PHB in soybean bacteroids, and that this became

the principle source of reducing power for nitrogen fixation when exogenous

carbon was withdrawn (Bergersen & Turner, 1990 a). Bergersen went on to show

that PHB in B. japonicum bacteroids was able to support nitrogen fixation in vitro

and reserves declined by 9.2% over 5 hours (Bergersen & Turner, 1992).

As in chickpea bacteroids (Lee & Copeland, 1994), we have shown that in pea

bacteroids PHB granules were not observed in transmission electron

micrographs of nodule sections (see section 3.5). However PHB has been shown

to be present in chickpea bacteroids by gas chromatographic analysis (Kim &

Copeland, 1996). The authors suggested the PHB was present in small amounts

such that substantial sized granules were not visable under the electron

microscope. Moreover, chickpea bacteroids retain the enzymatic capacity for

PHB synthesis and breakdown (Kim & Copeland, 1996). In R. etli free-living cells,

PHB is continually synthesised and degraded even under conditions in which the

polymer does not build up (Encarnacion et al., 1995). Therefore PHB may be

continually cycled in the bacteroids of indeterminate nodules such that a

significant reserve might not accumulate. It has been reported that for the same

specific rate, nitrogenase activity supported by endogenous substrates requires a

lower pO2 than when supported by malate (20% of the malate driven nitrogenase

182

activity) (Appels & Haaker, 1991). Therefore cycling carbon through PHB, rather

than cycling directly through the TCA might be a respiratory adaption in

bacteroids maintained under low oxygen.

To ascertain whether PHB metabolism in pea bacteroids was an important factor

in maintaining bacteroid nitrogen fixation, we conducted an experiment to assess

ammonium production by RU1328 (the PHB mutant) and the parental strain

under conditions of 0, 1, 2, 5, 10mM malate. If PHB metabolism in pea bacteroids

was required to support N2 fix under concentrations of reduced carbon supply

then the mutant would not be expected to fix N2 under conditions of no

exogenous malate. If carbon was cycled through PHB to effectively regulate

metabolism under microaerobic conditions then the mutant might be expected to

fix nitrogen less efficiently than the wild type on 1, 2, 5, 10mM malate. We were

also interested to see levels of alanine excreted from the mutant bacteroids.

Unfortunately we only had one greenhouse harvest with which to attempt this

experiment and on this particular day the experiment failed to work.

183

4.3 Ammonium and alanine are excreted from pea bacteroids as products of N2 fixation

Under the optimised conditions (1% O2, 2mM malate, pH7.4) alanine was not

excreted by bacteroid suspensions of 2 - 3 mg protein ml-1. Ammonium was the

only nitrogen compound excreted in significant amounts from bacteroids at an

average rate of 21.2 nmol hr-1 mg protein-1 (SE = 2.2, n = 14). In these

experiments the starting ammonium concentration was between 0.13mM and

0.16mM. The final concentration rose to between 2.4mM and 2.8mM over 1.75

hours. The in vivo bacteroid ammonium concentration has been estimated to be

12mM for B. japonicum bacteroids (Streeter, 1989). The purification procedure

washes away the majority of ammonium from the medium in which the bacteroids

are held, and would dilute out cytosolic concentrations. Therefore to compensate

for the removal of bacteroid ammonium, the initial bacteroid ammonium

concentration was raised by increasing the bacteroid concentration. This would

return the ammonium concentration experienced by the bacteroid to more

physiologically relevant ammonium concentrations at the start of the experiment.

The higher density of bacteroids would also lead to a more rapid accumulation of

ammonium in the medium. Waters et al. (1998) used 8mg dry weight bacteroids

per ml. This would be approximately 16mg protein ml-1, therefore concentrating

the bacteroids to approximately 10mg protein ml-1 in our experiments was

considered a valid approach. The time course was also extended as previous

experiments had shown isolated bacteroids were capable of linear nitrogen

fixation for up to 4 hours (data not shown). Under these modified conditions,

which approximately doubled the initial ammonium concentration, both

ammonium and alanine were excreted linearly over 4 hours from bacteroids at an

184

average rate of 19.7 nmol hr-1 mg protein-1 (SE = 2.0, n = 5) and 7.8 nmol hr-1 mg

protein-1 (SE = 0.6, n = 8) respectively (Fig 4.5). The rate of ammonium excretion

was not significantly different when bacteroids also secrete alanine. Alanine

excretion did not reduce the rate of ammonium excretion. This suggested alanine

excretion was not an alternative route for nitrogen transfer out of the bacteroid,

rather a supplementary one.

Fig 4.5 A representative bacteroid excretion assay showing both ammonium and alanine were produced at linear rates by high density bacteroids (9.6mg protein ml-1). Initial ammonium concentration = 0.28mM, final = 1.2mM.

time (hours)

0 1 2 3 4

prod

uct f

orm

ed (n

mol

mg

prot

ein-1

)

0

20

40

60

80

100

120

ammoniumalanine

185

4.3.1 Alanine is a product of de novo synthesis

It was confirmed by incubating a high density preparation of bacteroids under 1%

O2 and 99 atom percentage 15N2 that both the ammonium and alanine that

accumulated in the bacteroid supernatant came from nitrogen fixation. The initial

ammonium pool was 0.66mM, while the final ammonium pool was 0.99mM. After

subtraction of the initial ammonium concentration (which had a natural

abundance level of 15N label), newly released ammonium had an 15N enrichment

of 90.4 (SE = 9.8, n = 4) atom percentage excess confirming that it was the

product of N2 fixation. The alanine had an 15N enrichment of 21.5 atom

percentage excess (SE = 9.8, n = 4). The initial ammonium pool only labelled

with natural abundance 15N therefore the initial alanine synthesised would use

ammonium from this pool, and would gain higher levels of label over time as the

proportion of 15N-ammonium built up due to fixation of labelled N2. Therefore the

results were consistent with the alanine being a product of de novo synthesis, the

ammonium pool being used for amination of carbon.

4.3.2 Alanine is the sole amino acid excreted

Total amino acid analysis of the excretion products showed that alanine was the

only amino acid excreted in significant amounts from the bacteroids (data not

shown).

186

4.4 Alanine production is dependant on the ammonium concentration

The N2 fixation rate of bacteroids in low density and high density preparations is

comparable (approximately 20 nmol hr-1 mg protein-1 in each case). Therefore the

actual accumulation of ammonium in a given volume of bacteroid suspension,

hence the ammonium concentration, would rise more rapidly in the dense

preparations. Alanine was only excreted by high density bacteroids, therefore it

seemed plausible that its synthesis and excretion in this system was dependent

on ammonium concentration.

To test this, ammonium was titrated into low density bacteroid preparations

incubated at different malate concentrations (Fig 4.6). With no addition of malate

there were no significant levels of alanine produced, however the small amounts

produced were proportional to the NH4Cl concentration. With 1mM and 10mM

malate alanine was significantly produced by bacteroids supplemented with

NH4Cl and its production was proportional to the ammonium concentration in all

cases. Bacteroids produced no significant amounts of alanine when incubated

with either 1mM or 10mM malate and no ammonium chloride.

Bacteroids incubated with 10mM malate produced more alanine for a given

concentration of NH4Cl than those bacteroids incubated with 1mM malate (for

example, 79.5 nmole mg protein-1 compared to 46.6 nmole mg protein-1 after 2

hours for bacteroids given 10mM NH4Cl and incubated with 10mM malate and

1mM malate respectively). Therefore, excretion of alanine was enhanced by

higher malate concentrations, presumably by the increased provision of carbon

skeletons for alanine synthesis.

187

To confirm the nitrogen component of the alanine excreted had come from the

ammonium pool, low density bacteroids were incubated in 10mM and 20mM

15NH4Cl (10 atom percentage excess) and supernatant samples were removed

after an hour. The alanine formed was 9.6 and 10.1 atom percentage excess 15N

for 10mM and 20mM respectively confirming it was the product of de novo

synthesis.

