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CHARACTERIZATION, CLONING AND
EXPRESSION OF Bt GENE AGAINST
COTTON PESTS
By
SALAH UD DIN
CENTRE OF EXCELLENCE IN MOLECULAR
BIOLOGY
UNIVERSITY OF THE PUNJAB
LAHORE, PAKISTAN
(2018)
CHARACTERIZATION, CLONING AND
EXPRESSION OF Bt GENE AGAINST
COTTON PESTS
A THESIS SUBMITTED TO
THE
UNIVERSITY OF THE PUNJAB
In Partial Fulfillment of the requirement for the Degree of
DOCTOR OF PHILOSOPHY
In
MOLECULAR BIOLOGY
By
SALAH UD DIN
Supervisor:
Dr. ABDUL QAYYUM RAO
CENTRE OF EXCELLENCE IN MOLECULAR BIOLOGY
UNIVERSITY OF THE PUNJAB
LAHORE, PAKISTAN
2018
ii
I
AUTHOR’S DECLARATION
I hereby state that my Ph.D. thesis entitled “Characterization, Cloning and Expression of Bt
Gene against Cotton Pests” is my own work and has not been submitted by me previously
for taking any degree as research work, thesis or publication from University of the Punjab
or anywhere else in country/world.
At any time, if my statement is found to be incorrect even after my graduation, the
university has the right to withdraw my Ph.D. degree.
_______________
Signature of Deponent
Salah ud Din
June, 2018
II
CERTIFICATE
It is to certify that the research work described in this thesis is the original work of the author
Mr. Salah ud Din and has been carried out under my direct supervision. I have personally
gone through all the data reported herein and certify their correctness/authenticity. I have
found that the thesis has been written in pure academic language and is free from any typos
and grammatical errors. It is further certified that the data reported in this thesis has not been
used in part or full, in a manuscript already submitted or in the process of submission in
partial/complete fulfillment of the award of any other degree from any other institution. It is
also certified that the thesis has been prepared under my supervision according to the
prescribed format of the university and I endorse its evaluation for the award of Ph.D. degree
through the official procedures.
In accordance with the rules of the Centre, data books (1054 and 1261) are declared as un-
expendable document that will be kept in the registry of the Centre for a minimum of three
years from the date of thesis defense.
Signature of the Supervisor: __________________
Name: Dr. Abdul Qayyum Rao
Assistant Professor
III
PLAGIARISM UNDERTAKING
I solemnly declare that research work presented in the thesis titled “Characterization,
Cloning and Expression of Bt Gene against Cotton Pests” is solely my research work with
no significant contribution from any other person. Small contributions/help wherever taken,
has been duly acknowledged and that complete thesis has been written by me.
I understand the zero-tolerance policy of the Higher Education Commission of
Pakistan (HEC) and “University of the Punjab” towards plagiarism. Therefore, as an author
of the above titled thesis, I declare that no portion of my thesis has been plagiarized and any
material used as reference has been properly cited.
I undertake that if I am found guilty of any formal plagiarism in the above titled
thesis even after award of the Ph.D. degree, the university reserves the rights to withdraw/
revoke my Ph.D. degree and that HEC and the university has the right to publish my name
on the HEC/ university website on which names of students are placed who submitted
plagiarized thesis.
_______________
Signature of Deponent
Salah ud Din
July, 2018
IV
ACKNOWLEDGMENTS
In the name of ALLAH, the Most Gracious and the Most Merciful
All praises are for the Almighty “ALLAH” the ultimate source of knowledge, Who has
given me the consciousness to understand and the Holy Prophet “MUHAMMAD” (Peace
be upon him), the source of guidance for the whole mankind.
The work that I have presented here in my thesis could not have been accomplished
had it not been the guidance and help of many people that I am highly thankful to. First
of all, I am grateful to Professor Dr. Tayyab Husnain, Acting Director, Centre of
Excellence in Molecular Biology, University of the Punjab, Lahore, for providing me the
opportunity to complete my research. His support has been of great value in this study.
It seems impossible to pay thanks to my kind and worthy supervisor Dr. Abdul Qayyum
Rao, Assistant Professor and in-charge Plant Biotechnology Laboratory, National
Center of Excellence in Molecular Biology, University of the Punjab, who is indeed a
role model for me. His trust and support has always been the source of vitality and
energy which helped me to complete this task in-time.
I would like to express my sincere thanks to Dr. Ahmad Ali Shahid for his inspiring
guidance and unending support that I gained from him in theory and practical research
work. Special thanks are due to Dr. Idrees Ahmad Nasir for his technical guidance
during field studies of my project. I am deeply and truly thankful to Dr. Bushra Rashid
for her financial support.
I have been lucky enough for having cooperative seniors whose accommodative and
friendly behavior made my work less laborious. I wish to express my sincere gratitude
to Dr. Sameera Hassan, Dr. Naila Shahid, Ms. Ayesha Latif, Ms. Saira Azam, Mr. Tahir
Rehman Samiullah, Ms. Aneela Yasmeen, Dr. Kamran Shahzad Bajwa, Dr. Azmatullah
Khan and Mr. Adnan Muzaffar.
V
I am thankful to my colleagues/lab fellows Ms. Sidra Akhtar, Ms. Ammara Ahad, Ms.
Ambreen Gul, Mr. Muhammad Azam, Mr. Adnan Iqbal, Ms. Amina Yaqoob, Ms. Sana
Shakoor, Ms. Samina Hassan, Mr. Tahir Iqbal, Mr. Ibrahim Bala Salisu for their help and
support. I would like thank Mr. Nada e Hussain, Mr. Tanvir, Mr. Rasheed, Mr. Fayyaz,
and Mr. Nazir for timely assistance in lab/field work.
I am truly thankful to Mr. Mukhtar Ahmed, Mr. Fayyaz Ahmad, Mr. Muhammad Bilal
Sarwar, Mr. Anwar Khan and Mr. Mazhar Hussain for being true friends and
companions in times of need and in deed. I can never thank them much for their
unending support and love.
I would like to acknowledge Higher Education Commission of Pakistan for providing
me six month fellowship under International Research Support Initiative Program to
work in School of Biosciences Cardiff University (UK) to enhance my research
capabilities.
Last but certainly not the least; I am extremely grateful to my parents and my brothers,
who have done every possible and impossible thing just to make sure that I go through
this academic stage with success, satisfaction and ease.
Salah ud Din
VI
Dedicated To
My Family
Who Always Wish to See Me Successful
In Every Walk of Life
VII
LIST OF ABBREVIATIONS
ASAL Allium sativum leaf agglutinin gene
bp base pair
Bt Bacillus thuringiensis
β beta
CaCl2 Calcium chloride
CV Column volume
cm Centimeter
Cry Crystal
CTAB cetyltrimethylammounium bromide
dH2O Distilled water
DMSO Dimethyl sulfoxide
DNA Deoxyribonucleic Acid
dNTPs Dinucleotide Triphosphate
E. coli Escherichia coli
EDTA Ethylene Diamine Tetra Acetic Acid
ELISA Enzyme Linked Immune Sorbent Assay
et al. (et alii) and others
G Gram
GOT Ginning out-turn
GUS Glucuronidase gene
H2O Water
HCl Hydrochloric acid
HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
HgCl2 Mercuric chloride
VIII
IBA Indole Butyric Acid
i.e. That is
Kb Kilo base
Kg Kilo gram
KDa Kilo Daltons
KCl Potassium Chloride
L Liter
LB Luria Broth
Min Minutes
mm Millimeter
mg Milligram
mL Milliliter
mM Milli molar
M Molar
MS Murashige and Skoog
N2 Nitrogen
NaCl Sodium chloride
NaOH Sodium Hydroxide
No. Number
nm Nanometer
ng Nano gram
OD Optical Density
PCR Polymerase Chain Reaction
pmol Pico moles
pH Negative log of hydrogen ions
PBS Phosphate buffer saline
IX
PBST Phosphate buffered saline with Tween
Pfu Pyrococcus furiosus
qRT-PCR Quantitative Real Time Polymerase Chain Reaction
RNA Ribonucleic Acid
RNase Ribonuclease
RTBV Rice tungro bacilliform virus
rpm Rotations per minute
SDS Sodium dodecyl sulfate
Sec Seconds
TAE Tris-acetate EDTA
TMB 3,3',5,5'-Tetramethylbenzidine
Taq Thermus aquaticus
U Unit
UV Ultra violet
V Volts
YEP Yeast Extract Peptone
% Percent
C Degree centigrade
µg Microgram
µl Microliter
X
Summary
The efficacy of Bt toxins is diminishing because insects are becoming resistant to Bt
δ-endotoxins. Second generation Bt vegetative insecticidal proteins (VIPs) could be a
possible alternate of Bt crystal proteins. It has been demonstrated that Vip3Aa do not share
any sequence similarity with any known Cry proteins and bind with different receptors sites
in insect midgut. Plant lectins are carbohydrate-binding proteins that specifically recognize
glycan structures in brush border membrane vesicle receptors present on gut epithelial cells
of insects. In this study, codon optimized synthetic Bt Vip3Aa gene (2370 bp) driven by
constitutive CaMV35S promoter and Allium sativum leaf agglutinin (ASAL) gene (339 bp)
under phloem-specific RTBV promoter in a single 4870 bp (Vip3Aa+ASAL) cassette cloned
in plant expression vector pCAMBIA 1301 under XhoI and HindIII were transformed in a
local cotton variety (CEMB-33). For this purpose, the recombinant
pCAMBIA_Vip3Aa+ASAL plasmid was electroporated in Agrobacterium tumefaciens strain
LBA 4404 and subjected to inoculation with injured cotton embryos for introduction of
desired genes through Agrobacterium to develop high resistance against major chewing and
sucking insects. The putative transgenic cotton plants were confirmed through PCR by using
gene-specific primers. The amplification of 587 bp fragment of ASAL gene and 682 bp
fragment of Vip3Aa confirmed the successful introduction of desired genes in cotton. Total
eighteen plants were found positive out of total fifty-three plants that were shifted in the
field. The transformation efficiency was found to be 1.17%. The transgenic plants were also
screened through Vip3Aa specific dipsticks and expression of GUS marker gene. The mRNA
XI
expression of both the genes was studied in five T1 transgenic cotton lines through
quantitative real-time PCR (qRT-PCR). The comparative analysis of Ct values obtained by
qRT-PCR analysis revealed that the mRNA expression of Vip3Aa gene varied from 2-8.7
folds in the lines L3P2 and L6P3 respectively. Similarly, the comparison of Ct values showed
that the mRNA expression of ASAL gene ranged between 2-5 folds in transgenic lines L4P4
and L34P2. The transgenic line L6P3 showed significantly higher expression for both the
genes. The expression of Vip3Aa protein in T0 and T1 generations was quantified through
ELISA. The maximum protein concentration in both generations was seen in L6P3. The
transgene location and copy number in the cotton genome were determined through
Fluorescence in situ hybridization (FISH) in T2 generation. The transgenic cotton plant from
transgenic line L3P2 showed homozygosity (two copy numbers) on chromosome number 9 at
late telophase stage and one copy number at chromosome number 10 at prophase stage.
Different morphological and physiological parameters of transgenic cotton lines in T1
progeny were studied in comparison to non-transgenic cotton plants. Statistically significant
variation was observed in transgenic cotton lines for all the studied morphological
characteristics, whereas, no significant differences were observed for physiological
parameters. Similarly, the scanning electron microscopic (SEM) images of transgenic and
non-transgenic cotton plants showed no visible difference in fiber morphology between
transgenic and non-transgenic cotton plants which depicts no positive or negative correlation
between expression of insecticidal genes and cotton fiber quality. The final objective of the
current project was to evaluate transgenes efficacy against cotton bollworm (H. armigera)
and whitefly (Bamisia tabaci). The results showed that all the transgenic cotton lines were
XII
significantly resistant to H. armigera as compared to non-transgenic control showing
mortality rates between 78%-100%. Similarly, the transgenic cotton lines L3P2, L5P3, L6P3B
and L6P3C showed 95%; 89%, 89%, and 72% mortality rates respectively in the whitefly
bioassay. This study was unique in a sense that a combination of Bt and plant lectin genes
was used to control major chewing and sucking insects to delay resistance buildup and stable
insect control management.
TABLE OF CONTENTS
AUTHOR’S DECLARATION I
CERTIFICATE II
PLAGIARISM UNDERTAKING III
ACKNOWLEDGMENTS IV
LIST of ABBREVIATIONS VII
SUMMARY X
1. LIST OF FIGURES 1
1. INTRODUCTION 1
2. REVIEW OF LITERATURE 6
2.1. From Al Qutn to Cotton 6
2.2. Genome Evolution of Cotton 6
2.3. World Cotton Outlook 7
2.4. Role of Cotton in Pakistan Economy 8
2.5. Problems in Cotton Cultivation 9
2.6. Major Pests of Cotton Crop 10
2.6.1. Bollworms or Chewing insects 10
2.6.2. American Bollworm (Helicoverpa armigera) 11
2.6.3. Pink Bollworm (Pectinophora gossypiella) 11
2.6.4. Spotted Bollworms: (Earias insulana) 12
2.6.5. Armyworm (Spodoptera litura) 12
2.7. Sucking or Phytophagous Insects 13
2.7.1. Whitefly (Bemisia tabaci) 13
2.7.2. Dusky Cotton Bug (Oxycarenus hyalinipennis) 14
2.7.3. Mealybugs (Phenococcus gossypiphilous) 14
2.7.4. Leafhopper/ Jassids (Amrasca devastans) 14
2.7.5. Thrips (Thrips tabaci) 15
2.8. Conventional Breeding for Cotton Improvement 15
2.9. Discovery of Bacillus thuringiensis (Bt) and Its Toxic Crystal Proteins 16
2.10. Insecticidal Toxins of Bacillus thuringiensis 17
2.11. Crystal (Cry) Proteins or δ-endotoxins 18
2.12. Cytolytic (Cyt) Proteins 19
2.13. Mode of Action of Cry Toxins 19
2.13.1. Classical Model for Action of Cry Toxin 20
2.13.2. Sequential Binding Model 20
2.13.3. Signaling-pathway Model 21
2.14. Vegetative Insecticidal Proteins or VIPs 21
2.14.1. The Vip1/Vip2 (Binary) Toxins 22
2.14.2. Vip3 Toxins 22
2.14.3. Structure and Function of Vip3 Proteins 23
2.14.4. Mode of Action of Vip3 Proteins 23
2.14.5. Proteolytic Processing of Vip3 Toxin 24
2.14.6. Binding of Vip3A to the Midgut Epithelial Cells 25
2.15. Plant Lectins: Natural Plant Defense Proteins 26
2.15.1. Classification of Plant Lectins 26
2.15.2. Three-Dimensional Structure of Plant Lectins 27
2.15.3. Mode of Action of Plant Lectins 28
2.16. Risk Assessment of Plant Lectins in Non-Target Organisms 28
2.16.1. Risk Assessment in Honeybee (Apis mellifera) 29
2.16.2. Risk Assessment in Parasitoids 30
2.16.3. Evaluation of Lectin Based Risks on Vertebrates 31
2.17. Agrobacterium tumefaciens: A Machinery of Genetic Transformation 32
2.17.1. What is T-DNA and How It Is Transformed from Agrobacterium to Plant Cells? 33
2.18. Development of Bt-transformed Plants 35
2.19. The problem of Bt-resistance in Pests and Combating Strategies 36
2.20. Modified Expression of Vip3Aa Gene in Plants 37
3. MATERIALS AND METHODS 39
3.1. Codon Optimization and Chemical Synthesis of Insecticidal Genes 39
3.2. Preparation of Competent Cells 39
3.3. Transformation of Vip3Aa+ASAL Cassette in Top 10 40
3.3.1. Plasmid Isolation from Positive Clones 40
3.3.2. Confirmation of Positive Clones through PCR 41
3.3.3. Confirmation of Positive Clones through Restriction Digestion 42
3.3.4. Elution of DNA Fragments from Agarose Gel 42
3.4. Restriction Digestion of Plant Expression Vector (pCAMBIA 1301) 43
3.4.1. Ligation of Vip3Aa+ASAL Cassette into the pCAMBIA 1301 Vector 45
3.4.2. Transformation and Confirmation of Ligated Product in pCAMBIA 1301 45
3.5. Preparation of Agrobacterium Competent Cells 46
3.6. Transformation of Vip3Aa+ASAL plasmid in Agrobacterium tumefaciens 46
3.7. Confirmation of Electroporation through Colony PCR 47
3.8. Confirmation of Agrobacterium transformation through Restriction Digestion 47
3.9. Cotton Transformation 48
3.9.1. Selection of a Suitable Cotton Variety 48
3.9.2. Delinting of Cotton Seeds 48
3.9.3. Sterilization and Soaking of Seeds 48
3.9.4. Murashige and Skoog (MS) Medium Preparation 49
3.9.5. Bacterial Inoculum Preparation 49
3.9.6. Isolation of Embryos 49
3.9.7. Agrobacterium Mediated Transformation (Shoot Apex Method) 50
3.9.8. Co-cultivation 50
3.9.9. Transformation Efficiency 50
3.9.10. Transferring Transgenic Plants to Soil 51
3.10. Molecular Analyses of Transgenic Cotton Plants 51
3.10.1. Genomic DNA Isolation 51
3.10.2. Polymerase Chain Reaction 52
3.10.3. Transient Expression of GUS Gene 52
3.11. Expression Analysis of Transgenic Cotton Plants 53
3.11.1. Total RNA Extraction 53
3.11.2. Complementary DNA (cDNA) Synthesis 54
3.11.3. Quantitative Real Time qRT-PCR 55
3.11.4. Dipstick Analysis of Putative Transgenic Cotton Plants 55
3.11.5. Quantification of Vip3Aa Protein in Transgenic Cotton Plants through ELISA 56
3.12. Morphological Traits of Transgenic Cotton Plants 57
3.12.1. Plant Height (Inch) 57
3.12.2. Number of Monopodial and Sympodial Branches per Plant 57
3.12.3. Number of Bolls per Plant 58
3.12.4. Ginning out Turn Percentage (GOT %) 58
3.12.5. Scanning Electron Microscopic Analysis of Cotton Fiber 58
3.13. Physiological Traits of Transgenic Cotton Plants 58
3.14. Insect Bioassays 59
3.15. Fluorescence in situ Hybridization (FISH) 60
3.15.1. Preparation of Chromosome 60
3.15.2. RNase Treatment 61
3.15.3. Hybridization 61
3.15.4. Post Hybridization Washes 61
3.15.5. Color Detection Reaction 61
3.15.6. Counterstaining with DAPI 62
3.15.7. Counterstaining with Propidium Iodide (PI) 62
3.15.8. Fluorescent Signal Detection 62
4. RESULTS 63
4.1. Transformation of pUC57_Vip3Aa+ASAL Cassette in E.coli 63
4.2. Confirmation of Vip3Aa+ASAL Cassette in pUC57 Vector 64
4.3. Cloning of Vip3Aa+ASAL Cassette in Plant Expression Vector 65
4.4. Transformation and Confirmation of Ligated Product in pCAMBIA 1301 66
4.5. Electroporation of pCAMBIA1301_Vip3Aa+ASAL into Agrobacterium 67
4.6. Confirmation of Vip3A+ASAL Cassette in Agrobacterium through Restriction Digestion 68
4.7. Cotton Transformation 69
4.7.1. Shoot Apex Method of Agrobacterium Mediated Transformation 69
4.8. Molecular Analyses of Putative Transgenic Cotton Plants (T0 Progeny) 70
4.8.1. Confirmation of Putative Transgenic Cotton Plants through PCR (T0 progeny) 70
4.8.2. Quantification of Vip3Aa Protein through ELISA (T0 progeny) 71
4.9. Advancement of Next Generation (T1 Progeny) 74
4.9.1. Confirmation of Transgenic Cotton Plants through Polymerase Chain Reaction (T1 progeny) 74
4.9.2. The mRNA Quantification of Insecticidal Genes (Vip3Aa and ASAL) in Transgenic Cotton Plants 75
4.9.3. Quantification of Vip3Aa Protein through ELISA (T1 progeny) 76
4.10. Morphological and Physiological Characteristics of Transgenic Plants in T1 Progeny 78
4.10.1. Plant Yield 78
4.10.2. Plant Height 79
4.10.3. Number of Bolls per Plant 79
4.10.4. Number of Monopodial and Sympodial Branches per Plant 79
4.10.5. Ginning Out Turn Percentage (GOT %) 80
4.10.6. Scanning Electron Microscopic Analysis of Cotton Fiber 82
4.10.7. Photosynthetic Rate 83
4.10.8. Transpiration Rate 83
4.10.9. Gaseous Exchange Rate 83
4.11. Insect Bioassays 85
4.11.1. Laboratory Bioassay with Cotton Bollworm (Helicoverpa armigera) 85
4.11.2. Laboratory Bioassay with Whitefly (Bamisia tabaci) 87
4.12. Determination of Transgene Copy Number and Location 89
5. DISCUSSION 90
6. REFERENCES 97
7. APPENDICES 123
8. LIST OF PUBLICATIONS 133
1. LIST OF FIGURES
Figure 2.1 Mode of Action of Cry toxins (Pardo-Lopez et al. 2013) 21
Figure 3.1: Map of pCAMBIA1301_Vip3Aa+ASAL 44
Figure 3.2: Schematic diagram of Vip3Aa+ASAL cassette 44
Figure 4.1: Confirmation of Vip3Aa+ASAL cassette in pUC57 vector through colony PCR 63
Figure 4.2: Confirmation of Vip3Aa+ASAL cassette in pUC57 vector through restriction digestion 64
Figure 4.3: Digestion of plant expression vector (pCAMBIA 1301) 65
Figure 4.4: Digestion of plant expression vector (pCAMBIA 1301) ligated with Vip3Aa+ASAL cassette 66
Figure 4.5: Confirmation of Vip3A+ASAL cassette in Agrobacterium tumefaciens LBA4404 67
Figure 4.6: Confirmation of Vip3A+ASAL cassette in Agrobacterium through restriction digestion 68
Figure 4.7: A schematic diagram of Agrobacterium mediated transformation 69
Figure 4.8: Confirmation of putative transgenic cotton plants through PCR 70
Figure 4.9: Standard curve of known Vip3Aa standards 71
Figure 4.10: Quantification of Vip3Aa protein through ELISA 72
Figure 4.11: Transient expression of GUS gene 73
Figure 4.12: Dipstick analysis of transgenic cotton plants 73
Figure 4.13: Confirmation of transgenic cotton plants through PCR in T1 progeny 75
Figure 4.14: mRNA expression of Vip3Aa & ASAL genes through qRT-PCR 76
Figure 4.15: Quantification of Vip3Aa protein through ELISA in T1 transgenic lines 77
Figure 4.16: Dipstick analysis of transgenic cotton plants 77
Figure 4.17: Comparison of morphological characteristics between transgenic and non-transgenic lines 81
Figure 4.18: Scanning Electron Microscopic analysis of fiber surface 82
Figure 4.19: Comparison of physiological parameters between transgenic and non-transgenic cotton lines. 84
Figure 4.20: Leaf-detach bioassays of transgenic cotton lines with Cotton Bollworm (Helicoverpa armigera). 86
Figure 4.21: Mean percent mortality of Cotton Bollworm (Helicoverpa armigera) larvae 87
Figure 4.22: Pictures of bioassay performed with Bamisia tabaci 88
Figure 4.23: Mean percent mortality of whitefly (Bamisia tabaci) 88
Figure 4.24: Determination of copy number and transgene location through FISH 89
1
1. INTRODUCTION
Cotton is the foremost cash crop of Pakistan; usually known as “White gold” due to
its importance in the country’s economy (Puspito et al. 2015). Pakistan is the fourth leading
producer of cotton; the third major exporter and consumer and the largest exporter of cotton
yarn all over the world (Imran et al. 2018). Cotton shares 5.2% of value addition in
agriculture, 1% of GDP and adds over US$10 billion to the national economy (Saleem et al.
