Katholieke Universiteit Leuven · 2008-02-01 · op partikel beschieting (Particle...

ISBN 978-90-8826-044-5 Wettelijk depot D/2008/11.109/2

Katholieke Universiteit Leuven

Faculteit Bio-ingenieurswetenschappen

Departement Biosystemen

Afdeling Plantenbiotechniek

DISSERTATIONES DE AGRICULTURA

An Improved Agrobacterium-Mediated Transformation Method for Banana and Plantain

(Musa spp.)

Promotor:

Prof. R. Swennen, K.U.Leuven

Co-promotor:

Dr. L. Sági, K.U.Leuven

Leden van de examencommissie:

Prof. E. Decuypere, voorzitter

Prof. W. Keulemans, K.U.Leuven

Dr. M. De Bolle, K.U.Leuven

Prof. G. Angenon, V.U.B

Dr. W. Tushemereirwe, NARO, Uganda

Proefschrift voorgedragen tot het

behalen van de graad van

Doctor in de

Bio-ingenieurswetenschappen

door

Geofrey Arinaitwe

FEBRUARY 2008

This thesis is gratefully dedicated to my late mother Glady Komushoro, my wife Caroline

Asasira and my daughter Leticia Ayesiga

Acknowledgement

Acknowledgement

I would like to express my gratitude to all those who gave me the possibility to complete

this PhD thesis. I want to thank Prof. R. Swennen of the Laboratory of Tropical Crop

Improvement, Katholieke Universiteit Leuven (K.U.Leuven), Belgium, for allowing me

to conduct the research work in his laboratory and for his academic guidance throughout

my training at K.U.Leuven. Furthermore, I would also like to thank the former Director

of INIBAP (now Bioversity International), Dr. E. Frison, who spear-headed the Uganda

Banana Biotechnology Project under which this research was started.

I am deeply indebted to my immediate supervisor Dr. László Sági for his technical skills,

advice and encouragements. He changed my way to look at things and opened my eyes to

experimentation in plant molecular biology and genetic engineering. Lots of thanks go to

Dr. Remy Serge as well whose stimulating suggestions and encouragement helped me all

the time during the research and thesis write up.

This work would not have been possible without the availability of embryogenic cell

suspension. As this is a very time consuming process, I want to thank especially

Hannelore Strosse, Bart Panis, Francois Côte, Edwige André and Karen Reyniers for

making that material available.

My former colleagues from the Laboratory of Tropical Crop Improvement supported me

in my research work. Special thanks to Els Thiry, Saskia Windelinckx, Wim Dillemans,

Bert Coemans, and Efren Santos for the social and scientific interactions. Marleen

Stockmans and Suzy Voets made my travels possible and cleared all other administrative

issues. Also, I appreciate the company of other technical and scientific staff from the lab:

Alex Henneau, Ronald Boogaerts, Ines van den Houwe and Els Kempenaers.

I can not forget Dr. W. Tushemereirwe of the National Agriculture Research

Organization (NARO) and the staff at Kawanda Agricultural Research Institute (KARI).

Your support whenever I came to work at KARI was great. I deeply thank Prof. P. R.

Rubaihayo, who always reminded me to go for advanced studies.

I would like to give my special thanks to my wife Asasira Caroline and my children

Ayesiga Leticia and Ainebyona Niels for their patience, encouragement and allowing me

to be away for such a long time. Their great love enabled me to complete this work. My

Acknowledgement

ii

late son Mutatina Louis, who never lived long enough to see me back home, was a source

of joy in our family.

The research results presented in this thesis were generated within the framework of a

collaborative project ‘Novel Approaches to the Improvement of Banana Production in

East Africa – application of biotechnological methodologies’. This collaborative research

aimed at the development of Ugandan banana varieties with enhanced resistance to Black

Sigatoka, nematodes and banana weevils; and the development of embryogenic cell

suspensions in East African highland bananas. The development of banana lines with

these traits needs extensive research experience, considerable expertise, and permanent

technical support. This was achieved through collaboration with research organisations

that have vast experience in banana genetic improvement via both molecular and classical

breeding. The role played by these research organisations range from technical training,

provision of research materials, coordination, monitoring and evaluation. The main

collaborators were K.U.Leuven (Belgium), Makerere University (Kampala), Bioversity

International (France), NARO (Uganda), IITA (Uganda) and CIRAD (France).

The research work that involved the integration of transgenes (Cht-2, Cht-3 and Rs-afp2)

alone and in combination was supported by the Belgian Technical Co-operation

(BTC/CTB) via a scholarship to me through the Banana Biotechnology Project initiated

and additionally funded by the Government of Uganda. This project was coordinated by

the International Network for the Improvement of Bananas and Plantains (INIBAP), now

Bioversity International. This work contributes to the achievement of the aims and

objectives of the Program for the Modernization of Agriculture (PMA) that is politically

backed by Government of Uganda. The genetically modified banana plants developed

during the course of this PhD thesis research are now being field tested thanks to support

of ABSP-II and with support of USAID.

Finally I want to thank especially Dr. Frank Shotkoski, director of ABSP-II, and Dr.

Richard Markham, Bioversity International, for their coordinating role in bringing all

these partners together and finding additional financial resources.

List of acronyms

iii

List of acronyms 2,4-D 2,4-Dichlorophenoxyacetic acid AFP Anti-fungal peptide AMP Anti-microbial peptide AmT Agrobacterium-mediated transformation AS Acetosyringone bp Base pair BSV Banana Streak Virus CaMV Cauliflower Mosaic Virus cDNA Complementary DNA CHAPS 3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate Cht-2 Rice chitinase gene isolated from cDNA library Cht-3 Rice chitinase gene isolated from genomic DNA library CIRAD Centre de coopération Internationale en Recherche Agronomique pour le

Développement (France) CSPD Alkaline phosphatase substrate: Disodium 3-(4-methoxyspiro{1,2-dioxetane-

3,2´-(5'-chloro)tricyclo[3.3.1.13,7]decan}-4-yl)phenyl phosphate DNA Deoxyribonucleic acid EAHB East African Highland Banana EC Embryogenic Colony ECS Embryogenic Cell Suspension EDTA Ethylenediaminetetraacetic acid ET Ethylene FAOSTAT Online FAO Statistical Database containing statistics on agriculture, nutrition,

fisheries, forestry, food aid, land use and population. FHIA Honduran agricultural research foundation GFP Green fluorescent protein from jellyfish Aequorea victoria gfp GFP gene GM Banana cultivar ‘Gros Michel’ GN Banana cultivar ‘Grand Naine’ uidA β-glucuronidase gene from E. coli uidAINT Intron-interrupted uidA gene hpt Hygromycin phosphotransferase gene INIBAP International Network for the Improvement of Bananas and Plantains; currently,

List of acronyms

iv

Bioversity International JA Jasmonic acid kb Kilobase LB Left T-DNA border sequence M2 Cell suspension culture medium (immature flower method) MPCR Multiplex PCR mRNA Messenger RNA MS Medium after Murashige and Skoog NARO National Agricultural Research Organization (Uganda) nptII Neomycin phosphotransferase gene OD Optical Density OE Banana cultivar ‘Obino l’Ewai’ OR Banana cultivar ‘Orishele’ P35S CaMV 35S RNA promoter PCR Polymerase Chain Reaction PmT Particle bombardment-mediated transformation Pnos Nopaline synthase gene promoter PR Pathogenesis-related PUbi Maize polyubiquitin promoter and first intron RB Right T-DNA border sequence RD1 Somatic embryo induction medium (scalp method) RD2 Somatic embryo germination medium (scalp method) RNA Ribonucleic acid Rs-AFP2 AFP from Raphanus sativus Rs-afp2 Raphanus sativus AFP gene RT-PCR Reverse Transcription Polymerase Chain Reaction SA Salicylic acid SCV Settled Cell Volume SDS Sodium dodecyl sulphate sgfpS65T Codon optimised synthetic gfp gene with a mutation of serine to threonine at

position 65 SPD Spermidine SSC Saline Sodium Citrate Taq Thermus aquaticus bacterium TCaMV CaMV 35S RNA poly(A) region

List of acronyms

v

T-DNA Transferred DNA TGE Transient Gene Expression TGFPE Transient GFP expression

THP Banana cultivar ‘Three Hand Planty’

Tmas Mannopine synthase gene poly(A) region

Tnos Nopaline synthase gene poly(A) region

Tocs Octopine synthase gene poly(A) region

W Banana cultivar ‘Williams’

X-Gluc 5-bromo-4-chloro-3-indolyl-β-D-glucuronide

ZZ Cell suspension culture medium (scalp method)

Samenvatting

vii

Samenvatting

In Oeganda wordt banaan “Matooke” genoemd wat eigenlijk voedsel betekent. Meer dan

8 miljoen Oegandezen zijn afhankelijk van banaan voor voedsel, inkomen en werk. De

productie van banaan in Oeganda vermindert evenwel door de volgende ziekten en

plagen: Black Sigatoka Disease (BSD), plant parasitaire nematoden, de bananen

snuitkever, Fusarium verwelkingsziekte, en heel recentelijk, Banana Bacterial Wilt. Black

Sigatoka resistente hybriden van Oost-Afrikaanse Hoogland banaanvariëteiten (East

African Highland Banana cultivars (EAHB)) werden reeds ontwikkeld, maar de smaak en

na-oogst kenmerken waren veranderd. Via genetische manipulatie van EAHB

banaanvariëteiten kan echter resistentie ingebouwd worden zonder dat er veranderingen

optreden in smaak en culinaire kenmerken. Om dit te kunnen verwezenlijken zijn

embryogene celsuspensies (ECS) nodig en deze werden al ontwikkeld voor niet EAHB

banaanvariëteiten. Technologieën voor genetische manipulatie van banaan vallen terug

op partikel beschieting (Particle bombardment-mediated Transformation (PmT)) en

Agrobacterium-gemediëerde transformatie (Agrobacterium-mediated Transformation

(AmT)). Ten slotte werden er genen geïdentificeerd aan de Katholieke Universiteit

Leuven (K.U.Leuven) die mogelijk resistentie kunnen geven tegen BSD.

In deze thesis wordt een geoptimaliseerde Agrobacterium-gen transfer methode

beschreven. Vervolgens wordt de genetische transformatie van banaan met de rijst

chitinase genen Cht-2 and Cht-3 gerapporteerd, alsook de co-integratie van die chitinase

genen en Rs-AFP2 gen coderend voor het radijs defensine (antifungale eiwit) in

verschillende banaanvariëteiten. De gebruikte reportergenen zijn uidA en sgfpS65T, en de

Agrobacterium stammen EHA 101, AGLO, en EHA 105 werden getest. De gecloneerde

rijst chitinase genen waren aanwezig in de binaire vectoren pBI333-EN4-RCC2 en

pBI333-EN4-RCG3. Het “Green fluorescent protein” coderend gen sgfpS65T was

aanwezig in pBIN Ubi1-sgfpS65T, uidA in FAJ3000 en Rs-AFP2 in FAJ3494. De zes

banaanvariëteiten die getest werden waren ´Grand Naine´ (AAA), ´Gros Michel´ (AAA),

Óbino lÉwai´ (AAB), Órishele´ (AAB), ´Three Hand Planty’ (AAB) en ‘Williams’

(AAA).

Samenvatting

viii

Na vergelijking werd AmT superieur bevonden aan PmT. De resultaten toonden ook aan

dat de regeneratie capaciteit van transgene lijnen variëteit afhankelijk is. Er werd geen

correlatie gevonden tussen transiënte en stabiele genexpressie. Ten slotte werd

vastgesteld dat het aantal transgene scheuten per lijn variëteit afhankelijk is.

Verschillende parameters beïnvloeden de transformatie efficiëntie. Ten eerste is de

leeftijd van een ECS bepalend en idealiter moet de transformatie 4 tot 6 dagen na de

laatste subcultuur plaats vinden. Ten tweede werd de transformatie efficiëntie zeer sterk

verhoogd door een infectieduur van 8 uren. Ten derde werd de transformatie efficiëntie

verbeterd door het ECS volume tijdens de co-cultivatie te verlagen van het standaard

volume van 1200 µL naar 100 tot 300µL. Ten slotte werd aangetoond dat de vierde

parameter, namelijk het polyamine spermidine de scheut regeneratie erg bevorderde

alhoewel dit variëteit afhankelijk was.

De A-mT toegepast op de variëteiten ‘Grand Naine´ en ´Gros Michel´ leverde transgene

lijnen op met minimum 1 tot 4 integratieplaatsen. ´Gros Michel´ had een lager aantal

integratieplaatsen en aantal kopijen (1 tot 5) terwijl in ‘Grand Naine’ het transgen tot 7

maal kon ingebouwd worden. In beide variëteiten werd het transgen Cht-3 in een hoger

aantal ingebouwd dan het transgen Cht-2. Co-transformatie met één Agrobacterium stam

dat twee verschillende binaire vectoren bevatte met verschillende antibioticum

selectiemerkergenen, was zeer efficiënt. De transformatie frequenties geanalyseerd via

PCR en multiplex PCR, toonden de aanwezigheid aan van de twee verschillende

seletiemerkergenen in 90% tot 100% van de geanalyseerde lijnen indien selectie met

beide antibiotica werd doorgevoerd. Wanneer één selectief agens werd gebruikt,

variëerde de transformatie efficiëntie tussen 70% en 90%. ECSs van de variëteiten ‘Three

Hand Planty’ en ‘Orishele’ vertoonden een verschillende gevoeligheid tegenover

antibioticum selectie, waarbij al de geco-transformeerde ‘Orishele’ stalen afstierven op

het medium met de twee verschillende selectieve agentia. Integratie profielen bepaald via

Southern hybridizatie bevestigden dat de geregenereerde lijnen geco-tranformeerd waren.

Co-integratie van twee verschillende T-DNAs vertaalde zich niet in een hoger aantal

Samenvatting

ix

integratieplaatsen, wat doet veronderstellen dat het aantal integratieplaatsen niet

beïnvloed wordt door het type T-DNA dat geïntegreerd werd.

Momenteel worden 26 transgene lijnen, die de twee rijst chitinase genen bevatten, in het

veld getest tegen BSD. Tolerante lijnen zullen geselecteerd en geëvalueerd worden naar

productie toe. Deze veldresultaten zullen dan dienen als basis voor de ontwikkeling van

EAHB banaanvariëteiten met rijst chitinase genen voor een verhoogde BSD tolerantie

Summary

xi

Summary

Banana (commonly known as Matooke) is synonymous to food in Uganda. Over eight

million Ugandans depend on bananas as a source of food, income and employment.

Banana production has declined in Uganda due to biotic constraints. These are Black

Sigatoka Disease (BSD), plant parasitic nematodes, banana weevil, Fusarium wilt, and,

more recently, Banana Bacterial Wilt (BBW). Black Sigatoka resistant hybrids of East

African Highland Banana cultivars (EAHB) have been reported, but their culinary

attributes were inferior to the landraces. However, resistance in EAHB banana cultivars

could be improved via genetic engineering without altering their desirable culinary

characteristics. Technologies for banana genetic modification including tissue culture and

Embryogenic Cell Suspension (ECS) have been reported in non EAHB banana cultivars.

Genetic transformation systems optimised for banana are particle bombardment and

Agrobacterium-mediated gene transfer. Genes with potential resistance against BSD have

also been tested at the Katholieke Universiteit Leuven (K.U.Leuven).

This thesis reports on the optimisation of the Agrobacterium-mediated transformation

system, the transformation of banana with rice chitinase genes Cht-2 and Cht-3, and the

co-integration of these chitinase genes with a defensin (Rs-afp2) gene in several banana

cultivars. Reporter genes used were uidA and sgfpS65T. The three Agrobacterium strains

tested were EHA 101, EHA 105 and AGLO. Six banana cultivars were used and these

included ‘Grand Naine’ (AAA), ‘Gros Michel’ (AAA), ‘Obino l’Ewai’ (AAB), ‘Orishele’

(AAB), ‘Three Hand Planty’ (AAB) and ‘Williams’ (AAA).

The performance of the Agrobacterium-mediated transformation system (AmT) was

compared with the particle bombardment-mediated transformation system (PmT). The

AmT was found to be superior. Results indicated also that ECS competence and their

regenerability were cultivar dependent. Moreover there was no correlation between

transient and stable gene expression. The number of transgenic shoots regenerated

depended on the cultivar.

Several parameters were shown to affect the transformation efficiency. First, an ECS age

of 4 to 6 days after the last subculture was optimal. Second, an increased infection length

of up to 8 h dramatically improved transformation efficiency. Third, ECS volume, during

Summary

xii

cocultivation, of 100 to 300 µL had higher transformation frequencies than the frequently

used 1200 µL. The fourth parameter, the polyamine spermidine also contributed through

increased shoot regeneration, though its effects were cultivar dependant.

AmT of the cultivars ‘Grand Naine’ and ‘Gros Michel’ resulted in transformed lines with

integration loci varying from 1 to 4. In general ‘Gros Michel’ showed a lower number of

integration loci and copy numbers (1 to 5) while in ‘Grand Naine’ up to 7 integrated

transgene copy numbers were observed. In both cultivars the transgene Cht-3 was

integrated in more copy numbers than the transgene Cht-2. Co-transformation, using one

strain of Agrobacterium harbouring two different binary vectors was highly efficient (up

to 100%). Transformation frequencies, based on PCR and MPCR analyses, showed a

success rate of 90% to 100% with two different selective agents (antibiotics). When one

selection agent was used, transformation frequencies ranged between 70% and 90%.

ECSs of the two cultivars, ‘Three Hand Planty’ and ‘Orishele’, showed different

sensitivities towards antibiotic selection pressure, with all cotransformed ECSs of

‘Orishele’ dying on medium supplemented with the two selective agents. Integration

profiles as detected by Southern blot analysis, confirmed that the regenerated lines were

actually cotransformants. The cointegration of two different T-DNAs did not increase the

number of integration loci, implying that the number of integration loci were not

influenced by the variation or sources of T-DNAs integrated.

To assess whether rice chitinases could protect banana against BSD, 26 lines are now

being field tested in Uganda. Tolerant lines will be selected and further assessed for use

in banana production. Based on field evaluation results, rice chitinases genes will be

introduced into EAHB cultivars.

Table of contents

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Table of contents

ACKNOWLEDGEMENT..........................................................................................................................I

LIST OF ACRONYMS ...........................................................................................................................III

ABSTRACT............................................................................................................................................ VII

CHAPTER 1. GENERAL INTRODUCTION......................................................................................... 1

1.1. IMPORTANCE OF BANANA IN EAST AFRICA .......................................................................................... 1 1.2 BANANA PRODUCTION CONSTRAINTS.................................................................................................... 1 1.3. BANANA GENETIC IMPROVEMENT THROUGH CLASSICAL BREEDING..................................................... 2 1.4. IMPROVEMENT OF BANANA VIA GENETIC ENGINEERING........................................................................ 3 1.5. POTENTIAL GENES FOR FUNGAL CONTROL IN BANANA......................................................................... 4 1.6. RESEARCH OBJECTIVES ........................................................................................................................ 5 1.7. OUTLINE OF THIS THESIS ...................................................................................................................... 5

CHAPTER 2. LITERATURE REVIEW ................................................................................................. 7

2.1. FUNGAL DISEASES OF BANANAS........................................................................................................... 7 2.1.1. Black Sigatoka disease ................................................................................................................ 7

2.1.1.1. The pathogen ........................................................................................................................................7 2.1.1.2. Infection process and diseases development process ............................................................................8 2.1.1.3. Symptoms .............................................................................................................................................9 2.1.1.4. Distribution of Black Sigatoka disease ...............................................................................................10 2.1.1.5. Management of Black Sigatoka disease..............................................................................................11 2.1.1.6. Mechanisms of resistance against Black Sigatoka ..............................................................................12

2.2. PLANT GENETIC TRANSFORMATION AND PLANT REGENERATION........................................................ 13 2.2.1. Agrobacterium-mediated transformation .................................................................................. 13

2.2.1.1 Virulence gene expression ...................................................................................................................14 2.2.1.2. T-DNA transportation into plant cell ..................................................................................................15 2.2.1.3. Intracellular transport and T-DNA integration into plant cell genome ...............................................15 2.2.1.4. The structure of integration sites in plants ..........................................................................................16 2.2.1.5. Factors influencing Agrobacterium-mediated transformation.............................................................17 2.2.1.6. Agrobacterium-mediated co-transformation .......................................................................................18

2.2.2. Direct gene transfer................................................................................................................... 19 2.2.3. Polyamines and plant regeneration........................................................................................... 19

2.3. PLANT RESPONSES TO PATHOGEN INFECTION ..................................................................................... 20 2.3.1. Plant-pathogen interactions ...................................................................................................... 20

2.3.1.1. Pathogen recognition ..........................................................................................................................21 2.3.1.2. Signal transduction .............................................................................................................................23

2.3.2. Induced defence responses ........................................................................................................ 25 2.4. POTENTIAL GENETIC ENGINEERING STRATEGIES ................................................................................ 25

2.4.1. Hydrolytic enzymes.................................................................................................................... 26 2.4.1.1. Plant chitinases ...................................................................................................................................27

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2.4.1.1.1. Class I and II chitinases....................................................................................................................28 2.4.1.1.2. Class III chitinases ...........................................................................................................................33 2.4.1.1.3. Class IV-VII chitinases ....................................................................................................................35 2.4.1.1.4. Functions of plant chitinases ............................................................................................................35 2.4.1.2. Application of chitinases in plant genetic engineering........................................................................36 2.4.1.3. Rice chitinase genes ............................................................................................................................37 2.4.1.4. Resistance based on rice chitinases .....................................................................................................38

2.4.2. Plant defensins .......................................................................................................................... 40 2.4.2.1. Radish defensin (Rs-AFP2).................................................................................................................41 2.4.2.2. Plant genetic engineering with plant defensins ...................................................................................42

2.5. RESISTANCE THROUGH COMBINATORIAL EXPRESSION OF PLANT DEFENCE GENES ............................. 43 2.6. GENETIC MODIFICATION OF BANANA FOR BLACK SIGATOKA RESISTANCE ........................................ 44 2.7. CO-TRANSFORMATION IN BANANA .................................................................................................... 44

CHAPTER 3. MATERIALS AND METHODS.................................................................................... 47

3.1. GENETIC TRANSFORMATION SYSTEMS, BANANA CULTIVARS AND CELL CULTURES............................ 47 3.2. VECTORS AND BACTERIAL MANIPULATIONS ...................................................................................... 48

3.2.1. Agrobacterium strains, binary and expression vectors ............................................................. 48 3.2.2. Growth and preparation of competent bacterial cells .............................................................. 50 3.2.3. Plasmid DNA purification......................................................................................................... 50 3.2.4. Heat shock transformation of E. coli cells ................................................................................ 51 3.2.5. Electroporation of Agrobacterium cells.................................................................................... 51

3.3. AGROBACTERIUM-MEDIATED TRANSFORMATION OF BANANA............................................................. 52 3.3.1. The effect of physical parameters on transformation frequency ............................................... 53

3.3.1.1. Length of infection time......................................................................................................................53 3.3.1.2. Age of ECS .........................................................................................................................................53 3.3.1.3. ECS volume during co-cultivation ......................................................................................................53

3.4. PARTICLE BOMBARDMENT-MEDIATED TRANSFORMATION OF BANANA.............................................. 54 3.4.1. Preparation of ECS for particle bombardment ......................................................................... 54 3.4.2. Coating of microparticles and ECS bombardment ................................................................... 54

3.5. POLYAMINES AND PLANT REGENERATION.......................................................................................... 54 3.6. TRANSIENT AND STABLE UIDA GENE EXPRESSION, HISTOCHEMICAL GUS ASSAY .............................. 55 3.7. MOLECULAR CHARACTERISATION OF TRANSFORMANTS.................................................................... 56

3.7.1. PCR analysis ............................................................................................................................. 56 3.7.1.1. DNA isolation for PCR analysis .........................................................................................................56 3.7.1.2. PCR conditions ...................................................................................................................................56 3.7.1.3. Multiplex PCR (MPCR) analysis ........................................................................................................57

3.7.2. Reverse Transcriptase (RT)-PCR analysis................................................................................ 58 3.7.3. Southern hybridisation analysis ................................................................................................ 59

3.7.3.1. DNA isolation for Southern analysis...................................................................................................59 3.7.3.2. DNA digestion and one copy reconstruction.......................................................................................60 3.7.3.3. Blotting, hybridisation and detection with non-radioactive probes .....................................................61

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CHAPTER 4. COMPARISON OF TRANSFORMATION METHODS ............................................ 63

4.1. INTRODUCTION .................................................................................................................................. 63 4.2. TRANSIENT GENE EXPRESSION IN AMT AND PMT SYSTEMS ............................................................... 63 4.3. STABLE TRANSFORMATION FREQUENCIES IN AMT AND PMT SYSTEMS.............................................. 65

4.3.1. Embryogenic cell colonies......................................................................................................... 65 4.3.2. Regenerated plants .................................................................................................................... 66 4.3.3. Grouping banana cultivars based on transformation competence and regeneration ............... 67

4.4. CHARACTERISATION OF TRANSGENIC LINES FROM AMT AND PMT SYSTEMS ..................................... 67 4.4.1. Histochemical GUS assay of transformed lines ........................................................................ 67 4.4.2. PCR analysis in AmT and PmT generated transformants ......................................................... 69

4.4.2.1. PCR analysis in AmT system .............................................................................................................69 4.4.2.2. PCR analysis in P-mT system.............................................................................................................71

4.4.3. RT-PCR analysis of transformants generated via AmT and PmT systems ................................ 72 4.4.4 Southern analysis of transgenic lines from AmT and PmT systems............................................ 74

4.5. CONCLUSION...................................................................................................................................... 75

CHAPTER 5. OPTIMISATION OF AMT SYSTEM........................................................................... 77

5.1. INTRODUCTION .................................................................................................................................. 77 5.2. OPTIMISING PHYSICAL PARAMETERS FOR IMPROVED TRANSFORMATION FREQUENCY ....................... 78

5.2.1. Length of infection period ......................................................................................................... 78 ................................................................................................................................................................. 78

5.2.2. Effect of ECS age....................................................................................................................... 79 5.2.3. Effect of ECS volume................................................................................................................. 80

5.3. TRANSFORMATION OF FOUR BANANA CULTIVARS WITH GFP GENE..................................................... 81 5.3.1. Transient and stable gfp gene expression.................................................................................. 81

5.4. MOLECULAR ANALYSIS OF GFP GENE IN BANANA .............................................................................. 84 5.4.1. PCR analysis ............................................................................................................................. 84 5.4.2. Transcription of gfp gene .......................................................................................................... 85 5.4.3. Integration pattern of gfp transgene into banana genome ........................................................ 86

5.5. THE EFFECTS OF SPERMIDINE ON BANANA ECS REGENERABILITY ..................................................... 88 5.6. CONCLUSIONS AND PERSPECTIVES ..................................................................................................... 90

CHAPTER 6. TRANSFORMATION WITH RICE CHITINASE GENES........................................ 91

6.1. INTRODUCTION .................................................................................................................................. 91 6.2. PLANT MATERIAL AND BINARY VECTORS........................................................................................... 91 6.3. INDUCTION OF TRANSFORMED EMBRYOGENIC COLONIES AND PLANT REGENERATION ....................... 91 6.4. MOLECULAR ANALYSIS OF CHITINASE TRANSFORMANTS................................................................... 92

6.4.1. PCR analysis ............................................................................................................................. 92 6.4.2. Southern blot analysis of Cht-2 and Cht-3 genes ...................................................................... 94

6.4.2.1. DNA isolation and restriction digestion..............................................................................................95 6.4.2.2. Nucleotide sequence analyses of chitinase from banana and rice .......................................................96

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6.4.2.3. Comparisons of amino acid sequences of rice and banana chitinases .................................................98 6.4.2.4. Improved Southern blot analysis of rice chitinases genes .................................................................100

6.5. CONCLUSION ................................................................................................................................... 101

CHAPTER 7. CO-TRANSFORMATION OF RICE CHITINASE WITH A PLANT DEFENSIN103

7.1. INTRODUCTION ................................................................................................................................ 103 7.2. CO-TRANSFORMATION OF BANANA.................................................................................................. 103 7.3. EFFICIENCY OF CO-TRANSFORMATION IN BANANA ECS .................................................................. 105 7.4. MULTIPLEX PCR (MPCR) ANALYSIS OF CO-TRANSFORMANTS ....................................................... 109

7.4.1. Primer combinations and their concentrations....................................................................... 109 7.4.2. Effect of increased template DNA ........................................................................................... 111

7.5. SOUTHERN BLOT ANALYSIS OF CO-TRANSFORMED BANANA LINES .................................................. 112 7.6. CONCLUSION ................................................................................................................................... 114

CHAPTER 8. GENERAL CONCLUSION AND DISCUSSION ...................................................... 117

8.1. COMPARISON OF AMT AND PMT SYSTEMS ...................................................................................... 117 8.2 OPTIMISATION OF AMT SYSTEM ....................................................................................................... 119 8.3. INTEGRATION OF RICE CHITINASE IN BANANA.................................................................................. 122 8.4 CO-TRANSFORMATION WITH RICE CHITINASE AND A DEFENSIN........................................................ 123

REFERENCES ...................................................................................................................................... 127

LIST OF PUBLICATIONS .................................................................................................................. 155

General introduction

1

Chapter 1. General introduction

1.1. Importance of banana in East Africa

Bananas (Musa spp.) are among the most important crops in East Africa and constitute a

major staple food for millions of people in the region (INIBAP, 2000). There, around 15

million tonnes of bananas are produced annually, and the consumption rate is the highest

in the world. Over 90% of the crop is produced in smallholdings (0.25-1.0 ha) with

minimum inputs and consumed almost exclusively locally. Uganda is the leading producer

and consumer of bananas in Africa (FAOSTAT, 2004). East African Highland Bananas

(EAHB) serve as the principle staple food (‘matooke’) in Uganda with an average daily

consumption of 0.6 kg/capita (FAOSTAT, 2004). This is due to the continuous fruiting

habit of EAHB varieties, an ability that provides food to millions of families throughout

the year without hunger-gaps as opposite to cereal and root crop-based systems.

‘Matooke’ is the staple food for over 7 million people in Uganda (Karamura and

Karamura, 1994) with more than 66% of urban dwellers depending on it (Rubaihayo,

1991). Besides providing a source of income through local sales in urban centers, other

uses of bananas in Uganda include livestock feeds, mulch, medicine and fibre for

thatching and making crafts (Rubaihayo and Gold, 1993).

Bananas and plantains include diverse types, which are classified according to their end

uses and genome groupings. The former category includes dessert, cooking, roasting and

beer bananas (Simmonds, 1962). Genome groupings of cultivated bananas include a range

of diploids, triploids and tetraploids and are divided according to their morphology and the

origin of their genome(s), A and B representing Musa acuminata and M. balbisiana,

respectively. A wide range of banana varieties can be found in East Africa, but well over

75% of the crop consists of EAHB with an AAA genome. These are principally cooking

(‘matooke’) and beer bananas. Other varieties grown include plantains (AAB), the dessert

varieties ‘Cavendish’ (AAA) and ‘Gros Michel’ (AAA), other beer bananas (ABB), and

some diploid dessert varieties (AB) (Simmonds and Stover, 1987). More recently,

improved hybrids, mainly tetraploids, have been introduced from breeding programmes to

address problems of declining yields and pest/disease pressure (see also 1.3).

1.2. Banana production constraints

Between 1970 and 1990, banana and plantain yields significantly declined in Uganda from

8 million to 5 million tonnes (Ministry of Agriculture Report, 1991). This yield decline

has led to some replacement of the highland cooking bananas by exotic beer bananas

Chapter one

2

and/or by other crops such as cassava and sweet potatoes. As a result, many rural

communities in the region are now unable to meet their needs, resulting in food insecurity

and other poverty-related problems.

The production decline has so far been more or less compensated by opening up more

fresh land for banana cultivation. However, this extension frequently leads to

environmental degradation and exposure of more land to pest infestation and soil erosion.

Moreover, the 270 capita/square km of population density in Uganda, already the highest

of Africa, leaves not much room to further expansion, let alone leaving land under fallow

to restore fertility. The dimension of the problems facing the banana sector in Uganda can

best be illustrated by the expected fast increase of the country’s population in the next 25

years exceeding 100 million by 2050 compared to 28 million at present (UBOS, 2007).

Thus, banana production should increase with the same speed if current consumption rate

is to be kept.

Results from a rapid rural appraisal held in 1992 indicated that banana production in

Uganda is hampered by biotic and abiotic factors (Bekunda and Woomer, 1996), which

differ between regions and even farms (Gold et al., 1993). The biotic constraints primarily

include leaf diseases (Tenywa et al., 1999; Karamura et al., 1999; Gold et al., 2004),

followed by plant parasitic nematodes (Ssango et al., 2004), banana weevil (Gold et al.,

2004; Kiggundu et al., 2007), and, more recently, banana streak virus and bacterial wilt

(Tushemereirwe et al., 2001). Of the foliar diseases, Yellow and Black Sigatoka are the

major fungal ones, causing severe reductions in fruit quality and yield (Burt et al., 1997).

Black Sigatoka, which has gradually replaced Yellow Sigatoka, is the most aggressive,

and causes crop losses up to 30-50% (Stover and Simmonds, 1987; Mobambo et al., 1993;

Tushemereirwe et al., 2000). Declining soil fertility resulting from intensive land use

(Okech et al., 1996) or a reduction of farm inputs, such as mulch (Rubaihayo et al., 1994)

are the major abiotic constraints.

1.3. Banana genetic improvement through classical breeding

In response to the above constraints, attempts have been made to develop banana varieties

that are superior to the endemic varieties in terms of vigour, drought tolerance, disease,

and pest resistance and yield. However, progress in classical breeding is limited by

sterility and polyploidy in most edible bananas, relatively long generation times, and large

area requirements for field testing. Some of these obstacles have been overcome through

conventional methods by screening for seed fertility, and via ploidy manipulations as well


3

as interspecific hybridisation (Swennen and Vuylsteke, 1993; Rowe and Rosales, 1996;

Vuylsteke et al., 1997). As a result, several classical breeding programmes have generated

new hybrids that are widely distributed in Africa and elsewhere. For instance, Black

Sigatoka resistant hybrids have been produced at the International Institute of Tropical

Agriculture (IITA) and distributed in East and West Africa for evaluation (Vuylsteke et

al., 1995; Gallez et al. 2004). In Uganda, disease resistant hybrids were introduced from

the Fundacion Hondurena de Investigacion Agricola (FHIA), a breeding programme in

Honduras. However, the acceptance of these new hybrids has remained low because of

their inferior cooking quality. In addition, crosses between triploid EAHB and the fertile

diploid Calcutta 4 (Musa acuminata ssp. burmannicoides) resulted in hybrids with

moderate to high resistance to Black Sigatoka but of poor cooking qualities compared to

the triploid ‘matooke’ parents. Recently, 12 (secondary triploid) hybrids of Black Sigatoka

resistant EAHB were selected and are undergoing on farm evaluation (Tushemereirwe et

al., 2005).

1.4. Improvement of banana via genetic engineering

Most banana varieties do not produce seeds and can thus not be crossed but those that are

fertile produce just a few seeds per bunch (Swennen and Vuylsteke, 1993). This high

sterility calls for the integration of biotechnological tools into breeding programmes. In

addition, gene transfer offers the possibility to add just a few novel traits without altering

the genome of the preferred variety. This is very attractive as all Black Sigatoka resistant

hybrids currently available deviate from traditional varieties in taste, shelf life,

morphology, etc. making them less acceptable.

All technologies required for genetic engineering of banana have become available during

the past decade. Embryogenic cell suspensions were generated from male buds (Escalant

et al., 1994; Côte et al., 1996; Grapin et al., 1998) and scalps (Dhed'a et al., 1991; Strosse

et al., 2006). Cryogenic techniques aimed at preserving these cell suspensions (Panis et

al., 1990) reduced losses by contamination and made cell suspensions available relatively

quickly without the need to go repeatedly through the complicated induction procedure.

Several genetic transformation systems have been optimised for banana, which include

electroporation of protoplasts (Sági et al., 1994), particle bombardment of cell suspensions

(Sági et al., 1995a; Becker et al., 2000) and Agrobacterium-mediated gene transfer (May

et al., 1995; Hernández et al., 1999; Ganapathi et al., 2001; Hernández et al., 2006). The

potential of integrating multiple genes, via particle bombardment, required for durable

Chapter one

4

resistance in banana has also been investigated (Remy et al., 1998a) and later

demonstrated by using Agrobacterium-mediated transformation (Ahmed et al., 2002).

1.5. Potential genes for fungal control in banana

Fungal diseases affect all major banana organs and tissues (Sági, 2000b). Pathogenic fungi

primarily attack and necrotise foliage, roots, vascular tissues, and fruits. In their

interaction with banana, pathogenic fungi produce toxins (Hoss et al., 2000), and a mass

of mycelium in the intercellular space, block the vascular system and cause necrotic spots

on foliage and fruits (Jones, 1999; Sagi, 2000a; Ploetz, 1999). In susceptible banana

cultivars, the hemibiotroph Mycosphaerella fijiensis (Hadrami et al., 2005) colonizes, 3 to

4 weeks after infection, biotrophically the intercellular space followed by increased

synthesis of 2,4,8-trihydroxytetralone (2,4,8-THT) as the main avirulent factor (Hoss et

al., 2000). Continuous production and accumulation of 2,4,8-THT enhances extensive

mycelial growth within the leaf intercellular space leading to necrosis and advanced

disease symptom development. Thereon the fungus lives saprophytically on dead leaf

tissue. Thus the growth patterns displayed by this pathogen provide several potential

approaches to achieve resistance. These include prevention of pathogen entry into the

stomata, suppression of its establishment in the intercellular space, and finally restricting

its extensive growth after penetration or cell death.