188

Fig 4.6 Production of alanine from low density bacteroids bacteroids (3.0mg protein ml-1) supplemented with (A) no malate; (B) 1mM malate; and (C) 10mM malate and ammonium (at concentrations shown).

time (minutes)

0 20 40 60 80 100 120 140

alan

ine

(nm

ol m

g pr

otei

n-1)

0

20

40

60

80

100

time (minutes)

0 20 40 60 80 100 120 140

alan

ine

(nm

ol m

g pr

otei

n-1)

0

20

40

60

80

100

time (minutes)

0 20 40 60 80 100 120 140

alan

ine

(nm

ol m

g pr

otei

n-1)

0

20

40

60

80

100A

B

C

0mM NH4

2mM NH4

5mM NH4

10mM NH4

189

4.5 Alanine synthesis is not directly coupled to N2 fixation

The activity of L-alanine dehydrogenase in nodule tissue is exclusive to the

bacteroid cytoplasm (Dunn & Klucas, 1973). Its activity is high and increases

during the period of nodule development when nitrogenase also increases

(Werner & Stripf, 1978; Werner et al., 1980). The activity of L-alanine

dehydrogenase is 4 times higher in free-living B. japonicum cells that fix nitrogen

compared to cells that do not fix nitrogen, and is comparable to that found in

bacteroids (Werner & Stripf, 1978). This suggests that L-alanine dehydrogenase

activity is linked to nitrogenase activity. If alanine synthesis was dependent on

nitrogenase activity in R. leguminosarum, bacteroids isolated in air would be

expected to have lost the ability to synthesise and secrete alanine. Bacteroids

isolated under air did not fix nitrogen as measured by ammonium release.

However when bacteroids were incubated in the presence of ammonium, alanine

was excreted at 36.56 nmol mg protein-1 (bacteroids incubated with 2mM malate

and 10mM NH4Cl) (Fig 4.7). Thus alanine is an assimilation product of

ammonium but its synthesis was not dependant on nitrogen fixation.

190

Fig 4.7 Alanine production by bacteroids isolated in air incubated with 10mM NH4Cl or 2mM malate or both.

time (minutes)

0 20 40 60 80 100 120 140

alan

ine

(nm

ol m

g pr

otei

n-1)

0

10

20

30

40

50

60

no addition10mM NH4Cl2mM malate10mM NH4Cl & 2mM malate

191

4.6 Identification of the pathway of alanine synthesis in the bacteroid

It was apparent that alanine was excreted from bacteroids of pea at a significant

rate when presented with a suitable supply of carbon source and ammonium as

would occur in planta. As the products of N2 fixation by isolated bacteroids

depend on numerous factors, for example density of cells, malate concentration

and ammonium concentration, it would be incorrect to make physiological

conclusions from these in vitro biochemical experiments. Therefore it was

necessary to identify and mutate the pathway of alanine synthesis to determine

what occurs in planta.

If the alanine excreted by R. leguminosarum bacteroid preparations was

synthesised by L-alanine dehydrogenase then the affinity of this enzyme for

ammonium would be expected to be similar to that of alanine synthesis by

bacteroids. Using a crude total enzyme preparation, the apparent Km of L-alanine

dehydrogenase for ammonium was calculated to be 5.1mM. To obtain the

equivalent Km for alanine excretion by isolated bacteroids, a low density

preparation (1.7mg protein ml-1) was supplemented with a range (0 – 20mM

NH4Cl) of ammonium chloride concentrations and the production of alanine was

assessed at 1 hour (Fig 4.8). The apparent Km calculated for alanine excretion by

isolated bacteroids was similar to the enzyme preparation at 3.2mM.

192

L-Alanine dehydrogenase exists in several microorganisms but usually has a

high Km for NH4+, for example 30mM in Bacillus cereus (Porumb et al., 1987).

The results presented here are consistent with kinetic studies of L-alanine

dehydrogenase from other rhizobia. L-Alanine dehydrogenase from both R. lupini

and B. japonicum has an apparent Km for NH4+ under 10mM, and therefore within

the NH4+ concentration expected to be found in bacteroids (Dunn & Klucas, 1973;

Smith & Emerich, 1993 a & b). The kinetic data suggested that L-alanine

dehydrogenase is the primary route for alanine biosynthesis in the bacteroid and

therefore it was appropriate to isolate and mutate the enzyme in R.

leguminosarum.

Fig 4.8 Alanine excretion from a low density preparation of bacteroids incubated for 1 hr at various ammonium concentrations with malate at 2mM. The apparent Km for ammonium was 3.2mM and the Vmax was 95.8 nmol hr-1 mg protein-1.

Ammonia (mM)

0 5 10 15 20 25

Ala

nine

(nm

ole

((m

g pr

otei

n)-1

) h-1

)

0

20

40

60

80

100

Km 3.2 mMVmax 1.6 nmol (mg protein)-1 min-1

193

4.7 Isolation of mutants in alanine metabolism

The following work on isolation of mutants in alanine metabolism was done by

Philip Poole and David Allaway and is summarised here for understanding of the

work presented in the rest of this chapter and in chapter 5.

4.7.1 Isolation of mutant RU1275 unable to grow on alanine

To isolate a mutant in L-alanine dehydrogenase approximately 15,000 Tn5

mutants of 3841 were screened for growth on alanine as the sole carbon source.

One strain did not grow on alanine and was designated strain RU1275. The

transposon and flanking DNA was cloned as an EcoRI fragment into pBC

(pRU549) in order to identify the transposon insertion site by sequencing. Tn5

was located in a gene with high identity to a putative transcriptional regulator of

Rhizobium sp. NGR234 (89% amino acid identity, EMBL accession number

AE000097). The transposon was located in the N-terminus of the gene and had a

9 bp repeat of CATCAAGAT. The mutation was complemented for growth by

cosmid pRU3131. To identify what the gene might regulate, 3.6 Kb DNA was

sequenced directly from cosmid pRU3131. Divergent from the regulator were two

genes: the first showed 49% amino acid identity to dadX (alanine racemase) of

Rickettsia prowazekii (EMBL accession number AJ235270) and the second

showed 74% amino acid identity to the dadA (D-alanine dehydrogenase) gene of

Pseudomonas aeruginosa (EMBL accession number AE004943). The regulator

gene was therefore designated dadR. The dad operon is the principle alanine

catabolic operon in several bacteria (Wild & Klopotowski, 1981; McFall &

Newman, 1996; Reitzer, 1996; Janes & Bender, 1998). Indeed, the complete loss

of growth of RU1275 on alanine suggested that products of the dad operon

194

constituted the principal alanine catabolic pathway in R. leguminosarum strain

3841.

4.7.2 Isolation of multicopy aldA as a suppressor of strain RU1275

The screening of 3841 Tn5 mutants failed to isolate a mutant in L-alanine

dehydrogenase (AldA). However, the mutant strain RU1275 was rescued for

growth on alanine as the sole carbon source by a second cosmid pRU3135. To

identify the gene responsible, pRU3135 was mutagenised with TnB20.

Mutagenesis identified a cosmid (pRU3138) unable to rescue growth of RU1275

on alanine. To identify which gene the transposon was inserted in, and hence the

gene responsible for supressing the Dad mutation, DNA adjacent to the

transposon was cloned as a 6 Kb EcoRI fragment into pSK (pRU593). DNA was

sequenced using a primer that bound to the IS50 region of the transposon. The

transposon was inserted in a gene with high identity to L-alanine dehydrogenase

(aldA) of Vibrio proteolyticus (65% amino acid identity, EMBL accession number

AF070716). The transposon was located in the N-terminus of the gene and had a

9 bp repeat of CCTCGTGGC. It was orientated such that LacZ was inactive.

4.7.3 Construction of a mutation in L-alanine dehydrogenase (aldA)

An aldA mutant (RU1327) was constructed by recombination of pRU3138 into the

chromosome of 3841 using pPHJI1 (Ruvkun & Ausubel, 1981). The mutant

lacked any detectable AldA activity when grown on glucose, succinate, or alanine

(Table 5.1) and was confirmed genetically by Southern blot and hybridisation

(data not shown). The mutant grew as the wild type on alanine as sole carbon

source, indicating that AldA was not a catabolic enzyme in R. leguminosarum

strain 3841.

