Characterization of Transport Proteins on the Symbiosome Membrane of Glycine Max

Characterization of transport proteins on the symbiosome membrane of Glycine max

Siti Nurfadhlina Mohd Noor

A thesis submitted in fulfilment of the requirements for the degree of Doctor of Philosophy School of Environmental and Life Sciences The University of Sydney 2016 Statement of originality

This is to certify that to the best of my knowledge, the content of this thesis is my own work.

This thesis has not been submitted for any degree or other purposes.

I certify that the intellectual content of this thesis is the product of my own work and that all the assistance received in preparing this thesis and sources have been acknowledged.

Chapter 1 of this thesis contains some part of the book chapter published in “Mohd Noor, S.N.,

Day, D.A. and Smith, P.M. 2015. The Symbiosome Membrane. Biological Nitrogen Fixation.

John Wiley & Sons, Inc. p. 683-694”. The book chapter was written by me during PhD candidature, and was reviewed and edited by David Day and Penelope Smith.

Abstract

During the legume-rhizobia symbiosis, a new plant organ called the root nodule is induced and developed. Within the root nodule, rhizobia are enclosed in the symbiosome. The symbiosome is central to the interaction; it is an organelle where rhizobia fix atmospheric nitrogen and is surrounded by the plant-derived symbiosome membrane (SM). The SM regulates nutrient exchange between the symbionts, and thus effectively controls the symbiosis. It is therefore critical that we understand the properties of the SM and identify the transport proteins embedded in it.

GmABCA1 and GmABCA2 proteins have been identified by proteomic analysis on the SM of soybean. These proteins are part of the large ATP-binding cassette (ABC) superfamily that transports diverse substrates and plays essential roles in plant growth and development. ABC subfamily A proteins are of particular interest as many of them can transport lipids. The present work aims to characterize GmABCA1 and GmABCA2, and to explore potential function they may play in relation to fatty acid transport. Expression analyses showed that they are preferentially expressed in the nodule tissue, and have specific expression in infected cells. The localization of GmABCA2 on the SM was confirmed by FP tagging but that of GmABCA1 could not be determined by this method. Complementation of the yeast fatty acid uptake- deficient mutant, Δfat1, indicated that both proteins were able to transport oleic acid.

GmABCA1 and GmABCA2 also complemented the T-DNA mutant (Salk 058070) for the

Arabidopsis fatty acid transporter Atabca9 by restoring establishment of seedlings on half MS

ii without sucrose to varying degrees. The possible roles that GmABCA1 and GmABCA2 play as components of the SM is discussed.

The nitrate transporter 1/peptide transporter family (NPF) belongs to the major facilitator superfamily (MFS), and is mainly involved in the transport of nitrates and peptides in plants.

Complementation of eight soybean NPF members with enhanced expression in nodule:

GmNPF1.2, GmNPF5.24, GmNPF5.25, GmNPF5.29, GmNPF5.30, GmNPF5.2, GmNPF5.3 and GmNPF8.6 in a yeast peptide transport mutant, ptr2, was attempted in this study. Only

GmNPF8.6 was able to restore ptr2 yeast growth on minimal media containing tri-alanine

(AAA), glycine-proline (GP) and valine-leucine (VL) peptides as the sole source of nitrogen.

The significance of peptide transport across the symbiosome in unclear. This study provides the first evidence of such transport across the SM.

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Acknowledgement

I would like to express my special appreciation to my supervisor Associate Professor Penelope

Smith for her boundless patience and encouragement, and for her time editing this thesis. Her guidance and ideas to try new methods has pushed me to gather better results. Without her advice and help this thesis would not have been possible.

I am also indebted to several people who assisted my PhD work. I would like to thank Dr Ella

Brear for her help with many aspects of my experiments. It was my pleasure to share this PhD journey with her. I want to thank Professor David Day for his time editing this thesis. I am appreciative to Asmini Athman at The University of Adelaide for help and guidance with in situ PCR. I am grateful to Dr Debbie Barton, who willingly shared her expertise in microscopy.

I would also like to thank Dr Aleksandr Gavrin, Dr Amanda Huen, Dr Victoria Clarke, Dr

Patrick Loughlin, Yihan Qu and Dr Evgenia Ovchinnikova for their help with various lab protocols. I would like to acknowledge the Australian Centre for Microscopy Analysis at The

University of Sydney for their facility and guidance in using the confocal microscope and vibratome.

A special thanks to my family, especially my parents, who have been supporting me during my academic pursuit. I am thankful for my husband, Aiman, who has been very understanding and supportive, especially during the time writing this dissertation. I would also like to express my appreciation to my parents-in-law for their understanding while writing this thesis.

Abbreviations

ABA Abscisic acid cDNA complementary DNA

ABC ATP-binding cassette CDS Coding sequence

ABRC Arabidopsis Biological Resource CLE Clavata/ embryo surrounding region-

Center related protein

ACMM Australian Centre for Microscopy CmR Chloramphenicol resistance and Microanalysis CoA Coenzyme A

Ala Alanine Col-0 Colombia-0

AmpR Ampicillin resistance gene Ct Threshold cycle

AON Autoregulation of nodulation CTS Comatose

ARP Actin-related protein Dct Dicarboxylate transport

AS Air space DGDG Digalactosyldiacylglycerol

Asp Aspartic acid DIG Digoxigenin

B&D Burton and Dilworth DKP Diketopiperazine

BIC Bayesian Information Criterion DMA Deoxymugineic acid

BICINE N,N-Bis(2-hydroxyethyl)glycine DNA Deoxyribonucleic acid

BiFC Bimolecular fluorescence dNTP Deoxynucleotide triphosphate complementation DPI Days post inoculation

BlpR Basta/ phosphinothricin/ glufosinate DsRED Discosoma sp. red fluorescent resistance gene protein

BM Bacteriod membrane DVL1/ROT4 Devil/rotundifolia-Four-

CEP C-terminally encoded peptide Like

EDTA ethylenediaminetetraacetic acid IAA Indole acetic acid

EGFP Enhanced green fluorescent protein IBA Indole-3-butyric acid

ENOD Early nodulin ICs Infected cells

ER Endoplasmic reticulum IC Inner cortex or infected cell

Euroscarf European Saccharomyces ICL Intra-cytosolic loop

Cerevisiae Archive for Functional Analysis IRT1 Iron-regulated transporter 1

EV Empty vector JA Jasmonic acid

F Flower JA-Ile Jasmonoyl-isoleucine

FAs Fatty acids L Leaf

FAS Fatty acid synthase LB Left border

FP Fluorescent protein LbC3 Leghemoglobin C3

GA Gibberellin LCMS/MS Liquid chromatography

GC/MS Gas chromatography mass tandem mass spectrometry spectrometry Leu Leucine gDNA Genomic DNA LORE1 Lotus Retrotransposon 1

Gln Glutamine LP Left primer

Glu Glutamic acid LTPs Lipid transfers proteins

Gly Glycine Lys Lysine

GSA Glutamine synthetase activity MA Mugineic acid

GSH Glutathione MC Middle cortex

GUS β-glucuronidase gene MDR Multidrug resistance protein

HA hemaggluinin Met Methionine

His Histidine MFS Major facilitator superfamily

ML Maximum likelihood Ori Origin of replication

MRP Multidrug resistance-associated P Pod protein PACA Phosphotungstic acid-chromic acid

MS Murashige and Skoog PCR Polymerase chain reaction

N Nodule PDR Pleiotropic drug resistance

NA Nicotianamine PEG Polyethylene glycol

NBD Nucleotide-binding domain Peff PCR efficiency

NCBI National Center for Biotechnology PEP3 Peroxisome defective 3

Information PHB Ply-β-hydroxybutyrate

NCR Nodule-specific cysteine-rich Phe Phenylalanine

NEB New England Biolabs PM Plasma membrane

NIP/LATD Numerous Infections and POT Proton-coupled oligopeptide

Polyphenolics/Lateral root-organ Defective transporter

Nod Nodulation Pro Proline

NOS Nopaline synthase PS Phytosiderophore

NPF NRT1/PTR family PTR Peptide transporter

NRAMP Natural Resistance-Associated PXA1 Peroxisomal ABC transporter 1

Macrophage Protein QS Quorum sensing

NRT1 Nitrate transporter 1 R Root

NTR Nitrate transporter RALF Rapid alkalization factor

OC Outer cortex RB Right border

OD Optical density Ri Root-inducing

OPT Oligopeptide transporter RNA Ribonucleic acid

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RNAi RNA interference TE Transposable element

ROI Region of interest TGD Trigalactosyldiacyl glycerol

RP Right primer TGN Trans-golgi network

RT-qPCR Real time quantitative PCR TMD Transmembrane domain

SA Salicylic acid TMHs Transmembrane helices

SD Synthetic defined TRIS Tris(hydroxymethyl)aminomethane

SDS Sodium dodecyl sulfate UBI Ubiquitin

SLC15 Solute carrier 15 UN Uninfected cell/s

SM Symbiosome membrane UN Untransformed

SMP Symbiosome membrane peripheral URA3 Uracil biosynthesis

Sm/SpR Spectinomycin and streptomycin URGT UDP–L-Rha/UDP–D-galactose resistance genes transporter

SNARE SNAP receptor VAMPs Vesicle-associated membrane

SNF Symbiotic nitrogen fixation proteins

SP Signal peptide VB Vascular bundles

SS Symbiosome space WBC White brown complex

SylA Syringolin A WT Wild type

T35S Terminator of 35S X-Gluc 5-bromo-4-chloro-3-indolyl-β-D-

TAE TRIS acetic acid EDTA glucuronide cyclohexylammonium salt

TAG Triacylglycerol YEM Yeast extract mannitol

TAIR The Arabidopsis Information YNB Yeast nitrogen base

Resource YPD Yeast-peptone-dextrose

T-DNA Transfer-DNA YSL Yellow-stripe like

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Table of contents

Statement of originality………………………………………….….…………………i

Abstract……………………………………………………….….……………………ii

Acknowledgements……………………………………………………….….……….iv

Abbreviations………………………………………………………….….…………...v

1 General introduction…………………………………………………….….1

1.1 Symbiotic nitrogen fixation……………………………………….….…….….….1

1.1.1 Root nodule organogenesis……………………………………….……….….…2

1.1.2 Symbiosome formation and development……………………………...…….….2

1.1.2.1 Symbiosome initiation and formation…………………………………....….3

1.1.2.2 Symbiosome proliferation, maturation and senescence.……………...…….6

1.1.3 Transport activity of the symbiosome membrane………………………...….….9

1.2 ABC transporters……………………………………………….………….…….14

1.2.1 Domain organization and structure………………………………….….….….15

1.2.1.1 Nucleotide binding domains (NBDs)…………………………..…………16

1.2.1.2 Transmembrane domains (TMDs)……………………………….….…….17

1.2.2 Plant ABC transporters…………………………………………………..…....18

1.2.2.1 Subfamily A…………………………………………………….…….…...19

1.2.2.2 Subfamily B…………………………………………………………..…...21

1.2.2.3 Subfamily C……………………………………………………….……....21

1.2.2.4 Subfamily D…………………….…………………………………….…...22

1.2.2.5 Subfamily G……………………………….……………………………....22

1.2.2.6 Subfamily I………………………………………..………………………23

1.3 NPF Transporters…………………………………………………………..……23

1.3.1 NPF subfamilies…………………………………………………….……….…24

1.3.1.1 Subfamily 1……………………………………………………….….……24

1.3.1.2 Subfamily 2……………………………………………………….….……25

1.3.1.3 Subfamily 3……………………………………………………….….……25

1.3.1.4 Subfamily 4…………………………………………………………..……25

1.3.1.5 Subfamily 5………………………………………………………….….…26

1.3.1.6 Subfamily 6…………………………………………………………….….26

1.3.1.7 Subfamily 7………………………………………………………….……..27

1.3.1.8 Subfamily 8…………………………………………………………….….27

1.4 Oligopeptide transporters………………………………………………….……..28

1.4.1 Yellow stripe-like transporters…………………………………………….…..28

1.4.2 True OPTs………………………………………………………………….….30

1.5 Objectives of the present study…………………………………………...... 31

2 General materials and methods…………………………………….….…33

2.1 Growing wild type soybean plants………………………………………….…...33

2.2 Isolation of nucleic acids from plant tissue……………………………………….33

2.2.1 Genomic DNA extraction…………………………………………..…………34

2.2.2 Total RNA extraction and cDNA synthesis…………………………….….….34

2.3 Gene expression analysis………………………………………………..………35

2.3.1 Plant material, RNA isolation and cDNA synthesis…………………….…….35

2.3.2 Primer design and reference gene……….………………………….……..…..35

2.3.3 Validating primer specificity…………………………………………………..36

2.3.4 qRT-PCR conditions…………………………………………………………..36

2.3.5 Data analysis…………………………………………………………………..37

2.4 Gene cloning using Gateway® Technology………………..….…………………37

2.4.1 Vectors………………………………………….…………………….……….38

2.4.2 Designing attB PCR primers……………………………………………..…….38

2.4.3 Producing attB-PCR products………………………………………………....39

2.4.3.1 Agarose gel electrophoresis……………………………………………...... 39

2.4.4 Purifying PCR products………………………………………………………..40

2.4.4.1 PEG/ MgCl2 purification……………………………………………………..40

2.4.4.2 PCR clean-up………………………………………………………………...40

2.4.4.3 Gel extraction………………………………………………………………..40

2.4.5 Generating entry clones using the BP recombination reaction……………...….40

2.4.6 Generating expression clones using the LR recombination reaction…………...41

2.5 Preparation of E. coli competent cells……………………………….………..…41

2.6 Heat-shock transformation of chemically competent bacteria…………………...42

2.7 Screening for transformed E. coli cells…………………………………………...42

2.7.1 Colony PCR…………………………………………………………..……….42

2.7.2 Isolation of plasmid DNA………………………………………………..…….43

2.7.3 Restriction enzyme digest………………………………………………..…….43 xi

2.8 Preparation of Agrobacterium competent cells……………………………..……43

2.9 Agrobacterium transformation…………………………………………………...44

2.10 Screening for transformed Agrobacterium cells………………………………...44

2.10.1 Colony PCR……………………………………….…………………………44

2.10.2 Agrobacterium plasmid isolation and restriction digest…...………………….45

2.11 Gene transfer to plants cells via Agrobacterium rhizogenes-mediated

hairy root transformation……………………………………….………..…….45

2.11.1 Seed germination…………………………………………………….………45

2.11.2 Hairy roots induction……………………………………………………...….46

2.11.3 Removal of non-transgenic roots……………………………………………..46

2.11.4 Maintaining plants and screening for transgenic nodules……………………..47

2.12 Preparation of yeast competent cells…………….………………..……………47

2.13 Yeast transformation……………………………………………………...…….48

3 Characterization of GmABCA1 and GmABCA2 in soybean nodules…49

3.1 Introduction……………………………………………………………………...49

3.2 Materials and Methods…………………………………………………………..50

3.2.1 Protein alignment and phylogenetic analysis………………………………….50

3.2.2 Gene expression analysis………………………………………………………51

3.2.3 In planta localization…………………………………………………………..51

3.2.3.1 Amplification and cloning…………………………………………………51

3.2.3.2 Biolistic bombardment in Allium porrum L……………………………….52

3.2.3.3 Soybean transformation and growth………………………………………53

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3.2.3.4 Confocal microscopy………………………………………………………53

3.2.4 Bioinformatics analysis………………………………………………………..54

3.2.5 In situ PCR…………………………………………………………………….54

3.2.6 Heterologous complementation of Δfat1………………………………………55

3.2.7 Functional complementation of the Atabca9 mutant………………………….56

3.2.7.1 Plant material and growth conditions………………………………………56

3.2.7.2 Identification of homozygous abca9 mutant plants……………………….56

3.2.7.3 Vector construction…………………………………….………………….57

3.2.7.4 Transformation via the floral dip method………………………………….58

3.2.7.5 Maintaining plants and seeds harvest………………………………………59

3.2.7.6 Obtaining homozygous lines for the transgenes…………………………..59

3.2.7.7 Performing the complementation assay……………………………………60

3.3 Results…………………………………………………………………………...60

3.3.1 Isolation of GmABCA1 and GmABCA2 CDS…………………………………..60

3.3.1.1 GmABCA1…………………………………………………………………...61

3.3.1.2 GmABCA2…………………………………………………………………...62

3.3.2 Analysis of predicted protein structure ……………………….………………63

3.3.3 Phylogenetic analysis…………………………………………………………..64

3.3.4 Gene expression analysis………………………………………………………65

3.3.5 Spatial localization of expression………………………………………………66

3.3.6 Protein localization…………………………………………………………….67

3.3.6.1 GmABCA1………………………………………………………………..67

3.3.6.2 GmABCA2………………………………………………………………...68

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3.3.7 Yeast complementation…………………………………………………….….68

3.3.7.1 fet3fet4ftr complementation……………………………………………….68

3.3.7.2 ∆fat1 complementation…………………………………………………….69

3.3.8 Atabca9 complementation………………………………………………….….71

3.3.8.1 Identification of homozygous abca9 mutant plants…………………….…71

3.3.8.2 Complementation of abca9-1 mutant……………………………………...71

3.4 Discussion……………………………………………………………………….72

3.4.1 GmABCA1 and GmABCA2 have nodule enhanced expression…………………73

3.4.2 GmABCA1 and GmABCA2 are specifically expressed in the infected nodule

cells………...………………………………………………………………….75

3.4.3 Localization……………………………………………………………………75

3.4.3.1 GmABCA1 localization in G. max………………………………………..76

3.4.3.2 GmABCA1 localization in Allium porrum L………………………………77

3.4.3.2 GmABCA2 is localized on the SM………………………………………..78

3.4.4 Yeast complementation………………………………………………………..80

3.4.4.1 GmABCA1 and GmABCA2 could not transport iron into yeast………81

3.4.4.2 GmABCA1 and GmABCA2 could transport fatty acids into yeast……82

3.4.6 GmABCA1 and GmABCA2 complement Atabca9 mutant………………… ..84

3.4.7 Future characterization of GmABCA1 and GmABCA2………………………89

4 Investigation of putative peptide transporters in soybean nodules…….92

4.1 Introduction……………………………………………………………………...92

4.1.1 The roles of peptides…………………………………………………………..93

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4.1.2 Candidate proteins……………………………………………………………..95

4.2 Materials and methods…………………………………………………………..99

4.2.1 Phylogenetic analysis………………………………………………………….99

4.2.2 Heterologous complementation in yeast………………………………………100

4.2.2.1 Vector construction…………………………………………………..….100

4.2.2.2 Yeast growth, transformation and complementation……………………100

4.3 Results………………………………………………………………………….102

4.3.1 Phylogenetic analysis………………………………………………………...102

4.3.1.1 The NPF phylogeny………………………………………………………102

4.3.1.2 The OPT phylogeny………………………………………………………103

4.3.2 Complementation of yeast mutant ptr2………………………………………..103

4.3.3 Complementation of yeast mutant opt1……………………………………..104

4.4 Discussion………………………………………………………………………105

4.4.1 Only GmNPF8.6 is able to transport peptides………………………………..105

4.4.2 The other GmNPFs may not be able to transport peptides…………………..107

4.4.3 GmYSL7……………………………………………………………………..109

5 General discussion………………………………………………….…….113

5.1 Introduction…………………………………………………………………….113

5.2 Direction of transport…………………………………………………………..116

5.3 Alternative method for transport study….……………..………………………118

5.3.1 Reconstitution of transport proteins into liposomes…………………………118

5.3.2 Heterologous expression in Xenopus oocytes……………………………….119

5.3 Uncovering physiological roles through metabolic profiling………………….120

5.4 Stable mutants in other legumes………………………………………………..121

5.5 Conclusion………………………………………………………………...... 123

References…………………………………………………………………………..124

Appendices………………………………………………………………………….174

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Chapter 1

General Introduction

1.1 Symbiotic nitrogen fixation

Nitrogen is a key nutrient for plants that is most often the major limiting factor to crop productivity. The supply of nitrogen fertilizers to elevate nitrogen input for crop production has led to undesirable environmental impacts (Rockstrom et al., 2009). Symbiotic nitrogen fixation

(SNF) provides an alternative natural source of fixed nitrogen to plants, by converting

+ biologically inert atmospheric nitrogen, N2, to a usable reduced form of ammonium, NH4 . Soil bacteria collectively known as rhizobia are able to perform SNF when in a symbiotic association with members of the Fabaceae family. The association results in the formation of a new plant organ called the nodule that creates favorable conditions for the activity of the rhizobial nitrogenase enzyme, which catalyzes the reduction of nitrogen.

1.1.1 Root nodule organogenesis

Nodule organogenesis starts by a complex signal exchange between the symbiotic partners in the rhizosphere (Gibson et al., 2008). The host plant releases root exudates that attract rhizobia to the growing tip of the root hair via positive chemotaxis. The attachment induces the production of nodulation (Nod) factors by the bacteria, and these signal molecules are perceived in the root epidermis by compatible Nod factor receptors (Oldroyd and Downie, 2004). This triggers an early nodulation response that leads to a number of physiological and morphological changes such as root hair deformation, cytoskeleton reorganization and initiation of cell divisions in the root cortex (Oldroyd, 2013; Oldroyd and Downie, 2004).

Rhizobia subsequently enter legume roots via infection threads that are initiated when the rhizobia are entrapped by curling of the root hair (Brewin, 2004; Oldroyd, 2013). The local cell wall is hydrolyzed and degraded, while the plasma membrane (PM) invaginates leading to the formation of a tubular infection thread. Infection threads then grow onward into the root cortex, transporting bacteria into the nodule primordium cells that are formed from reprogrammed root cortical cells (Brewin, 2004; Rae et al., 1992). Ultimately, the bacteria are internalized into nodule primordium cells. The host membrane surrounds the enclosed bacteria and results in an organelle-like membrane compartment called the symbiosome (Roth et al., 1988; Figure 1.1), inside infected cells of the root nodule.

1.1.2 Symbiosome formation and development

The symbiosome is the fundamental cellular unit of symbiotic nitrogen fixation, in which the

Bacteroid

Symbiosome membrane UC IC

Symbiosome space AS Cytosol

Figure 1.1 Electron micrograph of symbiosomes within infected cell of a soybean nodule. In determinate soybean nodules, the symbiosomes housed several bacteroids. Abbreviations: IC – infected cell, UC – uninfected cell and AS – air space. Image is taken from Udvardi and Day (1997).

plant provides carbon and other essential nutrients to the bacteroid to support nitrogen fixation and bacteroid metabolism, and in return the bacteroid provides fixed nitrogen to the plant. The membrane around the symbiosome can be termed the symbiosome membrane (SM), and is where transport activities between the symbionts occur. The term SM is used here instead of the older term of peribacteroid membrane, which ignores the similarity of symbiotic structures in associations between a range of different organisms (Roth et al., 1988). The SM and the bacteroid membrane (BM) are physically separated by a symbiosome space (SS), and therefore their transport activities are distinct. Symbiosomes undergo four developmental phases, namely initiation, proliferation, maturity and degradation. The role and composition of the SM is considered to be different in the four phases (Brewin, 1991; Whitehead and Day, 1997).

1.1.2.1 Symbiosome initiation and formation

Symbiosome biogenesis begins with the formation of an infection droplet, a structure that represents an unwalled outgrowth from the infection thread. This is a region where the infection thread membrane invaginates and where rhizobia come into direct contact with the host PM without the cell wall as a barrier (Brewin, 2004; Rae et al., 1992). Bacteria are then released into the cytoplasm of cortical cells when they come into close contact with the host membrane, which results in encapsulation of bacteria by the plant cell membrane (Bassett et al., 1977;

Goodchild and Bergersen, 1966; Newcomb, 1976). Inside the symbiosome, bacteria differentiate into bacteroids, their symbiotic form, which enables nitrogen fixation to take place

(Vasse et al., 1990). In indeterminate nodules of temperate legumes (e.g. alfalfa, pea), each symbiosome typically houses a single large bacteroid, while in determinate nodules of tropical legumes (e.g. soybean), there are usually several bacteroids enclosed in a symbiosome

(Fedorova et al., 1999; Figure 1.1). Formation of symbiosomes involves a major reorganization of the host cytoskeleton and endomembrane systems (Davidson and Newcomb, 2001). To accommodate biogenesis of symbiosomes in the infected cells, massive amounts of protein and lipid are needed. Depending on the legume species, the total surface area of SM can exceed the

PM by approximately 30-fold (Verma et al., 1978). Although traditionally it is considered that the symbiosome is formed through endocytosis, more recent evidence suggests that the exocytotic pathway also plays a major role.

In endocytosis, internalized cargo molecules that are taken up by the PM are transported through the endocytic pathway (Šamaj et al., 2005). The endocytotic pathway involves a multifaceted network of membrane compartments with each compartment involved in different tasks. After internalization into the early endosome, cargo to be recycled is transported back to the PM whereas cargo for degradation is transported to the late endosome. The late endosome will finally fuse with the vacuole in plants or lysosomes in animals for degradation (Šamaj et al., 2005).

In Medicago truncatula, symbiosomes did not acquire the (late) endosomal marker Rab5 or early endosomal/trans-golgi network (TGN) marker, SYP4, at any stage during their development and only acquired Rab7, a late endosomal/ vacuolar marker, at a later stage of development when symbiosomes had stopped dividing (the fixation zone) (Limpens et al.,

2009). This marker was retained until senescence began. This suggests that the early formation of the symbiosome follows a Rab5-independent endocytic pathway and that Rab7 might be recruited directly from the cytoplasm to regulate symbiosome formation.

Additionally, symbiosomes did not acquire vacuolar SNAP receptor (SNARE), SYP22 and

VTI11 until the onset of senescence. However, the symbiosome membrane contains a PM t-

SNARE/ SYNTAXIN SYP132 throughout development (Catalano et al., 2007; Limpens et al.,

2009). The delay in acquiring vacuolar SNAREs might be required for maintenance and survival of the symbiosome as an individual nitrogen-fixing organelle. The presence of SYP132 and Rab7 showed that the SM has a unique composition where proteins that move from the secretory pathway to the PM and from the endocytic pathway to the vacuole are involved

(Limpens et al., 2009). However, there is no clear molecular evidence for involvement of the default endocytosis pathway in the early stages of symbiosome formation.

In contrast, evidence of the involvement of an exocytotic pathway is accumulating (Genre et al., 2012; Ivanov et al., 2012). Exocytosis involves fusion of transport vesicles with the PM.

This fusion is mediated by exocytic vesicle-associated membrane proteins (VAMPs) and in plants, members of the VAMP72 family. Silencing of two VAMP72 family members in M. truncatula prevented symbiosome formation, supporting the role of exocytosis in their formation. In the MtVAMP72 silenced nodules, infection droplets did not form properly as they were bound within a thin layer of cell wall. The presence of the cell wall prevents close contact between bacteria and host membrane, which prevents further release of bacteria (Ivanov et al.,

2012).

Recent evidence showed that the silencing of VAMP72d in Glycine max produced a similar phenotype as silencing of MtVAMP72, in which bacterial release and symbiosome formation are hampered (Gavrin et al., 2016). In addition, in GmVAMP721d-silenced nodules, the

infected cells were filled with clusters of bacteria embedded in a pectin matrix. GmVAMP721d- positive vesicles partially colocalized with pectate lyase GmNPL1. This suggests that

GmVAMP72 is involved in delivering the pectin-modifying enzymes to the rhizobia release site (Gavrin et al., 2016).

1.1.2.2 Symbiosome proliferation, maturation and senescence

The SM proliferates and differentiates to accommodate bacteroid growth and division until infected cells are filled with symbiosomes (Robertson and Lyttleton, 1984). Enlarged mature infected cells may house thousands of symbiosomes (Day et al., 2001). It is still unclear how the SM proliferates, but it is proposed that it may be the result of redirection of the host secretory pathway toward SM biogenesis (Leborgne-Castel et al., 2010). Proliferation of the SM is considered to occur independently of bacteroid division (Mellor and Werner, 1987). Within the developing infected cell, there is a specialized area adjacent to the nucleus that is devoid of symbiosomes where abundant membrane is observed associated with the endoplasmic reticulum (ER). This was suggested to serve as a membrane reservoir for the proliferating symbiosomes (Bulbul and Kaneko, 2009). Bundles of actin filaments that co-localized at the same site may function in delivering membranes to the developing symbiosomes (Bulbul and

Kaneko, 2009) and play a role in spatial organization of the symbiosomes, maintaining effective pathways for nutrient diffusion within the symbiosomes (Whitehead et al., 1998).

The SM consists of a membrane bilayer 9 to10 nm thick (Dart and Mercer, 1963; Mellor and

Werner, 1987) and is derived from the PM through the infection thread membrane. This is illustrated by staining by phosphotungstic acid-chromic acid (PACA) (Robertson et al., 1978;

Roth and Stacey, 1989; Verma et al., 1978), which is normally considered a PM specific stain.

A freeze-fracturing study showed that the particle profiles of SM surface in alfalfa, clover and soybean are highly similar to the PM surface (Tu, 1975). The syntaxin SYP132, as well as being present in the SM, labels the PM around the infection threads and infection droplets (Catalano et al., 2007), also linking the two membranes.

Notwithstanding the origin of the symbiosome, bacteroid division and SM proliferation results in a SM with distinct properties. Both the protein and lipid complement of the SM are unique and are supplied from a range of sources, including the ER and de novo membrane synthesis

(Roth and Stacey, 1989). Digalactosyldiacylglycerol (DGDG) lipids, which are normally associated with plastid membranes rather than PM, are present in the SM (Gaude et al., 2004), indicating its unique identity. The mature SM also has an exceptionally high lipid to protein ratio (6:1) (Robertson et al., 1978). The phospholipid and fatty acid content of the SM most closely resembles that of the endomembrane system (the ER and Golgi apparatus) (Bassarab et al., 1989; Mellor et al., 1985) rather than the PM. The SM also contains a high level of phophatidylcholine, but low levels of phosphatidylethanolamine compared to the PM

(Hernández and Cooke, 1996). The SM contains β-amyrin, a novel plant triterpenoid that is not accumulated in the root PM (Bassarab et al., 1989; Hernández and Cooke, 1996).

During symbiosome maturation to become nitrogen-fixing organelles, changes in vacuole features are induced in order to accommodate rhizobia (Gavrin et al., 2014). Expression of two members of the HOPS vacuole-tethering complex, VPS11 and VPS39, is suppressed in the infected cells resulting in contraction of the vacuole in the host cell, allowing expansion of the

symbiosomes (Gavrin et al., 2014). Moreover, retargeting of the aquaporin TIP1g from the tonoplast membrane to the SM occurs, which permits the maintenance of turgor pressure in the infected cells. These results show that the modifications in vacuole function in host cells are essential for the symbiosome to host rhizobia (Gavrin et al., 2014). Actin-related protein 3

(ARP3) is also essential for symbiosome maturation, in which ARP3 protein and actin were spatially associated with maturing symbiosomes in the infected cells (Gavrin et al., 2015).

Silencing of ARP3 upsets the final differentiation steps of symbiosome maturation into a nitrogen-fixing organelle (Gavrin et al., 2015).

ER and golgi are observed close to symbiosomes (Kijne and Pluvqué, 1979; Whitehead and

Day, 1997) suggesting they are involved in transport of newly synthesized proteins and lipids to the SM via vesicles from the Golgi, but there is little information available about how proteins are targeted to the SM. Targeting of two SS proteins has been investigated. For NOD25 from

M. truncatula, a 24 amino acid signal peptide was sufficient to target GFP to the SS across the

SM (Hohnjec et al., 2009). This suggests a post-translational pathway for its import. The peptide is conserved in two other proteins known to be targeted to the symbiosome. Similarly the early nodulin 8 (MtENOD8) signal peptide (SP) could direct GFP across the SM but two other domains in the protein also had this ability, suggesting redundancy in the targeting signals

(Hohnjec et al., 2009; Meckfessel et al., 2012). Further analysis of other SS and SM proteins is required to determine if N-terminal signal peptides are a common feature and to determine if different proteins follow a common pathway to reach these destinations.

In indeterminant nodules such as M. truncatula and Pisum sativum, the meristemic activity of

the apical meristem results in the formation of a gradient of developmental stages that are defined as four zones: the meristematic zone at the distal end, provides new cells to the nodules; the infection zone, where bacteria are released into the cytoplasm, forming the symbiosome; the fixation zone, where symbiosomes are completely differentiated and the nif genes, encoding the rhizobial nitrogenase complex, are induced (de Maagd et al., 1994) and the senescence zone at the proximal end where symbiosomes degrade (Gage, 2004). Determinate nodules such as

G. max and Lotus japonicus lack a persistent meristem (Brewin, 1991) and have a central infected zone surrounded by nodule parenchyma.