Genetic enhancing strategies could include the expression of genes encoding antifungal

plant proteins (Remy et al., 1998b) or hydrolytic enzymes of fungal origin. Genes

encoding elicitors of defense responses could also be used (Keller et al., 1999). In

addition, broad-spectrum antimicrobial peptides (AMP) have potential to control fungi and

bacteria. Reported examples of AMPs include magainin from the African clawed frog

(Bevins and Zasloff, 1990), cecropins from the giant silk moth (Boman and Hultmark,

1987) and plant defensins (Broekaert et al., 1995). Transgenic banana lines containing

several AMPs (Remy et al., 2000) showed differential disease response (Remy et al.,

1999). Cecropin (Alan and Earle, 2002) and its derivative D4E1 as well as its hybrid

peptide with melittin (Osusky et al., 2000) inhibited in vitro growth of a range of

pathogenic fungi. Expression of this synthetic peptide enhanced disease resistance in

transgenic tobacco and banana (Chakrabarti et al., 2003). Other strategies could include

expression of genes resulting in cell wall reinforcement and increased levels of

phytoalexins (Otalvaro et al., 2002). The use of plant chitinases and defensins is discussed

in details in sections 2.4.1.3 and 2.4.1.4, respectively. Based on the Mycosphaerella


5

fijiensis infection process, we propose to use two rice chitinases that are localised either

intra- or extracellularly. Rice chitinase gene Cht-2 is targeted intracellularly whereas Cht-3

is targeted to the intercellular space (apoplast). Considering the complexity of

Mycosphaerella fijiensis, a multi-line resistance approach could be more effective than

single gene-based host defence. Thus, a strategy for co-transformation of genes with

different modes of action could give effective and durable host protection if the products

of such genes were localised differently. A drawback is however that the in vitro activity

of purified chitinases against Mycosphaerella fijiensis has not been tested yet. An

overview of other genes that can be evaluated in banana genetic improvement via genetic

engineering was described by Sági (1999).

1.6. Research objectives

The general goal of this research was to generate transgenic lines containing genes with

potential to create resistance against Black Sigatoka disease preferably in EAHB. This

would then create the basis for field testing in Uganda. In addition, the acquired expertise

would create a platform in Uganda for the transformation of EAHB and evaluation of

transgenic EAHB plants under tropical field conditions. Specifically, this work aimed at

contributing to an improved transformation technology. First, the efficacy of

Agrobacterium-mediated and particle bombardment transformation systems was

compared. Further, the effect of several parameters on transformation efficiency was

tested in the Agrobacterium system. Parameters included infection length, co-cultivation

volumes, and embryogenic cell suspension (ECS) age. Finally, several candidate genes

alone or in combination were transferred to banana via Agrobacterium-mediated (co-)

transformation.

1.7. Outline of this thesis

This thesis contains eight chapters. The basis and the background rationale to the problem,

and objectives are presented in Chapter 1. Chapter 2 reviews established scientific facts

related to the objectives. Inter-related events of host-pathogen interaction that leads to

disease and/or resistance response are introduced. Such events link some resistance

mechanisms, enzymatic hydrolysis and pathogen inhibition that form the research basis of

this thesis. Two elements of these mechanisms, chitinases and plant defensins, are

discussed giving their classifications, modes of action, and their recent applicability in

plant genetic engineering.

Chapter one

6

Description of methods and molecular techniques employed is given in Chapter 3. This

chapter explains in details, the two gene transfer systems commonly used in the genetic

transformation of banana. It concludes with different aspects of ECS maintenance and

regeneration, and detection of transgenes in banana genomes.

Chapters 4, 5, 6 and 7 present results generated from different sets of experiments.

Chapter 4 presents results and comparative analyses of Agrobacterium- and particle

bombardment-mediated transformation systems. Analyses of transgenic lines derived from

each gene transfer system are given and their transformation efficiencies compared.

Chapter 5 assesses Agrobacterium-mediated transformation system using a synthetic gfp

gene (sgfpS65T). Possible optimisation steps are identified. The effects of physical

parameters including infection length, ECS age, and ECS volumes, on transformation

frequency are presented. This chapter concludes with a discussion on the effects of

polyamine spermidine on regenerability of transformed embryogenic cell clones.

Transformation and integration of rice chitinase genes Cht-2 and Cht-3 in banana are

presented in Chapter 6.

Chapter 7 presents results of Agrobacterium-mediated co-transformation of banana with

rice chitinase genes and a defensin (Rs-afp2). Transformation frequencies in combinations

Cht-2/Rs-afp2 and Cht-3/Rs-afp2 are evaluated and compared. Effects of using single

versus two different selectable marker genes are presented. Finally, the chapter presents

results on integration of co-transformed chitinase and defensin genes. The thesis ends with

general conclusions and future perspectives in Chapter 8.

Literature review

7

Chapter 2. Literature review

2.1. Fungal diseases of bananas

Fungal diseases have been one of the main causes of crop losses ever since mankind

started to cultivate plants (Oerke, 1994) and they also are a great challenge to the genetic

improvement of bananas. The most serious fungal diseases of bananas are caused by

Mycosphaerella fijiensis Morelet (Black Sigatoka), M. musicola (Yellow Sigatoka) and

Fusarium oxysporum f.sp. cubense (Panama disease or wilt). Among these pathogens, M.

fijiensis is the most aggressive species and (unlike Yellow Sigatoka) attacks almost all

types of bananas and plantains (Jones, 1993). In plantains (Mobambo et al., 1993) and

EAHB cultivars (Tushemereirwe et al., 2000), leaf necrosis caused by this pathogen was

reported to reduce fruit yield by 30-50%. Currently, Black Sigatoka is the major constraint

to cultivation in commercial banana plantations as well as for small-scale and subsistence

farmers growing plantain (Jones, 1993). In Uganda, Black Sigatoka was highlighted as the

most devastating banana disease (Tushemereirwe et al., 1996). It has gradually replaced

Yellow Sigatoka all over the country, therefore only Black Sigatoka will be discussed

from this point on. Black Sigatoka has also become a main target for banana breeding (De

Langhe, 1992; Ploetz, 1999) as well as for biotechnological research to improve this crop.

2.1.1. Black Sigatoka disease

2.1.1.1. The pathogen

The asexual form of M. fijiensis, the causal agent of Black Sigatoka, was first described as

Cercospora fijiensis Morelet, later renamed to Pseudocercospora fijiensis (Deighton,

1976) and subsequently transferred to the Paracercospora genus (Deighton, 1979). The

conidiophores of M. fijiensis emerge singly or in a small groups of two to eight stalks and

sporodochia are absent (Fullerton, 1994). The conidiophores are mainly confined to the

lower surface of the lesion (Meredith and Lawrence, 1970; Fullerton, 1994).

Conidiospores are obclavate to cylindro-obclavate and straight or slightly curved. The

conidial scars are thickened, conspicuous and confined to a narrow rim where the

conidium is attached to the conidiophore (Stover and Simmonds, 1987; Gaviria et al.,

1999).

Chapter two

8

During sexual reproduction, the fungus first develops many spermagonia on the lower

surface of the leaf, usually when lesions collapse but occasionally already during the

development of streaks (second stage, Figure 2.1) or even spots (first stage). Spermagonia

are dark, somewhat erupt and pear-like in shape, and frequently develop in the sub-

stomatal chamber of the stomata, from which one or more conidiophores emerge. These

structures may ooze large quantities of male reproductive cells (spermatia). Spermatia are

tiny and cylindrical, and will fertilise neighbouring female receptive hyphae, called

trichogynes. Once fertilisation is complete, pseudothecia are formed mainly on the upper

surface of mature lesions, with their ostioles poking through the leaf tissue. The oblong to

club-shaped sac-like structures (asci) have two cell walls (bitunicate), and contain eight

sexual spores (ascospores) that are lined up two-by-two. The ascospores are colorless and

have one septum. One cell of the spore may be slightly broader than the other one, and the

spore may be slightly constricted at the septum. Pseudothecia mature when dead leaf

tissues are saturated with water for approximately 48 hours (Stover, 1980, 1986). The

incidence and spread of Black Sigatoka disease is highly influenced by the phase of

reproduction as M. fijiensis forms relatively few conidia and it is mainly dispersed by

ascospores (Ploetz, 1999). The disease pressure and spread are intensified by infected

planting materials and leaves, which often are used as packaging materials. Hence,

ascospores are the primary means of long distance dispersal and the main means of

spreading during extended periods of wet weather (Thurston, 1998).

2.1.1.2. Infection process and diseases development process

Mycosphaerella fijiensis is a hemibiotrophic pathogen with a very high level of genetic

diversity (Carlier et al., 1996). In susceptible banana cultivars the fungus penetrates

banana leaf tissue exclusively through stomata (Hoss et al., 2000). Prior to stomata

penetration, the hyphae form swellings (stomatopodia) just above the stomata (Balint-

Kurti et al., 2001). After penetrating the leaf, the pathogen colonizes a few adjacent cells

for approximately 7 days without any evidence of cell disruption (Marin et al., 2003).

Appressoria enlarge and haustorial protrusions are observed at this stage. The vegetative

hyphae can emerge from the stomata, grow on the leaf surface, penetrate other adjacent

stomata or produce conidiophores and conidia (Marin et al., 2003). Three to four weeks

after penetration, extensive hyphal growth occurs in inter- rather than intra-cellular space

and the fungus enters its biotrophic phase (Hadrami et al., 2005). Microscopic analysis

shows fungal appressoria attached to cell walls. Points of attachment have been reported to

Literature review

9

exchange nutrients, elicitors or toxins (Heath, 2002). As infection and disease

development proceed, the fungus produces a wide range of secondary metabolites

(fijiensin, tetralone, and juglone) some of which are toxic to banana tissues. It is proposed

that these toxins facilitate extensive spreading of the mycelium within the intracellular

space (Hoss et al., 2000). Continuous hyphal growth, beyond 4-4.5 weeks, the further

accumulation of fungitoxins results in chlorosis, necrosis and cell death, and finally

saprophytic growth of the mycelium. Further disease development is advanced by

intensive chlorosis and necrosis followed by darkening and sporadic fungal growth on

dead leaf tissues (Hoss et al., 2000).

2.1.1.3. Symptoms

With Black Sigatoka, the first symptoms appear as dark brown specks on the lower surface

of the leaf (Stover and Simmonds, 1987). Leaves showing typical symptoms on bananas

are shown in Figure 2.1. The successions of symptoms produced by the pathogen were

described by Stover and Simmonds (1987) and are as follows:

1. Faint, minute, reddish-brown specks (spots) on the lower surface of the leaf.

2. Specks elongate, becoming slightly wider to form narrow reddish-brown streaks.

3. Streaks change colour from reddish brown to dark brown or black, sometimes with a

purplish tinge, clearly visible at the upper surface of the leaf.

4. The streaks broaden and become more or less fused or elliptical in outline, and a water

soaked border appears around them.

5. The dark brown or black centre of each lesion becomes slightly depressed and a water

soaked border becomes more pronounced.

6. The centres of the lesions dry out, become light grey or buff coloured and a bright

yellow transitional zone appears between them and the normal green colour of the leaf.

The lesions remain clearly visible after the leaf collapsed or withered because of their

light coloured centre and dark border.

Figure 2.1 Different developmental stages of characteristic symptoms of leaf streaks caused by M. fijiensis. (A) Spots develop (stage numbers 1-3), (B) lesions enlarge (stage number 4), and (C) lesions merge and reduce living leaf surface (stage numbers 5-6).

Chapter two

10

2.1.1.4. Distribution of Black Sigatoka disease

Black Sigatoka disease was first reported on the Fiji Islands of the South Pacific in 1963

(Rhodes, 1964) but examination of herbarium specimens indicated that it had probably

been present in other areas of Asia and the Pacific before 1963 (Stover, 1978; Long,

1979). The disease was first reported outside Asia in Honduras in 1972, in Costa Rica in

1979, in southern Mexico and throughout Central America by the 1980s (Stover, 1980). It

was later reported in Colombia in 1981 and Ecuador in 1986 (Stover and Simmonds,

1987). The disease is still spreading and was recently reported for the first time in Florida

(Ploetz, 1999). It has become the most important disease of bananas and plantains in South

and Central America, in Africa, in Asia and the Pacific Islands (Figure 2.2).

Figure 2.2 Current global distribution of Black Sigatoka disease (Jones 2000).

In Africa, the disease was first reported in 1973 in Zambia (Raemaekers, 1975).

Thereafter, the disease spread rapidly, initially in West and Central Africa: in Cameroon

and Gabon in 1980 (Frossard, 1980), and in Nigeria in 1986 (Mourichon and Fullerton,

1990). In East Africa, Black Sigatoka disease was first identified in Rwanda, Burundi and

Tanzania (Pemba) in 1987 (Sebasigari and Stover, 1988), and later confirmed in Kenya in

1988 (Kung’U et al., 1992) and in Uganda in 1990 (Tushemereirwe and Waller, 1993).

Literature review

11

2.1.1.5. Management of Black Sigatoka disease

In Uganda, host plant resistance and cultural practices, like crop rotation, are employed as

the main disease control measures (Stover, 1991; Bananuka and Rubaihayo, 1994).

However, in countries producing bananas for export such as those in Latin America, Black

Sigatoka disease is usually controlled by frequent application of fungicides. This is an

expensive practice usually including the use of an aircraft or helicopter, permanent landing

strips, and facilities for mixing and loading the fungicides. With the additional high

recurring expense of the spray materials it has been estimated that the costs ultimately

account for 25% of the final retail price of bananas in the importing countries (Ploetz,

2000). Although East Africa produces more than a third of the total world bananas, this

crop is not treated as it is unaffordable to smallholder farmers. Moreover, bananas are

grown near homesteads, which precludes widespread use of chemicals.

Given the high expense of fungicides and the recurring problem with fungicide resistance

in export plantations, it is clear that the main sustainable and practically effective control

measure is the use of Black Sigatoka resistant varieties. Resistant or more tolerant

varieties for subsistence agriculture are available, but are less productive and less

appealing to consumers than susceptible ones. The situation has slowly begun to change

and hybrids with some level of resistance to Black Sigatoka have been distributed

worldwide for evaluation through the National Research Centers including those in East

Africa. However, reports have already revealed low resistance to Black Sigatoka in

hybrids such as FHIA-01 and FHIA-03 (Daniells et al., 1995; Alvarez, 1997). Varieties

such as ‘Yangambi Km5’ known to be resistant (Fullerton, 1990; Fullerton and Olsen,

1995) showed resistance breakdown in Cameroon (Mouliom-Pefoura, 1998), though such

cases have not been again reported anywhere. In Honduras, where most hybrids were

developed, resistance was proven to M. fijiensis during selection and screening. The poor

resistance performance of these hybrids in other evaluation sites suggests the existence of

different populations of M. fijiensis. This is because of the nature of reproduction of the

fungus and the different environmental conditions to which the pathogen is adapted in

various countries. Since new isolates with different genetic make-up can be formed via

extensive recombination during the predominant sexual reproduction, the chance of these

isolates having different pathogenicity or virulence is real. Fungal populations that are

highly variable adapt very quickly through selection to any control measure be it

chemicals or resistant hosts (McDonald and Martinez, 1991). This therefore calls for

Chapter two

12

durable resistance breeding through gene stacking by either classical improvement or

genetic modification.

2.1.1.6. Mechanisms of resistance against Black Sigatoka

Although the precise mechanisms of resistance to Black Sigatoka are still unclear, various

processes associated with partial resistance such as phytoalexin production and

insensitivity to toxins produced by the fungus are thought to be involved (Beveraggi et al.,

1992). Phytoalexin production can be triggered after stomatal penetration of host tissue by

fungal hyphae. It is believed that the events at a very early stage of contact with the

pathogen determine the future of host-pathogen interaction (Beveraggi et al., 1992).

Mourichon et al. (1990) reported the extraction of toxic substances from two resistant

cultivars but none from a susceptible one. It is still unknown whether these fungitoxic

compounds are induced during infection and act in combination with other compounds

such as phytoalexins or via the modification of the pathogen’s enzyme systems. A recent

study indicated the importance of 2,4,8-trihydroxytetralone (2,4,8-THT) among other

secondary metabolites of the pathogen for host-specific reactions (Hoss et al., 2000). Early

activation of fungal 2,4,8-THT metabolism by resistant cultivars caused necrotic micro-

lesions and elicited defence reactions leading to incompatibility between pathogen and

host plant. Plant defence mechanisms of resistant cultivars in this case were first detected

as an activation of phenylalanine-ammonia lyase (PAL) and then subsequent accumulation

of substances that blocked fungal growth. The role of other secondary metabolites

produced by the fungus still needs to be investigated. Cytological and ultrastructural

studies on the infection process of M. fijiensis on a resistant cultivar (‘Yangambi Km5’)

revealed the accumulation of polyphenolics after fungal penetration (Salle et al., 1990;

Tapia et al., 1990). However, no evidence has been produced to demonstrate the role of

these phenolic compounds in host resistance to Black Sigatoka. Studies on the anatomical

features of leaf surfaces of ‘Grand Naine’, False Horn plantains and ‘Pelipita’ showed

variations in stomatal density (Tapia et al., 1990). A positive correlation between stomatal

density and susceptibility of the cultivars was reported; the most susceptible ‘Grand

Naine’ was also found to have the highest stomatal density. However, these findings may

be of no use to breeders, as younger leaves are usually more susceptible than older ones

while their stomata density is lower than that of older leaves. A thorough understanding of

the mechanism of resistance in the host and of critical factors that contribute to

Literature review

13

incompatible reactions between the pathogen and the host cultivars is vital to assist in the

development and selection of hybrids resistant to Black Sigatoka.

2.2. Plant genetic transformation and plant regeneration

2.2.1 Agrobacterium-mediated transformation

Agrobacterium tumefaciens, a soil bacterium that infects a wide range of dicot plant

species, has been utilised to transfer a DNA fragment (T-DNA) of its tumor inducing (Ti)

plasmid into the genomes of a wide range of organisms, including bacteria, fungi, plants

and even human cells (McCullen and Binns, 2006). The genes inserted into the T-DNA

region are co-transferred and integrated into the host genome. It is well established that

only the T-DNA borders and some flanking sequences are needed for DNA transfer. Thus,

by deleting the original genes that reside on the T-DNA and adding selectable marker

genes and other genes of interest, plasmid vectors without oncogenes can be used to

transfer foreign genes without disturbing the host’s endogenous hormone balance. The

method has been adopted successfully for transformation of numerous dicot species

(reviewed by Herrera-Estrella et al., 2005). Agrobacterium-mediated gene transfer method

has also been employed in transformation of agronomically important monocots like rice

(Hiei et al., 1994), maize (Ishida et al., 1996), wheat (Cheng et al., 1997), and banana

(May et al., 1995; Pérez Hernández et al., 1999, 2000).

Gene transfer in Agrobacterium-mediated transformation is executed through a cascade of

genetically regulated biochemical pathways. These pathways have been studied and their

molecular analyses indicate that several genes are induced and expressed under different

environmental and physiological conditions of both Agrobacterium and host plant cells

(McCullen and Binns, 2006). Agrobacterium tumefaciens has an exceptional genetic and

biochemical ability to transfer T-DNA from a Ti plasmid or genetically engineered

plasmids (binary vectors) into the nucleus of infected host cells (Zambryski, 1998). T-

DNA is later integrated into the plant host cell genome, transcribed and translated into

active proteins (Zupan and Zambryski, 1995). Details of the native A. tumefaciens biology,

its gene transfer and subsequent oncogenesis and tumorigenesis of host plant cells or

tissues were previously reviewed (Zamryski, 1998; Li et al., 2002; Gelvin, 2003).

The molecular basis of Agrobacterium-mediated gene transfer is facilitated by the

activities of a large, ~200 kb Ti plasmid that is resident in virulent Agrobacterium strains

Chapter two

14

(Zambryski, 1998). T-DNA is well defined and flanked by a 23 bp repeat segment on

either end, namely left and right border sequences (LB and RB). Virtually any DNA

fragment cloned within the T-DNA can be transferred into the host plant cell irrespective

of its composition or source. Based on this genetic property, the deletion of the T-DNA

genes responsible for tumorigenesis results into regeneration of a fertile plant that is able

to transmit the engineered DNA to the progeny (Newell, 2000). For convenience, de la

Riva et al. (1998) subdivided the events that lead to T-DNA integration and expression in

host plant cells into five steps. These include: (i) bacterial colonisation (ii) induction of

bacterial virulence system, (iii) generation of T-DNA transfer complex, (iv) T-DNA

transfer, and (vi) integration of T-DNA into the plant genome. Bacterial colonisation is

preceded by host recognition and takes place after the attachment process in a polar

fashion. The attachment of bacterial cells onto host plant cells is reported to be enhanced

by the production of acidic polysaccharides (de la Riva et al., 1998) and expression

products of the chromosomally located locus att (Bradley, 1997). In addition, McCullen

and Binns, (2006) reported the involvement of three chromosomally encoded genes chvA,

chvB and pscA (exoC) in the attachment process. Host recognition also involves virulence

gene activation (Tzfira et al. 2002).

2.2.1.1 Virulence gene expression

Virulence gene (vir) activation during the attachment process requires two genes, virA (a

membrane-bound sensor kinase) and virG (a cytoplasmic response regulator), which are

constitutively expressed at low level and are highly induced in an auto-regulatory fashion

(Winans et al., 1988). During virA/virG activation, VirA autophosphorylates at a

conserved histidine residue and transfers the active phosphate to a conserved aspartate

residue on the VirG. VirG-PO4 then binds at specific 12 bp DNA sequences of the vir

promoters (vir boxes) and activates transcription of vir genes. The phosphorylation of

VirA/VirG is signalled by polyphenols, aldose monosaccharides, low pH (5.5) (Yuan et

al., 2007) and low phosphate concentration (Palmer, 2004). It has been reported that virG

and the chromosomally encoded chvG/chvI genes are independently activated by low pH

(Yuan et al., 2007). The chvG/chvI genes are also involved in the induction of vir gene

expression (Li et al., 2002). The proposed interaction model of VirA/VirG, phenolic

substances and monosaccharides at low pH was recently reviewed by McCullen and Binns

(2006). The vir region is comprised of at least six essential operons, namely virA, virB,

virC, virD, virE, and virG (Tzfira and Citovsky, 2006). Two non-essential operons, virF

Literature review

15

and virH are also involved (De la Riva et al., 1998). These operons contain variable

number of genes. For example virA, virG and virF contain only one gene, and virE, virC,

virH contain two genes, while virD and virB contain four and eleven genes, respectively

(McCullen and Binns, 2006). After effective activation of the vir region the T-DNA

transfer process is initiated.

2.2.1.2. T-DNA transportation into plant cell

Following attachment and vir gene activation, A. tumefaciens transports single stranded T-

DNA (ssT-DNA) and several proteins into the plant cell. Transported components are

delivered into host plant cell through a specialised type IV secretion system transporter

complex (T4SS) made up of VirB and VirD4 protein units (Christie et al., 2004). T-DNA

complex transfer via the T4SS protein structure is facilitated by T4SS-targeting motifs in

VirD2 and VirE2 proteins (Ward et al., 2002). Alongside the T-DNA, several other

proteins are reported to be transported into host plant cells. These include VirD5, VirE3

and VirF, and their detailed roles in T-DNA transport and integration were reviewed by

Gelvin (2003) and Christie (2004).

2.2.1.3. Intracellular transport and T-DNA integration into plant cell genome

Across the cytosol, VirE coated ssT-DNA-VirD2 complex (T-complex) is transported with

the help of host plant cell proteins (Tzfira et al., 2002). It is now known that

Agrobacterium transfers VirD2-T-strand and VirE2 separately, and that the T-complex is

assembled within the host plant cell (Cascales and Christie, 2004). While inside the cell,

VirD2 covalently binds at 5’ end of the ssT-DNA and the rest of the T-stand is coated with

VirE2 to protect it from exonucleolytic degradation in planta (Tzfira and Citovsky, 2006).

T-complex movement across the cytosol is assisted by binding onto host cell protein

microtubules through the bipartite nuclear localisation signals (NLS) of VirD2 and VirE2

proteins (Zupan et al., 2000; Gelvin, 2000) and the dynein motors (Salman et al., 2005).

These NLS interact with NLS-binding proteins that are localised at several points along

nucleus leading microtubules (McCullen and Binns, 2006). T-complex is translocated into

the host cell nucleus through interaction with VirE2 interacting protein (VIP1, Tzfira et

al., 2002) and importinα (Bollas and Citovsky, 1997; Gorlich and Kutay, 1999).

Mayerhofer et al. (1991) proposed two models for T-DNA integration: (i) the double-

stand-break (DSB) repair and (ii) the single-stand-gap (SSG) repair. DSB repair model

requires the presence of a double-stranded (ds) break in the target DNA sequence for T-

Chapter two

16

DNA integration to occur. For a SSG repair model, previously proposed by Tinland

(1996), a single nick is along the T-DNA integration site is converted into a gap by a

5’→3’ endonuclease. The cut strand ends are then partially annealed to the target DNA

and the T-strand overhangs are trimmed. After the ligation of the T-strand strand to the

target DNA, a nick is introduced in the second target DNA and is extended into a gap by

exonucleases. The integration process is finished by the synthesis of a complementary

strand to the T-strand and the ligation of the 3’ end of this newly synthesized strand to the

target DNA (Tinland, 1996). This is the most preferred model for T-DNA integration in

host plant cell genome given the fact that T-DNA is transferred as a single strand (Fu et

al., 2002). However, this model does not explain well the formation of complex T-DNA

integration loci frequently observed in (co-)transformed plants. For illustration purposes,

this is explained further below.

According to the DSB repair model, after localisation of the T-complex in the nucleus

VirE2 is degraded by CSCVirF ubiquitin complex (Tzfira et al., 2004). The ssT-DNA is

recognised by proteins, such as H2A, converted to dsT-DNA by the host cell DNA repair

machinery (Mysore et al., 2000) and integrated at DSB sites within the host cell genome

(De Buck et al., 1999; Tzfira et al., 2002). Integration is assisted by a transcription

regulator VIP2, a second VirE2 interacting protein (Tzfira and Citovsky, 2006). In this

pathway, dsT-DNA intermediates are captured during the DSB repair process (Salomon

and Puchta, 1988). Plant protein KU80, known to be involved in the non-homologous-end-

joining (NHEJ) mechanism (van Attikum et al., 2001), is reported to play a key role

during T-DNA integration (Li et al., 2002). These authors proposed that KU80 is the first

point of contact between dsT-DNA and the host cell DNA repair mechanism. Several

DNA repair and packaging proteins are reported to be essential for T-DNA integration in

plant cells (Tzfira et al., 2004).

2.2.1.4. The structure of integration sites in plants

Integration of T-DNA in the host plant cell genome occurs at random throughout the

genome and is thought to occur via illegitimate recombination (Tinland, 1996; Tzfira et

al., 2004; Tzfira and Citovsky, 2006). Analysis of T-DNA/host DNA junction sequences

and integration loci sequences show that T-DNA RB termini are more conserved than the

LB termini after T-DNA integration (Tinland, 1996; Krizkova and Hrouda, 1998; Tzfira et

al., 2004; Fu et al., 2006). Moreover, these authors observed small deletions in T-DNA

and in the nearby host genomic DNA. Their results also showed that such deletions were

Literature review

17

more severe at the T-DNA 3’ end compared with its 5’ end. Integration sites also showed

the presence of microhomologies between T-DNA 3’ end and the pre-insertion sites. In

most cases, the integrated T-DNA was interspaced with filler DNA (Brunaud et al., 2002).

Such rearrangements are reported to occur before integration. In petunia (Jones et al.,

1987), Arabidopsis (Grevelding et al., 1993) and bentgrass (Fu et al., 2006) multiple T-

DNA copies were integrated at a single chromosome locus. Using two binary plasmids

(carried by two different bacterial strains) containing two different selectable markers on

their T-DNAs, doubly-transformed Arabidopsis plants were obtained in which both T-

DNAs, each derived from a different strain, were found to integrate at the same location

on the plant chromosome (De Block and Debrouwer, 1991). The integration patterns

showed head-to-head, tail-to-head, and other versions of T-DNA interspaced with host

DNA segments (Fu et al., 2006). T-DNA integration frequently occurred in A/T-rich (Fu

et al., 2006) and high gene density regions (Muller et al., 1999). However, no significant

differences were observed between coding exons versus introns (Brunaud et al., 2002) and

a low integration frequency was observed within repetitive sequences in rice (Chen et al.,

2003).

2.2.1.5. Factors influencing Agrobacterium-mediated transformation

Since the success of Agrobacterium-mediated transformation of rice in the early 1990s,

transgenic plants have been regenerated in more than a dozen monocotyledonous species,

ranging from the most important cereal crops to ornamental plant species. Many factors

influencing Agrobacterium-mediated transformation of monocot plants have been

investigated and elucidated. The effect of plant genotype (Carvalho et al., 2004), explant

types (Carvalho et al., 2004) and their transformation competence (Chateau et al., 2000), as

well as the influence of Agrobacterium strains and binary vectors have been reported

(Cheng et al., 2004; Khanna et al., 2004). In addition, a wide variety of inoculation and

co-cultivation conditions have been shown to be important for transformation of monocots.

These include antinecrotic treatments using antioxidants and bactericides, osmotic

treatments (Cheng et al., 2004), pre-culture with growth regulators (Chateau et al., 2000),

desiccation of explants before or after Agrobacterium infection, use of surfactants like

Pluronic F68 (Khanna et al., 2004), and composition of inoculation and co-cultivation

medium (Cheng et al., 2004). Transformation frequencies of wheat inflorescence tissue

were influenced by the duration of pre-culture, level of wounding, and amount of bacterial

cells infiltrated (Amoah et al., 2001). The effects of other physical parameters like

Chapter two

18

infection time and co-cultivation volume can also be investigated. Dillen et al. (1997) and

De Clercq et al. (2002) tested the influence of co-cultivation temperature and 22°C was

reported as the optimum. The effects of Agrobacterium cell density during infection,

medium pH, age and size of calli, density of calli during co-cultivation, and the

concentration of acetosyringone on transformation frequency were also studied (De Clercq

et al., 2002). All these reports highlight the importance of a complex and thorough

optimisation of Agrobacterium-mediated transformation procedures when dealing with

new crops or plant species.

2.2.1.6. Agrobacterium-mediated co-transformation

Genetic transformation with A. tumefaciens has become a vital research tool for gene

expression studies and improvement of crop plants. However, much of this success has

been achieved in cases when the examined trait was encoded by a single gene of interest.

This approach is constrained by the fact that most metabolites in plant cells are produced

in long and complicated pathways, which are composed of and regulated by multiple

genes. The availability of a system for transferring many genes in a single cell would

therefore make it possible to integrate into a single genome different genes with variable

modes of action, a desirable basis for durable resistance.

Problems associated with multiple gene transfer have been reviewed (Daniel and Dhingra,

2002; Francois et al., 2002). Co-integration of several foreign genes in a single genome

can be done via consecutive re-transformations or crosspollinations of lines containing

different transgenes. Co-transformation, the simultaneous transfer of several independent

genes, presents an alternative and promising approach. Co-transformation has frequently

been used with direct gene transfer as DNA constructs can be conveniently mixed before

transformation (Uchimiya et al., 1986; Hadi et al., 1996; Wu et al., 2002). An essential

drawback of this approach is the frequent occurrence of transgenic events with high copy

numbers of transgenes, which increases the risk and incidence of gene silencing.

Therefore, the method can not be routinely applied for studies on multiple gene integration

and their interaction in plant cells (Radchuk et al., 2005).

Agrobacterium-mediated gene transfer was also used for co-transformation with two

different vectors either in a single Agrobacterium strain (Depicker et al., 1985; de Block

and Debrouwer, 1991; Komari et al., 1996; Daley et al., 1998; Matthews et al., 2001;

Jacob and Veluthambi, 2002) or in different Agrobacterium strains (Komari et al., 1996;

McKnight et al., 1987; De Neve et al., 1997; De Buck et al., 1998; Slater et al., 1999) or

Literature review

19

as combination of these two methods (Poirier et al., 2000). These approaches yielded high

co-transformation frequencies of more than 60% in tobacco and 47% in Arabidopsis (De

Buck et al., 1998) and 39-85% in Brassica napus (De Block and Debrouwer, 1991).

2.2.2. Direct gene transfer

Although A. tumefaciens has been found to be a very effective DNA transfer system for

genetic transformation, direct DNA transfer methods have also been developed for

transformation of many plant species (Christou, 1997; Kohli et al., 1998; Ghareyazie et

al., 1997; Christou and Swain, 1990; Tu et al., 2000). This has been primarily in plant

species, which appeared recalcitrant to Agrobacterium infection.

The biolistic method by particle bombardment was a major development in direct gene

transfer (Ghareyazie et al., 1997; Nayak et al., 1997; Tu et al., 2000; Taylor and Fauquet,

2002). The method is considered to be genotype or plant tissue independent (Cheng et al.,

1998; Tu et al., 2000; Taylor and Fauquet, 2002). It consists of delivering DNA into cells

of intact plant organs or cultured tissues via microprojectile acceleration. In this

procedure, small and high density particles (microprojectiles) are accelerated to high

velocity by a particle gun apparatus to acquire sufficient kinetic energy to penetrate plant

cells and membranes thereby carrying foreign DNA into the interior of bombarded cells.

Genetic transformation through microprojectile bombardment has been reported in major

crop plants including barley, bean, canola, cassava, maize, cotton, papaya, peanut,

soybean, squash, sunflower, sugarbeet, wheat (Christou, 1996) and banana (Sági et al.,

1995a, 1995b, 1998).

2.2.3. Polyamines and plant regeneration

Regeneration of transformed banana plants remains the limiting phase in both

Agrobacterium-mediated and direct transformation systems. Though regeneration

frequency is appreciably high in some newly initiated banana ECS cultures, it

tremendously drops with increasing duration of maintenance in liquid media. The reduced

regeneration efficiency could also be caused by stress due to the selective agent(s),

physiological changes within cell clones during regeneration, and transgene expression.

The selective agent may adversely affect growth of transformed plant cells, either directly

or through the accumulation of oxidised polyphenolics and other toxic compounds from

necrotic untransformed tissue (Lindsey and Gallois, 1990). Several approaches have been

utilised to avoid these problems and to maintain or increase regeneration capacity. These

Chapter two

20

include the use of cryopreserved ECSs (Panis and Swennen, 1995), use of strong

antioxidants (Tang et al. 2004), and application of polyamines (Yadav and Rajam, 1998;

Minocha et al., 1999; Tang et al., 2004). Among these approaches, polyamines have been

reported to be effective in the improvement of regeneration capacity in a wide range of

crops (Kumar et al., 1997; Kumria and Rajam, 2002; Kakkar and Sawhney, 2002).

Polyamines are low molecular weight organic cations in living organisms and are involved

in a range of biological processes including growth, development and stress responses

(Kumar et al., 1997; Kakkar and Sawhney, 2002). In plants, the major polyamines are

spermidine [N-(3-aminopropyl) butane-1,4-diamine], spermine [NN´bis-(3-propyl) butane-

1,4-diamine], and their precursor putrescine (butane-1,4-diamine) (Kumar et al., 1997).

Though their biological functions are not well understood, several reports indicate that

they play a crucial role in somatic embryo development, stimulate cell division and

regulate rhizogenesis, embryogenesis, and senescence (Kakkar and Rai, 1993; Minocha

and Minocha, 1995). Strong positive correlation has been found between plant-forming

embryos and their endogenous spermidine levels (Minocha et al., 1999). However, the

effectiveness of polyamines appears to be specific to some key stages of somatic embryo

development (Yadav and Rajam, 1998). The positive effect of spermidine, at various

concentrations, has been reported in cell and tissue cultures of different plant species.

These include 0.1 mM in onion (Martinez et al., 2000), 0.5 mM in rice (Shoeb et al.,

2001); 0.1 M in wheat (Khanna and Daggard, 2003) and 1.5 mM in pine (Tang et al.,

2004).

2.3. Plant responses to pathogen infection

2.3.1. Plant-pathogen interactions

In the event of pathogen (fungi, bacteria, nematodes, and viruses) attack, plants first

reinforce structural barriers that prevent pathogen entry (Dangl et al., 1996; Schmelzer,

2002) and activate enzymatic and chemical defence responses that interfere with pathogen

metabolism (Glazebrook et al., 1997; Melchers and Stuiver, 2000). In brief, these

responses include the synthesis of reactive oxygen species and antimicrobial secondary

metabolites (Kombrink and Somssich, 1995; Dangl and Jones, 2001), lignification of cell

walls and activation of a wide range of genes for such as pathogenesis related (PR)

proteins, which include chitinases (PR-3, Datta and Muthukrishnan, 1999; Kasprzewska,

2003) and plant defensins (PR-12, Terras et al., 1995; Lay and Anderson, 2005). These

Literature review

21

genes, which provide basal defence, inhibit pathogen spread after infection (Dangl and

Jones, 2001). Defence mechanisms employed here include cell wall liginification and

fortification (Hammond-Kosack and Jones, 1996; Schulze-Lefert, 2004; Juge, 2006) and

production of phytoalexins as well as cell wall degrading enzyme inhibitors (Juge, 2006)

as illustrated on Figure 2.4B. Other genes, including resistance genes (R-genes) that are

involved in recognition-dependent or ligand-receptor mediated defence mechanisms

(Glazebrook et al., 1997; McDowell and Dangl, 2000; Dodds and Taylor, 2000; Dangl and

Jones, 2001), trigger a chain of signal transduction events that results into activation of

several defence mechanisms and arrest of pathogen growth. Avirulent pathogens

frequently trigger hypersensitive response (HR, Staskawicz et al., 1995; Cutt and Klessig,

1992; McDowell and Dangl, 2000; Dodds and Taylor, 2000; Dangl and Jones, 2001) and

systemic acquired resistance (SAR) or induced systemic resistance (ISR, Pieterse et al.,

2001). Defence signalling also involves the plant hormones salicylic acid (SA), jasmonic

acid (JA) and ethylene (ET) (reviewed by Pieterse et al., 2001). SA-dependent defence

pathways are induced by biotrophic invasion whereas necrotrophic or wound associated

attacks are more often associated with ET-JA-dependent pathways. Furthermore, SA

signalling pathway induces SAR whereas ET-JA signalling results in ISR. It is important

to note that SA- and ET-JA-dependent defence pathways induce different resistance

mechanisms involving different components, and the two pathways usually have negative

crosstalk between each other (Pieterse et al., 2001). However, both SA- and ET-JA-

induced SAR or ISR, respectively, involve a regulatory protein NPR1. Upon induction of

SAR or ISR, NPR1 activates PR-1 gene expression by physically interacting with a

subclass of basic leucine zipper protein transcription factors that bind to the promoter

sequences of genes required for SA or ET-JA-related PR protein synthesis (Pieterse et al.,

2001). Such defence responses are frequently implemented after a host plant has failed to

contain the pathogen’s invasion, growth and multiplication within its cells (Greenberg,

1996; Sticher et al., 1997; Dangl and Jones, 2001; Pieterse et al., 2001).

2.3.1.1. Pathogen recognition

Perception in specific resistance involves receptors with high degrees of specificity to

pathogen strains, which are encoded by constitutively expressed resistance (R) genes,

located either on the cell membrane or in the cytosol (Edreva, 1991; McDowell and Dangl,

2000). Recognition of pathogen invasion in host plants involves direct interaction between

host specific resistance proteins and the corresponding avirulence (Avr) gene products.