195

4.8 Effect of mutation RU1327 on plant symbiotic performance

Strain RU1327 was used to inoculate pea plants grown in 2 L pots in the growth

room. The strain nodulated pea plants forming healthy pink nodules, which were

of the same total mass and number as the wild type (data not shown).

Uninoculated plants did not become nodulated, and were yellow and stunted in

growth due to a deficiency in nitrogen. Bacteroids of RU1327 isolated from the

nodules excreted ammonium at normal rates (21.1 nmol hr-1 mg protein-1) but did

not secrete alanine from relatively high density preparations (3841 - 4.80 mg

protein ml-1, RU1327 - 4.03 mg protein ml-1) with exogenous ammonium present

(Fig 4.9).

Fig 4.9 Alanine produced by bacteroids of RU1327 and 3841 over 2 hours when supplemented with 10mM NH4Cl and 2mM or 10mM malate.

time (hours)

0 30 60 90 120

alan

ine

(nm

ol m

g pr

otei

n-1)

0

20

40

60

80

3841 10mM NH4/2mM malate3841 10mM NH4/10mM malateRU1327 10mM NH4/2mM malate RU1327 10mM NH4/10mM malate

196

Furthermore, AldA activity was undetectable in mutant bacteroid protein extracts,

whereas for the wild type the activity was 0.09µmol min-1mg protein-1. These data

confirmed that alanine biosynthesis was predominately via AldA in the bacteroid.

Plants were harvested at flowering and showed no significant difference in

acetylene reduction between the mutant and the wild type (11.1, SE=1.0 and 9.4,

SE=1.4 µmol g wet weight nodule-1 respectively). At the second harvest at six

weeks the peas inoculated with RU1327 were 20% reduced in total shoot dry

weight compared to those inoculated with 3841 (Fig 4.10).

Fig 4.10 Average dry weight (g) of peas inoculated with RU1327 (n=56) and 3841 (n=54) harvested at 6 weeks.

The plants inoculated with RU1327 were not unhealthy and showed no signs of

nitrogen deficiency. Therefore, transfer of ammonium alone appears to meet the

plants’ requirement for nitrogen. The rate of ammonium excretion was not

significantly different when bacteroids also excreted alanine (section 4.3).

Therefore alanine synthesis via AldA, and its excretion from the bacteroid, either

dry

wei

ght (

g)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

3841 RU1327

197

supplements the total nitrogen transported out of the bacteroid, and contributes to

the efficiency of symbiotic nitrogen fixation in pea, or has some other role (for

example regulation of metabolism).

198

4.9 Discussion

In this chapter we aimed to determine the nitrogenous compounds excreted from

pea bacteroids that had been isolated using Percoll gradients. We further aimed

to clarify the role of alanine excretion and to identify the conditions under which

alanine excretion may occur. We have shown that both ammonium and alanine

are excreted by bacteroids of R. leguminosarum in symbiosis with pea and the

major factor that determines the relative amounts of nitrogen compounds

excreted is the absolute ammonium concentration. At low concentrations of

ammonium, alanine is not produced in significant amounts but as the

concentration of ammonium is raised bacteroids excrete alanine. We have shown

the nitrogen component of the excreted alanine comes from the ammonium pool

and have confirmed that fixed nitrogen is directly incorporated into alanine using

labelling studies. The results presented here may explain the disparity between

the studies of Bergersen and Turner (1990) and Waters et al. (1998). In

Bergersen’s flow chamber experiments there was a constant removal of excretion

product from the bacteroids. Under these conditions the only excretion product of

nitrogen fixing soybean bacteroids is ammonium. We suggest the constant

removal of ammonium from the flow chamber results in insufficient ammonium

accumulation to activate alanine synthesis. Conversely, in the system studied by

Waters et al. (1998) where bacteroids are resuspended at very high densities

(approximately 16mg protein ml-1) the ammonium accumulation from nitrogen

fixation would be extremely high and in this situation alanine synthesis and

excretion may predominate. Therefore fundamental differences in assay

199

conditions between the studies of Bergersen (1990) and Waters et al. (1998) are

likely to have produced the differing results.

Total amino acid analysis of the excretion products from pea bacteroids showed

that alanine was the sole amino acid excreted. Rosendahl (1992) showed that

pea symbiosomes incubated with malate excreted a significant amount of alanine

together with aspartate (Rosendahl et al., 1992). As we did not detect aspartate

from bacteroids this suggests that aspartate might be produced in the

peribacteroid space then excreted. However R. lupini ‘naked’ bacteroids have

been shown to excrete both alanine and aspartate when incubated with

succinate, malate or fumarate (Kretovich et al., 1986). Therefore we are unsure

why aspartate was not detected in our study. It might be that the assay conditions

did not favour aspartate formation in these bacteroids. To ascertain whether

alanine and aspartate are excreted by symbiosomes formed by R.

leguminosarum strain 3841 similar experiments to those presented here will be

carried out using peribacteroid units. The PBM can be easily removed from

isolated symbiosomes by vortexing. Therefore the differences in compounds

excreted by the symbiosome and the bacteroid can be assessed in the same

experiment. However, the isolation procedure for symbiosomes requires

considerable adaptation to the current technique we use in the laboratory for

isolating bacteroids. For an equivalent yield of symbiosomes, a greater amount of

starting material (ie. nodules) is required because the procedure demands a

more delicate maceration of tissue. This means the scale of the pea harvest

would have to be much greater for these experiments. To circumvent the

problems of harvest size experimental conditions will also be modified so that

symbiosomes are incubated with 14C-malate. This will allow detection of products

200

from small volumes of symbiosomes. As there was not enough time during the

period of this research to carry out these experiments they are scheduled for the

summer period 2002.

We have shown that mutation of L-alanine dehydrogenase (aldA) abolishes AldA

activity in free-living cells and bacteroids and prevents alanine excretion by

isolated bacteroids. Therefore the pathway of alanine synthesis in pea bacteroids

is via L-alanine dehydrogenase. Peas inoculated with the aldA mutant are Fix+ as

measured by acetylene reduction, although six week old plants show a significant

(20%) decrease in dry weight. This shows that alanine cannot be the sole

nitrogen excretion product from pea bacteroids as suggested by Waters et al.

(1998) for soybean bacteroids. The reason for the 20% decrease in dry weight is

not certain. The data is consistent with alanine excretion contributing to the

transfer of nitrogen to the pea plant and hence the overall efficiency of the

symbiosis. However, we cannot rule out the possibility that the decrease in

symbiosis efficiency we have reported in the AldA mutant is the result of a

decrease in the efficiency of carbon metabolism, hence delivery of energy and

reductant to nitrogenase, in the bacteroid. Alanine synthesis via AldA consumes

NADH and the reaction might be involved in regulation of the TCA cycle by

removing reductant and carbon away from the cycle. To use dicarboxylates as

sole carbon source bacteroids have to make acetyl-CoA via pyruvate which is the

precursor of alanine. Therefore it would not be surprising if the consumption of

pyruvate by pyruvate dehydrogenase or L-alanine dehydrogenase were to play a

pivotal role in the direction of carbon flux to acetyl-CoA or alanine depending on

the status of the TCA cycle.

201

However, that alanine excretion supplements nitrogen export from wild type

bacteroids suggests a defined role for alanine synthesis and excretion in nitrogen

metabolism of bacteroids. It might be that alanine synthesis (and excretion)

serves two roles that are integrally linked: the regulation of the TCA cycle

maintains optimum functioning of nitrogenase and the provision of an alternative

route for nitrogen assimilation and export increases the transfer rate out of the

bacteroid.

A possible way to assess whether alanine synthesis and excretion is involved in

the regulation of the TCA cycle is to study alanine excretion in free-living cells

under a range of oxygen concentrations. Encarnacion (1995) showed that alanine

(along with a range of other amino and organic acids) is excreted from R. etli

after successive sub-cultures on minimal medium without biotin (Encarnacion et

al., 1995). This condition is thought to be a fermentative-like response that

mimics low oxygen environments by reducing TCA enzyme activity and flow of

carbon through the TCA cycle. However, under these circumstances PDH is

severely inhibited by the lack of biotin, so these results should be treated with

caution. In the same study the effect of low oxygen conditions on PHB was

studied, but no mention was made on alanine excretion under the same

conditions. We have not as yet looked at alanine excretion from free-living

bacteria the laboratory but this may add to our understanding of its role in the

bacteroid.