2018). Cotton production for the financial year 2016/17 was expected to be 10.671 million
bales registering an increase of 7.6% over the last year’s production which was 9.917 million
bales, still significantly behind the expected target of 14.1 million bales. The lower than
expected production is due to the considerable fall of 14.2% in cultivation area which was
2.49 million hectares as compared to 2.9 million hectares last year. Significant losses from
last year’s pink bollworm infestation and low profit margin shifted the farmer’s confidence to
other competitive crops. However, the average yield is increased to 730 kg/Hec, a 25.4%
increase, as compared to 582 kg/Hec last year. Being a developing country, Pakistan needs to
grow more cotton to meet export and domestic requirements (Economic Survey of Pakistan,
2016-17).
The cotton industry of Pakistan is facing several challenges in the global market that
include competition with synthetic fibers, low fiber quality, high production cost and low
yield mainly due to infestation of different insect pests (Bakhsh et al. 2005). The cotton crop
is attacked by more than fifteen insect species which can be divided into two categories:
sucking and chewing insects (Shah et al. 2017). Sucking insects (whitefly, thrip, aphid,
Jassid, and mealybug) damage the cotton crop in two ways. Firstly, these insects suck the
2
essential plant nutrients especially from leaf and develop a fungus named sooty mold on the
surface of the leaf that results in wilting, yellowing, drying of the plant and reduced fiber
quality (Chandrasekhar et al. 2014; Chaudhry et al. 2009). Secondly, these insects also act as
carriers of hazardous plant viruses such as Begomoviruses transmitted by whitefly (B. tabaci)
in a circulative manner (Ahmad et al. 2017). On the other hand, chewing insects like army
bollworm, American bollworm, pink bollworm and spotted bollworm attack directly on
cotton bolls. It is estimated that almost half of the cotton yield is lost due to the infestation of
sucking and chewing insect pests (Saini 2011).
Intensive use of chemical pesticides on cotton and other crop plants to inhibit
insect/pest population is not only harmful to plant and environment but also create an
economic pressure on farmer’s input. These synthetic pesticides adversely affect other non-
target beneficial insects directly or indirectly due to their non-selective properties, moreover,
insects have also developed resistance to these synthetic insecticides (Ali et al. 2016;
Wojciechowska et al. 2016). It has been estimated that the use of insecticides in Pakistan has
increased from 5000 tons to 44,872 tons during the last thirty years (Khan et al. 2015a).
The era of modern genetic engineering has started in the late 1970s, since then, using
sophisticated transformation tools, scientists are able to transform foreign genes from
different species or even from different kingdoms having better insect and herbicide
tolerance, enhanced nutritional qualities, drought and salt stress tolerance into major cash
crops (Chandan et al. 2017). The genetically transformed plants with enhanced
morphological, physiological and nutritional traits hold great promise for efficient use of
shrinking cultivable land that will ultimately help to feed the overspreading world population
(Park et al. 2017).
3
The most successful approach to develop inbuilt resistance against chewing or
lepidopteran insects is to engineer crop plants through the introduction of insecticidal crystal
proteins (ICPs) or simply crystal (Cry) proteins derived from Bacillus thuringiensis, a gram-
positive and spore-producing bacterium (Chandan et al. 2017; Javaid et al. 2016). Transgenic
crops expressing Bt crystal toxins in controlling insect pests have revolutionized global
agriculture since its commercial introduction in 1996 (Karthikeyan et al. 2012). Due to the
continuous plantation and poor insect pest management in the field, over the years, many pest
species have developed resistance against these toxins that may minimize the overall
performance of this technology. Therefore, to expand the insecticidal spectrum of pest
control programmes and to combat resistance build-up, novel insecticidal proteins with
greater toxicity are required (Javaid et al. 2016; Tabashnik et al. 2013).
The vegetative insecticidal protein 3A (Vip3A) from Bacillus thuringiensis (Bt),
secreted during the vegetative growth stage (Estruch et al. 1996), could be a best possible
candidate toxin due to its unique receptor binding sites and greater toxicity (Lee et al. 2006).
The vip3A proteins are toxic to broad spectrum lepidopteran insects, interestingly, to certain
species that have developed resistance against some Cry1A toxins due to the fact that these
proteins share no sequence homology with any known Bt crystal proteins and target different
receptors on midgut membrane of the insects (Chen et al. 2017a). Despite being structurally
different, both protein families exert their toxicity apparently through similar mode of action
i.e. the toxin is activated by midgut protease enzymes and penetrate through the peritrophic
membrane where it binds to specific receptors on the apical membrane and create pores in
the epithelial midgut cell line (Lee et al. 2003). The vip3Aa gene has been successfully
transformed in different crop plants like cotton, corn and cowpea (Bett et al. 2017; Burkness
4
et al. 2010) and in some studies, pyramided with different crystal toxin genes to enhance
broad range pest protection and delay resistance build-up (Chen et al. 2017a).
Sucking insect pests are another devastating group of insects belonging to the order
Homoptera, which cause severe damage to cotton crop by directly sucking the phloem sap
and indirectly by acting as vectors for more than 200 viral, bacterial and fungal diseases
(Mineau 2011). Insecticidal crystal proteins from Bacillus thuringiensis have not been
reported to be able to control this specific group of insects (Vajhala et al. 2013). These
insects have specialized mouth parts and use a unique feeding approach for sucking free
amino acids and sugars directly from phloem tissues (Alliaume et al. 2018). Therefore, a lot
of work has been done to explore possible substitutes of Bt insecticidal toxins to engineer
resistance against sucking insect pest. In the last three decades, many plant-derived lectins
have shown insecticidal activity (Macedo et al. 2015) The expression of these genes under
the influence of phloem-specific promoters could be a possible alternate of chemical
insecticides and Bt crystal proteins against major economically important sucking pests
(Javaid et al. 2016).
Plant lectins are mannose-binding proteins that specifically bind with glycan
structures with great affinity and thus involved in various biological functions (Lannoo and
Van Damme 2010). The entomotoxic activity of plant lectins has been well-reported in the
literature. The possible mechanism of action of plant lectins in target insects initiates with the
binding of lectin molecules with different glycan structures present in brush border
membrane vesicle receptors of gut epithelium cell line, resulting in disruption of cell
structure and function which ultimately leads to cell death (Vasconcelos and Oliveira 2004).
The exact mechanism of toxicity of plant lectins is difficult to predict due to the fact that
5
each lectin protein possesses at least one carbohydrate-binding domain that can bind with a
number of glycoproteins in insects. It is perhaps more difficult to understand why different
plant lectins when exposed to insects show variable insect mortality (Ghosh et al. 2015).
Recently, a 25-kDa mannose-binding homodimeric leaf agglutinin lectin isolated from
Allium sativum (ASAL) has been successfully transformed into different economically
important crops and shown to have entomotoxic effects on major sap-sucking pests of cotton,
mustard and rice (Vajhala et al. 2013; Yarasi et al. 2008).
The emergence of resistance build-up in major chewing insects against Bt insecticidal
Cry proteins and to widen the overall insect spectrum has encouraged the idea of stacking
different genes in the same plant. Three-gene cotton (Puspito et al. 2015); three-gene rice
(Das and Rao 2015) are a few examples of gene stacking. It seems that this “blind” stacking
has no end. Other strategies include fusing different Bt toxins together (Chen et al. 2017a;
Rani et al. 2017) by removing the stop-codon of the first gene and fusing it to the second
gene. The fused protein will translates in a single polypeptide and contains the functional
domains of both the proteins, thus increasing the number of binding sites on midgut receptors
(Ahmad et al. 2015).
In the current study, an effort was made to develop transgenic cotton plants through the
transformation of codon optimized synthetic Bt Vip3Aa gene driven by constitutive
CaMV35S promoter and leaf agglutinin gene from Allium sativum (ASAL) under phloem-
specific RTBV promoter to minimize the losses caused by the infestation of major sucking
and chewing insect pests.
6
2. REVIEW OF LITERATURE
2.1. From Al Qutn to Cotton
The English Word “Cotton” has been modified from an Arabic terminology “Al
Qutn” (Khaleequr et al. 2012); a plant that has been used in making clothes and ropes since
prehistoric times (Grömer 2016). Although the exact origin of cotton cultivation is still
unknown, Archeologists have found some 8000 years old cotton fabric traces while
excavating graves and city ruins in Mohenjo Daro (an ancient town in the Indus valley
civilization) (Bhat 2017). Similar aged examples have also been found from Egypt’s Nile
valley civilization and Tehuacán valley of Mexico (Beckert 2015). The Greek historian
Herodotus, after his visit to India in 5th
century B.C., wrote about a plant: "wild plant that
bears fleece as its fruit" (Box 2000). Arab merchants introduced cotton cloths to the
Europeans about 800 A.D. By the start of 16th
century, cotton was recognized all over the
world (Riello 2013). Due to the industrial revolution in England, cotton lint was first spun by
machines in the 1730s (Nisbet 2009). An American engineer, Eli Whitney, in 1793 filed a
patent on the structure of a cotton gin in America. The design was so effective that it
remained practically un-amended until today. The development of the cotton ginning
machines in the United States has paved the way to supply large quantities of cotton fiber to
meet the demands of flourisihing textile industry (Resis 2006).
2.2. Genome Evolution of Cotton
Cotton is a perennial crop which exists for two successive years; belongs to genus
Gossypium and family Malvaceae (Zhang et al. 2016). Genome-wide analysis of genus
Gossypium revealed that it has been originated from paleohexaploidy of an ancestral eudicot
7
progenitor followed by divergent evolution into eight diploid genomic groups i.e. A-G and K
(Paterson et al. 2012). The A-genome is African endemic and evolved from the eudicot
ancestor along with the Mexican origin D-genome about 5-10 million years ago (MYA) (Li
et al. 2014; Wang et al. 2012). These two genomes came together geologically around 1-2
MYA by the transoceanic dispersion of an A-genome progenitor (Gossypium arboreum AA)
and D-genome progenitor (Gossypium raimondii DD), on the basis of chromosome pairing
(Wendel and Grover 2015). The resultant allotetraploid cotton dissipated from the Americas
towards the western Pacific and divided into 45 diploid (2n=2x=26) and five allotetraploid
(AAxDD; 2n=4x=52) Gossypium species (Tang et al. 2016). Evolutionary studies placed the
diploid species in two lineage groups and the tetraploid species into one lineage (Zhang et al.
2015). Only four species, two allotetraploids (Gossypium hirsutum and Gossypium
barbadense) and two diploids (Gossypium herbaceum and Gossypium arboreum), are
commercially cultivated these days. Gossypium hirsutum produces almost 90% of the
commercial cotton worldwide. Gossypium barbadense (Egyptian cotton) contributes 8%;
Gossypium herbaceum and Gossypium arboreum together add 2% to the world’s cotton
(Chen et al. 2017b).
2.3. World Cotton Outlook
Cotton is considered an essential non-food and leading fiber crop that is also a major
source of foreign exchange earnings for more than 50 countries worldwide (Awan et al.
2015). In 2016/17, the global cotton cultivated area is predicted to reduce by 1% to 30
million hectares, which is the lowest cultivated area since 2009, when the cultivated area was
29.7 million hectares. The overall cotton production is forecasted to rise by 9% to 753 kg per
hectares, and the overall yield is predicted to improve by 7% to 22.6 million tons in 2016/17
8
(Arora et al. 2017). The area under cotton cultivation in India is reduced by 8%, to just less
than 11 million hectares, however, a 9% increase in the average yield to 526 kg per hectares
may likely compensate the losses but overall yield is anticipated to remain constant at 5.8
million tons (Reddy et al. 2017). Cotton production in China is expected to fall by 4% to 4.6
million tons in spite of a 3% increase in the average production to 1600 kg per hectares. The
total cultivated area has shrunken by 7% to 2.8 million hectares (Li 2017). Higher cotton
prices encouraged the farmers of the United States that eventually resulted in a 20% increase
in the cultivated area expected at 3.9 million hectares. Favorable weather conditions have
resulted in 5% increase in cotton production of 899 kg per hectares resulting in a 25% rise in
overall production to 3.5 million tons (Mathur et al. 2017). The total cultivated area under
cotton in Pakistan has fallen by 12% to 2.5 million hectares, but the average production for
the year 2016-17 is expected to increase by 26% to 1.9 million tons as the average yield
increased by 43% to 756 kg per hectares (James 2015; Solangi et al. 2017).
2.4. Role of Cotton in Pakistan Economy
Cotton is a major non-food cash crop and an essential source of raw material to the
textile industry in Pakistan. Moreover, its seeds are also multi-product such as hulls, oil, lint,
and food for animals (Ozyigit et al. 2007). Cotton production supports Pakistan’s largest
textile sector, which consists of around 400 textile mills, 7 million spindles, 27,000 looms in
the mill sector, over 250,000 looms in the non-mill sector, 700 knitwear units, 4,000 garment
units, 650 dyeing and finishing units, some 1,000 ginneries, 300 oil expellers and 15,000 to
20,000 indigenous small-scale oil expellers (kohlus) (Rehman et al. 2016). Cotton contributes
significantly to foreign exchange earnings of Pakistan. Cotton shares 5.2% of the value
addition in the agriculture sector and has a contribution of about 1% in the country’s overall
9
GDP. Cotton production for the financial year 2016/17 was recorded to be 10.671 million
bales registering a 7.6% rise over the last year’s production of 9.917 million bales, still
behind the estimated 14.1 million bales by a significant margin. The lower than expected
production is due to the considerable fall of 14.2% in cultivation area which was 2.5 million
hectares as compared to 2.9 million hectares last year. Significant losses from last year’s pink
bollworm attack and low market prices of fiber shifted the farmer’s confidence to other
competing crops. However, the average yield is increased to 730 kg/ha, a 25.4% increase, as
compared to 582 kg/ha last year (Farooq 2016).
2.5. Problems in Cotton Cultivation
Ancient Chinese and Egyptian literature described plant diseases which were most
likely to be caused by bacteria, fungi, and viruses. Similarly, Aristotle described the
foulbrood infection of Honeybee (Apis millifera) in his writings. With the passage of time,
understanding with these plant diseases has been increased and various control measures are
being used by farmers (Meixner et al. 2014). The cotton crop is also attacked by a number of
insect pests throughout the world and a lot of efforts and money has been spent to resolve
this issue (Pretty and Bharucha 2015). In Pakistan, sucking and chewing insects are the major
threatening pests to cotton. Whiteflies, Aphids, Thrips, Mealybugs, Jassids, and Mites are
categorized as sucking pests while American bollworms, Armyworms, spotted bollworms
and Pink bollworms are the chewing insects (Sarwar and Sattar 2016). It has been estimated
that almost 50-60% yield losses are caused by these two insect groups (Arora et al. 2017).
Sucking insects damage the crop by leaching plant nutrients, specifically from the leaf
portion resulting in deformed leaves and stem that eventually leads to fungal growth
characterized as sooty mold and dried or yellowish plant organs (Luo et al. 2017). Many bug
10
species are responsible for the damage to cotton fiber quality every year (Pulakkatu-Thodi et
al. 2014). Chewing insects, on the other hand, directly attack and destroy cotton bolls and
leaves (Ahmed et al. 2015).