Chapter two

22

This is the receptor/ligand-model, where the plant R genes encode putative receptors that

bind the products of matching Avr genes or race-specific elicitors (Glazebrook, 1999;

Figure 2.4). Plant R genes then encode proteins (Figure 2.3) that both determine

recognition of specific Avr proteins and initiate signal transduction pathways (Figure 2.4)

leading to complex defence responses (Zhou et al., 1998; Martin, 1999). A pathogen Avr

gene, thus, if expressed, causes the host plant to activate its defence responses against the

invading pathogen (Nimchuk et al., 2001).

In addition to the gene-for-gene recognition mediated by the R and Avr genes in host

plants, nonhost resistance is activated through recognition of specific pathogen or plant

cell wall derived signal molecules frequently referred to as exogenous or endogenous

elicitors, respectively. The chemical structure of different elicitors is of great variety, such

as glycoproteins, peptides (Zimmermann et al., 1997), oligosaccharides, and lipids (Ebel

and Cosio, 1994; Edreva, 2004). Some proteinaceous elicitors are directly produced by

bacterial or fungal pathogens, whereas biologically active oligosaccharides are released

from pathogens and plant cell walls by hydrolases secreted by two organisms. Non-

specific elicitors, such as cellulolytic enzymes (Edreva, 2004) can cause transmembrane

ion fluxes in artificial lipid bilayers and other non-specific proteinaceous elicitors, such as

cryptogenin, have been shown to have binding sites on plant membranes, which activate

multiple intracellular defence signalling pathways (Klüsener and Weiler, 1999).

Figure 2.3 Major classes of R proteins (Avr-receptors). A, extracellular Leucine-Rich Repeat (eLRR) linked to transmembrane protein domain and an intracellular protein kinase (KIN); B, eLRR and a transmembrane protein domain; C, intracellular LRR linked to a Nucleotide Binding (NB), and Toll and Interleukin-1 Receptor (TIR) or Coiled-Coil (CC) domain at the N-terminus; D, Intracellular protein kinase (KIN); E, CC linked to a transmembrane protein domain (McDowell and Woffenden, 2003). These elicitors are recognized by pathogen-race specific receptors located in the plant cell

wall and with variable domains inside the cell.

B

A

C

D

D

D

E

Literature review

23

Figure 2.4 Signalling pathways that are activated in response to pathogen attack (Glazebrook et al., 1997).

Five classes of R-proteins (Avr-receptors) (A-E in Figure 2.3) have been reported

(Glazebrook et al., 1997; McDowell and Dangl, 2000; Dodds and Taylor, 2000; Dangl and

Jones, 2001; Dodds et al., 2001). Recognition of Avr products (elicitors) by receptor

proteins encoded by R genes initiates signal transduction pathways (Figure 2.4A) resulting

into multiple gene activation, programmed cell death, SA production and finally SAR in

distal plant tissues. These pathways result in complex defence responses (Zhou et al.,

1998) among which chitinases are also produced (Ryan and Farmer, 1992).

2.3.1.2. Signal transduction

Receptor-mediated recognition at the infection site initiates cellular and systemic

signalling processes that activate multicomponent defence responses at local and systemic

levels resulting in rapid establishment of local resistance and delayed development of SAR

(Scheel, 1998, Dangl and Jones, 2001; McDowell and Dangl, 2000). The earliest reactions

of plant cells include changes in plasma membrane permeability leading to calcium and

proton influx and potassium and chloride efflux (McDowell and Dangl, 2000, Dangl and

Jones, 2001). Ion fluxes subsequently induce extracellular production of reactive oxygen

intermediates, such as superoxide (O.2), hydrogen peroxide (H2O2) and free hydroxyl

A B

Chapter two

24

radicals (OH-), catalyzed by a membrane-located NADPH oxidase and/or apoplastic-

localised peroxidases (Somssich and Halbrock, 1998). Heterotrimeric GTP-binding

proteins and protein phosphorylation/dephosphorylation are involved in transferring

signals from the receptor to calcium channels that activate downstream reactions

(Legendre et al., 1992). The changes in ion fluxes trigger localised production of reactive

oxygen intermediates and nitric oxide, which act as messengers for HR induction and

defence gene expression (Piffanelli et al., 1999). Other components of the signal network

are specifically induced phospholipases, which act on lipid-bound unsaturated fatty acids

within the membrane, resulting in the release of linolenic acid, which serves as a substrate

for the production of JA, methyl jasmonate and related molecules via a series of enzymatic

steps (Odjakova and Hadjiivanova, 2001). Most of the inducible, defence-related genes

are regulated by signal pathways involving one or more of the three regulators JA, ET and

SA (Van Loon, 1997; Ananieva and Ananiev, 1999; Pieterse et al., 2001). JA and ET are

reported to co-operate to regulate the expression of many defence related genes (Reymond

and Farmer, 1998).

An increase in the intracellular Ca2+ concentration induces β-1,3-glucan synthase, a

constitutive membrane-bound enzyme that converts UDP-glucose into β-l,3-glucan

polymers, and the subsequent local deposition of callose from the plasma membrane onto

the adjacent cell wall (Blumwald et al., 1998; McDowell and Dangl, 2000). Also, these

ion fluxes are prerequisite for the activation of specific Mitogen-Activated Protein (MAP)

kinases (Dangl and Jones, 2001). Substantial accumulation of callose at the sites of

potential fungal penetration functions as an effective early response for delaying pathogen

spread. This allows the plant to prepare complementary responses requiring the

transcriptional activation of defence genes.

Subsequently, among the major biochemical changes that are induced when plants

encounter pathogens are the accumulation of phytoalexins with microbial toxicity, the

synthesis of PR proteins and the production of protease inhibitors. Responses of plants to

pathogen invasion also result in the suppression of various housekeeping activities of the

cells, thus diverting the cellular resources to defence responses. In the initially attacked

cell(s), rapid responses ultimately lead to death (Narasimhan et al., 2001).

Literature review

25

2.3.2. Induced defence responses

Hypersensitive response (HR), the localised cell death, is induced by some pathogens and

elicitors (Greenberg, 1996, Glazebrook et al., 1997; McDowell and Dangl, 2000) and is

characterised by DNA breaks with 3’OH ends, loosening of the plasma membrane, nuclear

and cytoplasmic condensation, and activation of membrane Ca2+/K+ exchange (Baker et

al., 1987; Davis et al., 1991), all of which lead to localised cell and tissue death at the site

of infection (Van Loon, 1997). This response may be responsible for disease resistance or

for the activation of other plant development processes (Heath, 1980; Hammond-Kosack

and Jones, 1996; Greenberg, 1996). HR involves the rapid localised necrosis of the

infected plant cells or tissue or both (Pieterse et al., 2001). The purpose of this self-

sacrifice is to deprive the invading pathogen from an adequate nutrient supply (Greenberg,

1996; Glazebrook et al., 1997; McDowell and Dangl, 2000) or releasing microbiocidal

compounds from dying cells and to restrict the pathogen to small areas immediately

surrounding the initially infected cells (Dangl et al., 1996; Jackson and Taylor, 1996). HR

also results in the production of signalling molecules SA or nitric oxide, which may help

to transmit the signal to other parts of the plant and induce the systemic acquired

resistance (Pieterse, et al., 2001).

2.4. Potential genetic engineering strategies

Due to the biology of the banana plant and lack of knowledge of resistance genes against

Sigatoka in most cultivated and edible banana cultivars, classical breeding has achieved

limited success. There is a need, therefore, to exploit all available resistance strategies and

use available genes with a broad spectrum of antifungal activity. Several resistance

development approaches via plant genetic engineering have been undertaken and reviewed

(Punja, 2001) in other crops, and variable levels of resistance were observed in the

laboratory and the greenhouse. It is important to assess the suitability and efficacy of these

strategies for the induction of Black Sigatoka resistance. The approaches that have been

taken in other plants include:

(a) The expression of gene encoding products that are directly toxic to pathogens or

interfere with their growth. These include PR proteins such as hydrolytic enzymes

(chitinases, glucanases), and antifungal proteins.

(b) The expression of genes encoding products that destroy or neutralise a structural

component of the pathogen such as polygalacturonides, oxalic acid, and lipids.

Chapter two

26

(c) The expression of genes encoding enzymes that are involved in liginin biosynthesis.

These induce elevated levels of peroxidase and lignin.

(d) The expression of genes encoding products that regulate plant defence mechanisms.

These include the production of specific elicitors, hydrogen peroxide, salicylic acid

(SA), and ethylene.

(e) The expression of R genes encoding products involved in hypersensitive response

(HR) and in interactions with avirulence (Avr) factors.

In our research the first (a) strategy that involves the use of rice chitinase genes and a

radish defensin gene (Rs-afp2) are exploited. Thus, a detailed analysis on the scientific

basis and applicability of these strategies is presented.

2.4.1. Hydrolytic enzymes

The most widely used transgenic approach to create disease resistance has been the

overexpression of chitinases and glucanases, which belong to the group of PR proteins

(Neuhaus, 1999). These enzymes inhibit the growth of many fungi in culture, indicating

that they have a direct antifungal function (Mauch et al., 1988; Ji and Kuc, 1996; Boller,

1993; Yun et al., 1997). The antifungal activity is due to their ability to hydrolyse β-

glycosidic bond of different polymer-forming glucosamines such as chitin (N-acetyl-D-

glucosamine), chitosan or peptidoglycan (Schlumbaum et al., 1986), the main structural

components of the fungal cell wall (Legrand et al., 1987; Punja, 2001).

Chitinases (E.C. 3.2.1.14) are poly(1,4-(N-acetyl-β-D-glucosaminide))-glycanohydrolases.

They are widely distributed in nature, occurring in bacteria, fungi, animals, and plants.

Chitinases are classified into glycosyl hydrolase families 18 and 19, depending on their

amino acid sequences (Warren, 1996; Patil et al., 2000). All the known bacterial chitinases

belong to the family 18 glycosidase (Henrissat, 1991; Henrrisat and Bairoch, 1993).

Chitinases produced by bacteria appear to have a nutritional or scavenging role, while

those produced by filamentous fungi have been shown to be involved in a variety of

functions such as cell wall digestion, germination of spores, hyphal growth, hyphal

autolysis, differentiation into spores, assimilation of chitin and mycoparasitism (Gooday,

1990; Flach et al., 1992). Some chitinase genes that have been used in plant genetic

engineering are presented in section 2.4.1.2.

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27

2.4.1.1. Plant chitinases

Plant chitinases are usually endo-chitinases capable of degrading chitin (Graham and

Sticklen, 1994), via hydrolysis of β-1,4-linkage of the N-acetylglucosamine polymer.

Chitin is a major constituent of certain fungal cell walls as well as arthropodal and

nematodal exoskeletons and insect gut linings. In fungi, chitin occurs as a major

component in the cell walls of fungal divisions Basidiomycetes, Ascomycetes,

Deuteromycetes and Zygomycetes (Wessels and Sietsma, 1981; Gooday, 1990). Figure 2.5

shows the enzymatic reaction catalyzed by endochitinases. Multiple chitinase isoforms and

gene clusters have been detected in many plants analyzed to date. Plant chitinases are

generally small proteins of 25-35 kDa molecular weight (Shinshi et al., 1990), with wide

range of isoelectric points (3-10), and post-translational modifications such as

glycosylation and prolyl-hydroxylation (Sticher et al., 1992; Colinge et al., 1993; Nielsen

et al., 1994). Chitinases generally show wide pH optima (4-9) for activity. Some

chitinases, such as a yam class III chitinase, show two pH optima depending on the

substrate used (Tsukamoto et al., 1984).

Figure 2.5 Enzymatic hydrolysis of polymer chitin by an endo-chitinase. Chitin polymer is reduced to N,N’-diacetylchitobiose and higher oligomers of reduced lengths (www.sigmaaldrich.com/Chitinase).

This particular chitinase is also stable at 80°C, whereas other plant chitinases show

moderate temperature tolerance of up to 60°C. Chitinases contain several disulfide

linkages through conserved cysteine residues in their tertiary structure. Crystal structures

of heveamine, a class III chitinase/lysozyme from rubber tree (Hevea brasiliensis)

Chapter two

28

(Scheltinga et al., 1995), class II chitinases from barley (Hordeum vulgare) (Hart et al.,

1995), and jack bean (Canavalia ensiformis) (Hahn et al., 2000) have been determined.

Based on their primary structures, plant chitinases have been classified into seven classes,

class I through VII (Neuhaus, 1999). Different chitinase classes have no apparent

correlation to being present in a particular plant species and plant organ or tissue.

However, certain chitinase isoforms are sometimes induced by a particular elicitor. For

example, in potato, a class I basic chitinase was strongly induced by ethylene and

wounding whereas a class II acidic chitinase was induced by SA (Büchter et al., 1997).

Also, only particular isoforms show antifungal activities and certain isoforms have

additional novel functions such as antifreeze activity (Buurlage et al., 1993; Yeh et al.,

2000).

2.4.1.1.1. Class I and II chitinases

Class I and II chitinases belong to the PR-3 family of PR proteins with the tobacco

chitinases as the prototypical members (Neuhaus, 1999). All members of the PR-3 family

belong to family 19 of glycosyl-hydrolases, which catalyzes sugar hydrolysis with the

inversion of configuration at the anomeric carbon. Class I chitinases are synthesised as

precursors with an N-terminal propeptide. Class II chitinases are synthesized as

propeptides, directed to the secretory pathway and eventually directed to the vacuole by a

short C-terminal signal sequence. Their Chitin Binding Domain (CBD) is separated from

the catalytic domain by a proline- and glycine-rich hinge or spacer region, variable both in

size and composition. For a tobacco class I chitinase, the deletion of CBD and the spacer

region singly or in combination reduces the hydrolytic activity by 50%, whereas antifungal

activity is reduced by 80% (Suarez et al., 2001). Deletion of the C-terminal signal peptide

redirects a class I chitinase to the apoplast while retaining the enzymatic activity (Grover

et al., 2001). Addition to this six amino acid signal (GLLVDTM), an unrelated, usually

secreted class III chitinase redirects this protein to the vacuole in tobacco (Neuhaus et al.,

1991). Class II chitinases are similar to class I but they lack the N-terminal CBD and the

hinge region. In the catalytic domain they sometimes have a deletion as compared to class

I chitinases. Class II chitinases are usually secreted to the apoplast as they lack the C-

terminal vacuolar-targeting signal.

Literature review

29

Protein structure of class I and II chitinases

A barley class II monomeric 26 kDa chitinase (Hart et al., 1995) and a jack bean class II

chitinase (Hahn et al., 2000) are the only members of the PR-3 family whose crystal

structures are known. The structure of barley chitinase is mostly α-helical and forms a

globular structure of approximately 42 Å. The barley enzyme is composed of one short

antiparallel β-strand and ten α-helices occupying 47% of the primary structure. The jack

bean chitinase also shows similar structure with ten helices of 7-8 amino acids. These

structures have similarity to the lysozyme fold of hen egg-white lysozyme (HEWL) and

other family 19 glycosyl-hydrolases without sequence homology. In the barley enzyme

four loops have been identified. Loops 1 and 2 surround the catalytic site and are held by

conserved disulfide bonds, whereas loops 3 and 4 are located at the surface and are not

likely to participate in the catalysis. In the barley enzyme three disulfide bonds, between

cysteine residues 24-86, 98-104, and 203-222, are held by conserved cysteines. These

disulfide linkages are also found in jack bean chitinase at similar locations. Two active site

glutamic acid residues have been identified in the crystal structures. In the barley enzyme

these residues are identified at amino acid positions 67 and 89. Mutation of either active

site glutamate turned a class I chitinase into a chitin binding lectin (Iseli-Gamboni et al.,

1998). The precursor of stinging nettle (Urtica dioica) lectin sequence is related to

chitinases, but it has both the active-site Glu residues mutated and the protein does not

show chitinase activity. Jack bean chitinase also has catalytic site residues located at

similar positions. Interestingly, the HEWL Glu 35 is essential for catalysis and

superimposes with Glu 67 of barley chitinase upon superimposition of the crystal

structures. Other active site residues have been modified or mutated to show their

functions in catalysis. Mutation of Tyr 123 of Zea mays chitinase and a similar tyrosine of

Arabidopsis chitinase in the conserved NYNY catalytic site motif, present in most class I

and class II chitinases at similar locations, causes greatly reduced chitinase activities

(Verburg et al., 1992; 1993). Other residues such as Asn 124, 199, Trp 103, and Try 123

of barley chitinase are implicated in substrate binding and are conserved in other

chitinases.

Chitin binding to the barley enzyme has been proposed only hypothetically since chitinase

crystals dissolve upon substrate binding. The crystal structure shows the active site nicely

fitting the substrate in an elongated cleft running though the length of the protein. Six

sugar binding sites, labeled A-F, have been identified in this cleft (Honda and Fukamizo,

1998). Catalysis requires that an acid (Glu 67) attacks the glycosidic bond at C4 oxygen

Chapter two

30

and a base (Glu 89) activates a water molecule that attacks C1 position of the sugar. The

catalysis occurs between sugar binding sites D and E of the enzyme. NMR analysis of the

cleavage products has shown that this enzyme acts by inversion of the configuration at the

anomeric carbon (Hollis et al., 1997).

Class I and II chitinase gene structure and regulation

A large number of cDNAs, but fewer gene sequences, have been obtained for PR-3 family

chitinases (Kasprzewska, 2003). The genomic sequences of class I and II chitinases show

none, one, or two introns. The first intron is located after the position corresponding to the

conserved catalytic site motif SHETTG whereas the second intron is located just before

the conserved motif NYNY. An exception to these locations is a class II chitinase gene

from Bermuda grass which has two introns but the first intron of 94 bases is located

upstream of the normal first intron at the catalytic site SHETTG motif (de los Reyes et al.,

2001). The introns are usually small, ranging in size from approximately 50-200 bases. An

unusual Beta vulgaris chitinase gene, however, has two introns of 2.5 and 1.5 kb

(Bergland et al., 1995). This particular gene codes for a chitinase with a relatively short

CBD but an unusually long spacer region of 131 amino acids, of which 90 are proline

residues, as compared to 5-22 amino acid spacers in the majority of class I chitinases. Any

significance of such a structure is unknown. Genomic structures of various chitinases

show that they exist as single-copy to large multi-gene families. For example, potato class

I chitinase genes (Ancillo et al., 1999), strawberry class II genes, and maize class I genes

(Wu et al, 1994) exist as one or two copies per haploid genome. Cotton (Chlan and

Bourgeois, 2001) and rice (Takakura et al., 2000) show complex genomic organizations of

chitinase genes with 4 or 8-10 members, respectively and these genes appear in clusters as

shown for Arabidopsis, cucumber, and potato. The gene expression is complex and varies

among different plant species. Chitinase genes show differential induction in various

plants upon challenge with pathogens or treatment with ET, JA, SA, and fungal elicitors

such as chitosan (Kasprzewska, 2003).

Currently, the study of signaling events from signal perception to the transcription of PR-

genes is an intense area of research. Several elicitor molecules capable of inducing PR-

gene expression have been characterised. These elicitors include β-glucans, peptides, and

avirulence (Avr) gene products of the pathogens (Neuhaus, 1999). Various secondary

messengers, including SA, ET, JA, and nitric oxide have been shown to be required in the

signaling events leading to PR-gene activation (Grant and Loake, 2000; Feys and Parker,

Literature review

31

2000; Glazebrook, 2001; Nürnberger and Scheel, 2001; Wendehenne et al., 2001). In

Arabidopsis, NPR1 (non expresser of PR-genes), an important switch controlling a

number of PR-genes and responding to SA and pathogens has been identified. NPR1 is an

ankyrin repeat protein presumed to mediate protein-protein interactions (Cao et al., 1997)

and has been shown to translocate to the nucleus upon stimulation (Dong, 1998). NPR1

appears to work downstream of SA and acts to negatively regulate SA production.

Similarly to SA, NPR1 is required for the establishment of SAR and expression of PR-

genes (Cao et al., 1994). ET and JA appear to function in a pathway different from SA

leading to the induction of a variety of genes including basic PR-genes, thionins and

defensins (Dong, 1998; Feys and Parker, 2000; Glazebrook, 2001). There is considerable

cross-talk between different pathways and activation of a particular PR-gene by a

particular pathway utilising certain second messengers seems to be dependent upon

individual pathogen/elicitor recognition events (Feys and Parker, 2000).

The signaling pathway mediating elicitor-inducible gene expression appears to be

conserved in distantly related plant species. The promoter of a pine class II chitinase gene

which is responsive to chitosan, a deacetylated derivative of chitin, in pine suspension

culture cells, mediated chitosan induced expression in tobacco plants (Wu et al., 1997).

The fact that a chitinase gene promoter from a gymnosperm showed similar regulation in

an angiosperm illustrates that at least the major components of signaling events in plants

are conserved. Chitinase gene expression is regulated at the transcription level

(Kasprzewska, 2003). The most direct evidence came from promoter studies of a bean

class I chitinase gene. A bean chitinase gene is activated by ET (75-100 fold),

oligosaccharide elicitors, and fungal pathogens (Broglie et al., 1986). The 1.6 kb promoter

region of this gene was able to confer the ET regulation on a uidA (GUS) reporter gene

transformed into tobacco plants (Broglie et al, 1989). The ethylene responsive element

was localised to the -422 to -195 region from the transcription start site of the gene. The

region contains two DNA sequences imparting quantitative expression and ethylene

response for the gene. The uidA gene was also induced when tobacco plants were

challenged with fungal pathogens and GUS activity closely correlated with the induction

of endogenous tobacco chitinase activity (Roby et al., 1990). The promoter was only

active at the areas of infection and the signal was sharply reduced away from the infection

site. Such ET responsiveness has also been studied in a class I tobacco chitinase promoter

(Shinshi et al., 1995) and was localised to the -503 to -358 region from the transcription

start site. This sequence, in either orientation, was sufficient to confer ET responsiveness

Chapter two

32

in a heterologous construct containing a cauliflower mosaic virus (CaMV) 35S RNA

promoter, suggesting that this sequence also acts as an enhancer. Within this region a 71

base sequence was further localised (-480 to -410) for ET regulation. This sequence

contains two GCC boxes, (TAAGAGCCGCC), which are frequently found in other PR-

gene promoters responding to ET. One such gene is a basic class I β-1, 3-glucanase gene

(Hart et al., 1993; Ohme-Takagi and Shinshi, 1990).

Induction of class I chitinase synthesis has been demonstrated through Northern and/or

Western blot analyses in a number of plants and cell cultures in response to various biotic

and abiotic stress factors. However, relatively few studies have been done for chitinases

belonging to other classes. In cultured tobacco cells, induction of a class I basic chitinase,

a class II acidic chitinase, and a class I β-1,3-glucanase have been investigated in response

to fungal elicitors from Phytophthora infestans (Suzuki et al., 1995). The class II acidic

chitinase gene was induced rapidly within 30 min of treatment and the induction reached a

maximum level at 4-5 h. The expression decreased to background levels within 24 h. In

contrast, the basic class I chitinase and the glucanase genes were induced after a 2 h lag

period, reaching a maximum level at 6 h and maintaining that level over a 24 h period. The

induction was shown to be dependent on protein synthesis for class I chitinase and

glucanase genes but not for class II chitinase gene. Also, inhibition of protein

phosphorylation prevented induction of basic class I chitinase and glucanase but had no

effect on class II chitinase. These results demonstrate that regulation of the expression of

these genes, at least in response to the elicitors studied, is accomplished through separate

signal transduction pathways. In a similar study, Kim et al. (1998) showed that the

induction of a rice class II acidic chitinase in response to fungal elicitors was repressed by

protein phosphatase 1 and 2A. The investigators suggest that protein dephosphorylation

might be a key step in the regulation of class II acidic chitinases, which is in agreement

with the results reported by Suzuki et al. (1995). It should be noted that differences in the

activation of rice class II chitinase gene were observed in rice cell culture and rice leaves.

The gene was induced by ethephon and HgCl2 in leaves but not in suspension cells. It was

induced by glycol chitin and fungal elicitors in suspension cells but not in leaves. Salicylic

acid and β-1,3-glucan had no effect in either system. This demonstrates that cell culture,

although a simple and an effective system may not reproduce the same results as at the

level of the organism.

Literature review

33

2.4.1.1.2. Class III chitinases

Class III chitinases are unique in that they have a structure unrelated to any other class of

plant chitinases (Neuhaus, 1999). These chitinases belong to the PR-8 family and family

18 of glycosyl-hydrolases. Members of family 18 glycosyl-hydrolases catalyze sugar

hydrolysis with the retention of configuration at the anomeric carbon. Class III chitinases

generally have lysozyme activity and appear to be more closely related to the bacterial

chitinases. A class III chitinase enzyme was purified from the seeds of Benincasa hispida

(white gourd), a Chinese medicinal plant (Shih et al., 2001). The enzyme is a 29 kDa

protein with 27 amino acid N-terminal signal peptide (as deduced from N-terminal amino

acid and genomic DNA sequences), directing its secretion into the apoplast. The length of

signal peptides and molecular weights are similar in other class III chitinases such as a

pumpkin chitinase of 29 kDa with a 27 amino acid signal peptide (Kim et al., 1999) and a

sugar beet chitinase of 29 kDa with a 25 amino acid signal peptide (Nielsen et al., 1993).

The pumpkin class III chitinase was purified by chitin affinity chromatography, which

showed strong retention of this protein. This is unusual for a class III chitinase since they

do not have a chitin-binding domain. Class III chitinases show a wide range of isoelectric

points, activity over a wide range of pH, and temperature stability at 60-70oC. The B.

hispida chitinase has a pH optimum of 2 and retains approximately 50% activity at pH 8

(Shih et al., 2001). Some class III chitinases, such as a yam enzyme, show two pH optima

and heat stability at 80oC (Tsukomoto et al., 1984). The three-dimensional structure of

heveamine, a chitinase/lysozyme from rubber tree, and its complex with the inhibitor

allosamodin has been determined (Scheltinga et al., 1995). The structure is an (α/β) 8

barrel similar to the bacterial family 18 glycosyl-hydrolases without significant sequence

identity. These enzymes contain a substrate-binding cleft located at the C-terminal end of

the β-strand in the barrel structure. The active site residue Glu127 of heveamine is

required for activity whereas Asp125 allows a wider pH range for catalysis. Heveamine

requires chitopentose as minimum substrate. The catalysis occurs by retention of

configuration at the anomeric carbon, and is substrate assisted. Generally class III

chitinases also act as lysozymes. However, Bokma et al. (1997) showed that heveamine

hydrolyzes the glycosidic bond of the peptidoglycan between C-1 of N-acetylglucosamine

and C-4 of N-acetylmuramic acid as opposed to lysozyme which catalyzes the hydrolysis

of peptidoglycan by cleavage of C-1 of N-acetylmuramic acid and C-4 of N-

acetylglucosamine. Therefore, heveamine and possibly other class III plant chitinases are

Chapter two

34

not strictly lysozymes. Some class III chitinases such as a sugar beet enzyme do not

exhibit lysozyme activity (Nielsen et al., 1993). A recent study shows kinetic constants of

heveamine by an improved assay method (Bokma et al., 2000). The km and kcat for N-

acetylglucosamine-pentamer (GlcNac)5 and (GlcNac)6 were measured to be 13.8 µM,

0.355/s and 3.2 µM, 1.0/s, respectively. Allosamodin was found to be a competitive

inhibitor with a Ki of 3.1 µM.

Class III chitinase genes in Sesbania rostrata (Goormachtig et al., 2001), Beta vulgaris

(Nielsen et al., 1993), Lupinus albus (Regalado et al., 2000), and Cucurbita sp. (Kim et

al., 1999) exist as single copies. In contrast, heveamine from Hevea brasiliensis is

encoded by a small multigene family (Bokma et al., 2001). Also, class III chitinase genes

from B. hispida (Shih et al., 2001) and H. brasiliensis (Bokma et al., 2001) lack introns.

Various class III chitinase genes showed distinct regulation upon stress treatment. For

example, a L. albus gene was shown to be induced by infection with Colletotrichum

gloesporioides, by treatments with UV light, and by wounding (Regalado et al., 2000). No

antifungal activity was observed for Trichosanthes kirilowii class III chitinase, and it was

not induced by salicylic acid (Savary et al., 1997). Both acidic and basic isoforms of

tobacco class III chitinases were induced upon infection of plants with TMV (Lawton et

al., 1992). The level of induction was about 5-10 fold and was observed in the infected

leaves as well as in secondary non-infected leaves. This suggests that class III chitinases

act as a generalized systemic-acquired resistance (SAR) response instead of being induced

in response to pathogen as has been seen for class I or II chitinases. A grape class III

chitinase was also shown to be induced in infected and non-infected leaves upon fungal

infection (Busam et al., 1997). The induction showed two maxima at 2 d and 6 d in the

susceptible Vitis vinifera whereas the level was steeply induced up to 4 d and declined to

the basal level by day 7 in the resistant Vitis rupestris. This gene showed SAR whereas a

class I gene analyzed simultaneously did not. A pumpkin class III gene was also

responsive to the fungal elicitor and glycol chitin (Kim et al., 1999). This gene showed

maximal induction within 1 h of fungal elicitor treatment and the transcript disappeared

within 6 h. In contrast, glycol chitin induced this gene at 3 h and the expression gradually

decreased to background level at 24 h. The gene was not induced by salicylic acid or by

UV irradiation. Protein accumulation took 6-10 h after transcription.

Literature review

35

2.4.1.1.3. Class IV-VII chitinases

Class IV, V, VI, and VII chitinases belong to the PR-3 family of pathogenesis-related

proteins. The structure of class IV chitinases is similar to class I chitinases except that they

are shorter due to four deletions in both the catalytic and chitin binding domain (CBD)

(Neuhaus, 1999). Class V, VI, and VII chitinases have unique structures and are

represented by one or a few members in each class.

2.4.1.1.4. Functions of plant chitinases

Plant chitinases have been known to be induced upon fungal infection and inhibit fungal

growth in vitro, which was initially used to implicate chitinases in plant defence

(Schlumbaum et al., 1986; Mauch et al., 1988). The induction of chitinases was initially

shown in pea plants infected with Fusarium solani, or challenged with other biotic or

abiotic stress factors (Mauch et al., 1988). Protein extracts made from infected pea plants

were able to inhibit growth of 15 of the 18 fungal species tested in vitro. Purified chitinase

inhibited growth of only one fungal species whereas a combination of chitinase and

another PR-protein, β-1,3-glucanase, inhibited the growth of all fungi tested showing a

synergism in activities (Mauch et al., 1988). Subsequently, a number of studies verified

these results in tobacco (Yun et al., 1996), grapes (Derckel et al., 1998), chickpea (Giri et

al., 1998), rice (Velazhahan et al., 2000) and other plants. The current view is that only

specific isoforms are induced in response to a particular pathogen and only certain

isoforms are able to inhibit specific fungi (Ji et al., 2000; Sela-Buurlage et al., 1993). For

example, a class I chitinase from tobacco showed antifungal activity against Fusarium

solani, but class II chitinases showed only a slight growth inhibitory effect when used with

high concentrations of a β-1,3-glucanase (Jach et al., 1995). Constitutive chitinase

expression is higher and induction is stronger and quicker in the resistant varieties as

compared to the susceptible varieties in some plant-pathogen systems such as sugar beet

(Nielsen et al., 1993), wheat (Anguelova et al., 2001) and tomato (Lawrence et al., 2000).

However, contrary data also exist showing no difference in the timing, induction, or

intensity of PR-gene expression in susceptible and resistant cultivars, for example in

cotton (McFadden et al., 2001). Quick response in the resistant cultivars might affect the

cell wall of germinating fungal spores, releasing elicitors leading to the expression of PR-

genes and disease resistance. It was shown for Alternaria solani that a basic chitinase was

only active on the germinating spores and not on the mature fungal cell wall for generation

Chapter two

36

of elicitor molecules able to induce disease resistance (Lawrence et al., 2000). The

difference may be in the length of these fragments as it is known that four or five N-

acetylglucosamine residues are necessary for elicitation of defence (Staehelin et al., 1995;

Montesano et al., 2003). In an interesting study in potato, it was shown that chitinase and

osmotin-like proteins interact with actin filaments (Takemoto et al., 1997). Since actin

filaments show cytoplasmic aggregation at the site of fungal penetration, it was

hypothesized that PR-proteins are translocated to the site of fungal penetration for

effective blocking of the pathogen and/or for the release of elicitors.

2.4.1.2. Application of chitinases in plant genetic engineering

Chitinase is capable of inhibiting the growth of the pathogen by lysing its hyphal tips

(Schlumbaum et al., 1986; Broglie et al., 1991). In addition to this direct action, the

released oligomers of N-acetylglucosamine could function as elicitors to amplify the

defence response in cells surrounding a site of infection (Ren and West, 1992). Thus,

genes encoding chitinases are attractive candidates for improving disease resistance.

Based on the antifungal activity of chitinases in vitro and in planta, genes coding for

chitinases have been cloned and expressed in different plants. Resistance based on

chitinase genes has been demonstrated in several crops such as tobacco (Jach et al., 1995),

Brassica napus (Broglie et al., 1991) and rice (Nishizawa et al., 1999). It has been

reported that transgenic tobacco plants that constitutively expressed a bean chitinase gene

showed increased resistance to the fungal pathogen, Rhizoctonia solani (Broglie et al.,

1993). The source, type and efficacy of some chitinase gene used in plant genetic

engineering are shown in the Table 2.1 below.

Table 2.1 Plant species genetically engineered with chitinase genes resulting in enhanced resistance to fungal diseases Plant species Expressed gene product Effect on disease development Reference Apple (Malus ×domestica)

Trichoderma harzianum endochitinase

Reduced lesion number and lesion area of Venturia inaequalis

Wong et al. (1999), Bolar et al. (2000)

Bentgrass (Agrostis palustris)

American elm (Ulmus americana) chitinase

Reduced growth and spread of Rhizoctonia solani

Chai et al. (2002)

Canola (Brassica napus)

Bean chitinase Tomato chitinase

Reduced rate of total seedling mortality caused by Rhizoctonia solani Lower percentage of diseased plants by Cylindrosporium concentricum and Sclerotinia sclerotiorum

Broglie et al. (1991) Grison et al. (1996)

Carrot (Daucus carota)

Tobacco chitinase

Reduced rate and final incidence of disease by Botrytis cinerea, Rhizoctonia solani, and Sclerotium rolfsii; no effect on Thielaviopsis basicola and Alternaria

Punja and Raharjo (1996)

Literature review

37

radicina Cotton (Gossypium hirsutum)

Trichoderma virens endochitinase

Inhibited growth of Rhizoctonia solani and Alternaria alternate

Emani et al. ( 2003)

Chrysanthemum (Dendranthema grandiflorum)

Rice chitinase Reduced lesion development of Botrytis cinerea

Takatsu et al. (1999)

Cucumber (Cucumis sativus)

Rice chitinase

Reduced lesion development due to Botrytis cinerea Growth of gray mold (Botrytis cinerea) suppressed

Tabei et al. (1998) Kishimoto et al. (2002)

Grape (Vitis vinifera)

Rice chitinase Trichoderma harzianum endochitinase

Reduced development of Uncinula necator and fewer lesions of Elisinoe ampelina Reduction of Botrytis cinerea development in preliminary tests

Yamamoto et al. (2000) Kikkert et al. (2000)

Peanut (Arachis hypogaea)

Tobacco chitinase Delayed lesion development and smaller lesion size of Cercospora arachidicola

Rohini and Rao (2001)

Pigeonpea (Cajanus cajan)

Rice chitinase

Resistance not tested Kumar et al. (2004)

Potato (Solanum tuberosum)

Trichoderma harzianum endochitinase

Lower lesion numbers and size of Alternaria solani; reduced mortality of Rhizoctonia solani

Lorito et al. (1998)

Rice (Oryza sativa) Rice chitinase Delayed onset and reduced severity of symptoms by Magnaporthe grisea Fewer numbers of lesions and smaller size due to Rhizoctonia solani

Nishizawa et al. (1999) Lin et al. (1995); Datta et al. (2000, 2001)

Rose (Rosa hybrida)

Rice chitinase

Reduced lesion diameter of black spot disease (Diplocarpon rosae)

Marchant et al. (1998)

Strawberry (Fragaria ×ananassa)

Rice chitinase

Reduced development of powdery mildew (Sphaerotheca humuli)

Asao et al. (1997)

Tobacco (Nicotiana tabacum)

Bean chitinase Serratia marcescens chitinase Serratia marcescens chitinase Rhizopus oligosporus chitinase Trichoderma harzianum endochitinase Baculovirus chitinase

Lower seedling mortality of Rhizoctonia solani; no effect on Pythium aphanidermatum Reduced development of Rhizoctonia solani Reduced disease incidence of Rhizoctonia solani on seedlings; no effect on Pythium ultimum Reduced rate of development and size of lesions on leaves by Botrytis cinerea and Sclerotinia sclerotiorum Reduced symptoms of Alternaria alternata, Botrytis cinerea, and Rhizoctonia solani Reduced lesion development of brown spot (Alternaria alternata)

Broglie et al. (1991, 1993) Jach et al. (1992) Howie et al. (1994) Terakawa et al. (1997) Lorito et al. (1998) Shi et al. (2000)

Tomato (Lycopersicon esculentum)

Wild tomato (Lycopersicon chilense) chitinase

Reduced development of Verticillium dahliae races 1 and 2

Tabaeizadeh et al. (1999)

Wheat (Triticum aestivum)

Barley chitinase Barley chitinase

Reduced development of colonies of Blumeria graminis f. sp. tritici Reduced development of colonies of Blumeria graminis and Puccinia Recondite

Bliffeld et al. (1999) Oldach et al. (2001)

2.4.1.3. Rice chitinase genes

Rice chitinase genes were isolated and characterised from both cDNA and genomic DNA

clones (Nishizawa et al., 1993). Previously, a chitinase gene Cht-1 was isolated from rice

cDNA library (Nishizawa and Hibi, 1991). Rice chitinase gene Cht-2 was later also

isolated and characterised from cDNA. Using three flanking sequences of Cht-1 and Cht-

Chapter two

38

2, screening genomic DNA library (λgt11) generated three genes Cht-1g, Cht-2g and Cht-

3g. The sequence of Cht-1 completely matched that of Cht-1g except for a poly(A) tail.

Cht-2 also completely matched Cht-2g except for a poly(A) tail, but in Cht-2g a 130 bp

intron was found within the codon for 208 amino acid. Cht-1g and Cht-3g had no introns.