202

CHAPTER 5

203

5 REGULATION OF L-ALANINE DEHYDROGENASE

This chapter focuses on the regulation of L-alanine dehydrogenase, the enzyme

we have shown is operative during symbiosis in the synthesis of alanine (chapter

4). During root nodule development certain key genes involved in nitrogen

fixation and assimilation exhibit enhanced levels of expression. For example,

transcript levels of glutamate synthase and glutamine synthetase increase in the

plant cytosol during nodule development (Gregerson et al., 1993; Roche et al.,

1993). We have confirmed in section 4.5 that AldA activity is not dependent on

nitrogenase activity. To ascertain under what conditions L-alanine dehydrogenase

activity is induced we have studied its activity in free-living cells under a range of

growth conditions.

We have identified a putative regulator transcribed divergently to aldA and have

confirmed its role in the regulation of aldA. We have made transcriptional fusions

to aldA and aldR to confirm the transcriptional regulation of aldA by AldR and to

determine the levels of aldA induction on a range of carbon sources. The

transcriptional fusions were made in pJP2 which is derived from the plasmid

pTR101. This plasmid carries the GUS reporter gene and is stable in the nodule

even over prolonged incubations of more than 6 weeks (Boesten, pers. comm.).

Therefore its GUS marker activity and plasmid stability make this vector ideal for

examining the induction of aldA during the course of nodule development.

Indeterminate nodules, such as those of pea, have distinct zones of development

due to the nature of their growth (section 1.2.2.2) and we had hoped to establish

whether AldA activity is restricted to the mature nitrogen fixing zone of pea

nodules or whether activity is switched on earlier during bacteroid development.

204

However, results of experiments determining the levels of gene induction on

different carbon sources suggested we adopt an alternative strategy to carry out

these plant studies.

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5.1 Biochemical analysis of L-alanine dehydrogenase activity

To ascertain the induction pattern of aldA, L-alanine dehydrogenase activity was

assessed in 3841 grown on various carbon sources (Table 5.1). There was no

detectable activity in cells grown on glucose. Activity was induced on when the

cells were grown on the carboxylic acids succinate, malate or pyruvate. The rate

of enzyme activity on these carbon sources was similar to that seen in bacteroids

(0.09µmol min-1mg protein-1 from section 4.8) suggesting transcription of aldA

might be regulated in response to carbon source in planta.

Cells grown on alanine showed the highest level of induction. Cells grown on

glucose with alanine (10mM) as the nitrogen source were induced for AldA

activity but this was not as high as when grown on alanine (20mM) as the sole

carbon and nitrogen source. This high level of induction is surprising as the

primary physiological role for AldA is proposed to be that of aminating pyruvate.

Mutant RU1327 showed no L-alanine dehydrogenase activity when grown on

glucose, succinate or glucose/alanine, and RU1422 showed no L-alanine

dehydrogenase activity when grown on glucose/alanine. This indicated that there

was only one copy of the gene encoding L-alanine dehydrogenase in R.

leguminosarum.

To complement the aldA mutation, the aldA gene was amplified by PCR using

primers P199 and P200 giving a 1.45 Kb fragment that was cloned into pCR®2.1-

TOPO (plasmid pRU640). The gene was subcloned from pRU640 as a 1.45 Kb

SacI / KpnI fragment into the broad host range plasmid pTR101 producing

pRU708. This plasmid was conjugated into RU1327 to give strain RU1371. The

206

presence of a full length copy of aldA on a plasmid restored L-alanine

dehydrogenase activity in cells grown on succinate, glucose / alanine and 20mM

alanine (Table 5.1). However the level of activity was not restored to wild type

levels when grown on glucose / alanine or 20mM alanine. This suggests that

something becomes limiting and transcriptional work suggests it might be levels

of the regulator aldR (see section 5.4).

Table 5.1 L-Alanine dehydrogenase activity of 3841, RU1327, RU1371 and RU1422 grown on various carbon sources.

L-Alanine dehydrogenase activity (µmol min-1mg protein-1)

3841 RU1327 RU1371 RU1422

Glucose/NH4 nd nd 0.014 (0.004) -

Succinate/NH4 0.074 (0.014) nd 0.262 (0.023) -

Glucose/Alanine 0.494 (0.015) nd 0.204 (0.030) nd

20mM Alanine 0.770 (0.022) - 0.261 (0.021) -

Malate/NH4 0.084 (0.003) - - -

Pyruvate/NH4 0.079 (0.004) - - -

Glucose/Succinate/NH4

0.066 (0.006) - - -

nd – no activity detected

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5.2 Analysis of the sequence of aldA (L-alanine dehydrogenase)

Sequencing directly from pRU3135 (the wild type Ald cosmid) identified a putative

transcriptional regulator, designated aldR, transcribed divergent to aldA in 3841

(Fig 5.2). AldR showed 63% amino acid homology to a putative transcriptional

regulator in Streptomyces coelicolor (EMBL accession number AJ131213) and

58% amino acid homology to bkdR, the bkd operon transcriptional regulator, of

Pseudomonas putida (EMBL accession number AE004943). Both regulators

belong to the AsnC family of bacterial transcriptional regulators, which also

includes Lrp of E. coli (Willins et al., 1991). Bacterial transcription regulation

proteins which bind DNA through a helix-turn-helix motif can be classified into

subfamilies on the basis of sequence similarities. The helix-turn-helix DNA-

binding motif of proteins in the AsnC family is located in the N-terminal part of the

sequences. Analysis of the translated protein a putative helix-turn-helix motif at

22-48 amino acids (Fig 5.1).

Fig 5.1 Translation of aldR showing helix-turn-helix motif located in the N-terminus of the protein in red.

MADLDTIDLAILRVLQANARITNAELAERIGLSPSACSRRLDILERSGVIGGYHARLSHKALDYKMIAIVHISLSGQFAKTLAEFEAAVKLCPNVLVCYLMSGEYDYILRVAARDLEDYERIHRDWLSALPHVVKINSSFALREIIDKPNVGL*

208

Analysis of the DNA sequence highlighted two potential ATG start codons for

aldA which were separated by 15bp (Fig 5.2). Alignments of this sequence

against AldA from B. japonicum, M. loti and S. meliloti show the ATG giving the

shorter peptide is the most likely start codon (Fig 5.3). Primer extension analysis

would be required to confirm the correct transcriptional start site but it was

decided that this analysis was beyond the scope of this study. The intergenic

region of aldA and aldR contains an area of thymine rich residues (Fig 5.2). This

region might be important in binding of the AldR to DNA as has been shown in

the consensus sequence for the binding of Lrp to DNA (Calvo & Matthews, 1994).

209

Fig 5.2 Sequence obtained from pR3135. Restriction sites shown in pink (EcoRI); green (BamHI). 9bp repeat of RU1327 in red. P199, P200, P219 and P220 priming sites shown in yellow.