Various chemical insecticides have been used so far to control these pests which
indirectly affect human health and the environment (Oerke 2006). Apart from that, the
emerging insect resistance against these chemical pesticides is also a major threat. It is
reported that over 500 insect species have now become resistant to chemical insecticides
(Beaty et al. 2016). Moreover, a lot of money is spent every year on the production of
insecticides. Economic survey report of the financial year 2013-14 has shown that indigenous
pesticide industry in Pakistan has produced worth USD 227.88 million of pesticides and
annual import value accounted for more than USD 80.28 million (Farooq and Wasti 2015).
2.6. Major Pests of Cotton Crop
Insect attack on crops is not a new topic; it has been a major yield constraint since
ages. Some of the most devastating cotton pests in Pakistan are explained below.
2.6.1. Bollworms or Chewing insects
A number of bollworm species attacks the cotton crop at any growth stage, but a
reproductive stage is mostly affected (Mapuranga et al. 2015). On hatching, larvae bore in all
reproductive organs of the plant, i.e. bolls, flowers, squares and locules with their heads
inserted into the plant organ and remaining bodies protruded outward. Larval feeding
resulted in the excavation of inner tissues and formation of holes in bolls, locules, and
squares. After the destruction of squares, larvae survived by feeding on plant foliage and
eventually resulted in complete damage to the plant (Hu et al. 2018). The attack of bollworm
11
to squares and small bolls may cause pre-mature boll shedding while maturing bolls may
prevent normal growth which may also lead to secondary fungal infections such as boll rot.
Integrated pest management is available for some of these pests like armyworms. Chemical
insecticides are also administrated when other strategies do not work (Luo et al. 2014).
2.6.2. American Bollworm (Helicoverpa armigera)
Helicoverpa armigera species are still the major threat to conventional (non-
genetically modified) cotton. A clear and distinct damage can easily be observed in the attack
of cotton plants with these insects including Helicoverpa armigera, Helicoverpa punctigera
or related species (Shaw 2000). Even a single larva can cause complete destruction of up to
30-40 bolls per plant. H. armigera is found to be highly resistant to conventional insecticides.
It is alarming for the cotton growers of the world. In the temperate region, the attacking
season for H. armigera persists from Mid-March to October (Aggarwal et al. 2006).
The diseased plants are characterized by “damaged terminal buds” before their
opening and “tripped out” seedlings. Granular faecal pellets of insect larva can also be
observed near the boreholes. Affected squares have spread bracts with distinct flaring up
symptoms. Under favorable conditions, adults of Helicoverpa spp. lay a large number of eggs
leading to rapid population build up. However, the severity of insect attack is influenced by
various biotic and abiotic factors such as predators present in the field; wind; rainfall and
extreme weather conditions (Shah et al. 2017).
2.6.3. Pink Bollworm (Pectinophora gossypiella)
Pink Bollworm is considered to be the most noxious pest of the cotton crop in
Pakistan these days (Azfar et al. 2015). It attacks the plant only at the reproductive phase
12
when the female moths deposit eggs at the base of flower (Sridhar et al. 2017). The Larvae
tend to hibernate in seeds during boll formation. The most common indications of the attack
are a failure or abnormal boll opening; rosetted flowers; immature boll senescence and
discolored lint. The larvae usually enter through the tip of 14-28 days old bolls leaving a
yellowish spot on the lint at the entry point. For the first 24 hours, the larvae feed on the
contents of the boll, including the lint near the entry point. After that, It starts feeding on seed
coat and kernel and travel to the second seed. They make a hole in adjoining seeds forming
“Double seeds”. These joint seeds actually act as niches for the pupa (Sarwar 2017).
2.6.4. Spotted Bollworms: (Earias insulana)
Although the severity of spotted bollworm attack is low, still it may cause significant
damage to crop yield in the early season (Hasan 2010). The larvae feed not only on the
reproductive organs but also on the vegetative parts too; just after hatching, the larvae start
chewing tender plant organs (Behera 2015). The most important indications of the attack
include: the premature opening of the bolls; shedding of young bolls and squares; dried or
drooped terminal shoots at the pre-flowering stage; flared bract squares during boll formation
and discolored lint (Kanher et al. 2015).
2.6.5. Armyworm (Spodoptera litura)
Spodoptera litura is amongst the major threats to the cotton crop in the world (Khan
et al. 2015b). The mode of action is quite different as compared to other bollworms. The
affected plant is characterized by damaged leaves with empty areas because the larvae chew
the leaves from veins and veinlets. Leaf skeleton dries up due to incomplete photosynthesis
13
that leads to loss of plant vigor, damaged bolls, and ultimately lower cotton yield (Ahmad
and Gull 2017).
2.7. Sucking or Phytophagous Insects
Insects with specialized “piercing mouth-parts” that specifically feed upon phloem
sap are termed as phytophagous or sap-sucking insects (Bonaventure 2014). They can attack
the plant in different ways (1) sucking the plant fluids including amino acids that are building
blocks of proteins causing loss of plant vigour (2) secreting honeydew that can accumulate
on leave that inhibits the photosynthesis (3) some aphids and whiteflies act as a vector of
plant viruses (Stephens and Westoby 2015). Clear symptoms of the acute infestation include
plant stunting, plant wilting, leaf distortion and leaf yellowing (starting with the lower leaves
results in shedding of bolls due to early defoliation or improper boll opening) (Schoonhoven
et al. 2005). Some of the significant sucking pests are described below.
2.7.1. Whitefly (Bemisia tabaci)
Bemisia tabaci is one of the most notorious pests of cotton. It acts as a vector for
Cotton Leaf Curl Virus (CLCuV), a single-stranded DNA (ssDNA) virus belonging to
Geminivirus family, and causes almost 50% yield losses in Pakistan (Khan et al. 2015c). The
first epidemic of CLCuV in Pakistan was reported in 1992-93 when 7.9 million cotton bales
were lost and the textile industry faced a loss of worth the US$ 5 billion (Mansoor et al.
2003). Whitefly infected plant can be characterized by sooty molds, sticky black coloration
of leaves, reduced photosynthesis and decreased plant strength. Severe infestation results in
premature or poor boll formation (Maharshi et al. 2017).
14
2.7.2. Dusky Cotton Bug (Oxycarenus hyalinipennis)
The dusky cotton bug is considered to be the possible threats to the cotton crop in
Pakistan these days (Ahmed et al. 2015). Both adults and nymphs tend to feed upon sap from
reproductive organs results in poor seed quality. Moreover, Lint quality and quantity is also
affected (Zeilinger et al. 2016).
2.7.3. Mealybugs (Phenococcus gossypiphilous)
The mealybug is a major problem in almost all the cotton producing countries of
Asia, Africa, and the Pacific (Wang et al. 2010). The Mealybug was first observed in the
fields of Pakistan in 2005 when the entire cotton belt was greatly affected (Khan and Mathur
2015). Phenococcus spp. possess a very rapid life cycle. Nymphs readily inhabit the
underneath surface of leaves in the form of the thick mat and develop into adults by feeding
plant sap. Mealybugs excrete copious amount of honeydew on which the sooty mold fungus
grows resulting reduced fruiting capacity, curling, and drying of leaves, ultimately, a drastic
decrease in yield (Noureen et al. 2016).
2.7.4. Leafhopper/ Jassids (Amrasca devastans)
The population of leafhopper often remains under the threshold level in Pakistan and
no significant outbreak has been reported until now (Saeed et al. 2017). Cotton Jassid attacks
the plant at the six-leaf stage. Rainy and humid weather during the monsoon season is the
most favorable time for its attack characterized by the development of cup-shaped leaves.
The pest not only sucks the phloem sap but also releases toxic saliva to the vascular system
that affects the metabolic pathways of the host plant. The affected plant is distinguished by
yellow tender leaves, curled leaf margins, redness, and dryness. Red colored leaves are called
15
as “hopper burns” and considered to be the distinctive symptom of severe Jassid infestation
(Saha and Mukhopadhyay 2013).
2.7.5. Thrips (Thrips tabaci)
The extent and intensity of thrips attack may differ according to the season and
geographic locations (Steenbergen et al. 2018). Thrips attack the plant at the seedling stage
where it scrapes off foliar parts with their piercing mouthparts and feeds on the plant
nutrients (Steenbergen et al. 2018). The symptoms of attack include brownish/silvery or
deformed leaves. Minor attack of thrips tends to retard plant growth and delay maturity;
however, severe attack may kill the entire plant. Once cotton plants get 30-40 days old, they
can easily outgrow from thrips damage and recover by their own self-defense mechanism
(Miyazaki et al. 2017).
2.8. Conventional Breeding for Cotton Improvement
The conventional breeding efforts for the improvement of cotton were dated back to
the early 19th
century at the individual farmer level using visual selection procedures
(Bowman et al. 2004). The cultivation of tetraploid cotton was introduced in the late 14th
century when the Egyptian breeders developed many elite cultivars of G. barbadense and G.
hirsutum through selection and domestication (Abdalla et al. 2001). The Egyptian cotton is
well known for its long, strong and fine cotton fibers, whereas Upland cotton has the
properties of early maturation, high yield, and better stress tolerance (Zhang and Percy 2007).
Several studies have reported the use of different hybridization and backcrossing techniques
to develop elite cotton cultivars that could combine alleles of favorable traits from both the
species (Tang et al. 2015). The genetic breakdown in later generations of interspecific
16
hybrids was found showing a negative relationship between fiber yield and length (Zhang et
al. 2014). All the breeding efforts for the crop improvement are generally restricted to intra
and interspecific hybridization but the prolonged time period is required to achieve the
desired results (Marasek-Ciolakowska et al. 2016). With the advent of the latest
biotechnological approaches, the development of a first transgenic plant in the 1990s has
eliminated these genetic barriers even at kingdom level (Gosal et al. 2010).
2.9. Discovery of Bacillus thuringiensis (Bt) and Its Toxic Crystal Proteins
The era of Bacillus thuringiensis (Bt) has started in 1901 when Shigetane Ishiwata, a
Japanese microbiologist, identified a rod-shaped bacterium from dead silkworm larvae while
investigating the possible cause of “sotto disease” of silkworm larvae and coined the term
Bacillus sotto for that bacterium (Ibrahim et al. 2010). Later in 1911, Ernst Berliner, a
German Microbiologist isolated similar rods from dead Mediterranean flour moth larvae
while inspecting a flour mill in Thuringia. Berliner named these bacteria as Bacillus
thuringiensis (Bt) in 1915 (Lone et al. 2014). Further, Berliner also claimed the presence of
small inclusion bodies near endospores which were confirmed by Mattes in 1927. However,
it took almost three decades to postulate the insecticidal function to these “parasporal
crystals” a term coined by Christopher Hannay and his colleague Philip Fitz-James who also
revealed the protein nature of these toxic parasporal crystals (Upadhyay 2003). Later in 1956,
the work of Angus discovered that the crystalline inclusion bodies were formed during
sporulation and responsible for the insecticidal activity of these toxins (Bt). Various
experimental studies were performed thereafter to verify the pesticidal role of Bt protein
crystals (Sansinenea 2012). In 1982 Gonzalez and his co-workers reported that the genes
encoding crystal proteins were localized on transmissible plasmids using plasmid curing
17
technique (Abdoarrahem et al. 2009). Schnepf and Whiteley successfully characterized and
isolated the crystal genes from Bt subsp. Kurstaki HD-1 and cloned into E. coli to assess their
insecticidal role against Tobacco hornworm (Melo et al. 2016; Schnepf et al. 1998). In the
1980s, several research groups reported that plants can be genetically engineered through
different transformation tools, and finally first genetically modified (Bt cotton) got access to
the market in 1996 (Roh et al. 2007).
2.10. Insecticidal Toxins of Bacillus thuringiensis
A large number of Bt isolates secrete a range of insecticidal proteins that are toxic to
different insect orders, and in some cases, species from other phyla (Sheikh et al. 2017).
These proteins can be categorized as Crystal (Cry) and Cytolytic (Cyt) toxins (also known as
δ-endotoxins), that are secreted as parasporal crystals during sporulation phase. Additionally,
different Bt strains can also produce vegetative insecticidal proteins (VIPs) and the secreted
insecticidal protein (SIPs) into the culture medium during the vegetative growth phase
(Palma et al. 2014). Vegetative insecticidal proteins are further classified into four distinct
families i.e. Vip1, Vip2, Vip3, and Vip4, based on their sequence homology. The Vip1 and
Vip2 belong to binary toxin group (i.e. both toxins are required for insecticidal activity); SIPs
are active against some coleopteran insects (Donovan et al. 2006) while Vip3 proteins are
toxic to lepidopteran insects (Chakroun et al. 2016). The host spectrum of the Vip4Aa1 toxin
remains unknown to date (Palma et al. 2014). Recently, some parasporal crystal proteins with
unknown target, termed as parasporins, have been reported and exhibit strong cytocidal
activity against human cancer cells (Ohba et al. 2009).
18
2.11. Crystal (Cry) Proteins or δ-endotoxins
Crystal proteins commonly called δ-endotoxins (Cry and Cyt toxins) are released in
the form of parasporal crystals during the sporulation phase of growth cycle in Bacillus spp
(Federici et al. 2006; Höfte and Whiteley 1989). To date, the nomenclature committee for Bt
toxins has identified 75 families of Crystal proteins i.e. Cry1-Cry75 (Crickmore et al. 2017).
These Cry toxins are active against different insect orders like hemipterans; nematodes;
lepidopterans; coleopterans; dipterans; few snails and even some tumor cells of the human
body (de Maagd et al. 2003; Ohba et al. 2009; Van Frankenhuyzen 2009). Most of the crystal
proteins consist of the well-studied three-domain structure while others belong to different
protein groups e.g. Binary Bin, ETX_MTX2 like protein families (de Maagd et al. 2003).
The N terminal domain I is made up of a bundle of seven α-helices and wrapped around the
central conserved amphipathic α-helix (Deist et al. 2014). Structurally, the domain I is
responsible for the proteolytic activity when stimulated, and considered to be involved in
host membrane insertion and pore formation (Xu et al. 2014). Domain II, on the other hand,
has a central hydrophobic core around which three pairs of antiparallel β-sheets are wrapped
tightly, giving a prism-like structure. Functionally Domain II is involved in most important
toxin receptor binding with host cells (Jenkins and Dean 2000). Domain III is also called as
galactose-binding domain, making a sandwich of two antiparallel β-sheets. Domain III assists
in receptor binding as well as pore formation (Rausell et al. 2004; Xu et al. 2014). Multiple
sequence alignment searches of different cry proteins showed five conserved regions in the
external toxic core of protoxin. Additionally, three conserved amino acid blocks are also
present at the C terminal outside the active core, toward the C-terminal end. Some cry
19
proteins like Cry3 and Cry11 lack this additional C-terminal domain and form a relatively
shorter protoxin (70 kDa) (Jurat-Fuentes and Crickmore 2017).
2.12. Cytolytic (Cyt) Proteins
The Cytolytic toxins are a relatively smaller group of insecticidal proteins with a
molecular weight of around 25 kDa, mostly active against dipteran insects, but to some
extent, lepidopterans, coleopterans, some nematodes and cancer cells (Bravo et al. 2007).
These cytolytic proteins have been grouped into three families (Cyt1, Cyt2, and Cyt3) by the
Bacillus thuringiensis toxin nomenclature committee (Crickmore et al. 2015). The 3-
dimensional structure has shown that the activated toxin consists of a single domain with
three layers of β-strands placed in the central region and α1 and α2 helices flanking on one
side and α3 - α6 helices on the other (Xu et al. 2014). It is believed that α helix-rich N-
terminal is involved in oligomerization and the three β-strands in the C-terminal domain are
responsible for pore formation (Rodriguez-Almazan et al. 2010). Two different models have
been proposed for the mode of action of Cytolytic proteins. “Pore Model” states that the Cyt
toxins form cation-selective beta-barrel pores with the β5, β6, and β7 sheets (van der Hoeven
2014). According to the “Detergent Model,” Cyt toxin aggregates on the surface of the cell
membrane and disrupts the membrane lipid bilayer structure (Butko 2003).
2.13. Mode of Action of Cry Toxins
Three different models for insecticidal activity of three-dimensional crystal proteins
have been proposed.
20
2.13.1. Classical Model for Action of Cry Toxin
The most considered “Classical model” of Bt mode of action is further elaborated by
four simple steps. In the first step, the protoxin is synthesized, ingested and dissolved in the
alkaline midgut. In the next step, the protoxin is activated by host midgut proteases i.e.
appropriate cleavage of protoxin from N and/or C terminal to form a protease-resistant active
toxin. In the third step, the active toxin binds to the surface receptors of the midgut
epithelium followed by formation of the pore to penetrate into the cell membrane.
Eventually, the pores become more extended and permeable to inorganic molecules, amino
acids and sugars, which result in lysis of epithelial cell line and ultimately death of host
insect. Although the classical model has been accepted for a long time, this model still does
not elaborate pore structure and mechanism of pore assembly in detail and the more critical
investigation is required (Pardo-Lopez et al. 2013; van der Hoeven 2014).
2.13.2. Sequential Binding Model
The sequential binding model of Cry toxin activity states the binding of the activated
toxin to cadherin-receptors (alkaline phosphatase or Aminopeptidase-N) which are already
present on the cell membrane of the host cell. Following this binding and conformational
modification, the α1 helix of Domain I is sliced off the toxin in the presence of proteases and
led to the formation of a “pre-pore” like structure. The pre-pore oligomer further binds with
the second surface receptor and eventually insert into the epithelial membrane of the midgut
to cause the insect death (Bravo et al. 2007; Bravo et al. 2015).
21
Figure 2.1 Mode of Action of Cry toxins (Pardo-Lopez et al. 2013)
2.13.3. Signaling-pathway Model
This model proposes that cry toxin, after binding to midgut surface receptor, convey
signals to its associated G-protein from Cadherin receptor to activate membrane-bound
Protein Kinase A (PKA) and Adenylyl Cyclase (AC) in presence of Mg2+
ions that result in
membrane swelling and cell necrosis (Zhang et al. 2006).
2.14. Vegetative Insecticidal Proteins or VIPs
Some Bt strains also secrete additional insecticidal toxins into the medium during
vegetative or log phase called as vegetative (VIPs) and secreted (SIPs) insecticidal proteins
which are toxic against Lepidopterans and Coleopterans respectively, extending the overall
spectrum of this bacterium (Chakroun et al. 2016). The nomenclature committee for Bt toxin
has grouped VIPs into four families based on their amino acid sequence similarities i.e. Vip1,
Vip2, Vip3, and Vip4. Unlike Cry proteins, VIPs do not form a crystal and do not share any
sequence homology with any known Cry protein (Crickmore et al. 2015). First two groups,
22
Vip1 and Vip2 act synergistically to perform the toxic activity (Ribeiro et al. 2017). The host
range of Vip4 proteins still needs to be investigated (Crickmore et al. 2015). On the other
hand, Sip toxins possess pesticidal action against coleopterans (Donovan et al. 2006).
Various parasporal crystal proteins are produced by some Bt strains which have non-specific
targets and therefore characterized as parasporins (Ohba et al. 2009).
2.14.1. The Vip1/Vip2 (Binary) Toxins
The Vip1 and Vip2 are members of the binary toxin group of A+B type, i.e. both the
components are required for insecticidal activity, co-translated from a single operon of about
4 kb and encode ~100 and ~50 kDa proteins respectively (de Maagd et al. 2003). In the
proposed mechanism of action of Vip1/Vip2 complex, Vip1 acts as a receptor-binding
domain while Vip2 acts as the cytotoxic domain (Barth et al. 2004), which causes ADP-
ribosylation in cytoskeletal actin protein, blocking its polymerization. Ultimately,
cytoskeletal disarrangement may lead to insect death (Shi et al. 2007).