Nucleotide sequence alignments showed 77% (Cht-1g/Cht-2g), 78% (Cht-2g/Cht-3g) and

90% (Cht-1g/Cht-3g) similarities. Cht-3g encodes a gene similar to the gene reported by

Huang et al. (1991). Cht-2 product was reported to accumulate in the intracellular space

whereas the Cht-3g expression was targeted in the extracellular spaces (Nishizawa et al.,

1999).

Basal expression levels of all three genes Cht-1g, Cht-2g and Cht-3g were quite low in

young leaves, but higher in young roots. In rice cell suspensions, basic levels of Cht-1 and

Cht-3g transcripts were higher than in leaves, but no Cht-2g transcripts were detectable.

The mRNA transcripts of the three chitinase genes were all about 1.2 kb long. RNA blot

hybridisation with the coding region of rice chitinase cDNA clone Cht-1 as a probe

showed that the rice chitinase mRNA levels increased after treatment of leaves and

suspension-cultured cells with stress-inducing compounds (Nishizawa and Hibi, 1991).

For instance, mercury chloride induced the expression of all three genes strongly in leaves,

but the transcripts of the Cht-3g gene did not accumulate following wounding or UV light

treatment, whereas both Cht-1g and Cht-3g were activated. In suspension-cultured cells,

mRNA levels of Cht-1g and Cht-3g were increased by treatment with glycol chitin, glycol

chitosan or pectic acids. The expression levels of the Cht-2g gene were still too low to be

detected in suspension-cultured cells, even after stress treatment. In all treatments studied,

Cht-1g and Cht-3g had very similar responses to external stimuli (Nishizawa et al., 1993).

Throughout the text Cht-3g is presented as Cht-3. Antifungal resistance based on rice

chitinase genes has been demonstrated in a range of crop species (Table 2.2).

2.4.1.4. Resistance based on rice chitinases

Rice endochitinase belongs to class I chitinase (Cht2 and Cht3) (Nishizawa et al., 1993), a

member of PR-3 plant chitinases (Datta and Muthukrishnan, 1999). Rice chitinase genes,

mostly Cht2, have been used in several crops to control diseases caused by both biotrophic

and necrotrophic fungal species. Table 2.2 shows crops that have been engineered with

rice chitinases and the type of fungus controlled.

In all these plants, evaluation in the greenhouse showed resistance levels ranging from the

highest to less or more susceptible than untransformed controls. Kishimoto et al. (2002)

Literature review

39

characterised lines as highly resistant, tolerant and susceptible. Line CR32 that had the

highest resistance level showed reduced appressoria formation and hyphae penetration into

leaves. On the other the hand, line CR3 had intermediate resistance. Fungal growth within

the leaf epidermal cells was also suppressed in the resistant lines CR32 and CR3, which

had higher chitinase expression and intracellular localisation.

Table 2.2 Chitinase genes that suppressed growth of both biotrophic and necrotrophic fungi resulting into reduced diseases development Plant species Chitinase

gene

Pathogen Nature of parasitism Reference

Cucumber

(Cucumis sativus)

Cht-2 Botrytis

cinerea

Necrotrophic Tabei et al. (1998)

Chrysanthemum

(Dendranthema

grandiflorum)

Cht-2 Botrytis

cinerea

Necrotrophic Takatsu et al. (1999)

Grape (Vitis vinifera)

Cht-2 Uncinula

necator

Biotrophic Yamamoto et al. (2000)

Rice (indica)

(Oryza sativa)

Cht-7 Rhizoctonia

solani

Necrotrophic Datta et al. (2001)

Cucumber

(Cucumis sativus)

Cht-2 Botrytis

cinerea

Necrotrophic Kishimoto et al. (2002)

Bent grass

(Agrostis palustris)

Hs2 Rhizoctonia

solani

Necrotrophic Chai et al. (2002)

Italian ryegrass

(Lolium multiflorum)

Cht-2 Puccinia

coronata

Biotrophic Takahashi et al. (2005)

In grape, higher chitinase expressing lines were more resistant (Yamamoto et al., 2000).

These lines showed suppressed conidial germination and malformed hyphae. Formation of

conidiophores was also suppressed. Evaluations of transgenic bent grass lines showed

similar resistance trends with higher expressers being more resistant to a biotrophic fungus

Puccinia coronata. Interestingly, chitinase activity did not correlate with antifungal

activity. Similar resistance presentations were observed in other plants, and in most cases

the fungal growth was arrested after penetration or death of one or two host cells. In

conclusion, the resistance observed was partial and quantitative.

It has been reported that other PR-proteins also show similar resistance patterns, suppress

fungal growth but do not completely prevent the invasion of the attacking pathogen into

the host plant cells (Takahashi et al., 2005).

Chapter two

40

2.4.2. Plant defensins

Defensins are widespread in plants and are expressed in tissues that provide a first line of

defence against potential pests and pathogens (Lay and Anderson, 2005). Plant defensins

belong to PR-12 family (Datta and Muthukrishnan, 1999) and are characterised with

cysteine-stabilised αβ motif that has representatives in vertebrates, invertebrates, and

plants (Lay and Anderson, 2005). This feature underscores the importance of these

defence molecules as central components of a widespread strategy of multicellular

organisms (Conceicao and Broekaert, 1999). In plants, antimicrobial plant defensins can

be constitutively expressed during developmental stages and in particular cell types

(Vanoosthuyse et al., 2001). Other plant defensin genes are expressed upon microbial

infection (Penninckx et al., 1996), drought (Do et al., 2004), heavy metals (Mirouze, et al.,

2006), and cold (Koike et al., 2002).

Plant defensins are small (about 5 kDa, 45 to 54 amino acids), basic, cysteine-rich proteins

(Mendez et al., 1990; Lay and Anderson, 2005). They are encoded by small multigene

families and are expressed in various plant tissues, but are best characterised in seeds. The

first members of plant defensins were isolated from the endosperm of barley (Mendez et

al., 1990) and wheat (Colilla et al., 1990) and were proposed to form a novel class of

thionins family (γ-thionin) that was distinct from the α- and β-subclasses (Bohlmann,

1994; Mendez et al., 1990; Colilla et al., 1990). Their classification as γ-thionin subclass

of the thionin family was based on the similarities in size (5 kDa) and similar number of

cysteines (Mendez et al., 1990), however their structure was significantly different from

those of α and β-thionins (Mendez et al., 1990; Colilla et al., 1990; Bohlmann, 1994).

Later, numerous other γ-thionin-like proteins were identified, either as purified proteins or

deduced from cDNAs from both monocots and dicots (Broekaert et al., 1995, 1997).

Several peptides were purified from a wide range of plant tissues including seeds, stems,

roots, leaves, and floral organs (Thomma et al., 2002). The term ‘plant defensin’ was

coined by Terras et al. (1995) after isolation and purification of two antifungal proteins

from radish seeds (Rs-AFP1 and Rs-AFP2) and the discovery that at the level of primary

and three-dimensional structure they were more related to insect and mammalian defensins

than to plant thionins (Terras et al., 1995). Plant defensins are distributed throughout the

plant kingdom and are likely to be present in most, if not all, plants (Broekaert et al., 1995,

1997; Osborn et al., 1995; Shewry and Lucas, 1997). Due to their antimicrobial effects in

vitro, plant defensins have been reported to be involved in plant defence response (Terras

Literature review

41

et al, 1995). To date, many plant defensins have been reported in several plant species

(Lay and Anderson, 2005). In Arabidopsis alone, 300 putative plant defensins have been

described (Sels 2007). In the current study, radish defensin gene Rs-afp2 (Terras et al.,

1995) was used in co-transformation with rice chitinases genes (Cht-2 and Cht-3)

(Nishizawa et al., 1993).

2.4.2.1. Radish defensin (Rs-AFP2)

Many different proteins with antifungal and/or antibacterial activity have been identified

and reported in seeds (Terras et al., 1995). These include chitinase (Roberts and

Selitrennikoff, 1986), β-1,3-glucanases (Manners and Marshall, 1973), thionins

(Fernandez et al., 1972), permatins (Vigers et al., 1991), and ribosome-inactivating

proteins (Leah et al., 1991). For example, antimicrobial chitin-binding lectin-like peptides

were identified from amaranth seeds (Ac-AMPs) (Broekaert et al., 1992) and a new class

of antimicrobial peptides was isolated from Mirabilis jalapa seeds (Mj-AMPs) (Cammue

et al., 1992). The first four classes of the above antimicrobial proteins are induced in plant

vegetative parts infected with fungi, and bacteria (Hedrick et al., 1988).

Two new classes of antifungal proteins (Rs-AFP1 and Rs-AFP2) were later isolated from

radish, Raphanus sativus L. seeds (Terras et al., 1992). These were highly basic

oligomeric proteins composed of small (5 kDa) polypeptides that are rich in cysteine.

Their oligomeric structures are stabilised by intact disulfide bridges.

The antimicrobial proteins isolated from radish, Rs-AFP1 and Rs-AFP2, have a broad

antifungal spectrum and are among the most potent antifungal proteins so far characterised

(Terras et al., 1992). Their activity on fungal growth in transgenic tobacco was previously

reported (Terras et al., 1995). Compared to many other plant antifungal proteins, the

antifungal activity of Rs-AFPs are less sensitive to the presence of salts in fungal growth

media and their antibiotic activity shows a high degree of specificity to filamentous fungi.

The amino-terminal regions of the Rs-AFPs show homology with the derived amino acid

sequences of the two pea genes specifically induced upon fungal attack, to γ-thionins and

to sorghum α-amylase inhibitors. Rs-AFP2 showed higher antifungal activity than Rs-

AFP1 and such activity is even more pronounced in medium with added salts. Constitutive

expression of Rs-AFP2 demonstrated enhanced resistance of tobacco plants to the fungal

leaf pathogen Alternaria longipes (Terras et al., 1995). Fungal cultures treated with Rs-

AFPs show characteristic claws of branched swollen hyphae however, no spore

germination is observed at higher concentrations (Terras et al., 1992).

Chapter two

42

2.4.2.2. Plant genetic engineering with plant defensins

To date, a number of plants have been transformed with plant defensin genes (Lay and

Anderson, 2005). A list of these genes, their recipient plants and target pathogens is

presented in Table 2.3. Constitutive expression of radish defensin (Rs-AFP2) enhanced

resistance of tobacco plants to the fungal leaf pathogen Alternaria longipes (Terras et al.,

1995). Canola (Brassica napus) expressing pea defensin, constitutively, has slightly

enhanced resistance against blackleg (Leptsphaeria maculans) disease (Wang et al., 1999).

The effective resistance conferred by defensin was obtained by constitutive expression of

alfalfa defensin (alfAFP) in potato against Verticillium dahliae (Gao et al., 2000). In that

report, fungus levels in transformed plants were reduced by approximately six-fold

compared to the non-transformed plants (Gao et al., 2000).

Table 2.3 Plant defensins in transgenic plants Transgene Source plant Recipient

plant(s) Promoter Increased resistance

against test organism(s) Reference

Rs-AFP2 Radish Tobacco CaMV 35S Alternaria longipes Terras et al. (1995)

Rs-AFP2 Radish Tomato, oil rape CaMV 35S A.solani, Fusarium oxysporum, Phytophthora infestans, Rhizoctonia solani, Verticillium dahliae

Koike et al. (2002)

AlfAFP Alfalfa Potato Figwort mosaic virus 35S

Verticillium dahliae Gao et al. (2000)

Spi1 Norway spruce

Tobacco, Norway spruce ECS cultures

CaMV 35S Erwinia carotovora, Heterobasidion annosum

Elfstrand (2001)

DRR230-a Pea Canola CaMV 35S Leptosphaeria maculans Wang et al. (1999)

DRR230-a DRR230-c

Pea Tobacco Duplicated CaMV 35S

F.oxysporum, Ascochyta pinodes, Trichoderma resei, Ascochyta pisi, Alternaria alternata

Lai et al. (2002)

BSD1 Chinese cabbage

Tobacco CaMV 35S P. parasitica Park et al. (2002)

WT1 Wasabi Rice Maize Ubiquitin-1

Magnaporthe grisea Kanzaki et al. (2002)

The protection provided by alfAFP transgene was not only maintained under glasshouse

conditions, but also in the field and over several years in different geographical sites (Gao

et al., 2000). Furthermore, the level of resistance in the transgenic plants was equal or

greater to the level of resistance obtained with non-transgenic plants grown in fumigated,

non-infested soil.

Literature review

43

2.5. Resistance through combinatorial expression of plant defence genes

Several defence-related genes encoding chitinases, glucanases, peroxidases and PR-

proteins are either constitutively expressed or induced upon pathogen infection (Shah et

al., 1997). The proteins encoded by these genes display in vitro antifungal activity,

suggesting a direct role in plant defence. Individually, some of these proteins impart

partial resistance to fungal pathogens in transgenic plants, however, the level of resistance

appears to be insufficient for practical use (Cornelissen and Melchers, 1993; Broglie and

Broglie, 1993). The fungal cell wall degrading enzymes chitinases and glucanases have

been examined extensively for their potential to give durable resistance to fungal

pathogens in transgenic plants. Synergistic in vitro antifungal activity between the basic

isoforms of tobacco chitinase and glucanase has been previously reported (Sela-Buurlage

et al., 1993). The combined expression of chitinase and glucanase in transgenic carrot and

tomato too was much more effective in preventing development of disease due to a

number of pathogens than either one alone (Jongedijk et al., 1995; Zhu et al., 1994),

confirming the synergistic activity of these two enzymes reported from in vitro studies

(Sela-Buurlage et al. 1993; Melchers and Stuiver, 2000). A few examples of fungal

disease resistance provided by co-expression of two different transgenes are given below

(Table 2.4).

Table 2.4 Expression of combined gene products in plant disease resistance development

Plant species engineered

Expressed gene product Effect on disease development Reference

Carrot (D. carota)

Tobacco chitinase + ß- 1,3-glucanase/osmotin

Enhanced resistance to Alternaria dauci, Alternaria radicina, Cercospora carotae, and Erysiphe heraclei

Melchers and Stuiver (2000)

Tobacco (N. tabacum)

Barley chitinase + ß- 1,3-glucanase, or chitinase + RIP Rice chitinase + alfalfa glucanase

Reduced disease severity to Rhizoctonia solani Reduced rate of lesion development and fewer lesions by Cercospora nicotianae

Jach et al. (1995) Zhu et al. (1994)

Tomato (L. esculentum)

Tobacco chitinase + ß- 1,3-glucanase

Reduced disease severity by Fusarium oxysporum f.sp. lycopersici

Jongedijk et al. (1995)

As a general rule, the deployment of genetic engineering approaches that involve the

expression of two or more antifungal gene products in a specific crop should provide more

effective and broad-spectrum disease control than the single-gene strategy (Lamb et al.,

1992; Cornelissen and Melchers, 1993; Strittmatter and Wegner, 1993; Jach et al., 1995;

Chapter two

44

Shah, 1997; Evans and Greenland, 1998; Salmeron and Vernooij, 1998; Melchers and

Stuiver, 2000).

2.6. Genetic modification of banana for Black Sigatoka resistance

Genetic engineering, using single gene and multiple gene insertions, has been reported in

banana. Mainly, two genetic modification systems were used and these were

Agrobacterium-mediated transformation and particle bombardment (May et al., 1995; Sagi

et al., 1995a, 1995b, 1998). The opportunity to develop transgenic bananas with fungal

disease resistance using biotechnology is of great demand because of the importance of

these diseases, but the technology is still in its infancy in this area. This is because genes

to impart resistance to fungal diseases particularly Black Sigatoka are either not available

or have yet to be demonstrated to be effective (Dale, 1999). The approach tried so far has

been the incorporation of antifungal peptide genes into transgenic bananas. Cammue et al.

(1992) reported antifungal proteins Mj-AFP2 isolated from seeds of Mirabilis jalapa and

Rs-AFP2 to be active in vitro against M. fijiensis. Bioassays showed 50% growth

inhibition when 0.5 µgmL-1 and 4 µgmL-1 of Mj-AFP2 and Rs-AFP2, respectively, were

applied together. Another study by Sági et al. (1995) reported the introduction of several

genes encoding AMPs into banana embryonic cell suspension cultures via particle

bombardment in order to generate transgenic plants with resistance to Black Sigatoka. In

1999, infections under controlled conditions of leaf discs excised from transgenic bananas

with mycelia of a pathogenic test fungus identified independent transgenic lines with high

tolerance (Remy et al., 1999).

2.7. Co-transformation in banana

Co-transformation, which leads to co-integration, has been mainly done via particle

bombardment-mediated transformation. This is basically achieved by coating a mixture of

vectors carrying different genes onto micro-carriers (metallic particles). Bombardment of

combinations of unlinked and linked chimeric genes, which resulted in high

transformation frequencies, was already reported in banana (Remy et al., 1998). Marziah

and Sreeramanan (2002), further reported co-transformation of rice chitinase (Cht-2) with

β-1,3-glucanase in a test for their synergistic activity in banana. With increasing interest in

gene pyramiding for durable resistance various approaches using Agrobacterium-mediated

gene transfer have been reported. Agrobacterium mediated co-transformation can be

Literature review

45

accomplished by using one plasmid with multiple T-DNAs (Depicker et al., 1985; Komari

et al., 1996) or separate plasmids with different T-DNAs which are contained in either one

(de Framond et al., 1986; Daley et al., 1998) or more Agrobacterium strains (Depicker et

al., 1985; McKnight et al., 1987; De Block and Debrouwer, 1991). In these systems co-

transformation frequencies of 19% to 85% were reported (McKnight et al., 1987; De

Block and Debrouwer, 1991; Kamari et al., 1996). The potential of Agrobacterium-

mediated co-transformation in banana was reported by Ahmed et al. (2002). In this study,

ECS of four banana cultivars were infected with two different A. tumefaciens strains each

carrying a distinct disarmed T-DNA containing one of the three reporter genes luciferase

(luc), β-glucuronidase (uidA), or green fluorescent protein (gfp) as well as the nptII

selectable marker gene. Multicellular structures expressing multiple genes were recovered,

and co-transformation frequencies were measured. The co-transformation frequency was

far less than the transformation frequencies involving each of the two genes. Significant

differences in (co-)transformation frequency were detected among the cultivars tested.

Materials and methods

47

Chapter 3. Materials and methods3.1. Genetic transformation systems, banana cultivars and cell cultures

Two transformation systems, developed for genetic modification of banana at the

Katholieke Universiteit Leuven (KULeuven), were applied in this study. These were

Agrobacterium-mediated Transformation (AmT) and particle bombardment-mediated

transformation (PmT) using a modified particle gun (Sági et al., 1995a). The specific

transgenes used included uidA (Jefferson et al., 1986) encoding β-glucuronidase (GUS)

enzyme; sgfpS65T (Chiu et al., 1996) encoding a synthetic green fluorescent protein

(GFP); nptII, neomycin phosphotransferase; hpt, hygromycin phosphotransferase (Van

den Elzen et al., 1985); Cht-2 and Cht-3 (Nishizawa et al., 1993) coding for rice

chitinases, and Rs-afp2 (Terras et al., 1992) for a radish defensin. These transgenes were

introduced into the dessert (AAA) bananas ‘Grand Naine’ (GN, ITC.1256) ‘Williams’ (W,

QDPI Will.0385) and ‘Gros Michel’ (GM, CIRAD) as well as to the plantain (AAB)

cultivars ‘Three Hand Planty’ (THP, ITC.0185), ‘Obino l’Ewai’ (OE, ITC.0109), and

‘Orishele’ (OR, ITC.0517). These cultivars are amenable to generation of Embryogenic

Cell Suspensions (ECS) using the ’scalp’ method (Dhed’a et al., 1991) and their ECS lines

exert high regenerability (Strosse et al., 2006). Preliminary evaluation showed that these

cell lines were repeatedly transformable and thus, suitable for our experiments. The choice

of material during the experiments often depended on which cultivar had sufficient

amount of ECS at the beginning of the experiments. For instance, ECS of ‘Gros Michel’

was used in the rice chitinase gene experiment because it is free from banana streak virus

(BSV) and is also an important dessert cultivar in Uganda.

ECS of ‘Grand Naine’, ‘Williams’, ‘Gros Michel’, ‘Three Hand Planty’ and ‘Orishele’

were maintained in liquid ZZ medium (Dhed’a et al., 1991) while ‘Obino l’Ewai’ ECS

were maintained in M1 medium (Escalant et al., 1994). Liquid ZZ medium is half-strength

MS medium containing 5 μM 2,4-D and 1 μM zeatin. M1 medium contained MS salts and

MS vitamins (Murashige and Skoog, 1962), 7 gL-1 agarose and was supplemented with 4.5

μM of 2,4-dichlorophenoxyacetic acid (2,4-D). ‘Grand Naine’, ‘Williams’, ‘Three Hand

Planty’, ‘Obino l’Ewai’, and ‘Orishele’ ECS lines were initiated from in vitro proliferating

meristems and maintained as described by Dhed’a et al. (1991) while ‘Gros Michel’ was

initiated from male buds (Côte et al., 1996) and kindly provided by CIRAD (Montpellier,

France). Cells were maintained on a rotary shaker (70 rpm) at 26±2°C under fluorescent

light and subcultured at an interval of 2 weeks.

Chapter three

48

3.2. Vectors and bacterial manipulations

3.2.1. Agrobacterium strains, binary and expression vectors

The three Agrobacterium tumefaciens strains used were EHA101 (Hood et al., 1986),

EHA105 (Hood et al., 1993) and AGLO (Lazo et al., 1993). These strains have the same

genetic background (Hood et al., 1993), i.e. they are all derivatives of the C58 nopaline-

type strain (Watson et al., 1975), and they all contain pTiBo542, a disarmed supervirulent

Ti plasmid (Hood et al. 1986). Previous experiments showed that these strains infect ECS

cells of a broad range of banana cultivars (Pérez Hernández et al., 1999, 2000). The use of

these strains, therefore, would not be another source of variation in our experiments.

EHA101 harbored binary vector pFAJ3000 whereas AGLO contained pBINUbi-

sgfpS65T. EHA105, which was used for co-transformation, contained binary plasmids

pBI333-EN4-RCC2, pBI333-EN4-RCG3, and pFAJ3494 or the combinations of pBI333-

EN4-RCC2/pFAJ3494 and pBI333-EN4-RCG3/pFAJ3494 (Figure 3.1).

Plasmid pFAJ3000 (De Bondt et al., 1994) contains an intron-interrupted uidA (GUS)

gene driven by the cauliflower mosaic virus (CaMV) 35S RNA promoter, and the nptII

selectable marker gene under the control of nos gene promoter. Plasmid pBINUbi-

sgfpS65T (Elliot et al., 1999) contains a synthetic gfp gene (Chiu et al., 1996) driven by

the maize ubiquitin gene promoter plus first intron (Christensen and Quail, 1996) and the

same chimaeric selectable marker gene as in pFAJ3000. The synthetic gfp gene construct

is optimised for codon usage in plants, and contains a mutation at amino acid position 65

to replace a serine residue with a threonine residue as well as a deletion of a cryptic intron

site found in wild type gfp (Heim et al., 1995). These modifications resulted in a 120-fold

increase in fluorescence and a single 490 nm fluorescence emission peak (Maximova et

al., 1998).

Binary vectors pBI333-EN4-RCC2 and pBI333-EN4-RCG3 (Nishizawa et al., 1999)

contain the hygromycin phosphotransferase (hpt) gene (Bevan, et al., 1983; Van den Elzen

et al., 1985) driven by the CaMV35S promoter, and each of two rice chitinase (Cht-2 and

Cht-3, see 2.4.1.3) genes fused to enhanced CaMV35S promoter. Binary plasmid

pFAJ3494 contains the Rs-afp2 radish defensin gene (Terras et al., 1995) and the

neomycin phosphotransferase (nptII) gene, controlled by the CaMV35S and mas gene

promoter, respectively.


49

Two expression vectors were employed for particle bombardment into banana (Figure

3.2): pAct1F-neo contains the rice actin gene promoter fused to the nptII selectable marker

gene, and pMy-Gus has the uidA reporter gene under the control of a promoter of the

‘Mysore’ isolate of banana streak virus (Schenk et al., 2001). The activities of My

promoter were close to CaMV35S promoter as uidA expression levels were comparable

(Schenk et al., 2001). Thus, the use of My promoter in the comparison of AmT and PmT

would not be a source of significant variation.

RB Pnos nptII Tnos PUbi sgfpS65T Tnos LB

H P E E

(c) pBINUbi-sgfpS65T

(e) pFAJ3494

Tnos Rs-afp2 PUbi Pmas nptII Tmas LB RB

H E E H E Sm

(d) pFAJ3000

H

RB T35S uidA-Int P35S Pnos nptII Tocs LB

S B H H

(a) pBI333-EN4-RCC2

P35S hpt T35S PEN4 Cht-2 Tnos LB RB

E H E S E

P35S hpt T35S PEN4 Cht-3 Tnos LB RB

E H S S E(b) pBI333-EN4-RCG3

Figure 3.1 Schematic presentation of T-DNA regions in binary vectors: (a) pBI333-EN4-RCC2, (b) pBI333-EN4-RCG3, (c) pBINUbi-sgfpS65T, (d) pFAJ3000, and (e) pFAJ3494. P35S and T35S, cauliflower mosaic virus 35S RNA promoter and poly(A) region; uidA-Int, intron-interrupted β-glucuronidase (GUS) gene; Pnos and Tnos, nopaline synthase gene promoter and poly(A) region; nptII, neomycin phosphotransferase gene; Tocs, octopine synthase gene poly(A) region; pUbi, maize polyubiquitin gene promoter and intron; sgfpS65T, synthetic gfp gene; hpt, hygromycin phosphotransferase gene; PEN4, enhanced CaMV35S promoter; Cht-2 and Cht-3, rice chitinase genes; Rs-afp2, radish defensin gene; Pmas and Tmas, mannopine synthase gene promoter and poly(A) region; RB and LB, right and left T-DNA borders. B, BamHI; E, EcoRI; H, HindIII; P, PstI; S, SacI; Sm, SmaI.

Chapter three

50

3.2.2. Growth and preparation of competent bacterial cells

Bacterial cultures were plated aseptically on selective medium. For all E. coli cultures, LB

medium (10 gL-1 tryptone, 5 gL-1 yeast extract, 10 gL-1 NaCl, and pH 7.0) was used.

Agrobacterium tumefaciens strains were incubated for 48 h on solid yeast-mannitol (YM)

medium (0.4 gL-1 yeast extract, 10 gL-1 mannitol, 0.5 gL-1 K2HPO4, 0.2 gL-1 MgSO4, 0.1

gL-1 NaCl, pH 7.0) and in liquid yeast-peptone medium (10 gL-1 yeast extract, 10 gL-1

peptone, 5 gL-1 NaCl). Single colonies of E. coli were cultured overnight at 37°C and 210

rpm whereas liquid Agrobacterium cultures were incubated at 28°C and 210 rpm for 30 h.

The appropriate antibiotics for binary vectors pBINUbi-sgfpS65T, pBI333-EN4-RCC2

and pBI333-EN4-RCG3 were 50 mgL-1 kanamycin (Km50) while for pFAJ3000 and

FAJ3494 a combination of spectinomycin at 100 mgL-1 (Sp100) and streptomycin at 300

mgL-1 (Sm300).

For the preparation of electro-competent cells of A. tumefaciens, 50 mL of fresh medium

were inoculated with 500 μL of overnight bacterial culture. At OD600 = 0.5-0.7 units (3-4

x 108 cell mL-1), cells were harvested by centrifugation at 17,900 x g and 4°C for 10 min.

Cell pellets were gently resuspended in 40 mL of ice cold, sterile Milli-Q water,

centrifuged as above and supernatant discarded. This step was repeated. Cell pellets were

gently resuspended in 8 mL of ice cold 10% (v/v) glycerol solution in Milli-Q water,

centrifuged at 16,500 x g for 25 min at 4°C and supernatant discarded. The pellets were

gently resuspended in 200 μL of ice cold, sterile 10% (v/v) glycerol solution in Milli-Q

water, then in 50 μL of aliquots dispensed into pre-cooled cryotubes and flash frozen in

liquid nitrogen before storage in -80°C freezer.

3.2.3. Plasmid DNA purification

Each vector (including the binary ones) was heat-shock transformed (see 3.2.4) into E. coli

for increased amount of DNA. Single bacterial colonies were picked and cultured in 5 mL

of selective LB medium (see 3.2.2). Cultures were incubated at 37°C with shaking at 210

(a) pAct1F-neo

nptII Tnos PAct1F

(b) pMy-Gus

PBSV-My uidA Tnos

Figure 3.2 Schematic representation of plasmid constructs for particle bombardment: (a) pAct1F-neo, (b) pMy-Gus. PAct1F, rice actin gene promoter; nptII, neomycin phosphotransferase gene; Tnos, nopaline synthase gene poly(A) region; PBSV-My, promoter of banana streak virus isolate from the cultivar ‘Mysore’; uidA, β-glucuronidase (GUS) gene.


51

rpm overnight. Plasmid isolation was done with the QIAprep Spin Miniprep Kit. Buffer

composition, isolation procedure, and other necessary chemical products are given in the

user instructional manual (QIAGEN, 2005). Briefly, the 5-mL cultures were centrifuged at

3000 x g for 5 min and the supernatant discarded. The pelleted bacterial cells were

resuspended in 250 μL of P1 buffer and transferred to a microfuge tube. The lysis reaction

was initiated by the addition of 250 μL of P2 solution. After gently inverting the tube four

times, proteins and polysaccharides were precipitated by the addition of 350 μL of P3

buffer. This was followed by centrifugation at 13,700 x g for 10 min after which plasmid

DNA in supernatant was loaded onto QIAprep spin column by centrifugation for 2 min at

17,900 x g. The column was then washed with buffers PB (500 μL) and PE (750 μL) by

centrifugation at 17,900 x g for 1 min, in each case. To elute plasmid DNA, 50 μL of

sterile water was added at 70°C. The column was placed into a 1.5 mL microfuge tube,

left to stand for 5 min, and centrifuged at 17,900 x g for 1 min. The isolated plasmid DNA

was stored at -20°C.

3.2.4. Heat shock transformation of E. coli cells

An aliquot of 50 μL of competent E. coli (see 3.2.2) was left on ice for 10 min. One to five

μL containing about 100 ng of plasmid DNA was added to the competent bacteria and the

tube was gently swirled and tapped for thorough mixing. After 30 min of incubation on

ice, the tubes were placed in 42°C water bath for exactly 30 sec without mixing or

shaking. At the end of incubation, the tubes were immediately placed on ice for 1 to 2 min.

Then, 250 μL of LB medium were added to the transformation mix and the bacteria were

incubated for 2 h at 37°C and 210 rpm to allow recovery from the heat shock and start

expression of the selectable marker gene. After 2 h of incubation, 100 μL of the culture

was plated on selective LB medium pre-warmed to 37°C and incubated overnight at 37°C.

Single colonies were then picked to grow cultures for plasmid purification and making

glycerol stocks for long-term storage.

3.2.5. Electroporation of Agrobacterium cells

The concentration of purified binary vectors (see 3.2.3) was adjusted to 100 ngμL-1 in

sterile water prior to (re-)electroporation. One μL of each binary vector (alone or in

combination) was gently mixed with a 50 μL aliquot of compentent cells and incubated on

ice for 2 min. The mixture was gently transferred into ice-cold electroporation cuvette,

Chapter three

52

with gentle tapping to ensure complete filling of the gap between the electrodes at the

bottom of the cuvette. Pre-cooled and dry cuvette was quickly placed into cuvette holder

and an electric pulse of 12.5 kVcm-1 was applied for 4-6 msec. Immediately, 1 mL of

liquid YM medium (Sambrook and Russel, 2000) was added, the mixture mixed

thoroughly and transferred into 15-mL Falcon tube. The electroporated mixtures were

incubated for 3 h at 28°C and 180 rpm to allow expression of the introduced antibiotic

resistance gene(s). Then, 100 μL aliquots were plated on selective YM medium followed

by 2-3 days of incubation at 28°C. Single colonies were randomly picked for re-growth,

plasmid purification, and restriction analysis prior to transformation into banana.

3.3. Agrobacterium-mediated transformation of banana

ECS in liquid ZZ (Dhed’a et al., 1991; Strosse et al., 2006) medium (for ‘Grand Naine’,

‘Williams’, ‘Gros Michel’, ‘Three Hand Planty’ and ‘Orishele’) and M2 (Côte et al.,

1996) medium (for ‘Obino l’Ewai’) medium were used in transformation experiments.

ECSs (200 μL) of 33.3% of settled cell volume were infected with Agrobacterium

tumefaciens cells, harbouring one or two binary vectors (in co-transformation), adjusted to

0.4 units of OD600 with liquid ZZ or M2 medium containing 200 μM acetosyringone (AS).

One mL of induced-diluted bacterial suspension was mixed with ECS in 24-well titre

plates and the plates were incubated for 6 h at 25 rpm in the dark. The infected ECS,

drained on 50 μm nylon mesh, were then transferred onto 10 mL AS-containing semi-solid

ZZ or M2 medium in 5-cm Petri dishes and co-cultivated for 6 days.

After co-cultivation, the cells were transferred to semi-solid ZZ or M2 medium containing

timentin (200 mgL-1) to eliminate Agrobacterium cells and supplemented with the

selective agent for transformed plant cells. ECS transformed with binary vectors

containing the nptII selectable marker gene (pFAJ3000, pBINUbi-sgfpS65T, and

pFAJ3494, Figure 3.1) were selected on medium containing geneticin (50 mgL-1), and

ECS transformed with the hpt gene (pBI333-EN4-ECC2 or pBI333-RN4-RCG3, Figure

3.1) were selected with hygromycin (50 mgL-1). Petri plates (5 cm in diameter) containing

transformed ECS on 50 μM nylon mesh were incubated in the dark at 25±2°C for 2

months, with subcultures every 2 weeks. The regeneration process involved transfer of

single transformed cell colonies to RD1 medium (Dhed’a et al., 1991) supplemented with

timentin (200 mgL-1) and geneticin or hygromycin (50 mgL-1), non-selective RD2 (Dhed’a

et al., 1991) medium and finally to REG medium to produce multiple shoots. Solid RD1


53

medium contained half-strength MS salts supplemented with 2.0 mgL-1 glycine, 10 mgL-1 ascorbic acid,

100 mgL-1 myo-inositol, 0.4 mgL-1 thiamine-HCl, 0.5 mgL-1 nicotinic acid, 0.5 mgL-1 pyridoxine-HCl, 30

gL-1 sucrose, 2.5 gL-1 gelrite at pH 5.8. RD2 medium was half-strength MS medium with 1 µM

benzyladenine (BA) and 100 mgL-1 myo-inositol, and REG medium was MS medium with

1 µM indoleacetic acid (IAA) and 1 µM benzyladenine. For each construct, a maximum of

24 independent transgenic events per cultivar were maintained on REG medium.

Following selection and subsequent regeneration of putatively transformed plants,

randomly selected lines that had roots were transferred into the greenhouse. Because these

lines were maintained on REG medium, which gives adequate shoot and root growth, no

separate rooting phase was done. Prior to planting, plantlets were washed to remove in

vitro growth medium. Hardening was done under polythene sheet for 2-3 weeks in order

to increase and maintain humidity.

3.3.1. The effect of physical parameters on transformation frequency

3.3.1.1. Length of infection time

ECS of ‘Grand Naine’ and ‘Three Hand Planty’ were transformed with A. tumefaciens

strain EHA105 harbouring pFAJ3000 (Figure 3.1) and infection times of 4, 6, 8, 10, 12,

and 14 h were investigated. After 6 days of co-cultivation, histochemical GUS assay was

done (see section 3.6) and blue spots counted in four replicates. The average numbers of

blue spots per treatment were calculated and indicated as the frequency of transformation

in a given banana cultivar cell line. The experiment was repeated to evaluate the

reproducibility of obtained results.

3.3.1.2. Age of ECS

Batches of ‘Obino l’Ewai’ ECS with different ages after the last subculture were

transformed with A. tumefaciens strain EHA105 harbouring pFAJ3000 (Figure 3.1). ECS

ages of 1, 3, 5, 7 and 9 days were investigated and the experiment was repeated. After 6

days of co-cultivation, the average number of blue spots per treatment was counted.

3.3.1.3. ECS volume during co-cultivation

‘Three Hand Planty’ ECSs were infected for 6 h with A. tumefaciens strain AGLO

harbouring pBINUbi-sgfpS65T (Figure 3.1). Prior to co-cultivation, ECS volumes of 50,

100, 200, 300, 600, and 1200 μL were plated in replicates on non-selective ZZ medium.

After 6 days of co-cultivation, independent samples from the same ECS volume group

Chapter three

54

were combined to make a total volume to 1200 μL and transferred onto selective ZZ

medium. Transient GFP expression was quantified as green fluorescent spots under MZ

FLIII stereomicroscope (Leica) equipped with GFP3 plant fluorescence filter, averages

were calculated and compared among ECS volumes and this experiment was also

repeated.

3.4. Particle bombardment-mediated transformation of banana

3.4.1. Preparation of ECS for particle bombardment

ECS (200 μL) of 33.3% settled cell volume were pipetted onto 1 cm2 of 50 μM sterile

nylon mesh. The liquid medium from the ECS was drained and cells were placed on solid

ZZ or M2 medium. The meshes with cells were then transferred onto a metallic circular

basement of the particle gun for bombardment with DNA coated tungsten particles.

3.4.2. Coating of microparticles and ECS bombardment

Plasmid DNA (Figure 3.2) was purified from E. coli by Qiagen midi-prep procedure

(QIAGEN, 2003). Coating of tungsten microparticles with plasmid DNA was done as

reported by Sági et al. (1995a) and the efficiency of coating monitored by viewing

particles under a fluorescence microscope after staining with Hoechst No. 33258. Then, 8

µL of coated particles were pipetted onto 200 µm iron mesh and accelerated in a home-

made particle gun by helium at a pressure of 8 kilobars (Sági et al., 1995a). Selection and

regeneration of transgenic plants was done as described in section 3.3.

3.5. Polyamines and plant regeneration

ECSs of ‘Three Hand Planty’ and ‘Williams’ were used to study the effect of the

polyamine spermidine on plant regeneration from ECS. ‘Three Hand Planty’ ECS had a

medium regeneration frequency (±50% of selected putative transformants regenerated into

plantlets), whereas less than 40% of putative transformants regenerated in ‘Williams’.