Sequence: aldA_R sequence1.doc

210

211

1 60 B. jap. AldA (1) MKVGVPKEIKAHEYRVGLTPGAAREYVAAGHRVMIETNAGAGIGATDGDYRNAGATILTS R. leg. AldA (1) MRVGCPKEIKNHEYRVGLTPASVREYVAHGHEVWVETKAGVGIGADDAAYAAAGAKIAAS M. l. c AldA (1) MRVGCPKEIKNHEYRVGLTPGSVREYVAHGHEVLVEAGAGAGIGADDNAYRAAGATIAKT M. l. p AldA (1) MRVGCPKEIKNHEYRVGLTPGSVREYVAHGHEVLVETGAGAGIGADDNAYRAAGATIAKT S. mel. AldA (1) MRVGCPKEIKNHEYRVGLTPGSVREYVAHGHEVIVETKAGAGIGADDDSYRAAGARIVPT Consensus (1) MRVGCPKEIKNHEYRVGLTPGSVREYVAHGHEVLVET AGAGIGADD AYRAAGATIA T 61 120 B. jap. AldA (61) AAEVFASSEMIVKVKEPQPAEWSQLREDQILFTYLHLAPDPEQAAGLLKSGCIAIAYETV R. leg. AldA (61) AKDIFEKCDMIVKVKEPQPAEWAQLRDGQLLYTYLHLAPDPEQTKGLIASGVTAIAYETV M. l. c AldA (61) AADVFAKSDMIVKVKEPQPNEWVQLRDGQILYTYLHLAPDPEQTKGLLASGVTAIAYETV M. l. p AldA (61) AADVFAKSDMIVKVKEPQPNEWVQLRDGQILYTYLHLAPDPEQTKGLLASGVTAIAYETV S. mel. AldA (61) AREVFEKADMIVKVKEPQPSEWAQLREGQILYTYLHLAPDPEQTQGLLKSGVTAVAYETV Consensus (61) AADVFAKSDMIVKVKEPQPAEWAQLRDGQILYTYLHLAPDPEQTKGLLASGVTAIAYETV 121 180 B. jap. AldA (121) TDAHGGLPLLAPMSEVAGRLSIEAAGSALKRSTGGRGLLIGGVPGVQPARIVVIGGGVVG R. leg. AldA (121) TDERGGLPLLAPMSEVAGRLSIQAGATALQKANGGLGVLLGGVPGVLPAKVAVIGGGVVG M. l. c AldA (121) TDDRGGLPLLAPMSEVAGRLSIQAGATALQKANGGRGVLLGGVPGVLPGKVTVLGGGVVG M. l. p AldA (121) TDDRGGLPLLAPMSEVAGRLSIQAGATALQKANGGRGVLLGGVPGVLPGKVTVLGGGVVG S. mel. AldA (121) TDERGGLPLLAPMSEVAGRLAIQAGATSLQKANGGRGILLGGVPGVLPAKVAIIGGGVVG Consensus (121) TDDRGGLPLLAPMSEVAGRLSIQAGATALQKANGGRGVLLGGVPGVLPAKV VIGGGVVG 181 240 B. jap. AldA (181) THAARMAAGLGAEVTIIDRSIIRLRELDELFEGRVRTRFSTIESVEEEVFAADVVIGAVL R. leg. AldA (181) LHAARMAAGLGADVSILDKSLPRLRQLDDIFAGRIHTRYSSIQALEEEVFSADLIIGAVL M. l. c AldA (181) LHAARMAAGLGADVTIIDRSIPRLRQLDDLFAGRVHTRYSTVEALEEECFSADIVVGAVL M. l. p AldA (181) LHAARMAAGLGADVTIIDRSIPRLRQLDDLFAGRVHTRYSTVEALEEECFSADIVVGAVL S. mel. AldA (181) LHAAKMAAGLGADVSILDRSLPRLRQLDDIFNGRVHTRYSTIDALEEEVFSADMVIGAVL Consensus (181) LHAARMAAGLGADVTIIDRSIPRLRQLDDLFAGRVHTRYSTIEALEEEVFSADIVIGAVL 241 300 B. jap. AldA (241) VPGASAPKLVRRSMLSSMRKRAVLVDVAIDQGGCFETSRPTTHADPTYEVDGIIHYCVAN R. leg. AldA (241) IPGAAAPKLVTREMLSGMKKGSVIVDVAIDQGGCFETSHATTHSDPTYEVDGVVHYCVAN M. l. c AldA (241) IPGAAAPKLVTREMLSGMKKGSVLVDVAIDQGGCFETSHATTHAEPTYEVDGVIHYCVAN M. l. p AldA (241) IPGAAAPKLVTREMLSGMKKGSVLVDVAIDQGGCFETSHATTHAEPTYEVDGVIHYCVAN S. mel. AldA (241) IPGAAAPKLVTREMLSAMKKGAVIVDVAIDQGGCFETSHATTHSEPTYEVEGIVHYCVAN Consensus (241) IPGAAAPKLVTREMLSGMKKGSVLVDVAIDQGGCFETSHATTHAEPTYEVDGVIHYCVAN 301 360 B. jap. AldA (301) MPGAVPLTSSQALNNATLPFGLALANKGFSAVLENPHLRAGLNVHRGRLTYKAVAESLGL R. leg. AldA (301) MPGAVPVTSAHALNNATLVHGLALADRGLRAIAEDRHLRNGLNVHKGRITSKPVAEALGY M. l. c AldA (301) MPGAVPVTSAHALNNATLHYGLQLADKGLKALVDDHHLRNGLNVHKGKITNRAVAEALGY M. l. p AldA (301) MPGAVPVTSAHALNNATLHYGLQLADKGLKALVDDHHLRNGLNVHKGKITNRAVAEALGY S. mel. AldA (301) MPGAVPITSAHALNNATLQYGLQLADRGLKAIAEDRHLRAGLNVHRGRVTNAAVAEALGY Consensus (301) MPGAVPVTSAHALNNATL YGLQLADKGLKAIVED HLRNGLNVHKGRITNKAVAEALGY 361 372 B. jap. AldA (361) PFSPIEQAAA-- R. leg. AldA (361) EAFAPESVLNVA M. l. c AldA (361) ELVEPKAVLAA- M. l. p AldA (361) ELVEPKAVLAA- S. mel. AldA (361) DAHAPEAVLHVA Consensus (361) E PEAVLA

Fig 5.3 Amino acid alignment of L-alanine dehydrogenase from B. japonicum (B. jap.), R. leguminosarum bv. viciae (R. leg.), M. loti chromosome (M. l. c), M. loti pLMa (M. l. p) and S. meliloti (S. mel.). The amino acid identity is colour coded. black - identical, green - weakly similar, blue - conservative, grey - non homologous, pink - block of similar.

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5.3 Construction of a mutant in the putative regulator AldR

In order to confirm AldR was involved in the regulation of aldA, a mutant in aldR

was constructed to assess L-alanine dehydrogenase activity. A 3.1 Kb SalI

fragment containing aldR was cloned from cosmid pRI3135 into pSK- (to produce

plasmid pRU693). The DNA sequence of aldR was analysed for restriction sites

that could be used to insert the omega spcr cassette. The cassette is flanked by

BamHI, EcoRI, HindIII, SmaI and XmaI sites which can be used to clone the

DNA. AldR was found to have a single EcoRI site 250 bp from the putative start

codon (see Fig 5.2). A partial EcoRI digest of pRU693 produced a linear band of

6 Kb which was excised from an agarose gel and purified. The cassette was

excised from the vector pPH45Ω as an EcoRI fragment and cloned into the EcoRI

site within aldR to give pRU734 (Fig 5.4). The disrupted gene was subcloned into

the suicide vector pJQ200SK (genr) (Quandt & Hynes, 1993) as a SalI fragment

(plasmid pRU735) and this construct was conjugated into R. leguminosarum

strain 3841. Double recombinants were selected for by growth on 10% sucrose

(as for construction of the PHB mutant in chapter 3). L-alanine dehydrogenase

activity was assessed in five strr, spcr, gens isolates grown on alanine/glucose. In

each case there was no detectable AldA activity confirming that AldR was

involved in the regulation of aldA (Table 5.1). One isolate was designated strain

number RU1422 and was characterised by Southern blot and hybridisation using

a 2.5 Kb EcoRV fragment carrying the entire sequence of aldR from pRU693 as a

probe. This confirmed a 2 Kb increase in size of the hybridising SalI fragment due

to the insertion of the 2 Kb spcr cassette in aldR (Fig 5.5).

213

Fig 5.4 Vectors used in the construction of RU1422. Not to scale.

aldR construction.doc

214

Fig 5.5 Southern hybridisation to confirm AldR mutant indicated by increase of 2 Kb in aldR gene size. lane 1 - 1 Kb ladder; lane 2 - pRU693 SalI digest; lane 3 - RU1422 SalI digest; lane 4 - 3841 SalI digest; lane 5 - RU1422 EcoRV digest; lane 6 - 3841 EcoRV digest.