2.14.2. Vip3 Toxins
The 88.5 kDa Vip3 insecticidal toxin was first isolated in 1996 from the culture
medium during the vegetative growth phase of a Bt strain (Estruch et al. 1996). It is a well-
known fact that Vip3 shares no sequence similarity with any other Cry protein and targets
different receptors on the midgut epithelial cell line of target insects and has a broad
lepidopteran spectrum of activity (de Maagd et al. 2003). Recently a nomenclature system
has also been adopted for Vip toxins, like Cry and Cyt proteins. The nomenclature committee
has grouped Vip3 proteins into three subfamilies i.e. Vip3A, Vip3B, and Vip3C. To date,
there are 64 vip3Aa, 2 vip3Ab, 1 vip3Ac, 6 vip3Ad, 1 vip3Ae, 4 vip3Af, 15 vip3Ag, 2
23
vip3Ah, 1 vip3Ai, 2 Vip3Aj, 2 vip3Ba, 3 vip3Bb, and 4 vip3Ca genes reported (Crickmore et
al. 2015). Most of the research has been done on the more abundant Vip3Aa type proteins;
therefore, very little structural and functional information is available on less common
Vip3B, Vip3C proteins (Chakroun et al. 2016).
2.14.3. Structure and Function of Vip3 Proteins
The three-dimensional structure of the Vip3 protein has not been reported to date
(Palma et al. 2017). Only a partial tertiary structure corresponding to the last 200 amino acids
has been modeled by homology studies (Gayen et al. 2012). In Silico structure, the prediction
has suggested that the N-terminal region consists of α helices, whereas, the C-terminal region
contains β-helices and coils (Rang et al. 2005). Electron microscopic studies with Vip3Ag4
strongly proposed that the protein may be grouped together forming a tetrameric structure
(Palma et al. 2017). A highly conserved signal peptide sequence is present in the N-terminal
region of the mature peptide that is not processed during secretion and responsible for
translocation of the toxin into the periplasmic space across the cell membrane. The role of
this N-terminal region in protein folding and binding to membrane receptors is well
documented in the literature (Chen et al. 2003). It has been proposed that specific target
spectrum of Vip3 proteins is due to its high variability in the C terminal region, any mutation
to the last few amino acids of this region may completely disturb protein function (Chi et al.
2017).
2.14.4. Mode of Action of Vip3 Proteins
The mode of Vip3 action is poorly understood due to the unavailability of its 3-D
structure (Bel et al. 2017). It is assumed that the receptor binding and ion channel properties
24
of Vip3 are different as proposed for the Cry proteins, however, the sequence of events looks
presumably the same i.e. activation of toxin by midgut protease; crossing the peritrophic
membrane; the binding of the toxin to specific receptors on the surface of epithelial midgut
cell membrane and finally pore formation occur leading to insect death (Bravo et al. 2017).
The analysis of the gut cross sections of susceptible insects showed lethal damage in the
midgut area after ingestion of Vip3Aa protein with disrupted, swollen, and lysed epithelial
cells results in leakage of cellular material to the lumen (Hernández-Martínez et al. 2017).
2.14.5. Proteolytic Processing of Vip3 Toxin
In vitro proteolysis of the full-length Vip3A protein with insect midgut extract or
trypsin showed two major proteolytic sites of lysine-rich residues and four major proteolytic
polypeptide products of approximately 66, 45, 33 and 22 kDa (Abdelkefi-Mesrati et al. 2011;
Hamadou-Charfi et al. 2013). The first site is located at lysine 198 and thought to release a
22 kDa N-terminal fragment (from amino acids 1-198) of the protoxin from the 66 kDa
fragment (from amino acid 199 to the end) which is known as highly conserved activated or
trypsin-resistant core that retains toxicity. The second site is located at lysine 455 that
separates two fragments; a 33 kDa fragment (from amino acids 200 to 455) and a 45 kDa
fragment (from amino acids 456 to the end) derived from the 66 kDa portion (Estruch and Yu
2001). The activated 66 kDa fragment when expressed in E. coli and used for bioassay
experiments, no toxicity was observed against insects that were previously susceptible,
suggesting that 22 kDa fragment is necessary for proper protein folding and functioning
(Chakroun and Ferré 2014).
25
2.14.6. Binding of Vip3A to the Midgut Epithelial Cells
The Vip3Af and Vip3Aa labeled with biotin when locally injected to susceptible
insects showed specific binding with microvilli present on their midgut epithelium
(Chakroun and Ferré 2014). Brush border membrane vesicles (BBMVs) of some non-
susceptible insects are also considered to be the target sites for Vip3Aa. It means that Vip3Aa
binding to receptors is not always required for its toxicity in insects (Lee et al. 2006).
Binding of 125
I-labelled Vip3Aa to BBMVs of Spodoptera frugiperda showed a lower
affinity, but the higher number of binding sites as compared to Cry toxins. Binding
competition assays revealed that both trypsins activated 125
I-Vip3Aa fragments, i.e. 20kDa
and 62kDa fragments; compete with each other for the binding to BBMVs. Moreover, this
study also showed that any shared binding sites between Vip3 and Cry toxins, which were
previously confirmed in several insect species, are also absent (Chakroun and Ferré 2014).
Ligand blot analyses showed that specific molecules are associated with the binding sites of
BBMVs for the interaction of Vip3Aa, which were not recognized during Cry1A binding. In
Manduca Sexta, Vip3A recognized and bound to specific proteins of 80 and 110 kDa while
Cry1Ab undergo binding with 210 and 120 kDa protein molecules (Liu et al. 2011).
Only two binding molecules have been identified so far that bind with Vip3A protein
in insect midgut using “yeast two-hybrid system” i.e. 48 kDa from Agrotis ipsilon and S2
ribosomal protein from Spodoptera litura (Estruch and Yu 2001). Silencing of this S2
encoding gene has resulted in reduced toxicity both in fifth-instar S. litura larvae and the
Sf21 cell line, but the exact mechanism of cell lysis due to Vip3A-S2 interaction is still not
clear (Singh et al. 2010).
26
2.15. Plant Lectins: Natural Plant Defense Proteins
Agri-biotechnology has been revolutionized by the transformation of Bt insecticidal
genes into plants. Economic benefits like increased crop production and less usage of
pesticides are worth mentioning (Gupta et al. 2017). Transgenic crops are also environment-
friendly and no experimental study has reported a lethal effect of Bt toxin on non-target
organisms (Rao et al. 2017). However, the host range of Bt toxins is restricted to specific
insect groups and do not work significantly for sap-sucking pests like mirids, grasshoppers,
thrips, aphids, and bugs etc. Therefore, researchers are focusing on the possible alternatives
to control these insects (Gatehouse 2008). From the last two decades, many of the naturally
occurring plant lectins have been found to possess entomotoxic traits and could be the
potential candidate for its use in transgenic plants (Yarasi et al. 2008). Lectins were first
described by Peter Herman Stillmark in 1888 as a toxic element isolated from castor beans
(Ricinus communis L.) that can coagulate red blood cells in the animals (Stillmark 1992).
Concavalin A (ConA) from Jack bean (Canavalia ensiformis) seeds was the first lectin
protein that was crystallized through advanced protein isolation techniques (Grossi-de-Sá et
al. 2015). The plant lectins can be defined as the presence of at-least one non-catalytic
domain which can bind reversibly to a definite mono- or oligosaccharide. Some additional
domains are also present in lectins other than sugar-binding domains with different biological
activities (Van Damme 2008).
2.15.1. Classification of Plant Lectins
Different classification methods have been used to organize plant lectins. Initially,
lectins were classified on the basis of their carbohydrate binding specificity. However, such a
classification system was not relevant to the evolutionary relationships (Itakura et al. 2017).
27
Plant lectins can also be classified on the basis of overall domain architecture i.e. hololectins;
merolectins; superlectins and chimerolectins (Macedo et al. 2015). Merolectins contain only
one carbohydrate binding domain that lacks agglutination activity and considered to be the
simplest class of the lectins. Hololectins, on the other hand, possess more than two similar
carbohydrate binding domains which help them in cell agglutination. Many of the newly
identified plant lectins are placed in this group. Superlectins consist of two or more
carbohydrate binding domains that bind with structurally unrelated carbohydrate structures.
All the plant lectins comprised of one or more biologically active domains other than
carbohydrate binding domains are known as chimerolectins which is the most abundant
group in nature (Macedo et al. 2015). More recently, a novel classification system was
proposed on the basis of whole genome/transcriptomic data. According to this system, plant
lectins are grouped into twelve evolutionary diverse but structurally related lectin domains,
having distinct folding and binding pattern with one or more carbohydrate binding sites (Van
Holle et al. 2017).
2.15.2. Three-Dimensional Structure of Plant Lectins
The three-dimensional structure of lectins is mainly consists of interconnected
antiparallel β-sheets with some α-helices. The dimers and the tetramers are highly stable due
to the hydrophobic interactions and the hydrogen bonds. The central carbohydrate-binding
region contains amino acids residues that interact with metallic ions (Mn2+
and Ca2+
) and
provides necessary binding energy to specifically interact with carbohydrates (Lagarda-Diaz
et al. 2017). Some aromatic amino acids occupy variable positions in a central horseshoe
shape region which is involved in monosaccharide specificity and interaction of the lectin
molecule with larger oligosaccharide ligands (Benevides et al. 2012).
28
2.15.3. Mode of Action of Plant Lectins
The midgut epithelium cell line of many insects is covered with a physical barrier
called the peritrophic membrane (PM) consists of a grid-like network of chitins held together
by chitin-binding proteins such as neurotrophins that contain many glycan structures in the
interstitial spaces creating a molecular sieve (Hegedus et al. 2009). Since the peritrophic
membrane (PM) contains the chitin fibrils and many glycoproteins, this midgut structure is a
possible target for lectins. The effects of plant lectins have been extensively studied on the
structural arrangement of the insect midgut. Clear anomalies were observed in peritrophic
membrane through transmission electron micrographs from midgut of cotton leaf-worm
larvae fed on a Gleheda-containing diet (Li-Byarlay et al. 2016; Vandenborre et al. 2011).
Similarly, the midgut structures of European corn borer (O. nubilalis) fed on a WGA-mixed
diet showed hypersecretion of irregular PM layers into the lumen and disruption of microvilli
structures (Hopkins and Harper 2001).
2.16. Risk Assessment of Plant Lectins in Non-Target Organisms
Plant lectins are playing a significant role in the implementation of pest management
strategies for crops and other plants but the ectopic expression of lectin-related genes in
transgenic plants enlightened the issues of biosafety and global risk mitigation. Undesirable
side effects i.e. harmful effects on beneficial insects could disturb the natural ecosystem and
food web. Lectin expression in GM plant can impart either direct or indirect effect on non-
target insects when they consume the plant parts e.g. seed, leaf, stem, and root (Grossi-de-Sá
et al. 2015). Honeybees and butterflies are among the beneficial phytophilous insects which
are considered to have a very direct effect of such lectin expression while sucking sap and
nectar from transgenic plants (Ricroch et al. 2017). Likewise, indirect effects include the
29
predation or consumption of target insects e.g. aphids by beneficial insects e.g. lacewings,
wasps, and beetles. Both of the scenarios depict a doubtful picture of lectin expression in
transgenic plants and therefore critical risk assessment studies are needed before the proper
commercialization of any GM event. Several experimental studies have been conducted in
the last few decades. For this purpose, feeding trials of beneficial insects on lectin expressing
plant parts are done and physiological parameters of experimental insects are then assessed
(Domingo 2007).
2.16.1. Risk Assessment in Honeybee (Apis mellifera)
The honeybee is the most significant phytophilous insects of the ecosystem as it
carries out 90% of the total pollination of plants. In order to assess the potentially harmful
effects of PSA lectin on honey bee population, transgenic pollens from oilseed rape (Brassica
napus) with PSA expression were mixed into the insect diet and a feeding trial was done with
larvae for continuous seven days. No differences in survival, body mass; growth rate and
differentiation were observed when compared with those of control larvae which were fed
with non-transgenic pollens (Lehrman et al. 2007).
In contrarily, when GNA based feeding trial was done with bumble bees in micro-
colonies, strong negative effects were observed on survival rate and offspring production in
both drones and worker bees (Babendreier et al. 2008). Moreover, the probable effects of
GNA were also assessed on solitary bee (Osmia bicornis) larvae by giving them minimum
concentrations of GNA mixed larval diet (0.01% of total soluble protein) and no effect was
reported on larval development and differentiation. On the other hand, a high concentration
of GNA mixed larval diet when given to the same group (0.1% of total soluble protein);
lengthy differentiation phase and reduced body masses of larvae were observed. However, it
30
has been declared that bees are unlikely to be exposed with GNA doses of ⩾0.1% when fed
in fields naturally. Therefore, the risk of their exposure to high GNA concentration is
eliminated (Konrad et al. 2008).
2.16.2. Risk Assessment in Parasitoids
Predators are a very important part of the food chain in the ecosystem that plays a
significant role in the natural control of various pests and disease-causing insects (Birch et al.
1999; Gill and Garg 2014) performed an indirect feeding assay where common two-spot lady
beetle (Adalia bipunctata) population was allowed to feed on green peach aphid (Myzus
persicae) larvae which were reared on GNA-expressing transgenic potatoes and reported a
decrease in fertility, egg production, and female survival rate. However, later studies did not
support these findings probably due to a combination of direct and indirect feeding trials
(Down et al. 2003).
In two different studies, (Hogervorst et al. 2006) and (Li and Romeis 2009), studied
direct effect of GNA lectins on aphid larvae such as green lacewings (Chrysoperla carnea),
lady beetle (Adalia bipunctata) and seven-spot ladybird (Coccinella septempunctata). A
continuous feeding trial was performed with an artificial diet containing 1% of the GNA.
Results showed that no negative impact on body mass was detected but the survival rate was
affected badly for all the three predators.
The endophagous insects are another important group of parasitoids that lay eggs
within the body of the host insect. Developing larvae uses the same host as a food source on
a later stage (Zhu et al. 2016). The activity of parasitic wasp (Eulophus pennicornis) was
studied under artificial field settings with tomato moth (L. oleracea) larvae which were
reared on GNA expressing tomato plants. The parasitizing capability of the wasp on pest
31
larvae of L. oleracea was observed to be unchanged by the addition of GNA to the host diet.
Similarly, no alteration in fertility was reported when compared with that of the control
group (Bell et al. 2001).
Various parasitoids consume honeydew content as primary carbohydrate source
which is secreted by phloem-sucking insects. Such honeydew concentration could be an
indirect source of plant-based lectins to the beneficial insects. Therefore, the toxicity of
GNA-linked lectins was assessed for various parasitic wasps i.e. Trichogramma brassicae,
Aphidius colemani and Cotesia glomerata, where all of the three insect groups were allowed
to feed on sucrose containing GNA. A distinct reduction in survival rates of three populations
was reported (Romeis et al. 2003). However, their GNA concentration (0.1% or 1%) is again
a topic of dispute in this study. Moreover, white backed plant hopper (Sogatella furcifera)
when fed with honeydew drops from GM rice expressing GNA (0.3% of total soluble
protein) found to be negatively affected by lectins but surprisingly no definite concentration
of GNA was observed in honeydew with which feeding trial was done (Nagadhara et al.
2004).
2.16.3. Evaluation of Lectin Based Risks on Vertebrates
Humans and animals widely consume the plant lectins through vegetables and fruits
e.g. Tomato, Leek, Lentil, garlic, peanut, corn, pea, wheat, bean, soybean, mulberry, banana,
breadfruits etc. Since most of them are considered to be non-toxic to humans on their
consumption without cooking (Muramoto 2017). However, a few legume lectins e.g. ConA
and PHA are considered toxic for mammals. The raw or improperly cooked kidney beans
contain PHA and therefore reported harmful to humans. The PHA poisoning is characterized
by vomiting, nausea, and diarrhea. Basically, PHA toxin possesses binding domains for gut
32
epithelial cells and eventually causes changes in cellular morphology as well as metabolism
(Wiederschain 2009).
Several lectins have the ability to resist gastric juice digestion and rapidly undergo an
interaction with surface glycoproteins of the digestive tract. Local and systemic reactions
occur as a result of this binding (Vasconcelos and Oliveira 2004). Other well-known sources
of toxic lectins are castor beans (R. communis) with lectin “ricin” and Jequirity bean (Abrus
precatorius) for “abrin”. Both toxins are potentially lethal for mammals (Dickers et al. 2003).
Contrarily, Ricin-B type lectin from elderberry (Sambucus sp.) is not found to be as harmful
as ricin and abrin. Although considerable toxicity has been shown by various bean types,
some of these have been exploited for medicinal applications i.e. anticancer drugs (Girbes et
al. 2003).
A feeding trial of GM rice with PHA-E expression was done with rats to evaluate the
effect of lectins. The clear anomalies were reported in these experimental rats after a
continuous feeding for 90-days (Poulsen et al. 2007b). Contrarily, same experimental
approach when done with GNA-based lectins in the diet of rats, no adverse effects were
found. It clearly depicts the safety of GNA-based lectins in vertebrates (Poulsen et al. 2007a).
However, it is very critical to perform long-term toxicity assays on different non-target
organisms for biosafety prediction of GM plants with lectin expressions.
2.17. Agrobacterium tumefaciens: A Machinery of Genetic Transformation
Agrobacterium is a gram-negative pathogenic bacterium present in soil and causes
crown gall disease (formation of tumors) at injured sites in some monocotyledonous and
many dicotyledonous plants (Pitzschke and Hirt 2010). Crown gall disease was reported in
33
1907 for the first time (Smith and Townsend 1907) and later, it was concluded that A.
tumefaciens is proficient to transfer a precise DNA segment (Transfer DNA, T-DNA) into
the infected host cells through tumor-inducing (Ti) plasmid. Both the T-DNA and Ti plasmid
then subsequently get integrated into the host genome and resulted in crown gall formation
(Horsch et al. 1985). Tumor induction is initiated when auxins and cytokinins are synthesized
at an infection site due to the rapid expression of oncogenes in T-DNA (Zupan and
Zambryski 1995). Therefore, Agrobacterium has been vastly used in biotechnology to insert
foreign genes into the plants via tumor formation (Gelvin 2003). Cry genes isolated from Bt
have been incorporated into the cotton plant for insect resistance via Agrobacterium-
mediated transformation (Firoozabady et al. 1987; Umbeck et al. 1987).
2.17.1. What is T-DNA and How It Is Transformed from Agrobacterium to Plant Cells?
All virulent strains of Agrobacterium contain a large megaplasmid which is usually
200 to 800 kb in size. One or multiple fragments of (10-30 kb) of transferred DNA (T-DNA)
or virulence region (vir genes) also known as tumor-inducing plasmid; are located on this
megaplasmid and plays an important role in tumor induction (Zupan et al. 1996). The export
of T-DNA from Ti plasmid to the host genome depends on the activity of these virulence
(vir) genes (Garfinkel and Nester 1980; Horsch et al. 1986; Lundquist et al. 1984). The
VirD1/VirD2 are highly specific endonucleases that cleave 25 bp homologous flanking
border sequences (T-regions) making it single-stranded (Peralta and Ream 1985; Wang et al.
1984; Yadav et al. 1982). In the presence of appropriate phenolic and sugar molecules, the
VirA protein acts as a periplasmic antenna and transphosphorylates the VirG protein with the
help of a monosaccharide transporter ChvE protein (Doty et al. 1996; Jin et al. 1990) to
interact and activate Vir-box sequences more efficiently (Stachel and Zambryski 1986).