These two ECS were transformed with the Agrobacterium strain EHA101 harbouring the

binary plasmid pFAJ3000 (Figure 3.1). After 2 to3 months of selection on ZZ medium,

control untransformed and putative transformed cell colonies were individually transferred

onto RD1 medium and one month later to RD2 medium (RD1 medium without myo-inositol and

containing 1μM benzyladenine) supplemented with various spermidine concentrations in 24-

well plates. The SPD treatments and the number of cell colonies per treatment are


55

summarised in Table 3.1. After one month of culture on RD2 medium, the shoots

regenerated per spermidine treatment were counted and expressed as percentage of the

initial number of cell colonies transferred onto RD1 medium. Table 3.1 The number of cell colonies of ‘Three Hand Planty’ (THP) and ‘Williams’ (W) to be regenerated at various concentrations of spermidine (SPD) Cultivar Controls SPD concentration (mM) NT1 NT2 0.0 0.1 0.1 0.5 1.0 5.0 10.0 THP 96 96 96 96 96 96 96 96 96 W 96 96 96 70 70 70 70 70 70 NT1, untransformed colonies at 0 mM SPD; NT2, untransformed colonies at 0.1 mM SPD

3.6. Transient and stable uidA gene expression, histochemical GUS assay

Two or six days after co-cultivation for PmT and AmT, respectively, transformed ECS

were assayed for transient expression of the uidA gene. Three to four samples per

treatment were incubated in a staining solution containing 100 mM sodium phosphate (pH

7.0), 50 mM ascorbate, 0.1% Triton X-100, 0.4 mM potassium ferricyanide, 0.5 mM

potassium ferrocyanide and 1 mM 5-bromo-4-chloro-3-indolyl-ß-D-glucuronic acid (X-

Gluc) according to Jefferson (1987). Two thin sheets of sterile filter papers were placed at

the bottom of clean and transparent 15-cm diameter Petri dishes. A volume of 500-700 μL

of X-Gluc staining solution was added at the centre of the sterile filter papers.

Transformed or control ECS, plated on rectangular 50 μM nylon mesh, were transferred

onto wet filter papers. An additional 400 μL of X-Gluc staining solution was added onto

each sample, and then Petri dishes were covered, sealed with plastic film, and incubated

for 4-5 h at 37°C. Stained samples were left overnight at room temperature prior to

counting blue spots. Blue foci in each transformation system were counted and an average

of three to four plates per treatment was calculated. The quantitative data were analysed

with Statistix 8.0 software (Analytical Software, Tallahassee, FL, USA).

After 2 to 3 months of selection and after shoot regeneration transformed cell colonies and

leaf tissues, respectively, were tested for stable uidA expression. The final solution in this

case contained 100 mM Tris-HCl (pH 8.0), 0.5 mM potassium ferricyanide, 0.5 mM

potassium ferrocyanide, 1% (w/v) ascorbic acid, 10 mM Na2EDTA.2H2O, 0.2% (v/v) 3-

[(3-cholamidopropyl) dimethylammonio]-1-propane-sulfonate (CHAPS), and 1 mM X-

Gluc. Pieces of leaves, leaf sheaths and corms were incubated overnight at 37°C in 1 mL

of this solution. Two untransformed samples were stained as negative controls. ECS

competence to transformation was expressed as the average number of blue spots counted.

Chapter three

56

3.7. Molecular characterisation of transformants

3.7.1. PCR analysis

3.7.1.1. DNA isolation for PCR analysis

Total DNA was extracted from transformed plants and untransformed controls with the

modified miniprep protocol of Dellaporta et al. (1983). Thirty mg of leaf tissue, mixed

with 500 µL of extraction buffer [100 mM Tris-HCl/pH 8.0, 50 mM EDTA, 500 mM

NaCl, 10 mM β-mercaptoethanol, 2% (w/v) PVP (polyvinyl pyrrolidone, MW 10,000)]

was ground in a 1.5 mL microfuge tube. SDS was added to a final concentration of 1.32%

(w/v), the mixture vortexed for 30 sec, and incubated at 55°C for 10 min. Potassium

acetate was added to a final concentration of 1.17 M and the mixture again vortexed

followed by centrifugation at 17,900 x g for 10 min to precipitate cell debris, proteins, and

polysaccharides complexed with precipitated potassium dodecylsulphate. Without

destabilizing the precipitated pellet, the supernatant was transferred to a 1.5 mL microfuge

tube. The centrifugation was repeated to remove the remaining proteins and/or

polysaccharides. DNA precipitation was done by the addition of an equal volume of

isopropanol to the supernatant. The mixture was vortexed and the DNA collected by

centrifugation at 4500 x g for 10 min. The precipitated DNA was washed by brief

centrifugation (4500 x g, 5 min) in 70% ethanol and air dried for 30 min. The DNA pellet

was finally resuspended in 20 μL of Milli-Q water containing 1 mgmL-1 of RNase, treated

for 15 min at 37°C. DNA was either used directly in PCR analysis or stored at -20°C.

3.7.1.2. PCR conditions

All PCR reactions were performed in 0.2 mL microfuge tubes with the Mastercycler

GradientTM (Eppendorf) cycler in a final volume of 20 μL made up of 2 μL of plant DNA

template and 18 μL of master mix. The master mix consisted of 1 x MgCl2 containing

PCR buffer (QIAGEN), 0.25 mM dNTPs, the specific primers (Table 3.2) at 2 μM of final

concentration, and 0.025 UμL-1 Taq DNA polymerase. Eighteen microliters of this master

mix was transferred into 0.2 mL PCR tubes (Eppendorf) and mixed with 2 μL template

DNA from each of 10 putatively transformed lines. Reactions were started with initial

denaturation (94°C for 2 min) and subjected to 35 cycles as follows: 1 min at 94°C, 30 sec

at respective annealing temperature (Table 3.2), and 1-2 min at 72°C. The last extension


57

phase was prolonged to 7 min at 72°C. The plasmid vectors as positive controls as well as

two negative controls (water and untransformed plant DNA) were included in each

experiment. Gel electrophoresis was done with different agarose concentrations (1 to

1.3%, w/v) depending on the size of the expected PCR product (Bio-Rad, M1704400AB).

PCR gene specific probe DIG labeling was done as per manufacturer’s instructional

manual (Roche, 2003).

Table 3.2 List of gene-specific primer pair sequences and amplified fragment sizes

Gene Primer pair Expected fragment size (bp)

Annealing temp. (oC)

actin 5’-ACC GAA GCC CCT CTT AAC CC-3’ 5’-GTA TGG CTG ACA CCA TCA CC-3’

170 56

Cht-2 5´-GCG GGT TCT ACA CCT ACG AG-3´ 5´-GCG TCA TCC AGA ACC ACA-3´

405 56

Cht-3 5´-CCG CTA AGG GCT TCT ACA-3´ 5´-GCG TCA TCC AGA ACC AGA AC-3´

414 56

hpt 5´-ACT TCT ACA CAG CCA TCG GTC-3´ 5´-GAC CTG CCT GAA ACC GAA CTG-3´

668 56

nptII 5´-GAG GCT ATT CGG CTAT GAC TG-3´ 5´-GGC CAT TTT CCA CCA TGA TA-3´

554 58

sgfpS65T 5’-ATGGTCAGCAAGGGCGAGGAG-3’ 5’-TTACTTGTACAGCTCGT CCAT-3’

719 62

Rs-afp2 5´-TTA ACA AGG AAA GTA GCA GAT AC-3´ 5´-GAG GCT ATT CGG CTA TGA CTG-3´

180 56

uidA-Intron

5’-CAACAGACGCGTGGTTACAG-3’ 5’-GTTCGCCGATG CAGATATTC-3’

632 56

uidA 5’-CAACAGACGCGTGGTTACAG-3’ 5’-GTTCGCCGATG CAGATATTC-3’

443 56

3.7.1.3. Multiplex PCR (MPCR) analysis

Pure DNA for MPCR analyses was isolated using the modified protocol of Dellaporta et

al. (1983) combined with the DNeasy Plant Mini Kit. The details of the buffers, and other

necessary chemical products are given in DNeasyR Plant Handbook (QIAGEN, 2006).

Fresh leaf samples of 300 mg were collected from Cht-3/Rs-afp2 co-transformants in the

greenhouse, homogenised in liquid nitrogen with mortar and pestle, and treated as

described in 3.7.1.1. The supernatant treated with RNase (200 µgmL-1 final concentration)

was loaded, in several folds of 500 μL, into QIAshredder spin columns, centrifuged at

17,900 x g for 2 min and flow-throughs collected in 15 mL tubes. DNA binding buffer

AP3 was added, tubes inverted a few times before loading the mixture onto DNeasy Mini

Spin columns. The DNA on the columns was washed a few times with wash buffer AW,

air dried and eluted twice with 100 μL of preheated (55°C) sterile Milli-Q water. Quality

of DNA was observed after electrophoresis on 0.8% (w/v) agarose gel prior to PCR

amplification.

Chapter three

58

The transgenes in the putative Cht-2 or Cht-3/Rs-afp2 co-transformants were hpt, nptII,

Cht-2 or Cht-3, and Rs-afp2. Primer pairs, specific for these genes (Table 3.2) were

factorially combined in duplexes and triplexes to evaluate the formation of primer dimers

as well as competition of primers and amplified fragments during MPCR cycles. MPCR

was initiated with a denaturation at 94°C for 4 min and then subjected to 35 cycles as

follows: 1 min at 94°C, 1 min at 56°C, and 2 min at 72°C. The last extension phase was

prolonged to 7 min at 72°C. Primers of hpt and nptII were at 0.5 μM equimolar

concentrations whereas primers for Cht-3 were used at 0.75 μM. Specificity of primers

was controlled with equimolar mixture of plasmids pBI333-EN4-RCG3 and pFAJ3494 as

positive controls while water and untransformed plant DNA were used as negative

controls. MPCR products were separated on 1.3% (w/v) agarose gels.

3.7.2. Reverse Transcriptase (RT)-PCR analysis

3.7.2.1. RNA isolation and DNase treatment

RNA was isolated from leaf tissue of in vitro cultured transformants using RNeasy Plant

Mini Kit (Qiagen) with some modifications. Three hundred mg of leaf tissue was

homogenized with mortar and pestle in liquid nitrogen. The fine powder was transferred

using a fine spatula to pre-cooled 15 ml tubes, which were kept on ice to reduce RNA

degradation. Pre-cooled 900 μL of lysis buffer RLT supplemented with 20 mg of PVP

and 10 μL of β-mercapto-ethanol was immediately added. After 1 min of vortexing, the

samples were blended twice for 2 sec with Ultraturrax. After completing the procedure,

6.7 μL of 10 mM Na-citrate was added and the RNA solution was stored at -80°C.

RNA yield was determined fluorometrically using SYBR Green II (Molecular Probes).

Briefly, 200 μL of SYBR Green solution [1:10,000 dilution in 1 x TAE (0.04 M Tris-

acetate, 0.001 M EDTA)] was mixed with 4 μL of 1:10 diluted RNA sample and

fluorescence was measured at 527 nm after excitation at 497 nm. An RNA molecular

weight marker with known concentration (RNA molecular weight marker III, Invitrogen)

was used as standard. A denaturing RNA gel was run to monitor the integrity of RNA

samples.

For DNase treatment, the following components were added to the volume of a sample

containing 20 μg RNA, 5 μL of Mn-buffer (0.66 mM MnCl2 and 10 mM Tris-HCl/pH

7.8), 10 μL of 2 UμL-1 DNase I (Ambion) and the final volume was adjusted to 50 μL with

RNase-free water. Samples were incubated at 37°C for 30 min. Immediately after the

incubation, 10 μL of 15 mM EDTA was added and the RNA solution was incubated at


59

75°C for 5 min to de-activate or denature DNase I (Ambion). DNase treatment was

verified by performing PCR with actin-specific primers. Negative PCR results indicated

complete DNA removal.

3.7.2.2. cDNA synthesis

First strand cDNA synthesis, using RNA template was done using the OmniscriptTM RT

Kit (QIAGEN). Briefly, 6 µL of DNase-treated and denatured RNA was added to 14 µL of

master stock containing 2 µL of RT buffer, 2 µL of 5 mM dNTPs, 10 μM dT12-18 primer, 1

μL of 26 UμL-1 Anti-RNase, 1 μL of 4 UμL-1 Omniscript RT, and the volume was

adjusted to 14 μL with RNase-free water. The reaction was incubated in a 37°C water bath

for 1 h.

3.7.2.3. PCR amplification of transcripts

All RT-PCR reactions were performed in 0.2 mL microfuge tubes with the Mastercycler

Gradient (Eppendorf) cycler in a final volume of 20 μL consisting of 2 μL of the first

strand cDNA reaction and 18 μL of master mix. The master mix contained the gene

specific primers (final concentration of 2 μM) and Hot Start Taq polymerase (0.5 U per

reaction). The following cycling program was followed: initial heating at 95°C for 15 min

to denature the RNA-cDNA duplex and to inactivate the reverse transcriptase followed by

35 cycles consisting of denaturation at 94°C for 30 sec, primer annealing at 56°C (uidA

and nptII) or 62°C (gfp) for 30 sec and elongation at 72°C for 1 min (uidA and nptII) or 2

min (gfp) before a final elongation at 72°C for 4 min. A positive control (plasmids

pFAJ3000 and pBINUbi-sgfpS65T) and two negative controls (water and cDNA from an

untransformed plant) were included in each analysis.

3.7.3. Southern hybridisation analysis

3.7.3.1. DNA isolation for Southern analysis

Total banana DNA was isolated using a combined protocol (Dellaporta et al., 1983;

Aljanabi and Martinez, 1997), and purification on Qiagen minicolumns. Leaf tissues of

approximately 3 g were harvested in the greenhouse, wrapped in labeled aluminium foil,

and quickly dipped in liquid nitrogen. About 1 g of sample was homogenised in liquid

nitrogen with mortar and pestle, and transferred into 50 ml centrifuge tubes. The plant

powder was then mixed with 4 mL of extraction buffer [100 mM Tris-HCl/pH 8.0, 50 mM

EDTA, 500 mM NaCl, 10 mM β-mercaptoethanol, and 2% (w/v) PVP] and vortexed

Chapter three

60

briefly. Then, 429 µL of 20% (w/v) SDS was added and the mixture was incubated at

55°C for 10 min. Proteins and carbohydrates were later precipitated by adding 1.3 mL of 5

M potassium acetate. All centrifugations were performed at 4°C and 16,500 x g. After

vortexing the mixture was centrifuged for 10 min. Then, 3 mL of 6 M NaCl was added to

the supernatant, followed by 30 sec of vortexing. The mixture was centrifuged again and

the supernatant treated with RNase (200 µgmL-1 final concentration), incubated for 15 min

at 37°C, and centrifuged for 30 min. Centrifugation was repeated when the supernatant

contained visible plant debris. An equal volume of chloroform-isoamylalcohol (24:1) was

added and tubes were gently inverted a few times before centrifugation for 5 min. The

upper aqueous phase was transferred to a new tube. This step was repeated when the two

phases were not separated. An equal volume of isopropanol was finally added to the

supernatant and DNA was precipitated at -20°C overnight followed by 30 min of

centrifugation. The DNA pellet was air-dried and dissolved in 1 mL sterile water. The

DNA solution was mixed with column binding buffer AP3 and loaded onto the DNeasy

Mini spin column. The DNA-bound columns were washed with buffer AW (QIAGEN,

2006), air-dried and DNA was eluted with 100 µL of pre-heated sterile water. DNA

concentration was determined by ND-1000 spectrophotometer (NanoDrop Technologies,

Wilmington, DE, USA) or in Spectra Fluor Reader (Tecan) using Pico-Green (Molecular

Probes, Inc.) and adjusted to 0.5 μgμL-1. DNA integrity was then observed by

electrophoresis on 0.8% (w/v) agarose gel.

3.7.3.2. DNA digestion and one copy reconstruction

Restriction digestion of individual DNA samples was carried out under the conditions

suitable for the respective restriction enzyme(s). The choice of the enzyme depended on

which restriction enzyme cuts the vector once. For Southern analysis of a given transgene,

10 μg of total banana DNA were digested with 50 units of restriction enzyme (New

England Biolabs) overnight at 37°C. The enzymes were: BamHI for pFAJ3000; HindIII

for pBINUbi-sgfpS65T, pBI333-EN4-RCC2, pBI333-EN4-RCG3 and pMy-Gus; and

SmaI for pFAJ3494 (see Figure 3.1). The number of integration sites was estimated by

counting the number of band signals detected per transgenic line and the number of

transgene copies estimated by comparing band intensities with specific copy number

standards.

In the determination of transgene copy numbers, a specific gene fragment was excised

from the integrated T-DNA by double restriction digestion, which was then hybridised


61

with a gene-specific probe. For copy numbers of Cht-2 and Cht-3, double digestions were

performed with the following enzymes: HindIII and SacI for pBI333-EN4-RCC2, HindIII

and EcoRI for pBI333-EN4-RCG3.

The number of transgene copies integrated per transgenic line was calculated based on

established genome sizes and weight equivalents of DNA. In this approach, the haploid

genome size of banana is approximated to be 600 Mbp (Lysák et al., 1999), and 1 pg DNA

to be equivalent with 103 Mbp. Thus, 1 µg DNA from a triploid banana (with a genome

size of 1800 Mbp) contains 109 Mbp divided by 1800 Mbp = 5.55 x 105 genomes. The

amount of vector DNA to be added per µg of untransformed banana DNA for the

establishment of a copy number standard sample is then calculated as follows:

No. of genomes x vector size (bp) x 1 pg

µg DNA 109 bp .

From this relationship, the amount of vector DNA equivalent to one copy in a plant transformed with pFAJ3000 (12 kb) will be:

5.55 x 105 genomes x 12,000 (bp) x 1 pg

µg DNA 109 bp

Therefore, in 10 µg total DNA of a transformed triploid banana 66.6 pg of vector DNA

corresponded to one copy.

3.7.3.3. Blotting, hybridisation and detection with non-radioactive probes

Southern hybridisation analysis was performed according to standard protocols, including

(i) electrophoresis on 0.8% (w/v) agarose gel for 5 h at 40 V to separate the digested total

DNA fragments, (ii) the transfer of separated fragments to a positively charged nylon

membrane (Roche) by downward capillary blotting; (iii) labelling of probes with

digoxigenin-dUTP (Roche) by PCR, and (iv) prehybridisation, hybridisation and detection

of hybridised fragments on the nylon membrane with the CSPD chemiluminescent

substrate according to the manufacturer’s instructions (Roche) and by image capturing

with a cooled CCD camera (Roper Scientific). For estimation of copy numbers, digital

images were analyzed by Image J software (http://rsb.info.nih.gov/ij). Grey intensities and

grey areas in transgenic banana lines were compared to established and reconstituted

vector DNA copy standards.

= 6.66 pgµg-1 DNA.

Comparison of transformation methods

63

Chapter 4. Comparison of transformation methods

4.1. Introduction

A comprehensive comparison of Agrobacterium-mediated (AmT) and particle

bombardment-mediated transformation (PmT) was carried in several banana cultivars. The

choice of these transformation methods depended on their reported use and efficiency in

banana and their simplicity for plant genetic engineering. For example, genetic

engineering via electroporation is limited by the need to isolate and regenerate protoplasts,

and resulted in lower transient transformation frequency (Sági et al., 1994, 1995).

Moreover, its use has not been reported in many banana cultivars. In the current study, the

presence, integration, transcription and translation of the introduced genes were verified

by various techniques.

4.2. Transient gene expression in AmT and PmT systems

In the Agrobacterium-mediated transformation system, ECSs of cultivars GN, THP, OE,

and OR were co-cultivated for 6 days with the EHA101 strain harbouring pFAJ3000.

Then, samples were selected randomly and histochemically stained for the expression of

uidA gene (Figure 4.1).

ECSs bombarded with plasmid DNA-coated particles (pMy-Gus + pActin1Fneo) were

immediately transferred onto non-selective ZZ medium, supplemented with 5 µM 2,4-D

(section 3.1) to enhance cell division and thus facilitate transgene integration. Two days

after bombardment, transient expression of the uidA reporter gene was assayed

histochemically in the embryogenic cells (Figure 4.2).

Figure 4.1 Histochemical assay for transient expression of the uidA gene (EHA101/pFAJ3000) in Agrobacterium-transformed ECSs of ‘Grand Naine’, ‘Three Hand Planty’, ‘Obino l’Ewai’, and ‘Orishele’ after 4-hr incubation at 37°C followed by overnight incubation at room temperature. The scale bar is equivalent to 1mm.

‘Grand Naine’ ‘Three Hand Planty’ ‘Obino l’Ewai’ ‘Orishele’

Chapter four

64

Histochemical staining usually showed high levels of GUS expression with the AmT

system. This could be attributed to the presence of an intron in the N-terminal region of

the uidA coding sequence of pFAJ3000. The presence of introns resulted in efficient

splicing of pre-mRNA and increased stability of mRNA, which enhanced gene expression

levels in various plants such as rice (Kyozuka et al., 1990). Tanaka et al. (1990) reported

that a uidAINT gene increased the level of GUS enzyme activities 10- 40-fold and 80-90-

fold compared with the intron-less plasmid in transformed protoplasts and tissues,

respectively.

To effectively compare gene transfer efficiency associated with AmT and PmT systems in

banana, quantitative analysis of blue foci obtained by histochemical GUS assay of

transformed ECSs was performed. The graphical presentation of transient GUS expression

results, expressed as the number of blue foci, is shown in Figure 4.3. Though there was no

overall statistical difference (P≤ 0.3314) between the two gene transfer systems, for GN

only, transient GUS expression was significantly higher (P≤ 0.0002) for AmT than with

the PmT system. Transient GUS expressions did not differ significantly (P≤ 0.2541) in

THP, while PmT was slightly more efficient in OE and OR (P≤ 0.05).

‘Grand

‘Orishele’ ‘Obino l’Ewai’ ‘Three Hand Planty’ ‘Grand Nain’

0200400600800

1000120014001600

GN THP OE OR

Banana cultivar

No.

of b

lue

foci

A-MTP-MT

Figure 4.3 Transient GUS expression in ECSs of ‘Grand Naine’ (GN), ‘Three Hand Planty’ (THP), ‘Obino l’Ewai’ (OE), and ‘Orishele’ (OR), transformed with pFAJ3000 via Agrobacterium (AmT) or with pMy-Gus by particle bombardment (PmT). Mean±SE of at least four replications.

Figure 4.2 Histochemical assay for transient expression of the uidA gene (pMy-Gus) in bombarded ECS of four cultivars after 4-hr incubation at 37°C followed by overnight incubation at room temperature. The scale bar is equivalent to 1mm.


65

4.3. Stable transformation frequencies in AmT and PmT systems

4.3.1. Embryogenic cell colonies

Transformed ECSs obtained with both transformation processes, were transferred to

selective culture media and incubated in the dark at 25±2°C for 2 to 3 months. After 3

weeks in culture, ECSs turned brown due to necrosis and massive death of untransformed

embryogenic cells. One month later, numerous whitish cell clumps (embryogenic cell

colonies) appeared on the surface. This response occurred in all cultivars with increasing

time in culture. Embryogenic cell colonies were quantified, picked, and transferred onto

selective RD1 medium. Significant differences in the number of surviving embryogenic

cell colonies was observed between gene transfer systems and among the cultivars used

(Figure 4.4).

The selection regime used was highly effective since no surviving colonies were ever

observed on plates of untransformed ECS controls. The overall number of surviving

colonies significantly (P≤ 0.001) showed that AmT was superior to PmT. The biggest

significant difference (P≤ 0.003) between both transformation systems was observed in

GN and OR, with lower but still significant difference (P≤ 0.05) in the cultivars THP and

OE.

Within each gene transfer system, cultivars showed variable numbers of surviving

embryogenic cell colonies (Figure 4.4). In the AmT system, GN exerted the highest

survival with OR giving the lowest response. In the PmT system, which generally gave a

much lower colony survival, THP exhibited the highest colony survival followed by OE,

with the lowest response again in the cultivar OR. GN, which had the highest colony

survival in the AmT system, responded significantly (P≤ 0.001) lower with PmT than

THP.

0

1020

3040

50

6070

8090

100

GN T HP OE OR

Cultivars

No.

of c

olon

ies

A-MTP-MT

Figure 4.4 Number of colonies surviving after a 2-month selection of GN, THP, OE, and OR, transformed via Agrobacterium (AmT) or particle bombardment (PmT). Mean±SE of 10 replications.

Chapter four

66

A smaller amount of transformed cells expressing a given transgene is frequently observed

with the PmT system. This feature is reported to be associated with shock waves, sound

waves, and cellular membrane injuries caused by the particle gun treatment (Houllou-Kido

et al., 2005), which then accounts for a reduced number of shoots regenerated (Russel et

al., 1992).

4.3.2. Regenerated plants

Variable numbers of plantlets were recovered after AmT and PmT of ECSs of GN, THP,

OE and OR. The number of plantlets regenerated was influenced by cell line

characteristics and the gene transfer system used (Figure 4.5). In all experiments, 24

plantlets were retained. Based on the number of transgenic colonies (Figure 4.4) and the

number of shoots regenerated (Figure 4.5), AmT was significantly (P≤ 0.000) more

efficient than PmT system except in THP where slightly more shoots were observed in

PmT system.

One of the critical points in plant genetic engineering is the selection and recovery of

transgenic shoots (Joersbo, 2001). This is because embryogenic cell colonies appearing

during the selection procedure must survive the stress induced by the combination of

media and antibiotics used to kill both Agrobacterium and untransformed plant cells. In

this study, embryogenic cell colonies that survived the selection procedure were quantified

0

10

20

30

40

50

60

70

GN THP OE OR

Cultivar

No.

of

shoo

ts r

egen

erat

ed

AmT

PmT

Figure 4.5 Regeneration of putatively transformed ECS clones of ‘Grand Naine’ (GN), ‘Three Hand Planty’ (THP), ‘Obino l’Ewai’ (OE), and ‘Orishele’ (OR) after 2 months on selective media.


67

and analysed (Figure 4.5). However, such embryogenic cell colonies showed lower

regeneration frequencies than those from untransformed control ECS. Low regeneration

frequencies in both genetic transformation systems are reported to be due to accumulation

of toxic compounds from necrotic untransformed tissue or ECS (Lindsey and Gallois,

1990). Though the selection pressure was identical in both the AmT and PmT systems,

significantly higher (P≤ 0.004) regeneration frequencies were observed in the AmT

system. Such higher survival of colonies on selective medium (Figure 4.4) and higher

number of plants regenerated in AmT than PmT system (Figure 4.5) indicates that the

AmT as a method of choice for transforming banana.

4.3.3. Grouping banana cultivars based on transformation competence and

regeneration

Based on the quantifiable transient reporter gene expression data (influenced by gene

transfer efficiency and ECS competence) and the number of cell colonies after two months

on selection medium (indicative for regeneration potential or regenerability), the ECS

lines of the four banana cultivars were categorised (Table 4.1).

Table 4.1 Categories of ECS lines of the four banana cultivars after 2 months on selection medium Transformation competence

Regeneration competence

High Medium Poor High ‘Grand Naine’ Medium ‘Obino l’Ewai’ ‘Three Hand Planty’ Poor ‘Orishele’

On the basis of the above data OR regenerative response is inferior, while OE appears to

have superior regenerability among the cultivars tested. GN and THP are intermediate in

performance. The transformation competence of GN is considered to be highest.

4.4. Characterisation of transgenic lines from AmT and PmT systems

4.4.1. Histochemical GUS assay of transformed lines

Plantlets transformed with the uidA gene via the AmT or PmT system were regenerated

and tested for histochemical localisation of the ß-glucuronidase (GUS) enzyme. The GUS

assay substrate penetrates and diffuses at variable rates depending on the type and age of

tissues. Two types of solutions were used depending on the banana tissue tested. For

Chapter four

68

transient gusA expression, a standard GUS assay solution (Jefferson, 1987) was employed

on the transformed ECS. However, this had very low diffusion and penetration rates in

organised or differentiated plant tissues. To enhance the level of GUS assay substrate

penetration into regenerated banana tissues, the CHAPS detergent, which solubilises

membrane proteins and breaks protein-protein interactions (Herbert, 1999), was included.

The different banana tissues tested (leaves, corm and leaf sheathes) showed different

patterns of blue staining (Figure 4.6).

Expression of the uidA gene, indicative of correct transcription and translation, showed

variations between the two gene transfer systems. In both systems, intense blue staining

was readily observed in the GUS assay of leaf pieces, leaf sheaths, and segments of corms

(Figure 4.6). Lines generated through the AmT system showed higher and consistent

frequencies of uidA expression with cultivars GN and OE at 95%, THP at 83%, and OR at

81% (Table 4.2). Variations in transformation frequencies based on uidA expression are

frequently reported to be cultivar or genotype dependent as for example in Texas rice

cultivars, where expression frequencies ranged between 0 and 87% (Dong et al., 1996).

For Agrobacterium-mediated transformation of Brassica napus L. cultivars ‘Sarow-4’ and

‘Semu-249’, a difference of 50% was observed in GUS expression with cultivar ‘Sarow-4’

showing the highest expression frequency of 61% (Moghaieb et al., 2006). This variation

is attributed to differential necrosis, hypersensitive response, and subsequent cell death in

host plant species (Hansen et al., 2000; Khanna et al., 2004) and reported to account for

varied plant transformation efficiencies (Potrykus, 1990; Goodman and Novacky, 1994).

The PmT system, on the other hand, showed rather low and more varied expression of the

uidA gene. Frequencies of uidA expression in the PmT system were in all cultivars about

50% lower: GN showing only 57%, THP 33%, OR 0%, and OE, which had 95% uidA

expression frequency in AmT, exerting 47% only (Table 4.2). Low frequency of GUS

GN, AmT GN, PmT GN, AmT

ABB

CC

Figure 4.6 Histochemical GUS assay with X-Gluc-CHAPS of pieces of leaf (A), corm (B) and leaf sheaths (C) from regenerated transgenic ‘Grand Naine’ (GN) lines after AmT (pFAJ3000) and PmT systems (pMy-Gus). The scale bar is equivalent to 1mm.


69

expression could be due to non-uniform coating of microparticles and death of

bombarded cells due to injuries. The PmT system has sometimes been reported to be

genotype independent (Dai et al., 2001). Table 4.2 Regenerated banana plantlets after Agrobacterium (AmT) and particle bombardment-mediated transformation (PmT) tested for histochemical uidA gene expression

AmT PmT Cultivar TP (%) +ve TP (%) +ve ‘Grand Naine’ 24 95 07 57 ‘Three Hand Planty’ 24 83 24 33 ‘Obino l’Ewai’ 23 95 21 47 ‘Orishele’ 11 81 - -

However, genetic engineering research on wheat (Sonriza et al., 2001), maize (Kennedy et

al., 2001) and Chinese rice cultivars (Tang et al., 1999) generated variable gene

expression frequencies and numbers of putative transformants. The gene transfer might be

genotype independent but cell division and cycling (Asako et al., 1991), expression, and

plant cell regeneration (Bailey et al., 1993; Droste et al., 2001; Sakhonokho et al., 2004)

are genotype and/or cultivar dependent. Evidences of genotype and/or line dependence on

cell regeneration have also been reported in banana ECS regeneration (Strosse et al.,

2006). Genotype dependence of transformed banana ECSs in expressing GUS was earlier

reported by Sági et al. (1995a) in PmT lines of ‘Williams’, THP and ‘Bluggoe’. Though

no regeneration data was reported, transient GUS expression revealed genotype

dependence with cultivar ‘Bluggoe’ showing the highest number of blue foci (over 800),

followed by ‘Williams’ with 400-500 and the least (100) blue foci observed in THP.

4.4.2. PCR analysis in AmT and PmT generated transformants

4.4.2.1. PCR analysis in AmT system

A 554-bp fragment representing the coding region of the neo gene and a 632-bp part of the

uidA gene was amplified by PCR among putatively transformed banana lines using PCR.

These lines included cultivar THP lines Ab.3.2.08, Ab.3.4.09, Ab.3.4.15, Ab.3.4.30,

Ab.3.4.31, Ab.3.4.34, Ab.3.4.37, Ab.3.4.64, Ab.3.4.76, Ab.3.4.94, and Ab.3.4.96. For

‘Obino l’Ewai’ lines were Ab.3.2.19, Ab.3.2.46, Ab.3.2.51, Ab.3.2.54, Ab.3.2.55,

Ab.3.2.60, Ab.3.2.61, Ab.3.2.68, Ab.3.2.72, and line Ab.3.2.91.

TP, Total Plants analysed; (%) +ve, percentage of plants with positive GUS

Chapter four

70

Ten putatively transformed lines (Fig.4.7) of THP and OE showed 100% co-existence of

both uidA and neo genes, which is indicative of the effective co-integration into the

banana genome. This is expected since nptII and uidA are on the same T-DNA. Similarly

high co-occurrence of genes in transgenic plant lines has been observed and reported in

many crops including rice (Dai et al., 2001; Al-Forkan et al., 2004), banana (Sági et al.,

1995a; Remy, 2000), and switch grass (Somleva et al., 2002).

Putatively transformed lines from GN and OR were also analysed for the presence of both

uidA and neo genes (Figure 4.8). Randomly selected lines of GN were included Ab.3.1.08,

Ab.3.1.14, Ab.3.1.26, Ab.3.1.41, Ab.3.1.45, Ab.3.1.50, Ab.3.1.53, Ab.3.1.72, Ab.3.1.93,

and line Ab.3.1.94. Lines Ab.3.1.50 (GN) and Ab.3.3.83 (OR) gave negative results both

for gusA and nptII genes (Figure 4.8). These lines could be escapes or the PCR

amplification could have been inhibited by contaminants in their template DNA samples.

Figure 4.7 PCR analysis of plants regenerated after AmT of ‘Three Hand Planty’ (A) and ‘Obino l’Ewai’ (B). Upper panel and lower panel are for nptII and uidA genes, respectively; M, 1-kb ladder; +Co1 and +Co2, positive controls (pFAJ3000 and pFAJ3006, both containing an gusA gene with and without an intron, respectively); -Co and W, negative controls (untransformed plant and water, respectively); lanes 1 to 10 are independent regenerants which included Ab.3.4.34, Ab.3.4.94, Ab.3.4.31, Ab.3.4.64, Ab.3.4.96, Ab.3.4.76, Ab.3.4.15, Ab.3.4.37, Ab.3.4.09, and Ab.3.4.30 for THP; and Ab.3.2.91, Ab.3.2.55, Ab.3.2.60, Ab.3.2.51, Ab.3.2.19, Ab.3.2.68, Ab.3.2.72, Ab.3.2.61, Ab.3.2.54, Ab.3.2.08, and line Ab.3.2.46 for OE.

A Three Hand Planty

M 1 2 3 4 5 6 7 8 9 10 +Co1 W +Co2

632 bp

gusA

M 1 2 3 4 5 6 7 8 9 10 W +Co1 -Co

B Obino l’Ewai (OE)M 1 2 3 4 5 6 7 8 9 10 +Co1 W +Co2

nptII 554 bp

M 1 2 3 4 5 6 7 8 9 10 W +Co1–Co+Co2

G1 G2


71

Putative transformants of the banana cultivar OR also showed the presence of both uidA

and neo. The other nine regenerated lines (Ab.3.3.05, Ab.3.3.6, Ab.3.3.30, Ab.3.3.50,

Ab.3.3.71, Ab.3.3.73, Ab.3.3.80, Ab.3.3.91, and Ab.3.3.94) all showed positive signals for

both genes with a co-transformation rate of 90% (Figure 4.8B).

4.4.2.2. PCR analysis in P-mT system

Results from histochemical gusA gene expression (Table 4.2) were confirmed by PCR

analysis of the gusA and nptII genes. The cultivar OE, showed a transformation frequency

of 80% among randomly selected lines. Transformed lines Pb.1.2.04, Pb.1.2.10, Pb.1.2.15,

Pb.1.2.17, Pb.1.2.18, Pb.1.2.25, Pb.1.2.26, Pb.1.2.28, Pb.1.2.35, and Pb.1.2.36 were

analysed with PCR for the presence of both gusA and nptII genes (Figure 4.9). PCR

analysis of all these putatively transformed lines gave 100% presence of gusA gene

compared to 80% frequency for the nptII gene with lines Pb.1.2.28 and Pb.1.2.35 showing

negative signals. PCR results of putatively transformed lines Pb.1.4.08, Pb.1.4.11,

Pb.1.4.39, Pb.1.4.42, Pb.1.4.44, Pb.1.4.54, Pb.1.4.55, Pb.1.4.62, Pb.1.4.89, Pb.1.4.90

showed 100% presence of the gusA gene. However, nptII was not detected (50%) in lines

Pb.1.4.44, Pb.1.4.55, Pb.1.4.62, Pb.1.4.89, and line Pb.1.4.90.

Figure 4.8 PCR analysis of plants regenerated after AmT of ‘Grand Naine’ (A) and ‘Orishele’ (B). Upper panel and lower panel are nptII and uidA genes, respectively; M, 1-kb ladder; +Co, positive control (pFAJ3000 containing an gusAgene with an intron); W, water as negative control; lanes 1 to 10 areindependently regenerated lines and included Ab.3.1.45, Ab.3.1.72, Ab.3.1.14, Ab.3.1.50, Ab.3.1.08, Ab.3.1.26, Ab.3.1.53, Ab.3.1.94, Ab.3.1.41, and line Ab.3.1.93 for GN; Ab.3.3.94, Ab.3.3.30, Ab.3.3.80, Ab.3.3.05, Ab.3.3.91, Ab.3.3.73, Ab.3.3.50, Ab.3.3.83, Ab.3.3.71 and Ab.3.3.6 for OR.

gusA

632 bp

A Grand Nain B Orishele M 1 2 3 4 5 6 7 8 9 10 +Co M 1 2 3 4 5 6 7 8 9 10 W +Co

nptII

554 bp

M 1 2 3 4 5 6 7 8 9 10 W +Co M 1 2 3 4 5 6 7 8 9 10 +Co

Chapter four

72

Depending on what is coated on the micro-carriers (linked transgenes or transgenes on

separate vectors), variable transformation frequencies have been reported. During

transgene stacking in potato, 45% of transgenic lines had all transgenes when transgenes

were located on separate vectors but the frequency reached 70-80% when genes were

linked on the same vector (Romano et al., 2003). Though higher transformation

frequencies have been reported in cases where linked transgenes were used, the results are

highly variable. Transformation frequencies of 17-33% were reported in oat (Cho et al.,

1999); 43% for linked gusA and nptII genes in Phaseolus vulgaris (Kim and Minamikawa,

1996), 90% for linked uidA and hpt in Catharanthus roseus (Hilliou et al., 1999), 40% in

barley (Koprek et al., 1996), and 67-79% for unlinked uidA and nptII genes in sugarcane

(Bower et al., 1996). The efficiency of PmT system shown here, therefore, is in the range

of what has been reported in other plant species.