In both the fixation zone of indeterminant nodules and the infected zone of determinant nodules, large infected cells are interspersed with smaller uninfected cells. The uninfected cells are thought to accumulate reduced carbon such as sucrose and convert these to organic acids which are then transported to the infected cells through symplastic transport (Peiter et al., 2004; White et al., 2007). In determinant nodules they are also important in assimilation of fixed nitrogen and are the site of ureide synthesis, which in most determinant nodules is the form in which nitrogen is transported to the shoot (Smith and Atkins, 2002). Senescence of symbiosomes usually begins at week five after inoculation (Vasse et al., 1990). The symbiosomes are targeted and fused to the lytic vacuole for degradation.

1.1.3 Transport activity of the symbiosome membrane

A key role of the SM is to mediate and regulate the exchange of nutrients and metabolites between the symbiotic partners in a way that optimizes nitrogen fixation (Brewin, 1996; Clarke et al., 2014). The SM may also play a role in protecting the bacteria from plant defense

mechanisms (Mellor and Werner, 1987; Vance, 1983) and plants from any pathogenic behaviour of the rhizobia (Brewin, 1996). Since the bacteroid is enveloped by the SM as a result of an endocytosis-like process of the PM, the SM is inverted compared to the PM and hence metabolite movement from the infected cell cytosol to the SS (across the SM) is actually export from the plant cell; conversely, efflux of metabolites from the symbiosome represents uptake into the plant cytoplasm (White et al., 2007).

Information on the transport activity on the SM is essential to determine what regulates the amount of nitrogen fixed and how bacteroids retain a symbiotic relationship with the host plant

(Rosendahl et al., 1991). Our knowledge of the transport activity of the SM mostly results from biochemical studies and transport assays using isolated symbiosomes. Transport assays using radiolabeled substrates and patch clamp techniques have provided insights into the nature of the metabolites that cross the SM, but the molecular identities of most of the transporters involved remain largely unknown. With the availability of complete genome sequences of several legumes (http://phytozome.net/soybean: (Schmutz et al., 2010); http://www.kazusa.or.jp/lotus: (Sato et al., 2008); http://medicago.org/genome: (Young et al.,

2011)), molecular characterization of these transporters is now possible.

To support nitrogen fixation, the plant must provide a carbon source to the bacteroid, in exchange for fixed nitrogen. The principal carbon and energy supply for nodule metabolism is derived from recently fixed plant photosynthetic carbon compounds in the form of sucrose.

There is a large body of evidence that bacteroids derive their energy from oxidative respiration of dicarboxylate acids, principally malate (Day and Copeland, 1991; Figure 1.2). Bacteroids

that are mutant for the dicarboxylate transporter encoded by dctA are unable to fix nitrogen

+ (Yurgel and Kahn, 2004). Ammonia, NH3, or ammonium ions, NH4 , are believed to be the most likely form of fixed nitrogen provided to the plant (Day et al., 2001; Udvardi and Poole,

2013). Reduced carbon for fixed nitrogen is the principal metabolite exchange between the symbiotic partners, but the bacteroid is dependent on the plant for many other (micro)nutrients that include iron, molybdenum, vanadium, nickel cobalt, sulphur, selenium, phosphate and homocitrate (Hakoyama et al., 2009; Rosendahl et al., 1991).

A number of transport activities present on the SM have been identified, with the genes encoding some of the transporters identified (Figure 1.2). The transport processes occurring on the SM include: (i) dicarboxylic acid transport with specificity for univalent malate anions

+ (Herrada et al., 1989; Udvardi, 1988), (ii) NH4 transport by a monovalent cation channel

(Mouritzen and Rosendahl, 1997; Roberts and Tyerman, 2002; Tyerman et al., 1995), (iii) NH3 diffusion through a channel, proposed to be Nodulin 26 (Niemietz and Tyerman, 2000), (iv) proton pumping via an H+-ATPase (Blumwald, 1985; Fedorova et al., 1999), (iv) Fe2+ transport, which maybe mediated by GmDMT1 in soybean (Kaiser et al., 2003; Moreau et al., 1998)

(although there is some controversy as to its direction of transport (Brear et al., 2013)), (v) Fe3+- chelate transport (LeVier et al., 1996; Moreau et al., 1995), (vi) zinc transport by GmZIP1 in soybean (Moreau et al., 2002), (viii) nitrate transport by GmN70 in soybean and LjN70 in L. japonicus (Vincill et al., 2005), (ix) sulphate transport by LjSST1 in L. japonicus (Krusell et al., 2005), and (x) branched-chain amino acid transport in pea (Prell et al., 2009; Udvardi et al.,

1990).

Figure 1.2 Diagrammatic representation of transport processes in the symbiosome of infected cell. Sucrose derived from photosynthesis is the major reduced carbon supply for nodule metabolism. Dicarboxylate acids (principally malate) are supplied to the bacteroids for oxidative respiration and nitrogen fixation. The gene encoding the dicarboxylate transporter on the SM has not been identified. On the BM, a dicarboxylate transport (Dct) system is responsible for importing dicarboxylates into the bacteroids. The product of nitrogenase enzyme, NH3 is transported out of the bacteroid, probably by simple diffusion. NH3 is either + protonated into NH4 and leaves the symbiosome via a monovalent cation channel or remains unprotonated and leaves the symbiosome via NH3 channel proposed to be Nodulin 26. Fe is supplied to the symbiosome as either Fe(II) via DMT1 or Fe(III)-citrate via an uncharacterized Fe(III) transporter which will then be chelated by ferric chelate reductase, which is located on 2- 2+ the SM side. Sulphate, SO4 is transported into the symbiosome via SST1 and zinc, Zn via - ZIP1. An anion transporter with a preference for nitrate, NO3 is encoded by NOD70

In addition to the transporters mentioned above, the SM contains more proteins that have yet to be characterized and that are possibly involved in the transport of various compounds across the SM and/or in the process of maintaining the symbiosis. A recent proteomic analysis of G. max SM has identified 197 proteins as constituents of the SM (Clarke et al., 2015). Two previous proteomic studies of G. max SM had limited success as hydrophobic protein did not resolve well in 2D-PAGE and the unavailability of complete G. max genome sequence, at that time, to match peptides hindered their identification (Panter et al., 2000; Winzer et al., 1999).

The availability of the complete G. max genome sequence, and the use of non-gel proteomic methods and additional techniques to enhance detection of hydrophobic proteins have contributed to the success of identifying more components of the SM (Clarke et al., 2015).

Many of the identified SM proteins of G. max are proteins involved in regulatory processes such as protein folding, RNA regulation, and membrane trafficking, but a portion are predicted transport proteins that are likely to be involved in transport processes across the SM (Clarke et al., 2015). Some of the transport proteins identified can be correlated with known functions.

For example, Glyma09g32110 and Glyma07g09710 are homologous to LjSST1 which is a sulfate transporter on the SM in L. japonicus (Krusell et al., 2005). Other proteins identified on the SM are related to P-type H+-ATPases (Glyma04g34370, Glyma06g20200, and

Glyma19g02270) (Clarke et al., 2015). A P-type H+-ATPase was detected previously on the SM of G. max using specific antibody labeling (Fedorova et al., 1999) and was shown to be involved in energization of the SM. ATPases generate a pH gradient across the SM by pumping protons into the SS while the bacteroid respiratory electron transport chain pumps protons out of the bacteroid into the SS. Consequently, the SS is acidified and this creates SM electrochemical

gradients, which drive exchange of malate and ammonium across the SM (Udvardi and Day,

1997).

Some other transporter types found in the SM proteome are the ATP-binding cassette (ABC) transporters, nitrate transporter 1/ peptide transporter family (NPF) and yellow stripe-like

(YSL) family transporter (Clarke et al., 2015). Five putative proteins Glyma04g34130.1,

Glyma04g34140.1, Glyma02g10530.1, Glyma07580.1 and Glyma10g34700.1 discovered on the SM have homology to ABC transporters (Clarke et al., 2015). This family is of particular interest because the family is very large in plants, and transports a wide range of substrates, primarily out of the cytoplasm (Verrier et al., 2008).

NPF are proton-coupled symporters, which transport their substrates in the cytosolic direction

(Léran et al., 2014). Five NPF members were found in the SM proteome, Glyma08g04160.1

(GmNPF1.2), Glyma11g34613.1 (GmNPF5.24), Glyma11g34600.1 (GmNPF5.25),

Glyma18g03790.1 (GmNPF5.29), and Glyma02g38970.1 (GmNPF8.6). Members of the NPF subfamilies 5 and 8 can transport peptides. A member of the Yellow stripe-like (YSL) family:

Glyma11g31870 (GmYSL7) was also discovered on the SM. Most members of the YSL transport iron (Curie et al., 2009), but an attempt to complement GmYSL7 in a yeast iron transport mutant failed (Qu, unpublished). There is a possibility that GmYSL7 could transport peptides, as its closest homologue in A. thaliana, AtYSL7 can transport an oligopeptide derivative Syringolin A and its transport is inhibited by oligopeptides (Hofstetter et al., 2013).

Two G. max transcriptome atlases have been published, describing gene expression profiles in tissues including leaf, flower, seed, pod, shoot, root and nodule (Libault et al., 2010; Severin et

al., 2010). The data gives an insight into gene expression patterns in specific soybean tissues and possible functions of the predicted genes. Combining all the resources available has provided an essential framework to mine for candidate genes that are possibly essential for symbiosis.

This thesis describes the further characterization of three classes of transport proteins that were identified in the SM proteome (Clarke et al., 2015): ABC transporters, NPF transporters and

YSL family transporter. These proteins were chosen because of their strong and specific expression in the nodule based on the two soybean transcriptome studies (Libault et al., 2010;

Severin et al., 2010) and the roles of these families of transporter are discussed further below.

1.2 ABC transporters

The ATP-binding cassette (ABC) superfamily is one of the largest and most functionally diverse protein families found in both prokaryotes and eukaryotes (Higgins and Linton, 2003).

They are named on the basis of their ATP-binding domain (Hyde et al., 1990), which binds to

ATP and uses the energy from ATP hydrolysis to fuel a variety of biological processes. The majority of ABC proteins are transporters, mediating import or export of substrates across membranes, often against a concentration gradient (Veen and Callaghan, 2003). The substrates transported by ABC transporters are very diverse, ranging from small molecules to macromolecules such as inorganic ions, peptides, polysaccharides and proteins (Higgins, 1992).

ABC transporters are classified as primary transporters, with greater capacity as they have a direct ATP energization mechanism; unlike secondary transporters, which need to establish an

electrochemical potential difference across the membrane for transport (Rea et al., 1998;

Sánchez-Fernández et al., 2001).

1.2.1 Domain organization and structure

An ABC transporter typically consists of four core domains – two cytosolic ATP-binding domains, also known as nucleotide-binding domains (NBDs), and two transmembrane domains

(TMDs) (Higgins et al., 1986). Substrate translocation, in general, requires cooperation and interaction of all four domains (Linton et al., 2003). A topology of a transporter with TMDs preceding the NBDs is considered as a forward orientation, whilst the alternate is referred to as a reverse orientation (Kretzschmar et al., 2011; Rea, 2007).

In bacterial systems, each of the four core domains is often encoded by a separate polypeptide with genes for each arranged in an operon (Hyde et al., 1990). In other systems, the four domains are fused into multi domain polypeptides in a number of ways (Higgins and Linton,

2003; Hyde et al., 1990). Where all four domains are fused to yield a single polypeptide they are termed full-size transporters. This arrangement is often found in eukaryotic ABC transporters such as the multidrug resistance (MDR) proteins (Endicott and Ling, 1989; Higgins and Linton, 2003). Where one of the four domains is absent (half-size transporter), the proteins need to form homodimers or heterodimers to form a functional transporter (Higgins et al.,

1988). For instance, in A. thaliana, half transporter AtABCG11 was demonstrated to form homodimers and also heterodimers with AtABCG12 (McFarlane et al., 2010). Interestingly, the different dimer combinations were implicated in the different behavior of these transporters during trafficking to the PM (McFarlane et al., 2010).

1.2.1.1 Nucleotide-binding domains (NBDs)

In contrast to the TMDs, the structures of the NBDs are highly conserved. This suggests that the NBDs share a common evolutionary origin and the same mechanism of transport (Linton and Higgins, 2007). The NBD is defined by six short sequence motifs: Walker A, Q-loop,

Signature, Walker B, D-loop and H-loop (Ames et al., 1992; Dassa, 2011; Higgins et al., 1988;

Jones and George, 2002; Linton and Higgins, 1998; Walker et al., 1982; Figure 1.3).

Structural analyses have shown that all six motifs are necessary to form a complete ATPase active site (Hopfner and Tainer, 2003; Procko et al., 2009). The Walker A motif, also known as the P-loop, has a consensus sequence of GxxGxGKT/S, where x represents any amino acids.

The Walker A motif interacts directly with ATP. Biochemical and structural evidence suggest that the highly conserved lysine residue of the Walker A motif is crucial – it forms hydrogen bonds with phosphates of ATP, fixing ATP in a defined orientation (Davidson et al., 2008). The

Walker B motif (hhhhDE, where h represents a hydrophobic residue) also interacts with nucleotides and may coordinate the Mg2+ ions required for ATP hydrolysis (Hopfner et al.,

2000).

Whilst the functions of the Walker motifs are reasonably well understood, the individual functions of the other four motifs are ambiguous: (i) the Q-loop is presumed to participate in the interaction between the NBD and the TMD during nucleotide hydrolysis (Procko et al.,

2009); (ii) the Signature motif (LSGGQ), located upstream of the Walker B motif, is unique to

Figure 1.3 Linear representation of the six characteristic motifs, which defined the NBD found in ABC transporters. The NBD is a 200‒250 amino acids stretch of polypeptides located in the cytoplasm. Please refer to the text for the individual functions of the six motifs. The one-letter amino acids code is used, where x represents any of the 20 standard amino acids and h stand for hydrophobic amino acids. Diagram reproduced from Dassa (2011).

ABC proteins and is thought to be involved in ATP hydrolysis (Gaudet and Wiley, 2001); (iii) the D-loop that contains aspartate is postulated to stabilize the Walker A motif in a position favorable for catalysis (De La Rosa and Nelson, 2011); (iv) the H-loop (also referred to as the switch region), is thought to interact with the nucleotide γ-phosphate, coordinating the bound nucleotides (Davidson et al., 2008).

1.2.1.2 Transmembrane domains (TMDs)

The TMDs represent the translocation conduit of ABC transporters, consisting of multiple membrane-spanning α-helices and include the substrate-binding sites and intra-cytosolic loop

(ICL) domains (Higgins and Linton, 2001; Jones and George, 2013). Many ABC transporters are predicted to form six hydrophobic transmembrane α-helices within each module, though this number can vary in some cases (Holland and Blight, 1999). It is intriguing that the different numbers of α-helices between TMDs and the low sequence similarity they share might reflect the diverse substrates ABC transporters can transport. Substrate crosses the lipid bilayer of a cellular membrane via the translocation pathway formed by lined residues from the α-helices

(Dawson and Locher, 2006). ABC transporters may contain a single broad substrate-binding site or multiple sites with selective substrate specificity, in which both TMDs are necessary for substrate recognition and binding (Arora et al., 2001; Veen and Callaghan, 2003). The ICL domains form a physical interface with the NBDs and are implicated in coordinating and coupling the energy from ATP binding and hydrolysis for substrate transport (Dawson and

Locher, 2006).

1.2.2 Plant ABC transporters

ABC transporters were first extensively characterized in bacteria, and then in humans as they are of considerable medical importance. Defects or mutations in half of the 48 ABC transporters identified in humans have been linked to human diseases, mostly caused by abnormal lipid transport (reviewed in Štefková et al., 2004; Tarling et al., 2013). In addition, multidrug resistance occurs as the result of increased activity of drug efflux by MDR1. Consequently, cancer cells become resistant to chemotherapeutic drugs as this lowers the concentration of intracellular drugs to a non-toxic level (Gottesman et al., 2002; Ueda et al., 1987).

Plants have more ABC transporters than human and non-plant taxa (Jasinski et al., 2003;

Sánchez-Fernández et al., 2001). The genomes of A. thaliana and Oryza sativa encode more than 120 ABC proteins compared with 50 to 70 proteins encoded by the human, Drosophila melanogaster and Caenorhabditis elegens genomes (Sánchez-Fernández et al., 2001). The high number of ABC transporters in plants might be attributed to their sessile nature, that does not allow them to quickly avoid environmental and biotic stress (Rea et al., 1998). Plants need to transport exogenous toxins (xenobiotics), heavy metals, secondary metabolites and photo oxidative products from photosynthesis out of the cytosol or out of the cell for detoxification.

The lack of excretory organs places great demand on the plant's detoxification machinery

(Sánchez-Fernández et al., 2001). In fact, the first plant ABC transporter discovered mediated transport of xenobiotic glutathione S-conjugates into barley vacuoles (Hortensteiner et al.,

1993). Though plant ABC transporters were first implicated in detoxification processes, numerous studies have now shown that ABC transporters are involved in diverse physiological processes such as plant growth, development and interaction with the environment.

The nomenclature approved by the Human Genome Organization (HUGO) has been adopted as a unified naming system for plant and vertebrate/invertebrate ABC transporters (Verrier et al., 2008). The nomenclature clusters ABC transporters into eight subfamilies (A-H) and for plants, an additional subfamily (I) was created to accommodate 'prokaryotic'-type ABCs. The classification into subfamilies is based on amino acids alignment of NBDs and phylogeny, domain organization and content, e.g. full-size versus half-size transporters, and presence of additional domains (Verrier et al., 2008). This nomenclature will be used throughout this thesis.

In recent years, concurrent heterologous expression and forward and/or reverse genetics have uncovered many functional roles of plant ABC transporters (Verrier et al., 2008). Plant ABC transporters have been implicated in auxin transport, plant-pathogen interactions, stomatal regulation and many other important physiological functions (summarized in Appendix 1).

They are mainly localized to the PM, but are also found on the membrane of organelles such as the vacuoles, plastids, peroxisomes and mitochondria (reviewed in Kretzschmar et al., 2011;

Verrier et al., 2008; Yazaki et al., 2009). Subfamilies of ABC transporters characterized to date are discussed further below.

1.2.2.1 Subfamily A

The majority of subfamily A transporters are half-size transporters and are present in the forward orientation (TMD-NBD). In human, many ABC subfamily A transporters transport lipids and lipid-related compounds such as phospholipids and sterols to diverse organs and cell- types (Tarling et al., 2013; Vasiliou et al., 2009), and mutations in the transporters are causative of many diseases, including Tangier disease, Alzheimer’s disease and Stargardt disease

(Bodzioch et al., 1999; Macé et al., 2005; Tsybovsky et al., 2010).

There are twelve predicted ABCA transporter genes in A. thaliana, eleven of them are half-size transporters whereas G. max has seven predicted ABCA transporter, six of which are half-size transporters (Takanashi and Yazaki, 2014; Verrier et al., 2008). Two members of ABCA subfamily in G. max were identified in the proteomic analysis of the SM: Glyma04g34130.1

(now named Glyma04G172900.1 in genome assembly version 2.0) and Glyma04g34140.1

(now named Glyma04G17300.1 in genome assembly version 2.0) (Clarke et al., 2015).

Only two members of the plant ABCA transporters are characterized so far – AtABCA2

(Biedrzycki et al., 2011) and AtABCA9 (Kim et al., 2013). AtABCA2 is expressed in the root

(Badri et al., 2008), and is involved in kin (from the same mother plant) recognition in A. thaliana, showing changing expression when exposed to kin or stranger (from a different ecotype) root secretions (Biedrzycki et al., 2011). The compound it transports has not been characterized. AtABCA9 transports fatty acids for triacylglycerol (TAG) biosynthesis to the ER during seed-filling stage (Kim et al., 2013). abca9 mutant seeds have lowered TAG content and radioactive feeding experiment showed that AtABCA9 transports both acyl-CoAs and free fatty acids. In plants, fatty acids are synthesized in the plastids from acetyl-Coenzyme A (CoA), which are then converted to acyl-CoAs in the chloroplast envelope (Chapman and Ohlrogge,

2012). Acyl-CoAs are subsequently transported through the cytosol to the ER for biosynthesis of TAG, which is the final product in the formation of seed storage lipids (Chapman and

Ohlrogge, 2012). It was unclear how acyl-CoA was exported into the ER until AtABCA9 was found to be responsible for the transport (Kim et al., 2013).

1.2.2.2 Subfamily B

Subfamily B represents the second largest ABC subfamily in A. thaliana and G. max with 29 members and 49 members, respectively (Takanashi and Yazaki, 2014). ABCB transporters have the forward topology, TMD-NBD. The most studied ABCB transporters in plants are

AtABCB1, AtABCB4 and AtABCA19, which are implicated in auxin or indole acetic acid

(IAA) transport on the PM (reviewed in Xu et al., 2014). Interestingly, AtABCB4 and Coptis japonica ABCB1 are some of the few plant ABC importers, a characteristic not common in other eukaryotic ABC transporters (Santelia et al., 2005; Shitan et al., 2003; Verrier et al., 2008).

A member of the ABCB subfamily was identified in the proteomic analysis of the SM:

Glyma02g10530.1 (Glyma.02G094800.1, version 2.0) (Clarke et al., 2015).

1.2.2.3 Subfamily C

All members of plant ABCC are full-size transporters often with a hydrophobic N-terminal extension in the forward orientation (Andolfo et al., 2015; Rea et al., 1998). The prototype from human, multidrug resistance-associated protein (MRP) 1 (HsMRP1) is involved in export of glutathione conjugates from drug-resistant cell lines. Characterization of HsMRP1 led to identification of plant MRPs (also called GS-X pumps), which also sequester glutathione- conjugates into the vacuole (Ishikawa et al., 1997; Verrier et al., 2008). Though plant ABCCs were considered classical GS-X pumps (reviewed in Rea et al., 1998), subsequent studies demonstrated that they are involved in vacuolar anthocyanin transport, heavy metal sequestration, detoxification of chlorophyll catabolites and glucuronides, and inositol hexakisphosphate transport.

1.2.2.4 Subfamily D

The plant ABC subfamily D is represented by two members in Arabidopsis and L. japonicus, and five members in G. max. ABCD transporters are in the forward orientation. The most studied member in the ABCD subfamily is AtABCD1, also known as Peroxisomal ABC transporter 1 (PXA1), Peroxisome defective 3 (PEP3) or Comatose (CTS). ABCD1 is an importer, mediating the import of acyl-CoA fatty acid into peroxisomes (reviewed in Linka and

Theodoulou, 2013). Additionally, AtABCD1 is required for transport of auxin precursors, indole-3-butyric acid (IBA) and jasmonic acid (JA) precursors into the peroxisomes (Strader and Bartel, 2011; Theodoulou et al., 2005). A recent study in Hordeum vulgare showed involvement of HvABCD1 and HvABCD2 in IBA import and jasmonate biosynthesis, suggesting possible conservation of functions of monocot and dicot ABCD transporters

(Mendiondo et al., 2014).

1.2.2.5 Subfamily G

ABCG forms the largest subfamily of ABC transporters with 43 members in Arabidopsis, 36 members in L. japonicus and 108 members in G. max (recently reviewed in Banasiak and

Jasiński, 2014). The expansion of the G subfamily in plants seems to have been associated with taxon-specific functional diversification (Verrier et al., 2008). A distinct feature of this subfamily is that the domains are in reverse orientation, with the NBD domain located at the amino terminal end of the protein. ABCGs can be divided into two groups: (i) half-size transporters also known as white brown complex (WBC), homologues of the prototypical member from Drosophila melanogaster, which contain one NBD and TMD; and (ii) full-size transporters also known as pleiotropic drug resistance (PDR) transporters, which comprise of

two NBDs and TMDs (Banasiak and Jasiński, 2014). ABCGs have diverse roles in plants and transport a wide range of substrates such as abscisic acid (ABA), wax, cutin, IBA and terpenoids. Glyma08g07580.1 (Glyma.08G071200.1, version 2.0) and Glyma10g34700.2

(Glyma.10G203000.2, version 2.0) were identified through proteomic analysis of the soybean

SM (Clarke et al., 2015) as ABCG transporters.

1.2.2.6 Subfamily I

Plant possesses an additional subfamily, designated subfamily I which bear similarities to components of the prokaryotic multisubunit ABC transporters (Verrier et al., 2008). The most studied members are the AtABCI4, AtABCI5 and AtABCI3 also known as trigalactosyldiacyl glycerol 1 (TGD1), TGD2 and TGD3, respectively. They are found in the chloroplast inner envelope and were shown to play a role in the transport of ER-derived lipids to the plastid (Awai et al., 2006; Lu and Benning, 2009).

1.3 NPF Transporters

In plants, the nitrate transporter 1/ peptide transporter family (NPF) includes transporters previously named nitrate transporter (NTR) and peptide transporter (PTR) (Léran et al., 2014).

The NPF belong to the major facilitator superfamily (MFS) and are secondary active transporters, utilizing an inwardly directed electrochemical proton gradient to drive transport of substrates across cell membranes (Daniel et al., 2006; Newstead, 2015). The NPF in plants is large, with 53 members in Arabidopsis and 134 members in G. max, whereas in non-plant organisms there are only a few members. The knowledge gained about NPF transporters is primarily from investigations in A. thaliana, with a few exceptions in other plant species A

unified nomenclature for naming these proteins was recently developed in plants (Léran et al.,

2014). Members are named based on phylogenetic relationships between the family members, dividing them into eight major subfamilies (NPF1-8). Names are designated accordingly:

NPFX.Y, where X represents the subfamily or clade number, and Y denotes the individual member within the species (Léran et al., 2014).

Plant NPF members have been demonstrated to transport a wide range of substrates including amino acids, dicarboxylates, glucosinolates, ABA, auxin and nitrite. Intriguingly, a number of plant NPF transporters can also transport more than one substrate. For example, AtNPF2.10 can transport nitrate and glucosinolates while AtNPF6.3 is capable of transporting nitrate and auxin. Evidence to date suggests that sequence homologies between family members do not reflect the substrates they can transport. The subfamilies of NPF members characterized to date are discussed below.

1.3.1 NPF subfamilies

1.3.1.1 Subfamily 1

MtNPF1.7 also known as NIP/LATD (for Numerous Infections and Polyphenolics/Lateral root- organ Defective) is a high affinity nitrate transporter and an ABA transporter, involved in nodulation and regulating root architecture (Bagchi et al., 2012; Zhang et al., 2014). Using a yeast two-hybrid approach, AtNPF1.1 and AtNPF1.2 were shown to transport gibberellin (GA) and the plant hormone, jasmonoyl-isoleucine (JA-Ile) (Chiba et al., 2015). One of the NPF candidates in this study, GmNPF1.2, is found in this subfamily and was identified in the SM proteome (Clarke et al., 2015).

1.3.1.2 Subfamily 2

This subfamily predominantly consists of nitrate transporters. Eight NPF2 members function as nitrate transporters: AtNPF2.3 (Taochy et al., 2015), AtNPF2.7 (NAXT1, Segonzac et al.,

2007), AtNPF2.9 (NRT1.9, Wang and Tsaya, 2011), AtNPF2.12 (NRT1.6, Almagro et al.,

2008), AtNPF2.13 (NRT1.7, Fan et al., 2009), OsNPF2.2 (Li et al., 2015c) and OsNPF2.4 (Xia et al., 2015). Two glucosinolates transporters, AtNPF2.10 (GTR1) and AtNPF2.11 (GTR2) that control the loading of glucosinolates from the apoplasm into the phloem have also been identified (Nour-Eldin et al., 2012).

1.3.1.3 Subfamily 3

NPF3 subfamily has the least members. Only three members have been functionally characterized so far. The nitrite transporter Cucumis sativus NPF3.2 (CsNitr1) is localized in the chloroplasts where it is involved in the removal of toxic nitrite from the cytosol (Sugiura et al., 2007). Two other NPF3 transporters: AtNPF3.1 and Vitis vinifera NPF3.2 displayed nitrite and nitrate transport activities in Xenopus oocytes (Pike et al., 2014). A recent study showed that AtNPF3.1 is also able to transport GA (David et al., 2016).

1.3.1.4 Subfamily 4

AtNPF4.6 (NRT1.2/AIT1) is a low affinity nitrate transporter and has recently been demonstrated to mediate uptake of cellular ABA to regulate stomatal aperture in inflorescence stems (Huang et al., 1999; Kanno et al., 2012). Another member AtNPF4.1 (AIT3) transports

ABA, GA and JA-Ile when expressed in yeast (Chiba et al., 2015; Kanno et al., 2012). ABA

transport activity was also observed in AtNPF4.5 (AIT2), whilst AtNPF4.2 showed GA transport activity (Chiba et al., 2015).

1.3.1.5 Subfamily 5

This subfamily represent the largest subfamily with 663 members from the 33 plant genomes

(Léran et al., 2014). The first characterized member, AtNPF5.2 (AtPTR3) mediated transport of di- and tripeptides in yeast that include His-Leu, His-Phe, Leu-Met, Leu-His-Leu and Met-

Leu-Gly (Karim et al., 2007). AtNPF5.2 is implicated in defense against a virulent bacterial pathogen by transporting defense-related peptides (Karim et al., 2007). AtNPF5.2 was recently shown to transport ABA and GA1, while both AtNPF5.7 and AtNPF5.1 were demonstrated to transport ABA, GA1 and JA-Ile (Chiba et al., 2015). AtNPF5.5 and AtNPF5.10 have recently been identified as nitrate transporters (Léran et al., 2015). Another NPF5 member in rice,

OsNPF5.5 (OsPTR3) did not confer di- and tripeptide uptake activity in yeast (Ouyang et al.,

2010). Six of the NPF candidates in this thesis, GmNPF5.2, GmNPF5.24, GmNPF5.25,

GmNPF5.29, GmNPF5.3 and GmNPF5.30, belong to the NPF5 subfamily, and three of them were identified on the SM (Clarke et al., 2015).

1.3.1.6 Subfamily 6

This subfamily contains the first nitrate transporter identified in plants: AtNPF6.3 (NRT1.1).

AtNPF6.3 acts as dual-affinity nitrate transporter by adapting to fluctuations in soil nitrate levels (Huang et al., 1996; Liu et al., 1999; Wang et al., 1998). AtNPF6.3 also facilitates the uptake of auxin and at low nitrate availability, AtNPF6.3 represses lateral root growth by stimulating auxin transport out of the roots (Krouk et al., 2010). Three other members are also

nitrate transporters: AtNPF6.2 (NRT1.4, Chiu et al., 2004), AtNPF6.4 (NRT1.3, Okamoto et al.,

2003) and MtNPF6.8 (NRT1.3, Morère-Le Paven et al., 2011). MtNPF6.8 is a dual-affinity nitrate transporter and also transports ABA when expressed in Xenopus oocytes (Pellizzaro et al., 2014).

1.3.1.7 Subfamily 7

Two nitrate transporters: AtNPF7.2 (NRT1.8, Li et al., 2010) and AtNPF7.3 (NRT1.5, Chen et al., 2012; Drechsler et al., 2015; Lin et al., 2008) were identified in this subfamily. When expressed in yeast, OsNPF7.3 (PTR6) transports the di- and tripeptides, Gly-His and Gly-His-

Gly (Ouyang et al., 2010). OsNPF7.3 over expression is suggested to enhance plant growth by increasing ammonium transporter expression and glutamine synthetase activity (GSA) (Fan et al., 2014).

1.3.1.8 Subfamily 8

Three members of this subfamily: AtNPF8.1 (PTR1), AtNPF8.2 (PTR5) and AtNPF8.3 (PTR2) have been identified as dipeptide transporters (Komarova et al., 2008; Rentsch et al., 1995).

AtNPF8.1 and AtNPF8.2 are able to transport dipeptides such as Ala-Ala, Ala-Asp, Ala-Lys, and His-Ala when expressed in yeast and Xenopus oocytes (Dietrich et al., 2004; Komarova et al., 2008). AtNPF8.1 mediates uptake of dipeptides into root cells, whereas AtNPF8.2 facilitates transport of peptides into germinating pollen (Komarova et al., 2008). AtNPF8.3 is localized to the vacuole and seemed to transport a wider range of dipeptides (Chiang et al., 2004; Rentsch et al., 1995) compared to AtNPF8.1 and AtNPF8.2. Antisense AtNPF8.3 showed delay in flowering, and has reduced seed numbers per silique. This suggests an important role of

AtNPF8.3 in peptide transport during seed development and flowering (Song et al., 1997). A member in Z. mays, ZmNPF8.8, is also a peptide transporter that mediate uptake of Ala-Ala from the growing media (Tnani et al., 2013). OsNPF8.9 (OsNRT1.1) from rice, encodes a constitutively expressed low-affinity nitrate transport system (Lin et al., 2000), while

OsNPF8.20 (OsPTR9) did not show nitrate, di- or tripeptide uptake activity when expressed in yeast or Xenopus oocytes (Fang et al., 2013). GmNPF8.6, an SM protein that is a candidate in this study is in this sub-family (Clarke et al., 2015).