34
The type IV secretion system composed of eleven VirB proteins along with the VirD4
protein is necessary for the transfer of the T-DNA and other Vir proteins, including VirE2
and VirF (Vergunst et al. 2000). The VirD4 acts as a “linker protein” to promote the
interaction of the T-DNA/VirD2 complex with the VirB-encoded secretion apparatus
(Christie 1997). Most VirB proteins like VirB2; VirB5 and possibly VirB7 form the
membrane channel for T-DNA and Vir protein transfer or serve as ATPases to provide
energy for the channel assembly or export processes (Eisenbrandt et al. 1999; Jones et al.
1996; Lai and Kado 1998).
The VirD2 and VirE2 proteins bind with the T-DNA region forming a “T-complex”,
and this binding is believed to be the most essential step in Agrobacterium mediated
transformation. The VirE2 possibly forms a pore in the host cytoplasmic membrane to
facilitate the passage of the T-strand (Howard and Citovsky 1990). The VirD2 protein binds
with 5’ end of the T-strand with its nuclear localization signal (NLS) sequences that guide it
through to the plant nucleus (Citovsky et al. 1994; Herrera-Estrella et al. 1990; Howard et al.
1992). However, recent literature suggests that for the transport of larger linked T-strands to
the nucleus, the VirD2 may not be sufficient (Ziemienowicz et al. 2001), for that, VirE2 had
to be accompanied with the T-strands. Eventually, VirD2 plays its role in the successful
incorporation of Transfer-DNA into the plant genome (Tinland et al. 1995). The VirE2 can
alter the conformation of ssDNA and prevents degradation of T-strand that occurs perhaps in
the cytoplasm or in nucleus (Rossi et al. 1993). Recently, many other bacterial chromosomal
genes are also found to play an essential role in the transformation process. These genes are
involved in production of exopolysaccharides (Cangelosi et al. 1987; Cangelosi et al. 2007;
Thomashow et al. 1987); attachment of Agrobacterium to the plant cells (Matthysse et al.
35
1996); regulation and induction of Vir genes (Kemner et al. 1997; Shimoda et al. 1993; Liu et
al. 1999) and transport of T-DNA fragment into plant cell (Gede Putu Wirawan and Kojima
1996; Wirawan and Kojima 1996).
2.18. Development of Bt-transformed Plants
Sporine was the first commercialized Bt-based bio-insecticide, containing a mixture
of spores and insecticidal crystals, developed in France in 1938, to control the flour moths
(Sansinenea 2012). The United States of America introduced and commercialized about 182
different Bt-based bio-insecticides from 1958-1995 which were registered by the US
Environmental Protection Agency (Carpenter and Gianessi 2001). Insect-resistant transgenic
plants have been developed globally since 1996 to improve the plant defense mechanism and
replace the chemical sprays or insecticide uses (Hartman et al. 2016). Recent figures revealed
that over 70% of total Bt-corn and over 90% of total Bt-cotton grown in the US is genetically
modified to produce Bt toxin proteins (Jin et al. 2015).
The interaction of plants with insects and microbes is very complex and considered to
be very important for sustained ecosystems (Stirling 2017). Although, plants have natural
defense mechanisms to defend the harmful effects of disease-causing microbes, arthropods,
and herbivores; sometimes their immune mechanism fails to resist the immense attack of
external agencies. Naturally, Plants release secondary metabolites or certain chemicals to
protect themselves from pests and other stress factors (Mithöfer and Maffei 2017). Similarly,
conventional breeding and in-vitro strategies were also being used to strengthen plant defense
mechanism i.e. interspecific breeding and electrofusion of protoplasts respectively.
With the advent of genetic engineering tools, transgenic plants that express
insecticidal proteins are developed which are more reliable then conventionally developed
36
plants. The most important Bt toxins that are expressed by transgenic plants are lepidopteran-
active Cry1Ab, Cry1Ac and coleopteran-active Cry3Bb in maize (Zea mays L.) and cotton
(Gossypium spp.). Transgenic plants with two or more different Bt transgenes have been
developed and commercialized (Hutchison et al. 2015).
2.19. The problem of Bt-resistance in Pests and Combating Strategies
Recently, field-evolved resistance buildup has been reported against major Bt toxins
in cotton bollworm (Helicoverpa spp), pink bollworm (Pectinophore gosspiella), western
corn rootworm (Diabrotica virgifera) and armyworm (Spodoptera frugiperda) in major
cotton growers like the US and India . Similarly, reduced filed performance of Bt maize
against Spodoptera frugiperda and Busseola fusca respectively, were also reported in South
Africa and Puerto Rico (Storer et al. 2012).
The most common mechanism of insect resistance development is due to disturbance
in the binding pattern of the Bt toxin with the midgut membrane receptors of the host. This
disruption is most likely due to the structural alterations in the receptor domains or variations
in expression of the receptor genes (Castagnola and Jurat-Fuentes 2016). The resistance in
Helicoverpa armigera, Heliothis virescens, and Pectinophora gossypiella is reported to be
the result of mutations in cadherin genes (Morin et al. 2003; Xiao et al. 2017). Another
possible mechanism of resistance associated with the mutations in ABC transporter locus that
is reported in three lepidopteran insects (Xiao et al. 2016).
Two main approaches are used to overcome the insect resistance in Bt crops i.e.
pyramiding and refuge. Delay in resistance evolution is the refuge method where farmers are
instructed to keep abundant non-Bt crops in surroundings of their Bt crops as a refuge
(Gryspeirt and Grégoire 2012). The principle behind this policy is that Bt-resistant larvae
37
when arise in Bt crops undergo a mating with susceptible individuals from non-Bt crops in
surroundings. The offspring will be susceptible to Bt crops as long as the inheritance of
resistance remains recessive (MacIntosh 2010). Gene pyramiding i.e. stacking two or more
genes in the same plant could be another possible approach to combat with the development
of Bt resistance, such as the introduction of second-generation Bt cotton with at least two new
Bt toxins i.e. Monsanto Bollgard II cotton variety and six Cry toxins are the example of gene
pyramiding (Carrière et al. 2016). Another approach used for resistance management is to
use insecticidal genes with a different mode of action e.g. proteinase inhibitors (Schlüter et
al. 2010).
Farmers are using integrated pest management (IPM) system in addition to above-
mentioned resistance management policies in order to achieve a sustainable control against
all insects. IPM combines the biological, chemical, biotechnological and cultural control
methods to prevent pests. These strategies are (1) maintaining the beneficial insect
populations, (2) regular handling of the weed hosts, (3) confirming healthy plant growth or
development, (4) monitoring pest populations regularly. These additional approaches will be
beneficial in advanced insect control and prevention of insect resistance to Bt (Chandler et al.
2011).
2.20. Modified Expression of Vip3Aa Gene in Plants
Cotton and Maize have been successfully transformed with Vip3A genes both
individually as well as in combination i.e. fusion with Cry genes to develop insect resistance
and improved defense mechanism (Meade et al. 2015). Genetically modified cotton
(COT102) and corn (MIR162) expressing Vip3Aa19 and Vip3Aa20 respectively were
registered in the USA in 2008-09 (Chen et al. 2017a; Raybould and Vlachos 2011). Both
38
these events have been pyramided first with cry1Fa (VipCot= Vip3Aa + Cry1Ac + Cry1Fa,
and Agrisure Viptera= Vip3Aa + Cry1Ab + Cry1Fa) and later with Cry1Ab (VipCot=
Vip3Aa + Cry1Ab, and Agrisure Viptera= Vip3Aa + Cry1Ab) for the broad range protection
against lepidopterans (Yang et al. 2015). The corn event MIR162 was further stacked with
coleopteran specific Cry3A and eCry3.1Ab genes to confer resistance against Coleopterans
and Lepidopterans (Graser et al. 2017). After the field performance trials of VipCot plants
against H. armigera attacks for consecutive three years, it was observed that the high
insecticidal efficiency of the toxin tends to decrease as the season progresses (Llewellyn et
al. 2007). In another study, Cotton plants have been successfully transformed with a codon-
optimized, chemically synthesized vip3A gene fused with a chloroplast transit peptide. The
Vip3A concentration in the chloroplast was found three-fold higher than the transgenic
cotton without signaling peptide (Wu et al. 2011).
39
3. MATERIALS AND METHODS
3.1. Codon Optimization and Chemical Synthesis of Insecticidal Genes
Full length nucleotide sequences of Vegetative insecticidal protein 3A (accession
number JQ946639.1) and Allium sativum leaf agglutinin genes (accession number
EU252577.1) were retrieved from GeneBank (Coordinators et al. 2014). The gene sequences
were checked for complete open reading frames (ORFs) by using online tool available on
Expert Protein Analysis System (ExPASy) (Gasteiger et al. 2003). The codon usage was
optimized according to cotton (Gossypium hirsutum) to get high transgene expression
through a web-based tool freely available on Integrated DNA Technologies (IDT) website
(Owczarzy et al. 2008). The Vip3Aa gene under CaMV35S and ASAL gene under phloem
specific RTBV promoter was chemically synthesized by BioBasic Inc. in a single 4870 bp
gene cassette (Vip3Aa+ASAL) cloned in the pUC57 vector under XhoI and HindIII
restriction sites (https://www.biobasic.com/us/gene-splash-gene-in-vector/).
3.2. Preparation of Competent Cells
A well separated single colony of Escherichia coli strain Top 10 from freshly
streaked (1-5 days old) Luria-Bertani (Bertani 1951) (LB) agar plate (Appendix I) was
inoculated into 5 mL of Luria-Bertani broth (Appendix I) with 100 µg/mL tetracycline
(Appendix III) in a 15 mL culture tube and incubated overnight at 37° C in a rotary shaker
with the shaking speed of 250 rpm. Next morning, the starter culture was diluted at the ratio
of 1/100 and incubated again at 37° C for four hours or until the optical density (OD600) of
the culture reached to 0.6-0.8. The cell suspension was then incubated at ice for 10 min and
40
centrifuged at 4000 rpm for 30 min using a JA-14 rotor (Beckman) to get the cell pellet. The
supernatant was discarded and the cells were resuspended in 200 mL of 10% glycerol
(Appendix VI). The cell suspension was incubated at ice for 15 minutes and centrifuged
again at same conditions. Finally, the bacterial cells were washed with 100mM ice-cold
CaCl2 (Appendix VI) and resuspended in 1.5 mL CaCl2 (100mM) + 1.5 mL 10% glycerol.
The aliquots of 80 µL were made and stored at -70° C for future use.
3.3. Transformation of Vip3Aa+ASAL Cassette in Top 10
Chemically synthesized gene cassette (Vip3Aa+ASAL) cloned in pUC57 vector was
transformed into freshly prepared competent cells of E. coli (Top 10) through heat shock
method. Total 2µL of chemically synthesized plasmid DNA (100 ng/µL) was added in an 80
µL vial of competent cells, mixed thoroughly by tapping and incubated at ice for 30 minutes.
The mixture was then subjected to a heat shock of 60-90 seconds at 42° C and immediately
transferred to ice for 5 minutes for cell wall recovery. After that, 1 mL of LB broth was
added in the cell mixture and incubated at 37° C with continuous shaking (250 rpm) for 1
hour. The transformed bacterial cells were centrifuged, resuspended again in 200 µL of LB
broth and spread on LB agar plates containing 100 µg/mL Tetracycline+Ampicillin
(Appendix III) at the rate of 50 and 80 µL per plate and incubated for 16-18 hours at 37° C.
3.3.1. Plasmid Isolation from Positive Clones
The plasmid DNA was isolated using GeneJet Plasmid Miniprep Kit (Catalogue #
K0503) by following the manufacturer’s instructions. The well-separated single colonies
were inoculated in 3 mL of LB broth containing Tetracycline and Ampicillin (100 µg/mL) as
selection drugs and incubated overnight at 37° C in a shaking incubator. Next morning, the
41
culture was centrifuged at 13000 rpm for 5 minutes in the 1.5 mL tube. The harvested cells
were resuspended in 250 µL of RNase A added resuspension solution by vortexing. Total
250 µL of lysis solution was added and properly mixed until the solution became clear. Then
350 µL of neutralization solution was added and gently mixed by inverting the tubes. The
tubes were then centrifuged at 13000 rpm for 5 minutes. The supernatant was loaded onto the
GeneJet spin column (provided with the kit) and centrifuged for 2 minutes. The flow-through
was decanted and the column was washed twice with 500 µL of wash solution (diluted with
ethanol). The column was placed in a new 1.5 mL tube and 20 µL of elution buffer was
added carefully to the center. The tubes containing columns were centrifuged for 5 minutes
to elute the purified plasmid DNA and stored at -20 C.
3.3.2. Confirmation of Positive Clones through PCR
Positive clones were confirmed through polymerase chain reaction (PCR) using
isolated plasmid DNA as a template using following gene-specific primers:
Vip3Aa (F) 5’-ATCACAGAACGGAGATGAGG-3’
Vip3Aa (R) 5’-GTGTACCTCCCGATCTAGTAAC-3’
ASAL (F) 5’-GGAGAAGGTCTTTATGCTGGTC-3’
ASAL (R) 5’-GAAGTATGCAGTCCCCGATCTA-3’
Total 100 ng of the plasmid DNA; 10 pmol of forward and reverse primers; 200 µM
dNTPs, 10X pfu Buffer (with 20mM MgSO4) and 1 unit Taq DNA polymerase enzyme was
used in a 20 µL PCR master mixture. The initial denaturation was done at 94º C for 5
minutes followed by 40 cycles of denaturation at 94º C for 45 seconds, annealing of primers
at 55º C for 45 seconds and extension at 72º C for 45 seconds. The final extension was given
42
at 72º C for 10 minutes. The amplified products were resolved on 1.5% agarose gel
containing 1.0 µg/mL ethidium bromide (Appendix II) and visualized under the UV light.
3.3.3. Confirmation of Positive Clones through Restriction Digestion
Recombinant plasmid containing Vip3A+ASAL cassette was also confirmed through
restriction digestion by using FastDigest XhoI and HindIII restriction enzymes and to
produce sticky ends in the cassette to use further in the cloning experiments. The reaction
mixture was prepared as:
Plasmid DNA (100 ng) 6μL
Green Buffer 2μL
FastDigest HindIII (10 U/μL) 1 μL
FastDigest XhoI (10 U/μL) 1 μL
Sterile H2O up to 20 μL
The reaction mixture was incubated at 37° C for 10 minutes. The digested products
were resolved on 0.8% agarose gel along with 1kb DNA ladder. The required bands were
carefully cut from the gel using a sterilized surgical blade.
3.3.4. Elution of DNA Fragments from Agarose Gel
The required digested bands were purified from the agarose gel using GeneJet Gel
Extraction Kit (Catalogue# K0692) following manufacturer’s manual. The gel slices were
weighed and incubated with DNA binding solution in a water bath preset at 55° C to
completely dissolve the gel slice. The binding solution was used in 1:1 (Volume: Weight)
ratio i.e. 100 μL binding solution for every 100 mg slice of the gel. The gel mixture was
43
dissolved completely by vortexing before loading onto the purification column. The
solubilized gel solution was centrifuged at 13000 rpm for 2 minutes, the flow through was
decanted and the column was washed twice with 700 μL wash buffer (diluted with ethanol).
Finally, total 20 μL of elution buffer was added in the column and centrifuged for two
minutes to elute the DNA and stored at -20° C for further use.
3.4. Restriction Digestion of Plant Expression Vector (pCAMBIA 1301)
The binary vector pCAMBIA-1301 (11837 bp) was used as a plant expression vector
for the cloning of Vip3Aa+ASAL cassette. The map of the vector and gene cassette is shown
in Figure 3.1 and 3.2 respectively. The vector was also digested with the FastDigest XhoI and
HindIII restriction enzymes to produce sticky ends in the pCAMBIA-1301 vector. Reaction
mixture was prepared as:
Plasmid DNA (100 ng) 10μL
Green Buffer 2μL
FastDigest HindIII (10 U/μL) 1μL
FastDigest XhoI (10U/μL) 1μL
Sterile H2O up to 20 μL
44
Figure 3.1: Map of pCAMBIA1301_Vip3Aa+ASAL
Figure 3.2: Schematic diagram of Vip3Aa+ASAL cassette
45
3.4.1. Ligation of Vip3Aa+ASAL Cassette into the pCAMBIA 1301 Vector
The overhangs of the gene cassette (insert) and pCAMBIA 1301 (vector) were ligated
together using T4 DNA Ligase enzyme (Thermo Scientific Rapid DNA Ligation Kit, Cat#
K1422) following the manufacturer’s instructions. The vector to insert ratio was adjusted to
be 1:1 and 1:3 respectively. The Ligation mixture was incubated at 22° C for one hour.
Vector: Insert: 1:1 Vector: Insert: 1:3
Reagent Quantity Reagent Quantity
Vector (85 ng/µL) 1 µL Vector (85 ng/µL) 1 µL
Insert 65.7 ng/µL) 1.3 µL Insert (65.7 ng/µL) 4 µL
Buffer 4 µL Buffer 4 µL
Ligase 1 µL Ligase 1 µL
ddH2O 12.7 µL ddH2O 10 µL
Final volume 20 µL Final volume 20 µL
3.4.2. Transformation and Confirmation of Ligated Product in pCAMBIA 1301
The ligated product was transformed into competent cells of E. coli strain Top10 as
described previously in section 3.3. The mixture was spread on LB agar plates containing
100 µg/mL tetracycline and 50 µg/mL kanamycin (Appendix III) for selection and incubated
overnight at 37° C. Next day, the well-separated single colonies were inoculated in 3 mL LB
broth containing selection drugs in a 15 mL culture tube. The plasmid DNA was isolated and
used as a template in a PCR reaction as explained in section 3.3.1 & 3.3.2 respectively. The
same plasmid DNA was also evaluated through restriction digestion using FastDigest XhoI
46
and HindIII enzymes (section 3.3.3). Glycerol stocks of the positive clones were made for
future use.
3.5. Preparation of Agrobacterium Competent Cells
The competent cells of Agrobacterium tumefaciens strain LBA 4404 were prepared
following the same method as described by (Fisher and Guiltinan 1995). A well separated
single colony was inoculated in 5 mL Yeast Extract Peptone (YEP) broth (Vervliet et al.
1975) (Appendix I) with 5 µL Rifampicin (100µg/mL) (Appendix III) as a selection
antibiotic and incubated at 30° C overnight in a shaking incubator. The overnight grew starter
culture was then diluted in 50 mL YEP broth containing selection drug and incubated at 30°
C on an orbital shaker (300 rpm) until the OD595 reached up to 0.8. The culture was
incubated on ice for 15 minutes and shifted to a 50 mL conical tube. The cells were harvested
by centrifugation of 10 minutes at 4000 rpm (4° C). The cells were washed twice with ice-
cold 10mM HEPES solution (Appendix VI). Finally, the cells were resuspended in 1 mL of
10% glycerol solution (Appendix VI). The aliquots of 80 µL were prepared and stored at -
70° C for future use.
3.6. Transformation of Vip3Aa+ASAL plasmid in Agrobacterium
tumefaciens
Total 2µL of recombinant pCAMBIA 1301 vector harboring genes of interest
(Vip3Aa and ASAL) was transformed into 100 µL freshly prepared chemically competent
cells of Agrobacterium tumefaciens strain LBA4404 through electroporation by using
BioRad Gene Pulser Electroporator (Model 165-2105) at the capacitance of 25 µF; voltage of
2.20 kV with a resistance of 200Ω for 2.6 milliseconds or until a beep was heard.
47
Immediately after electroporation, 1 mL YEP broth was added in transformed cell culture
and incubated at 30° C for 2 hours in a shaking incubator. The cells were spread on YEP-agar
plates (Appendix I) containing 50µg/mL rifampicin and kanamycin for screening of positive
colonies and incubated at 30° C for 36-48 hours.