4.4.3. RT-PCR analysis of transformants generated via AmT and PmT systems

Though stable integration of transgenes into the plant host genomes is indicated by PCR

analyses, expression of such transgenes needs to be further confirmed. Transgene

expression is first assessed by performing RT-PCR, which confirms effective

transcription.

Six putatively transformed lines from AmT and PmT systems were selected and analysed

for effective transcription of the gusA gene. In all cases tested, the amplification product

obtained by RT-PCR analysis was identical to the one of the positive PCR plasmid control

Figure 4.9 PCR analysis of plants regenerated after PmT of ‘Obino l’Ewai’ (A) and ‘Three Hand Planty’ (B). Upper panel and lower panel are neo and gusA genes, respectively; M, 1-kb ladder; +Co1 and +Co2, positive controls (pFAJ3000 and FAJ3006 both containing an gusA gene with and without intron, respectively); W, water as negative control; lanes 1 to 10 are independently regenerated lines which included Pb.1.2.25, Pb.1.2.15, Pb.1.2.04, Pb.1.2.10, Pb.1.2.18, Pb.1.2.36, Pb.1.2.26, Pb.1.2.28, Pb.1.2.17, and Pb.1.2.35 for OE; and Pb.1.4.42, Pb.1.4.55, Pb.1.4.90, Pb.1.4.11, Pb.1.4.39, Pb.1.4.54, Pb.1.4.08, Pb.1.4.62, Pb.1.4.89, and Pb.1.4.44 for THP.

W

nptII

554 bp

gusA 632 bp

M 1 2 3 4 5 6 7 8 9 10 +Co

B Three Hand Planty

M 1 2 3 4 5 6 7 8 9 10 +Co1 +Co2

A Obino l’Ewai

M 1 2 3 4 5 6 7 8 9 10+Co1 +Co2 M 1 2 3 4 5 6 7 8 9 10 +Co


73

(632 bp for gusA with intron, and 443 bp without intron) indicating that the gusA gene was

properly transcribed in the tested transformants. The RT-PCR reaction performed on the

control PCR master mix yielded no product. Of equal importance, PCR analysis of

DNase-treated RNA samples with actin-specific primers (Act1F and Act1R) did not result

in any signal, indicating that the samples were not contaminated with DNA (data not

shown). However, in Figure 4.10B where RNA samples were not treated with DNase,

amplification products of different sizes were observed. The amplification products

included 443 bp and 632bp fragments. Thus, depending on the case, correct transcriptions

products (small PCR product of 443 bp) were found alone or in combination with genomic

DNA (632 bp); i.e. correct transcription processes occurred with effective intron clipping

in the transformed lines Ab.3.3.05, Ab.3.3.30, Ab.3.3.91 and Ab.3.3.94) but in Ab.3.3.71

there was contamination with genomic DNA.

In Figure 4.10A, Particle-mediated transformed lines Pb.1.2.04, Pb.1.2.15, Pb.1.2.18,

Pb.1.2.25 and line Pb.1.2.36 were positive for correct and effective gusA transcription,

whereas line Pb.1.2.28, which gave a negative signal with PCR analyses, was again

negative indicating that it could have been an escape. RT-PCR analysis in PmT system

was done among selected lines of OE because no transformants were regenerated from

OR.

Chapter four

74

4.4.4 Southern analysis of transgenic lines from AmT and PmT systems

Integration patterns of the introduced gusA gene into the genome of geneticin-resistant and

GUS-expressing banana plantlets derived from AmT and PmT systems were analysed by

Southern blot assays (Figure 4.11). To detect the number of integrations, genomic DNA

was digested with HindIII, which cuts once within the T-DNA of the binary vector

pFAJ3000. Digests of respective genomic DNA samples were electrophoretically

separated, transferred onto nylon membrane, and probed with 0.6 kb fragment from gusA

coding region. The mobility of bands differed in most transgenic lines, indicating that

these lines represent different transgenic events. Number of integrations varied between

two and six in the PmT system. Line Pb.1.1.18 showed six integrations whereas lines

Pb.1.1.06 and Pb.1.1.19 showed four integration loci. The integration pattern of Pb.1.1.06

and Pb.1.1.19 was similar so that they are presumed to originate from the same

transformation event. The lowest integration number (two integrations) in PmT system

was detected in line Pb.1.1.13. Variable integration and copy numbers in PmT system

have been observed in other crops as well. A range of one to four integrations were found

in japonica rice (Dai et al., 2001). On the other hand, Wakita et al. (1998) reported over

seven copies and integration loci in the same crop. Inglis et al. (2000) observed over 14

Figure 4.10 RT-PCR analysis of gusA gene expression in transgenic lines of ‘Obino l’Ewai’ (A) generated by PmT (upper panel) and ‘Orishele’ (B) by AmT (lower panel). M, 1-kb ladder; +Co1 and +Co2, positive controls (pFAJ3000 and FAJ3006 both containing a gusA gene with and without intron, respectively); W, water as negative control.

B Orishele

+Co2

Ab.3.3.30

Ab.3.3.50

Ab.3.3.05

Ab.3.3.91

Ab.3.3.94

Ab.3.3.71

+Co1

M

632 bp

443

443 bp

+Co2

W

Pb.1.2.25

Pb.1.2.04

Pb.1.2.28

Pb.1.2.18

Pb.1.2.36

Pb.1.2.15

M

A Obino l’Ewai


75

integration loci in biolistic transformation of Metarhizium anisopliae whereas a maximum

of five gusA integration loci were reported in wheat (Sonriza et al., 2001).

The integration loci varied from one to four in the AmT system with line Ab.3.1.14 giving

the highest number of integrations. Lines Ab.3.1.50 and Ab.3.1.26 both had one

integration locus of gusA. In general, the AmT system resulted in a simpler integration

pattern than the PmT system, which is in agreement with other authors. Independent of

host plant and/or transgene used, simple gene integration patterns have been reported and

these include in grapes, Cht-2 (Yamamoto et al., 2000); in garlic, gusA (Zheng et al.,

2004); cotton, gusA/nptII Haq, 2004); and in chickpea, Cry1Ac (Sanyal et al., 2005).

4.5. Conclusion

The absence of biological constraints (host-range) and the ability to target any cell type

make the PmT system uniquely versatile (Altpeter et al., 2005). However, the associated

physical injuries to cells and tissues (Houllou-Kido et al., 2005) tremendously reduce the

frequencies of subsequent regenerated transgenic lines. Our results with the PmT system

showed also low transformation frequencies in the banana cultivars GN, THP, OE and

21 kb

5 kb

2 kb

MW

MIII

-VeC

o

Pb.1.1.19

Pb.1.1.11

Pb.1.1.13

Pb.1.1.06

Pb.1.1.18

Ab.3.1.50

Ab.3.1.26

Ab.3.1.14

Figure 4.11 Southern blot analysis showing integration patterns of the gusA gene in transgenic lines of ‘Grand Naine’ from PmT (Pb) and AmT (Ab) systems. For each sample, 10 μg genomic DNA was digested with HindIII and probed with 0.6 kb gusA fragment. For PmT, genomic DNA was digested with SmaI. MW III, DIG labelled DNA sizing marker; and -VeCo, non transformed control.

Chapter four

76

OR. Regeneration and PCR analysis of AmT transformed banana cultivars (the same as

for the PmT) showed higher transformation frequencies and lower number of integration

sites (one to four). Though the PmT system will continue to play an important role in plant

biology and crop biotechnology (Altpeter et al., 2005), regeneration, integration and

expression results from the current study indicate that the AmT is the method of choice for

the genetic modification of a wide range of banana (Musa spp.) cultivars.

Optimisation of AmT system

77

Chapter 5. Optimisation of AmT system

5.1. Introduction

Gene transfer mechanisms with Agrobacterium are complicated (Zambryski, 1992;

McCullen and Binns, 2006), genotype specific (Bauer et al., 2002) and influenced by a

range of physical and biological factors (Wang et al., 2005; Yong et al., 2006). The

Agrobacterium-mediated transformation (AmT) system is frequently improved for

increased transformation frequency in many crops by optimising parameters like the

Agrobacterium strain (Cheng et al., 2004; Yong et al., 2006), bacterial cell density

(Amoah et al., 2001; Yong et al., 2006), length of pre-culture and co-cultivation phase

(Wang et al., 2005). Other factors investigated are: vector type (Amoah et al., 2001),

osmotic treatment (Uze et al., 1997), infection period and acetosyringone concentration

(Amoah et al., 2001; Srivatanakul et al., 2001; Clercq et al., 2002), co-cultivation

temperature (Dillen et al., 1997), and wounding method (Yong et al., 2006). Some of

these factors were previously considered in the development of an AmT procedure for

banana (Perez Hernandez et al., 2006). There is a need to evaluate parameters that could

increase the applicability of this procedure to a wide range of banana cultivars. Such

parameters would, specifically, be targeted at increasing both T-DNA transfer and enhance

its integration into the banana genome.

In the current study, physical parameters expected to affect T-DNA transfer and

integration in the AmT system were optimised for banana cells. Investigated physical

parameters included infection length, ECS volume during the co-cultivation phase (to

enhance T-DNA transfer), and age of ECS (influences T-DNA integration). The effect of

the polyamine spermidine on shoot regeneration was also evaluated. These studies were

performed primarily with the β-glucuronidase (uidA) gene (Jefferson et al., 1987). Making

use of the improved protocol, four banana cultivars (GN, THP, OE, and OR) were

transformed with a modified green fluorescent protein (sgfpS65T) gene.

Chapter five

78

5.2. Optimising physical parameters for improved transformation frequency

5.2.1. Length of infection period

The effect of infection length on transformation frequency was investigated in GN and

THP. ECSs were infected with Agrobacterium tumefaciens strain EHA105 containing

binary vector pFAJ3000 (Figure 3.1, section 3.2.1). In both cultivars an increase in

transient GUS expression was observed with prolonged infection time. However, beyond

10 h (GN) and 6 h (THP) it was difficult to distinguish positive single cells from stained

cell clumps, which precluded quantification (Figure 5.1, Table 5.1). A consistently higher

transient GUS expression was observed in THP than in GN (Table 5.1), which is likely to

be a characteristic of the particular cell line.

Table 5.1 Number of blue foci counted after different infection times in Agrobacterium-mediated transformed ‘Grand Naine’ (GN) and ‘Three Hand Planty’ (THP) ECS

Infection length (h) 4 6 8 10 12 14 GN 209.7 ± 24 272.0 ± 41 365.7 ± 28 922.7 ± 21 >1500 >1500 THP 1169.3 ± 150 1311.7 ± 95 >1500 >1500 >1500 >1500

4 h 6h 8 h

10 h 12 h 14 h

Figure 5.1 Schematic presentations of the effects of infection length during AmT in ‘Grand Naine’. The experiment was repeated twice. The scale bar is equivalent to 1 mm.

Mean±SE of at least four replications


79

5.2.2. Effect of ECS age

In this experiment also the uidA gene was used and similar rationale, explained in section

5.2.1, was applied in choosing a suitable reporter gene. In general, ECS competence to

AmT increased with the age of ECS (i.e. the number of days after subculture, Figure 5.2).

Transformation frequency, expressed as the number of blue foci observed at the transient

level, significantly increased from day 1 till day 7 beyond which it dropped (Figure 5.2

and 5.3). High transformation frequencies were also observed in 7 days old ECSs of

‘Rasthali’ (AAB) (Ganapathi et al., 2001). The highest competence phase is thought to

coincide with the exponential growth phase in the ECS growth curve (Sági et al., 1995a,

1995b).

Figure 5.3 Representative images on the effects of ECS age (days after subculture) on AmT frequencies in ‘Obino l’Ewai’. The scale bar is equivalent to 1mm.

3 d 5 d 9 d

Figure 5.2 Transient GUS expression (indicated as number of blue foci) in 'Obino l'Ewai' transformed with Agrobacterium (EHA 105, pFAJ3000). Means of at least three replications are presented in two independent trials.

0

500

1000

1500

2000

2500

1 3 5 7 9

ECS age (days)

No.

of b

lue

foci

/ pl

ate

1st trial2nd trial

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5.2.3. Effect of ECS volume

Synthetic gfp gene (sgfpS65T) was optimised for use in monocots and provides a non

destructive assay or physical marker for transformed plant cells (section 3.2.1). This

characteristic allows monitoring of transformed cell or cell clusters from transformation,

through selection to regeneration phases. Thus, the use of sgfpS65T expression enabled

the monitoring of cells and cell clusters during the selection process of reduced ECS

volumes.

Transient GFP Expression (TGFPE) was quantified as green fluorescent spots and was

determined at variable ECS volumes in two independent experiments. The mean numbers

of GFP spots per experiment were plotted against their respective ECS volumes (Figure

5.4). The mean TGFPE increased with ECS volume between 50 and 100 µL with the

highest expression observed at 100 µL of ECS volume. Then with increasing ECS volume,

the TGFPE decreased. The lowest TGFPE was observed at 1200 μL, which is commonly

used in transformation experiments. Higher TGFPE in smaller volumes can be attributed

to increased exposure of ECS to Agrobacteria since efficient spreading of thin layers of

cells is achieved during the co-cultivation period. During the selection phase, cells in the

50 μL ECS treatment turned brown and died. This was not observed in control 50 μL ECS.

The level of browning decreased with increasing ECS volumes. However, more

fluorescent spots could still be observed in reduced ECS volumes after 1 month on

Figure 5.4. Transient GFP expression in 'Three Hand Planty' transformed via Agrobacterium (AGLO, pUbi-sgfpS65T). Mean±SE of two independent trials each with at least 3 replications.

0

500

1000

1500

2000

2500

50 100 200 300 600 1200

ECS volume ( l)

No.

of G

FP sp

ots

1st trial2nd trial


81

selective medium. Whether reduced ECS volumes increase transformation efficiency

needs to be confirmed by quantification of stable transformants.

5.3. Transformation of four banana cultivars with gfp gene

AmT system for banana embryogenic cells involves different steps. These steps consist of

both physical and physiological parameters that influence subsequent transformation of

plant cells. It is important to identify the main parameters that reduce/increase the

transformation frequency of banana ECS. To assess the efficiency of transformation, and

later determine limiting physical parameters, ECS of four banana cultivars GN, THP, OE

and OR were transformed with the synthetic sgfpS65T gene. The use of gfp gene enabled

monitoring variations between transient and stable gene expression.

5.3.1. Transient and stable gfp gene expression

Green fluorescence, though faint just after co-cultivation, was visible in banana

embryogenic cells. Untransformed ECS, used as negative control, were not fluorescent.

Immediately after co-cultivation, numerous fluorescent spots could be viewed in a weakly

fluorescing background. Transient expression was completely distinguishable from the

background after two to three weeks on selective medium. The magnitude of fluorescent

spots decreased with increasing time during selection. Cell clusters or ECS groups, which

initially fluoresced highly, turned faint and finally became non fluorescent. Though this

was observed in green fluorescing ECS of all banana cultivars used, such decline in

fluorescence was cultivar dependent (Figure 5.5). These observations suggest a highly

efficient delivery of T-DNA into the ECS with a low efficiency of stable T-DNA

integration into the genome.

Highly fluorescing colonies were picked under the GFP microscope and transferred onto

RD1 for further selection. One week after selection, GFPE was quantified as the number

of green fluorescent spots per plate. Stable GFP expression was defined as the number of

fluorescing colonies per plate, after 2-3 months on selection medium. Continuous GFP

monitoring was done so as to study expression during the transition from transient to

stable gene expression.

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82

The change from transient to stable GFP expression was quantified as the ratio of the

number of GFP expressing cell colonies (stable gfp gene expression) over the total number

of transiently fluorescing cells or cell clusters counted. Variations in GFP expression at

both transient and stable levels were observed among the four banana cultivars used

(Table 5.2).

The highest TGFPE (996 spots) was observed in ‘Grand Naine’ with the least (335)

observed in cultivar ‘Orishele’ (Table 5.2). GFP expression in THP (557) and OE (546)

did not differ significantly. Though appreciable TGFPE were observed in all the four

banana cultivars, low and variable embryogenesis rate was observed. The highest number

of colonies per plate, was counted in OE (179) and the least (9) in GN. These differences

were probably caused by the variable regeneration capacity of the cell lines used.

Variances among the cultivars’ transient and stable GFP expression were analysed.

Figure 5.5 Green fluorescent protein (GFP)-expressing banana ECS and embryos. A, GFP expression in ‘Grand Naine’ 1 month after transformation; B, GFP expressing embryos in ‘Grand Naine’ 2.5 months after transformation; C, GFP expression in ‘Obino l’Ewai’ 1 month after transformation; D, GFP expressing embryos in ‘Obino l’Ewai’ 2.5 months after transformation. Size bar = 1 mm.

B

D

A

C


83

Table 5.2 Analysis of transient and stable GFP expression in cells and colonies of four banana cultivars after AmT with the synthetic gfp gene

Cultivar Expression of synthetic gfp gene (sgfpS65T) TGFPE SGFPE SGFPE/TGFPE (%) GN 996.3 ± 315 a 8.6 ± 5.40 b 0.86 THP 556.5 ± 108 b 17.6 ± 8.20 b 3.16 OE 545.9 ± 185 b 178.8 ± 23.7 a 32.75 OR 334.8 ± 89 c 12.3 ± 04.8 b 3.67

The data of two independent experiments were used to generate statistically acceptable

deductions. The four banana cultivars differed highly (P≤ 0.000) in transient and stable

GFP expression (P≤ 0.000). This indicates that observed variations in transient expression

and the number of GFP expressing colonies were greatly influenced by intrinsic

differences among cultivars. High transient GFP expressing cells did not (P≤ 0.000)

indicate high stable expression. Nevertheless any strategy that increases the survival of

colonies during selection should be pursued. Though appreciably high numbers of

colonies were obtained in all four banana cultivars, notable (P≤ 0.016) differences were

observed in their shoot proliferation ability (Figure 5.6).

Though cultivars GN and OE had the highest number of GFP expressing colonies, the

regenerated shoots were smaller, especially with GN. Interestingly, THP and OR that had

reasonable numbers of GFP expressing ECS generated higher numbers of shoots.

TGFPE, Transient GFP Expression when measured as expressed by the number of fluorescing cells per plate; SGFPE, Stable GFP Expression when measured as the number of GFP expressing colonies on the same plates. Mean ± SE, mean and standard error of 10 replicates per cultivar. Entries in the same column followed by the same letter are not significantly different from each other at P=0.05. GN, ‘Grand Naine’;THP, ‘Three Hand Planty’; OE, ‘Obino I’Ewai’, OR,’ Orishele’.

OE ORGN THP

0 20

40

60

80

100

120

140

160

Banana cultivars

Col

onie

s/Sh

oots

ColoniesShoots

Figure 5.6 Number of colonies and shoots regenerated in four banana cultivars after 2-3 month of selection. Colonies/Shoots, mean number of colonies per plate and total number of shoots regenerated per cultivar. GN, ‘Grand Naine’; THP, ‘Three Hand Planty’; OE, ‘Obino I’Ewai’, OR,’ Orishele’. Mean±SE of 10 replications.

Chapter five

84

Results generated in this experiment shows variable numbers of GFP expressing colonies

(Figure 5.6). This could mean that the used ECS had variable competencies to

Agrobacterium infection.

5.4. Molecular analysis of gfp gene in banana

5.4.1. PCR analysis

‘Obino l’Ewai’ lines analysed were: Ab.GA.2.2.01, Ab.GA.2.2.09, Ab.GA.2.2.11,

Ab.GA.2.2.12, Ab.GA.2.2.14, Ab.GA.2.2.17, Ab.GA.2.2.35, Ab.GA.2.2.49,

Ab.GA.2.2.53, and line Ab.GA.2.2.67. To compare transformation frequencies among the

cultivars, representatives of transformed ‘Orishele’ lines were analysed as well. These

lines included Ab.GA.2.3.25, Ab.GA.2.3.26, Ab.GA.2.3.34, Ab.GA.2.3.38, Ab.GA.2.3.40,

Ab.GA.2.3.46, Ab.GA.2.3.51 Ab.GA.2.3.57, Ab.GA.2.3.60 and line Ab.GA.2.3.61. All

lines but two of OE and OR gave positive signals for the gfp gene. The lines which were

negative are Ab.GA.2.2.12 and Ab.GA.2.3.38. The presence of the nptII gene needs

confirmation by Southern blot analysis.

A B

719 bp

M 1 2 3 4 5 6 7 8 9 10 +Co -Co

sgfpS65T

Figure 5.7 PCR analyses of regenerated lines from OE (A) and OR (B) containing the gfp gene (sgfpS65T). M, 1-kb ladder; +Co, positive plasmid control (pUbi-sgfpS65T); -Co, negative plant control; lanes 1 to 10 are randomly selected lines analysed and they included Ab.GA.2.2.01, Ab.GA.2.2.53, Ab.GA.2.2.67, Ab.GA.2.2.12, Ab.GA.2.2.11, Ab.GA.2.2.35, Ab.GA.2.2.09, Ab.GA.2.2.14, Ab.GA.2.2.49 and Ab.GA.2.2.17; and Ab.GA.2.3.61, Ab.GA.2.3.26, Ab.GA.2.3.34, Ab.GA.2.3.25, Ab.GA.2.3.51, Ab.GA.2.3.38, Ab.GA.2.3.46, Ab.GA.2.3.57, Ab.GA.2.3.40, and line Ab.GA.2.3.60 for OR.

M 1 2 3 4 5 6 7 8 9 10 -Co+Co

sgfpS65T

M 1 2 3 4 5 6 7 8 9 10 -Co +Co

nptII

M 1 2 3 4 5 6 7 8 9 10 +Co -Co

nptII

554 bp


85

PCR analysis (Figure 5.7) and GFP microscopic assessment (Table 5.3) of the two banana

cultivars showed 80-100% transformation frequencies with respect to the gfp gene and

90% for the nptII gene. These results were consistent with observations in section 4.2.2.1

where over 90% presence of uidA and nptII were detected in the AmT system.

Table 5.3 Microscopic GFP test and PCR analysis of putative Agrobacterium-transformed ‘Grand Naine’,

‘Three Hand Planty’, ‘Obino l’Ewai’ and ‘Orishele’ plantlets

Cultivar Samples GFP test PCR analysis +ve nptII Gfp ‘Grand Naine’ 07 07 07 07 ‘Three Hand Planty’ 10 08 08 08 ‘Obino l’Ewai’ 10 10 10 10 ‘Orishele’ 10 09 09 09

Plantlets were viewed under a GFP microscope before DNA isolation. Total DNA extracted from transformed plantlets was screened by PCR with nptII or gfp gene specific primers. In each reaction PCR reaction mix and genomic DNA sample of an untransformed plantlet were used as negative control. These results confirmed the high plant transformation efficiency reported in several other

crops (Roy et al., 2000; Dai et al., 2001). Based on our analyses and reported literature,

the AmT system was thus used for the rest of our experiments.

5.4.2. Transcription of gfp gene

Total RNA was isolated from selected putative transformed THP lines Ab.GA.2.4.04,

Ab.GA.2.4.13, Ab.GA.2.4.39, Ab.GA.2.4.57, Ab.GA.2.4.80, and Ab.GA.2.4.82. In all

cases tested the amplification product obtained by gfp gene specific primers was identical

to the one of the positive PCR plasmid control (719 bp) indicating that the gfp gene was

properly transcribed in the tested transformants. The RT-PCR reaction performed on the

control, non-transformed lines yielded no product. PCR analysis of DNaseI-treated RNA

samples with actin-specific primers did not result in any signal indicating that all RNA

samples used did not contain DNA. The results of RT-PCR analysis for gfp gene in

selected banana cultivars are shown Figure 5.8.

Chapter five

86

M 1 2 3 4 5 6 –Co W +Co

5.4.3. Integration pattern of gfp transgene into banana genome

Genomic DNA was isolated from transformed banana plantlets grown in the greenhouse.

GFP expressing lines used included Ab.GA.2.2.01, Ab.GA.2.4.04, Ab.GA.2.4.13,

Ab.GA.2.2.17, Ab.GA.2.3.25, Ab.GA.2.1.36, Ab.GA.2.2.45, Ab.GA.2.2.49, and line

Ab.GA.2.4.57. Hybridisation using a 719-bp PCR probe resulted in various gfp integration

loci among different transformed lines analysed (Figure 5.9). Most of the transformants

displayed different integration profiles (Figure 5.9) implying that they were independent

transformed lines from different transformation events (Table 5.4). Integration events

varied among different banana cultivars. A range of one to five integration events was

observed with the highest detected in OR. Numbers of gfp integration loci detected in the

four cultivars are shown in Table 5.4.

719bp

gfp

Figure 5.8 RT-PCR analysis of gfp expression in transgenic lines of ‘Three Hand Planty’ generated by AmT. M, 1-kb ladder; -Co, RNA from untransformed THP; W, water as negative control; +Co, pUbi-sgfpS65T as positive control; lanes 1 to 6, are putatively transformed lines and included Ab.GA.2.4.57, Ab.GA.2.4.39, Ab.GA.2.4.82, Ab.GA.2.4.04, Ab.GA.2.4.80, and Ab.GA.2.4.13.


87

Table 5.4 Transgene gfp integration profiles among selected transgenic lines of ‘Grand Naine’, ‘Three Hand Planty’, ‘Obino l’Ewai’ and ‘Orishele’

Transgenic line Banana cultivar No. of integrated loci -Co (Control) OE 0 0 Ab.GA.2.2.01 OE 1 2 Ab.GA.2.2.17 OE 1 1 Ab.GA.2.2.45 OE 4 3 Ab.GA.2.2.49 OE 4 3 Ab.GA.2.1.36 GN 2 2 Ab.GA.2.3.25 OR 5 5 Ab.GA.2.4.04 THP 3 3 Ab.GA.2.4.13 THP 4 3 Ab.GA.2.4.57 THP 1 1

Integration events in transgenic lines of cultivars OE and THP ranged between 1 and 4.

Two integration sites were observed in line Ab.GA.2.1.36, the only GN transgenic line

analysed. Four transgenic lines of OE, Ab.GA.2.2.01, Ab.GA.2.2.17, Ab.GA.2.2.45, and

Ab.GA.2.2.49 were characterised. Lines Ab.GA.2.2.45 and Ab.GA.2.2.49 showed similar

integration profiles indicating that they were most likely clones from one transformation

event and hence derived from a single transformed embryogenic cell (Figure 5.9)

Transgenic line Ab.GA.2.3.25 of OR had five integration sites which was the highest

among the transgenic lines analysed. Transgenic lines of cultivar THP, Ab.GA.2.4.04,

Ab.GA.2.4.13 and Ab.GA.2.4.57, displayed specific integration patterns with

Ab.GA.2.4.57 showing one integration event.

Ab.G

A.2.2.45

Ab.G

A.2.2.49

Ab.G

A.2.1.36

Ab.G

A.2.2.17

Ab.G

A.2.2.01

Hybridised with sgfpS65T probe

Ab.G

A.2.4.57

+CO

Ab.G

A.2.3.25

Ab.G

A.2.4.1 3

MW

III

-Co

5 Kb

21 Kb

3.5Kb

Figure 5.9 Southern blot analysis of transgenic lines of ‘Grand Naine’, ‘Three Hand Planty’, ‘Obino l’Ewai’ and ‘Orishele’. HindIII digested DNA was hybridised with a PCR probe from the gfp coding sequences. The fourth position in the line identification code indicates the name of the cultivar where 1=‘Grand Naine’, 2=‘Obino l’Ewai’, 3=‘Orishele’ and 4=‘Three Hand Planty’; MW III, DIG labeled molecular weight marker; -Co, untransformed ‘Obino l’Ewai’ plant as negative control; +Co, pUbi-sgfpS65T as positive control.

The fourth position in the line identification code indicates the name of the cultivar where 1=‘Grand Naine’ (GN), 2=‘Obino l’Ewai’ (OE), 3=‘Orishele’ (OR), 4=‘Three Hand Planty’ (THP).

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5.5. The effects of spermidine on banana ECS regenerability

Several strategies have been reported to increase the survival of infected and transformed

plant cells and hence increase the number of regenerable shoots. Treatments include

increased antioxidant concentrations (Cheng et al., 2004), heat shock prior to

Agrobacterium infection (Hansen et al., 2000; Khanna et al., 2004), incorporation of

polyamines such as spermidine (Khanna and Daggard, 2003) and inhibition apoptosis or

programmed cell death (Dickman, 2001). Thus the incorporation of the polyamine

spermidine in selective media could increase the regenerability of transformed banana

cells.

After 1 month culture on spermidine (SPD)-supplemented selective RD1 medium, EC

clones were transferred onto non-selective RD2 medium. EC clones remained for

approximately 2 months in culture, till shoots appeared. Spermidine-containing media had

negative effects on regeneration of ‘Williams’ EC clones at all concentrations tested

(Figure 5.10).

Regeneration of THP, however, significantly increased at low SPD concentrations,

particularly at 0.1 mM (Figure 5.10). The amount of polyamines required during

embryogenesis depends on exogenously and intrinsically supplied polyamine biosynthesis

ability (Shoeb et al., 2001). Spermidine and other polyamines are reported to behave like

antioxidants and improve plant regeneration by acting as plant growth substances (Tang et

al. 2004). Different SPD concentrations have been reported to influence plant cell

embryogenesis and regeneration. These are 0.1 mM in onions (Martinez et al., 2000), 0.5

mM in rice (Shoeb et al., 2001), 100 mM in wheat (Khanna and Daggard, 2003), and 1.5

mM in pine (Tang et al., 2004). On the basis of shoot quantity and quality, SPD at 0.1 mM

(Figure 5.11A) gave the best response. The shoots looked more vigorous than transformed

(Figure 5.12E) and non-transformed (Figure 5.11F) EC clones that were regenerated on

RD1 and RD2 without SPD. Responses observed in Figure 5.11A could be due to the

reported crucial role in somatic embryo development, stimulation of cell division,

regulation of rhizogenesis and embryogenesis (Kakkar and Shawney, 2002).


89

Figure 5.11 Influence of varied SPD concentrations, exogenously added, on the regeneration of putatively transformed ECS clones of ‘Three Hand Planty’. Shoot growth on RD2 supplemented with SPD concentrations: A, at 0.1 mM; B, at 0.5 mM; C, at 1 mM; D, at 5 mM; E, transformed ECS clones at 0 mM; and F, untransformed ECS clones at 0 mM. Similar SPD concentrations were maintained at RD1 and RD2 regeneration steps. RD1 is additionally supplemented with antibiotics geneticin and timentin. The scale bar is equivalent to 1mm.

E

A C

D F

B

0

20

40

60

80

100

120

NT TC NT2 0.1 0.5 1 5 10

SPD in mM

Reg

ener

atio

n fr

eque

ncy

THPW

Figure 5.10 The effects of exogenously added spermidine (SPD) on the regeneration frequency of transformed colonies (shoots/total colonies per cultivars) of ‘Three Hand Planty’ (THP) and ‘Williams’ (W). NT, untransformed colonies at 0 mM SPD; TC, transformed colonies at 0 mM SPD; NT2, untransformed colonies at 0.1 mM SPD determined previously in preliminary experiments (data not shown). Regeneration frequencies are calculated as percentages of shoots obtained from 70 to 96 colonies at a given SPD concentration (in mM).

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90

Further increased SPD concentrations from 0.5 mM to 5 mM (Figures 5.11B-D) however,

suppressed shoot regeneration and at 10 mM SPD only two shoots were observed out of

70 EC clones evaluated. The results obtained show a clear optimum concentration of 0.1

mM SPD for THP transformed EC clones. At that concentration, the regeneration

frequency increased from 70% to 95% of the control. However, optimum concentrations

appear to be genotype-dependent and thus a need to be optimised for each cultivar.

5.6. Conclusions and perspectives

Experimental assessment of banana transformation using the AmT system indicated

potential areas where optimization could further increase transformation frequencies.

Using the modified GFP gene sgfpS65T, factors that influence transient expression, stable

gene expression and subsequent shoot regeneration were analyzed. Molecular analyses

(PCR, RT-PCR and Southern blot analyses) confirmed that the regenerated banana lines

from cultivars GN, OE, OR and THP were transgenic. Results indicated that the observed

variations at both transient and stable gene expression depended on the banana cultivar.

Though the observed transient expression varied with the cultivar, there was no correlation

with the amount of shoots regenerated. All parameters investigated show that

transformation frequencies can be increased by optimising the ECS age, the ECS volume

during co-cultivation, the infection length and the EC regeneration via spermidine

application. An ECS age between 5 and 7 days gave the highest amount of transformation

frequencies and an infection length up to 6 hours seems optimal. Also an ECS volume

between 100 and 300 μL during co-cultivation seems preferable. Finally a spermidine

concentration of 0.1 mM resulted in the highest number of transgenic plants regenerated.

Hence, these parameters deserve to be combined and tested in an improved AmT system.

Other parameters like antioxidants, temperature, light, and auxin concentrations need to be

tested as well.

Transformation with rice chitinase genes

91

Chapter 6. Transformation with rice chitinase genes

6.1. Introduction

Chitinases are known to exert antifungal activity (Broglie et al., 1991; Neuhaus, 1999) and

were the first genes reported to enhance resistance against a range of fungal diseases

(Legrand et al., 1987; Broglie et al., 1991). Rice chitinase genes have then been

extensively studied (Zhu and Lamb, 1991; Nishizawa et al., 1991, 1993, 1999; Tabei et

al., 1998; Takatsu et al., 1999; Yamamoto et al., 2000; Datta et al., 2001; Kishimoto et al.,

2002; Takahashi et al., 2005) and applied to confer protection against fungal diseases

(Asao et al., 1997; Nishizawa et al., 1999; Kishimoto et al., 2002). This chapter presents

results on the transformation of two banana cultivars with rice chitinase genes (Cht-2 and

Cht-3), regeneration of transformed embryogenic colonies, PCR and Southern blot

analysis of putative transgenic lines.

6.2. Plant material and binary vectors

ECSs of the banana cultivars ‘Gros Michel’ (GM) and ‘Grand Naine’ (GN) were infected

and cocultivated with the Agrobacterium strain EHA105 as described (section 3.3). Strain

EHA105 contained the binary vectors pBI333-EN4-RCC2 or pBI333-EN4-RCG3 (section

3.2.1). To enable selection of transformed banana cells, both binary vectors contained the

hpt gene that codes for the hygromycin phosphotransferase enzyme. Banana cells

expressing hpt are able to survive on medium supplemented with the selective antibiotic

hygromycin.

6.3. Induction of transformed embryogenic colonies and plant regeneration

After 2-3 months of incubation in the dark on ZZ medium supplemented with 50 mgL-1

hygromycin and 200 mgL-1 timentin, surviving and proliferating masses of cells (colonies)

were picked and individually transferred onto 1 mL of selective semi-solid RD1 medium

in 24-well plates (Table 6.1). Embryogenic colonies were transferred onto non-selective

RD2 medium and incubated at 26°C till shoots appeared. For each transgenic event, three

plants were maintained on REG medium.

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92

Table 6.1 Number of colonies transferred onto selective RD1 medium after Agrobacterium-mediated transformation of two rice chitinase genes

Chitinase genes Cultivar Cht-2 Cht-3 ‘Grand Naine’ 87 38 ‘Gros Michel’ 120 120

The embryogenic capacity was different in the two banana cultivars and the vector

constructs used: GM had more embryogenic colonies than GN, however, their

regenerability was lower than that of GN (Table 6.2).

Table 6.2 Number and regeneration frequency (%) of independent transgenic lines in two banana cultivars transformed by Agrobacterium

Chitinase genes Cultivar Cht-2 Cht-3 ‘Grand Naine’ 45 (51.7) 13 (34.2) ‘Gros Michel’ 26 (21.7) 39 (32.5)

Regeneration frequency (percentage, in brackets) is the number of regenerated plants divided by the number of embryogenic colonies (Table 6.1), multiplied by 100

The results generally indicate that the cultivar is not a bottleneck for the introduction of

either hpt or rice chitinase genes into banana. However, numbers of putative transformants

clearly depended on the cultivar. This could be due to variations in transformation

competence and/or regenerability of the specific ECS lines used.

6.4. Molecular analysis of chitinase transformants 6.4.1. PCR analysis

Transformation of the two cultivars was first indicated by the survival of putative

transgenic plants on selective media for a period of 3 to 4 months. While PCR analysis

does not confirm stable transgene integration, it is an initial indicator of the presence of

these transgenes in the host plant genome. Total DNA was therefore extracted (section

3.7.1.1) from transgenic plants and untransformed controls, and PCR analysis was

performed as detailed (section 3.7.1). A fragment of hpt, Cht-2 or Cht-3 genes was

amplified by specific oligonucleotide primer pairs (Table 3.2). Binary vector constructs

pBI333-EN4-RCC2 and pBC333-EN4-RCG3 (Figure 3.1) were used as positive controls.

As the hpt selective marker gene and Cht-2 or Cht-3 rice chitinase genes are linked on the

same T-DNA, the expectation is of 100% co-transformation frequency. To test this, PCR

analysis of hpt in combination with each of the chitinase genes was performed on 9 to 10


93

independent transformants of each banana cultivar and construct. For GM putatively

transformed lines, GM.rcc2.02, GM.rcc2.03, GM.rcc2.05, GM.rcc2.06, GM.rcc2.10,

GM.rcc2.11, GM.rcc2.14, GM.rcc2.15, GM.rcc2.23 and GM.rcc2.24.were analyzed for

the hpt and Cht-2 genes (Figure 6.1A). The presence of hpt and Cht-3 genes was analyzed

among another 10 randomly selected lines, which included GM.rcg3.43, GM.rcg3.08,

GM.rcg3.09, GM.rcg3.10, GM.rcg3.11, GM.rcg3.13, GM.rcg3.20, GM.rcg3.21,

GM.rcg3.22, and GM.rcg3.35 (Figure 6.1B). The PCR analysis results in ‘Gros Michel’

(Figure 6.1) demonstrate 100% co-transformation frequency between Cht-3 and hpt, and

100% co-transformation between Cht-2 and hpt. These data also indicate that 90-100% of

the regenerated plants are likely true transgenic.

The results of PCR analysis did slightly differ in the regenerated transgenic lines of GN.

Analysis of Cht-2 in lines GN.rcc2.01, GN.rcc2.05, GN.rcc2.24, GN.rcc2.26, GN.rcc2.33,

GN.rcc2.35, GN.rcc2.36, GN.rcc2.40, and GN.rcc2.41, and GN.rcc2.83 gave a (co-)

transformation frequency of 90% (Figure 6.2A).