1.4 Oligopeptide transporters

1.4.1 Yellow stripe-like transporters

The Yellow stripe-like (YSL) family is part of the wider oligopeptide transporter (OPT) superfamily. Members of the YSL family are involved primarily in the transport of metals, complexed with nicotianamine (NA) or phytosiderophores (PS) (reviewed in Conte and Walker,

2012). Both NA and PS are metal chelators, but only grasses produce PS, where its secretion by the plant increases solubility of metals, especially iron, in the rhizosphere. All plants produce

NA which binds to metals (Higuchi et al., 1994), forming metal-NA complexes for movement of metals throughout the plant. Formation of these complexes is essential in their long-distance transport (Schuler et al., 2012).

The founding member of the YSL family, Zea mays yellow stripe 1 (ZmYS1) (Curie et al.,

2001; Von Wiren et al., 1994) has a wide substrate specificity, mediating transport of Fe(II)-

NA, Fe(III)-NA, Ni(II)-NA, Ni(II)-deoxymugineic acid (DMA, a PS derived from NA), and

Cu-mugineic acid (MA) (Roberts et al., 2004; Schaaf et al., 2004). Eight YSL members in A.

thaliana were identified from sequence similarity with ZmYS1 (Curie et al., 2001). A. thaliana

YSL members are mostly implicated in the transport of iron chelated with NA. For instance,

AtYSL1 functions in mediating the transport of Fe-NA to developing seeds (Jean et al., 2005),

AtYSL2 is involved in the lateral movement of metals within the vasculature (DiDonato et al.,

2004) while AtYSL4 and AtYSL6 are involved in detoxifying iron during plastid dedifferentiation (Chu et al., 2013).

A study showed that Arabidopsis ysl3 is more susceptible to pathogen Pseudomonas syringe pv. tomato compared to the wild type (WT) suggesting that AtYSL3 is involved in pathogen defense (Chen et al., 2014) . AtYSL3 was positively regulated by salicylic acid (SA) signaling and was negatively regulated in the coi1 mutant defective in the jasmonic acid (JA) receptor.

These results indicated that YSL3 is involved in SA and JA signaling for involvement in pathogen-induced defense (Chen et al., 2014).

Until recently there was no information on the substrates for AtYSL7 and AtYSL8. However, a recent study showed that AtYSL7 and AtYSL8 mediated the transport of a peptide derivative

Syringolin A (SylA), which is a virulence factor secreted by plant pathogen Pseudomonas syringae pv. syringae (Hofstetter et al., 2013). When expressed in yeast, AtYSL7 and AtYSL8 mediated growth inhibition of yeast on a medium containing SylA and this inhibition was blocked if oligopeptides were included in the medium. The Arabidopsis ysl7 mutant had reduced SylA sensitivity in a root growth inhibition assay and in leaves both ysl7 and ysl8 mutants showed reduced effects of SylA treatment. These results suggested that the physiological substrate for AtYSL7 and AtYSL8 may be oligopeptides and that P. syringae

hijacked them for transport of SylA into host cells (Hofstetter et al., 2013).

One YSL protein, GmYSL7, was identified in the proteomic analysis of SM (Clarke et al.,

2015). GmYSL7A is part of a family consisting of 15 members in soybean (Schmutz et al.,

2010). Initial analysis suggested this protein could not transport Fe2+-NA (Qu, unpublished).

On the basis of this result and the report by Hofstetter et al. (2013) it was chosen for further analysis as an oligopeptide transporter in this study.

1.4.2 True OPTs

Nine members of the true OPTs were identified in the model species A. thaliana (AtOPT1 to

AtOPT9), exhibiting 49 to 53% sequence similarity to the initially characterized yeast OPTs

(Koh et al., 2002). Plant OPT members are also identified in O. sativa (OsGT1; Vasconcelos et al., 2008; Zhang et al., 2004) and Brassica juncea (BjGT1; Bogs et al., 2003). In yeast, OPT members Candida albicans CaOPT1, Schizosaccharomyces pombe Isp4p, and S. cerevisiae

ScOPT1 and ScOPT2 were shown to transport tetra- and pentapeptides such as KLGL, KLLG and KLLLG (Hauser et al., 2000; Lubkowitz et al., 1998; Lubkowitz et al., 1997). ScOPT1 can also mediate the uptake of GSH, Met-enkephalin (YGGFM) and Leu-enkephalin (YGGFL)

(Bourbouloux et al., 2000; Hauser et al., 2000).

Similarly, plant OPT members are able to transport the tripeptide GSH, tetra- and pentapeptides but with different affinities. For example, AtOPT4 have been shown to rescue yeast leucine auxotroph on media containing KLLG, KLGL and KLLLG peptides as the sole leucine source

(Koh et al., 2002). In addition, methionine-containing peptides of GGFM, IIGLM and YGGFM

were transported by AtOPT4 in a yeast methionine auxotroph (Osawa et al., 2006). Some members are implicated in the transport of iron in different forms. For instance, AtOPT3 mediates iron transport in long-distance signaling between shoots and roots, and to developing seeds (Stacey et al., 2008; Zhai et al., 2014), whilst OsOPT1, OsOPT3, OsOPT4, OsOPT5 and

OsOPT7 could transport ferrous and/or ferric iron chelated to nicotianamine, Fe(III)-NA

(Vasconcelos et al., 2008). Expression analyses of AtOPTs mRNA showed various expression patterns, but all are strongly expressed in the vasculature of different tissues including cotyledons, hypocotyls, leaves, roots, flowers and siliques (Koh et al., 2002; Stacey et al.,

2006). This suggests that they might have distinct but broad and overlapping function in nitrogen distribution to various tissues.

1.5 Objectives of the present study

Considerable advances have been made in the proteomic study of the soybean SM, which identified a comprehensive array of proteins that are constituents of the SM (Clarke et al., 2015).

Only a small fraction of the identified proteins have been characterized to date, either by biochemical methods or by genetic and/or molecular characterization. Clearly, there is a need for better understanding of the physiological roles these SM proteins play, as they are likely to be involved in the transport processes across the SM, and regulation and maintenance of the rhizobia-symbiosis in general.

This study seeks to functionally characterize two ABC subfamily A proteins, GmABCA1 and

GmABCA2 that were identified in the SM proteome of G. max (Clarke et al., 2015). Some of the initial work on these transport proteins has been performed by Clarke (2013) and where

applicable will be compared to results in the present study. GmABCA1 and GmABCA2 are hypothesized to possess the ability to transport fatty acids or lipids, based on functional homologies with other ABCA members. Heterologous complementation in yeast and reverse genetic approaches were used to test this hypothesis. The results of the investigations are presented in Chapter 3.

GmNPFs and GmYSL7 were also identified in the SM proteome of G. max (Clarke et al., 2015).

Initial characterization suggested that these proteins were essential for the symbiosis (Gavrin and Smith, unpublished data). Given that many members of the NPF transport peptides and that

AtYSL7 appears to transport oligopeptides, Chapter 4 utilizes heterologous complementation in yeast to examine the ability of these proteins to transport peptides.

Chapter 2

General materials and methods

2.1 Growing wild type soybean plants

G. max cv. Stephens seeds were inoculated with a peat-slurry inoculant of B. japonicum

USDA110 (Soybean Nodulaid® Group H, Becker Underwood, USA), and grown in river sand under natural light in a temperature controlled glass house set to long-day conditions (16 h day length, 26°C day/20°C night). Seedlings were re-inoculated at seven days post-planting. Plants were irrigated with B&D nutrient solution (Broughton and Dilworth, 1971) without nitrogen twice a week.

2.2 Isolation of nucleic acids from plant tissue

2.2.1 Genomic DNA extraction

Genomic DNA (gDNA) was extracted from leaves for promoter amplification and genotyping

PCRs. Fresh leaves were frozen in liquid nitrogen and ground into fine powder using a plastic pestle. The homogenate was lysed by adding 600 µl lysis buffer (10 mM Tris-HCl, 1 mM

EDTA, 1% (w/v) SDS). The lysate was incubated at 65°C for 1 h with occasional mixing followed by incubation at 4°C for 5 min. 2 M ammonium acetate was added to the lysate and the mixture was vortexed thoroughly to precipitate proteins and other contaminants from the cell lysate. The precipitant was removed by centrifugation at 16,000 x g for 10 min. The supernatant was transferred to a new tube and DNA was recovered by adding 0.6-0.7 volumes of ice-cold 100% (v/v) isopropanol. The supernatant was carefully decanted and the DNA pellet was washed with ice-cold 70% (v/v) ethanol followed by centrifugation for 5 min. The DNA pellet was air dried and re-dissolved in 30-50 µl TE buffer. DNA yield and purity was determined spectrophotometrically using a NanoDrop® (Thermo Scientific, USA).

2.2.2 Total RNA extraction and cDNA synthesis

Purification of total RNA from plant tissue was carried out using RNeasy® Plant Mini Kit

(QIAGEN, Germany) according to manufacturer’s protocols. Optional on-column DNase digestion was performed to remove contaminating DNA. The integrity and quantity of purified

RNA was assessed using NanoDrop® spectrophotometer and gel electrophoresis. RNA was reversed transcribed to cDNA using iScript™ cDNA Synthesis Kit (BioRad, USA) following the manufacturer’s instructions. 1 µg RNA was used as the template.

2.3 Gene expression analysis

Quantitative real time PCR (qRT-PCR) assay was performed to quantify the expression of candidate genes in four soybean tissues and during six stages of nodule development.

2.3.1 Plant material, RNA isolation and cDNA synthesis

Soybean plants were grown as described in Section 2.1. Mature tissues from leaf, stem, root and nodule (32 DPI) and nodules of different developmental stages (11, 14, 19, 32 and 42 DPI) were harvested and pooled from a minimum of two plants for each biological replicate. Two independent biological replicates were sampled simultaneously, and immediately flash-frozen in liquid nitrogen before storage at -80°C. Total RNA extraction and cDNA synthesis were carried out by Ella Brear (The University of Sydney) as outlined in Section 2.2.2.

2.3.2 Primer design and reference gene

Specific qRT-PCR primers were designed for each gene based on sequence prediction in soybean genome assembly Glyma1.1 using the Primer3 program (Untergasser et al., 2012). The following parameters were used to design the primers: primer size 17-25 bp long, GC content between 50-60%, primer Tm 50-60˚C with compatible melting temperatures (within 5˚C), and amplicon size between 150 and 300 bp that spanned exon-exon junctions and flanked a long intron to avoid contaminating gDNA amplification.

GmUBI (Glyma.20G141600) was chosen as the reference gene for normalizing gene expression as its expression is stable and ubiquitous. Expression stability of GmUBI was confirmed by

Patrick Louglin (The University of Sydney) using geNorm software (Vandesompele et al.,

2002), when compared with other proposed soybean reference genes (Libault et al., 2008).

GmUBI primer sequences were obtained from Trevaskis et al. (2002).

2.3.3 Validating primer specificity

The specificity of the primer sets was evaluated via standard PCR amplification on nodule cDNA and post qRT-PCR melt curve analysis. A single gene-specific PCR product of the correct size is expected when visualized by agarose gel electrophoresis. The PCR product was subsequently purified (§2.4.4), sequenced (Macrogen, South Korea) and analyzed using

SnapGene® software to confirm only a single amplicon was amplified in the reaction. Melt curve analysis following reaction completion was undertaken by analyzing the dissociation curve profile of each gene.

2.3.4 qRT-PCR conditions

The qRT-PCR assay was performed using a LightCycler® 480 instrument (Roche Life Science,

Germany), and SYBR Green I (Roche) was used as the real-time molecular reporter. All reactions were set up manually in 384-well plate format (Roche), with each reaction performed in triplicates. PCR mix was prepared following manufacturer’s protocols, but the reactions were scaled down to 5 µl. Each reaction was composed of 1x SYBR Green I Master Mix (contains

FastStart Taq DNA Polymerase, reaction buffer, dNTP mix, SYBR Green I dye, and MgCl2),

0.5 µM of each PCR primer, and approximately 10 ng cDNA. cDNA was replaced by PCR grade water in the negative control reaction. Cycling parameters used for real-time PCR

reaction were pre-incubation at 95°C for 5 min, 45 cycles of denaturation at 95°C for 10 s; annealing at 56°C for 20 s; extension at 72°C for 30 s. After the completion of the amplification program, melting curve analysis was performed with the following settings: denaturation at

95°C for 10 s, annealing at 55°C for 1 min and gradual temperature increment to 95°C at a ramp rate of 0.06°C/s. The amount of florescence was recorded during both programs.

2.3.5 Data analysis

The resulting qPCR data was analyzed using LightCycler® 480 system software version 1.5

(Roche). The non-baseline corrected data was imported into LinReg software (Ramakers et al.,

2003) to calculate PCR efficiency (Peff) for each primer pair, which is represented by the slope of the linear regression line (Eff10slope). The value was used to calculate the threshold cycle (Ct) value for each gene, followed by normalization to the GmUBI reference gene (ΔCt), through subtracting the reference gene Ct value (Ctreff) from each test gene Ct value (Ctgene). The average

−ΔΔCt of the expression levels between the technical triplicates was calculated using the 2 method

(Livak and Schmittgen, 2001), with outliers more than one Ct value different within the triplicates removed from the calculation. Final results were represented as means of expression levels between the two biological replicates.

2.4 Gene cloning using Gateway® Technology

Gateway® Technology takes advantage of the bacteriophage lambda site-specific recombination system. The recombination facilitates transfer of DNA sequences flanked by att sites between

Gateway® vectors via lysogenic pathway (catalyzed by BP Clonase™ enzyme) and lytic pathway (catalyzed by LR Clonase™ enzyme) (Hartley et al., 2000; Landy, 1989).

2.4.1 Vectors

Gateway® vectors were propagated and maintained in DB3.1 E. coli strain (genotype F- gyrA462 endA1 Δ (sr1-recA) mcrB mrr hsdS20 (rB-, mB-) supE44 ara14 galK2 lacY1 proA2 rpsL20 (Smr) xyl5 Δleu mtl1). DB3.1 is resistant to the effects of a product produced by the ccdB gene present on Gateway® donor and destination vectors and thus used to propagate and maintain vectors harboring the ccdB gene. Gateway® vectors have a chloramphenicol resistance gene (CmR), allowing counter selection of plasmid on LB (1% (w/v) tryptone, 0.5% (w/v) yeast extract, 1% (w/v) NaCl, pH 7.0) plate containing 15 µg/ml chloramphenicol.

2.4.2 Designing attB PCR primers

To produce PCR products that could be used as substrates for BP recombination reaction with the donor vector, primers of the PCR products need to incorporate attB sites. The forward primer was designed to have four guanine (G) residues at the 5’ end followed by the 25 bp attB1 site and 18-25 bp gene specific sequences. To fuse PCR product in frame with an N-terminal reporter protein and attB1 region, two nucleotides were added to the attB1 site end. The reverse primer was designed to contain four guanine residues at the 5’ end followed by the 25 bp attB2 site and 18-25 bp gene specific sequences. To fuse PCR product in frame with a C-terminal reporter protein and attB2 region, one additional nucleotide was added at the end of attB2 site.

A stop codon was added to the reverse primer if fusion of the PCR product and the C-terminal tag was not required. Primers were designed using Primer3 software (Untergasser et al., 2012)

with the following parameters; primer size 18-27 bp (optimum 20 bp), primer Tm 57-63°C

(optimum 60°C), GC content 40-60%. MOPC™-purified oligonucleotides were synthesized by

Macrogen (South Korea).

2.4.3 Producing attB-PCR products cDNA or gDNA were used as the template for amplifying attB-PCR products. Reactions contained 1x Phusion High-Fidelity (HF) Buffer, 200 µM dNTPs, 0.5 µM of each forward and reverse primers, 50-250 ng DNA template and 0.02 U/µl Phusion® HF DNA polymerase.

Cycling parameters used were initial denaturation at 98°C for 30 s, 35 cycles of denaturation at

98°C for 30 s; annealing at primers specific Tm for 30 s; extension at 72°C for 15-30 s/kb and final extension at 72°C for 5 min.

2.4.3.1 Agarose gel electrophoresis

Gel electrophoresis was used to determine yield and quality of PCR products. PCR products were resolved by gel electrophoresis on a 1-2% (w/v) TAE (40 mM TRIS, 20 mM acetate, 1 mM EDTA, pH 7.6) agarose gel with 1x SYBR® Safe DNA gel stain (Invitrogen, USA). 0.8-1

µg 2-log DNA ladder (New England Biolabs) was used as the standards for size determination and quantification of PCR products. PCR products were mixed to a concentration of 1x loading gel buffer and reactions were run in 1x TAE buffer at 110V. Gels were visualized and photographed under UV light using G:BOX (Syngene, UK) operated by GeneSnap software version 7.09.

2.4.4 Purifying PCR products attB-PCR products were purified to remove attB primers and attB primer-dimer products as these products can also recombine with donor vector resulting in increased background after transformation.

2.4.4.1 PEG/ MgCl2 purification

PEG/ MgCl2 purification is recommended by the manufacturer as large primer-dimer products and DNA less than 100 bp can be effectively removed. Purification was performed based on manufacturer’s protocols. The integrity and quantity of the recovered attB-PCR products were assessed using agarose gel electrophoresis and NanoDrop® spectrophotometer.

2.4.4.2 PCR clean-up

Alternatively, attB-PCR products were purified using NucleoSpin® Gel and PCR Clean-up kit

(Macherey-Nagel, Germany) following the manufacturer’s instructions.

2.4.4.3 Gel extraction

When multiple bands were acquired, all PCR products were loaded on agarose gel and a single band was excised and purified using the same kit as above.

2.4.5 Generating entry clones using the BP recombination reaction

BP recombination reactions were performed between an attB-PCR product and an attP- containing donor vector to generate an attL-containing entry clone. The reaction was catalyzed

by BP Clonase™ enzyme mix. 8-25 fmol attB-PCR product, 37.5 ng pDONR™221 vector, and

1 µl BP Clonase™ enzyme mix were mixed, and TE buffer added to bring the reaction to 5 µl.

The reaction was incubated at 25ºC overnight. 2 µg Proteinase K solution was added and incubated at 37ºC for 10 min to terminate the reaction. Transformation was performed as described in Section 2.6. Transformants were screened via colony PCR for colonies with the desired plasmid (§2.7.1). Plasmids from positive entry clones were isolated (§2.7.2), analyzed by restriction enzyme digest (§2.7.3) and sequenced (Macrogen, South Korea) to confirm sequence integrity of the cloned DNA.

2.4.6 Generating expression clones using the LR recombination reaction

LR recombination reactions were performed between an attL substrate (entry clone) and an attR-containing destination vector, yielding an attB-containing expression clone. The reaction was catalyzed by LR Clonase™ enzyme mix. 25-75 ng entry clone, 37.5 ng destination vector, and 1 µl LR Clonase™ enzyme mix were mixed and TE added to bring the reaction to 5 µl. The reaction was incubated at 25ºC overnight. Reaction termination, transformation into E. coli and screening for positive transformants were as described above.

2.5 Preparation of E. coli competent cells

DH5α E. coli strain (genotype F- recA1 endA1 hsdR17 (rk-, mk+) supE44 λ- thi-1 gyrA96 relA1) were used for transformation. Competent cells were prepared using a calcium chloride

(CaCl2) protocol as outlined in Sambrook et al. (1989). A single colony was inoculated in 5 ml

LB and grown overnight at 37°C. 1 ml overnight culture was diluted in 100 ml LB and left

shaking for 2-3 h at 37°C. Cells were incubated on ice for 10 min before centrifugation at 3000 x g for 10 min at 4ºC. The cell pellet was gently resuspended in 10 ml ice-cold sterile 0.1 M

CaCl2 followed by incubation on ice for 20 min. Cells were centrifuged again and the cell pellet was gently resuspended in 5 ml ice-cold sterile 0.1 M CaCl2 containing 15% (v/v) glycerol.

Aliquots of 50 µl were frozen in liquid nitrogen and stored at -80°C until use.

2.6 Heat-shock transformation of chemically competent bacteria

Transformation of CaCl2-treated cells was performed according to Sambrook et al. (1989) with a few modifications. An aliquot of 50 µl chemically competent cells was thawed on ice, and

2.5-5 µl reaction mix or plasmid DNA was added to the thawed cells. Cells were incubated on ice for 30 min, followed by heat shock at 42°C for 30 s. Tube was immediately placed on ice for 5 min. 450 µl S.O.C (2% (w/v) peptone, 0.5% (w/v) yeast extract, 10 mM NaCl, 2.5 mM

KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM glucose) was added to the tube and cells were grown for 1 h at 37°C with shaking. 50-100 µl of the transformed culture was spread onto a selection plate containing appropriate antibiotics. The plate was incubated at 37°C overnight.

2.7 Screening for transformed E. coli cells

2.7.1 Colony PCR

Colony PCR was employed to rapidly screen transformants for the presence or absence of insert in plasmid DNA. An individual colony was added directly to the PCR reaction and lysed during denaturation step of PCR. Using a pipette tip, a small patch of a single colony was streaked onto LB plate and then resuspended in the PCR reaction mix containing the following

components; 0.1 µl NEB Taq DNA polymerase, 1x ThermoPol Reaction Buffer, 200 µM dNTPs and 200 nM gene specific forward and reverse primers. Cycling parameters used were based on the manufacturer’s recommendation for Taq DNA polymerase. PCR products were visualized using agarose gel electrophoresis (§2.4.3.1) and plasmids were extracted from colonies that had the correct size amplification product.

2.7.2 Isolation of plasmid DNA

Plasmid DNA was isolated using UltraClean® 6 Minute Mini Plasmid Prep Kit (MO BIO

Laboratories, USA) according to the manufacturer’s protocol. Plasmid was eluted in 20 µl

Milli-Q water and the concentration was measured using NanoDrop® spectrophotometer.

2.7.3 Restriction enzyme digest

Restriction digests were used to verify vector backbone and insert. Plasmid was digested with appropriate enzymes to produce fragments of defined sizes that were diagnostic for a particular vector/ insert combination. The resulting fragments were resolved by gel electrophoresis.

Digestion was carried out following manufacturer’s recommendation specific to the restriction enzyme type. Plasmid that contained all the expected fragments was sequenced (Macrogen,

South Korea) and resulting sequences were analyzed using SnapGene® software.

2.8 Preparation of Agrobacterium competent cells

Agrobacterium competent cells were prepared based on protocol described in Hofgen and

Willmitzer (1988). A 5 ml overnight culture was diluted in 200 ml YEM and grown for 3-4 h at

28ºC with shaking. Cells were incubated on ice for 5 min followed by centrifugation at 3000 x g for 20 min at 4ºC. The cell pellet was washed in ice-cold TE and centrifuged again. The pellet was gently resuspended in 10 ml YEM medium. Aliquots of 50 µl were frozen in liquid nitrogen and stored at -80°C until use.

2.9 Agrobacterium transformation

Expression clones were transformed into Agrobacterium using the freeze-thaw method (Hofgen and Willmitzer, 1988). Plasmid DNA of 0.5-1.0 µg was added to thawed competent cells. The cells were incubated successively on ice, in liquid nitrogen and at 37ºC, each for 5 min. 1 ml

LB was added and left shaking for 2-4 h at 28ºC. Aliquot of 150 µl was plated on LB agar with appropriate antibiotics and grown for two days at 28ºC. Colonies were screened for positive transformants as described below.

2.10 Screening for transformed Agrobacterium cells

2.10.1 Colony PCR

For Agrobacterium colony PCR, individual transformants were lysed in water with a brief heating step rather than added directly into the PCR reaction. Individual colonies were picked using a pipette tip and suspended in sterile 50 μl Milli-Q water. Resupended colonies were incubated at 100ºC for 2 min followed by 5 min on ice. Cell debris was spun down for 1 min using a microcentrifuge. 1.5 µl of the heat treated colony was used as the template and PCR was performed as described in Section 2.7.1.

2.10.2 Agrobacterium plasmid isolation and restriction digest

Colonies for which the correct size amplification product was obtained were analyzed further to verify the presence of the right plasmid. Plasmid was extracted from Agrobacterium and restriction digest was performed. UltraClean® 6 Minute Mini Plasmid Prep Kit was used with a number of modifications to the protocol. A 5 ml overnight culture was harvested, and 150 µl

Solution 1 and two tungsten carbide beads (3mm diameter, QIAGEN) were added into the tube.

Disruption and homogenization was performed using TissueLyser LT (QIAGEN), operating for

2 min at 30 Hz. 250 µl Solution 2 and 450 µl Solution 3 were added and lysate was centrifuged for 10 min at full speed. Clear supernatant was transferred to a spin filter and centrifuged again.

Plasmid DNA was washed with 300 µl Solution 4 and eluted in 23 µl Milli-Q water. Restriction enzyme digests were performed as described in Section 2.7.3.

2.11 Gene transfer to plants cells via Agrobacterium rhizogenes-mediated

hairy root transformation

An A. rhizogenes-mediated hairy root transformation procedure was adapted from Kereszt et al. (2007) and Mohammadi-Dehcheshmeh et al. (2014). This procedure was used to introduce expression vectors containing the gene of interest to soybean roots. A. rhizogenes induces hairy roots at the site of infection through the transfer of its root-inducing (Ri) plasmid along with a binary vector containing gene of interest into the plant genome.

2.11.1 Seed germination

G. max seeds were surface-sterilized in 70% (v/v) ethanol for 2 min by gentle inversion and rinsed a few times with sterile water. The sterilized seeds were placed into wet sterile 1:1 vermiculite: perlite at a depth of 1-2 cm and grown under artificial lighting (OSRAM, 28W/840) set to long-day condition (16 h day length, 26°C day/20°C night) for 5 days.

2.11.2 Hairy roots induction

Two days prior to plant inoculation, a single colony of A. rhizogenes harboring the desired plasmid was inoculated in LB medium with antibiotic selection and grown for 48 h at 28ºC. On the day of inoculation, cells were harvested at 3000 rpm for 6 min, and washed in sterile water.

The pellet was resuspended in sterile water. Healthy plantlets with unfolded cotyledons, 3–5 cm length were used for inoculation. Approximately 3 µl of a bacterial suspension was injected into the hypocotyl close to the cotyledonary node using a 1 ml, 16 G syringe needle. Seedlings were placed into a 100-1000 µl sterile pipette tip (with its end cut off) to prevent the seedlings being exposed to the nutrient solution. Seedlings were transferred into plastic jars (around 15 cm height x 6 cm width), containing 3-4 cm sterile sand, which were subsequently sealed to maintain high humidity. 40 ml B&D nutrient solution with nitrogen (Broughton and Dilworth,

1971) was supplied to the plants. Seedlings were grown under artificial lighting with the same condition as above for 2.5 weeks.

2.11.3 Removal of non-transgenic roots

After 2.5 weeks, 2-5 cm hairy roots should emerge from the inoculation site. Stems below the infection site were removed and plants were transferred into a 30-cell tray filled with sand.

Hairy roots were left to grow for another week. After a week, some or all untransformed hairy

roots were cut off to prevent formation of nodules on untransformed roots. Transformed hairy roots fluoresce green or red when irradiated with excitation light (450-490 nm for GFP, 540-

580 nm for DsRED) due to the expression of either a GFP or DsRED reporter gene. Non- fluorescent untransformed roots were removed using a scalpel (Figure 2.1A). Fluorescence was visualized under a Fluorescence Stereo Microscope Leica M205 FA with 470/40 nm excitation filter for GFP (emission wavelength of 500-550 nm) or 545/30 nm excitation filter for DsRED

(emission wavelength of 593-667 nm). Images were captured using a digital color camera Leica

DFC310 FX.

2.11.4 Maintaining plants and screening for transgenic nodules

Following removal of untransformed roots, plants were transferred into pots with river sand

(Narellan Sand, Soil & Garden Supplies) and grown in the glasshouse (16 h day length, 26°C day/20°C night). Plants were inoculated with B. japonicum USDA 110 (Nodulaid Group H,

Becker Underwood, USA) and were re inoculated a week afterwards. B&D nutrient solution

(Broughton and Dilworth, 1971) without nitrogen was provided once per week and plants were watered every 2-3 days. At 23-28 DPI, plants were screened for transgenic nodules that fluoresced green or red (Figure 2.1B) using the same microscopy settings as described above.

Only transgenic nodules were selected for further experimentation.

2.12 Preparation of yeast competent cells

Yeast competent cells were prepared based on a modified protocols by Dohmen et al. (1991).

To make yeast competent cells, a single yeast colony was grown in 100 ml YPD (1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) glucose) to an OD600 of 0.6-1.0, followed by

Figure 2.1 Screening for transformed hairy roots and nodules. (A) The left panel shows transformed hairy roots that fluoresced green when irradiated with excitation light while untransformed roots did not fluoresce (indicated by arrows). Images to the right shows only transformed roots were remained as untransformed roots were removed using scalpel. (B) Transgenic roots and nodules expressing another reporter gene DsRED, fluoresced red when irradiated with excitation light (left). Non-transgenic nodules (indicated by arrows) did not fluoresce and were not used for further experimentation (right). Scale bars represent 1 mm.

centrifugation at 3000 x g for 10 min. The resulting pellet was washed in 20 ml Solution A (10 mM BICINE (N,N-Bis(2-hydroxyethyl)glycine), 1 M sorbitol, 3% (v/v) ethylene glycol, pH

8.35) and centrifuged again. The yeast pellet was resuspended in 2 ml Solution A. Aliquots of

100 µl were stored at -80°C until use.

2.13 Yeast transformation

Yeasts were transformed with the desired yeast expression vectors as described in Dohmen et al. (1991). 50 ng of the desired plasmid, 25 µg heat-denatured salmon sperm DNA and 2.5 mM histamine were added to the frozen yeast competent cells, followed by rapid agitation at 37°C for 5 min. 500 µl Solution B (200 mM BICINE, 40% (w/v) PEG 1000, pH 8.35) was then added and left shaking at 30°C for 1 h. Cells were pelleted by centrifugation at 3000 x g for 5 min, washed in 400 µl Solution C (10 mM BICINE, 150 mM NaCl, pH 8.35) and centrifuged again.

The pellet was resuspended in 50 µl Solution C and plated on a selective transformation plate.

Chapter 3

Characterization of GmABCA1 and GmABCA2 in soybean nodules

3.1 Introduction

Members of the ABC transporters are integral membrane proteins, and in plants localize to various subcellular membranes as well as to the PM and transport a wide variety of substrates.

Two integral membrane proteins with homology to ABC subfamily A transporters were discovered on the SM of G. max, GmABCA1 (Glyma04g34130.1 v1.1, Glyma.04G172900.1 v2.0) and GmABCA2 (Glyma04g34140.1 v1.1, Glyma.04G173000.1 v2.0) (Clarke et al.,

2015). These assigned systematic names as proposed by Verrier et al. (2008) will be used to

refer to these ABCA proteins henceforth. Expression data from the two soybean transcriptome studies showed that GmABCA1 and GmABCA2 have nodule-enhanced expression and very little expression in other tissues (Libault et al., 2010; Severin et al., 2010). In search for proteins that are particularly important for the symbiosis, they were chosen for further study because of their upregulated expression within the nodules.

GmABCA1 and GmABCA2 are members of the ABC transporter subfamily A which mainly transports lipids and lipid-related compounds such as phospholipids, fatty acids and sterols

(Tarling et al., 2013). An ABCA member in plants, AtABCA9, has been demonstrated to mediate the transport of fatty acids and acetyl-CoA for TAG biosynthesis in the ER during seed-filling stage (Kim et al., 2013). Based on the functional homologies with other ABCA members, GmABCA1 and GmABCA2 could potentially transport lipids or fatty acids across the SM. Aside from the requirement for lipids for symbiosome biogenesis during the early stages of nodule development, to my knowledge, other roles for lipids in the symbiosome have not been proposed. It is hypothesized that within the symbiosome, lipids or fatty acids either from the plants or the bacteroids might function as a signaling molecules, to co-ordinate the behavior of the symbionts. They might also be an alternative form of carbon supplied to the bacteroid.