1 2 3 4 5 6

5.1kb

3.1kb

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5.4 Growth of mutant in AldR

RU1422 was further characterised by growth on plates to compare the phenotype

with RU1327 and the wild type 3841. As the aldA mutant, RU1327, did not

prevent growth on alanine it was expected the mutation in the putative regulator

gene would also allow growth on alanine. Indeed, RU1422 grew as the wild type

and RU1327 on alanine as sole carbon source (Table 5.2).

Table 5.2 Growth of 3841 and RU1422 on different carbon sources on agar plates. +++ good growth, ++ poor growth.

Carbon source 3841 RU1327 RU1422

Glucose +++ +++ +++

Succinate +++ +++ +++

Alanine ++ ++ ++

Pyruvate ++ ++ ++

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5.5 Transcriptional analysis of aldA and aldR

To confirm that AldR was a transcriptional regulator we constructed GUS fusions

to aldA and aldR in plasmid pJP2 and assessed GUS activity in the aldR mutant

background.

The intergenic region of aldA / aldR was amplified by PCR using primers P219

and P220 to give a 0.7 bp fragment which was cloned into pCR®2.1-TOPO

(pRU679). To construct a GUS fusion to aldR the region was subcloned into

pJP2 as a HindIII / XbaI fragment (pRU730 see Fig 5.6). To construct a GUS

fusion to aldA the region was first subcloned into pSK- as a EcoRI fragment

(pRU701) in order to switch the orientation of the aldR / aldA promoter regions

with respect to the HindIII and XbaI cloning sites in the vector. The region was

further subcloned from pRU701 into pJP2 as a HindIII / XbaI fragment to

construct the aldA fusion (pRU731 see Fig 5.6).

Plasmids pRU730 and pRU731 were conjugated into the 3841 and the AldR

mutant. There was no GUS activity from either fusion in the AldR mutant

background when cells were grown on glucose, succinate, 20mM alanine or

glucose / alanine whereas there was considerable activity in the wild type

backgound (see below). This indicated that in the absence of the AldR protein

there was no transcription of aldA confirming AldR was the transcriptional

regulator of aldA. Furthermore, the complete loss of GUS activity from the aldR

fusion in the aldR mutant background indicated that AldR was autoregulated.

217

Fig 5.6 Plasmids pRU731 (aldA GUS fusion) and pRU730 (aldR GUS fusion). Not to scale.

ald GUS fusions1.doc

Pictures of fusions from vector nti

218

L-alanine dehydrogenase activity was induced by growth on alanine and

carboxylic acids (see section 5.1). Therefore, GUS activity was assessed in

RU1414 (aldR-GUS fusion in 3841) and RU1415 (aldA-GUS fusion in 3841) to

monitor the levels of transcription in cells grown on these carbon sources.

There was a high level of GUS activity when grown on 20mM alanine or glucose /

alanine, but this level showed a high degree of variability over at least 3

independent assays carried out on different days. Within each independent assay

there was little variability (each done in x6). There was a moderate induction

when cells were grown on succinate. This level of induction was also inconsistent

over at least 3 assays (each done in x6) (Table 5.3). This moderate induction was

also seen on pyruvate and malate in one independent trial (in x6 data not shown).

There was a background level of GUS activity from both fusions when the cells

were grown on glucose (Table 5.3).

219

Table 5.3 GUS activity of RU1414, RU1415 and RU1416 grown on various C/N sourcesmeans for total assays carried out, standard error of the mean, total number assays carrrange in activity for these assays.

GUS activity (nmol min-1 mg protein-1

strain description Glucose/NH4 Alanine/Glucose 20mM Alanine

RU1414 AldR GUS fusion in 3841

163.40

20.28, n=15

555.47

145.00, n=16

range=171.50-1972.12

1143.77

244.28, n=18

range=218.38-2939.50

RU1415 AldA GUS fusion in 3841

104.41

9.09, n=15

349.12

100.05, n=17

range=121.01-1682.19

757.36

152.00, n=18

range=93.20-1872.22

RU1416 pJP2 in 3841 nd nd nd

nd – GUS activity not detected

219

220

It was hypothesised that the great variability in GUS activity on carbon sources

expected to induce L-alanine dehydrogenase activity and hence switch the fusion

on arose from a block in the regulatory pathway for AldA. The presence of the

intergenic region in multicopy on plasmid pJP2 would lead to the regulator being

sequestered by the plasmid. But the absence of a full length copy of regulator

gene aldR on the plasmid would prevent synthesis of AldR. This block in the

regulatory pathway would cause the differing levels of induction seen between

the independent assays. The level of induction would depend on the relative level

of AldR binding to the intergenic region of aldA and aldR on the chromosome

versus the plasmid.

It was necessary to assay actual enzyme activity to determine whether strains

RU1414 and RU1415 had a reduced level of L-alanine dehydrogenase activity

compared to 3841 due to the presence of the intergenic region in multicopy. L-

Alanine dehydrogenase activity was measured in RU1414 and RU1415 at the

same time as GUS activity was assayed in the strains grown on glucose,

succinate and alanine/glucose. L-Alanine dehydrogenase activity was measured

in 3841 grown on the same broth to give the control level of activity (Table 5.4).

221

Table 5.4 GUS activity vs. L-alanine dehydrogenase activity in cultures grown on glucose, succinate, and alanine/glucose.

Glucose/NH4 Succinate/NH4 Alanine/Glucose

GUS activity (nmol min-1 mg protein-1)

RU1414 320.495 283.503 1664.965

RU1415 202.935 198.610 888.286

L-Alanine dehydrogenase activity (µmoles min-1 mg protein-1)

RU1414 nd nd 0.219

RU1415 nd nd 0.232

3841 (control) nd 0.052 0.148

nd – not detected

There was no significant difference in the level of GUS activity in the strains

grown on glucose compared to succinate. At the same time there was no

detectable L-alanine dehydrogenase activity from the strains grown on glucose or

succinate whilst the level of activity in 3841 grown on succinate (0.052 µmoles

min-1 mg protein-1) was comparable to that seen in previous experiments (see

Table 5.1). Conversely, in the case of alanine / glucose, there was both high GUS

activity from strains RU1414 and RU1415 and induction of enzyme activity. This

confirms the presence of the intergenic region on plasmid pJP2 prevents proper

induction of the enzyme.

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5.6 Analysis of aldA mRNA levels

As we were unable to confirm the level of aldA transcription using the GUS

fusions constructed in pJP2 we decided to analyse aldA mRNA levels from 3841

grown on a range of carbon sources. Total RNA was extracted from cultures of

3841 grown in broth containing glucose, succinate, glucose / alanine and 20mM

alanine. RNA was probed with a 1.46 Kb EcoRI fragment from plasmid pRU640,

which carried a full-length copy of aldA. The resultant blots showed the aldA

transcript was estimated to be between 1.1 and 1.2 Kb, consistent with the length

of the aldA gene (1133 bp).

Fig 5.7 Representative northern analysis of aldA mRNA levels from 3841 grown on alanine (A), alanine/glucose (A/G), succinate (S), and glucose (G). L – RNA ladder.

L A1 A2 A/G S G L

estimated size of transcript = 1.1 to1.2Kb

223

DNA hybridized to mRNA from cultures grown on 20mM alanine with the greatest

intensity, confirming previous biochemical data that 20mM alanine results in the

highest level of enzyme activity. RNA analysis confirmed that aldA is transcribed

when cells were on glucose. With respect to the mRNA level when grown on

glucose, there was 2.1 fold induction of aldA mRNA levels on succinate; a 2.0

fold induction on alanine / glucose; and a 3.8 fold induction on 20mM alanine (Fig

5.7). These results were consistent over 3 independent trials of RNA extraction,

blotting and hybridisation. The pattern of induction was consistent with previous

biochemical data but the levels of induction were not. This may be due to the

difficulty in obtaining quantitative data from northern analysis.

