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)
vi
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),
´Obino l´Ewai´ (AAB), ´Orishele´ (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
x
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
xiii
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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xiv
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
General introduction
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
General introduction
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.
Literature review
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.
46
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.
Materials and methods
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.
Materials and methods
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
Materials and methods
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
Materials and methods
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
Materials and methods
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
Materials and methods
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
Materials and methods
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.
62
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.
Comparison of transformation methods
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.
Comparison of transformation methods
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.
Comparison of transformation methods
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
Comparison of transformation methods
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
Comparison of transformation methods
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
Comparison of transformation methods
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
Optimisation of AmT system
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
Optimisation of AmT system
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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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
Optimisation of AmT system
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.
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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
Optimisation of AmT system
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.
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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.
Optimisation of AmT system
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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88
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).
Optimisation of AmT system
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
Transformation with rice chitinase genes
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
Transformation with rice chitinase genes
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.
Transformation with rice chitinase genes
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).
Transformation with rice chitinase genes
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
Transformation with rice chitinase genes
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
Co-transformation of banana with chitinase genes and a plant defensin
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 + + + +
Co-transformation of banana with chitinase genes and a plant defensin
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
Chapter seven
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
Co-transformation of banana with chitinase genes and a plant defensin
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
Co-transformation of banana with chitinase genes and a plant defensin
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.
Co-transformation of banana with chitinase genes and a plant defensin
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
Chapter seven
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
Co-transformation of banana with chitinase genes and a plant defensin
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.
116
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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118
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
General conclusion and discussion
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.
Chapter eight
120
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
General conclusion and discussion
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
Chapter eight
122
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
General conclusion and discussion
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
Chapter eight
124
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
General conclusion and discussion
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.
.
126
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127
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List of publications
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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.