3.2 Materials and Methods

3.2.1 Protein alignment and phylogenetic analysis

A maximum likelihood (ML) tree was constructed for ABC subfamily A proteins in G. max and A. thaliana using Molecular Evolutionary Genetics Analysis (MEGA) version 6.0 software

(Tamura et al., 2013). Amino acid sequences of G. max ABCA proteins were obtained from

Phytozome (Goodstein et al., 2012) and SoyBase (Grant et al., 2010). Arabidopsis ABCA proteins were downloaded from The Arabidopsis Information Resource (TAIR, Lamesch et al.,

2012). Sequence alignment of the amino acid sequence dataset containing 19 ABCA proteins was built by MUSCLE aligner (Edgar, 2004). The software constructed the ML tree from the aligned sequences using the best substitution model, computed to be the Neighbor- Joining (NJ) algorithm with a matrix of pairwise distances of Jones–Thornton–Taylor (JTT) (Jones et al.,

1992) plus discrete gamma (+G) model. Bootstrap tests of 1000 replicates were evaluated, with a bootstrap value more than 40 percent labeled at the corresponding nodes. Optimal global sequence alignment of two proteins was examined using EMBOSS Needle (Li et al., 2015b).

3.2.2 Gene expression analysis

The steady state of GmABCA1 and GmABCA2 mRNA levels in different tissues, and also during nodule development were estimated and analyzed using real-time quantitative PCR (RT- qPCR). Gene expression in four soybean tissues (leaf, stem, root and nodule at 32 DPI), and throughout nodule development (at 11, 13, 19, 32 and 42 DPI) was measured and the data normalized to the soybean reference gene, GmUBI. Procedures were carried out as detailed in

Section 2. Primers used for the assay are shown in Table 3.1.

3.2.3 In planta localization

3.2.3.1 Amplification and cloning

Both GmABCA1 and GmABCA2 full-length CDS were amplified and cloned as outlined in

Table 3.1 Oligonucleotide primers used for RT-qPCR of GmABCA candidates and ubiquitin reference gene. Primers were designed using the Primer3 tool with the following parameters: primer size 17-25 bp long, GC content between 50-60%, Tm 50- 60˚C with no more than 5˚C difference between the Tm of primer pairs, and amplicon size between 150-300 bp that spans different exons. GmUBI primer sequences were obtained from Trevaskis et al. (2002). GENE FORWARD PRIMER REVERSE PRIMER SEQUENCE (5'→3') SEQUENCE (5'→3') GmABCA1 AAGTGGTTTATATGGACGAG CATTTGGGAAGAGCTGTCG GmABCA2 GCGCTCCTGAAAAATTTC GAACCAAGTAGGGTTTTCA GmUBI GTGTAATGTTGGATGTGTTC ACACATTGAGTTCAACACAA CC ACCG

Section 2.4 based on sequence predictions in the soybean genome version 1.1. Oligopeptides used for the amplifications are as listed in Table 3.2. GmABCA1 CDS was first subcloned into pGEM®-T Easy (Promega) as the PCR product was unable to be directly cloned into pDONR™221. Entry and expression clones were generated by Gateway® BP and LR recombination reactions, respectively. pENTR clones were recombined into p35S-pK7FWG2

(Figure 3.1) for C-terminal GFP fusion, and pGmLbC3-pK7WGF2-R (Figure 3.2) for N- terminal GFP fusion.

GmABCA1 and GmABCA2 genes were expressed under the control of the G. max leghemoglobin C3 (Glyma10g34260, now named Glyma.10G198800.1 in Glyma2.0) promoter for localization studies. The expression level of GmLbC3 is highly enhanced in nodules according to the two soybean transcriptome studies (Libault et al., 2010; Severin et al., 2010), where it is primarily expressed in the infected cells (Nguyen et al., 1985) and is active five to six days following infection until nodule senescence (Cheon et al., 1993; Franche et al., 1998).

Its high expression has successfully driven sufficient expression of number of SM transporters studied in our lab such as GmNPF5.29 (Clarke et al., 2015) and GmZIP1 (Chen, 2012). For these reasons, GmLbC3 promoter was chosen.

3.2.3.2 Biolistic bombardment in Allium porrum L.

As attempts to express GFP-tagged GmABCA1 in soybean failed to show any fluorescence,

GmABCA1 was transiently expressed in Allium porrum L. (leek) epidermal cells via biolistic bombardment according to Collings et al. (2002) with a few modifications. Mature leek plants obtained from a local grocer were bombarded with 1 µm gold particles (Bio-Rad Laboratories,

Table 3.2 Primer sequences for Gateway® cloning of GmABCA1 and GmABCA2 CDS. The attB2 and attB2 recombination sites are underlined. Primer sequences were designed by Clarke (2013). DNA FRAGMENT FORWARD PRIMER REVERSE PRIMER SEQUENCE (5'→3') SEQUENCE (5'→3') GmABCA1 CDS GGGGACAAGTTTGTACAAA GGGGACCACTTTGTACAAG AAAGCAGGCTTAATGGAA AAAGCTGGGTATTATGACA AATGGAAACACAGCC AGGTGTTAAA TGCCTGA GmABCA2 CDS GGGGACAAGTTTGTACAAA GGGGACCACTTTGTACAAG AAAGCAGGCTTAATGGCGA AAAGCTGGGTATCATTGAA CCACTATCAGC AATCCACTGAACTAACTT

Figure 3.1 Plasmid map of destination vector, p35S-pK7FWG2 (Karimi et al., 2002) used for C-terminal GFP fusion driven by 35S promoter. Abbreviations: ori – origin of replication, RB – right border, LB – left border, CaMV 35S promoter – constitutive promoter from cauliflower mosaic virus, CmR – chloramphenicol resistance gene, ccdB – ccdB death gene, EGFP – enhanced GFP, and Sm/SpR – spectinomycin and streptomycin resistance genes. Vector map created using SnapGene® software.

Figure 3.2 Plasmid map of destination vector, pGmLbC3-pK7WGF2-R (Gavrin et al., 2016) used for N-terminal GFP fusion driven by GmLbC3 promoter. Abbreviations: ori – origin of replication, RB – right border, LB – left border, LbC3 promoter – Leghemoglobin C3 promoter, EGFP – enhanced GFP, CmR – chloramphenicol resistance gene, ccdB – ccdB death gene, T35S – terminator of 35S, Ubi promoter – Ubiquitin promoter, DsRED – Discosoma sp. red fluorescent protein, NOS terminator – nopaline synthase terminator, and Sm/SpR – spectinomycin and streptomycin resistance genes. Vector map created using SnapGene® software.

USA) coated with 250 ng of each plasmid DNA of p35S-pK7FWG2-GmABCA1 and PM- mCherry marker (Nelson et al., 2007). Leek squares were bombarded at approximately 400 kPa helium gas and under 550 mm Hg vacuum. Bombarded leek squares were placed on Murashige and Skoog (MS) (Murashige and Skoog, 1962) agar and incubated overnight in the dark, at room temperature. Microscopic analysis was carried out the next day as described in Section

3.2.3.5.

3.2.3.3 Soybean transformation and growth

Soybean transformation and growth were carried out as detailed in Section 2.11.

3.2.3.4 Confocal microscopy

Fluorophores were visualized and photographed using a Leica TCS SPII Multi-photon at The

Australian Centre for Microscopy and Microanalysis (ACMM), University of Sydney and a

Zeiss Confocal LSM Pascal 510 at The Plant Molecular Biology Lab, University of Sydney.

Transgenic soybean nodules were sectioned by hand using a double edge knife and were then

® placed in phosphate buffer (0.1 M NaH2PO4-Na2HPO4). 35 µg/ml FM 4-64 (Invitrogen) was added to the sections and they were incubated for 1 h at 4°C prior to imaging. FM®4-64 is a membrane stain, goes into the endomembrane system and fluoresces when bound to the membrane. Bombarded leek squares were viewed directly under the microscope. GFP fluorophore was excited with a 488 nm argon laser and emission was collected through a 505-

530 nm filter. FM®4-64 and mCherry fluorophores were excited using a 543 nm HeNe laser and detected through a 560-615 nm emission filter.

3.2.4 Bioinformatics analysis

Bioinformatics analysis was performed for the CDS of both GmABCA1 and GmABCA2. The

BLASTP program (Altschul et al., 1997) was used to examine the translated CDS against a protein database for sequence similarities and conserved domains. Membrane protein topology were predicted by four bioinformatic tools: Phobius (Käll et al., 2004), SOSUI (Hirokawa et al., 1998), TMHMM (Krogh et al., 2001) and HMMTOP (Tusnády and Simon, 2001). Signal peptides were predicted by SignalP 4.1 (Petersen et al., 2011).

3.2.5 In situ PCR

The localization of the GmABCA1 and GmABCA2 transcripts in tissues and cell types was examined in soybean nodule cells via the in situ PCR method. The experiment was performed at The University of Adelaide, with the guidance of Asmini Athman. Procedures for in situ mRNA detection on a fixed nodule sections were performed as described in Athman et al.

(2014) with a few modifications. Soybean nodules of 32 DPI were halved and placed in fixative solution (63% (v/v) ethanol, 5% (v/v) acetic acid, 2% (v/v) formaldehyde) and kept at 4°C overnight. The next day, fixed nodules were embedded in 8% agarose for sectioning, with vibratome settings: 1 mm sec-1 (speed), 1 mm (amplitude), and 50 µm (section thickness).

Reverse transcription was done using Phusion® High Fidelity DNA polymerase. GmUBI with ubiquitous expression and GmYSL7 with infected cell specific expression acted as controls.

GmYSL7 forward and reverse primers were designed by Patrick Loughlin (The University of

Sydney) with the following sequences: 5’-TCTCATGGGCTATCATGTGG-3’ and 5’-

ATGTGCCCAAGCTTTTTCTG-3’, respectively.

3.2.6 Heterologous complementation of Δfat1

The ability of GmABCA1 and GmABCA2 to transport fatty acids were assessed through complementation of a fatty acid uptake-deficient yeast mutant, Δfat1 (Færgeman et al., 1997).

Growth complementation assay protocols outlined by Li et al. (2015a) were followed, with

ScFAT1 as the positive control. The antibiotic cerulenin was supplied in the assay, which irreversibly inhibits fatty acid synthase (FAS) and resulted in blockage of the de novo pathway for fatty acid biosynthesis (Vance et al., 1972). Yeast is consequently auxotrophic for fatty acids and media needs to be supplemented with exogenous fatty acids (C12-C18) for yeast to be viable. Δfat1 had reduced growth even after exogenous fatty acids were supplied (Færgeman et al., 1997).

The Δfat1 yeast deletion mutant (W303; MATa; ura3-1; trp1Δ2; leu2-3_112; his3-11; ade2-1; can1-100; YBR041w(43, 847)::kanMX4), and ScFAT1 in pRS416 were obtained from

European Saccharomyces Cerevisiae Archive for Functional Analysis (Euroscarf). pENTR

GmABCA1 and GmABCA2 full-length CDS were LR cloned into a Gateway® compatible yeast expression vector, pDR196-GW (Figure 3.3). ScFAT1 in pRS146 and expression clones

GmABCA1 and GmABCA2 in pDR196-GW, along with the empty vector pDR196 were transformed into Δfat1. Ura+ transformants were selected on SD agar containing 0.67% (w/v)

YNB without amino acids, 2% (w/v) glucose, 20 mg/l adenine, histidine, leucine and tryptophan.

Figure 3.3 Plasmid map of Gateway®-compatible yeast expression vector, pDR196- GW, modified from Rentsch et al. (1995). Abbreviations: 2µ ori – yeast 2μ plasmid origin of replication, CmR – chloramphenicol resistance gene, ccdB – ccdB death gene, AmpR – Ampicillin resistance gene, URA3 promoter – uracil biosynthesis promoter, URA3 – encodes orotidine-5'-phosphate decarboxylase, required for uracil biosynthesis, lac promoter - promoter for the E. coli lac operon. Vector map created using SnapGene® software.

A single colony of Δfat1 yeast harboring either ScFAT1, GmABCA1, GmABCA2 or empty vector were grown in liquid SD minus uracil (2% (w/v) glucose, 0.5% (w/v) Brij 58, 0.7% (w/v)

KH2PO4) until non-exponential phase at 30°C. Cells were harvested and diluted to an OD600 of

0.03 in SD minus uracil supplemented with 10 µM cerulenin and 100 µM oleate and grown until log phase. Cell densities were measured using UV-VIS spectrophotometer at multiple time points.

3.2.7 Functional complementation of the Atabca9 mutant

3.2.7.1 Plant material and growth conditions

A. thaliana ecotype Colombia-0 (Col-0) was used in all experiments. Seeds were germinated and cultivated either in solid MS (Murashige and Skoog, 1962) or in soil. For sterile growth conditions in MS media, seeds were surface sterilized for 5 min in 70% (v/v) ethanol, for 3 min in 5% (w/v) bleach, 0.1% (v/v) Triton X-100 and were finally rinsed 4-5 times in sterile water.

For growing seeds in non-sterile growth conditions, seeds were evenly distributed on the surface of potting mix (Debco plugger) supplemented with Osmocote® slow release fertilizer. Plates and pots were stratified at 4°C for 3 days prior to transfer to the growth room. Plants were grown under artificial lighting (OSRAM, 28W/840) at 22°C set to long-day condition (16 h light and 8 h dark).

3.2.7.2 Identification of homozygous abca9 mutant plants

Seeds for At5g61730 transfer-DNA (T-DNA) insertion line, Salk 058070, were obtained from

TAIR (Alonso et al., 2003). Seeds were segregating T3 or T4 generations, thus genotyping PCR

was employed to identify individuals homozygous for the mutant alleles. SIGnAL T-DNA primer design at signal.salk.edu/tdnaprimers.2.html was used to generate primers for the genotyping PCR. The Left Primer (LP) and Right Primer (RP) were used to amplify the wild type allele, while Left Border Primer (LB), LBb1.3 and RP were used to amplify the T-DNA insertion allele. Primer sequences are listed in Table 3.3. Genomic DNA was extracted from at least seven Salk 058070 plants at 4 weeks old (§2.2.1). The genotyping PCR was carried out as detailed in Section 2.7.1 using approximately 100 ng gDNA as template with 60°C annealing temperature for all reactions. Col-0 was used as the control.

3.2.7.3 Vector construction

PCR amplification and Gateway® cloning were done as described in Section 2.4. The

AtABCA9 native promoter from 2 kb upstream of the ATG start codon and the full-length

AtABCA9 gDNA of 4.3 kb were amplified using primers with attached att sites (Table 3.4) and

Col-0 gDNA as the template. GmABCA1 and GmABCA2 primers were designed as detailed in Section 2.4.2 and incorporated specific att sites, an ER retention signal HDEL (Denecke et al., 1992) and also a stop codon (Table 3.4). Both CDS were amplified from vectors containing the previously cloned full-length CDS.

To generate complementation lines of atabca9, three constructs were generated: (i) AtABCA9 native promoter fused to full-length AtABCA9 gDNA (positive control), (ii) AtABCA9 native promoter fused to GmABCA1 CDS-HDEL and (iii) AtABCA9 native promoter fused to

GmABCA2 CDS-HDEL. The two DNA fragments were fused and cloned using MultiSite

Table 3.3 Primer sequences for genotyping PCR of Arabidopsis Salk 058070. Primers were designed by SIGnAL T-DNA primer design with the following default parameters: primer size 18-28 bp (optimum 21 bp), primer Tm 53-71°C (optimum 61°C), GC content between 20-80%, Max N 300 bp, Ext5 300 bp, Ext3 300 bp, primer zone 200 bp and BPos 110 bp. N is the difference between the actual insertion site and the flanking sequence position. Max N is the maximum difference between the actual insertion site and the sequence. Ext5 and Ext3 are the regions between the Max N to primer zone. The primer zone is the region used to design primers. BPos is the distance between BP and the insertion site. PRIMER PRIMER SEQUENCE (5'→3') DESCRIPTION LP CGGAGACTCTGGAATCAGTTG Left genomic primer RP AAAGAGGTGGAGGTGCTCTTC Right genomic primer LBb1.3 ATTTTGCCGATTTCGGAAC Left border primer of the T- DNA insertion

Table 3.4 Primer sequences for MultiSite Gateway® cloning of Atabca9 complementation constructs. Specific att sites flanking insert were incorporated into forward and reverse primers to allow recombination with pDONR™221 P1-P5r or pDONR™221 P5-P2. Primer sequences for AtABCA9 were obtained from Kim et al. (2013). The att recombination sites are underlined. An ER retention signal, HDEL (Denecke et al., 1992) was added to GmABCA1 and GmABCA2 reverse primers for ER targeting of the proteins (highlighted in blue). A stop codon was added after the retention signal (highlighted in red). DNA att FORWARD PRIMER REVERSE PRIMER FRAGMENT SITES SEQUENCE (5'→3') SEQUENCE (5'→3') AtABCA9 attB1, GGGGACAAGTTTGTACA GGGGACAACTTTTGTATA native attB5r AAAAAGCAGGCTTAGC CAAAGTTGTGATCACAGA promoter ACGGTGTGAACATTAAT GGAAGAAGAAGATTAGA TGGA G AtABCA9 attB5, GGGGACAACTTTGTATA GGGGACCACTTTGTACAA gDNA attB2 CAAAAGTTGTAATGACT GAAAGCTGGGTATTAATT CTGCGAGAAGGCTT GTTAGATTCATAATCAAT GmABCA1 attB5, GGGGACAACTTTGTATA GGGGACCACTTTGTACAA CDS attB2 CAAAAGTTGTAATGGAA GAAAGCTGGGTATTAGAG AATGGAAACACA CTCATCATGTGACAAGGT GT GmABCA2 attB5, GGGGACAACTTTGTATA GGGGACCACTTTGTACAA CDS attB2 CAAAAGTTGTAATGGCG GAAAGCTGGGTATTAGAG ACCACTATCAGCGGC CTCATCATGTTGAAAATC CACTGAAC

Gateway® Pro 2-fragment recombination technology (Invitrogen). In this system, two entry clones and a destination vector are recombined in a LR reaction to generate an expression vector consisting the two DNA fragments in the desired order and orientation. To create the two entry clones, two separate BP reactions were performed. Entry clone I was generated from recombination of attB1 and attB5r-flanked AtABCA9 native promoter with pDONR™221 P1-

P5r. While entry clone II was produced from recombination of attB5 and attB2-flanked

AtABCA9 gDNA/ GmABCA1 CDS-HDEL/ GmABCA2 CDS-HDEL with pDONR™221 P5-P2.

Together, entry clone I and II and a destination vector, pB7WG∆2 (Figure 3.4), were recombined in a LR reaction to create the expression vector for transformation into A. tumefaciens strain GV3101.

3.2.7.4 Transformation via the floral dip method

A single colony of A. tumefaciens that contained the desired expression vector was inoculated in 5 ml YEP media (1% (w/v) peptone, 1% (w/v) yeast extract, 0.5% (w/v) NaCl, pH 7.0) containing 100 µg/ml spectinomycin and was grown overnight at 28°C. The starter culture was diluted approximately 1:1000 into YEP medium containing 100 µg/ml spectinomycin and 100

µg/ml gentamicin. The culture was grown until midlogarithmic phase, OD600 0.6-0.8. The density was measured using a UV-Vis Spectrophotometer (Shimadzu). 0.02-0.05% (v/v) Silwet

L-77® and 0.06% (w/v) freshly made sucrose were added to the culture before dipping.

Only healthy plants that were bolting and had many unopened flower buds were used for the transformation. The Agrobacterium culture was transferred to a 500 ml measuring cylinder to enable plant dipping. Plants were inverted into the culture until the above-ground tissues were

Figure 3.4 Plasmid map of Gateway® binary T-DNA destination vector, pB7WG∆2 (Karimi et al., 2002), for complementation with genomic sequences. Abbreviations: ori – origin of replication, RB – right border, LB – left border, CmR – chloramphenicol resistance gene, ccdB – ccdB death gene, T35S – terminator of 35S, BlpR – Basta/ phosphinothricin/ glufosinate resistance gene, NOS – nopaline synthase and Sm/SpR – spectinomycin and streptomycin resistance genes. Vector map created by SnapGene® software.

fully submerged. Plants were removed from the culture after 20-30 s, and then gentle agitation was applied to remove excess culture from the dipped plants. At least 8 plants were dipped per construct (AtABCA9 gDNA, GmABCA1 CDS-HDEL, GmABCA2 CDS-HDEL and empty vector). Because the plants were tall, they were positioned on their side in trays. They were placed under a dome covered with aluminum foil to maintain high humidity and to keep them away from direct light. Domes were removed 12-24 h after dipping.

3.2.7.5 Maintaining plants and seeds harvest

Plants were grown for 3-5 weeks after transformation. Watering was stopped when most of the siliques turned yellow. During seed harvesting, siliques and stems were gently pulled through fingers over a piece of paper. Plant debris was separated from the seeds by sifting through a sieve. Seeds harvested from transformed plants or T0 plants are designated T1 seeds.

3.2.7.6 Obtaining homozygous lines for the transgenes

T0 progeny (T1 generation) were sown on soil for Basta selection. Selection was done on emerging seedlings by spraying 200 mg/l Basta once every 3 days for approximately 2 weeks.

Individual resistant seedlings of primary transformants (T1 plants), which expressed the Basta resistance gene were grown to maturity and their progeny (T2) were collected.

Twenty T2 seeds were spread on half MS agar. Genomic DNA was extracted from leaves

(§2.2.1), and PCR was conducted to detect the presence of transgene (§2.7.1). The ones with the expected PCR band were transferred to soil and seeds were harvested (T3 seeds). Four individual T3 lines for each construct were further selected on half MS agar containing 5 mg/l

BASTA to select for T3 lines that were homozygous for the transgenes. At least 40 seeds from each T3 lines were placed onto Basta selection plate. Col-0 seeds were placed on the same plate acting as the negative control.

3.2.7.7 Performing the complementation assay

Between 120-150 seeds were assayed for each homozygous T3 line from the two independent

T1 lines (designated C1 and C2). At least 100 seeds were placed on half MS without sucrose, while no less than 20 seeds were sown on half MS with 1% (w/v) sucrose (positive control plate). Col-0 was used as the control for both treatments. Plates were incubated as described in

Section 3.2.7.1. Seedlings that grew normally on half MS without sucrose were scored 14 days after incubation. Three independent assays were performed.

3.3 Results

3.3.1 Isolation of GmABCA1 and GmABCA2 CDS

Two integral proteins with homology to ABC transporter subfamily A were identified in the

SM proteome of G. max (Clarke et al., 2015). Although Clarke et al. (2015) referred to these proteins as GmABCA7 and GmABCA2, I use the unified nomenclature proposed by Verrier et al. (2008) and refer to them as GmABCA1 and GmABCA2, respectively. Based on CDS predictions in the soybean reference genome Glyma1.1, full-length CDS of GmABCA1 was called Glyma04g34130.1 in version 1.1 of the soybean genome but is now named

Glyma04G172900.1 in genome assembly Glyma2.0 and GmABCA2 was Glyma04g34140.1 but is now named Glyma04G17300.1.

3.3.1.1 GmABCA1

The cDNA cloned for GmABCA1 by Clarke (2013) was not complete and did not encode a complete ABC protein as it did not contain exons 12 through 15 based on the soybean genome prediction in version 1.1. The protein encoded by the truncated GmABCA1 cDNA did not include four amino acid regions identified in the SM proteome of G. max (Clarke, 2013). In this study, a full-length 2850 nucleotide ORF of GmABCA1 encoding all amino acid regions identified on the SM was successfully amplified and cloned after multiple attempts. The attB- flanked product was first ligated into pGEM®-T Easy vector and subsequently cloned into pDONR™221 as the product could not be cloned directly into pDONR™221. The isolated full- length GmABCA1 CDS is structured in 18 exons based on the prediction in the reference genome version 1.1 and had the same nucleotide sequence as predicted, except with one nucleotide different located in exon 15 (Figure 3.5A). However, this difference did not change the predicted amino acid sequence.

The GmABCA1 CDS isolated in this study encodes a putative protein of 950 amino acids with a calculated molecular mass of 106.6 kDa. BLASTP analysis identified conserved regions including one TMD and all six conserved motifs of an NBD in the forward orientation and revealed that the translated CDS encodes an ABC subfamily A transporter (Figure 3.6A). As there is only a single transmembrane domain and NBD the encoded protein is a half-size transporter. The translated CDS also contains all six amino acid sequences identified in the SM proteome (Clarke et al., 2015; Figure 3.6A).

Figure 3.5 Representation of GmABCA1 and GmABCA2 coding sequences that were isolated and cloned in this study. (A) The 2850 bp amplified GmABCA1 CDS had a nucleotide different in exon 15, as indicated. This however did not change the amino acid sequence. (B) The 2889 bp amplified GmABCA2 CDS had two nucleotides (in exon 1 and 2) different compared to sequence in genome prediction. Phenylalanines were produced instead of leucines in the predicted amino acid sequence. Image generated using the SnapGene® software.

A GmABCA1 version 1.1 1 950

B GmABCA1 version 2.0 1 729 a.a

C GmABCA2 version 1.1 and version 2.0

1 963

Figure 3.6 Representation of primary protein structure of GmABCA1 and GmABCA2. (A) Glyma04g34130.1 protein based on sequence prediction in the earlier genome released version 1.1 (isolated in this study). (B) Glyma04G172900.1 protein in the updated genome assembly version 2.0. (C) Glyma04g34140.1 (version 1.1) and Glyma04G17300.1 (version 2.0) have the same amino acid sequence. The six conserved domains are represented by coloured regions (Walker A: brown, Q-loop: red, Signature: orange, Walker B: blue, D-loop: green, Switch/H-loop: violet), determined by BLASTP tool. The triangles indicate amino acid regions identified in the SM proteome (Clarke et al., 2015).

Since completing this experiment, an updated version of the genome (version 2.0) was released.

The predicted amino acid sequence of the new gene model, Glyma.04G172900.1 is shorter (729 residues) in version 2.0 and BLASTP analysis showed that the protein is missing Walker A and

Q-loop motifs (Figure 3.6B). An amino acid region identified in the SM proteome (Clarke et al., 2015) is also missing (Figure 3.6B). Based on my cloning results and the amino acid data from the SM proteome (Clarke et al., 2015) it seems likely that the gene model in genome assembly version 2.0 incorrectly predicts the GmABCA1 CDS. In addition, BLASTP of

Genbank identified a gene model (KHN45210.1) almost the same as the translated cDNA cloned in this study, except that the encoded protein contains an extra amino acid (valine) inserted after amino acid 785.

3.3.1.2 GmABCA2

The GmABCA2 CDS of 2889 bp isolated in this study had the same sequence as previously cloned CDS (Clarke, 2013). Two nucleotides were different from the predicted CDS in genome assembly version 1.1 (Figure 3.5B). Nucleotide bases at position 253 bp (in exon 1) and 539 bp

(in exon 2) were T, instead of C, resulting in phenylalanines instead of leucines in the amino acid sequence. PCR amplification was carried out twice and the sequencing peaks of those nucleotides were well formed with good quality scores, demonstrating that the differences in nucleotides did not stem from PCR error or sequencing error. This indicated that the base differences are due to SNPs between sequences in G. max cv. Stephen and the reference genome

G. max cv. Williams 82.

The isolated GmABCA2 CDS is divided into 16 exons based on genome Glyma1.1, and encodes a putative protein of 962 amino acids with a theoretical molecular mass of 106.9 kDa. BLASTP analysis revealed that the translated CDS encodes a half-size member of an ABCA transporter that encompassed all six conserved domains of an NBD and a single transmembrane domain in a forward topology (Figure 3.6C). The translated CDS also contained all three amino acid regions identified in the SM proteome (Figure 3.6C). The protein in the updated genome assembly version 2.0 (Glyma04G17300.1) had the same amino acid residues for GmABCA2 as in version 1.1 (Glyma04g34140.1).

GmABCA1 and GmABCA2 have an amino acid identity of 24.4% and a similarity of 38.7%.

These genes are located next to each other on chromosome 4. GmABCA1´s closest homologue in soybean is Glyma06g20370 (now named Glyma.06G191400 in Glyma2.0) with 80.7% protein identity while GmABCA2´s closest homologue in soybean is Glyma06g20360 (now named Glyma.06G191300 in Glyma2.0) sharing 91.4% protein identity.

3.3.2 Analysis of predicted protein structure

Four membrane topology programs (HMMTOP, Phobius, SOSUI and TMHMM) were used to assess if GmABCA1 and GmABCA2 are integral membrane proteins, as they were identified in the SM fraction in the proteomic analysis (Clarke et al., 2015). All four programs detected transmembrane helices (TMHs) within both GmABCA1 and GmABCA2 and thus it can be concluded that both proteins are likely integral membrane proteins. Two programs HMMTOP and TMHMM predicted GmABCA1 to contain seven TMHs, while Phobius and SOSUI predicted eight and six TMHs, respectively. No signal peptide was predicted in GmABCA1 by

SignalP and Phobius. The secondary protein structure representation of GmABCA1 is shown in Figure 3.7.

GmABCA2 is predicted to contain six TMHs by three programs (Phobius, HMMTOP and

TMHMM), while SOSUI predicted seven TMHs. The first 17 amino acid residues are predicted to be a signal peptide by Phobius while SignalP detected no signal peptide in GmABCA2.

GmABCA2´s secondary protein structure representation is shown in Figure 3.8. An NBD sequence alignment of GmABCA1 and GmBCA2 with the previously characterized AtABCA9

(Kim et al., 2013) demonstrated that there is a high degree of conservation of the NBD structure within the ABC proteins (Figure 3.9).

3.3.3 Phylogenetic analysis

The phylogenetic relationships of GmABCA1 and GmABCA2 in relation to other ABCA members in G. max and A. thaliana were analyzed by inferring sequence distances between them. A maximum likelihood tree was constructed and three distinct clades were identified, supported by good bootstrap values (Figure 3.10). GmABCA1 and GmABCA2 clustered in different clades, with GmABCA2 clustering closer to the characterized ER-localized fatty acid transporter AtABCA9 (Kim et al., 2013), located in Clade II (Figure 3.10). GmABCA2 and

AtABCA9 have an amino acid identity of 62.9%, whereas GmABCA1 and AtABCA9 shared an amino acid identity of 24.3%. Within the ABCA family, GmABCA1 is most closely related to AtABCA7 (At3g47780.1) showing the shortest sequence distance (Figure 3.10), and shared

65.9% protein identity. GmABCA2’s closest structural homologue in A. thaliana is AtABCA2

(At3g47730.1) (Figure 3.10), sharing 64.9% protein identity. None of the predicted proteins

1 2 3 4 5 6 7 8 cytoplasm

Signal peptide Walker A Q-loop ABC signature motif Walker B D-loop Switch region/H-loop Peptides identified in SM proteome

Figure 3.7 Representation of secondary protein structure of GmABCA1 isolated in this study. GmABCA1 is a half-size transporter that can form homodimers or heterodimers, and is present in the forward orientation (TMD-NBD). The NBD is located in the cytosol, at the carboxyl end of the protein. The image was generated using the Protter program (Omasits et al., 2013) with the default setting: Phobius (Käll et al., 2004) to predict TMHs and signal peptide. The TMHs are labeled by red numbers. GmABCA1 is predicted to contain eight TMHs with no signal peptide. The six NBD domains, as determined by BLAST alignment with other known ABC proteins, are shown in color as detailed above. The light blue region indicates amino acid regions identified by LCMS/MS in the SM proteome (Clarke et al., 2015).

1 2 3 4 5 6 cytoplasm

Signal peptide Walker A Q-loop ABC signature motif Walker B D-loop Switch region/H-loop Peptides identified in SM proteome

Figure 3.8 Secondary protein structure representation of GmABCA2 isolated in this study. GmABCA2 is also a half-size transporter, with the forward topology (TMD-NBD). The image was generated using the Protter program (Omasits et al., 2013) using the default setting: Phobius (Käll et al., 2004) to predict TMHs and signal peptide. The TMHs are labeled by red numbers. Phobius predicted GmABCA2 to contain six TMHs and a signal peptide in the first 17 amino acids. The six NBD domains, as determined by BLAST alignment with other known ABC proteins, are shown in color as outlined above. The light blue region denotes amino acid regions identified by LCMS/MS in the SM proteome (Clarke et al., 2015).

Walker A GmABCA1 MRKVYPGRDGN------PEKLAVRGLSLALPQGECFGMLGPNGAGKTSFINMMIGLT 682 GmABCA2 LAKTYPGTRSIGCCFKCKRTSPYNAVKGLWVNFAKDQLFCLLGPNGAGKTTAINCLAGIT 577 AtABCA9 LAKTYPGTTKLGCC-KCTKTSPFHAVKGLWMNIAKDQLFCLLGPNGAGKTTTISCLTGIN 583 : *.*** **:** : : : : * :*********: *. : *:.