3.7. Confirmation of Electroporation through Colony PCR
The colony PCR was done to confirm the successful transformation of recombinant
pCAMBIA 1301 vector harboring the cassette of interest in Agrobacterium tumefaciens by
using gene specific primers. The transformed Agrobacterium colonies were dissolved in 10
µL of autoclaved distilled H2O in a 0.2 mL tube. The cell suspension was given a heat shock
at 95° C for 10 minutes and immediately transferred to ice for 1 minute. The lysate was
centrifuged at 13000 rpm for 2 min and a total of 3 µL was used as template in a 20 µL PCR
reaction mixture following the same methods as described in section 3.3.2. For the long-term
storage, glycerol stocks of the positive colonies were prepared and preserved at -70º C.
3.8. Confirmation of Agrobacterium transformation through Restriction
Digestion
Positive clones were also confirmed through restriction digestion analysis using XhoI
and HindIII restriction enzymes. Plasmid DNA was isolated from positive colonies using the
same methods as described in section 3.3.3. The reaction mixture was incubated for 10
minutes at 37° C. The digested samples were resolved on 0.8% agarose gel.
48
3.9. Cotton Transformation
3.9.1. Selection of a Suitable Cotton Variety
Locally developed and commercially approved cotton (Gossypium hirsutum L.)
variety of Centre of Excellence in Molecular Biology namely CEMB-33 was selected for
transformation on the basis of better germination rate and its overall performance in the field.
The seeds were obtained from Seed Biotechnology laboratory of the Centre.
3.9.2. Delinting of Cotton Seeds
The cotton seeds of selected variety were delinted by treating with commercially
available concentrated sulphuric acid (100 mL/kg) for complete removal of the lint. The
seeds were mixed vigorously to remove the lint completely. The seeds were then washed
thoroughly under tap water to remove the acid because it may affect the seed germination.
The sinker seeds were collected and used for transformation experiments.
3.9.3. Sterilization and Soaking of Seeds
The cotton seeds were surface sterilized with the addition of 5% HgCl2 (2 mL) and
10% SDS (1 mL) solutions. During the sterilization process, the cotton seeds were
continuously mixed by vigorous shaking for 5 minutes and then rinsed thoroughly with
autoclaved distilled water to completely remove the detergents. Approximately 500 seeds
were soaked in each experiment in an autoclaved flask with almost 10 mL of autoclaved
distilled water to keep the seeds moist. The flask containing the seeds were covered with
black cloth and incubated at 30° C for 1-2 days to get maximum germination. During the
sterilization process, aseptic conditions were strictly maintained. The germination index was
calculated was follows:
49
3.9.4. Murashige and Skoog (MS) Medium Preparation
Murashige and Skoog medium (Murashige and Skoog 1962) (Appendix I) was used
to culture the transformed embryos. After autoclaving, the medium was allowed to cool
down to 50° C and cefotaxime (250µg/mL) (Appendix III) was added in the medium. The
medium was divided in two portions, in half of the medium, kinetin (1 mg/mL) (Appendix
VI) and B5 vitamin complex (1 mL/L) (Appendix I) was added and poured in glass culture
tubes. The other half was poured in petri plates without any drug and hormone.
3.9.5. Bacterial Inoculum Preparation
The transformed Agrobacterium cells containing Vip3Aa+ASAL cassette were
streaked on YEP agar plate containing 50 µg/mL kanamycin and 100 µg/mL rifampicin and
incubated at 30° C for two days. A well separated single colony was inoculated in 10 mL of
YEP broth (Appendix I) containing selection drugs and incubated for two days in a rotary
shaker at 30º C. The bacterial cells were pellet down by centrifugation at 4000 rpm for 15
minutes and dissolved in 10 mL of MS broth (Appendix I).
3.9.6. Isolation of Embryos
The testa and the cotyledonary leaves of the well-germinated seeds were removed
carefully with the help of a surgical blade; a sterilized forcep and a scalpel handle to isolate
mature cotton embryos and kept on moist filter paper to keep them wet.
50
3.9.7. Agrobacterium Mediated Transformation (Shoot Apex Method)
The shoot-apex-cut method was used for genetic transformation of cotton as
described by (Rao et al. 2011b). The shoot tips of the isolated mature cotton embryos were
subjected to tip injury at epicotyl-end using a sharp double-edged blade held vertically in a
petri plate. After injury, the embryos were shifted immediately to the MS broth containing
Agrobacterium tumefaciens cells transformed with Vip3Aa and ASAL genes, for bacterial
inoculation and incubated at 30° C on a shaking incubator for 1 hour. Total 4500 mature
cotton embryos were used in all transformation experiments.
3.9.8. Co-cultivation
After Agrobacterium treatment the embryos were taken out of the suspension; the
embryos were kept on autoclaved filter papers for some time to completely absorb the
bacterial culture and then shifted on MS medium plates containing 250 µg/mL cefotaxime
(Appendix III) as an antibiotic to avoid bacterial contamination and co-cultivated for the next
3 days. The plates were sealed with parafilm and kept in a growth room adjusted at 25 ± 2° C
and 16: 8 hours light: dark cycle. After two to three days the plantlets were shifted to glass
test tubes containing MS media supplemented with kinetin and B5 vitamin complex.
3.9.9. Transformation Efficiency
The transformation efficiency was calculated by the number of plants surviving after
4-6 weeks on MS medium and initial screening through transient expression of β-
glucuronidase (GUS) reporter gene in comparison with total number of embryos co-
cultivated in all the experiments as described by (Satyavathi et al. 2002).
51
3.9.10. Transferring Transgenic Plants to Soil
After 4-6 weeks, the rooted plantlets were taken out of the glass tubes carefully, roots
were washed with autoclaved water, dipped into Indole Butyric Acid (IBA 1mg/mL)
(Appendix VI), dried with tissue-paper and transferred to sigma pots containing sterilized
soil containing thoroughly mixed, equal proportion of clay, sand and peat moss (1:1:1)
(Rashid et al. 2004). These plants were covered with plastic bags to maintain proper humidity
and were kept in growth room at 28±2° C and a photoperiod of 16 hours light and 8 hours
dark under light intensity of 250-300 µmol m-2
S-1
. The plants were exposed to sunlight for
acclimatization after 2-3 days, the polythene covers were removed two times a day. The
duration of exposure time increased gradually until it reached 4-6 hours a day. The stem of
cotton plants were lignified due to newly formed secondary tissues after exposure to sun
light, it became hard. The putative transgenic cotton plants were then shifted to the field in
containment for better growth and then these plants were subjected to molecular analysis.
3.10. Molecular Analyses of Transgenic Cotton Plants
3.10.1. Genomic DNA Isolation
A combination of two protocols (Sukumar et al. 1997; Zhang et al. 2000) with some
modifications was followed for isolating genomic DNA from the cotton leaves. Total 1 gram
of the terminal leaf of cotton plant was ground to a fine powder in liquid nitrogen with the
help of a chilled pestle and mortar. Approximately 100 mg grounded powder was transferred
to a 1.5 mL tube and 750 µL of CTAB extraction buffer (Appendix IV) was added. The tubes
were mixed vigorously and incubated at 65º C in a water bath for about 30-60 minutes.
During the incubation, tubes were inverted 3-4 times for complete mixing. After the
incubation, tubes were brought back to room temperature and an equal volume of
52
chloroform: isoamyl alcohol (24:1) was added. Tubes were gently inverted or vortexed
several times to mix the ingredients completely. This mixture was centrifuged for 20 minutes
at 13000 rpm. The aqueous phase was carefully transferred to new 1.5 mL tubes and the step
was repeated twice. An equal volume of ice chilled isopropanol was added and tubes were
inverted 2-3 times till the entire DNA was aggregated. The tubes were kept at -20° C for
overnight. Next morning, the tubes were centrifuged at 13000 rpm for 30 minutes. The
supernatant was discarded and the DNA pellet was washed twice with 1 mL of 70% Ethanol.
The DNA pellet was resuspended in autoclaved distilled water or injection water after the
pellet was completely air-dried. The DNA concentration was estimated by both
spectrophotometer and resolving the samples on 0.8% agarose gel with λ/HindIII marker.
3.10.2. Polymerase Chain Reaction
Polymerase chain reaction was done for the detection of Vip3Aa and ASAL genes in
putative transgenic cotton plants with the same conditions as mentioned in the section 3.3.2
by using gene specific detection primers at 64°C annealing temperature. The DNA extracted
from untransformed plants was used as a negative control and the plasmid DNA as a positive
control. The amplified PCR fragment was resolved on 1.5 % agarose gel and observed under
UV light.
3.10.3. Transient Expression of GUS Gene
The transient expression of β-glucuronidase (GUS) reporter gene in young leaves of
putative transgenic cotton plants was taken through histochemical GUS assay. The GUS
solution was prepared by adding 25 mg/L X-gluc, 10mM EDTA, 100mM NaH2PO4, 0.1%
Triton X-100 and 50% methanol. The pH of the solution was adjusted to 8.0. The young
53
leaves from each transgenic and non-transgenic cotton plants were inundated with GUS
solution in 1.5 mL tubes and kept at 37° C overnight and observed under compound
microscope for appearance of blue colour.
3.11. Expression Analysis of Transgenic Cotton Plants
3.11.1. Total RNA Extraction
Total RNA from transgenic cotton leaves was extracted using a modified protocol as
described by (Jaakola et al. 2001). Newly emerged cotton leaves were ground in liquid
nitrogen with the help of a chilled pestle and mortal. For each 100 mg of ground sample, 750
µL of RNA extraction buffer (Appendix IV) was added. The tubes were thoroughly mixed by
inverting the tubes several times and incubated at 65° C for 30 minutes. Equal volume (750
µL) of chloroform: isoamylalcohol was added in the tubes and centrifuged twice at 13000
rpm and 4° C for 30 minutes. The supernatant was transferred to a new 1.5 mL tube and 10M
LiCl (60%) was added. The samples were incubated at 4° C overnight for complete
precipitation of the RNA. Next morning, the tubes were centrifuged at 13000 rpm (4° C) for
30 minutes. The supernatant was discarded and the pellet was washed twice with 70%
ethanol. The pellet was air-dried and dissolved in 100 µL of preheated SSTE buffer
(Appendix IV). The purification was done by mixing the contents of tube with an equal
volume (100 µL) of phenol: chloroform: isoamylalcohol (25: 24: 1) and then with an equal
volume of chloroform: isoamylalcohol (24:1). The supernatant was taken and two volumes of
absolute ice cold ethanol were added. The samples were incubated at -20° C overnight. Next
morning, the tubes were centrifuged at 13000 rpm for 20 minutes at 4° C. The pellet was
washed with 70% ethanol; air-dried and resuspended in DEPC treated water.
54
3.11.2. Complementary DNA (cDNA) Synthesis
The cDNA was synthesized through one-step reverse transcriptase RT-PCR with
random hexamers using RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific,
K1622). Following reaction mixture was prepared:
Reagent Quantity
Template RNA 1µg
Random Hexamers 1µL
Nuclease-Free water 10 µL
Total volume 12 µL in a 200 µL tube
The template RNA was linearized by first incubating it at on 65° C for 5 minutes in a
thermocycler and then on ice for 2 minutes. The ingredients of the master mix were added to
the linearized RNA as follow:
Reagent Quantity
5X Reaction Buffer 4 µL
10 mM dNTP Mix 2 µL
RiboLock RNase Inhibitor 1 µL
RevertAid Reverse Transcriptase 1 µL
Total Volume 8 µL
The cDNA was prepared by a single step reaction in a thermocycler by incubating it at 25° C
for 5 minutes, followed by 60 minutes at 42° C. The reaction was terminated by heating at
70° C for 5 minutes. The cDNA was preserved at -20° C for further use in real time
experiments.
55
3.11.3. Quantitative Real Time qRT-PCR
The quantitative Real-time qRT-PCR reaction was carried out in an iQ5 cycler (BIO-
RAD) with a 96-well plate by using the Maxima SYBR Green/ROX qPCR Master Mix (2X)
(Thermo Scientific, K0221). The housekeeping gene, Glyceraldehyde 3-phosphate
dehydrogenase (GAPDH), was used as a comparative control to normalize the data. Two
hundred ng of cDNA was used as template in qRT-PCR. The following gene specific and
GAPDH primers were used:
R-Vip (F) 5’-CGGCTCCCTTAATGATTTGA-3’
R-Vip (R) 5’-CGATCTGCAATGAAAGAGCA-3’
R-ASAL (F) 5’- ATGCTGGTCAATCACTCGAT -3’
R-ASAL (R) 5’- AACAGAGTTCGAAGCCCAAA -3’
GAPDH (F) 5’-AGGAAGAGCTGCTTCGTTCA-3´
GAPDH (R) 5´-CCGCCTTAATAGCAGCTTTG-3´
In the real time (RT-PCR) reaction, initial denaturation was done at 95˚ C for 4
minutes, followed by 40 cycles of denaturation at 95˚ C for 45 seconds, 61˚ C for 45 seconds,
and 72˚ C for 45 seconds, final extension was given at 72˚ C for 10 minutes. The Ct values
obtained from different transgenic cotton plant samples were statistically analyzed using iQ5
software (Bio-Rad) version 1.0.
3.11.4. Dipstick Analysis of Putative Transgenic Cotton Plants
Total crude protein from the fresh leaves of putative transgenic cotton plants was
isolated by grinding the leaves in liquid N2 using a chilled pestle and mortar. Total 700 µL of
protein extraction buffer (Appendix IV) was added. The tubes were vortexed briefly and left
56
overnight at 4° C. Next day, the tubes were centrifuged at 13000 rpm (4° C) for 15 minutes
and supernatant was used a template for the assay. The Vip3Aa specific dipsticks
(ImmunoStrip Cat# STX 83500/0050 Agdia) were dipped into the isolated crude protein
samples for 30 minutes and left to be dried for 15 minutes. The dipsticks were observed for
appearance of bands on the specific place.
3.11.5. Quantification of Vip3Aa Protein in Transgenic Cotton Plants through ELISA
Enzyme Linked Immunosorbent Assay (ELISA) was performed for the detection and
quantification of Vip3Aa protein in transgenic cotton plants using Agdia Vip3Aa ELISA kit
(Cat# PSP 83500) following manufacturer’s manual. Fresh leaf samples of transgenic cotton
plants were ground to a very fine powder with the help of liquid nitrogen in chilled pestle
mortar and dissolved in 700 µL protein extraction buffer. Tubes were mixed and incubated at
4° C overnight. Next morning, the tubes were centrifuged for 25 min at 4° C and 13000 rpm.
Total 100 µL supernatant was used for ELISA in comparison to non-transgenic cotton plants
as negative control (total crude protein extracted from non-transgenic cotton plant) and
enzyme conjugate was dispensed in appropriate test wells of ELISA plate. The contents of
the wells were mixed thoroughly with the help of a strip holder in a rapid circular motion for
20-30 seconds. The ELISA plate was covered with aluminium foil and incubated for one
hour at room temperature. After the incubation, the contents of the wells were discarded and
washed thoroughly with 1X PBST and air dried followed by addition of 100 µL TMB
substrate to each well of ELISA plate preceded by incubation at room temperature for 30
minutes. The optical density was measured at 650 nm on an ELISA plate reader
(ELx800/Micro Plate ELISA Reader).
57
The known standards of Vip3Aa protein (4.52, 5.23, 7.45, 9.95, 10.94, 13.1, 14.47,
15.43 and 16.91 μg/mL) were prepared. Total 200 μL Bradford dye (Bradford 1976) was
added in each well and absorbance was taken on Backman spectrophotometer reader at 650
nm wavelength. A standard curve was prepared by plotting XY (scatter) graph on an excel
sheet. A trend line was added to find out linear regression and R2 values on the graph. The
standard curve directly quantified unknown protein concentrations expressed in transgenic
cotton plants with reference to known standards of Vip3Aa.
3.12. Morphological Traits of Transgenic Cotton Plants
Different morphological and physiological traits were also studied in transgenic
cotton lines in T1 progeny in triplicates. Ordinary one-way Analysis of Variance (ANOVA)
was performed to compare the significance level between transgenic cotton lines with control
(Dunnett’s test) at 95% (P<0.05) confidence level using GraphPad Prism software version 5
for Windows (Prism 2007).
3.12.1. Plant Height (Inch)
Plant height of the transgenic cotton plants was recorded in comparison to non-
transgenic control plants from the base to the apex of the plant by using a measuring tape.
3.12.2. Number of Monopodial and Sympodial Branches per Plant
Average number of monopodial (indirect fruiting branches) and sympodial or direct
fruiting branches of transgenic and non-transgenic control plants were counted at the onset of
maturity.
58
3.12.3. Number of Bolls per Plant
Average number of mature and immature bolls on each transgenic plant line was
counted in comparison to non-transgenic control cotton plants.
3.12.4. Ginning out Turn Percentage (GOT %)
Average ginning out turn GOT% in different transgenic lines was calculated by using
the formula:
3.12.5. Scanning Electron Microscopic Analysis of Cotton Fiber
Surface morphology of fiber samples from transgenic and non-transgenic cotton
plants was examined by using scanning electron microscope (Model SU8010 Hitachi Japan).
Each fiber was excised into 3 parts i.e. tip, middle and base. The screw pitch of fiber and the
distance of fiber rotation in 360 degrees were measured three times for each sample by SEM
with an accelerating voltage of 20 kV and 10 µA current under 400X, 1000X and 4000X
magnifications.
3.13. Physiological Traits of Transgenic Cotton Plants
Different physiological parameters like net photosynthetic rate (A); transpiration rate
(E) and gaseous exchange rate (GE) were measured on a fully expanded cotton leaves of
each transgenic and control (non-transgenic) plant lines in triplicates using CIRAS-3 portable
photosynthesis system infrared gas analyzer (PP Systems, USA). Measurements were
performed from 1000 to 1300 hours with the specific adjustments of molar flow rate of air at
403.3 µmol m-2
s-1
, atmospheric pressure at 99.9 kPa, water vapor pressure in chamber at 6.0-
59
8.9 mbar, PAR at leaf surface at 1000-1711 µmol m-2
s-1
, temperature of leaf at 28.4-32.4° C,
ambient temperature 22.4-27.9° C and ambient CO2 concentration of 352 µmol mol-1
.
3.14. Insect Bioassays
The transgenic cotton plants were subjected to the standard laboratory leaf-detach
bioassays against American bollworm (Heliothis armigera) and clip bioassay against
whitefly (Bamisia tabaci) to assess the efficacy of insecticidal toxins.
Laboratory bioassay with American bollworm was performed in three replicates i.e.
three plants from each of the transgenic and non-transgenic control cotton lines in T2
generation were selected; one fresh leaf per plant was taken and placed on a wet sterile filter
paper in a 20 cm petri dishes. Three pre-fasted 2nd
instar larvae were released in each of the
plate and allowed to feed on the leaf. The plates were sealed with parafilm and kept at 25±2º
C, 70% relative humidity and 16:8 light and dark cycle in a growth room. The percent
mortality rate was recorded on third day of infestation using the following formula:
On the basis of mortality, the plants were categorized as resistant (40% or more insect
mortality) or susceptible (less than 40% insect mortality).
In planta insect bioassay with Bamisia tabaci was also performed under controlled
conditions. The transgenic cotton plants of T2 generation (6-10 leaf stage) were grown in a
glass house at 37 ± 2° C, 14L/10D and ≈60% humidity. The plants were left un-infested for
one week by carefully isolating them in a net cage. The whitefly culture was maintained on
non-transgenic cotton plants in a separate greenhouse at same conditions. Before the start of
60
bioassay, the 0-24 hour old whitefly adults were captured using a manual aspirator and kept
on ice to minimize environmental stress. The ice-inactivated whiteflies were carefully
released into the clip cage facing the under-side the leaf. Three biological replicates from
each transgenic and control cotton lines were used in the bioassay and to each replicate, three
clip cages having four whiteflies (2M/2F) were released. The mortality data was recorded on
third day of infestation.