5.7

224

Discussion

In chapter 4 we had shown that L-alanine dehydrogenase was responsible for the

synthesis of alanine in bacteroids. The main aim of this chapter was to look at the

regulation of L-alanine dehydrogenase.

Analysis of the Ald DNA sequence revealed a putative regulator, designated

aldR, transcribed divergent to aldA. Construction of a mutant in this gene

abolished L-alanine dehydrogenase activity confirming its role in the regulation of

aldA. The complete loss of activity from the aldA-GUS fusion in the aldR mutant

background confirmed this regulation was transcriptional. Furthermore, loss of

activity aldR-GUS fusion in the aldR mutant background indicated that expression

of aldR is regulated by the aldR gene product. Such autoregulation is the case for

many transcriptional regulators, for example BkdR of P. putida (Madhusudhan et

al., 1995) and Lrp of E.coli (Wang et al., 1994).

Enzyme data showed that L-alanine dehydrogenase was induced by growth on

alanine and dicarboxylates. This data was confirmed by the GUS fusion data and

by Northern analysis of aldA mRNA levels. L-Alanine dehydrogenase is most

strongly induced by growth of cells on alanine. We are uncertain why this should

be the case given its biosynthetic function and even though aldA is upregulated

by growth on alanine this level of enzyme does not allow growth on alanine. Only

when AldA is present in the cell in multiple copies (from cosmid pRU3135) does it

provide a catabolic function. This suggests that AldA may provide a biosynthetic

role, which may be regulatory, even in free-living cells.

225

The level of AldA activity in the bacteroid (0.09 µmoles min-1 mg protein-1) was

similar to that when cells were grown on dicarboxylates (0.07 µmoles min-1 mg

protein-1) suggesting transcription of aldA might be regulated in response to

carbon source in planta. This level of L-alanine dehydrogenase activity in

bacteroids is lower than that previously reported. Smith and Emerich (1993)

reported a much higher rate of AldA activity (1.0µmol min-1 mg protein-1) in

bacteroids of B. japonicum. This high enzyme activity might be a contributing

factor to the high level of alanine excreted from bacteroids of this strain of B.

japonicum (Waters et al., 1998). The reason for this difference in activity is not

known but perhaps significant is the fact that in some species of Rhizobium the

gene encoding L-alanine dehydrogenase is duplicated. There are two L-alanine

dehydrogenase genes in M. loti, one on the chromosome and the other on a

plasmid (pMLa) (Kaneko et al., 2000). The chromosomal copy is associated with

a regulator as in R. leguminosarum but the plasmid copy is not associated with a

regulator. This contrasts with R. leguminosarum where no residual activity of L-

alanine dehydrogenase could be detected in either strain RU1327 (aldA-) or

RU1422 (aldR-) strongly suggesting a single copy of the gene encoding this

dehydrogenase. A single copy of aldA and aldR is also found in S. meliloti

(http://sequence.toulouse.inra.fr/meliloti.html, unpublished data, 2001). This

raises the possibility that the slow and medium (meso) growing rhizobia have two

copies of aldA resulting in both higher AldA activity and nitrogen partitioning to

alanine in the nodule. So far one copy of aldA has been sequenced in the

symbiosis specific region of B. japonicum (Gottfert et al., 2001), but it is not yet

apparent whether other copies exist. This copy, like that in pMLa of M. loti, does

not have an associated aldR. In view of this data we aim to overexpress L-alanine

226

dehydrogenase to determine whether higher amounts of alanine are excreted

from bacteroids overexpressing aldA. In this chapter we showed that the aldA

mutant could be complemented by a full length copy of aldA (pRU708). L-alanine

dehydrogenase activity was restored in this strain (RU1371) but the level of

activity was not as high as wild type levels. Transcriptional work suggested that

this occurred because the levels of AldR were limiting the induction of the

enzyme (see below). Therefore it was decided that plasmid pRU708 would not be

used for overexpression studies and that a full length copy of aldA and aldR in

pTR101 should be constructed for these studies. This work is currently being

done in the laboratory.

To monitor the induction of aldA and aldR transcription, gene fusions were made

to the GUS reporter gene. We used pJP2, which is derived from pTR101,

because this plasmid is stable in the nodule and we hoped to examine the

induction of aldA during the course of nodule development. However we found

that the strains constructed gave inconsistent levels of induction due to a block in

the regulatory pathway. Because AldR is autoregulatory, an excess of AldR

binding sites that are not linked to a full length copy of the aldR gene will titrate

out the regulator. In RU1414 and RU1415 the presence of multiple intergenic

regions present on plasmids pRU730 and pRU731 would lead to an unbalanced

ratio of AldR binding sites to regulator and hence block induction of transcription.

Likewise, in RU1371 the presence of multiple binding sites not linked to aldR on

pRU708 would lead to a block in induction of L-alanine dehydrogenase activity.

From this data it is apparent that the aldA and aldR GUS fusions constructed

would not be useful in determining the induction of L-alanine dehydrogenase in

227

the nodule and an alternative strategy should be adopted. This new strategy

involves the construction of single copies of the aldA-GUS and aldR-GUS fusions

integrated into the chromosome of 3841 to overcome complications in the

malfunctioning of regulation experienced here. This work is currently being done

in the laboratory.

228

CHAPTER 6

229

6 GENERAL DISCUSSION

The fuelling of nitrogen reduction by bacteroids via the provision of dicarboxylic

acids is now widely accepted for legume nodules. However, our knowledge of

bacteroid metabolism is still limited. The aim of this thesis was to study pathways

that might regulate carbon metabolism within the bacteroid and subsequently

affect the efficiency of nitrogen fixation such as previously reported for PHB and

glycogen metabolism in bean (Cevallos et al., 1996; Marroqui et al., 2001). It has

been proposed that, as the bacteroid is subject to low oxygen concentrations,

normal aerobic metabolic pathways are modified to maintain efficient ATP and

reductant synthesis for nitrogen fixation to occur. The synthesis both of carbon

storage compounds and of amino acids has been suggested to be involved in this

regulation of carbon metabolism. The basic premise being that carbon and

reductant are drawn away from the TCA cycle by these pathways thus alleviating

TCA cycle enzyme inhibition under microaerobic conditions. Thus these

pathways operate as overflow metabolism for a TCA cycle that cannot operate to

its full aerobic capacity. However these pathways may not operate universally

throughout all Rhizobium – legume symbioses. For example, bacteroids in

indeterminate nodules do not synthesise large pools of PHB as seen in

bacteroids of determinate nodules. Moreover it has been argued that these

pathways might operate at the expense of nitrogen fixation and preventing PHB

synthesis in the R. etli – bean symbiosis results in an increase in the efficiency of

nitrogen fixation (Cevallos et al., 1996). Therefore we have studied PHB

metabolism in an in a determinate (bean) versus indeterminate (pea) symbiosis

230

using near isogenic strains of R. leguminosarum that differ only in symbiotic

plasmid.

The reason for the difference in accumulation of PHB in bacteroids of

determinate and indeterminate nodules is not known but recent work has

suggested different rates of malate consumption (MDH activity) by the TCA cycle

(Kim & Copeland, 1996; Kim & Copeland, 1997). Unfortunately we did not carry

out a full comparative study of enzyme activity between bacteroids of pea and

bean. But these factors would be important as differing enzyme rates will

determine the relative flux of carbon through different pathways and might reflect

different modes of regulating carbon metabolism in these different bacteroids.

Why these activities should differ between indeterminate and determinate

symbioses has not directly been questioned. In general slow growing rhizobia

develop determinate nodules whereas fast (and medium???) growing rhizobia

develop indeterminate nodules. Therefore one might first hypothesise that innate

difference in the rate of carbon metabolism by the bacterial partner leads to the

difference in accumulation of PHB in the bacteroid. However, this simple idea is

complicated when one considers the case of Rhizobium leguminosarum that is

able to form indeterminate nodules on pea and determinate nodules on bean,

depending on the Sym plasmid present. R. leguminosarum bv. viciae and bv.

phaseoli (example strains A34 and A5 respectively) have the same growth rate in

laboratory culture yet only bv. phaseoli accumulates PHB in bacteroids.