Q-loop GmABCA1 KPTSGTAYVQGLDLRT--HMDGIYTSMGVCPQHDLLWESLTGREHLLFYGRLKNLKGSAL 740 GmABCA2 PVTDGDALIYGHSIRSSSGLSNIQKLIGVCPQFDILWDALSGQEHLQLFATIKGLSPSSI 637 AtABCA9 PVTGGDAKIYGNSIRSSVGMSNIRKMIGVCPQFDILWDALSSEEHLHLFASIKGLPPSSI 643 * * * : * .:*: :. * . :*****.*:**::*:..*** ::. :* * *::

Signature Walker B D-loop GmABCA1 TQAVEESLKSVNLFHGGVADKQAGKYSGGMKRRLSVAISLIGDPKVVYMDEPSTGLDPAS 800 GmABCA2 KSITQTSLAEVRL--TDASKVRAGSYSGGMKRRLSFAIALIGDPKLVILDEPTTGMDPII 695 AtABCA9 KSIAEKLLVDVKL--TGSAKIRAGSYSGGMKRRLSVAIALIGDPKLVFLDEPTTGMDPIT 701 .. .: * .*.* :. :**.**********.**:******:* :***:**:**

Switch region GmABCA1 RKNLWNVVKRAKQDRAIILTTHSMEEAEVLCDRLGIFVDGGLQCIGNPKELKARYGGTYV 860 GmABCA2 RRHVWDIIENAKRGRAIVLTTHSMEEADILSDRIGIMAKGSLRCIGTSIRLKSRFGAGFI 755 AtABCA9 RRHVWDIIQESKKGRAIILTTHSMEEADILSDRIGIMAKGRLRCIGTSIRLKSRFGTGFV 761 *:.:*::::.:*: ***:*********::*.**:**:..* *:***. .**:*:* ::

Figure 3.9 Sequence alignment of nucleotide binding domain (NBD) of GmABCA1 and GmABCA2 with the characterized AtABCA9. The NBD consists of six motifs (indicated by shading) that are characteristic of an ABC transporter. The ATP binding sites are in solid red. An asterisk (*) indicates the position of a fully conserved residue, a colon (:) indicates the position of amino acids with strongly similar properties, and a period (.) denotes the position of amino acids with weakly similar properties. The alignment was produced using Clustal Omega aligner (Sievers et al., 2011) and the motifs were identified by BLASTP.

At3g47790.1 100 At5g61740.1 100 At5g61700.1 100 At3g47780.1 At3g47740.1 94 At3g47770.1 100 Clade I 99 At3g47750.1 44 88 At3g47760.1 100 Glyma17g10670.1 Glyma05g01230.1 98 Glyma04g34130.1 (GmABCA1) 99 Glyma06g20370.2 99 Glyma04g34140.1 (GmABCA2) 100 Glyma06g20360.1 At3g47730.1 (AtABCA2) Clade II

98 At5g61730.1 (AtABCA9) 100 100 At5g61690.1 Glyma03g29230.1 Clade III 100 At2g41700.1

0.2

Figure 3.10 Phylogenetic analysis of G. max and Arabidopsis ABC subfamily A proteins. The tree was constructed using MUSCLE (Edgar, 2004) alignment of 19 amino acid sequences: seven G. max proteins and twelve Arabidopsis proteins. The evolutionary relationships were inferred using the Maximum Likelihood method based on the JTT model (Jones et al., 1992). The tree is drawn to scale, in which the branch lengths represent the number of amino acid substitutions per site. The bootstrap (1000 replicates) values represented as percentages of which the associated taxa are clustered together are indicated at the corresponding nodes. GmABCA1 and GmABCA2 were highlighted in the green boxes. Analysis was conducted in MEGA6 (Tamura et al., 2013).

belonging to Clade I and III have been characterized so far. Clade III contained representatives of the only full-size member of the ABCA transporter family in both plants: At2g41700.1 and

Glyma03g29230.1.

3.3.4 Gene expression analysis

The expression levels of GmABCA1 and GmABCA2 in soybean tissues (leaf, stem, root and nodule) were analyzed in this study to confirm expression profiles of both transcripts from the two soybean transcriptomes (Libault et al., 2010; Severin et al., 2010). This assay was previously performed by Clarke (2013), but had to be repeated as GmABCA1 and GmABCA2 paralogs (Glyma06g20370, now Glyma.06G191400 and Glyma06g20360, now

Glyma.06G191300 in Glyma2.0, respectively) might be amplified due to the nonspecific primer sets used originally. In this study, paralogs were distinguished by designing primers with at least five nucleotide mismatches between the paralogs, as opposed to only one or two mismatches in the previous primer sets. Where possible, regions of dissimilarity between the paralogs were chosen. Post-run melting curve analysis confirmed that the primers used are specific for the genes (Appendix 2).

Among all the ABCA members of G. max, only GmABCA1 and GmABCA2 showed nodule- enhanced expression based on the two soybean transcriptome studies (Libault et al., 2010;

Severin et al., 2010; Table 3.5). Glyma.06G191300 (the GmABCA2 paralog) has a more diverse expression pattern across different tissues, where it is mostly expressed in leaf. Whereas

Glyma.06G191400 (the GmABCA1 paralog) is also expressed in all tissues except in flower, but its overall expression is considerably lower than Glyma.06G191300 (Table 3.5).

Table 3.5 Tissue-specific gene expression profiles of G. max ABCA genes, compiled from the two transcriptome studies: Libault et al. (2010) and Severin et al. (2010). Normalized expression data across five different soybean tissues: leaf (L), flower (F), pod (P), root (R) and nodule (N) were presented and is reads by the number of sequence reads/million reads aligned. PROTEIN NAME SYSTEMATIC Libault et al. (2010) Severin et al. (2010) (V1.1/ V2.0) NAME L F P R N L F P R N Glyma04g34130/ GmABCA1 0 1 1 3 138 0 0 0 1 205 Glyma.04G172900 Glyma04g34140/ GmABCA2 1 1 1 1 35 0 0 0 0 52 Glyma.04G173000 Glyma03g29230/ 9 46 9 11 14 1 10 3 5 6 Glyma.03G136000 Glyma05g01230/ 7 9 10 2 3 3 2 3 1 1 Glyma.05G028500 Glyma06g20360/ 16 32 18 47 11 1 7 5 16 8 Glyma.06G191300 Glyma06g20370/ 1 0 4 8 1 0 0 0 2 1 Glyma.06g191400 Glyma17g10670/ 1 2 10 1 2 1 2 3 0 1 Glyma.17G098300

Using the new primer sets, an RT-qPCR assay for GmABCA1 and GmABCA2 was conducted.

Across the four soybean tissues tested, both GmABCA1 and GmABCA2 showed enhanced expression in the nodule tissue, and very low expression in other tissues (Figure 3.11). The nodule expression level of GmABCA1 was 2.1-fold higher than GmABCA2. In contrast, Clarke

(2013), using less specific primers for RT-qPCR, showed expression of GmABCA2 in other tissues, although expression in nodules was much higher (Appendix 3).

The expression level of both transcripts was investigated further in tissue from roots (6 DPI), roots with nodule initials (11 DPI) and nodules throughout nodule development (14-42 DPI).

GmABCA1 and GmABCA2 exhibited a similar expression pattern, with higher expression levels observed for GmABCA1 at all time points (Figure 3.11). The largest difference in expression levels between the two transcripts was detected at 11 DPI, with 8.2-fold lower expression of

GmABCA2. The GmABCA1 expression level peaked at 14 DPI and decreased henceforth, while

GmABCA2 peaked later at 19 DPI during nodule development.

3.3.5 Spatial localization of expression

As both GmABCA1 and GmABCA2 have nodule-enhanced expression, the cell types in which

GmABCA1 and GmABCA2 are active within the nodule tissue were investigated. A construct with the GmABCA2 promoter driving expression of a GUS/GFP fusion was transformed into nodules via A. rhizogenes-mediated hairy root transformation. No GUS activity was detected in either the infected or the uninfected cells (Appendix 4).

In situ RT-PCR was undertaken as an alternative method to the GUS/GFP-promoter fusion

2.5E-02 GmABCA1 02 -

GmABCA2 2.36E

2.0E-02 02

1.5E-02 - 1.11E 1.0E-02

RELATIVE RELATIVE EXPRESSION 5.0E-03

1.16E

7.43E

5.14E

3.37E

3.20E 2.38E 0.0E+00 LEAF STEM ROOT NODULE TISSUE

4.5E-02 GmABCA1 4.0E-02 GmABCA2 3.5E-02 3.0E-02 2.5E-02 2.0E-02 1.5E-02 1.0E-02

RELATIVE RELATIVE EXPRESSION 5.0E-03 0.0E+00 0 5 10 15 20 25 30 35 40 45 DAYS POST INOCULATION (DPI)

Figure 3.11 Relative expression levels of GmABCA1 and GmABCA2 in (A) four different soybean tissues at 32 DPI, (B) and across six stages of nodule development (6, 11, 14, 19, 32 and 42 DPI). Tissue was taken from roots at 6 DPI as nodules have not yet formed. GmUBI was used to normalize expression levels of both genes. Results were represented as means of expression levels between two biological replicates.

because the GmABCA1 promoter was unable to be cloned. The transcripts were reverse transcribed and amplified directly on the nodule sections, and PCR products were then detected using colorimetric detection (Athman et al., 2014). The expression of the GmUBI positive control was seen in all cell types (Figure 3.12A). Another positive control, GmYSL7, was specifically expressed in the infected cells (Figure 3.12B), in agreement with previous GUS assay results (Chen, 2012; Appendix 5). Both GmABCA1 and GmABCA2 were strongly expressed in the infected cells, but transcripts were not detected in the uninfected cells (Figure

3.12C & D). The observed expression of GmABCA1 and GmABCA2 was similar to the GmYSL7 infected cell control. Negative control with no reverse transcriptase was not included and thus this experiment would need to be repeated to confirm the result.

3.3.6 Protein localization

3.3.6.1 GmABCA1

A construct with an N-terminal GFP fusion under the control of the GmLbC3 promoter was initially prepared from the amplified GmABCA1 and was expressed a number of times in soybean nodules, but no GFP fluorescence was observed. The A. rhizogenes transformation of roots relies on having constructs that include a visible marker such as DsRed. No such construct was available to express the candidate genes with a C-terminal GFP fusion in infected cells of nodules (the 35S promoter is not expressed in nodule infected cells (Chen, 2012)). For this reason, an attempt to transiently express GmABCA1 with a C-terminal GFP tag driven by the

35S promoter in Allium porrum L. (leek) was undertaken in this study. Confocal microcopy of bombarded leek epidermal cells showed fluorescence of GFP-tagged GmABCA1 that appeared

A GmUBI VB C GmABCA1

IC MC ICs OC

B GmYSL7 D GmABCA2 UC

ICs IC UC

Figure 3.12 In situ RT-PCR in cross-sectioned soybean nodules of 32 DPI. Reverse transcription and in situ PCR were carried out directly on nodule tissue using gene-specific primers. PCR products were detected using colourimetric detection (Athman et al., 2014). Dark purple staining using BM purple (Roche) shows the presence of the cDNA. (A, B) Positive controls of the in situ PCR. GmUBI is expressed in all cell types. GmYSL7 localized to the infected cells (ICs), consistent with GUS results (Appendix 5). (C, D) GmABCA1 and GmABCA2 are expressed specifically in the infected cells. Magnified images are shown on the left. Abbreviations: UC – Uninfected cells, IC – inner cortex, MC – middle cortex, OC – outer cortex, VB – vascular bundle. Scale bars represent 250 µm.

diffuse in the cytoplasm and nucleus (Figure 3.13, left panel). The mCherry-PM marker was most probably in the ER and nuclear envelope (Figure 3.13, middle panel).

3.3.6.2 GmABCA2

A construct with an N-terminal GFP fusion was prepared from the amplified GmABCA2 CDS.

Subcellular localization of GmABCA2-GFP under the control of GmLbc3 promoter was investigated in mature transgenic nodules at 23 DPI. Confocal microscopy of transgenic nodules expressing ProGmLbc3:GFP-GmABCA2 showed that GFP-tagged GmABCA2 formed ring-like structures, typical of symbiosome silhouettes within infected nodule cells (Figure

3.14, left panel). The pattern is consistent with the symbiosome membrane which forms the major internal membranes in the infected cell and that the pattern is not consistent with other organelles in infected cells such as ER, Golgi, vacuole, plastid or mitochondria. Localization of GmABCA2 on the SM was confirmed by counterstaining of the cell membranes by FM®4-

64, which showed a similar pattern to the GFP labeling (Figure 3.14, middle panel). The SM labelling is also similar to that seen for other proteins showing SM localization (Appendix 6).

Pixel intensity profiles of both fluorophores demonstrated that the two probes overlapped proportionally at the SM and did not overlap at the PM (Figure 3.15A-C), indicating

GmABCA2 specific localization to the SM.

3.3.7 Yeast complementation

3.3.7.1 fet3fet4ftr complementation

GmABCA1 mCherry Overlay A

Figure 3.13 Cellular localization of GmABCA1 under the control of 35S promoter in the epidermal cells of Allium porrum L. (A) to (C) Confocal images of leek epidermal cells expressing C-terminal GFP fusion of GmABCA1. The left panel shows GFP-tagged GmABCA1, the middle panel shows mCherry-tagged PM stain, and the right panel shows the overlay of GFP and mCherry channels. Open arrowhead indicates the cytoplasm; closed arrowhead indicates the cytoplasmic strand, double arrowhead indicates nucleus envelope and an arrow indicates the nucleus. Scale bars represent 25 µm.

GmABCA2 FM®4-64 Overlay A IC

IC IC

B IC

Figure 3.14 Subcellular localization of GmABCA2 driven by GmLbc3 promoter in the 23 DPI infected nodule cells of G. max. (A) to (C) Confocal images of transgenic nodules expressing N-terminal GFP fusion of GmABCA2. The left panel shows GFP-tagged GmABCA2 within the infected cells (IC), the middle panel shows FM®4-64 counterstaining the SM and PM, and the right panel shows the merged of GFP and FM®4-64 channels. Closed arrowhead indicates the SM, and open arrowhead indicates the PM. Scale bars represent 10 µm.

Overlay ROI 1 ROI 2 A 30 SM 60 PM ROI 1 SM 0 µm 2.7 µm 20 40 SM 0 µm 10 20

ROI 2 2.9 µm

Gray value Gray value

0 0 0.0 1.0 2.0 3.0 0.0 1.0 2.0 3.0 Distance (µm) Distance (µm)

FM4-64 GFP FM4-64 GFP

B 0 µm 60 PM 50 SM ROI 2 40 2.2 µm SM 40 SM 30 20

20 Gray value Gray value 10 0 0 0 µm 0.0 0.8 1.6 2.4 3.2 0.0 0.5 1.0 1.5 2.0 2.5

ROI 1 3.2 µm Distance (µm) Distance (µm)

FM4-64 GFP FM4-64 GFP

C 25 80 PM SM 20 SM 60 1.9 µm 15 0 µm 40 ROI 1 10

Gray value 20 0 µm 1.4 µm Gray value 5 ROI 2 0 0 0.0 1.0 2.0 0.0 0.5 1.0 1.5 Distance (µm) Distance (µm)

FM4-64 GFP FM4-64 GFP

Figure 3.15 Intensity profile graphs of GPF-GmABA2 and FM®4-64 in the region of interests (ROI), across SM and PM. Superimposed images of both probes were shown in the left panel. Fluorescence colocalization of the two fluorescent labels were analyzed and evaluated quantitatively by plotting pixel intensities of each color against the ROI (beginning at 0 µm). The two probes overlapped relatively proportionally with one another at the SM, but not at the PM, indicating GmABCA2 specific localization to the SM. Scale bars represent 10 µm.

The yeast fet3fet4 mutant is defective in both low-affinity (FET4) and high-affinity iron uptake system (FET3) (Askwith et al., 1994; Dix et al., 1994). fet3fet4 grows poorly on iron limited media and needs to be supplemented with iron for growth. A previous pilot study by Clarke

(2013) complementing fet3fet4 (DEY1453) with GmABCA1 (truncated) and GmABCA2 on a iron limited plate resulted in improved growth of both constructs when compared to the negative control, but no positive control was included in the experiment.

In this study, full-length GmABCA1 and GmABCA2 were expressed in yeast fet3fet4ftr1

(DEY1530) which has an additional mutation in the FTR1 gene, encoding a transporter protein that forms a complex with FET3 for high-affinity iron uptake (Spizzo et al., 1997). The

Arabidopsis iron-regulated transporter 1 (AtIRT1, Eide et al., 1996) positive control restored growth in the mutant yeast grown on iron-limiting media (0 and 2 µM FeSO4) but expression of GmABCA1 and GmABCA2 did not (Figure 3.16). This failure to complement DEY1530 suggests neither of the proteins encode by GmABCA1 and GmABCA2 could transport iron.

3.3.7.2 Δfat1 complementation

The ability of GmABCA1 and GmABCA2 to transport fatty acids (FAs) was investigated through complementation of a fatty acid uptake-deficient yeast mutant, Δfat1 (Færgeman et al.,

1997). Attempts to complement the Δfat1 mutant with GmABCA1 and GmABCA2 full-length using a formerly developed fatty acid assay (Færgeman et al., 1997; Johnson et al., 1994) were undertaken. However, no differences in growth were observed for all constructs including the positive control (ScFAT1) and negative control (empty vector) when plated on SD

A 0 µM FeSO4 C 20 µM FeSO4

AtIRT1 Empty

vector

GmABCA1 GmABCA2

B 2 µM FeSO4

Figure 3.16 Complementation of iron-deficient yeast mutant strain, fet3fet4ftr1 with GmABCA1 and GmABCA2 CDS. Full-length CDS of AtIRT1, GmABCA1 and GmABCA2 in pDR196gw along with the empty vector were transformed into DEY1530. Yeasts harboring the plasmids were streaked onto iron-free media supplemented with (A) 0, (B) 2 and (C) 20 µM

FeSO4 and grown for 3 days at 30°C. Photos were taken 5 days after plating.

supplemented with 25 µM cerulenin and 50 µM oleic acid (data not shown). A possible reason for the lack of noticeable growth differences for all constructs is that yeast growth is rapid in exponentially growing cells, where cell densities increased to a sufficient level to overcome growth inhibition of the antibiotic cerulenin. Another reason could be due to unknown yeast intrinsic transporters other than Fat1p that may function by mediating fatty acid import into yeast cells or though passive diffusion.

A recent paper by Li et al. (2015a) performed a similar assay but used non-exponentially and exponentially growing yeast for the complementation assay. Growth promotion was observed for Δfat1-AtFAX1 when using a slow growing yeast culture, possibly because strong cerulenin inhibition pushed the free FAs transported by FAX1 into the FA-metabolism pathways (Li et al., 2015a). In a rapidly growing yeast culture, growth inhibition of Δfat1-AtFAX1 was observed probably because the FAs transported by AtFAX1 cannot be efficiently esterified to

CoA as AtFAX1 could not interact with the endogenous yeast acyl-CoA synthetases (Li et al.,

2015a).

An attempt to complement the Δfat1 mutant based on protocols described in Li et al. (2015a) using non-exponential phase yeast cells was undertaken in a growth curve assay. GmABCA2 transported oleic acid almost as well as positive control ScFAT1 when compared to the empty vector (GmABCA2 with 15.2-fold and ScFAT1 with 15.4-fold growth rates, Figure 3.17).

GmABCA1 transported oleic acid slower than ScFAT1 (GmABCA1 growth rate was 12.7-fold higher when compared to the empty vector, Figure 3.17).

1.2 ScFAT1 pDR196 1.0 GmABCA1 15.4 0.8 GmABCA2 15.2 600 0.6

OD 12.7 0.4

0.2

0.0 0 25 50 75 100 125 150 TIME (H)

Figure 3.17 Complementation of fatty acid transporter (FAT1p) deletion mutant (Δfat1) with GmABCA1 and GmABCA2 CDS in a growth curve assay. The mature ScFAT1 in pRS416 (a positive control obtained from Euroscarf), GmABCA1 and GmABCA2 full-length CDS in pDR196gw and the empty vector, pDR196, were introduced into Δfat1 yeast mutant. The assay was performed according to Li et al. (2015a) by measuring fat1 cells optical density (OD) at 600 nm in liquid SD minus uracil (2% (w/v) glucose, 0.5% (w/v) Brij 58, 0.7%

(w/v) KH2PO4) containing 100 µM oleate and 10 µM cerulenin, at the indicated time points.

Cells were harvested from non-exponentially growing cultures, diluted to an OD600 of 0.03 and were grown at 30°C for 143 h. Growth promotion was observed for the positive control ScFAT1 (15.4-fold), GmABCA2 (15.2-fold) and GmABCA1 (12.7-fold) in comparison to the empty vector.

3.3.8 Atabca9 complementation

3.3.8.1 Identification of homozygous abca9 mutant plants

Salk 058070 (abca9-1) obtained from Arabidopsis Biological Resource Center (ABRC) carries a T-DNA insertion in exon 9 of AtABCA9 gene (Figure 3.18A) that encodes an ER localized fatty-acid transporter. Ten segregating T3 or T4 seedlings were PCR genotyped to identify homozygous individuals. All plants tested were homozygous for the T-DNA insertion as demonstrated by the presence of an amplification product corresponding to the T-DNA insertion and the absence of a wild type amplification product (Figure 3.18B). The T-DNA insertion product amplified using T-DNA and gene-specific primers (LB + RP) yielded an expected product size of around 600 bp. Reactions using Col-0 as a control template produced a wild type product of 1180 bp when using gene-specific primers (LP + RP) and no band for

LB + RP reaction as expected.

3.3.8.2 Complementation of abca9-1 mutant

Kim et al. (2013) previously described the abca9-1 mutant phenotype, displaying impaired seedling establishment on half MS without sucrose. abca9-1 seeds contain lower TAG content when compared to the WT. Early seedling growth of oilseed plants depends on storage lipids in the absence of sucrose, and thus media needs to be supplemented with sucrose for normal seedling growth of abca9-1 (Kim et al., 2013). Genetic complementation of the abca9-1 T-

DNA insertion mutation with the wild type gene driven by its native promoter rescued the mutant phenotype on half MS without the need for sucrose (Kim et al., 2013).

A LBb1.3 T-DNA RB LB LP RP

ATG At5g61730 gDNA Stop

Col-0 Salk 058070 B LP + ‒ + ‒ RP + + + + LB ‒ + ‒ +

1.5 kb — 1.2 kb —

700 bp — 600 bp —

Figure 3.18 (A) T-DNA insertion location in Salk 058070 (abca9-1) mutant line. The T- DNA is integrated in exon 9 at position 2478 bp relative to the start codon of At5g61730. (B) Genotyping of Salk 058070 to identify homozygous individuals for the insertion. A combination of gene-specific primers (LP + RP), as indicated in (A), were used to characterize WT allele while T-DNA and gene-specific primer (LBb1.3 + RP) were used to characterize T- DNA insertion. Only one plant genotyping was shown here and was found to be homozygous for the insertion. Five µl PCR products were loaded on 1% TAE gel. NEB 2-log ladder was used as the standards for size determination and quantification of PCR products. Abbreviations: LP – left primer, RP – right primer, LB – left border and RB – right border.

In this study, an attempt to complement the abca9-1 mutant background (Salk 058070) with the full-length CDS of GmABCA1 and GmABCA2 under the control of the AtABCA9 native promoter was undertaken based on the assay described above. Two independent lines were generated for each soybean gene. The AtABCA9 full-length gDNA, including the 2000 bp upstream of the start codon, was expressed in another two lines as a positive control. Each T3 independent line used for the assay was homozygous for the transgene, as evidenced by resistance of all seedlings tested on a Basta selection plate (Appendix 7).

Complementation of AtabcA9 (Salk 058070) with GmABCA1 and GmABCA2 restored seedling establishment on half MS without sucrose to varying degrees (Figure 3.19 and 3.20).

The percentage of seedlings that grew normally on half MS without sucrose was calculated and results were represented as a mean of percentages from three trials minus EV negative control

(Figure 3.20). AtABCA9 C2 had the highest mean percentage of phenotype recovery of 30%, followed by GmABCA2 C2 with 28% and GmABCA1 C1 with 21% (Figure 3.20). A student’s t-test performed to compare growth of the complementation lines to that of the empty vector control showed that the three complementation lines AtABCA9 C2, GmABCA1 C1 and

GmABCA2 C2 had significantly different mean percentages of seedling recovery on half MS without sucrose (Figure 3.20).

3.4 Discussion

In this study, two ABC subfamily A transporters, GmABCA1 and GmABCA2 that have enhanced expression within the nodule have been characterized as fatty acid transporters. Both

1% sucrose No sucrose 1% sucrose No sucrose A C

AtABCA9 WT C1 WT C1 WT C1 WT C1

GmABCA1

C2 Positive control control Positive

WT C2 WT C2 WT C2 WT C2

B D

abca9

WT abca9-1 WT abca9-1 WT C1 WT C1

GmABCA2

ctor

Negative controls Negative

C2 Empty ve Empty WT EV WT EV WT C2 WT C2

Figure 3.19 Representatives of abca9 complementation plates. Wild type (WT) Col-0 and AtABCA9 full-length gDNA fused to its native promoter were used as the positive control while lines transformed with an empty vector (EV) and untransformed (UN) Salk 058070 lines were used as the negative controls. The controls along with GmABCA1 and GmABCA2 CDS under the control of the AtABCA9 native promoter were expressed in Salk 058070. Two independent complementation lines were generated, represented by C1 and C2. Seeds were grown on half MS media with and without sucrose and seedling growth was monitored for 14 days.

60 *

50 54 * 40 * 30 * 30 28 20 21 17 17 10 13 0 3

0 % seedling germination and establishment germination % seedling

GENOTYPE

Figure 3.20 Complementation of GmABCA1 and GmABCA2 in abca9-1 mutant background (Salk 058070). The percentage of seedlings able to germinate and grow normally on half MS media without sucrose were calculated. The mean of percentages from three trials minus EV control germination rate are shown above. SD of the mean represented by the error bar. Student’s t-test was performed between complementation line and empty vector control. An asterisk (*) indicates that the difference between the two lines is significant (P-value <0.05). Abbreviations: EV – empty vector, UN – Unstransformed Salk 058070.

appear to be expressed specifically in the infected nodule cells. The localization of GmABCA2 on the SM has been confirmed. These results suggest that both proteins play an important role in the interaction between the symbionts. Although how fatty acid transport across the SM is important has not been established, the present study describes for the first time fatty acid transport activity across the SM through these transport proteins.

3.4.1 GmABCA1 and GmABCA2 have nodule enhanced expression

Both GmABCA1 and GmABCA2 were highly expressed within the nodule tissue, but have barely detectable expression in other tissues (Figure 3.11A). These results were broadly in agreement with the two soybean transcriptome profiles (Libault et al., 2010; Severin et al.,

2010; Table 3.5). The higher nodule expression of GmABCA1 in comparison to GmABCA2 also coincided with the data from the two soybean transcriptomes (Libault et al., 2010; Severin et al., 2010; Table 3.5). GmABCA1 was expressed 2.1-fold higher than GmABCA2 in nodule tissue in the present study, whereas Libault et al. (2010) and Severin et al. (2010) detected 3.6- fold and 3.9-fold higher expression, respectively. The variability of the expression data was possibly due to differences in soybean cultivar used, growth condition, age of the samples at time of harvesting, and the RT-qPCR assay. In soybean, the nodule possessed the largest number of tissue-specific genes (Libault et al., 2010). This might reflect functional specialization of nodule tissue, where GmABCA1 and GmABCA2 were among those highly upregulated genes detected.

Most soybean genes have duplicated copies due to the two duplication events in the soybean genome (Roulin et al., 2013). Approximately 50% of paralogs have different expression across different soybean tissues, and have undergone subfunctionalization or neofunctionalization

(Roulin et al., 2013). Both GmABCA1 and GmABCA2 have a paralog: Glyma06g20370 (now named Glyma.06G191400 in Glyma2.0) and Glyma06g20360 (now named Glyma.06G191300 in Glyma2.0), respectively. In this study primers that were more specific to GmABCA1 and

GmABCA2 were used in real-time qPCR to confirm the expression pattern of the two genes.

Expression of GmABCA1 was similar to that determined by Clarke (2013) but the new primer set showed much more specific expression of GmABCA2 in nodules compared to other tissues as seen in both transcriptome studies (Libault et al., 2010; Severin et al., 2010) suggesting that the primers used by Clarke (2013) may have amplified the Glyma.06G191300 as well as

GmABCA2 transcripts (Appendix 3). Comparing the GmABCA1 and GmABCA2 to their paralogs suggests that there has been subfunctionalization of the paralogs as Glyma.06G191400 and Glyma.06G191300 both have low expression in a number of different tissues including the nodule (Libault et al., 2010; Severin et al., 2010).

Examination of GmABCA1 and GmABCA2 expression across nodule development showed that both genes exhibited a similar expression pattern. Their expression was low in young nodules, increased as the nodule began to mature and started to fix nitrogen (nitrogen fixation starts at around 14 DPI based on the previous assay by Clarke (2013)) and then began to decline at 19 DPI (Figure 3.11B). The high expression of both genes during SNF and nodule maturation suggested that both genes are involved in the process of nitrogen fixation or have roles in nodule maintainence and maturation.

3.4.2 GmABCA1 and GmABCA2 are specifically expressed in the infected

nodule cells

In situ RT-PCR was utilized as an alternative method that allows detection of specific mRNA in situ on a fixed tissue section. The method provides information on gene activity at the transcript level in specific cell-types, without the need for an appropriate promoter to be cloned as required for the GUS assay (Athman et al., 2014). Using this technique, GmABCA1 and

GmABCA2 mRNAs showed infected cell-specific expression (Figure 3.12C and D), in a similar fashion to the GmYSL7 positive control which has been shown previously to be specific to the infected cells (Figure 3.12B, Chen, 2012; Appendix 5). The infected cell expression of GmABCA1 and GmABCA2 agrees with the proteomic data of Clarke et al. (2015), where GmABCA1 and GmABCA2 were identified in the SM fraction of the symbiosomes, which are only present in the infected cells (Clarke et al., 2015). The specific infected cell expression supports a role of these proteins in the symbiosis.

3.4.3 Localization

Determination of the subcellular localization of GmABCA1 and GmABCA2 proteins is important in order to understand their function in the symbiosis and to confirm their localization on the SM from proteomic analysis. A fluorescent protein (FP) tagging approach was employed to determine the location of GmABCA1 and GmABCA2 proteins in living cells. Together with the previous proteomic analysis, these two approaches would serve as the main tools to provide complementary and independent information on the location of both proteins within nodules

(Tanz et al., 2013).

3.4.3.1 GmABCA1 localization in G. max

An N-terminal GFP fusion construct driven by the GmLbC3 promoter was prepared for

GmABCA1 and was expressed three times in soybean but with no success in obtaining GFP fluorescence. A possible reason for why no fluorescence was detected is that the fused GFP caused improper folding or instability of the chimeric GFP-GmABCA1 (Hanson and Köhler,

2001). These conformational changes in GFP-tagged GmABCA1 may have resulted in production of a mis-folded or unstable protein, which was targeted for degradation by the proteasome through the proteolytic pathway (Goldberg, 2003; Schrader et al., 2009). An additional reason is that the N-terminal GFP tag might abrogate or interfere with many targeting signals such as the ER signal peptides and posttranslational modification sites which are essential for membrane targeting (Tian et al., 2004).

Confirming the localization of GmABCA1 to the SM in soybean has proved to be a challenging task. The method of A. rhizogenes-mediated hairy root transformation is a laborious process, and possesses some limitations that include low transformation efficiency and the lack of selection in the transformation system. The visual reporter marker of GFP or DsRED used in this study does not provide the same advantages as a traditional selectable marker (e.g. herbicide resistance) in removing untransformed roots. Producing stable transformants is beneficial, but it is time consuming and was beyond the scope and timeframe of my PhD.

Besides, the use of FP tagging to determine the localization of GmABCA1 in nodules is not entirely optimized. Not much is currently known about proteins targeted to the SM, hence whether the fused GFP has interrupted or masked essential protein targeting signals is unclear.

Although most SM transporters studied in our lab could be localized to the SM when the N- terminus of the protein was labeled with GFP (Clarke et al., 2015; Gavrin, unpublished), this did not seem to be the case for GmABCA1. This may indicate that different mechanisms for protein import into the SM are operating, as proposed by Catalano et al. (2004).

3.4.3.2 GmABCA1 localization in Allium porrum L.

GmABCA1 tagged with GFP localized to the cytoplasm as well as the nucleus of leek epidermal cells (Figure 3.13, left panel), while the PM-mCherry marker labeled the nuclear envelope (Figure 3.13, middle panel). This suggests that the PM-mCherry marker was most probably localized in the ER, which is connected to the nuclear envelope. The mCherry marker contain PM targeting sequence and thus it should be localized to the PM (Nelson et al., 2007).