One-way Analysis of Variance (ANOVA) was performed to compare the significance
mortality levels between transgenic and non-transgenic control lines (Dunnett’s test) at 95%
(P<0.05) confidence level using GraphPad Prism software version 5 for Windows (Prism
2007).
3.15. Fluorescence in situ Hybridization (FISH)
3.15.1. Preparation of Chromosome
Root tips measuring 1-2 cm long were collected from mature embryos of transgenic
and non-transgenic cotton plants and fixed in fixative solution (3 volumes Ethanol and 1
volume Glacial Acetic Acid) for overnight. Roots were washed thrice with distilled water
after removal of fixative solution. Meristematic portion of roots (1-2 mm) was cut and
incubated in an enzyme solution containing 2% Pectolyase (Sigma cat# P 3026) and 3%
Cellulase (Sigma cat# C 1184) at 37° C for 6 hours and washed carefully with distilled water.
Chromosomes were spreaded on microscopic glass slides with a drop of fixative solution and
air-dried. The slides were observed under phase contrast microscope (Olympus Model BX51)
and the best slides were selected for further analysis. The slides were then dehydrated in a
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serial dilution of ethanol (70%, 95% and 100%), for five minutes in each solution
respectively. The slides were then stored at room temperature.
3.15.2. RNase Treatment
The working solution of 1% RNase was prepared by diluting the stock solution 100
times with 8 µL of 1% RNase A, 10 µL 1M Tris HCl (pH 7.5), 3 µL of 15 mM NaCl and 987
µL of ddH2O. Total 100 µL of RNase working solution was added on each slide and covered
with a glass coverslip and incubated in water bath for 45 minutes at 37° C. Slides were gently
washed three times with 2X SSC solution (Appendix V) at room temperature and then
dehydrated in each dilution of ethanol (70%, 95% and 100%) for 5 minutes.
3.15.3. Hybridization
The hybridization solution (Appendix V) was denatured at 80° C for 10 minutes
followed by quick chilling on ice for 5 minutes. Total 35 µL of hybridization solution was
added on each slide, covered with a cover slip and air dried. The chromosomes were then
denatured in 2X SSC solution at 80° C for 10 minutes in a water bath. Finally, the slides
were incubated at 37° C overnight in a wet chamber.
3.15.4. Post Hybridization Washes
Next morning, the slides were washed twice with 2X SSC solution and once with 4X
SSC solution at 42° C for 10 minutes.
3.15.5. Color Detection Reaction
The slides were then washed thrice (5 minutes each) in TBS buffer (Appendix V) and
incubated for 30 minutes in blocking solution (TBS; 0.1 % Triton X-100; 1.0% Blocking
62
Reagent). The slides were then incubated with anti-DIG antibody (diluted 400 times with
blocking solution) for minimum 4 hours at room temperature. The slides were washed thrice
in TBS for 5 minutes each and incubated overnight in color substrate NBT/BCIP solution.
The reaction was stopped by rinsing the slides under tap water.
3.15.6. Counterstaining with DAPI
The stock solution of DAPI stain was ice-thawed and diluted 250 times by adding 8
µL DAPI (100 µg/µL) and 1992 µL Mellavaine buffer (Appendix V). Each slide was covered
with total 500 µL of diluted DAPI stain solution and incubated for 5 minutes at room
temperature. Slides were then washed with 3 mL Mellavaine buffer, covered with glass cover
slip and stored in dark at 4° C.
3.15.7. Counterstaining with Propidium Iodide (PI)
Propidium Iodide (PI) stock solution was thawed on ice and diluted 2500 times by
adding 0.8 µL PI and 2 mL IX PBS (Appendix V). Total 500 µL of diluted PI solution was
added on each slide and incubated at room temperature for 5 minutes. The slides were then
washed with 3 mL of 1X PBS buffer, covered with glass slip and stored in dark at 4° C.
3.15.8. Fluorescent Signal Detection
The fluorescent signals were detected by Fluorescent microscope (Olympus Model
BX6l) on Blue Filter for DAPI and Red Filter for PI. The picture of fluorescence signal was
taken by CCD camera attached with microscope and analyzed/enlarged by using Adobe
Photoshop 7.0.
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4. RESULTS
4.1. Transformation of pUC57_Vip3Aa+ASAL Cassette in E.coli
The chemically synthesized pUC57_Vip3Aa+ASAL cassette transformed in E.coli
strain Top10 was confirmed through polymerase chain reaction (PCR) by using gene specific
detection primers as mentioned in section 3.3.2. Plasmid DNA was isolated from positive
clones and used as a template in a PCR reaction. Amplification of 587 bp internal fragment
as shown in Figure 4.1 confirmed the presence of Vip3Aa+ASAL cassette in pUC57 vector
according to the suppliers (Bio-Basic Inc.) information.
Figure 4.1: Confirmation of Vip3Aa+ASAL cassette in pUC57 vector through colony PCR
Lane 1, 2, 3: Plasmid DNA isolated from positive colonies
Lane 4: Blank
Lane 5, 6, 7, 8, 9, and 10: Plasmid DNA isolated from positive colonies
Lane 11: 50 bp DNA ladder
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4.2. Confirmation of Vip3Aa+ASAL Cassette in pUC57 Vector
The chemically synthesized gene cassette (Vip3Aa+ASAL) cloned in the pUC57
vector was also confirmed through restriction digestion using FastDigest HindIII and
FastDigest XhoI enzymes. The appearance of 4870 bp fragment of Vip3Aa+ASAL cassette
and 2710 bp of pUC57 vector on 0.8% agarose gel confirmed the successful release of
required gene cassette as shown in Figure 4.2. The required band of Vip3Aa+ASAL was cut
with a sterile surgical blade and eluted from agarose gel for further ligation into pCAMBIA
1301 plant expression vector.
Figure 4.2: Confirmation of Vip3Aa+ASAL cassette in pUC57 vector through restriction digestion
Lane 1, 2, 4, 5: Plasmid DNA isolated from positive colonies
Lane 3: 1 kb DNA ladder
65
4.3. Cloning of Vip3Aa+ASAL Cassette in Plant Expression Vector
Once the gene cassette was confirmed in pUC57 vector through restriction digestion
and polymerase chain reaction, the next step was to clone the gene cassette in plant
expression vector. For this, pCAMBIA 1301 binary vector was also digested with FastDigest
XhoI and FastDigest HindIII enzymes to create sticky ends. The 4870 bp fragment of
Vip3Aa+ASAL gene cassette (insert) excised from pUC57 vector was ligated with eluted
fragment (10756 bp) of pCAMBIA 1301 in (1:1) vector to insert ratio using T4 DNA ligase
enzyme. Figure 4.3 showed the digestion of pCAMBIA 1301 vector.
Figure 4.3: Digestion of plant expression vector (pCAMBIA 1301)
Lane 1-3: Restriction digestion of plant expression vector
Lane 4: 1 kb DNA ladder
66
4.4. Transformation and Confirmation of Ligated Product in pCAMBIA
1301
The ligated product was transformed into competent cells of E. coli strain Top10. The
proper ligation of Vip3Aa+ASAL cassette in plant expression vector was confirmed through
restriction digestion using FastDigest XhoI and HindIII enzymes. The 4870 bp fragment
excised from pCAMBIA 1301 vector confirmed the successful ligation of gene cassette as
depicted from figure 4.4.
Figure 4.4: Digestion of plant expression vector (pCAMBIA 1301) ligated with Vip3Aa+ASAL cassette
Lane 1: 1 kb DNA ladder
Lane 2-14: Restriction digestion of ligated pCAMBIA 1301
67
1 2 3 4 5 6 7 8 9 10
4.5. Electroporation of pCAMBIA1301_Vip3Aa+ASAL into Agrobacterium
After the confirmation of successful ligation of Vip3Aa+ASAL gene cassette, the
recombinant vector pCAMBIA1301_Vip3Aa+ASAL was then electroporated in to the
competent cells of Agrobacterium strain LBA4404. The transformation of Vip3Aa+ASAL in
Agrobacterium cells was confirmed through colony PCR by using gene specific internal
primers of Vip3Aa gene. The amplification of 682 bp fragment confirmed the presence of
gene cassette in Agrobacterium cells (Fig 4.5).
Figure 4.5: Confirmation of Vip3A+ASAL cassette in Agrobacterium tumefaciens LBA4404
Lane 1: 1 kb DNA ladder
Lane 2-6, 8-10: Screened colonies
Lane 7: Negative Colony
68
4.6. Confirmation of Vip3A+ASAL Cassette in Agrobacterium through
Restriction Digestion
The presence of Vip3A+ASAL gene cassette in Agrobacterium was also confirmed
through restriction digestion of the pCAMBIA1301_Vip3Aa+ASAL plasmid with FastDigest
HindIII and FastDigest XhoI enzymes. The release of 4870 bp fragment confirmed the
presence of pCAMBIA1301_Vip3Aa+ASAL vector in Agrobacterium positive colonies
(Figure 4.6).
Figure 4.6: Confirmation of Vip3A+ASAL cassette in Agrobacterium through restriction digestion
Lane 1: 1 kb DNA ladder
Lane 2-11: Positive clones
69
4.7. Cotton Transformation
4.7.1. Shoot Apex Method of Agrobacterium Mediated Transformation
After the confirmation of construct in Agrobacterium, a commercially approved
locally developed cotton variety CEMB-33 (Gossypium hirsutum) was transformed with
Vip3Aa+ASAL construct as described by (Rao et al. 2011b). Approximately 4500 mature
embryos were treated in ten cotton transformation experiments. After eight weeks of
regeneration on MS medium, fifty three plants were able to survive and shifted in loamy soil.
The initial screening of plants was done with the help of GUS marker gene (Satyavathi et al.
2002). The overall transformation efficiency remained 1.17%. The steps of transformation
are shown in figure 4.7.
Figure 4.7: A schematic diagram of Agrobacterium mediated transformation
70
4.8. Molecular Analyses of Putative Transgenic Cotton Plants (T0 Progeny)
4.8.1. Confirmation of Putative Transgenic Cotton Plants through PCR (T0 progeny)
The successful integration of Vip3Aa+ASAL cassette in putative transgenic cotton
plants was confirmed through PCR using genomic DNA extracted from fresh cotton leaves
as mentioned in section (3.4.1.). The appearance of 587 bp fragment on 1.5% agarose gel
using gene specific internal detection primers confirmed the successful integration of genes
in cotton genome as shown in figure 4.8 (A & B). Total eighteen plants (L1P3; L1P5; L2P3;
L3P2; L4P2; L4P4; L5P2; L5P3; L6P2; L6P3; L34P1; L34P2; L34P4; L35P1; L35P2; L35P3; L36P1;
L38P4) were found positive out of fifty three plants that were shifted in field.
Figure 4.8: Confirmation of putative transgenic cotton plants through PCR
A B
Lane 1: 50 bp DNA ladder
Lane 2: Positive control (plasmid DNA)
Lane 3-16: Putative transgenic plants
Lane 1: 50 bp DNA ladder
Lane 2, 4-6: Putative transgenic plants
Lane 3: Negative control (non-transgenic plant)
B
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4.8.2. Quantification of Vip3Aa Protein through ELISA (T0 progeny)
The protein expression analysis of Vip3Aa in transgenic cotton plants was studied
through quantitative ELISA test. Total crude protein from fresh leaves of each transgenic
and non-transgenic cotton (control) plants was used as an antigen against Vip3Aa specific
antibodies coated in ELISA plate wells. The average optical density (O.D650) for each
protein sample was measured on an ELISA plate reader. The concentration of Vip3Aa
protein expressed in transgenic cotton plants was calculated with the help of a standard
curve of known standards of Vip3Aa protein (figure: 4.9). It was found that the maximum
concentration of Vip3Aa protein (4.26 µg/mL) was observed in plant L6P3 and minimum
protein concentration was seen in plant L34P1 when compared with negative control (non-
transgenic cotton plant) as depicted in figure 4.10.
Figure 4.9: Standard curve of known Vip3Aa standards
72
Figure 4.10: Quantification of Vip3Aa protein through ELISA
4.8.3. Transient Expression of GUS Gene (T0 progeny)
The transgenic cotton plants were further screened through histochemical staining of
β-glucuronidase (GUS) gene. The 5-bromo 4-choloro 3-indolyl glucuronide (X-gluc) when
hydrolyzed by β-glucuronidase enzyme within transformed plant tissues gives an insoluble
blue precipitation by oxidative dimerization of indolyl derivative. Fresh leaves of putative
transgenic cotton plants were incubated with X-gluc solution and kept at 37º C overnight.
The appearance of blue color in putative transgenic cotton plants determined the success of
transformation in these plants as compared to non-transgenic cotton plants where appearance
of blue colour was not visible (Figure 4.11).
73
Figure 4.11: Transient expression of GUS gene
4.8.4. Dipstick Analysis of Transgenic Plants (T0 progeny)
The transgenic cotton plants were also analyzed through Vip3Aa specific immunostrips/
dipsticks. The upper band of coated antibody of GAPDH housekeeping gene served as the
positive control to assure the accuracy of the test, while the lower coated band is of anti-
Vip3Aa antibody that specifically detects the presence of Vip3Aa protein in the sample.
Dipstick analysis for some of the transgenic plants is shown in Figure 4.12.
Figure 4.12: Dipstick analysis of transgenic cotton plants
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4.9. Advancement of Next Generation (T1 Progeny)
Transgenic cotton plants namely L1P3; L1P5; L2P3; L3P2; L4P2; L4P4; L5P2; L5P3; L6P2;
L6P3; L34P1; L34P2; L34P4; L35P1; L35P2; L35P3; L36P1 and L38P4 which were analyzed for the
integration of genes of interest through PCR, transient expression of GUS gene, expression
and quantification of Vip3Aa protein through dipstick assay and ELISA. The transgenic
cotton plants that showed better protein expression in T0 generation having good
morphological/physiological characteristics and showing better yield performance in field
conditions namely (L3P2; L4P4; L5P3; L6P3; L34P2) were further selected for the advancement
of T1 generation. The successful inheritance of gene cassette in advanced transgenic cotton
lines was confirmed through PCR. The mRNA expression levels of Vip3Aa, ASAL
insecticidal genes and quantification of Vip3Aa protein was also studied in T1 transgenic
lines. Morphological and physiological parameters were compared with non-transgenic
control plants and analyzed statistically (ANOVA; Dunnet test). Insect bioassays with cotton
bollworm and whitefly were conducted to evaluate the efficacy of transgenes.
4.9.1. Confirmation of Transgenic Cotton Plants through Polymerase Chain Reaction
(T1 progeny)
The transgenic cotton plants in T1 progeny were screened through PCR using gene
specific internal primers. The genomic DNA was isolated from fresh cotton leaves according
to previously described protocol (3.4.1.). The amplification of 587 bp fragment in the
following transgenic cotton plants (L3P2-A; L3P2-C; L3P2-D; L4P4-B; L4P4-D; L5P3-A; L5P3-
B; L5P3-E; L6P3-B; L6P3-C; L34P2-B) showed the successful inheritance of gene cassette in T1
generation. The PCR results for some of the transgenic cotton plants are shown in figure
4.13.
75
Figure 4.13: Confirmation of transgenic cotton plants through PCR in T1 progeny
Lane 1: Negative Control (DNA from non-transgenic cotton plants)
Lane 2: Positive control (plasmid DNA containing Vip3Aa+ASAL cassette)
Lane 3: 50 bp DNA ladder
Lane 4-8: Genomic DNA extracted from transgenic plants of T1 progeny
4.9.2. The mRNA Quantification of Insecticidal Genes (Vip3Aa and ASAL) in
Transgenic Cotton Plants
The mRNA expression of Vip3Aa and ASAL insecticidal genes was quantified
through quantitative real time qRT-PCR. The mRNA transcripts were first reverse
transcribed into complementary DNA (cDNA) using oligo (dt) random oligomers. The
cDNA of transgenic cotton plants from each transgenic line (L3P2; L4P4; L5P3; L6P3; L34P2)
expressing Vip3Aa and ASAL insecticidal genes were then exponentially amplified by a real
time thermocycler machine using gene specific real time primers as mentioned in Section
3.5.3 of methodology. The real time increase in concentration of amplicons in the reaction
was monitored with cyber green dye. The housekeeping gene Glyceraldehyde 3-phosphate
76
dehydrogenase (GAPDH) was used as an internal control for the normalization of data. The
maximum mRNA expression level of Vip3Aa gene (8.7 fold) was observed in transgenic line
L6P3 (Figure 4.14 A) and the maximum expression level of ASAL gene (5 fold) was seen in
the line L4P4 (Figure 4.14 B).
Figure 4.14: mRNA expression of Vip3Aa & ASAL genes through qRT-PCR
4.9.3. Quantification of Vip3Aa Protein through ELISA (T1 progeny)
Transgenic cotton plants that were confirmed through PCR and real time PCR were
then subjected to be evaluated through quantitative ELISA for the expression and
quantification of Vip3Aa protein inT1 transgenic lines. Maximum protein concentration was
seen in transgenic cotton plants of L6P3 line when compared with non-transgenic cotton
plants (negative control) as exhibited in figure 4.15.
77
Figure 4.15: Quantification of Vip3Aa protein through ELISA in T1 transgenic lines
4.9.4. Dipstick Analysis of Transgenic Plants (T1 progeny)
The transgenic cotton plants were also analyzed through Vip3Aa specific dipsticks in
T1 transgenic lines as described in section 4.8.4. Dipstick analysis for some of the transgenic
plants is shown in Figure 4.16.
Figure 4.16: Dipstick analysis of transgenic cotton plants
78
4.10. Morphological and Physiological Characteristics of Transgenic Plants
in T1 Progeny
Different morphological parameters (plant height; number of bolls per plant; number
of monopodial and sympodial branches per plant; overall yield per plant (lint + seed);
ginning out turn percentage (GOT%); scanning electron microscopic analysis of fibre) and
physiological parameters e.g. photosynthetic rate; transpiration rate; gaseous exchange rate of
transgenic cotton lines in T1 progeny were studied in comparison to non-transgenic (negative
control) plants. The ordinary one-way Analysis of Variance (ANOVA) was performed to
compare the significance level between transgenic cotton lines with control i.e. (Dunnett’s
test) at 95% (P<0.05) confidence level. All the transgenic cotton lines have shown significant
variation in morphological traits and less significant variation among physiological
parameters when compared with non-transgenic cotton plants.
4.10.1. Plant Yield
Improvement of cotton yield was the ultimate goal of this study by acquiring insect
resistance against major cotton pests. Average cotton yield (seed and lint) was calculated in
triplicates from each transgenic cotton line in T1 progeny after harvesting the cotton plants
from mature opened bolls. Maximum cotton yield (129 g) was observed in transgenic cotton
line L6P3 that is statistically significant at 95% confidence interval or probability value of P ≤
0.001 as compared to non-transgenic control plant; total yield in transgenic cotton lines L3P2
and L5P3 (99 g and 97 g respectivily) was significantly higher than non-transgenic control
plants at P ≤ 0.01; similarly, the yield in transgenic cotton line L4P4 (92 g) was significantly
higher than non-transgenic control plants at probability level P ≤ 0.05. No significant
79
difference in yield was recorded between L34P2 and non-transgenic control as shown in
Figure 4.17 (A).