Figure 6.1 PCR analysis of representative transgenic plants from ‘Gros Michel’ containing rice chitinase genes Cht-2 (A) or Cht-3 (B) together with hpt. M, 1-kb DNA ladder; +Co, positive controls (plasmids pBI333-EN4-RCG3 or pBI333-EN4-RCC2); -Co, negative plant control; lanes 1 to 10, independent transgenic lines GM.rcc2.15, GM.rcc2.14, GM.rcc2.02, GM.rcc2.06, GM.rcc2.24, GM.rcc2.11, GM.rcc2.03, GM.rcc2.05, GM.rcc2.23, and GM.rcc2.10 for Cht-2; GM.rcg3.11, GM.rcg3.35, GM.rcg3.21, GM.rcg3.10, GM.rcg3.13, GM.rcg3.06, GM.rcg3.09, GM.rcg3.22, GM.rcg3.08, and GM.rcg3.20 for Cht-3.

hpt

M 1 2 3 4 5 6 7 8 9 10 +Co Co

B Cht-3

M 1 2 3 4 5 6 7 8 9 10 +Co-Co

668bp

hpt

-Co+Co 1 2 3 4 5 6 7 8 9 10 M

A Cht-2

M 1 2 3 4 5 6 7 8 9 10 +Co -Co

405 bp

414bp

Chapter six

94

Similarly, lines GN.rcg3.03, GN.rcg3.04, GN.rcg3.05, GN.rcg3.06, GN.rcg3.10,

GN.rcg3.11, GN.rcg3.13, GN.rcg3.15, and GN.rcg3.25 showed 90% positive signals for

the presence of transgene Cht-3 (Figure 6.2B). The only negative line (GN.rcg3.06) was

also negative by Southern analysis (Figure 6.7).

The consistence in the high co-occurrence of hpt gene and Cht-2 or Cht-3 indicates that

both transgenes were transferred into the banana genome.

Even though higher plants themselves do not contain chitin (Graham and Sticklen, 1994),

chitinases are present in many plant species, including banana. Indeed, a class III chitinase

homolog was previously reported in young banana fruits, which is presumed to function as

a vegetative storage protein in developing fruits (Peumans et al., 2002). Yet, none of the

untransformed plant controls ever gave amplification products with the Cht-2 or Cht-3

gene-specific primers. Similarly high transformation frequencies were also observed in the

previous experiments indicating the consistence and efficiency of the established

Agrobacterium-mediated transformation system.

6.4.2. Southern blot analysis of Cht-2 and Cht-3 genes

The survival of transformed ECS on selective media (section 6.2) and subsequent PCR

analysis of regenerated putatively transformed plants (section 6.4.1) were indicative of a

successful transfer of transgenes Cht-2 and Cht-3 into banana. However, transgenic plants

need more characterisation with regard to transgene integration patterns and transgene

Figure 6.2 PCR analysis of representative transgenic plants from ‘Grand Naine’ containing rice chitinase genes Cht-2 (A) or Cht-3 (B) together with hpt. M, 1-kb DNA ladder; -Co, negative plant control; +Co, positive controls (plasmids pBI333-EN4-RCC2 or pBI333-EN4-RCG3); lanes 1 to 10, independent transgenic lines GN.rcc2.40, GN.rcc2.36, GN.rcc2.01, GN.rcc2.35, GN.rcc2.26, GN.rcc2.83, GN.rcc2.05, GN.rcc2.33, GN.rcc2.41, and GN.rcc2.24 for Cht-2; GN.rcg3.15, GN.rcg3.03, GN.rcg3.04, GN.rcg3.05, GN.rcg3.11, GN.rcg3.25, GN.rcg3.13, GN.rcg3.06, and GN.rcg3.10 for Cht-3.

hpt

405 bp

hpt M 1 2 3 4 5 6 7 8 9 10 -Co +Co

668 bp

M 1 2 3 4 5 6 7 8 9 +Co-Co

M 1 2 3 4 5 6 7 8 9 10 -Co +Co A Cht-2

414 bp

M 1 2 3 4 5 6 7 8 9 +Co-Co

B Cht-3


95

21kb

5 kb

2 kb

MII

I

GN

.co.

1

G

M.rc

c2.2

4

G

M.rc

c2.0

2

GM

.rcc2

.05

G

M.rc

c2.1

4

GN

.rcc2

.33

G

N.rc

c2.3

5

GN

.rcc2

.40

G

N.rc

c2.4

3

GN

.rcc2

.06

+Co

Figure 6.3 Southern blot analysis of transgenic lines of ‘Gros Michel’ (GM) and ‘Grand Naine’ (GN). HindIII digested total DNA was hybridised with a PCR probe from the Cht-2 coding sequence. M, molecular weight marker III (Roche); GN.co.1, untransformed GN control; +Co, pBI333-EN4-RCC2 as positive control.

copy numbers. Thus, transgenic lines of GN and GM, harboring the Cht-2 or Cht-3 rice

chitinase genes, were analyzed by Southern blot hybridisation.

6.4.2.1. DNA isolation and restriction digestion

The lines GM.rcc2.24, GM.rcc2.02, GM.rcc2.05, GM.rcc2.14 of ‘Gros Michel’, and lines

GN.rcc2.33, GN.rcc2.35, GN.rcc2.40, GN.rcc2.43, and GN.rcc2.06 of ‘Grand Naine’ were

used for the analysis of Cht-2 (Figure 6.3). For Cht-3, GM.rcg3.39, GM.rcg3.30,

GM.rcg3.20, GN.rcg3.02, GN.rcg3.03, GN.rcg3.04, GN.rcg3.05, and GN.rcg3.06 were

used (Figure 6.7). Ten micrograms of each sample were digested with HindIII, which cuts

once through the T-DNA of binary vector pBI333-EN4-RCC2 or pBI333-EN4-RCG3

(Figure 3.1).

Southern blot analysis with the Cht-2 (Figure 6.3) and Cht-3 (data not shown) probes gave

similar hybridisation patterns in all transformed lines as well as in the untransformed

controls suggesting the existence of highly complementary chitinase sequences in banana.

To confirm the existence of homologous endogenous banana chitinase sequences,

BLASTn searches for banana chitinase sequences and sequence alignments were done.

Chapter six

96

6.4.2.2. Nucleotide sequence analyses of chitinase from banana and rice

The rice chitinase genes Cht-2 and Cht-3 were isolated from a cDNA library and a

genomic DNA library (Nishizawa et al., 1993), respectively, and they are 78% identical.

When their amino acid sequences were compared to class I chitinases in tobacco (Shinshi

Cht-2 (X56787) AACACCGAGACGCGGAAGCGGGAGGTCGCCGCGTTCCTGG 456 Cht-3 (D16223) gACgaCGccACGaaGAAGaGGGAGaTCGCgGCtTTCtTGG 144 AJ277278 AACACCGAGACGCGGAAGCGGGAGGTCGCCGCGTTCCTGG 114 AJ277278 gACgaCnccACGaaGAAGaGGGAGaTCGCgGCtTTCtTGG 393 AF416677 gACgCCGAcACctGcAAGCGcGAGGTCGCCGCcTTCCTGG 2240 Z99966 gACgCCGAcACctGcAAGCGcGAGGTCGCCGCcTTCCTGG 120 Cht-2 (X56787) GCCAGACCTCCCACGAG.ACCACCGGCGGGTGGCCGACCG 495 Cht-3 (D16223) cgCAGACgTCtCACGAG.ACgACaGGtGGGTGGgCGACgG 183 AJ277278 GCCAGACCTCCCACGAG.ACCACCGGCGGGTGGCCGACCG 153 AJ277279 cgCAnACgTCtCACnAanACnACaGGtGGGTGGgCGACgG 433 AF416677 cgCAGACgTCCCACGAG.ACCACCGGCGGcTGGCCcACgG 2279 Z99966 cgCAGACgTCCCACGAG.ACCACCGGCGGcTGGCCcACgG 159 Cht-2 (X56787) GGAGCAGAAC..CCGCCGTCCGAC.TACTGCCAGCCCTC 571 Cht-3 (D16223) GGAaCAGAAC..CCcCCaTCgGAC.TACTGCgtcgCCag 259 AJ277278 GGAGCAGAAC..CCGCCGTCCGAC.TACTGCCAGCCCTC 229 AJ277279 aaAaCAGAAC..CCcCCaTCgGAC.TACTGCgtcnCCag 510 AF416677 GGAGaAcAACggCaaCgccCCcACaTACTGCgAGCCCaa 2358 Z99966 GGAGaAcAACggCaaCgccCCcACaTACTGCgAGCCCaa 238 Cht-2 (X56787) GCCGGAGTGGCCGTGCGCCCCCGGCCGCAAGTACTACGGC 611 Cht-3 (D16223) ctCGcAGTGGCCGTGCGCtgCaGGCaagAAGTACTACGGC 299 AJ277278 GCCGGAGTGGCCGTGCGCCCCCGGCCGCAAGTACTACGGC 269 AJ277279 ctCGcAnTGGCCGTGCGCtgCaGGCaanAAGTACTACGGC 550 AF416677 GCCGGAGTGGCCGTGCGCCgCCGGCaagAAGTACTACGGC 2398 Z99966 GCCGGAGTGGCCGTGCGCCgCCGGCaagAAGTACTACGGC 278 Cht-2 (X56787) CGCGGCCCCATCCAACTCTCCTTCAACTTCAACTACGGGC 651 Cht-3 (D16223) CGaGGCCCCATCCAAaTCTCaTTCAACTaCAACTACGGGC 339 AJ277278 CGCGGCCCCATCCAACTCTCCTTCAACTTCAACTACGGGC 309 AJ277279 CGaaGCCCCATCCAAaTCTCaTTCAACTaCAACTACnGGg 590 AF416677 CGgGGaCCCATCCAgaTCaCCTaCAACTaCAACTACGGcC 2438 Z99966 CGgGGaCCCATCCAgaTCaCCTaCAACTaCAACTACGGcC 318 Cht-2 (X56787) CGGCGGGGAGGGCGATCGGGGTGGACCTGCTGAGCAACCC 691 Cht-3 (D16223) CGGCcGGGAGaGCcATCGGctccGACCTGCTcAaCAACCC 379 AJ277278 CGGCGGGGAGGGCGATCGGGGTGGACCTGCTGAGCAACCC 349 AJ277279 CcGgccGGgaaaCcATCGGctccGAC.TGCTcAaCAACCC 629 AF416677 CGGCGGGGcaGGCcATCGGctccGACCTGCTcAaCAACCC 2478 Z99966.1 CGGCGGGGcaGGCcATCGGctccGACCTGCTcAaCAACCC 358 Cht-2(X56787) GGACCTGGTGGCGACGGACGCGACGGTGTCGTTCAAGAC 730 Cht-3(D16223) aGACCTGGTGGCcACcGACGCGACcaTcTCGTTCAAGAC 418 AJ277278 GGACCTGGTGGCGACGGACGCGACGGTGTCGTTCAAGAC 388 AJ277279 caaACCTGGTGGCcACcGACGCcAacaTcTCnTTCAAaA 669 AF416677 GGACCTGGTGGCGtCGGACGCcACcGTcTCcTTCAAGAC 2517 Z99966.1 GGACCTGGTGGCGtCGGACGCcACcGTcTCcTTCAAGAC 397

Figure 6.4 Nucleotide sequence comparisons (Multiple alignments, DNAMAN Version 4.13, Biosoft, Quebec, Canada) of rice chitinase genes Cht-2 and Cht-3 with banana chitinase sequences identified in GenBank. Gaps are introduced by the software for optimum alignment. Homologous nucleotides in all the six chitinase sequences are indicated in blue colour and Cht-2 (reference sequence) is indicated as dark blue.


97

et al., 1990), potato (Gaynor, 1988), and bean (Broglie et al., 1986; Zhu and Lamb, 1991),

a similarity of over 62% (Cht-2/potato chitinase) was observed (Nishizawa et al., 1993).

BLASTn searches (www.ncbi.nlm.nih.gov/Blast) were therefore made in the GenBank

database for homologous sequences to Cht-2 only, which resulted in three high hits

(Figure 6.4). These were the nucleotide accessions AJ277278 (Musa acuminata mRNA for

putative chitinase isoform 1), AJ277279 (M. acuminata mRNA for putative chitinase

isoform 2), and AF416677 (partial sequence of M. acuminata endochitinase mRNA).

BLASTn searching with banana chitinase gene (AJ277278), identified another banana

chitinase (GenBank accession number Z99966) (Figure 6.4), which was 91% identical to

accession AJ277278 and 79% to Cht-3 (E = 3e-142). Alignment of these sequences with

the Cht-2 gene by BLAST2 and ClustalW (www.clustalw.genome.jp) showed 77%

identity for AJ277278 (E = 6e-74), 78% identity for AJ277279 as well as for AF416677

(both at E = 3e-79). Comparing sequence AJ277278 with the three identified banana

chitinase sequences gave sequence similarities of 97%, 98%, and 91% for AJ277279,

AF416677 and Z99966, respectively. The nucleotide sequence comparisons of rice Cht-2

and Cht-3 probes and the banana chitinase sequences are given in Table 6.3. Table 6.3 Nucleotide sequence homology between rice chitinase gene probes (Cht-2 and Cht-3) and banana chitinase sequences

Regions of banana chitinase sequences homologous to the probe sequences are shown in brackets.

In all cases, sequence alignments resulted in over 70% sequence similarities (Table 6.3).

This explained why hybridisation of these probes (Cht-2 and Cht-3) gave background

signals in untransformed controls (Figure 6.3). However, since the homology between the

probes and the above banana sequences was much less than 100%, modifications of

hybridisation stringency could remove imperfectly matched probe-target hybrid sequences

and this would improve the quality of Southern blots. The specificity of the probe is

affected by the conditions of post-hybridisation washes. The critical parameters are the

ionic strength of the final wash solution and the temperature at which this wash is done.

Sequence source

Banana (Musa acuminata)

AJ277278 AJ277279 AF416677 Z99966

Probe Cht-2

77.36%

(304 -705)

78.11%

(320-721)

78.10%

(37-438)

70.68%

(288-688)

Ric

e (O

ryaz

a sa

tiva)

Probe Cht-3

78.54%

(294-703)

78.78%

(310-719)

78.77%

(29-438)

71.81%

(278-686)

Chapter six

98

Highly stringent wash conditions destabilise all mismatched heteroduplexes, so that

hybridisation signals are obtained only from sequences that are perfectly homologous to

the probe.

To reduce the background signals, both hybridisation and post-hybridisation washes were

performed at increased stringency. Hybridisation was performed at 70°C and the two

washing steps consisted of 1xSSC at 25°C, followed by 0.05xSSC at 70°C. These

hybridisation and washing conditions resulted in reduced background signals in the

untransformed control (cf. Figure 6.3 and 6.6) and the integration patterns of Cht-2 and

Cht-3 were thus revealeded. Though the use of more stringent washes enabled us

obtaining differential integration patterns for transgenic lines, it should be noted that weak

signals could also have been washed off the blot.

6.4.2.3. Comparisons of amino acid sequences of rice and banana chitinases

Although increased stringency washes could reduce the background, a few questions

remained unanswered. For example, do these sequences encode any chitinase protein? If

yes, how similar are they to rice chitinases? These questions lead to amino acid sequence

searches and alignments.

With five chitinases (Cht-2, Cht-3, AJ277278, AJ277279 and AF416677) the following

respective proteins were identified: CAA40107, BAA03751, CAC81811, CAC81812 and

AAL05705988. These proteins are 340, 320, 318, 317 and 229 amino acids long,

respectively, and also include the N-terminal signal peptides. The signal peptide sequence

in Cht-2 is 32 amino acids compared to Cht-3 with only 18 amino acids. The amino acid

sequence similarities among these chitinases were analysed and revealed high similarities

(70% and above) with the three banana chitinases (Table 6.4).

Table 6.4 Amino acid homologies of two rice chitinases with three banana proteins

Plant species Banana (Musa acuminata)

CAC81811 CAC81812 AAL05705988

Cht-2 70.22 70.12 76.25

Ric

e (O

ryza

sa

tiva)

Cht-3 78.18 72.81 78.06

Cht-2, CAA40107; Cht-3, BAA03751. Analysis was done with DNAMAN (Version 4.13, Lynnon Biosoft, Quebec, Canada).


99

1 PNCLCCSRWGWCGTTSDFCGDGCQSQCSG-CGPTPTPTPPSPSDGVGSIVPRDLFERLLL| CAA40107 1 --------------**DY**A****Q***G**GG*P*SSGGGSGVASI*S*SLFDQM***| BAA03751 1 -------------N*DPY**Q****Q*G*S**SG--------GGSVA**ISSS***QM*K| CAC81811 1 ---------GWCGN*DPY**KD***Q*G*S**SG-----GGSGGSV***ISSS***QM*K| CAC81812 1 ------------------------------------------------------------| AAL05705988 60 HRNDGACPARGFYTYEAFLAAAAAFPAFGGTGNTETRKREVAAFLGQTSHETTGGWPTAP| CAA40107 47 ****Q**A*K*****D**V***N*Y*D*AT**DAD*C********A**************| BAA03751 40 ****A***GK*****N**I***N**SG**T**DDAKK***I****A**********A***| CAC81811 47 ****A***GK*****N**I***N**SG**T**DDA*K***I****A**********A***| CAC81812 1 ------------------------------------------------------------| AAL5705988 120 DGPFSWGYCFKQEQN-PPSDYCQPSPEWPCAPGRKYYGRGPIQLSFNFNYGPAGRAIGVD| CAA40107 107 ***Y*******E*N*GNAPT**E*K******A*K**********TY*Y******Q***S*| BAA03751 100 ***Y******V****-*S****VA**Q****A*K*************Y******R***S*| CAC81811 107 ***Y******V****-******VA**Q****A*K*************Y******R***S*| CAC81812 1 -----------------------------------------******Y******R***S*| AAL05705988 179 LLSNPDLVATDATVSFKTALWFWMTPQGNKPSSHDVITGRWAPSPADAAAGRAPGYGVIT| CAA40107 167 **N**O***S*********F*******SP***C*A****Q*T**AD*Q****V****EI*| BAA03751 159 **N**********I*************SP***C*D****S*T**N**Q****L*****T*| CAC81811 166 **N**********I*************SP***C*D******T**N**R****L*****T*| CAC81812 20 **N**********I*************SP***C*D******T**N**R****L*****T*| AAL05705988 239 NIVNGGLECGHGPDDRVANRIGFYQRYCGAFGIGTGGNLDCYNQRPFNSGSSVGLAEQ | CAA40107 227 **I***V*****A**K**D*****K***DML*VSY*D*********YPPS-------- | BAA03751 219 **I*******K*Y*A***D*****K***DLL*VSY*D*********FASTAAT--*TF | CAC81811 226 **I*******K*S*A***D*****K***DLL*VSY*D*******S*FT---------- | CAC81812 80 **I*******K*S*A***D*****K***DLL*VSY*D*******S*FT---------- | AAL05705988

Figure 6.6 Comparisons of amino acid sequences of rice and banana chitinases, as generated by DNAMAM (Version 4.13, Lynnon Biosoft, Quebec, Canada). Amino acid residues identical to rice chitinase (Cht-2, GenBank accession number CAA40107) are indicated by asterisks. Gaps are introduced by the software for optimum alignment Rice chitinase (Cht-2, CAA40107) amino acid sequence is used is used here as a reference sequence.

Based on detailed protein characterisation of CAA40107 (Cht-2) (Nishizawa et al., 1993),

specific regions and sites can be identified in the banana proteins (Figure 6.5). Region 1

(1-18, blue letters), region 2 (33-62, green colour), and region 3 (93-223, grey colour) are

the signal peptide, chitin binding domain, and glycosidase hydrolase family 19 chitinase

domain, respectively. Sites B (154), C (176), D (205), E (207), F (210-211) and G (286)

are putative sugar binding sites. Chitinase catalytic residues are indicated by A (151), C

(176) and E (207) sites. Region 3 has more homology in banana chitinases than regions 1

and 2. Sites B, C, D, E, F and G also have homologous sites in banana chitinase amino

acid sequences (Figure 6.5).

G

D E F

C A B

2

3

1

Chapter six

100

6.4.2.4. Improved Southern blot analysis of rice chitinases genes

Southern blot analysis for transgenes Cht-2 and Cht-3 in transformed lines of GM and GN

showed different integration profiles. Negative signals in untransformed controls confirm

that the observed signals truly indicate the integration of these transgenes. This is further

corroborated by positive signals from the vector constructs (pBI333-EN4-RCC2 or

pBC333-EN-RCG3), implying that the probes correctly detected integration events

(Figure 6.6 and 6.7). Transgenic lines containing either Cht-2 or Cht-3 showed one to four

integration events, typical of Agrobacterium-mediated transformation events. For both

transgenes, putatively transformed lines of GM showed one to two integration sites

compared to GN where integration sites ranged from one to four (Figures 6.6 and 6.7).

Though these results suggest cultivar dependent transgene integration, it remains to be

confirmed in other banana cultivars.

Integration profiles of Cht-2 in GM lines (Figure 6.6) of GM.rcc2.05, GM.rcc2.14, and

GM.rcc2.24 showed single insertions each, with two integrations in line GM.rcc2.02. In

contrast, GN lines had higher numbers of integration events. Transformed lines

Figure 6.6 Improved Southern blot analysis of transgenic lines of ‘Gros Michel’ (GM) and ‘Grand Naine’ (GN). HindIII digested total DNA was hybridised with a PCR probe from the Cht-2 gene coding sequence. Doubling the stringency of the second wash improved the probe-hybridisation specificity (compare with Figure 6.4). Thus only highly complementarily bound probe fragments remained, specifically indicating the presence of integrated rice chitinase gene. Arrows indicate the approximate fragment sizes, based on DNA molecular weight marker III (Roche).GN.co.1, untransformed GN control; +Co, pBI333-EN4-RCC2 as positive control.

21 kb

5 kb

2 kb

GN

.co.

1

G

M.rc

c2.2

4

G

M.rc

c2.0

2

GM

.rcc2

.05

GM

.rcc2

.14

GN

.rcc2

.33

G

N.rc

c2.3

5

GN

.rcc2

.40

G

N.rc

c2.4

3

GN

.rcc2

.06

+Co


101

GN.rcc2.06 and GN.rcc2.35 contained three integration loci each, with the highest number

of four integration sites observed in line GN.rcc2.40. The least number of integration

events in GN was in GN.rcc2.43 with two insertions.

Integration profiles of Cht-3 in GM and GN (Figure 6.7) again showed slightly more

integration events in GN. Two GM lines GM.rcg3.30 and GM.rcg3.39 had single

insertions, with two events observed in GM.rcg3.20. In GN, the number of integration loci

ranged from one to three. A single transgene locus was detected in GN.rcg3.03 whereas

lines GN.rcg3.02, GN.rcg3.04 and GN.rcg3.05 had two integration events each.

GN.rcg3.06 was negative (see also Figure 6.2B), which could be an escape.

These transgene integration profiles described here for Cht-2 and Cht-3 containing lines of

GM and GN did not differ from what had been reported in a different cultivar ’Rasthali’

(AAB, Ganapathi et al., 2001).

6.5 Conclusion

PCR analysis of selected transgenic lines from both cultivars showed the presence of both

chitinase genes. Southern blot analyses confirmed that the lines regenerated were actually

putatively transformed. In most cases, PCR analyses were consistent with transgene

integration analyses. Line GN.rcg3.06 that had a negative PCR signal was also negative

for Southern blot analysis. For both genes, transgenic GM lines showed one to two

integration sites, whereas in ‘Grand Naine’ the number of integration sites ranged from

21 kb

5 kb

2 kb

GN

.co.

1

G

M.rc

g3.3

9

G

M.rc

g3.3

0

GM

.rcg3

.20

G

N.rc

g3.0

2

GN

.rcg3

.03

G

N.rc

g3.0

4

GN

.rcg3

.05

G

N.rc

g3.0

6

+Co

Figure 6.7 Improved Southern blot analysis of transgenic lines of ‘Gros Michel’ (GM) and ‘Grand Naine’ (GN). HindIII digested total DNA was hybridised with a PCR probe from the Cht-3 gene codingsequence. Hybridisation conditions described in Fig.6.5 were used. Arrows indicate the approximate fragment sizes, based on DNA molecular weight marker III (Roche). GN.co.1, untransformed GN control; +Co, pBI333-EN4-RCG3 as positive control.

Chapter six

102

one to four. Southern blot analyses showed the presence of homologous chitinase

sequences in banana. BLASTn searches generated four different chitinases sequences from

Musa acuminata (dessert banana). Nucleotide and amino acid sequence comparisons of

these banana and rice chitinases showed that they were highly homologous with

similarities of over 70% in each case.

In summary, more than 50 rice chitinase-containing transgenic lines were generated in

each of the banana cultivars GM and GN. Their molecular characterisation confirmed that

the rice chitinase genes Cht-2 and Cht-3 were stably integrated in the banana genome

(Table 6.5). Table 6.5 PCR analysis and integration patterns of selected transgenic lines of ‘Gros Michel’ and ‘Grand Naine’ containing rice chitinase genes (Cht-2 or Cht-3) and hygromycin phosphotransferase (hpt) gene

Line/Construct PCR analysis

Southern blot analysis

hpt Cht-2 Cht-3 No. of Integration loci

GM.rcc2.05 + + 1 GM.rcc2.14 + + 1 GM.rcg3.20 + + 2 GM.rcg3.30 + + 1 GN.rcc2.35 + + 3 GN.rcc2.40 + + 4 GN.rcg3.02 + + 2 GN.rcg3.05 + + 3 GN.rcg3.06 - - -

GM and GN refer to ‘Gros Michel’ and ‘Grand Naine’, respectively. + and - denote absence and presence of PCR (or Southern blot signal), respectively.

Co-transformation of banana with chitinase genes and a plant defensin

103

Chapter 7. Co-transformation of banana with chitinase genes

and a plant defensin

7.1. Introduction

Plant co-transformation refers to the simultaneous introduction of multiple transgenes into

a plant cell followed by co-integration of all these transgenes into the host genome. In case

of gene transfer by Agrobacterium, transgenes can be either linked on the same T-DNA or

located on separate individual T-DNAs within one or more binary vectors that are

harbored in one or more Agrobacterium strains.

Co-transformation provides the technical basis for the manipulation of complex traits, or

when several steps of a biosynthetic pathway have to be modified; for example flower

colour (Tanaka et al., 1998; Rasati et al., 2003) and Golden rice (Ye et al., 2000; Bayer et

al., 2002; Datta et al., 2003). Also, co-transformation is an important tool for ‘gene

pyramiding’, which means the combination of different traits in one plant (e.g. herbicide

and insect resistance in parallel) or expressing several genes to confer the same phenotype

more efficiently (e.g. for durable resistance to a pathogen). The potential of co-

transformation is even more pronounced for banana because cross-fertilisation is excluded

and thus multiple (trans)genes cannot be combined otherwise.

In this study, separate binary vectors, which carried different classes of antifungal genes –

namely the PR-3 rice chitinases (Cht-2 or Cht-3) and a defensin gene from radish (Rs-

afp2, see section 2.4.2) were used. These genes linked to either of two selectable marker

genes were consecutively introduced into a single Agrobacterium strain and then

transferred to two plantain-type banana cultivars. Co-transformation frequencies were

compared between combined selection and when single selective agents were employed.

7.2. Co-transformation of banana

Binary vectors (Figure 3.1) pBI333-EN4-RCC2 or pBI333-EN4-RCG3 (containing Cht-2

and Cht-3, respectively, together with the hpt gene) and pFAJ3494 (carrying Rs-afp2 and

nptII) were introduced into the Agrobacterium strain EHA105 by retransformation. Strains

containing either pBI333-EN4-RCC2 or pBI333-EN4-RCG3 were used to make

competent cells, which were then transformed with pFAJ3494 by electroporation (see

section 3.2.5). Binary vector pFAJ3494 contained aminoglycoside 3-adenylyltransferase

(aad) that confers resistance against streptomycin/spectinomycin (Hollingshead and

Chapter seven

104

Vapnek, 1985). Binary vectors pBI333-EN4-RCC2 and pBI333-EN4-RCG3 had nptII that

provides bacterial cells with resistance against kanamycin. Resulting bacterial

transformants contained either pBI333-EN4-RCC2/pFAJ3494 or pBI333-EN4-

RCG3/pFAJ3494 plasmid combination and were selected on medium supplemented with

kanamycin (50 mgL-1), spectinomycin (100 mgL-1) and streptomycin (300 mgL-1).

Restriction digestion of plasmid preparations purified from re-electroporated

Agrobacterium cells with HindIII gave the four expected bands. These bands included

single, linearised plasmids of either pBI333-EN4-RCC2 (10.5 kb) or pBI333-EN4-RCG3

(10.5kb) and three fragments (2.649, 3.3399 and 6.736kb) from pFAJ3494 (data not

shown). Agrobacterium strains with confirmed plasmid combinations were then used for

banana transformation.

ECS of the plantains THP and OR were transformed (see section 3.3) with pBI333-EN4-

RCC2, pBI333-EN4-RCG3 or pFAJ3494 alone or with the combinations described above.

Table 7.1 Selection schemes applied in single transformation and co-transformation experiments

Genes/gene combinations Cultivar Control Cht-2 Cht-3 Rs-afp2 Cht-2/Rs-afp2 Cht-3/Rs-afp2 THP NA H H G H, G, H+G H, G, H+G OR NA H H G H, G, H+G H, G, H+G

THP, ‘Three Hand Planty’; OR, ‘Orishele’; H, hygromycin; G, geneticin; NA, not applicable

Transformed cells were cultured on selective ZZ medium containing either a combination

of geneticin and hygromycin or separately (Table 7.1). After three months of selection,

150-200 embryogenic colonies were usually transferred for each construct or combination

onto RD1 medium with the same selection regime (Table 7.2).

Table 7.2 Number of colonies transferred onto selective RD1 medium after Agrobacterium-mediated transformation with single genes/gene combinations and selected with different schemes

Gene/gene combinations Cht-2 Cht-3 Rs-afp2 Cht-2/Rs-afp2 Cht-3/Rs-afp2 Cultivar Control H H G H G H+G H G H+G THP 100 288 192 240 240 264 168 240 288 166 OR 100 144 148 264 336 144 00 168 216 00

THP, ‘Three Hand Planty’; OR, ‘Orishele’; H, hygromycin; G, geneticin

Regeneration frequencies for each selection scheme are given in Table 7.3. Analysis of

variance (Statistix 8.0 software) did not show substantial (P≤ 0.9503) differences between

cultivars. The effect of single selection schemes, however, differed significantly between

cultivars. Recognisable variability (P≤ 0.0852) in response to selection schemes was


105

observed in THP whereas slight differences (P≤ 0.1833) were noted in OR where EC

clones were selected with a single selective agent (G or H). Initial regeneration abilities of

the non transformed ECS of both cultivars were higher than what was observed for

transgenic colonies, which is in accordance with the effect of factors associated with

reduced regenerability of selected plant cells or tissues (see section 4.1.5).

Table 7.3 Regeneration frequencies of different selection schemes after Agrobacterium- mediated co-transformation of banana ECS and 2-3 months of selection

Gene/gene combinations

Cht-2 Cht-3 Rs-afp2 Cht-2/Rs-afp2 Cht-3/Rs-afp2 Cultivar Control H H G H G H+G H G H+G THP 50 13 11 09 08 05 18 7.5 04 35 OR 65 14 18 10 11 11 00 14 07 00

THP, ‘Three Hand Planty’; OR, ‘Orishele’; H, hygromycin; G, geneticin

7.3. Efficiency of co-transformation in banana ECS

PCR was used to screen 10-20 regenerated co-transformed THP lines for the presence of

all transferred genes, i.e. Cht-2 or Cht-3 as well as Rs-afp2, nptII and hpt (Table 7.4). All

co-transformants of THP gave positive signals for all transgenes. This result is highly

expected because a stringent combined selection pressure was employed to obtain these

lines. However, the same combined selection regime resulted in zero survival of

embryogenic colonies in OR (Table 7.2), thus no plants could be analysed.

More varied co-transformation frequencies were observed between cultivars on simple

selection schemes (Table 7.5). All co-transformants of Cht-2/Rs-afp2 (A.2) and Cht-3/Rs-

afp2 (B.2), i.e. all transformants of OR, gave co-transformation frequencies of between

80-100% with respect to all transgenes. This consistence in response of OR to single

selection is illustrated by PCR analysis of co-transformants of Cht-3/Rs-afp2 (B.2.H in

Table 7.5) selected by hygromycin alone, when 95% of lines were co-transformed with

respect to all four genes studied (Figure 7.1). Putatively co-transformed lines B.2.H.02,

B.2.H.03, B.2.H.04, B.2.H.05, B.2.H.06, B.2.H.07, B.2.H.09, B.2.H.16, B.2.H.17,

B.2.H.23 and B.2.H.25 were analysed. Line B.2.H.17 in lane 9 had negative signals for

both Cht-3 and hpt implying that it was an escape in respect to hygromycin selection.

However, such claim needs also to be confirmed by other gene detection procedures.

Chapter seven

106

Table 7.4 PCR profile of putatively co-transformed ‘Three Hand Planty’ lines containing Cht-2, Rs-afp2, nptII, and hpt or Cht-3, Rs-afp2, nptII, and hpt genes and obtained by combined selection

Amplified DNAs were present (+) or absent (-); A, combination of Cht-2/Rs-afp2; B, Cht-3/Rs-afp2. A.1, Cht-2/Rs-afp2 in ‘Three Hand Planty’; B.1, Cht-3/Rs-afp2 in ‘Three Hand Planty’

Alternatively, the hpt gene (or the T-DNA itself) could have been excised after the

selection phase. By screening Arabidopsis and tobacco shoots for the presence of hpt, 16

shoots did not contain the hygromycin resistance marker on which they were selected. Out

of these shoots 4 were containing the non-selected T-DNA (De Neve et al., 1997). These

lines did not contain the T-DNA that contains the selectable marker gene selected for. The

authors proposed that the T-DNA could have become removed from plant cells during

shoot regeneration (De Neve et al., 1997).

Though ECS were transformed with Agrobacterium cells co-transformed with 2 T-DNAs,

not all ECS cells had both T-DNA inserted. For example, line B.2.H.25 in lane 11 has only

positive signals for hpt and Cht-3. This implies that the two heterogeneous T-DNAs were

independently transferred to banana cells, both single and co-transformants are generated

in this co-transformation approach, and that selection with one selective agent allows the

generation of single transformants.

Line Cht-2 Rs-afp2 nptII hpt Line Cht-3 Rs-afp2 nptII hpt A.1.01 + + + + B.1.01 + + + + A.1.03 + + + + B.1.03 + + + + A.1.04 + + + + B.1.05 + + + + A.1.05 + + + + B.1.06 + + + + A.1.06 + + + + B.1.07 + + + + A.1.07 + + + + B.1.09 + + + + A.1.08 + + + + B.1.10 + + + + A.1.09 + + + + B.1.11 + + + + A.1.10 + + + + B.1.12 + + + + A.1.11 + + + + B.1.13 + + + + A.1.12 + + + + B.1.14 + + + + A.1.13 + + + + B.1.15 + + + + A.1.14 + + + + B.1.17 + + + + A.1.15 + + + + B.1.18 + + + + A.1.16 + + + + B.1.19 + + + + A.1.17 + + + + B.1.26 + + + + A.1.18 + + + + B.1.29 + + - + A.1.19 + + + + B.1.31 + + + + A.1.23 + + + + B.1.33 + + + + A.1.25 + + + + B.1.34 + + + +


107

Increased co-transformation frequency in OR could be due to its higher sensitivity to the

selective agents applied. This can explain why the combined selection pressure may have

been too stringent and resulted in zero plants (Table 7.3). Lower co-transformation

frequencies of 60% to 80% were observed for both gene combinations (Cht-2/Rs-afp2, A.1

and Cht-3/Rs-afp2, B.1) in THP (Table 7.5). Reduced sensitivity of THP cells to the

selective agents could account for the lower co-transformation frequencies as this was

100% under the combined selection scheme for both gene combinations (Table 7.4). In

summary, a 60-100% frequency of co-transformation can be achieved in banana even

when only single selection is employed. This rate is quite high in comparison to other data

available. For instance, De Block and Debrouwer (1991) studied co-transformation of

Brassica napus hypocotyl explants by Agrobacterium strains. After selection for the first

T-DNA marker (hpt), the plants were screened for the presence of the second, unlinked T-

DNA marker (nptII), which was present at a frequency of 39% to 85%. In studies based

on Arabidopsis leaf disks, tobacco callus, and the same unlinked T-DNA marker genes

(hpt and nptII) co-transformation frequencies between 21% and 47% were reported among

lines selected with one selective agent (De Neve et al., 1997).

Using the A. tumefaciens C58 strain for co-transformation of tobacco, Li et al. (2003)

obtained 48 independent kanamycin resistant lines. Of these transgenic plants, 35%, 27%,

19% and 19% contained one, two, three and four unlinked transgenes (lignin biosynthesis

genes), respectively, based on PCR analysis.

Figure 7.1 PCR analysis of representative transgenic plants of ‘Orishele’ selected on hygromycin alone and containing hpt, Cht-3, nptII and Rs-afp2 genes. M, 1-kb DNA ladder; -Co, negative plant control; +Co, positive control (mixed plasmids pBI333-EN4-RCG3 and pFAJ3494); lanes 1 to 11, samples from independent co-transformants: B.2.H.02, B.2.H.03, B.2.H.04, B.2.H.05, B.2.H.06, B.2.H.07, B.2.H.09, B.2.H.16, B.2.H.17, B.2.H.23 and B.2.H.25

nptII M 1 2 3 4 5 6 7 8 9 10 11 -Co+Co

hpt M 1 2 3 4 5 6 7 8 9 10 11 -Co+Co

668 bp

M 1 2 3 4 5 6 7 8 9 10 11 -Co+Co Cht-3

414 bp

Rs-afp2M 1 2 3 4 5 6 7 8 9 10 11 -Co+Co

554bp 180

bp

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108

Linked genes, i.e. Cht-2 and hpt, Cht-3 and hpt, and Rs-afp2 and nptII, on the other hand

showed almost perfect co-transformation, as expected: see e.g. B.1 (G) for Rs-afp2 and

nptII and A.2 (H) for Cht-2 and hpt (Table 7.5). Table 7.5 Transformation frequency of Cht-2 or Cht-3 as well as Rs-afp2, nptII and hpt in 20-20 putatively transformed lines of ‘Three Hand Planty’ and ‘Orishele’ as determined by PCR after single selection

Target DNA absent (-ve) or present (+ve); A, combination of Cht-2/Rs-afp2; B, Cht-3/Rs-afp2. A.1, Cht-2/Rs-afp2 in ‘Three Hand Planty’; A.2, Cht-2/Rs-afp2 in ‘Orishele’; B.1, Cht-3/Rs-afp2 in ‘Three Hand Planty’; B.2, Cht-3/Rs-afp2 in ‘Orishele’; (G), geneticin; (H), hygromycin

The consecutive introduction of multiple genes demands the use of multiple selectable

marker genes, which induced public concerns about their release into the environment

(Halpin, 2005). Co-transformation by using a single selectable marker gene may thus help

to reduce these concerns. Results from this study show that this approach could be

successfully used in banana with several genes of agronomic importance.