It is possible that the mCherry marker was not able to exit the ER as it does not meet the ER quality control before being secreted through the secretory pathway (Vitale and Boston, 2008).

The PM-mCherry marker was also diffusely located in the cytoplasm. These results suggested that the PM marker and possibly GmABCA1 were not localized correctly.

No localization for GmABCA1 could be predicted in leek, but the chimeric C-terminal GFP construct did at least produce detectable GFP fluorescence. This suggests that GFP positioned at the C-terminal end of ABCA1 allows folding of the fusion protein such that it can be detected by its fluorescence. It follows that ABCA1 may have an N-terminal targeting sequence similar to that seen for M. truncatula NOD25 (Hohnjec et al., 2009). Future experiments should examine the localization of GmABCA1 with a C-terminal fusion of GFP in soybean.

If the C-terminal fusion is not effective, GFP could be inserted internally within GmABCA1 that might prevent interference of targeting signals by N- or C-terminal GFP (Tian et al., 2004).

This method has worked well for GmZIP1, which is one of the transport proteins on the SM

(Chen, 2012). Aside from FP tagging, immunogold labelling using an antibody raised against

GmABCA1, which was used for localizing H+- ATPase to the SM (Fedorova et al., 1999) could be attempted. Small hemagglutinin (HA) tags could also be added to the C-terminal of

GmABCA1. An advantage of this tag is that it does not alter the structure of the tagged protein to a large extent. This was demonstrated in the localization study of M. truncatula Natural

Resistance-Associated Macrophage Protein 1 (Tejada-Jiménez et al., 2015).

3.4.3.3 GmABCA2 is localized on the SM

GmABCA2 localized to the SM in the infected cells of soybean nodules, supporting its identification in the SM proteome (Clarke et al., 2015). The results of FP tagging and proteomic analysis have provided complementary information on the location of GmABCA2 within nodules. GFP-tagged GmABCA2 was detected at relatively high levels, forming clear ring-like structures within the infected cells (Figure 3.14, left panel). Since SMs are the major internal membranes of the infected nodule cells, this demonstrates that GmABCA2 is localized on the

SM. Moreover, a similar labelling pattern was observed in previous experiments that labeled the SM (Clarke et al., 2015; Appendix 6; Fedorova et al., 1999). The intensity plots of GFP and

FM®4-64 suggested that GmABCA2 is specifically localized on the SM, and not at the PM of infected cells (Figure 3.15). Colocalization of both fluorophores at the SM could be drawn with confidence, depicted by the repeated concurrence of overlapping signals of both probes at the

SM (Figure 3.15). The discovery of GmABCA2 on the SM also supports that GmABCA2 is an

integral membrane protein as predicted by all four membrane topology programs (Section

3.3.2).

The detection of GFP-tagged GmABCA2 suggests that GFP was optimally placed within the folded GmABCA2 protein and that the N-terminal GFP tag did not interfere with endogenous targeting signals. A number of SM proteins were reported to be successfully targeted using N- terminal fusions of GFP. This includes M. truncatula TIP1g (Gavrin et al., 2014), M. truncatula

Rab7 (Limpens et al., 2009), GmNPF5.24 and GmNPF5.25 (Clarke et al., 2015). Mistargeting may occur through the use of a strong promoter instead of the native promoter because expression of fusion protein is higher than the endogenous protein (Tanz et al., 2013). Given the GmABCA2 promoter did not work for GUS assay, another promoter was needed and the use of GmLbc3 promoter seems to have worked.

GmABCA2 is a half-transporter and would be assembled as a complex of either homodimers or heterodimers to form a functional protein. Different dimer combinations were observed for

AtABCG11, which is capable of forming homodimers or heterodimers with AtABCG12

(McFarlane et al., 2010). Dimerization of AtABCG12 with AtABCG11 is required for

AtABCG12 to exit the ER and localize to the PM. However, AtABCG11 trafficking to the PM is independent of AtABCG12 (McFarlane et al., 2010).

It remains to be determined whether GmABCA2 forms a homodimer and/or heterodimer. If

GmABCA2 forms a homodimer, GmABCA2 will be able to exit the protein maturation machinery in the ER independent of the presence of another protein for its localization to the

SM. Alternatively, if GmABCA2 forms an obligate heterodimer, similar to AtABCG12, its secretion from the ER will require dimerization with another protein. A study conducted on mouse ABCG5 and ABCG8 introducing missense mutations into either transporter resulted in retention and degradation of the transporters in the ER (Graf et al., 2004). Therefore, if

GmABCA2 forms a heterodimer, mutations introduced into either GmABCA2 or its partner (if it can be identified) can be tested to assess whether heterodimerization is required for SM localization. There is a possibility that GmABCA2 can form a heterodimer with GmABCA1.

The bimolecular fluorescence complementation (BiFC) method could be used to test whether they physically interact with each other in vivo. This method was successfully applied to

AtABCG11 and AtABCG12 (McFarlane et al., 2010).

3.4.4 Yeast complementation

The baker’s yeast Saccharomyces cerevisiae mutants that are defective in specific transport processes are a useful tool for studying transport activity of plant transporters through functional complementation assays. Complementation of the mutation provides evidence that the transporter possesses transport ability for that particular substrate. However, as the SM is inverted compared to the PM, import of substrates in yeast by SM transporters suggest the transporter would export its substrate out of the symbiosome into the infected cell cytosol

(White et al., 2007).

Another consideration that needs to be given when interpreting yeast complementation assay results is the localization of the plant transporters in yeast. The transporters are either recognized by yeast targeting machinery or via an unknown mechanism to reach the final

destination (Frommer and Ninnemann, 1995; Rentsch et al., 1998). They are usually assumed to be localized to the PM if complementing a yeast mutant located on the PM. The localization in yeast is not usually performed together with the complementation assay, even when complementation takes place (Komarova et al., 2008; Moreau et al., 2002; Rentsch et al., 1995).

Nevertheless, determining the localization of plant transporters in yeast is important as it provides a direct proof that if complementation occurs, the direction of transport is the same as the yeast intrinsic transport direction and that it implies that the transporters are on the PM of yeast. Unfortunately, none of the localization of the candidates was assessed in this study due to shortage of time.

3.4.4.1 GmABCA1 and GmABCA2 could not transport iron into yeast

The ability of GmABCA1 and GmABCA2 to transport iron was reinvestigated in this study, with appropriate controls prepared. Neither GmABCA1 nor GmABCA2 could rescue the growth of fet3fet4ftr1 on iron-limiting media, suggesting they were unable to transport iron into yeast cells (Figure 3.16). No known ABCA transporters have been shown to transport iron before, but a plant ABC transporter of subfamily B, AtABCB25, have been implicated in the biogenesis of iron-sulfur clusters in plants (Bernard et al., 2009).

Since the localization of GmABCA1 and GmABCA2 in fet3fet4ftr1 was not assessed in this study, it is assumed that they are located on the PM of yeast. If they have the same orientation in the two membranes (SM and yeast PM), and are bidirectional transporters, it can be concluded they could not transport iron as growth of fet3fet4ftr1 yeast cells expressing

GmABCA1 and GmABCA2 could not be restored.

3.4.4.2 GmABCA1 and GmABCA2 could transport fatty acids into yeast

A fatty acid consists of a long hydrocarbon chain and a terminal carboxylate group (Berg et al.,

2002). It serves four main purposes: (i) as building blocks of cellular lipids, essentially as constituents of the biological membranes, (ii) as reserve lipids, stored in the form of TAG, (iii) as precursors for signaling molecules and hormones, and (iv) as a component for protein modification by the covalent attachment of fatty acids for membrane targeting of the protein

(Berg et al., 2002; Li et al., 2016). Synthesis of fatty acids initially occurs by a combination of the action of acetyl-CoA carboxylase and fatty acid synthase (Gurr et al., 2008). In plants, de novo fatty acids are synthesized in the plastids and transported to the ER for modification and lipid assembly, necessitating fatty acid or lipid transport (Li et al., 2016). Several types of transport exist including diffusion, vesicular transport, protein-mediated transport, flip transfer, and membrane contact sites (Li et al., 2016).

In yeast, uptake of exogenous long chain fatty acids is mediated by the fatty acid transport protein (Fat1p) (Færgeman et al., 1997). Fat1p works in concert with the fatty acyl-CoA synthetases Faa1p and/or Faa4p for fatty acid transport and subsequent activation of the imported fatty acids in S. cerevisiae in a process called the vectorial acylation (Zou et al., 2003).

Mutants lacking either Faa1p/ Faa4p or Fat1p are unable to grow when FAS is inhibited by cerulenin and supplementation of exogenous long chain fatty acids is required for cell viability

(Duronio et al., 1992; Færgeman et al., 1997; Johnson et al., 1994).

A drop test assay of Δfat1 cells expressing ScFAT1, empty vector, GmABCA1 and GmABCA2 showed that all constructs grew at a similar rate on SD containing 25 µM cerulenin and 50 µM

oleic acid (data not shown), demonstrating that the assay produced inconclusive results. A possible reason for the lack of noticeable growth differences for all constructs is that yeast growth is rapid in exponentially growing cells, where cell densities increased at a sufficient level to overcome growth inhibition of the antibiotic cerulenin. Another reason could be due to unknown yeast intrinsic transporters other than Fat1p that may function by mediating fatty acid import into yeast cells or through a passive diffusion of fatty acids (Li et al., 2015a).

The complementation assay of the yeast mutant Δfat1 was repeated in a growth curve assay.

GmABCA1 and GmABCA2 were able to complement Δfat1, transporting oleic acid into Δfat1 cells with different rates. Though GmABCA2 transported oleic acid with a similar rate to

ScFAT1, growth promotion in GmABCA2 occurred later than in the ScFAT1 expressing line

(Figure 3.17). This later growth promotion was also observed for GmABCA1 (Figure 3.17).

This is likely to be because the imported oleic acids cannot be properly esterified with CoA into acyl-CoA thioesters upon transport in the vectorial acylation process, as GmABCA1 and

GmABCA2 cannot properly interact with Faa1p or Faa4p (Li et al., 2015a; Zou et al., 2003).

Replication of this Δfat1 yeast assay, is recommended to ensure the results are reproducible.

The ability of GmABCA1 and GmABCA2 to complement yeast mutant Δfat1 on the PM suggests that they are involved in the import of fatty acids into the cell. This raises question about the orientation of these transporters in yeast. Transport from the symbiosome into the infected cell cytosol is equivalent to uptake into the yeast cytosol. This implies that if they localized on the PM of yeast and have the same physical orientation as in the symbiosome, they

would be capable of mediating export of oleic acid, or similar FA, out of the symbiosome across the SM.

3.4.6 GmABCA1 and GmABCA2 complement Atabca9 mutant

Expression of GmABCA1 and GmABCA2 under the control of the AtABCA9 native promoter in the abca9-1 mutant background restored establishment of seedlings on half strength MS without sucrose (Figure 3.19), but to different degrees (Figure 3.20). This is likely due to the varying strength of expression of the transgenes driven by the AtABCA9 native promoter. A previous study by Cernac and Benning (2004) also observed a varying degree of complementation between two WRI1 transgenic lines 106 and 107. Line 106 restored seed oil amounts closed to the WT levels, but line 107 did not. The result correlated with the higher abundance of the WT copy of the WRI1 mRNA in line 106 compared to line 107 (Cernac and Benning, 2004). Kim et al. (2013) did not determine the extent of phenotype recovery, but RT-qPCR showed comparable AtABCA9 transcript abundance between the complementation lines and WT Col-

0. The expression of the transgenes in the complementation lines in the present study was not examined due to lack of time. Further studies are necessary to confirm whether the higher percentage of phenotype recovery of some complementation lines (especially AtABCA9 C2,

GmABCA1 C1 and GmABCA2 C2) correlates with stronger expression of the transgenes driven by the AtABCA9 promoter.

That GmABCA1 and GmABCA2 complemented Atabca9 suggests that both transporters localized to the ER and could transport free fatty acids and acyl-CoAs, which are the substrates transported by AtABCA9 (Kim et al., 2013). The higher mean percentages of phenotype

recovery in GmABCA2 C2 compared to GmABCA1 C1 (Figure 3.20) seemed to be in agreement with the Δfat1 yeast complementation results where higher yeast growth rate was observed for GmABCA2 than GmABCA1. These results may suggest that although both proteins can transport fatty acids, GmABCA2 has a higher rate for fatty acid transport in comparison to GmABCA1. GmABCA2 may be more highly expressed in the heterologous systems.

The localization of GmABCA1 and GmABCA2 in the transgenic Arabidopsis was not assessed in this study. AtABCA9 is an ER-localized exporter, transporting acyl-CoAs and free fatty acids from the cytosol into the ER. Its direction of transport is parallel to the direction of transport into the symbiosome. GmABCA1 and GmABCA2 are assumed to localize to the ER when the

ER retention signal, HDEL (Denecke et al., 1992) was added to the C-terminus of both proteins

(which do not have a natural ER retention signal). That GmABCA1 and GmABCA2 complemented AtABCA9 suggests that their direction of transport when expressed in

Arabidopsis is the same as AtABCA9. However, the direction of transport is the opposite in yeast, suggesting that GmABCA1 and GmABCA2 could possibly function as bidirectional transporters.

The likely presence of lipid or fatty acid transporters on the SM suggests that lipids or fatty acids must be transported from the site of synthesis (chloroplast) into the symbiosome. The mechanism of lipid trafficking to the SM is currently unknown, but two modes of lipid transport to the symbiosome have been proposed: vesicular transport and non-vesicular transport

(Hurlock et al., 2014). SNARE proteins are involved in vesicle targeting to the SM (described in Section 1.1.2), while lipid transfers proteins (LTPs) are involved in non-vesicular transport 85

(Hurlock et al., 2014). It is very likely that GmABCA1 and GmABCA2 are involved in the non- vesicular transport of fatty acids since they are integral membrane proteins. AsE246 LTP of the

Chinese milk vetch (Astragalus sinicus) localized to the SM and is involved in lipid transport for symbiosome membrane biogenesis (Lei et al., 2014). Another LTP, M. truncatula N5

(MtN5) was expressed in the root hairs and nodule primordia during early stages of symbiosis, and is required for interaction with Sinorhizobium meliloti and regulation of root tissue invasion

(Pii et al., 2009; Pii et al., 2013).

The ability of GmABCA1 and GmABCA2 to transport fatty acids raises interesting questions about the role of fatty acids or lipids in the rhizobia-symbiosis. A large amount of lipids is needed to accommodate symbiosome biogenesis during the early stages of nodule development

(Verma et al., 1978). Lipids such as lipo-chitooligosaccharides (Nod factors) produced by rhizobium act as signaling molecules, essential for the host plant to recognize rhizobium as a symbiotic microorganism and not a pathogenic microorganism (den Hartog et al., 2003).

However, not much is known about the role of lipids or fatty acids during later stages of nodule development (when expression of GmABCA1 and GmABCA2 is highest).

A possible function of lipids during later stages of nodule development is that the carbon supplied by the plant to the bacteroid could be in the form of lipid. Lipid droplets or bodies were found in peanut nodules, where they could act as a carbon storage compound (Siddique and Bal, 1991). The nature of carbon storage compounds in bacteroids seems to vary between different bacterial-plant symbioses, as glycogen and poly-β-hydroxybutyrate (PHB) have also been detected (Lodwig and Poole, 2003; Udvardi and Poole, 2013). Another possible role is

that lipids could act as signaling molecules, mediating interactions between host symbiont and the bacteroid. Lipids have long been established as vital signaling molecules, where lipid- mediated signaling have the capacity to control essential cellular processes such as cell proliferation in humans and stress responses in plants (Okazaki and Saito, 2014; Wymann and

Schneiter, 2008). To our knowledge, a role for lipid as a signaling molecules in the later stages during nodule maturation and nitrogen fixation has not been proposed before. Lipid as signaling molecules may be essential to maintain legume-rhizobia symbiosis, and may be required for the bacteroid to fix nitrogen. The biological roles of lipids or fatty acids during these later stages of nodule development warrants further study.

The present study describes for the first time the discovery of protein-mediated fatty acid transport on the SM. GmABCA1 and GmABCA2 were able to rescue growth of fatty acid uptake-deficient yeast mutant (Δfat1) and the Atabca9 T-DNA mutant, demonstrating that they are able to transport fatty acids. Whether GmABCA1 and GmABCA2 possess the same substrate specificity and the range of substrates that could be transported by both transporters have yet to be resolved. Transport studies in yeast and Xenopus oocyte (discussed in Section

5.3) could be carried out to determine substrate specificity. However, the results so far suggest that there is overlap in substrate specificity between GmABCA1 and GmABCA2, though

GmABCA2 appears to have higher rate for fatty acid transport compared to GmABCA1 from the data of the two complementation experiments.

Functional redundancy has occurred between GmABCA1 and GmABCA2 for the transport of fatty acids across the SM, evidenced from the overlapping substrate specificities. Whether the

functional redundancy is complete or partial is unclear and will require further study (Pickett and Meeks-Wagner, 1995). Gene members from the same family may display functional redundancy and incomplete diverged functions. This situation may have risen as the result of gene duplication, where redundancy is not lost by genetic drift and both copies of the gene remain functional (Thomas, 1993). GmABCA1 and GmABCA2 lie next to each other on the chromosome, and although they only share 24.4% amino acid identity, the two genes may have arisen by duplication before the soybean whole genome duplication occurred (Roulin et al.,

2013). Presumably they would subsequently undergo divergence from a single precursor gene and retain the same overlapping function, but may also have other unknown distinct roles

(Thomas, 1993). The functional redundancy of GmABCA1 and GmABCA2 for the transport of fatty acids across the SM may have a significant biological implication in the overall process of maintaining symbiosis.

Functional redundancy between two ABC transporters also occurred in human ABC transporters HsABCD1 and HsABCD2 (van Roermund et al., 2011). Both transporters possess distinct but overlapping substrate specificities when complemented and restored β-oxidation of different fatty acids in the yeast mutant pxa1/pxa2 (van Roermund et al., 2011), suggesting partial redundancy of function. High-affinity sulfate transporters AtSULTR1;1 and

AtSULTR1;2 also displayed redundancy of function (Barberon et al., 2008). Interestingly,

AtSULTR1;2 is able to completely complement sultr1;1 T-DNA mutant, but AtSULTR1;1 could only partially complement sultr;2 T-DNA mutant, suggesting unequal functional redundancy between the two transporters, and an additional uncovered role of AtSUTR1;1

(Barberon et al., 2008).

3.4.7 Future characterization of GmABCA1 and GmABCA2

Analogous to many ABC subfamily A members, both GmABCA1 and GmABCA2 can mediate the transport of lipids, or more specifically fatty acids. Complementation in the fatty acid uptake-deficient yeast mutant, Δfat1 and in Atabca9 T-DNA Arabidopsis mutant has shown that both proteins possess the ability to transport fatty acids. Additional experiments can be performed to provide further evidence of their fatty acid transport function.

Both GmABCA1 and GmABCA2 contain multiple TMDs that are necessary for substrate binding and recognition. The hydrophobic cavity of the TMDs represent the substrate-binding site, which can either be a single broad substrate-binding site or multiple sites with selective substrate specificity (Arora et al., 2001; Veen and Callaghan, 2003). Determining the type of substrate-binding sites by resolving their structures is beyond the scope of this project, but assessing whether GmABCA1 and GmABCA2 possess lipid binding activity may be useful to show that they can bind and transport lipids. For this, an in vitro assay for lipid binding activity could be utilized, which measures the fluorescence intensity emitted by fatty acid aromatic side chains when bound to a protein of interest in a hydrophobic environment. The lipid binding activity in vitro has been successfully observed for SM LTP AsE246 (Lei et al., 2014) and nodule LTP MtN5 (Pii et al., 2009).

The directionality of GmABCA1 and GmABCA2 transporters can be confirmed by complementation in yeast mutant pxa1/pxa2. This yeast mutant has defects in Pat1p and Pat2p transport proteins required for the import of long-chain fatty acids into the peroxisomes

(Hettema et al., 1996; Swartzman et al., 1996). The direction of fatty acid import into the

peroxisome is equivalent to the direction of transport into the symbiosome. If GmABCA1 and

GmABCA2 could complement yeast mutant pxa1/pxa2, this suggests that both proteins are bidirectional transporter that can mediate the import and export of fatty acids across the SM.

To perform this experiment, a peroxisome targeting signal need to be added to GmABCA1 and

GmABCA2 proteins for correct targeting of the proteins when expressed in yeast mutant pxa1/pxa2.

RNA interference (RNAi) was utilized to reduce the expression of GmABCA2. Nodules expressing GmABCA2 RNAi did not produce any obvious nodule phenotype and nitrogen fixation was not affected (data not shown). These results supported the functional redundancy occurring between GmABCA2 and GmABCA1 in the transport of fatty acids and that both of them may need to be knockdown together for detectable phenotypes. An RNAi construct was designed to reduce expression of both of them, but no transgenic roots formed, and thus RNAi experiments cannot proceed. This experiment warrants another attempt and should be the focus of future experiments.

Additionally, mass spectrometry-based lipidomic analysis to see whether the lipid composition of the symbiosome in the RNAi nodules is altered could be performed (Brügger, 2014). This approach would be especially informative to determine if reduced amounts of certain lipids are related to the reduced transport activity of GmABCA1 and GmABCA2. An RNAi approach has been used effectively to study a number of nodule proteins including GmNPF8.6, GmYSL7

(Gavrin and Smith, unpublished), GmUPS1 (Collier and Tegeder, 2012) and LTP AsE46 (Lei et al., 2014).

Extraction and isolation of symbiosomes and bacteroids from nitrogen-fixing nodules and subsequent transport assays on the isolated structures using radioactively labeled fatty acids for

GmABCA1 and GmABCA2 could also be attempted. The method is useful for determining whether bacteroids can also transport fatty acids. The method has effectively detected the presence of a number of transport processes across the SM and BM. Some examples include a dicarboxylate transport (Udvardi, 1988), ammonia transport (Udvardi and Day, 1990), ATPase activity (Udvardi, 1989), and glutamate and aspartate transport across the BM (Kouchi et al.,

1991). These experiments and their use will be discussed in further detail in the general discussion.

Chapter 4

Investigation of putative peptide transporters in soybean nodules

4.1 Introduction

The activity of peptide transporters is required for acquisition and translocation of peptides across cell membranes. Three peptide transporter families have been identified in plants, each transporting peptides of different lengths (Rentsch et al., 2007). Small di- and tripeptides are transported by members of the nitrate transporter 1/ peptide transporter (NRT1/PTR) family

(NPF), while the oligopeptide transporter (OPT) family mediates transport of larger peptides of

3-8 amino acids. The OPT family can be grouped into two subfamilies; (i) the true OPTs, and

(ii) the yellow stripe-like transporters (YSL). The third peptide transporter family is the

multidrug resistance-associated proteins (MRPs, ABCC), which is part of the wider ABC superfamily (Rentsch et al., 2007). AtMRP1-4 in A. thaliana have been shown to transport conjugates of the tripeptide glutathione (GSH) (reviewed in Bachhawat et al., 2013).

4.1.1 The roles of peptides

Transport of peptides across the cell membrane has been well documented in bacteria, fungi, animals and plants (Daniel et al., 2006; Payne and Smith, 1994; Tsay et al., 2007). A wide range of important cellular processes involve the transport of peptides. These include chemotaxis and quorum sensing (QS) in bacteria (Kumar and Engelberg-Kulka, 2014; Manson et al., 1986), transport of peptide-derived antibiotics in humans (Dantzig et al., 1992), and remobilization of nitrogen throughout the plant (Higgins and Payne, 1982). Nitrogen mobilization in the form of peptides, instead of in the form of individual amino acids, has been suggested to be a more efficient means for bulk movement and rapid distribution of nitrogen via long-distance transport, especially during seed germination and leaf senescence (Higgins and Payne, 1982;

Koh et al., 2002).

It is believed that the primary function of peptides is their utilization as amino acid, carbon or nitrogen sources (Koh et al., 2002; Steiner et al., 1995). However, recent studies reveal that peptides also function as important signaling molecules. In plants, as well as phytohormones

(e.g. auxin, cytokinin, and ABA) that have been recognized as the main intercellular signalling molecules, accumulating evidence suggest that peptides play crucial roles as signalling molecules in intercellular plant communication and development (Fukuda and Higashiyama,

2011). Intriguingly, some peptides are involved in the phytohormone signaling pathways, regulating and altering plant development (Ferguson and Mathesius, 2014).

The first characterized plant peptide is systemin, an 18 amino acid peptides involved in the wound response following mechanical damage in leaves by regulating the synthesis of defensive proteins (Pearce et al., 1991). A number of peptides are involved in the legume- rhizobia symbiosis, and are active at different stages of the symbiosis. These include nodule- specific cysteine-rich (NCR) peptides, clavata/ embryo surrounding region-related protein

(CLE) peptide, early nodulin 40 (ENOD40), rapid alkalization factor (RALF) peptide, C- terminally encoded peptide (CEP) and devil/rotundifolia-four-like (DVL1/ROT4) (reviewed in

Djordjevic et al., 2015; Ferguson and Mathesius, 2014).

The NCR peptides (30-50 amino acid residues) are particularly interesting as they play an essential role in controlling terminal differentiation of bacteroids, that occurs specifically in the inverted repeat-lacking clade (IRLC) legumes such as M. truncatula (Horváth et al., 2015; van de Velde et al., 2010; Wang et al., 2010). Additionally, CLE peptides (12 to 13 amino acid residues) of GmRIC1/2 in G. max and MtCLE12/13 in M. truncatula are involved in the systemic autoregulation of nodulation (AON) (Mortier et al., 2010; Reid et al., 2011a; Reid et al., 2013; Saur et al., 2011). AON is a mechanism that regulates the number of nodules the plant forms. These CLE peptides are negative regulators of AON that suppress further nodulation events (Reid et al., 2011b; Reid et al., 2012).

Most of these signaling peptides are larger than those transported by the peptide transporter families mentioned above, which transport small peptides. Most of the reported signaling peptides bind to or are perceived by receptors on the cell membranes. For example,

MtCLE12/13 and GmRIC1/2 peptides are perceived by leucine-rich repeat receptor kinases

MtSUNN and GmNARK that induces the downstream signaling pathway (Elise et al., 2005;

Searle et al., 2003).

Small peptides have roles in other symbiotic systems, for example, the cyclic dipeptides (e.g. cyclo-L-prolyl-L-leucine) in marine sponge Dysidea avara: Vibrio sp. symbiosis. The surfaces and internal spaces of marine sponges house commensally or symbiotically associated bacteria and fungi (De Rosa et al., 2003). It has been reported that bacteria communities in marine sponges communicate with each other through small cyclic dipeptides, also known as diketopiperazines (DKPs). These peptides function as QS signals, which mediate a cell-to-cell signaling mechanism in bacteria (Abbamondi et al., 2014). Nodules also contain other peptide derivatives such a glutathione (Frendo et al., 2013; Matamoros et al., 1999) that are transported by oligopeptide transporters (Zhang et al., 2016) and phytochelatins (PCs) are also peptide derivatives composed of three amino acids (Glu, Cys and Gly) that are glutathione-derived metal-binding peptides. In plants, PCs are involved in heavy metal stress tolerance and detoxification by acting as high affinity metal chelators (reviewed in Yadav, 2010).

4.1.2 Candidate proteins

Six proteins that have the potential to transport peptides were identified in the soybean SM proteome, including a number of members of the NPF family and a YSL family member. Three

other members of the NPF family (as well as those on the SM) have transcripts that are upregulated in soybean nodules (Libault et al., 2010; Severin et al., 2010; Table 4.1). The transporters that I am studying transport much smaller peptides than the peptides that are established signaling molecules in nodules and so the role of these di-, tri- and oligopeptide transporters is unclear. The primary aim of this chapter is to test whether these transporters have a peptide transport function. Peptides that have been used in yeast assays for other peptide and

YSL transporters (Hofstetter et al., 2013; Osawa et al., 2006; Pike et al., 2009) will be tested in heterologous complementation in yeast.

The NPF transporters, GmNPF1.2 (Glyma08g04160.1 v1.1, Glyma.08G037200.1 v2.0),

GmNPF5.24 (Glyma11g34613.1 v1.1, Glyma.11G224400.1 v2.0), GmNPF5.25

(Glyma11g34600.1 v1.1, Glyma.11G224200.1 v2.0), GmNPF5.29 (Glyma18g03790.1 v1.1,

Glyma.18G033900.1 v2.0), and GmNPF8.6 (Glyma02g38970.1 v1.1, Glyma.02G224600.1 v2.0), are part of sub-families 1, 5, and 8.

In A. thaliana, NPF members of subfamily 5 (AtNPF5.2) and 8 (AtNPF8.1, AtNPF8.2 and

AtNPF8.3) have been demonstrated to transport di- and tri-peptides when expressed in yeast and Xenopus oocytes (Karim et al., 2007; Komarova et al., 2008; Rentsch et al., 1995). Substrate specificity is quite conserved, at least within subfamily 8, hence GmNPF8.6 proteins found on the SM of G. max could potentially transport peptides which might play an important role in the symbiosis because of their specific expression in the nodule (Libault et al., 2010; Severin et al., 2010). The ability of members of the NPF5 family to transport peptides is more controversial as only AtNPF5.2 (AtPTR3) has been recorded as a peptide transporter

Table 4.1 Tissue-specific expression of GmNPF candidates based on the two soybean transcriptome studies: Libault et al. (2010) and Severin et al. (2010). Normalized expression data across five different soybean tissues: leaf (L), flower (F), pod (P), root (R) and nodule (N) were presented and is reads by the number of sequence reads/million reads aligned. The proteins identified in the SM proteome (Clarke et al., 2015) are indicated by (Y).

PROTEIN NAME SYSTEMATIC SM Libault et al. (2010) Severin et al. (2010) (v1.1/v2.0) NAME L F P R N L F P R N Glyma08g04160/ GmNPF1.2 Y 0 0 0 3 198 0 0 0 0 155 Glyma.08G037200 Glyma06g15020/ GmNPF5.2 0 0 0 0 67 0 0 0 0 74 Glyma.06G145200 Glyma11g34613/ GmNPF5.24 Y Glyma.11G224400 Glyma11g34600/ GmNPF5.25 Y 0 0 0 1 110 0 0 0 0 61 Glyma.11G224200 Glyma18g03790/ GmNPF5.29 Y 0 0 0 4 413 0 0 0 0 160 Glyma.18G033900 Glyma11g34580/.1 GmNPF5.30 0 0 0 0 318 0 0 0 0 96 Glyma.11G223900 Glyma04g39870/.1 GmNPF5.3 0 0 0 0 259 0 0 0 0 139 Glyma.04G220700 Glyma02g38970/ GmNPF8.6 Y 0 2 0 1 27 0 0 0 0 31 Glyma.02G224600 Glyma11g31870/ GmYSL7 Y 0 0 0 2 7 0 0 0 0 25 Glyma.11G203400

(Karim et al., 2007). More recent studies have suggested it transports hormone ABA and GA1, whereas AtNPF5.7 and AtNPF5.1 were able to transport ABA, GA1 and JA-Ile (Chiba et al.,

2015).

In this chapter I have tested each of the NPFs from the SM and also other NPF5 proteins upregulated in nodules: GmNPF5.2 (Glyma06g15020.1 v1.1, Glyma.06G145200.1 v2.0),

GmNPF5.3 (Glyma04g39870.1 v1.1, Glyma.04G220700.1 v2.0), and GmNPF5.30

(Glyma11g34580.1 v1.0, Glyma.11G223900.1 v2.0) for their ability to transport peptides in yeast. The systematic names used for GmNPFs are as listed in Léran et al. (2014) and will be used to refer to these proteins henceforth.

A potential oligopeptide transporter, GmYSL7 (Glyma11g31870.1 v1.1, Glyma.11G203400.1 v2.0) was also identified in the SM proteome (Clarke et al., 2015) and has specific expression in the nodule (Libault et al., 2010; Severin et al., 2010; Table 4.1). It is a member of the yellow stripe-like (YSL) family, which is part of the larger OPT family that transports larger peptides up to at least eight amino acids in length. This protein is most closely related to AtYSL7, with

73.9% protein identity, and hence we decided to use the systematic name GmYSL7. GmYSL7 was initially tested for its ability to complement iron-deficient yeast mutant strain (fet3fet4ftr1) for iron transport (Qu, unpublished). No complementation was observed, indicating that

GmYSL7 could not transport iron unlike many members of the YSL transporters (Curie et al.,

2009). Recent evidence showed that AtYSL7 is able to transport SylA, a peptide derivative that is a virulence factor secreted by a plant pathogen (Hofstetter et al., 2013). Thus, GmYSL7 could potentially be a peptide transporter, analogous to its homologue in A. thaliana.