Therefore the differences in carbon metabolism (dominant dicarboxylate

delivered to the bacteroid / rate of carbon supply) by the pea and bean host

plants might determine the relative flux of carbon through different pathways in

the microsymbiont (hence accumulation of PHB). Consideration of plant carbon

231

metabolism and its impact on bacteroid carbon metabolism (efficiency of nitrogen

fixation) is beyond the scope of this study. However we have hypothesised in

chapter 3 that growth conditions might have a significant effect on the fixation

efficiency of a PHB synthase mutant in the pea symbiosis. The two greenhouse

experiments carried out with this mutant only differed in time of planting (May

versus July) which would have affected the rate of photosynthesis of the plant

(Wheeler, pers. comm.) and hence the carbon supply to nodules.

It has often been assumed that PHB metabolism is not important in indeterminate

(pea) nodules as bacteroids do not accumulate pools of PHB. However, it might

be that carbon is cycled through PHB continuously (Bergersen & Turner, 1992;

Encarnacion et al., 1995) such that no pool accumulates and that the amount of

polymer present at any one time is negligible. While it is possible to assay for

pools of metabolites and the presence of enzymatic steps we do not know the

actual metabolite fluxes within the bacteroid. Even when a large carbon flux is

obvious, such as is the case with PHB in determinate nodules we cannot say that

this storage pool is important or not. This is exemplified here by the results for the

PHB synthase mutant on bean plants and glycogen synthase mutant on pea

plants (chapter 3). In neither study was there a significant difference in

nitrogenase activity (acetylene reduction) or dry weight of plants inoculated with

the respective mutants. However, simple knockout mutations that block one

pathway may not give a true indication of importance as other pathways might

compensate for the defect. In this manner the flux of carbon in bacteroids might

be considered plastic and several pathways may be involved in regulation such

as PHB, glycogen and alanine synthesis. This raises the possibility that bacteroid

carbon pools are inter-regulated in which case it would be interesting to study the

232

effect of a double PHB and glycogen synthase mutant (as stated in chapter 3)

and alanine excretion from a PHB mutant, a glycogen mutant, or the double

mutant for instance. Unfortunately we were unable to study these factors during

the time of this project. This also raises the possibility that mutations that have an

effect in plants grown optimally in growth rooms or greenhouses may not relate to

plants that are environmentally stressed in a field situation. Indeed there are

numerous reports of rhizobial inoculants selected for enhanced nitrogen fixation

in controlled experiments that do not respond in the same way when transferred

to a field trial (REFS?????).

Until now the possible role of PHB in bacteroid development has not been

suggested. Previous ultrastructural studies have noted the presence of PHB in

the indeterminate infection thread and its absence in bacteroids (Paau & Cowles,

1978; Hirsch et al., 1983; Vasse et al., 1990). Also it is known that bacA mutant

cells, that abort bacteroid development, contain considerable reserves of PHB

(Glazebrook et al., 1993). The evidence for the role of PHB metabolism in

bacteroid differentiation we have presented (chapter 3) remains circumstantial

however it is worth noting that this hypothesis is also being studied in the S.

meliloti – alfalfa symbiosis (Charles, pers. comm.).

Early work showed that ammonium was the only nitrogen secretion product of N2

fixing bacteroids [Bergersen, 1967 #7553; Kennedy, 1966 #5272; Bergersen,

1990 #1256]. This was challenged by Waters et al. (1998) who claimed that

alanine, not ammonium, was the major nitrogenous compound excreted from

isolated soybean bacteroids. We have shown that isolated pea bacteroids

excrete both ammonium and alanine and that formation of alanine depends on

233

the relative concentration of ammonium. Activity of L-alanine dehydrogenase,

which is responsible for synthesis of alanine, is not dependent on nitrogenase

activity (chapter 4) but the enzyme is induced by growth on dicarboxylates to

levels seen in pea bacteroids (chapter 5) suggesting transcription of aldA might

be regulated in response to carbon source in planta. We have made some

ground on the relevance of L-alanine dehydrogenase during symbiosis by

showing that peas inoculated with a mutant in L-alanine dehydrogenase fix

nitrogen but that 6 week old plants show a significant (20%) reduction in dry

weight (chapter 4). This shows that alanine cannot be the sole excretion product

as suggested by Waters et al. (1998) but the data is consistent with alanine

excretion contributing to the net transfer of nitrogen to the plant during symbiosis.

The data is also consistent with a role for alanine synthesis in regulating carbon

metabolism within the bacteroid. It is possible that the decrease in symbiosis

efficiency we have reported in the L-alanine dehydrogenase mutant is the result

of a decrease in the efficiency of dicarboxylate metabolism, hence delivery of

energy and reductant to nitrogenase, in the bacteroid. This role need not be

independent of a role for nitrogen assimilation during symbiosis. Likewise the

decrease in plant biomass of 40% in the PHB synthase mutant strain (chapter 3)

might be due to a decrease in efficiency of carbon metabolism by the bacteroid.

Both alanine and PHB synthesis consume carbon and reductant therefore can

feasibly contribute to regulation of TCA cycle function in bacteroids.

If alanine excretion does contribute to the net transfer of nitrogen to the plant then

further questions to ask would be (1) how is alanine transported across the

symbiotic membranes? and (2) what happens to the alanine in the plant cytosol?.

Transport systems for alanine have not been defined on the bacteroid or

234

peribacteroid membranes. However this probably reflects a direction of research

that has not previously questioned the need to have such transport systems and

that has therefore not looked for them. Recent sequencing projects of the S.

meliloti genome shows that there are xxxx uncharacterised ABC transport

systems in S. meliloti (http://sequence.toulouse.inra.fr/meliloti.html, unpublished

data, 2001). Therefore there it is clear that a candidate for alanine export from the

bacteroid might lie undiscovered.

Waters et al. (1998) suggested that alanine was a transport species and would

be de-aminated in the cytosol, then ammonium would be re-assimilated by GS-

GOGAT [Waters, 1998 #8115]. The fact that in this study by Waters et al.

ammonium was only detected when plant cytosol was added to the incubation

mixture of bacteroids strongly suggests that alanine is de-aminated in the plant

cytosol. To study whether alanine serves to transport fixed nitrogen across

symbiotic membranes, or whether it persists in the plant cytosol, the composition

of xylem sap could be studied. Plants incubated in the presence of 15N2 have

been shown to have label incorporated into asparagine or ureides (depending on

the symbiosis type) in the xylem sap (REFS?). If alanine persists after being

exported from the bacteroid and contributes to the total fixed nitrogen transported

out of the nodule we would expect to see labelled alanine in xylem sap. However,

if it is de-aminated upon excretion from the symbiosome, and the ammonium thus

formed rapidly re-assimilated by GS-GOGAT, then alanine would not be detected

in the xylem sap. In this instance experiments using methionine sulphoximine

(MSX) as an inhibitor of GS might be useful in tracking the fate of excreted

alanine. Also, by comparing the composition of labelled nitrogen within xylem sap

from nodules infected with wild type and L-alanine dehydrogenase mutant we

235

would also be able to address whether bacteroid alanine excretion significantly

alters nitrogen metabolism in the plant.

Immediate threads for future research in relation to this thesis are laid out within

the independent chapters. Clearly the number of questions that can be asked of

bacteroid metabolism in general are innumerable as it is apparent from the

genome sequencing projects of Sinorhizobium meliloti and Mesorhizobium loti

that there are large numbers of metabolic genes within the rhizobia, some of

which are linked to symbiotic regions. The role of the majority of these genes is

not yet known and therefore their analysis is likely to be a fruitful area of

investigation and will stimulate research for the foreseeable future.

Transcriptomic and proteomic investigations will hopefully provide expression

maps of when and where certain genes are switched on. This should provide an

enormous spur to this field but it will only provide us with a framework unless an

equivalent metabolomic profile can be built up. A great deal of the metabolic

diversity of the rhizobia will probably relate to survival, growth and colonization of

the plant rhizosphere as well as development in the infection thread. However,

there will undoubtedly be many surprises relating to metabolism of the mature

bacteroid.

236

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