Figure 7.2 presents results of PCR analyses of putatively co-transformed lines of ´Three

Hand Planty´. Co-transformants, which contain nptII, Cht-3, Rs-afp2, and hpt, were

selected on a combination of geneticin and hygromycin.

The PCR analyses included co-transformants lines B.1.01, B.1.05, B.1.07, B.1.09, B.1.12,

B.1.26, B.1.29, B.1.33, B.1.34 and line B.1.31. In this category (Cht-3/Rs-afp2-THP:

H&G) all putative co-transformants were positive for transgenes Cht-3, nptII and hpt and

Rs-afp2, except for co-transformant line B.1.33 that showed negative signal for transgene

Rs-afp2. Negative signal in this particular line could have been due to the presence of PCR

amplification inhibitors in that specific reaction tube.

Transgenes in co-transformants Cht-2 Cht-3 Rs-afp2 nptII hpt Category (selection)

-ve +ve -ve +ve -ve +ve -ve +ve -ve +ve

A.1 (G) 20 80 NA NA 30 70 20 80 20 80 A.1 (H) 10 90 NA NA 00 100 00 100 20 80 B.1 (G) NA NA 30 70 30 70 30 70 30 70 B.1 (H) NA NA 00 100 40 60 40 60 05 95 A.2 (G) 10 90 NA NA 10 90 10 90 10 90 A.2 (H) 00 100 NA NA 05 95 05 95 00 100 B.2 (G) NA NA 20 80 10 90 15 85 00 100 B.2 (H) NA NA 05 95 00 100 00 100 05 95


109

7.4. Multiplex PCR (MPCR) analysis of co-transformants

7.4.1. Primer combinations and their concentrations

Traditional PCR has a very high sensitivity and specificity and therefore has routinely

been used for the detection of transgenic plants (see also chapters 4-6). However, usually

one target gene is detected in a single PCR reaction. For several genes, multiple reactions

need to be set up, and finding the optimal conditions for each gene may be time

consuming, expensive and labour intensive.

Multiplex PCR (Chamberlain et al., 1988) relies on more than one primer pair, and thus,

under optimised conditions, allows detection of several target sequences in a single PCR

reaction. In this study, MPCR was adapted to detect multiple transgenes in the generated

co-transformed lines. Specifically, the procedure was developed to detect nptII, hpt, Cht-2,

Cht-3 and Rs-afp2 genes in duplex, triplex or tetraplex combinations in co-transformed

lines of ‘Orishele’ and ‘Three Hand Planty’.

Upon inclusion of multiple pairs of gene-specific primers, several factors were tested.

These included combinatorial analysis of primer pairs to assess the rate of primer-dimer

formation. Other parameters were: gradient PCR to establish a suitable common annealing

temperature, variation of primer and template DNA concentrations.

Figure 7.2 PCR analysis of representative putative co-transformant lines of ‘Three Hand Planty’ selected on a combination of geneticin and hygromycin, containing hpt, Cht-3, nptII and Rs-afp2 genes. M, 1-kb DNA ladder; -Co, negative plant control; +Co, positive control (mixed plasmids pBI333-EN4-RCG3 and pFAJ3494); 1 to 10, DNA samples from independent co-transformant lines including B.1.12, B.1.26, B.1.29, B.1.33, B.1.01, B.1.34, B.1.07, B.1.09, B.1.05 and line B.1.31; Co- and Co+, negative control (DNA from an untransformed plant) and positive control (mixed pBI333-EN4-RCC2 and pFAJ3494), respectively.

414 bp

Cht-3 M 1 2 3 4 5 6 7 8 9 10 -Co +Co

554 bp

nptII

M 1 2 3 4 5 6 7 8 9 10 -Co +Co

Rs-afp2

180 bp

M 1 2 3 4 5 6 7 8 9 10 -Co +Co

`Three Hand Planty’ hpt

M 1 2 3 4 5 6 7 8 9 10 -Co +Co

668 bp

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110

Combinations of a maximum of four primer pairs (hpt, nptII, Cht-2 or Cht-3, and Rs-afp2,

see Table 3.2) were evaluated. Duplex assessments were hpt/nptII, and Cht-2 or Cht-3/Rs-

afp2, whereas triplex amplification of hpt/nptII/Rs-afp2 and hpt/nptII/Cht-2 or Cht3 were

analysed. Duplex amplification was effective at equimolar primer concentration of 0.5

µM. Slightly reducing or increasing primer concentrations in cases where higher and

lower amplification was observed, respectively, gave balanced amplification in a few

primer combinations. Of the triplex combinations, only hpt/nptII/Cht-3 gave consistent

results. All tetraplex combinations consistently gave PCR products only for hpt and nptII

with inconsistent occurrence of PCR products from other genes.

Amplification reactions with primer concentrations of hpt and nptII both at 0.5 μM and

Cht-3 at 0.75 µM, gave visible signals for all the three genes. However, signals for hpt,

were weaker. Adjusting the primer concentrations to 0.5, 0.43, and 0.4 µM of Cht-3, hpt

and nptII, respectively, gave signals of almost equal strength. These primers were

commonly annealed at 56°C for 1 min; sequences elongation was done at 72°C for 2 min;

and final sequence elongation at 72 °C for 7 min in a PCR cycle with initial denaturation

temperatures of 94 for 4 min and 94°C for 0.5 min with 35 repeated cycles of

amplification.

Products of this triplex PCR amplification (Figure 7.3) were even more intense than what

was observed in parallel single primer-pair PCR reactions (Figure 7.2). Putatively co-

Figure 7.3 Triplex (A) and single PCR (B-D) analysis of 10 representative transgenic plants of ‘Three Hand Planty’ selected on hygromycin alone and containing hpt (A), nptII (B), Cht-3 (C) and Rs-afp2(D) genes. M, 1-kb DNA ladder; -Co, negative plant control; +Co, positive control (mixed plasmidspBI333-EN4-RCG3 and pFAJ3494); lanes 1 to 10, samples from independent co-transformants: B.1.H.04, B.1.H.09, B.1.H.03, B.1.H.15, B.1.H.11, B.1.H.34, B.1.H.21, B.1.H.24, B.1.H.39, and B.1.H.01.

D

M 1 2 3 4 5 6 7 8 9 10 -Co W+Co

A B

C

M 1 2 3 4 5 6 7 8 9 10 -Co W+Co

668 bp554 bp

414 bp

414 bp

180 bp

554 bp


111

transformed lines of ‘Three Hand Planty’ with Cht-3/Rs-afp2 selected on hygromycin

alone (B.1.H, Table 7.5) were: B.1.H.01, B.1.H.03, B.1.H.04, B.1.H.09, B.1.H.11,

B.1.H.15, B.1.H.21, B.1.H.34, B.1.H.24, and B.1.H.39.

All lines analysed showed positive signals for nptII and Rs-afp2 in single primer pair PCR

analyses (Figure 7.3 B, D). Co-transformant line B.1.H.15 showed negative signal for Cht-

3 and positive signal for nptII and Rs-afp2 in both single PCR (Figure 7.3 B-D) and triplex

PCR (Figure 7.3 A) analysis indicating that it contained no Rs-afp2 though a positive

signal was shown with nptII primers. In this particular case, triplex PCR analysis was

equally specific, though with clearer signals. In general, prominent amplification was

observed in triplex analysis results compared to single primer pair PCR, especially for

Cht-3 and Rs-afp2 reactions, where weak signals were observed.

7.4.2. Effect of increased template DNA

Several attempts were done to increase the yield of both single PCR and MPCR products.

These included running gradient PCR, using different quantities of Taq polymerase, varied

concentrations of MgCl2, and finally doubling the concentration of template DNA. Of all

these tests, only doubling template DNA was effective. Doubled template DNA

concentration gave more intense signals in both MPCR and single PCR analysis. Co-

transformant lines in Figure 7.3 were also used here (Figure 7.4).

M 1 2 3 4 5 6 7 8 9 10-CoW+Co

DC

Figure 7.4 Triplex and single PCR analysis with doubled concentrations of template DNA samples from 10 representative transgenic plants of ‘Three Hand Planty’ selected on hygromycin alone and containing hpt (A), nptII (B), Cht-3 (C) and Rs-afp2 (D) genes. M, 1-kb DNA ladder; -Co, negative plant control; +Co, positive control (mixed plasmids pBI333-EN4-RCG3 and pFAJ3494); lanes 1 to 10, samples from independent co-transformants: B.1.H.04, B.1.H.09, B.1.H.03, B.1.H.15, B.1.H.11, B.1.H.34, B.1.H.21, B.1.H.24, B.1.H.39, and B.1.H.01.

BA

M 1 2 3 4 5 6 7 8 9 10-Co W+Co

414bp

668bp

554bp

414bp

Chapter seven

112

7.5. Southern blot analysis of co-transformed banana lines

Southern blot analysis was performed with total DNA samples from putatively co-

transformed lines of ‘Three Hand Planty’ with Cht-2 or Cht-3/Rs-afp2 obtained by

combined selection.

Co-transformed lines A.1.01, A.1.03, A.1.07 and A.1.23 showed three integration loci.

The highest number of integration loci with four was observed in lines A.1.25 and A.1.19

while one insertion was found in line A.1.13. Line A.1.02 did show negative signal for

Cht-2 probe. A similar number of one to four integrations were observed in

transformations with single vectors (see section 6.4.2.2). The different integration profile

of each co-transformants indicates that all these lines represent independent transformation

events.

Positive signals for hpt transgene shown in Figure 7.6 are a good indicator that there was

integral introduction of T-DNAs containing hpt linked to Cht-2 as constructed in the

binary vector pBIN333-EN4-RCC2.

Integration profiles of transgene hpt in all lines that showed positive signals within a range

of 1 to 4 as observed previously. The highest number of integration loci was observed in

line A.1.07 with the least observed in line A.1.03. Co-transformant line A.1.02 gave

negative signal for the integration of hpt.

To evaluate whether these lines were co-transformants in respect to Rs-afp2 and nptII

transgenes, the same lines were evaluated with southern blot analysis. Genomic DNA of

these lines was digested overnight with 50U of BamHI followed by other southern analysis

procedural steps and finally probed for the integration of transgenes Rs-afp2 (Figure 7.7)

21.0 kb

5.0 kb 4.2 kb

3.5 kb

A.1.23

A.1.19

A.1.13

A.1.02

A.1.03

A.1.07

A.1.01

A.1.25

THP.C

o

MW

MIII

Figure 7.5 Southern blot analysis of co-transformed lines of ‘Three Hand Planty’ (THP). HindIII digested total DNA was hybridised with a PCR probe from the Cht-2 gene coding sequence. M, molecular weight marker III (Roche); THP.Co, untransformed THP control.


113

and nptII (Figure 7.8). All co-transformant lines showed simple integration profiles as

compared to single-transgene transformants.

For each particular gene, integration loci between 1 and 3 were observed in co-

transformant lines A.1.03, A.1.07, A.1.13 and A.1.19. Lines A.1.01, A.1.23 and A.1.25

had higher numbers of integration loci. Line A.1.01 had 5 integration loci for each of the

transgenes Rs-afp2 and nptII. Four integration loci of Cht-2 and 5 of transgene Rs-afp2

were observed in line A.1.23 where line A.1.25 had 4 integration loci for each of

transgenes nptII, Cht-2 and Rs-afp2. Transformed line A.1.02 which gave a negative

signal for Cht-2 (Figure 7.5), hpt (Figure 7.6), and nptII (Figure 7.8) had a positive signal

for Rs-afp2 (Figure 7.7), implying that it was a single-gene transformant. Integration

profiles of Cht-2, hpt, Rs-afp2 and nptII transgenes in most of the evaluated co-

transformant lines are in agreement with the observed high transformation frequencies

shown by PCR analyses.

Figure 7.6 Southern blot analysis of co-transformed lines of ‘Three Hand Planty’ (THP). BamHIdigested total DNA was hybridised with a PCR probe from the hpt gene coding sequence. M, molecular weight marker III (Roche); THP.Co, untransformed THP control.

A.1.19

A.1.23

A.1.13

A.1.02

A.1.07

A.1.03

A.1.25

A.1.01

21.0 kb

5.1 kb

4.2 kb

3.5 kb

MW

MIII

THP.C

o

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114

7.6. Conclusion

High co-transformation frequencies were achieved using a single Agrobacterium strain

containing two binary vectors. Co-transformation frequencies were higher compared to

what has been obtained using several Agrobacterium strains each containing a different

binary vector.

Analyses of co-transformants using PCR showed that up to three transgenes can be

detected when their suitable primer pairs are mixed in a single PCR reaction (Multiplex

PCR). Co-transformed lines of Cht-2/Rs-afp2 transformation category proved that lines

Figure 7.8 Southern blot analysis of co-transformed lines of ‘Three Hand Planty’ (THP). HindIII digested total DNA was hybridised with a PCR probe from the nptII gene coding sequence. M, molecular weight marker III (Roche); THP.co, untransformed THP control.

21.0 kb

5.1 kb

4.2 kb

3.5 kb

MW

MIII

THP.C

o

A.1.01

A.1.25

A.1.03

A.1.07

A.1.02

A.1.13

A.1.19

A.1.23

Figure 7.7 Southern blot analysis of co-transformed lines of ‘Three Hand Planty’ (THP). BamHIdigested total DNA was hybridised with a PCR probe from the Rs-afp2 gene coding sequence. M, molecular weight marker III (Roche); THP.co, untransformed THP control.

A.1.23

A.1.19

A.1.13

A.1.02

A.1.07

A.1.03

A.1.25

A.1.01

21.0 kb

5.1 kb

4.2 kb

3.5 kb

THP.C

o

MW

MIII


115

that gave positive signals with PCR were actually transgenic since respective transgene

integration was confirmed by Southern blot analysis. Integration patterns of individual

genes varied in different lines analysed. For all genes, integration loci ranged from 1 to 5

with the highest integration loci observed lines A.1.01 and A.1.25. Lines A.1.3 and A.1.13

had lower integration loci (1 to 2) for all the four genes integrated.

Table 7.6 PCR analysis and integration patterns of four different genes in selected co-transformant lines of

‘Three Hand Planty’

Line PCR analysis Southern blot analysis No. of integration loci

hpt Cht-2 nptII Rs-afp2 hpt Cht-2 nptII Rs-afp2 A.1.01 + + + + 1 2 5 5 A.1.03 + + + + 1 2 1 2 A.1.07 + + + + 3 3 1 3 A.1.13 + + + + 1 1 2 1 A.1.19 + + + + 3 3 1 3 A.1.23 + + + + 2 4 4 4 A.1.25 + + + + 2 4 2 5

Target DNA absent (-) or present (+); A, combination of Cht-2/Rs-afp2; these lines were subjected to combined selection.

These results show random gene integration patterns and the numbers of integration loci

were not different from those observed in single T-DNA integration (Figure 6.6 or 6.7).

From these results co-transformation approach using Agrobacterium has high potential for

transgene stacking, a future desirable approach for multigene traits in banana.

General conclusion and discussion

117

Chapter 8. General conclusion and discussion

Bananas and plantains are important staple food crops, which are difficult to breed due to

high sterility of commercial breeding parents. Caused by the lack of an overwintering

period, this crop suffers from numerous diseases and pests, in particular from fungal

diseases. Hence the need to generate novel transgenic bananas and plantains with genes

which might confer elevated resistance against pathogenic fungi including Mycosphaerella

fijiensis.

In this work, the efficiency of Agrobacterium-mediated (AmT) and particle bombardment-

mediated (PmT) transformation systems were first compared. Then the AmT system was

improved and used to develop transgenic banana and plantain with single candidate genes.

Finally, the same genes were combined in co-transformation experiments to generate

transgenic bananas as a proof of concept for generating durable resistance.

8.1. Comparison of AmT and PmT systems

AmT system was first established for a broad range of dicotyledonous plants. In the

assumption that A. tumefaciens has a narrow host range in monocots, PmT system became

routine in genetic engineering of these species. Transgenic lines in banana were first

produced by PmT of ECS from several cultivars. Sági et al. (1995) reported PmT of

‘Bluggoe’ (ABB), ‘Three Hand Planty’ (AAB) and ‘Williams’ (AAA) followed later by

genetic modification of the commercial dessert (AAA) banana ‘Grand Naine’ (Becker et

al., 2000). As AmT appears to generate lines with simple and well defined integrations

with single or few copies per insertion (De la Riva et al., 1998), more attention was given

to the development of an efficient AmT system in banana. Infection of banana cells with

A. tumefaciens was indicated by chemotactic movement and polar attachment of bacterial

cells to ECS of banana (Hernandez et al., 1999). Also, effective AmT of ‘Grand Naine’

was reported (May et al., 1995) using corm slices. Due to associated chimerism, corm

slices were replaced by ECS (Ganapathi et al., 2001) in ‘Rasthali’ (AAB), and ‘Grand

Naine’ and ‘Lady Finger’ (Khanna et al., 2004). In the current study, we used AmT and

PmT of a wide range of cultivars and we considered differences in regeneration, gene

transfer efficiency (based on PCR results), and transgene integrations at the same time.

Our results showed variable gene transfer rates (indicated by transient GUS expression)

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highlighting the influence of culture conditions x cultivar interactions. Higher gene

transfer rates were observed in ‘Grand Naine’ with 1400 blue foci per Petri dish. Further,

both systems had similar gene transfer rates in ‘Three Hand Planty’ with around 600 foci.

Significant variations, however, appeared in selection phase and shoot regeneration. For

stable transformation AmT system had significantly higher numbers of recovered cell

colonies in all the four tested cultivars. This indicates extensive loss of transient GUS

expressing cells in PmT, which could be due to cell death caused by injuries. Houllou-

Kido et al. (1992) also observed similar trends and proposed that the associated shock

waves, sound waves and cellular membrane injuries caused by particles account for such

difference. Shoot regeneration again showed that AmT was better in all the cultivars.

These experiments also showed higher transformation efficiencies in AmT than in PmT,

based on PCR data. In all the cultivars AmT showed over 90% transformation efficiency

whereas for PmT it fell between 50-80%. Moreover, AmT transformed lines showed

sharper and deeper blue spots with the histochemical GUS assay. This is reported to be

due to the presence of an intron within the uidA gene used in the AmT system.

The number of integration loci of the uidA gene was lower in banana in the AmT

generated lines than with the PmT system. Comparing the two gene transfer systems in

rice, Dai et al. (2001) reported 7% and 22% transformation efficiencies for AmT and

PmT, respectively. Similar experiments in barley reported 25% transgenic lines showing

stable transformation in PmT system. In contrast, using AmT a stable transformation

efficiency of 71.5% to 100% was detected (Travella et al., 2005). Moreover, Becker et al.

(1995) reported a transformation frequency by PmT of only 11% in ‘Grand Naine’.

AmT system is frequently associated with low transgene insertions and copy numbers (De

la Riva et al., 1998; Gelvin, 2003). Preliminary analysis showed slightly more integration

events in PmT than AmT system. The five lines analysed (Figure 4.11) for PmT had two

to five integrations whereas a few from AmT had one to three insertions. Of the barley

transgenic lines, analysed while comparing AmT and PmT systems, 60% of the PmT

transformed lines had more than eight transgene insertions whereas all lines generated

with AmT system had one to three integration events (Travella et al., 2005). Furthermore,

transgene integration analyses in European elite wheat varieties (Rasco-Gaunt et al., 2001)

gave more details of transgene integration patterns in PmT system. In this study, of the 25

lines analysed 32% had one-two insertions, 52%, had three-five, and 16% had six-eight

insertions per line. In another wheat study, Cheng et al. (1997) reported that presence of a

single insertion per line was in 25% of all lines and 85% of the lines contained four or five


119

insertions. In banana, Becker et al. (2000) reported five to nine integrations per line in

‘Grand Naine’ transformed via PmT system. AmT system in banana shows lower number

of transgene integrations. Ganapathi et al. (2001) reported one to four whereas Khanna et

al. (2004) reported only one to three per line in all transgenic lines analysed. These reports

support our observations in these experiments and present AmT system as a method of

choice for gene transfer in banana.

8.2. Optimisation of AmT system

To extend the use of AmT several optimisations have been done so far. These include

desiccation of explants, osmotic treatment, use of anti-necrosis agents, application of

surfactants, modification of infection and co-cultivation media, use of different selectable

marker genes, and a wide range of antibiotics (Opabode, 2006 and references therein). To

optimise the AmT system for banana cells, factors that influence transient expression,

stable gene expression and subsequent shoot regeneration were analysed with the modified

GFP gene sgfpS65T. Transformation of banana ECS with the sgfpS65T gene resulted in

variable numbers of putative transformants in different banana cultivars. Molecular

analysis (PCR, RT-PCR and Southern blot hybridisation) confirmed that the regenerated

banana and plantain lines from cultivars ‘Grand Naine’, ‘Obino l’Ewai’, ‘Orishele’ and

‘Three Hand Planty’ were transgenic. The relatively low number of embryogenic colonies

surviving and shoot regeneration after AmT indicated potential areas where optimisation

could increase transformation frequency. Results also indicated that the observed

variations at both the transient and stable gene expression level depended on the cultivar

tested.

Transformation of the four cultivars ‘Grand Naine’, Three Hand Planty’, Obino l’Ewai’,

and ‘Orishele’ showed a high number of green fluorescent cells 2 weeks after

transformation (Figure 5.5A). With increasing time in culture, the number of such cells

dropped significantly (Table 5.2). Death of cells after Agrobacterium infection has been

reported to be due to necrosis and induced hypersensitive response (Hansen, 2000). This

phenomenon has been reported in banana cells (Khanna et al., 2007) and is significantly

reversed when plant cells are transformed with anti-apoptosis genes (Dickman, 2001,

2004; Khanna et al., 2007). Although this approach is interesting, plant cells expressing

anti-apoptosis genes do not respond to biotic stress. In future this approach could be

improved by transient expression of these genes only during Agrobacterium infection.

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Here we discuss the manipulation of ECS age, infection length, ECS volume during co-

cultivation and the use of spermidine to increase shoot regeneration of cell colonies after

selection. Manipulation of ECS age entails the determination of the suitable age of banana

cells at which there is enhanced gene transfer. Our results show that transforming banana

cells 6-7 days after the previous subculture enhances gene transfer efficiency.

Optimisation of ECS age has deeper effect on gene transfer be it AmT or PmT. This is

clear when we consider the ECS growth or multiplication curve. In stable and established

ECS cultures of banana, subculture is done after 12 to 14 days (Strosse et al., 2006) and

this is the time when cell growth rate flattens. The period of the first 6-7 days coincides

with exponential growth phase of ECS, which is characterised with high cell division

rates. How is this related to increase gene transfer? There is increasing evidence that

young and fast growing plant cells are highly susceptible to Agrobacterium infection and

hence gene transfer. Exploitation of the use of highly dividing cells has been reported by

several authors. These include reports of increased gene transfer by wounding, pre-culture

on auxin-rich media, and use of previously subcultured plant cells. Pre-culture of explants

prior to Agrobacterium is frequently reported to increase gene transfer efficiency

(Sangwan et al., 1992; Weir et al., 2001). The observed increase in gene transfer

efficiency was attributed to be due to stimulation of cell division (Sangwan et al., 1992)

and activation of DNA replication machinery during pre-culture of plant cell. Chateau et

al. (2000) observed similar effects in Arabidopsis. Recently, several Agrobacterium gene

transfer system reviews have highlighted the importance of cell division during gene

transfer (Tzfira et al., 2002; Gelvin, 2003; Arias et al., 2006). In these reviews, authors

report that actively dividing cells are required for efficient gene transfer and integration.

These reports are based on the findings that intracellular T-DNA transport and subsequent

T-DNA integration is facilitated by host plant cell proteins (Tzfira et al., 2002 and

references therein). Arias et al. (2006) further emphasized the importance of cell division,

explaining that cell cycle phases S-M were important for plant cell transformation. They

added that cell cycle phase S is important in transient expression or gene transfer whereas

cell cycle M could be important for integration. The reasoning is based on the fact that

plant cell DNA repair machinery is more active during cell division due to on-going DNA

replication processes (Tzfira et al., 2002).

Transformation frequencies could be increased by fine-tuning the infection length (4 to 5 h

at 5-7 days after subculture), choosing a suitable ECS volume (200 to 300 µL) during co-

cultivation, and by improving the regeneration capacity of the ECSs via the inclusion of


121

the polyamine spermidine. Contrary to our observations, subculturing period did not have

a significant effect in AmT of beans (De Clercq et al., 2002). This could be because

subculturing did not significantly increase cell division in the tested regeneration-

competent callus.

Observed increase in gene transfer by manipulating infection length and ECS volumes

during co-cultivation could be due to increased access of agrobacteria to banana cells. In

Centrifugation Assisted Agrobacterium gene Transfer (CAAT) procedure (Khanna et al.,

2004), gene transfer is improved by the increased access of agrobacteria to plant cells and

the use of low ECS density during infection and co-cultivation. Hernandez et al. (2006)

reported an AmT system in which ECS are infected for 6 h prior to co-cultivation. To

investigate whether this infection length was appropriate, variable infection lengths were

evaluated. Our results show continuous increase in gene transfer (measured by

quantification of blue foci after histochemical GUS assay) with increasing infection time.

In ‘Grand Naine’ gene transfer increased from 209 blue foci per Petri dish (after 4 h of

infection) to 922 blue foci after 10 h. In the case of ‘Three Hand Planty’, frequently used

as a model cultivar for AmT system, 1169 blue foci were counted after 4 h and which

further increased to 1311 after 6 h. Whereas the results show 6 h infection length as

optimum for ‘Three Hand Planty’, infection length could be up to 10 h in ‘Grand Naine’.

This also points to a previous remark that optimisation will remain genotype or cultivar

dependent. Although results on gfp expression indicated that transient gene expression

may not be directly related to stable transformation, it has been reported that conditions

that enhance transient expression do actually result in a higher number of transformed

plant lines (Cao et al., 1998; Trifonova et al., 2001; Suziki and Nakana, 2002). With

emerging evidences that perfect synchrony of transformation and cell cycle (Arias et al.,

2006) would result in enhanced integration, a combination of these gene transfer

optimisations become crucial for an efficient AmT system.

Addition of polyamine spermidine, at low concentration increased shoot regeneration of

cell colonies after AmT. For instance, at 0.1 mM spermidine, a regeneration frequency of

over 80% was observed in ‘Three Hand Planty’ and regenerated plants were more

vigorous. Thus, though the number of independent transformants depended primarily on

the cultivar, the modifications resulted in an increased number of high-quality shoots. Our

observations are supported by observations in other crops where spermidine was used. In

onions, at 0.1mM (Martinez et al., 2000), rice at 0.5 mM (Shoeb et al., 2001), wheat at

100 mM (Khanna and Daggard, 2003), and pine at 1.5 mM (Tang et al., 2004) spermidine

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significantly increased shoot regeneration. We propose that the improvement of gene

transfer by targeting actively dividing cells and increasing shoot regeneration from cell

colonies thereafter strongly improve AmT system of banana cells.

8.3. Integration of rice chitinase in banana

Rice chitinase genes (Cht-2 and Cht-3) were introduced into ‘Grand Naine’ and ‘Gros

Michel’. Since we had planned to evaluate the performance of these genes in the field, it

was a pre-requisite to introduce these genes in banana cultivars that are important in

Uganda and are genetically related to the EAHB (East African Highland Bananas)

cultivars. ‘Grand Naine’ being a commercial variety would complicate the intellectual

property management in case a highly resistant line was identified. Hence, ‘Gros Michel’

was proposed for field evaluations in Uganda. Rice chitinase genes have been integrated

and expressed in many plant species to provide protection against fungal diseases. These

crops include cucumber (Tabei et al., 1998; Kishimoto et al., 2002), chrysanthemum

(Takatsu et al., 1999), grape (Yamamoto et al., 2000), rice (Datta et al., 2001) and Italian

ryegrass (Takahashi et al., 2005). In all these cases increased protection to symptom

development was observed. In the current study we introduced two rice chitinases genes in

an approach to protect banana against Mycosphaerella fijiensis the causative agent of

Black Sigatoka Disease.

In these experiments different lines were regenerated and these included 45 lines of

‘Grand Naine’ containing Cht-2; 13 lines with Cht-3; and 26 and 39 ‘Gros Michel’ lines

containing Cht-2 and Cht-3, respectively. PCR screening of these lines, showing over 90%

transformation efficiency, confirmed that they contained rice chitinases genes.

Southern blot analysis revealed the presence of banana genomic sequences with high

homology to rice chitinase genes. Alignment analysis combined with BLASTn search

indicated that these sequences were possibly banana chitinases as reported before

(Clendennen et al., 1997; Peumans et al., 2002). These sequences are indicated as BanChi-

1 (GenBank accession number AJ277278), BanCht-2 (AJ277279), BanCht-3 (AF416677)

and BanCht-4 (Z99966.1). Their nucleotide sequences showed over 70% similarities,

confirming the observed signals in the Southern blots (Figure 6.4). Note that the codes

BanCht-1 to BanCht-4 are arbitrarily given and do not in anyway refer to chitinase

standard classifications. Hence, it would be interesting to understand the characteristics of

these chitinase homologues in banana. Southern blot analysis also detected stable


123

integration of rice chitinase genes Cht-2 and Cht-3 in the banana genome. Integration

profiles of Cht-2 and Cht-3 showed a range of one to four random insertion loci for both

genes. However, in ‘Gros Michel’ a lower number of integration loci and transgene copy

numbers were observed as compared with ‘Grand Naine’. Transgenic ‘Gros Michel’ lines

showed one to two integration sites whereas in ‘Grand Naine’ lines the number of

integration sites ranged from one to four. For Cht-3 the copy numbers ranged from one to

seven with higher copy numbers observed in ‘Grand Naine’ lines. The results suggest a

cultivar effect on the observed integration patterns. However, this needs verification as

only two banana cultivars were tested.

Twenty-six lines of transgenic ‘Gros Michel’ lines were selected for field evaluation

against Black Sigatoka. Leaf discs of mature leaves will be tested in vitro against the

causal pathogen M. fijiensis and the response compared to the performance in the field in

Uganda. The expression levels of the transgenes will be followed during growth and

development of the plant while at the same time resistance will be evaluated. This

information will be of high value for the development of the next generation of transgenic

banana and plantain with genes of agronomic interest.

8.4. Co-transformation with rice chitinase and a defensin

Agrobacterium-mediated co-transformation presents a suitable approach in which the most

desirable traits like durable resistance in fertile and diploid plants like banana can be

introduced into highly developed and unique banana cultivars, for example EAHB

cultivars. Different approaches have been used in Agrobacterium-mediated co-

transformation of plant cells. Frequently used approaches include one Agrobacterium

strain/ one binary vector containing two distinct T-DNAs (Depicker et al., 1985), two

Agrobacterium strains/ two binary vectors (De Block and Debrouwer, 1991; De Block et

al., 1998), and two binary vectors/one Agrobacterium strain (De Framond et al., 1986;

McKnight et al., 1987; Daley et al., 1998). In this experiment we introduced two different

binary vectors in a single Agrobacterium strain EHA105 and used it to infect ECS lines of

cultivars ‘Three Hand Planty’ and ‘Orishele’. Each Agrobacterium cell contained two

binary vectors pBI333-EN4-RCC2/pFAJ3494 (denoted A) and PBI333-EN4-

RCG3/pFAJ3494 (denoted B). Components of these binary vectors are presented in

section 3.3.1. Transformed ECS were selected on media either containing single or double

selective agents. Selection and maintenance of two different and independently integrated

transgenes was independent of whether two or one selectable marker genes were used to

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select embryogenic colonies. After the selection process variable number of shoots were

regenerated and later analysed for the presence of four different transgenes.

PCR analysis of co-transformants showed that up to three transgenes can be detected

simultaneously when suitable primer pairs are mixed in a single PCR reaction (Multiplex

PCR). The efficiency of MPCR was influenced by the type and quantity of primers and the

quality of the DNA template. Increasing the template quantity generated intense

amplification signals, implying that MPCR amplification required more template-DNA

than single PCR. Amplification of selectable marker genes was reproducible and efficient

over a wide range of annealing temperatures. Given the time to obtain results, chemical

components involved, and detection sensitivity, MPCR was superior to normal PCR.

MPCR analysis will become a necessity when the presence of multiple transgenes has to

be confirmed. Hence a rather simple tool has been adapted for the evaluation of transgenic

banana with stacked genes for providing different new traits to banana and plantain. Co-

transformation frequencies of over 80% were obtained with respect to four transgenes

studied. Selection for a single selectable marker gene was effective in getting co-

transformants containing either Cht-2/Rs-afp2 or Cht-3/Rs-afp2. PCR results showed

higher transformation frequencies compared to what had been reported by using two

Agrobacterium strains each containing one or more different binary vectors. Regenerants

from single selection schemes showed co-transformation frequencies of 60-90%. Higher

co-transformation frequencies in a single selective agent scheme are due to the fact that

co-transformation frequencies do not change whether or not any of the T-DNA is selected

for (De Block et al., 1998). Analyzing these lines using simple PCR would have been time

consuming and expensive.

Our co-transformation frequencies are much higher than what was previously reported

using one Agrobacterium strain/two binary vectors co-transformation approach. Although

co-transformation frequencies are higher the use of two vectors per Agrobacterium cell did

not increase the amount of cells transformed or shoots regenerated as one would otherwise

expect. De Block et al. (1998) reported that this could be due to the following reasons: i)

during transformation or co-transformation a limited number of plant cells are accessible

or competent, ii) not all Agrobacterium cells are induced or physiologically able to

transfer T-DNA, and iii) some cells surfaces could contain chemical substances that inhibit

T-DNA transfer. Different co-transformation frequencies have been reported in other

crops. Using one strain/ one binary vector containing two distinct T-DNAs, Depicker et al.

(1985) reported co-transformation frequencies of 60-70% in tobacco. De Block and


125

Debrouwer (1991) employed a two plasmid/two strain approach in Brassica napus, to

obtain co-transformation frequencies of 39-85%. In a similar approach McKnight reported

co-transformation frequencies of 19% when tobacco leaf explants were co-cultivated with

two strains that carried different T-DNAs on similar plasmids. In Arabidopsis, six

different transgenes were integrated and expressed using one strain/one binary vector

containing six distinct T-DNAs (Goderis et al., 2002). Transgenic lines containing Cht-

3/Rs-afp2 were PCR positive and were confirmed by Southern blot analysis. Although,

simple integration patters are observed, most integration loci in co-transformed lines are

reported to be complex containing a few T-DNAs interspaced with host plant DNA. It is

important therefore, that these lines will be analysed further to understand the nature of

integration loci in co-transformed lines.

.

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List of publications

155

List of publications Articles in Scientific Journals

(i) Hakiza, J.J., Kakuhenzire, R., Kankwatsa, P., Arinaitwe, G., Rukuba, D., and Ngombi, B.F. 2003. Dissemination of knowledge and skills of potato crop management through farmers field schools in Uganda. Uganda Journal of Agricultural Sciences 8: 443-448.

(ii) Arinaitwe G., Rubaihayo P.R., and Magambo M.J.S. 2000. Proliferation rate

effect of cytokinins on shoot proliferation rates in AAA-EA (Musa spp.) cultivars. Scientia Horticulturae 86: 13-21.

(iii) Arinaitwe G., Rubaihayo P.R., and Magambo M.J.S. 1999. Effects of

auxin/cytokinin combinations on shoot proliferation in banana cultivars. African Crop Science Journal 7: 605-612.

Chapter in Books (i) Arinaitwe G., Remy S., Strosse H., Swennen R. and Sági L. 2004. Agrobacterium- and

particle bombardment-mediated transformation of a wide range of banana cultivars. In: Mohan Jain S., Swennen R. (ed.). Banana Improvement: Cellular, Molecular Biology, and Induced Mutations. Science Publishers Inc., Enfield, NH, USA: pp. 351-357. http://www.scipub.net/agriculture/banana-improvement-induced-mutations.html

Articles in Proceedings

i) Arinaitwe G., Kiggundu A., Lamwaka P., Namanya P. and Tushemereirwe W. 2006. Genetic engineering of East African highland banana (EA-AAA) cultivars. Abstract of paper presented in the Tropical Crop Biotechnology Conference (TCBC). 16-20 August 2006, Cairns, Australia.

ii) Arinaitwe G., Remy S., Thiry E., Sági L. and Swennen, R. 2005. Integration of

rice chitinase genes in banana (Musa spp.). 9th International Conference on Agricultural Biotechnology: Ten Years After. 6-10 July 2005, Ravello, Italy.

iii) Swennen R., Arinaitwe G., Cammue B.P.A., François I., Panis B., Remy S., Sági

L., Santos E., Strosse H. and Van den houwe I., 2003. Transgenic approaches for resistance to Mycosphaerella leaf spot diseases in Musa spp. In: Jacome L., Lepoivre P., Marin D., Ortiz R., Romero R., Escalant J.V. (ed.). Mycosphaerella leaf spot diseases of bananas: present status and outlook. Proceedings of the 2nd International workshop on Mycosphaerella leaf spot diseases of bananas. 20-23 May 2002, San José, Costa Rica. INIBAP, Montpellier, France: pp. 209-238. http://www.inibap.org/pdf/IN030306_en.pdf

List of publications

156

iv) Arinaitwe G., Remy S., Strosse H., Swennen R. and Sági L. 2002. Agrobacterium- and particle bombardment transformation of a wide range of banana cultivars. Abstract of paper presented during the 4th and final FAO/IAEA research coordination meeting. 24-28 September 2001, Leuven, Belgium. PROMUSA 11: 9.

v) Arinaitwe G. and Rubaihayo P.R. 1997. Preliminary evaluation of somaclonal variation among East African (AAA-EA) banana cultivars. African Crop Science Proceedings 3: 99-102.

vi) Arinaitwe G., Hakiiza J.J. and Kankwatsa P. 2000. Farmers’ Field School, a

participatory implementation of Integrated Pest Management of Late Blight (IPM-LB). APA Proceedings 2: 254-260.

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