Initial work on GmNPFs and GmYSL7 was previously conducted by Yihan Qu, Alex Gavrin and Penelope Smith (The University of Sydney) and is discussed further here. Nodule-enhanced expression of the GmNPF candidates and GmYSL7 was confirmed by an RT-qPCR assay (Qu,

2011; Qu, unpublished), consistent with the two published soybean transcriptome studies

(Libault et al., 2010; Severin et al., 2010). In a time-course RT-qPCR experiment to determine how the transcript abundance changes across nodule development, GmNPFs and GmYSL7 expression were broadly upregulated in mature nodules and during nitrogen fixation (Qu, 2011;

Qu, unpublished). Additionally, promoters of GmNPF1.2, GmNPF5.24, GmNPF5.25,

GmNPF8.6 and GmYSL7 assayed for histochemical GUS analysis showed that they are specifically expressed in infected cells of the nodule (Clarke et al., 2015; Gavrin and Smith, unpublished; Appendix 4), supporting their roles in the symbiosis. GFP tagging of GmNPF5.25,

GmNPF5.29, GmNPF8.6, GmNPF1.2 and GmYSL7 at the N-terminus driven by the GmLbC3 promoter indicated that they are localized to the SM (Clarke et al., 2015; Gavrin and Smith, unpublished) confirming the discovery of these proteins in the SM proteome (Clarke et al.,

2015).

RNAi-mediated gene silencing of GmNPF8.6 and GmYSL7 under the control of the GmLbC3 promoter was undertaken to investigate whether the proteins are essential for symbiosis.

Nodules with reduced expression of either GmNPF8.6 or GmYSL7 showed decreased nitrogen fixation, but produced different phenotypes. Nodules from GmYSL7 RNAi had blocked development of the infected cells at an early stage of development where symbiosomes have single rather than multiple bacteroids, suggesting that GmYSL7 is required for normal symbiosome development (Gavrin and Smith, unpublished). Nodules expressing GmNPF8.6

RNAi showed fairly normal morphology of the infected cells, although there are more vacuoles compared to the WT (indicative of early senescence), which suggests that GmNPF8.6 is required for nodule function rather than development (Gavrin and Smith, unpublished).

4.2 Materials and methods

4.2.1 Phylogenetic analysis

A phylogeny for NPF members was constructed using Phylogeny.fr with the default advanced settings (Dereeper et al., 2008). Protein sequences of 67 soybean NPF members (subfamily 1,

5 and 8), MtNPF1.7 (NIP/LATD, Medtr1g009200.1) and ZmNPF8.8 (GRMZM5G867390) were obtained from Phytozome (Goodstein et al., 2012). A. thaliana NPF amino acid sequences were obtained from TAIR (Lamesch et al., 2012). OsNPF5.5 (Os10g33210), OsNPF8.9

(Os03g13274) and OsNPF8.20 (Os06g49250) amino acid sequences were downloaded from

Plant Membrane Protein Database (http://aramemnon.uni-koeln.de/index.ep). Amino acid sequence alignment was built using MUSCLE (Edgar, 2004), and PhyML (Guindon and

Gascuel, 2003) algorithm was used to construct the ML tree. Bootstrapping test of 100 replicates was performed and the bootstrapping values were labeled at the corresponding nodes.

A ML tree was constructed for OPT superfamily proteins using MEGA6 (Tamura et al., 2013).

Amino acid sequences of G. max OPT proteins were downloaded from Phytozome and SoyBase while A. thaliana OPT proteins were obtained from TAIR. ScOPT1 (accession NP_012323),

CaOPT1 (accession XP_718267), BjGT1 (accession CAD91127), OsOPT3 (accession

AAQ56008) amino acid sequences were obtained from National Center for Biotechnology

Information (NCBI). A total of 37 protein sequences were aligned using MUSCLE aligner

(Edgar, 2004). The evolutionary model with the lowest Bayesian Information Criterion (BIC) score is considered the best substitution model. The best model was computed to be the Le

Gascuel model (Le and Gascuel, 2008) plus discrete Gamma distribution (+G), and the model was used to construct the neighbor-joining ML tree. The robustness of the nodes was assessed by bootstrapping replicates of 100. Sequence alignment of two proteins was examined using

EMBOSS Needle (Li et al., 2015b).

4.2.2 Heterologous complementation in yeast

4.2.2.1 Vector construction

Entry clones of GmNPF1.2, GmNPF5.2, GmNPF5.24, GmNPF5.25, GmNPF5.29, GmNPF5.3,

GmNPF5.30, GmNPF8.6, AtNPF5.2 (AtPTR3) were made by Yihan Qu, Alex Gavrin and

Muhammad Nazim Uddin (The University of Sydney). These entry clones were subsequently

LR cloned into yeast destination vector, pDR196-GW (Figure 3.5) as described in Section 2.4.6 to generate expression clones. Positive controls pDR195-AtNPF8.1 (AtPTR1) and pDR196-

AtOPT4, and negative control pDR196 empty vector were provided by Prof. Doris Rentsch

(The University of Bern).

4.2.2.2 Yeast growth, transformation and complementation

To examine whether GmNPFs are able to transport di- and tripeptides, their full-length CDS in pDR196-GW (expression clones) were transformed into LR2 yeast (MATa; hip1-614; his4-

401; can1; ino1; ura3-52; ptr2Δ::hisG; Rentsch et al. (1995)) along with controls AtNPF8.1

100

(AtPTR1), AtNPF5.2 (AtPTR3) and the empty vector pDR196. Ura+ transformants were selected on SD agar (0.67% YNB without amino acids) supplemented with 20 mM His and

0.02 g/l inositol. To test for peptide transport activity, transformants were tested on minimal medium (0.17% YNB without amino acids and ammonium sulfate, 167 µM histidine, and 20 mg/l inositol) containing 10 mM NH4Cl (positive control) or 100 µM Ala-Ala-Ala (AAA), Ala-

Ala (AA), Val-Leu (VL), Gly-Ala (GA), Gly-Glu (GE), Gly-His (GH), Gly-Leu (GL), Gly-Pro

(GP) and Gly-Gln (GQ) as sole source of nitrogen. For drop test assays, 3 µl of ten-fold serial dilutions were spotted on the test plates. All plates were incubated at 30°C and grown for 6 days.

A growth rescue assay was undertaken to examine whether AtOPT4, GmYSL7, AtYSL7,

AtYSL5 and AtYSL8 are able to transport leucine-containing peptides (4 to 6 amino acids) to satisfy an auxotrophic requirement of yeast mutant opt1 for nitrogen. S. cerevisiae strain

Y01213 (BY4741; MATa; ura3Δ0; leu2Δ0; his3Δ1; met15Δ0; YJL212c::kanMX4) from

Euroscarf was transformed with expression clones of AtOPT4, GmYSL7, AtYSL5, AtYSL7 and AtYSL8 along with the empty vector pDR196. Ura+ transformants were selected on SD agar containing 0.67% (w/v) YNB without amino acids, 2% (w/v) glucose, 30 mg/l leucine, 20 mg/l histidine, and 100 mg/l methionine. To test for peptide transport activity, transformants were serially spotted in a ten-fold dilutions on minimal medium (0.17% YNB without amino acids and (NH4)2SO4, and amino acids as required) containing 10 mM NH4Cl (positive control), or 100 µM ALAL, LSKL, IIGLM, and KLLLLG as sole source of nitrogen.

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4.3 Results

4.3.1 Phylogenetic analysis

4.3.1.1 The NPF phylogeny

A maximum likelihood analysis was performed for plant NPF subfamily 1, 5, and 8 proteins.

The number of proteins analyzed for the three subfamilies vary, with subfamily 5 making up the majority of proteins examined. Members of subfamily 1, 5, and 8 are grouped closely together, except for GmNPF5.34, which is a much smaller protein than most family members that clustered with subfamily 1 (Figure 4.1). GmNPF8.6 is most closely related to previously characterized di-, tripeptide transporter, AtPTR8.1 (AtPTR1, Komarova et al., 2008), among other characterized members of subfamily 8. They share 65.5% amino acid identity and 78.0% similarity, depicted by the shortest sequence distance (Figure 4.1).

Within the NPF5 subfamily, GmNPF5.13 has the highest sequence homology to characterized

AtNPF5.2 (AtPTR3, Karim et al., 2007) in comparison to other G. max NPF5 proteins, with a pairwise amino acid identity of 68.3% and 81.5% sequence similarity (Figure 4.1). Meanwhile, candidate proteins GmNPF5.24, GmNPF5.25 and GmNPF5.30 clustered away from AtNPF5.2, located in different clades. NPF subfamily 1 includes candidate protein GmNPF1.2, and three characterized members MtNPF1.7 (Bagchi et al., 2012; Zhang et al., 2014), AtNPF1.1 and

AtNPF1.2 (Chiba et al., 2015). Among these three transporters, GmNPF1.2 is most closely related to AtNPF1.1 with 43.7% protein identity and 62.5% similarity.

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0.2

Figure 4.1 Phylogenetic analysis of NPF proteins in G. max, A. thaliana and several characterized members in other plant species from subfamily 1, 5 and 8. 96 NPF proteins were aligned using MUSCLE (Edgar, 2004) and the alignment was used to produce a maximum likelihood tree using PhyML algorithm (Guindon and Gascuel, 2003) in Phylogenetic.fr online program (Dereeper et al., 2008). Bootstrapping tests of 100 replicates were conducted.

4.3.1.2 The OPT phylogeny

The OPT phylogenetic relationships were inferred by analyzing 37 proteins. Two distinct robust clades: YSL clade and true OPT clade were identified within the OPT phylogeny, with good bootstrap values (Figure 4.2). GmYSL7 is most closely related to AtYSL7 with the shortest distance (73.9% protein identity, 85.1% similarity), and clustered closely with AtYSL5,

AtYSL8, Glyma.16G212900 and Glyma.09G164500 (Figure 4.2). The true OPT clade contains the first characterized member of the OPT family: C. albicans CaOPT1, and also contains S. cerevisiae ScOPT1, AtOPT4 and AtOPT6 which are able to transport tetra- and pentapeptides

(Hauser et al., 2001; Lubkowitz et al., 1997; Osawa et al., 2006; Pike et al., 2009). Other members B. juncea BjGT1 and O. sativa OsGT1 transport the tripeptide GSH (Bogs et al.,

2003; Zhang et al., 2004).

4.3.2 Complementation of yeast mutant ptr2

Putative peptide transporters expressed in soybean nodules were tested to see if they could complement the S. cerevisiae mutant ptr2 (LR2 strain). This strain is defective in peptide transport across the PM (Rentsch et al., 1995; Tanaka and Fink, 1985) and as a consequence is unable to grow on media containing peptides as the sole nitrogen source or on media containing histidine exclusively in the form of a di- or tripeptide (Rentsch et al., 1995). LR2 yeast will only grow if the cells are able to transport and utilize the di- or tripeptides provided as the sole source of nitrogen.

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100 AtYSL5 64 AtYSL8 Glyma.16G212900 62 100 Glyma.09G164500 100 Glyma.11G203400 (GmYSL7) 100 AtYSL7 Glyma.09G281500 Glyma.20G004200 94 100 75 Glyma.20G004300 95 AtYSL4 AtYSL6 100 Glyma.19G094800 YSL 100 Glyma.16G054200 100 clade 100 Glyma.20G217400 88 Glyma.10G172900 Glyma.20G068300 100 100 Glyma.13G001500 AtYSL1

98 54 AtYSL2 AtYSL3

99 Glyma.17G191300

100 Glyma.06G133000 100 Glyma.04G231900 82 ScOPT1 CaOPT1 100 AtOPT1 100 AtOPT5 100 AtOPT3 100 BjGT1 100 AtOPT2 True 80 AtOPT4 OPT 100 AtOPT7 clade 67 OsOPT3 (OsGT1)

100 AtOPT8

100 AtOPT6 100 AtOPT9

0.5

Figure 4.2 Phylogenetic analysis of OPT proteins in G. max, A. thaliana and several

characterized members in other species. Amino acid sequences were aligned using MUSCLE (Edgar, 2004). The maximum likelihood tree was constructed using the neighbor- joining method, and the LG+G model. The tree is drawn to scale, in which the branch lengths represent the number of amino acid substitutions per site. Bootstrap values calculated for 100 replicates are indicated at the corresponding nodes. GmYSL7 is highlighted in the green box. Analysis was conducted in MEGA6 (Tamura et al., 2013).

Only LR2 cells expressing GmNPF8.6 and control AtNPF8.1 mediated growth on minimal media containing the tripeptide Ala-Ala-Ala and Gly-Pro (Figure 4.3). All yeast constructs including the negative control were able to grow on minimal media containing the dipeptide

Ala-Ala, Gly-Ala, Gly-Glu, Gly-Leu and Gly-Gln (only Gly-Leu plates as representatives were shown; Figure 4.3), demonstrating LR2 yeast may have another intrinsic transport system for transporting these dipeptides into the cell. No conclusion can be drawn for yeast growth on the other two selective plates supplemented with Gly-His and Val-Leu (data not shown). This is because either the controls did not grow as expected or yeast growth was so little that no conclusion could be reached. Since the first assay only showed complementation by

GmNPF8.6, this gene was tested in a second assay using a drop test.

Positive control AtNPF8.1, empty vector pDR196, and GmNPF8.6 were spotted on minimal media containing Ala-Ala-Ala, Gly-His, Gly-Pro, Val-Leu. Ala-Ala-Ala supported growth of

GmNPF8.6 as observed previously, having a similar growth rate to AtNPF8.1 (Figure 4.4).

GmNPF8.6 also mediated growth on Gly-Pro and Val-Leu, but transported Val-Leu at a lower affinity than AtNPF8.1 (Figure 4.4). Growth of LR2 cells expressing GmNPF8.6 and AtNPF8.1 was not observed on Gly-His (Figure 4.4).

4.3.3 Complementation of yeast mutant opt1

To investigate whether GmYSL7 and GmNPFs could transport peptides larger than four amino acids they were tested for complementation of the S. cerevisiae mutant opt1 (Y01213 strain).

Without OPT1 this strain cannot grow when oligopeptides of four or more amino acids are provided as the sole nitrogen source.

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A Template

EV AtNPF5.2 EV AtNPF5.7

AtNPF8.1 AtNPF5.2 AtNPF8.1 GmABCA2 GmNPF5.29 AtNPF5.1 GmNPF8.6 GmABCA1 GmNPF5.3 GmNPF5.25

GmNPF5.24 GmNPF1.2 GmNPF5.2 GmNPF5.30

B NH4Cl

C Ala-Ala-Ala

D Gly-Pro

E Gly-Leu

Figure 4.3 Complementation of yeast mutant ptr2 (LR2 strain) deficient in the uptake of di- and tripeptides on a streaked plate. Growth of LR2 cells expressing AtNPF8.1, AtNPF5.2, AtNPF5.1, all GmNPFs and the empty vector pDR196 was assessed on minimal medium supplemented with 10 mM NH4Cl (positive control plate), 100 µM Ala-Ala-Ala, Gly- Pro or Gly-His as the sole source of nitrogen. Yeast cells were grown at 30°C for six days.

0 -1 -2 -3 0 -1 -2 -3 OD600 10 10 10 10 10 10 10 10

GmNPF8.6

AtNPF8.1 EV

NH4Cl Ala-Ala-Ala

GmNPF8.6

AtNPF8.1

Gly-Pro Val-Leu

GmNPF8.6 AtNPF8.1

Gly-His

Figure 4.4 Complementation of yeast mutant ptr2 (LR2 strain) deficient in the uptake of di- and tripeptides. Growth of LR2 cells expressing GmNPF8.6, AtNPF8.1, and empty vector pDR196 was assessed on minimal medium supplemented with 10 mM NH4Cl (positive control plate), 100 µM Ala-Ala-Ala, Gly-His, Gly-Pro, and Val-Leu as the sole source of nitrogen. Ten-fold serially diluted cells (starting at OD600 1.0) were spotted on the selective plates and grown for six days at 30°C.

Only opt1 cells expressing AtOPT4 and AtYSL7 could grow on ALAL and KLLLLG as a sole nitrogen source (Figure 4.5). Growth on LSKL and IIGLM in cells expressing AtYSL7 was also observed. GmYSL7 did not grow on any of the media tested (Figure 4.5). Moreover, none of the GmNPFs grew and thus, they could not transport peptides larger than three amino acids

(data not shown).

4.4 Discussion

4.4.1 Only GmNPF8.6 is able to transport peptides

Of the eight GmNPFs tested in the complementation assay, only GmNPF8.6 supported the growth of yeast ptr2 on minimal media containing Ala-Ala-Ala, Gly-Pro and Val-Leu (Figure

4.4), indicating that GmNPF8.6 has a peptide transport function. GmNPF8.6 and AtNPF8.1

(AtPTR1) did not mediate growth on minimal media containing Gly-His. Transport of Gly-His dipeptide by AtNPF8.1 has not been investigated before, thus the lack of growth could be due to its inability to transport Gly-His, or its affinity for Gly-His is low hence a higher concentration of the dipeptide is needed for growth. Another possible reason could be due to insufficient amount of His rather than not enough nitrogen in the media. This reasoning could also apply for GmNPF8.6. Moreover, there is also possibility that the clones may have mutated and thus resequencing is needed to confirm the integrity of the clones.

Many peptide transporters, such as A. thaliana AtNPF8.1 (Dietrich et al., 2004), AtNPF8.3

(Rentsch et al., 1995), rabbit small intestine oligopeptide PepT1 (Fei et al., 1994) and S. cerevisiae PTR2 (Perry et al., 1994) have a low selectivity towards amino acid side chains. As

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0 -1 -2 -3 0 -1 -2 -3 OD600 10 10 10 10 10 10 10 10 AtOPT4 EV

AtYSL7

GmYSL7

AtYSL5

AtYSL8 NH4Cl ALAL

AtOPT4

AtYSL7

GmYSL7

AtYSL5

AtYSL8 KLLLLG LSKL AtOPT4

AtYSL7

GmYSL7

AtYSL5 AtYSL8

IIGLM Figure 4.5 Compleme ntation of yeast mutant opt1 (Euroscarf Y01213 strain) deficient in the uptake of 4-8 peptides. Growth of LR2 cells expressing AtOPT4, empty vector pDR196, AtYSL7, GmYSL7, AtYSL5 and AtYSL8 were assessed on minimal medium supplemented with 10 mM NH4Cl (positive control plate), 100 µM ALAL, LSKL, IIGLM, and

KLLLLG as sole source of nitrogen. Ten-fold serially diluted cells (starting at OD600 1.0) were spotted on the selective plates and grown for six days at 30°C.

a result, these transporters could recognize and transport a number of different peptides.

Transporters with low selectivity have been suggested to be required for mediating bulk transport of peptides during protein proteolysis (Dietrich et al., 2004). This is especially true as rapid distribution or remobilization of nitrogen in the form of peptides is important, for instance during germination, where hydrolyzed storage protein needs to be rapidly distributed to the growing parts of the seedling (Higgins and Payne, 1981). Only a small range of peptides were examined in the present study, thus GmNPF8.6 selectivity towards amino acid side chains, whether low or high selectivity has yet to be determined. Nevertheless, this study showed that

GmNPF8.6 has substrate selectivity at least within the di- and tripeptides tested.

GmNPF8.6 affinity for Ala-Ala-Ala and Gly-Pro seemed to be at a similar level to AtNPF8.1, but Val-Leu appeared to be transported at a lower affinity than AtNPF8.1, depicted by the slower yeast growth expressing GmNPF8.6 in comparison to AtNPF8.1 (Figure 4.4). Varying affinities for different peptides have been observed for AtNPF8.1, where AtNPF8.1 has a higher affinity for Ala-Ala and Ala-Lys, and a lower affinity for Ala-Asp when expressed in Xenopus oocytes (Dietrich et al., 2004). This demonstrated that the assay was not performed at saturating concentrations (Dietrich et al., 2004), which may also be the case in the present study. The concentration of peptides used in a transport assay is particularly important to assess substrate selectivity of a transporter. Future studies using different concentrations of peptides may reveal uptake affinity for GmNPF8.6.

It is unclear whether GmNPF8.6 is targeted to the PM of yeast, and its orientation when inserted into the membrane, although the fact that complementation was achieved suggests this is so.

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Assuming that GmNPF8.6 is localized on the PM of yeast and has the same physical orientation in the yeast PM and the symbiosome, complementation of GmNPF8.6 in yeast mutant ptr2 suggests that GmNPF8.6 is mediating export of peptides out of the symbiosome and into the infected cell cytosol. The role of peptide transport across the symbiosome is unclear, but based on the direction of transport, peptides could be the form of fixed N provided to the plant, in addition to ammonia or ammonium ions, which are the main form of fixed N.

4.4.2 The other GmNPFs may not be able to transport peptides

Other GmNPFs (GmNPF1.2, GmNPF5.2, GmNPF5.24, GmNPF5.25, GmNPF5.29,

GmNPF5.3, and GmNPF5.30) did not complement the ptr2 yeast mutant (Figure 4.3). A possible reason for why no complementation was observed is that they could possibly mediate transport of other peptides aside from the tested peptides, and might only transport a limited range of peptides. They could also have low affinities for the peptides tested in the present study as described above, hence no complementation was observed. Therefore, a wider range of tested peptides and higher concentration of peptides used are recommended in future complementation assays.

Two interesting examples showed that the concentration of peptide used is important in order to observe complementation. An oligopeptide transporter from A. thaliana, AtOPT4, restored growth on media containing GSH as the sole sulfur source when expressed in the yeast double mutant met15 opt1 (Zhang et al., 2016). This conflicted with the results obtained from a previous study which did not observe complementation (Osawa et al., 2006). A reason for the contradictory results is that the GSH concentration used in Osawa et al. (2006) was much lower

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(100 µM) compared to Zhang et al. (2016), which used up to 400 µM GSH. Another example is ScOPT1, where its growth on leucine-containing peptide was shown to be concentration- dependent. Growth is most robust when the highest concentration of YGGFL peptide (1 mM) was supplied, whilst no complementation was observed on media containing 10 µM peptide and little growth on 100 µM peptide (Hauser et al., 2000). Only 100 µM peptides were used in this study, and hence the use of higher concentration of peptides could be attempted.

The complementation assay used relies on the protein of interest being targeted to the PM so targeting of the heterologously expressed GmNPFs to another membrane in yeast could be reason for the lack of complementation. Some of the GmNPFs may lack the required sequences for assembly and targeting to the PM of yeast (Frommer and Ninnemann, 1995). In some cases, the vacuole has been shown to be the default yeast compartment for mistargeted proteins

(Cooper and Bussey, 1992; Roberts et al., 1992). Hence, the localization of transporters in yeast needs to be examined using fusions with markers such as the GFP or the immunogold labeling method to provide direct evidence that the GmNPF heterologous proteins are targeted to the

PM (Rentsch et al., 1998).

In our hands, AtNPF5.2 (AtPTR3, Karim et al., 2007) did not transport any of the peptides tested in this study (Figure 4.3) although we did not test the peptides tested by Karim et al.

(2007). AtNPF5.1 also did not support the growth on any peptides. These results suggest that the notion that NPF transporter subfamily 5 is able to transport peptides should be reevaluated.

A recent study showed that AtNPF5.2 can mediate the transport of ABA and GA1 using a yeast two hybrid approach, while both AtNPF5.7 and AtNPF5.1 were demonstrated to transport

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ABA, GA1 and JA-Ile (Chiba et al., 2015). This may also suggest that AtNPF5.2 is involved in the crosstalk between peptide and ABA/GA signaling, similar to AtNPF6.3 that is involved in the crosstalk between nitrate and ABA signaling (Krouk et al., 2010). Nonetheless, the roles of

GmNPF5s and GmNPF1.2 remain undetermined, but the fact that their expression is greatly enhanced within the nodule suggests that they play important roles in the symbiosis.

4.4.3 GmYSL7

In the present study, GmYSL7 did not transport any of the peptides tested when expressed in yeast mutant opt1. Only yeast mutant opt1 expressing AtYSL7 mediated yeast growth on all the peptides tested (ALAL, LSKL, KLLLLG, and IIGLM), whereas AtOPT4 only supported growth on ALAL and KLLLLG (Figure 4.5). The present results contrasted the results of a pilot study conducted by Penelope Smith, which observed complementation of GmYSL7 on media containing ALAL and LSKL (Smith, unpublished).

A possible reason for why no complementation was observed for GmYSL7 in the present study is that insufficient level of functional GmYSL7 product was produced instead of its inability to transport peptides. For this, the transcript level of GmYSL7 should be examined and analyzed by RT-qPCR assay to confirm that GmYSL7 transcript is actually present in the Ura+ transformants. Another method that could be tried is extracting yeast microsomal proteins, and performing PAGE and immunoblot analysis using the microsomal protein to confirm the presence of GmYSL7 protein. This method was used for UDP–L-Rha/UDP–D-galactose transporter 1 (URGT1) to provide evidence that the URGT1 protein is present for the yeast transport assay (Rautengarten et al., 2014).

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Although maybe only a fraction of the protein reaching the PM is sufficient for function and complementation (Rentsch et al., 1998), this may not be the case for GmYSL7. A portion of the A. thaliana H+-ATPase AHA2 and AHA3 proteins was adequate to observe complementation of a yeast mutant PM H+-ATPase (pma1), but only partial complementation was observed. A large proportion of the proteins were retained in the ER of the transformed yeast cells (Palmgren and Christensen, 1993; Palmgren and Christensen, 1994). There is a possibility that most of GmYSL7 protein may be trapped in the ER because it is not recognized correctly by yeast targeting machinery (Rentsch et al., 1998). Complementation is not observed since only a fraction of GmYSL7 is present on the PM of yeast, unlike AtAHA2 and AtAHA3, which only need a fraction of the proteins for partial complementation. This also stresses the importance of assessing the localization of GmYSL7 in yeast, whether it is retained in the ER or localized to the PM or elsewhere in the yeast.

New substrates transported by AtYSL7 were discovered in this study. AtYSL7 and AtYSL8 have been demonstrated previously to transport a peptide derivative SylA when expressed in yeast (Hofstetter et al., 2013). AtYSL7 was able to transport all the peptides used in the present study whereas AtYSL8 and AtYSL5 did not supported growth on any of the peptides tested

(Figure 4.5). These results suggest that AtYSL7 has low selectivity towards amino acid side chains and could probably transport a wide range of peptides. In comparison, AtYSL8 may have high substrate selectivity, but it seemed to have specificity for tetrapeptide AAAA, similar to AtYSL7, where addition of this peptide was able to inhibit transport of syringolin (Hofstetter et al., 2013). The results also suggest that substrate selectivity among AtYSLs may vary.

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To my knowledge, a role for transport of small peptides across the SM has not been proposed previously. The complementation shown in yeast suggest that these peptide transporters are likely to transport peptides from the bacteroid side to the cytoplasm. It is possible that peptides from the bacteroids could act as QS signals to communicate with each other and to behave coordinately with the legume hosts. Cyclic dipeptides produced by bacterial symbionts in marine sponges are involved in the QS mechanism, which regulated the interaction between bacterial communities and the host (Abbamondi et al., 2014). Cyclic dipeptides have also been shown to be involved in the switch in behavior between symbiosis and virulence in bacteria

(Bellezza et al., 2014; Scopel et al., 2013; Vikram et al., 2014).

The amino acids supplied by plants for bacteroids might be in the form of peptides, hence peptide transporters on the SM may be required to facilitate the process. After peptide uptake into the symbiosomes, peptides may be rapidly hydrolyzed by peptidases and used as a source of amino acids for the bacteroids. Various amino acids have been considered as the carbon source for that bacteroid that is supplied by the plant, but substantial evidence suggests that dicarboxylates are the main form of carbon source (Day and Copeland, 1991; Prell et al., 2009).

Evidence in pea (P. sativum) demonstrated that leucine, isoleucine, and valine (LIV) are required for effective nitrogen fixation, and that they needed to be supplied by the plant to prevent symbiotic auxotrophy for LIV by the pea bacteroids (Prell et al., 2009). Transport of these amino acids across the bacteroid membrane was essential but direct transport across the

SM was not assessed and the requirement did not appear to be universal for legumes (Prell et al., 2010). Additionally, glutamate and aspartate have been shown to support a high rate of

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nitrogen fixation and respiration in isolated bacteroids from soybean nodules (Bergersen and

Turner, 1988; Kouchi et al., 1991), although there is little evidence for uptake of amino acids across the SM (Udvardi and Day, 1997). However, given the probable direction of transport

(from the bacteroid side to the cytoplasm), the plausible function of peptide as an amino acid source is unlikely.

These different transporters are likely to have different roles since the phenotypes of the RNAi analyzed for GmNPF8.6 and GmYSL7 are different. Although both of these transporters could transport peptides, they have been shown in this study to possess different substrate specificities and may be required at different stages of nodule development. The GmYSL7 RNAi phenotype suggests involvement in the earlier stage of symbiosome development, whereas the GmNPF8.6

RNAi phenotype suggests it is required for mature nodule function.

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Chapter 5

General discussion

5.1 Introduction

SNF is of great importance to world agriculture, providing an alternative natural source of nitrogen inputs to plants. As well as decreasing the negative environmental impacts caused by the use of artificial nitrogen fertilizers, SNF also reduces the cost of crop production associated with the increased price of nitrogen fertilizer. SNF is calculated to contribute approximately 21 million tonnes (Tg) of fixed nitrogen annually, with soybean representing 77 percent of the total nitrogen fixed by crop legumes (Herridge et al., 2008). Given its importance, the genetic and molecular basis of legume SNF has been the focus of intense research for decades with the ultimate goal to improve existing symbioses and possibly to extend symbiosis to cereals and

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other major non-legume crops (Cocking et al., 2005; Oldroyd and Dixon, 2014; Udvardi and

Poole, 2013).

Elucidation of transport processes and metabolism of legume nodules is crucial to gain a better understanding of the molecular and genetic basis of legume-rhizobia symbioses (Clarke et al.,

2014; Udvardi and Poole, 2013). Early research to understand the transport processes on the

SM and BM are limited to biochemical and physiological approaches, where many studies have focused on the principal carbon and nitrogen metabolism as they are responsible for fueling

SNF and nitrogen assimilation (Udvardi and Poole, 2013). Other essential transport processes have also been characterized (reviewed in Clarke et al., 2014; White et al., 2007), but this may only represent a small portion of transport processes that occur on these membranes. A recent comprehensive proteomic analysis of the soybean SM has identified various transport proteins that might be involved in the transport processes on the SM and is undoubtedly an invaluable resource to facilitate the identification of novel transporters essential for SNF (Clarke et al.,

2015).

In this study, I have further characterized three classes of transporters that were identified in the proteomic analysis of soybean SM: ABC transporters, NPF transporters and a YSL transporter. GmABCA1 and GmABCA2 were chosen for further study because of their nodule- enhanced expression based on the two soybean transcriptomes (Libault et al., 2010; Severin et al., 2010). They have been identified as fatty acid transporters in this study, agreeing with the hypothesis that they are able to transport fatty acids or lipids based on their functional homologies with other ABCA members (Kim et al., 2013; Tarling et al., 2013; Vasiliou et al.,

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2009). GmNPFs and GmYSL7 were also chosen because of their enhanced expression within the nodule (Libault et al., 2010; Severin et al., 2010). Their homology with known peptide transporters in plants such as AtNPF8.1 (Komarova et al., 2008) and AtYSL7 (Hofstetter et al.,

2013) are suggestive that they might be peptide transporters on the SM. Only GmNPF8.6 was discovered to function as a peptide transporter in the present study. GmYSL7 could also transport peptides (Smith, unpublished), but the results could not be reproduced in this study.

The roles of others, however, remain undetermined but their enhanced expression at the onset of nitrogen fixation (Qu, unpublished) suggests important roles for these proteins in this symbiosis.

Several important questions remain unanswered, particularly the physiological role played by the transport processes I studied. Limitations associated with working with the polyploidy legume, soybean, with nodules, and particularly infected cells, as the study material and the requirement to use the hairy root transformation method to produce transgenic nodules make determining this role difficult. The direction of transport across the SM of the transporters studied is not yet resolved. For instance, for GmABCAs, results from the complementation experiments in planta and in yeast point to two possible transport directions. Secondly, while the ability of GmABCAs to transport fatty acids and that of GmNPF8.6 in peptide transport has been confirmed, their exact physiological roles in the symbiosis remain elusive. Their enhanced expression during nitrogen fixation suggests an important role in symbiosis, but the reasons why these transport activities are important for nitrogen fixation remain unclear. This chapter will address the limitations I have faced in this study and suggestions for new methods or techniques to overcome these problems will also be discussed.

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5.2 Direction of transport

Determination of transport direction of SM transporters, whether out of the symbiosome or into the symbiosome (or bidirectional) is essential to elucidate the physiological role these transporters may have in the symbiosis. The indecisive results gathered from this study stress the importance that the direction of transport is known so that their functions can be drawn with confidence. Assessing the localization in yeast is required but is insufficient to ascertain their transport direction. The SM is inverted compared to the PM of yeast, and the physical orientation in which the SM transporter is inserted into the membrane is uncertain, which complicates matters. Although it is likely that the SM proteins will have the same physical orientation as the PM proteins since SM proteins may also be inserted into the SM through the secretory pathway (Leborgne-Castel et al., 2010), no strong evidence has been presented to confirm this. Notwithstanding the above, determining the localization of transporters in yeast could indicate that the direction of transport is the same as the yeast intrinsic transport direction if complementation occurs and if localization to other yeast membrane compartments is ruled out.

Although transport assays with isolated symbiosomes can be used to study exporters that transport substrates from the cytosol to the symbiosome space (see Chapter 3), it is more difficult to study importers that transport from the bacteroid side of the SM to the plant cytosol.

Isolation of the symbiosomes and subsequent production of sealed inside-out SM vesicles

(where the SM is inverted) from soybean symbiosome could serve as a tool to study these importers (Christiansen et al., 1995). The method utilizes aqueous polymer two-phase fractionation to isolate the inside-out SM vesicles with fixed orientation (Christiansen et al.,

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1995; Larsson et al., 1988). Transport assays for uptake of radiolabeled compounds or using stopped-flow fluorimetry in the SM vesicles (Rivers et al., 1997) could be performed. This method has worked well for characterizing some transport processes on the SM including aspartate transport (Rudbeck et al., 1999), and ammonia transport (Niemietz and Tyerman,

2000), which are mediated by transport directed from the bacteroid to the host plant cytosol.

Based on the data from the two complementation experiments for GmABCAs, they could either function as bidirectional transporters or importers. It is generally accepted that ABC transporters could only operate in a unidirectional movement (either import or export) upon

ATP hydrolysis (Wilkens, 2015). However, a study in Lactococcus lactis challenges this view and showed that transport in both directions are possible both mechanistically and thermodynamically (Balakrishnan et al., 2004). Drug efflux pump LmrA mediates the export and the reverse transport of ethidium depending on the ATP levels (Balakrishnan et al., 2004).

Hence, the possibility that GmABCAs could function as bidirectional transporters, similar to

LmrA, cannot be ruled out.

Eukaryotic ABC proteins are mostly exporters, transporting substrates out of the cytoplasm

(Verrier et al., 2008; Wilkens, 2015). For instance, human ABCA1 and ABCA7 exporters are crucial participants in the export of lipids that include cholesterol and phospholipids (Abe-

Dohmae et al., 2004; Smith and Land, 2012). Only a few eukaryotic ABC transporters are importers and these include AtABCB4, AtABCB14 and CjABCB1 (Lee et al., 2008; Santelia et al., 2005; Shitan et al., 2003). Import of fatty acids into yeast to complement Δfat1 suggests

GmABCAs transport fatty acids out of the symbiosome and into the cytoplasm as importers.

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Alternatively, based on their similarity with the other ABCA members that also transport lipids,

GmABCAs could function in vivo as exporters, mediating transport out of the cytoplasm into the symbiosome. Further study with the isolation of SM vesicles and transport assays on these transporters would provide new insights into the direction of transport of GmABCA1 and

GmABCA2 across the SM.

NPF and OPT transporters in S. cerevisiae and other systems are electrochemical potential- driven transporters, catalyzing uptake of substrates by a cation: solute symport mechanism

(Daniel et al., 2006; Gomolplitinant and Saier, 2011; Hauser et al., 2001). It is likely that the

GmNPFs and GmYSL7 would transport substrates out of the symbiosome according to the direction of transport of other members, as well as the potential electrochemical differences across the SM. This places the substrate-binding sites of GmNPFs on the inside of the symbiosome. The inversion of the SM sidedness implies that the hypothesized substrate- binding sites of GmNPFs and GmYSL7 located on the inside of the symbiosome will be inverted to the outside of the inside-out SM vesicles. The export of peptides, out of the symbiosome (if it occurs as predicted) could then be assessed and peptide transport activity could also be measured. Inhibitors of proton motive force (Rudbeck et al., 1999) could be added to the transport assay to ascertain that GmNPFs function as proton symporters that link their transport to a proton gradient across the SM.

5.3 Alternative method for transport study

5.3.1 Reconstitution of transport proteins into liposomes

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The complexities of the SM, and most cell membranes have made it difficult to examine transport proteins in situ (Rigaud and Lévy, 2003). Membrane reconstitution of purified native proteins by insertion into an artificial lipid membrane (liposome) could serve as an alternative method for studying the structure and function of transport proteins (Geertsma et al., 2008;

Rigaud and Lévy, 2003). The method of membrane reconstitution was successfully utilized for

A. thaliana UDP–L-Rha/UDP–D-galactose transporter 1 (URGT1), which was shown to transport nucleotide sugar UDP–D-Gal when expressed in yeast (Rautengarten et al., 2014).

Following Rautengarten et al. (2014)’s novel method, transporters are firstly heterologously expressed in yeast. Yeast microsomal proteins are subsequently extracted and reconstituted into liposomes for transport assays. Proteoliposomes could be incubated with various fatty acids while nonincorporated fatty acids are removed by gel filtration. LC-MS/MS can be used to analyze the content of the proteoliposomes to determine the substrates transported, which could be compared to the empty vector control (Rautengarten et al., 2014). This method is useful to test a range of different substrates, instead of using growth complementation in yeast which may be substrate limited.

5.3.2 Heterologous expression in Xenopus oocytes

Electrophysiological studies or uptake studies of a radiolabeled substrate on cloned membrane transporters in Xenopus oocytes may serve as an effective alternative method for functional analysis as analysis in S. cerevisiae can be somewhat limiting (Miller and Zhou, 2000; Rentsch et al., 1998). Information on the activity of membrane transport proteins including substrate specificity, substrate affinity, sensitivity to inhibitors, stoichiometry and ion coupling can be

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obtained from the use of heterologous expression in Xenopus oocytes (Conte and Walker,

2012). A small number of nodule transport proteins have been successfully characterized using these methods, including nitrate transporter GmN70 (Vincill et al., 2005), aquaporins NOD26 and TIP1g (Gavrin et al., 2014; Rivers et al., 1997), and L. japonicus multidrug and toxic compound extrusion (MATE) protein 1 (LjMATE1) (Takanashi et al., 2013).

5.4 Uncovering physiological roles through metabolic profiling

SNF involves tight association and integration of metabolism between the host plant and the bacteroid. A number of studies have investigated the metabolite profiles of nodulated roots in response to rhizobia infection. These studies used gas chromatography coupled with mass spectrometry (GC/MS) or ultraperformance liquid chromatography-quadrupole time of flight- mass spectrometry (UPLC-QTOF-MS) to measure and analyze the metabolites. Numerous metabolites such as sugars, amino acids and organic acids were more enriched in nodules when compared to the corresponding uninoculated roots. (Barsch et al., 2006; Brechenmacher et al.,

2010; Colebatch et al., 2004; Desbrosses et al., 2005). The increased abundance of these metabolites was also reflected by the upregulation of the relevant genes in the nodules, for instance genes encoding proteins important for glycolysis and amino acid biosynthesis

(Colebatch et al., 2004). However, these studies extracted metabolites from whole nodules, revealing little information on specific metabolites located in different cells and their compartments in the nodules.

Our capability to isolate and extract different nodule compartments including the SS and the bacteroid means that metabolite changes occurring in these compartments can be analyzed.

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Moreover, soybean has larger nodules compared to other legumes such as L. japonicus and M. truncatula, allowing easier extraction of the symbiosomes and its constituents (Clarke et al.,

2014). The symbiosome from soybean also contains several bacteroids, which eases their separation from free bacteroids (White et al., 2007). It may also be valuable to examine metabolite changes in the cytoplasm of host infected cells, but cytoplasm would be difficult to isolate.

Examining the abundance of metabolites in normal nodules compared to RNAi nodules or nodules in which the transporter is overexpressed could provide an insight into the physiological roles these SM transporters play in symbiosis by addressing metabolic changes that occur as the result of changes in expression of the transporters. An RNAi approach would be particularly useful for importers, which mediate transport from the symbiosome to the cytoplasm, whereas generation of overexpression lines might be beneficial for exporters (that transport substrates out of the cytoplasm into the symbiosome). Both these assays would result in a build-up of the substrate of the transporter in symbiosomes from transgenic nodules.

The recent proteomic analysis of the soybean SM has identified vital steps involved in the nitrogen fixation (Clarke et al., 2015). Thus, analyzing the metabolome in response to reduce expression of these transporters can also tell us how metabolic and transport pathways are interconnected and can affect nitrogen fixation. Analyzing lipid and peptide containing metabolites should be the focus of this metabolomic study.

5.5 Stable mutants in other legumes

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It is unlikely that I would obtain mutants in soybean for functional analysis of these transporters.

The use of mutagenesis tools such as RNAi or gene editing techniques to generate transgenic soybeans maybe the best available option, however, stable lines are needed and A. rhizogenes- mediated hairy root transformation cannot provide this. Producing stable transgenic soybean is a challenging and time consuming task. Consequently, this makes it difficult to analyze the phenotype of a soybean mutant. It would be better to try and find mutants in legumes with smaller genomes. Generation of mutants have been useful in other legumes, but are not that beneficial in soybean due to the palaeopolyploidy nature of the genome and its relatively large genome size (Cooper et al., 2008; Schmutz et al., 2010). Thus, the use of other legumes might avoid problems of having to knockdown multiple genes with the same or overlapping functions to obtain a phenotype.

If orthologues of the soybean transporters are present in other mutant legumes, their mutant phenotype could therefore be analyzed. The recent availability of insertional mutagenesis tools in determinate legume (L. japonicus) and indeterminate legume (M. truncatula) using transposable elements (TE) has now made it possible to analyze phenotypes of soybean orthologues in these model legumes and may facilitate functional analysis of these soybean genes (Tadege et al., 2008; Urbański et al., 2012).

Orthologues of soybean candidates can be searched easily in Lotus (Verdier et al., 2013) and

Medicago (Benedito et al., 2010) gene expression atlases and also in LegumeIP (Li et al., 2012).

Once the soybean orthologue in L. japonicus or M. truncatula is identified, phenotypic analysis for any nitrogen fixation or nodule phenotypes can be carried out and complementation with

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soybean genes will be the next step. L. japonicus LORE1 mutants are available for GmABCA1,

GmABCA2, GmNPF8.6 and GmNPF1.2.

5.6 Conclusion

In this study, I have investigated and characterized two ABCA transporters, GmABCA1 and

GmABCA2 that are capable of transporting fatty acids in yeast and in planta, supporting the original hypothesis. This work has shed some light into the function of novel ABCA transporters that are constituents of the SM. However, further research is warranted on the role of fatty acids or lipids especially during the process of nitrogen fixation and as nodule matures, as well as the mechanism of which GmABCA1 and GmABCA2 transport fatty acid and their precise physiological roles in the symbiosis. This study has opened a new avenue for further characterization of fatty acid transport on the SM and the roles they might play.

The study has also demonstrated the role of GmNPF8.6 as a peptide transporter on the SM.

Evidence from its RNAi phenotype clearly shows that it is essential for symbiosis. Assessing its function in planta to confirm its role as a peptide transporter on the SM and to understand how this peptide transport is essential for symbiosis will be the next challenge.

GmYSL7 is also evidently important for symbiosis based on its RNAi phenotype. Whether like its homologue AtYSL7 that transports peptides, its physiological role for oligopeptide transport in the symbiosis will require further investigations. This will help us to understand how blocking this transport stops the development of multi-bacteroid symbiosomes and reduces nitrogen fixation.

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APPENDICES

174

Appendix 1 Plant ABC transporters characterised to date. Plant ABC Transporters whose functions/activities have been determined are listed according to the subfamily they reside.

SUBFAMILY GENE NAME SUBCELLULAR FUNCTION SUBSTRATE PLANT DOMAIN REFERENCE LOCALISATION SPP. Subfamily A AtABCA2 Kin recognition A. thaliana (TMD- Biedrzycki et al.

NBD)2 (2011) AtABCA9/ Endoplasmic Fatty acids Acyl-CoA, A. thaliana Kim et al. (2013) AtATH11 reticulum transport to ER free fatty acids Subfamily B AtABCB1/ Plasma Auxin export IAA A. thaliana (TMD- Geisler et al. AtPGP1/ membrane NBD)2 (2005), Murphy AtMDR1 et al. (2002), Petrášek et al. (2006) AtABCB4/ Plasma Auxin import IAA A. thaliana Cho et al. (2007), AtPGP4 membrane Kubeš et al. (2012), Santelia

et al. (2005), Terasaka et al. (2005) AtABCB14 Plasma Stomatal Malate A. thaliana Lee et al. (2008)

membrane regulation AtABCB19/ Plasma Auxin transport, IAA A. thaliana Christie et al. AtPGP19/ membrane cytoplasmic (2011), Murphy et AtMDR11 streaming and al. (2002), Noh et gravitropism of al. (2003), inflorescence Okamoto et al. (2016),

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stem Petrášek et al. (2006), Sukumar et al. (2013) AtABCB21/At Plasma Auxin import/ IAA A. thaliana Kamimoto et al.

PGP21 membrane export (2012) AtABCB23/ Mitochondrial Iron ‒ A. thaliana Chen et al. (2007)

AtATM1 membrane homeostasis AtABCB24/ Mitochondrial Iron ‒ A. thaliana Chen et al. (2007)

AtATM2 membrane homeostasis AtABCB25/ Mitochondrial Iron-sulfur Glutamine A. thaliana Bernard et al. AtATM3 membrane clusters export, synthetase- (2009), Bhuiyan heavy metal conjugated et al. (2011),

tolerance Cd(II)? Chen et al. (2007), Kim et al. (2006) AtABCB27/ Vacuolar Aluminium Chelated Al? A. thaliana Jaquinod et al. AtTAP2 membrane sequestration (2007), Larsen et al. (2007) LjABCB1 ‒ Auxin transport Auxin? L. Takanashi et al.

japonicus (2012) CjABCB1/ Plasma Berberine Berberine Coptis Shitan et al. CjMDR1 membrane import japonica (2003), Yazaki et al. (2001) CjABCB2 Plasma Alkaloid Alkaloid C. japonica Shitan et al.

membrane translocation (2013) OsABCB14 Plasma Auxin transport IAA O. sativa Xu et al. (2014)

membrane OsALS1 Tonoplast Aluminium Al O. sativa Huang et al.

detoxification (2012) ZmABCB/ Plasma Auxin export IAA Zea mays Knöller et al. ZmBR2 membrane (2010), Multani et al. (2003) 176

SbABCB1/ Plasma Auxin export IAA? Sorghum Multani et al.

SbDWF3 membrane bicolor (2003) TmABCB1 Plasma Berberine Berberine Thalictrum Shitan et al.

membrane transport minus (2015) TmABCB2 Plasma Berberine Berberine T. minus Shitan et al.

membrane transport (2015) Subfamily C AtABCC1/ Vacuolar Metal/metalloid Abscisic acid A. thaliana (TMD- Burla et al. AtMRP1 membrane detoxification, glucosyl ester NBD)2 (2013), Park et al. ABA-GE (2012), transport, Raichaudhuri arsenic stress (2016), Song et tolerance al. (2014b), Song et al. (2010) AtABCC2/ Vacuolar Metal/metalloid Abscisic acid A. thaliana Burla et al. AtMRP2 membrane detoxification, glucosyl ester, (2013), Frelet- ABA-GE chlorophyll Barrand et al. transport, catabolites (2008),Park et al.

vacuolar (2012), Song et transport of al. (2014b), Song chlorophyll et al. (2010), Su catabolites et al. (2007) AtABCC3 Vacuolar Phytochelatin- Cadmium A. thaliana Brunetti et al. membrane mediated Cd (2015) tolerance AtABCC5/ Plasma Guard cell Inositol A. thaliana Nagy et al. (2009) AtMRP5 membrane signaling- hexakisphosp stomata hate regulation, phytate storage ZmMRP3 Vacuolar Anthocyanin Anthocyanin Z. mays Goodman et al. membrane transport (2004)

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ZmMRP4 Vacuolar Anthocyanin Anthocyanin Z. mays Goodman et al. membrane transport (2004) VvABCC1 Tonoplast Anthocyanin Anthocyanin Vitis Francisco et al. transport vinifera (2013) OsABCC1 Tonoplast Arsenic Arsenic? O. sativa Song et al. detoxification (2014a) Subfamily D AtABCD1/ Peroxisomal β-oxidation β-oxidation A. thaliana (TMD- Bussell et al. CTS/ PXA1/ membrane products import substrates NBD)2 (2014), Footitt et PED3/PMP2/ to peroxisome, al. (2002), ACN2 production of Hayashi et al.

benzoylated (2002), Nyathi et metabolites in al. (2010), seeds Zolman et al. (2001) HvABCD1 ‒ β-oxidation β-oxidation Hordeum Mendiondo et al. products import substrates vulgare (2014) to peroxisome HvABCD2 ‒ β-oxidation β-oxidation H. vulgare Mendiondo et al. products import substrates (2014) to peroxisome Subfamily G AtABCG9 Plasma Vascular Pollen coat A. thaliana Full size, Choi et al. (2014), membrane development, NBD- Le Hir et al.

pollen coat TMD (2013) deposition AtABCG11/ Plasma Cuticular lipid Cutin and wax A. thaliana Bird et al. (2007),

AtWBC11 membrane secretion, organ components? fusion Lacey Samuels et prevention, al. (2010), Le Hir

suberin et al. (2013), Luo formation et al. (2007),

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Panikashvili et al. (2007), Panikashvili et al. (2010), Ukitsu et al. (2007) AtABCG12/ Plasma Cuticular lipid Wax? A. thaliana McFarlane et al.

AtWBC12 membrane export (2010) AtABCG13 ‒ Flower cuticle Cuticular A. thaliana Panikashvili et al. secretion, lipid? (2011)

patterning of petal AtABCG14 Plasma Cytokinin Cytokinin? A. thaliana Ko et al. (2014), membrane translocation to Le Hir et al.

shoot (2013), Zhang et al. (2014) AtABCG15 ‒ Post-meiotic Wax, cutin A. thaliana Qin et al. (2013) anther, pollen monomers? development AtABCG19/ Vacuolar Kanamycin Kanamycin? A. thaliana Mentewab and

AtWBC19 membrane resistance Stewart Jr (2005) AtABCG22 Plasma Guard cells ABA? A. thaliana Kuromori et al.

membrane functions (2011b) AtABCG25 Plasma ABA export ABA A. thaliana Kuromori et al. membrane (2010), Kuromori et al. (2016) AtABCG26 Plasma Pollen exine Sporopollenin A. thaliana Choi et al. (2011), membrane formationa precursors? Kuromori et al.

(2011a),

Quilichini et al.

(2010)

179

AtABCG31 Plasma Pollen coat Pollen coat A. thaliana Choi et al. (2014)

membrane deposition AvABCG1 Cell membrane Aluminium ‒ Andropogo Ezaki et al. tolerance n virginicus (2015) L. OsABCG5/ ‒ Involved in ‒ O. sativa Matsuda et al. OsRCN1 accumulation of (2014), Matsuda ABA, essential et al. (2016), and Yasuno et al. nonessential (2009)

minerals, Na/K homeostasis, and salt tolerance, root shoot branching OsABCG15 ‒ Pollen exine Sporopollenin O. sativa Niu et al. (2013),

formation ? Qin et al. (2013) StABCG1 Plasma Suberin ‒ Solanum Landgraf et al.

membrane formation tuberosum (2014) GpWBC1 Plasma Cotton fiber ‒ Gossypium Zhu et al. (2003)

membrane elongation hirsutum NtWBC1 Plasma Possible role in ‒ Nicotiana Otsu et al. (2004)

membrane? reproduction tabacum MtSTR1 Periarbuscular Arbuscule ‒ Medicago Zhang et al.

membrane development truncatula (2010) MtSTR2 Periarbuscular Arbuscule ‒ M. Zhang et al.

membrane development truncatula (2010) OsSTR1 Periarbuscular Mycorrhizal ‒ O. sativa Gutjahr et al. membrane arbuscule (2012) formation OsSTR2 Periarbuscular Mycorrhizal ‒ O. sativa Gutjahr et al.

membrane arbuscule (2012)

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formation formation OsABCG5/ Plasma Suberization of ‒ O. sativa Shiono et al.

OsRCN1 membrane roots (2014) GmPDR12 ‒ Involved in ‒ G. max Eichhorn et al. salicylic acid (2006)

signaling pathway AtABCG29/ Plasma Monolignol Monolignol A. thaliana (NBD- Alejandro et al. AtPDR1 membrane transport, lignin TMD)2 (2012) biosythesis AtABCG30/ ‒ Root exudates ‒ A. thaliana Badri et al. AtPDR2 (2008), Badri et al. (2009) AtABCG31 Plasma Pollen coat ‒ A. thaliana Choi et al. (2014)

membrane deposition AtABCG32 Plasma Cuticle Cutin A. thaliana Bessire et al.

membrane formation precursors? (2011) AtABCG36/ Plasma Pathogen Cadmium, A. thaliana Kim et al. (2007), AtPDR8 membrane defense, IBA Kim et al. (2010), cadmium Kobae et al. resistance, (2006), Stein et drought and salt al. (2006), Strader stress and Bartel (2009) resistance, auxin homeostasis AtABCG37 Plasma Auxin precursor Indole-3- A. thaliana Fourcroy et al. membrane exporter, butyric acid (2014), Růžička

secretion of (IBA), et al. (2010) scopoletin scopoletin AtABCG30/ Plasma Paraquat Paraquat? A. thaliana Xi et al. (2012)

AtPDR11 membrane transport

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AtABCG40/ Plasma ABA import ABA A. thaliana Kang et al. (2010)

AtPDR12 membrane MtABCG10 Plasma Modulate ‒ Medicago Banasiak et al. membrane isoflavonoid truncatula (2013) levels HvABCG31 ‒ Leaf water ‒ Hordeum Chen et al. (2011)

conservation vulgare SpTUR2/ Sp Plasma Antifungal Sclareol? Spirodela Van Den Brûle et PDR5 membrane diterpene polyrrhiza al. (2002) sclareol resistance NtPDR1 Plasma Diterpene Diterpenes N. tabacum Crouzet et al. membrane transport (2013) NtPDR6 ‒ Regulate shoot ‒ N. tabacum Xie et al. (2015) branching process NpABCA1/ Plasma Antifungal Terpenoid Nicotiana Jasiński et al. NpPDR1 membrane terpenoid sclareolide plumbagini (2001), Stukkens secretion, folia et al. (2005) pathogen defense LjABCG1 Plasma Plant-microbe ‒ L. Sugiyama et al. membrane interactions japonicus (2015) PaPDR1 Apical membrane Strigolactone Strigolactone Petunia Sasse et al. (2015) transport axillaris Subfamily I AtABCI14/ Chloroplast Lipid formation Phosphatidic A. thaliana TMD Xu et al. (2005)

AtTGD1 membrane acid? AtABCI15/ Chloroplast Lipid Phosphatidic A. thaliana Substrate Awai et al. AtTGD2 membrane biosynthesis acid? binding (2006), Roston et protein al. (2011)

182

AtABCI13/ Chloroplast Lipid Phosphatidic A. thaliana NBD Lu et al. (2007) AtTGD3 membrane biosynthesis acid?

AtABCI16/ Plasma Aluminium Al? A. thaliana TMD Larsen et al. AtALS3 membrane tolerance (1997), Larsen et

al. (2005), Sawaki et al. (2009) OsSTAR1 Vesicle Aluminium UDP-Glucose O. sativa NBD Huang et al.

membrane? tolerance (2009) OsSTAR2 Vesicle Aluminium UDP-Glucose O. sativa TMD Huang et al.

membrane? tolerance (2009) AtSTAR1 ‒ Aluminium UDP- A. thaliana NBD Huang et al.

detoxification Glucose? (2010)

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Appendix 2 Melting curves for four different amplicons – GmABCA1, GmABCA2,

GmNPF8.6 and GmUBI3. The single peaks observed for all transcripts demonstrated that only single amplicons were amplified in the reactions and hence confirmed that the primers used were specific for each transcript.

GmABCA1 melting curve

GmABCA2 melting curve

184

GmUBI3 melting curve

185

0.20 0.18 0.177 0.16

0.14 0.12

0.10 02

0.08 02

- 5.41E

0.06 -

0.04 3.63E 2.11E

EXPRESSION RELATIVE 0.02 0.00 LEAF STEM ROOT NODULE TISSUE Appendix 3 Tissue-specific expression of GmABCA2 across four different tissues (leaf, stem, root and nodule) at 35 DPI. The assay was performed by Clarke (2013).

186

Appendix 4 GUS assay of GmABCA2 promoter

Introduction

To examine the location in which GmABCA1 and GmABCA2 promoter are active within the nodule, the histochemical localization of GUS reporter activity fused to the promoter was investigated. GmABCA1 promoter was unable to be cloned, thus a GUS assay for GmABCA1 promoter could not be attempted. GmABCA2 promoter managed to be cloned and the construct was expressed in soybean nodules twice, and the same results were obtained.

Materials and methods

Amplification and cloning

Both GmABCA1 and GmABCA2 promoters were amplified and cloned as outlined in section

2.4 based on sequence prediction in the soybean genome version 1.1. Promoters were amplified

2 kb upstream the start codon. Oligopeptides used for the amplifications are as listed in Table

1. Only GmABCA2 promoter was successfully cloned. Entry and expression clones were generated by Gateway® BP and LR recombination reactions, respectively. pENTR clones were recombined into pKGW-GGRR (Figure 1) for GUS assays.

Soybean transformation and growth

Soybean transformation and growth were carried out as detailed in section 2.11.

187

Table 1 Primer sequences for Gateway® cloning of GmABCA1 and GmABCA2 promoters. The attB2 and attB2 recombination sites are underlined. DNA FRAGMENT FORWARD PRIMER REVERSE PRIMER SEQUENCE (5'→3') SEQUENCE (5'→3') GmABCA1 promoter GGGGACAAGTTTGTACAAA GGGGACCACTTTGTACAAG AAAGCAGGCTTAATGAACA AAAGCTGGGTAGGCTTTTG TATGTCCGCACGA TGTGGTCTGCTC GmABCA2 promoter GGGGACAAGTTTGTACAAA GGGGACCACTTTGTACAAG AAAGCAGGCTTAATCCGGT AAAGCTGGGTACGCCATGC CCTCACTTTCACC TATGCTGTCTGTG

Figure 1 Plasmid map of destination vector, pKGW-GGRR (Gavrin et al., 2014) used for GUS study. Abbreviations: ori – origin of replication, RB – right border, LB – left border, CmR – chloramphenicol resistance gene, ccdB – ccdB death gene, EGFP – enhanced GFP, GUS – β-glucuronidase gene, T35S – terminator of 35S, Ubi promoter – Ubiquitin promoter, DsRED – Discosoma sp. red fluorescent protein, NOS terminator – nopaline synthase terminator, and Sm/SpR – spectinomycin and streptomycin resistance genes. Vector map created using SnapGene® software.

GUS assay

Nodules were harvested at 32 dpi, and were screened for transformed nodules as described in section 2.11.4. Transformed nodules were embedded in 8% agar for sectioning, with the following vibratome settings: 1 mm sec-1 (speed), 1 mm (amplitude), and 50 µm (section thickness). Sections were placed in GUS buffer (100 mg/ml X-GLUC (5-bromo-4-chloro-3- indolyl-β-D-glucuronide cyclohexylammonium salt), 0.061 M NaH2PO4 (pH 7), 0.039 M

NaH2PO4 (pH 7), 0.01 M EDTA, 0.1% Triton X-100, 1 mM potassium ferrocyanide, 1 mM potassium ferricyanide) and vacuum infiltrated for 30 min. Sections were washed in water and de-stained in 20% (v/v) ethanol for 20 min. Sections were visualized using a Fluorescence

Stereo Microscope Leica M205 FA under a bright field light.

Results

A 2 kb region upstream of the start codon of GmABCA2 was cloned and expressed in soybean plants as a fusion with a GUS/GFP coding region for histochemical analysis of GUS activity.

A GUS assay on the transgenic nodule sections showed no detectable GUS activity of the

GmABCA2 promoter in the infected cells and the uninfected cells (Figure 2A & B). However, a low GUS signal was detected in the vascular bundles (Figure 2A). The same transgenic nodule section expressing the GmABCA2 promoter showed GFP expression in the infected cells

(Figure 2A & B).

188

A V B

ICs

B UC

ICs

Figure 2 Analysis of GUS activity of GmABCA2 promoter in mature nodule at 32 DAI. (A, B) No GUS activity was detected in the infected cells (images to the left), but some appeared in the vascular bundles (VB). Enlargement of images from the left (middle images). GFP fluorescent in the infected cells (images to the right). Abbreviations: ICs – infected cells, UC – Uninfected cells. Scale bars represent 100 µm (A) and 250 µm (B).

Discussion

No detectable GUS activity was observed in infected cells or uninfected cells in the assay to test the activity of the GmABCA2 promoter, although a weak GUS activity was detected in the vascular bundles (Figure 2). Other soybean promoters tested for GUS activity also showed

GUS staining in the vascular bundles (Clarke et al., 2015). The GUS gene in the assay was fused to GFP and GFP expression was observed in the infected cells of the same transgenic nodule sections (Figure 2). GFP fluorescence was not examined in untransformed nodule sections, so it cannot be ruled out that the GFP signals detected is not caused by inherent autofluorescence.

If the fluorescence observed is autofluorescence, the absence of GUS activity of GmABCA2 promoter may mean that the 2 kb region upstream of the initiation codon of GmABCA2 is insufficient to drive expression of the β-glucuronidase gene. Important regulatory elements may be positioned upstream (or downstream) of the region tested and a larger promoter region may need to be tested. For other SM proteins, the 2 kb region upstream the start codon was sufficient for detectable GUS activity (Clarke et al., 2015; Appendix 5) of transcripts such as

GmNPF5.24 (Clarke et al., 2015) with a similar expression level to that seen for these

GmABCA genes. For GUS analyses of ABC transporters LjABCB1 and PdPDR1 the 1.7 kb and 1.8 kb region uptream the start codon were adequate to drive expression of β-glucuronidase

(Kretzschmar et al., 2012; Takanashi et al., 2012).

GmABCA1 promoter was unable to be cloned after multiple attempts. The 2 kb region upstream the initiation codon of GmABCA1 was different between soybean genome assembly version

189

1.1 and the newly released genome assembly version 2.0. A 750 bp region was absent in the previous version 1.1. As the result, new primers sets were designed based on genome prediction in version 2.0, but the amplified product still could not be cloned, even after a different cloning system was used.

190

A B C

D E

Appendix 5 GUS expressions driven by different GmNPF and GmYSL promoters in 30 DAI G. max nodules. Around 2 kb region upstream the start codon in (A) GmNPF1.2, (B) GmNPF5.24, (C) GmNPF5.25, (D) GmNPF8.6, and (E) GmYSL7 (Chen, 2012) were fused to GUS reporter gene and subsequently expressed in G. max. GUS staining of the cells indicated that the promoter is active within that cell. GmNPF5.24 and GmNPF5.25 figures were published in Clarke et al. (2015). GmNPF1.2 and GmNPF8.6 promoter work were unpublished results of Alex Gavrin and Penelope Smith (The University of Sydney). All scale bars represent 500 µm, except in (E) which represent 100 µm.

191

GFP-GmNPF5.24

GFP-GmNPF5.25

Appendix 6 GFP localization of GmNPF5.24 and GmNPF5.25. These images were published in Clarke et al. (2015). Scale bars represent 20 µm.

192

A B

Col-0 Col-0 Appendix 7 Representatives of Basta selection plates to select for T3 lines that are homozygous for the transgenes. At least 40 T3 seeds and 7 WT Col-0 seeds (boxed in red) were placed on half MS agar supplemented with 5 mg/l Basta. (A) All seedlings were resistant and grew on the Basta selection plate except for the Col-0 negative control, demonstrating that the T3 line is homozygous for the transgene. (B) Some seedlings did not survive indicating that the T3 line is heterozygous for the transgene.

193

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