4.10.2. Plant Height
Maximum plant height was observed in transgenic lines L6P3 and L3P2 (96 inches and
94 inches respectively) that is significantly higher than non-transgenic control at 99%
confidence interval (P≤0.01). While the height of transgenic line L34P2 was found to be (52
inches) which was also significantly higher than non-transgenic control at 95% confidence
interval (P≤0.05). Transgenic lines L4P4 and L5P3 have shown non-significant variation in
plant height as compared to non-transgenic control (Figure 4.17 B).
4.10.3. Number of Bolls per Plant
The maximum number of bolls per plant (108) was observed in transgenic line L6P3
that is almost double as compared to non-transgenic control (58). Statistical data showed that
the increase in number of bolls in this transgenic line is significantly higher than non-
transgenic control cotton line at 99% confidence interval (P ≤ 0.01). Similarly, the transgenic
line L3P2 showed increased number of bolls that was significant at 95% confidence interval
(P ≤ 0.05). Transgenic lines L4P4; L5P3 and L34P2 showed non-significant difference in
number of bolls as compared to non-transgenic control (Figure 4.17 C).
4.10.4. Number of Monopodial and Sympodial Branches per Plant
A number of monopodial (black bars) and sympodial branches (colored bars) are
graphically represented in the figure 4.17 D. The number of monopodial branches per plant
ranged from 4-8 in transgenic lines as compared to 7 in non-transgenic control. The number
of sympodial branches has been found to increase up to 16 in transgenic lines as compared to
80
12 in non-transgenic cotton control. An ordinary two-way ANOVA was applied to this
grouped data. Statistical analysis showed that the interaction between transgenic lines and
number of branches per plant did not show any significant difference from control. On the
other hand, the number of branches per plant have been found to be significantly variable
among each other at 99.9% confidence interval (P ≤0.001) as depicted in Figure 4.17 D.
4.10.5. Ginning Out Turn Percentage (GOT %)
The lint and seeds were separated by a ginning machine and ginning out turn
percentage (GOT %) was calculated by dividing the weight of lint over the weight of seeds
multiplied by 100 for all the transgenic cotton lines. The GOT percentage in transgenic line
L6P3 was significantly higher than non-transgenic cotton plants at 99% confidence interval (P
≤ 0.01). All other transgenic cotton lines were found to be non-significantly different when
compared with non-transgenic control plants as evident from figure 4.17 E.
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Figure 4.17: Comparison of morphological characteristics between transgenic and non-transgenic lines
A) Plant Yield; B) Plant Height; C) Number of Bolls per Plants; D) Number of Branches per Plant; E) Ginning
out Turn. Each bar in graphs represents mean of three plants (n=3). Asterisk (*) shows significant variation at
95% confidence interval (P ≤0.05); **Significant at 99% confidence interval (P ≤ 0.01); ***Significant at
99.9% confidence interval (P ≤0.001); ns= Non-Significant (P>0.05).
82
4.10.6. Scanning Electron Microscopic Analysis of Cotton Fiber
The scanning electron microscopic (SEM) images showed that the fiber samples from
both, the transgenic and non-transgenic control cotton plants, were flat with a twisted ribbon-
like structure and natural folds running parallel along the length of cotton fibre. There was no
visible difference in fiber morphology (fineness, smoothness, number of twists and presence
of ribbon like structures) was seen between transgenic and non-transgenic control cotton
plants which depicts that there is no positive or negative correlation exists between transgene
expression and cotton fibre quality as evident from Figure 4.18.
Figure 4.18: Scanning Electron Microscopic analysis of fiber surface
View of cotton fiber at different magnifications (400X, 1000X and 4000X)
Non-transgenic control (A, B, C) Transgenic cotton plant (D, E, F)
A B C
D E F
83
4.10.7. Photosynthetic Rate
Net photosynthetic activity in fully expanded cotton leaves of transgenic and non-
transgenic control lines was observed through portable infrared gas analyzer (IRGA).
Statistical analysis showed that only the transgenic line L6P3 was found to have significantly
higher photosynthetic rate as compared to non-transgenic control plants at 95% confidence
interval (P ≤0.05), while no significant difference in photosynthetic rate was observed among
other transgenic lines with respect to non-transgenic cotton plants as evident from Figure
4.19 A.
4.10.8. Transpiration Rate
Figure 4.19 B represents the transpiration rate of transgenic cotton lines in
comparison to non-transgenic control line. Only the transgenic line L4P4 has been found to
show significantly higher transpiration rate as compared to control non-transgenic cotton
plants at 95% confidence interval (P ≤ 0.05).
4.10.9. Gaseous Exchange Rate
Rate of gaseous exchange was also calculated in transgenic and non-transgenic
control lines and graphically represented in figure 4.19 C. Statistical data showed that the
gaseous exchange rate of transgenic cotton lines L3P2 and L34P2 was significantly higher than
non-transgenic control cotton plants at confidence interval of 99% (P ≤ 0.01). While the
gaseous exchange rate of transgenic cotton line L5P3 was found to be significantly higher
than control at 95% confidence interval (P ≤ 0.05). No significant increase in gaseous
exchange rate was seen in transgenic cotton lines (L4P4 and L6P3) when compared with non-
transgenic cotton plants.
84
Figure 4.19: Comparison of physiological parameters between transgenic and non-transgenic cotton
lines.
A) Net Photosynthetic Rate; B) Transpiration Rate; C) Gaseous Exchange Rate. Each bar in graphs represents
mean of three plants (n=3). Asterisk (*) shows significant variation at 95% confidence interval (P ≤0.05);
**Significant at 99% confidence interval (P ≤ 0.01); ***Significant at 99.9% confidence interval (P ≤0.001);
ns= Non-Significant (P>0.05).
85
4.11. Insect Bioassays
4.11.1. Laboratory Bioassay with Cotton Bollworm (Helicoverpa armigera)
Transgenic cotton lines expressing Vip3Aa+ASAL insecticidal proteins were
evaluated for their toxicity to cotton bollworm larvae in T2 generation by standard laboratory
leaf detach bioassay. The mortality data was recorded after three days of infestation. The
results showed that the transgenic cotton plants were highly resistant to H. armigera as
compared to non-transgenic control. All the leaves of non-transgenic control plants had been
consumed, while the transgenic leaves were barely fed. Similarly, the larvae feeding on non-
transgenic control leaves were significantly larger and developed into next instar. Besides the
minor damage to the leaves of transgenic cotton plants, significant delay in the larval growth
was observed. All the larvae that survived after the third day of bioassay were extremely
weak as compared to the larvae fed on non-transgenic cotton plants as shown in figure 4.20.
In terms of mortality, transgenic cotton lines L6P3B; L6P3C and L34P2 showed 89%; 100%
and 89% mortality respectivily and were significantly resistant against H. armigera at
99.99% confidence interval (P≤0.0001) as compared to non-transgenic control cotton line.
Similarly, the transgenic cotton lines L3P2 and L5P3 showed 78% mortality against tested
insect and were resistant at 99.9% confidence interval (P≤0.001) when compared with non-
transgenic control (figure 4.21).
86
Figure 4.20: Leaf-detach bioassays of transgenic cotton lines with Cotton Bollworm (Helicoverpa
armigera).
(A) Leaf damage in non-transgenic control cotton plants. (B-F) Leaf damage in transgenic cotton lines
87
Figure 4.21: Mean percent mortality of Cotton Bollworm (Helicoverpa armigera) larvae
****Significant at 99.99% confidence interval (P ≤0.0001)
***Significant at 99.9% confidence interval (P ≤0.001)
4.11.2. Laboratory Bioassay with Whitefly (Bamisia tabaci)
After testing the transgenic cotton plants against cotton bollworm the same plants
were also exposed to whitefly feeding in specialized clip cages as shown in Figure 4.22 (A).
After 24 hours of feeding on transgenic plants the insects started showing notable behavior
changes like abnormal stretching of the body (Figure 4.22 B). The insects on non-transgenic
control cotton plants remained alive and active as depicted in figure 4.22 (C). The mortality
data was collected after three days of infestation. Transgenic cotton lines L3P2, L5P3, L6P3B
and L6P3C showed 95%; 89%, 89% and 72% mortality respectivily. The asterisks (****) and
(***) above the bars highlight the transgenic cotton lines with significant statistical
difference [ANOVA, 99.99% confidence interval (P ≤0.0001) and (P≤0.001)] respectivily
using the non-transgenic cotton line as the control (Figure 4.23).
88
Figure 4.22: Pictures of bioassay performed with Bamisia tabaci
(A) Transgenic plant with clip cages (B) Dead whiteflies on transgenic leaf (C) Whitefly feeding on non-
transgenic control leaf
Figure 4.23: Mean percent mortality of whitefly (Bamisia tabaci)
****Significant at 99.99% confidence interval (P ≤0.0001)
***Significant at 99.9% confidence interval (P ≤0.001)
A B C
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4.12. Determination of Transgene Copy Number and Location
One plant from each transgenic cotton line in T2 generation was subjected for
determination of copy number and transgene location at different stages of cell division;
prophase; metaphase and interphase using ASAL specific probe. The transgenic cotton plant
from line L3P2 showed homozygosity at telophase stage, showing two copy numbers at
chromosome number 9 and one copy number at chromosome number 10 at prophase stage
whereas, no signal was observed in non-transgenic control cotton plant as shown in figure
4.22 (A & B).
Figure 4.24: Determination of copy number and transgene location through FISH
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5. DISCUSSION
Cotton is the most important cash crop of Pakistan and contributes substantially to the
national economy (Akhtar et al. 2018). The average cotton yield in Pakistan is considerably
low as compared to other cotton growing countries (Basit 2018). Various abiotic factors like
drought, salinity and biotic stress factors like insects, weeds and virus attack are involved in
low cotton yield and poor fiber quality. It is estimated that sucking and chewing insects
collectively cause almost 50-60% yield losses. Despite the fact that conventional cotton has
been replaced with genetically modified Bt cotton, expressing insecticidal toxins from
Bacillus thuringiensis, that has effectively controlled major chewing insects. There are still
some concerns over the narrow insect spectrum and emergence of resistance build up in most
of the insect species. Therefore, it is the need of time to characterize and transform novel
insecticidal toxins with broad insect spectra (Bravo et al. 2017). The vegetative insecticidal
protein 3Aa (Vip3Aa) from Bacillus thuringiensis could be a possible alternate to overcome
the limitations of first-generation Bt toxins because it shares no sequence similarity with any
known crystal δ-endotoxin (Estruch et al. 1996) and displays a wider insecticidal spectrum
against a broad range of chewing insects (Song et al. 2016).
Sucking insects are another notorious group of insects that contribute to significant
cotton crop yield losses and are not controlled by Bt endotoxins (Sharma et al. 2017). The
carbohydrate binding plant lectins infer toxicity to agronomically important sap-sucking
insects by specifically binding with glycoproteins in the insect epithelial gut receptors. Lectin
gene isolated from Allium sativum (ASAL) has already been reported to be a successful
remedy against phytophagous insect pests in crops like tobacco, rice and mustard (Ahmed et
al. 2017; Chandrasekhar et al. 2014; Rani et al. 2017; Vajhala et al. 2013).
91
In the current study, an effort was made to develop transgenic cotton plants through
the introduction of Bt Vip3Aa and Allium sativum leaf agglutinin (ASAL) genes to minimize
losses caused by the infestation of major sucking and chewing insect pests. Full-length
nucleotide sequences of Vip3Aa (2370 bp) and ASAL (339 bp) genes were retrieved from
GeneBank and their codon usage was optimized through freely available web-based tools to
get maximum gene expression in cotton (Gossypium hirsutum). It is apparent from the
literature that prokaryotic genes are rich in AT bases as compared to eukaryotes that are rich
in GC bases. Transformation of prokaryotic genes into eukaryotic expression system leads to
low expression, possibly due to premature or aberrant mRNA splicing or mRNA instability
(Jabeen et al. 2010).
The constitutive promoters express in almost all cell types of the plant and during all
developmental stages, however, the low expression is reported in the phloem cells (Obertello
et al. 2005). As sucking insects feed exclusively on phloem sap, these insects can be
effectively controlled by expressing insecticidal genes under phloem-specific promoters. The
codon-optimized Vip3Aa and ASAL genes under a constitutive promoter (CaMV35S) and
phloem-specific promoter from Rice tungro bacilliform virus (RTBV) respectively were
chemically synthesized in a single 4870 bp (Vip3Aa+ASAL) cassette by Bio Basic Inc.
Canada. A similar approach was used by (Javaid et al. 2018) where they translationally fused
two insecticidal toxins (ω-atracotoxin from Hadronyche versuta and Allium sativum leaf
lectin) and evaluated the toxicity on Phenacoccus solenopsis (mealybug). In another study,
(Wu et al. 2011) developed transgenic cotton lines by transforming two modified Vip3A
genes (codon optimized vip3A* and chloroplast transit peptide added tvip3A* gene) through
Agrobacterium mediated transformation system and compared the insecticidal activity of
92
both the genes against cotton bollworm. The results indicated that the expression level of
Vip3A protein was three-fold higher in transgenic lines tvip3A* and showed 100% mortality
of cotton bollworm.
The cloning and orientation of synthetic gene cassette (Vip3Aa+ASAL) in binary
plant expression vector pCAMBIA 1301 was verified through polymerase chain reaction and
restriction digestion as shown in figures 4.3 and 4.4. The recombinant vector
(pCAMBIA1301_Vip3Aa+ASAL) was then successfully electroporated in Agrobacterium
tumefaciens and confirmed through PCR and restriction analysis as depicted in figures 4.5
and 4.6. (Muzaffar et al. 2015) used similar cloning procedure to localize the expression of
Cry1Ac and Cry2A proteins in the chloroplast.
The recombinant vector was transformed into a locally developed cotton variety
(CEMB-33) through shoot apex cut method of Agrobacterium mediated transformation.
Shoot apical meristems isolated from 4500 mature cotton seedlings as explants were co-
cultivated with Agrobacterium tumefaciens strain LBA 4404 harboring synthetic gene
cassette. A total of fifty-three putative transgenic cotton plants were obtained and shifted to
the field after initial screening through the expression of GUS marker gene (Figure 4.7). The
overall transformation efficiency remained at 1.17%. The similar approach was also used by
(Bajwa et al. 2015; Lei et al. 2012; Puspito et al. 2015; Rao et al. 2011b) in cotton and got
transformation efficiencies in the range of 1-1.2%.
The successful transformation in putative transgenic cotton plants was confirmed
through different molecular techniques. The amplification of 587 bp fragment of ASAL gene
and 682 bp fragment of Vip3Aa gene in eighteen putative transgenic cotton plants namely
(L1P3; L1P5; L2P3; L3P2; L4P2; L4P4; L5P2; L5P3; L6P2; L6P3; L34P1; L34P2; L34P4; L35P1; L35P2;
93
L35P3; L36P1; L38P4) out of total fifty three plants that were shifted to field, using gene
specific internal primers, confirmed the successful introduction of VIP3A and ASAL genes
in cotton (Figure 4.8). The results were in correlation with (Bajwa et al. 2013). The quantity
of Vip3Aa protein in transgenic cotton plants was determined through quantitative ELISA
test. The maximum concentration of Vip3Aa protein (4.26 µg/mL) was observed in plant
L6P3 and minimum protein concentration was seen in plant L34P1 when compared with
negative control (non-transgenic cotton plant) where no expression of Vip3A was seen as
depicted in figure 4.10. The results are in accordance with (Wu et al. 2011) where they
reported three-fold higher expression of chimeric Vip3Aa protein in the chloroplast of
transgenic cotton lines as compared to control. The expression of GUS marker gene was also
obtained through histochemical GUS assay. The appearance of blue color in transgenic
cotton leaf (Figure 4.11) in comparison to non-transgenic control confirmed the successful
expression of gene cassette in plant genome as previously done by (Bakhsh et al. 2012).
The mRNA expression of Vip3Aa and ASAL insecticidal genes was taken through
quantitative real-time qRT-PCR in five T1 transgenic cotton lines (L3P2; L4P4; L5P3; L6P3;
L34P2). The comparative analysis of Ct values obtained by qRT-PCR analysis revealed that
the mRNA expression of Vip3Aa gene varied from 2-8.7 folds in the lines L3P2 and L6P3
respectively. Similarly, the comparison of Ct values showed that the mRNA expression of
ASAL gene ranged between 2-5 folds in transgenic cotton lines L4P4 and L34P2. The
transgenic line L6P3 showed significantly higher expression for both the genes as depicted in
Figure 4.13 (A & B). The results were in consistent with (Wu et al. 2011) in which the
authors produced transgenic cotton plants with synthetic Vip3A gene fused with chloroplast
94
transit peptide and achieved three-fold increase in expression level of Vip3A and hence,
exhibited high entomotoxic activity against wide range of chewing insects.
Fluorescent in situ hybridization (FISH) was preferred than Southern hybridization in
determining the transgene location and copy number in the cotton genome due to its
reliability and accuracy (Tsuchiya and Taga 2001). The transgene expression is directly
related to different factors like promoter, insertion point of genes, copy number and location
on host chromosome (Rao et al. 2011a). One plant from each transgenic cotton line in T2
generation was subjected for FISH at different stages of cell division using ASAL specific
probe following the same procedure as explained by (Rao et al. 2013). The transgenic cotton
plant from transgenic line L3P2 showed homozygosity (two copy numbers) on chromosome
number 9 at late telophase stage and one copy number at chromosome number 10 at prophase
stage whereas, no signal was observed in the non-transgenic control cotton plant as shown in
figure 4.22 I&II. The results are in accordance with (Ali et al. 2016) who evaluated two
cotton varieties (CRSP-I and CRSP-II) for the expression and location of Cry1Ac+Cry2A,
GTG-gene and compared the genes expression level and resistance against insects relative to
copy number of genes.
Different morphological characteristics (plant height; number of bolls per plant; number
of monopodial and sympodial branches per plant; overall yield per plant (lint + seed);
ginning out turn percentage (GOT%); scanning electron microscopic analysis of fibre) and
physiological parameters (photosynthetic rate; transpiration rate; gaseous exchange rate) of
transgenic cotton lines in T1 progeny were studied in comparison to non-transgenic (negative
control) plants (Figures 4.17 A-E; 4.18; 4.19 A-C). The results indicated that different
transgenic lines showed significant variation in studied morphological characteristics as
95
previously published by (Raybould and Vlachos 2011; Romeis et al. 2008) who claimed that
statistically significant differences were observed between transgenic cotton plants
expressing Vip3Aa in comparison to non-transgenic controls. On the basis of the phenotypic
data, the authors also negated the hypothesis that the introduction of the Vip3Aa protein had
any unintended impact on the plant morphology or phenotype, besides conferring insect
resistance to lepidopteran pests. The scanning electron microscopic (SEM) images of
transgenic and non-transgenic cotton plants showed flat, twisted ribbon-like structure and
natural folds running parallel along the length of the cotton fiber. There was no visible
difference in fiber morphology (fineness, smoothness, number of twists and presence of
ribbon-like structures) between transgenic and non-transgenic control cotton plants which
depicts no positive or negative correlation between expression of insecticidal genes
(Vip3Aa+ASAL) and cotton fiber quality. Similar findings reported by (Raybould and
Vlachos 2011) also confirmed our results that expression of Vip3Aa gene does not have any
unintended impact on overall morphology and phenotypic characteristics of the plant.
The final objective of the current study was to evaluate transgenic cotton plants
expressing Bt Vip3Aa gene under a constitutive promoter (CaMV35S) and Allium sativum
leaf agglutinin (ASAL) gene under RTBV phloem-specific promoter against cotton bollworm
(H. armigera) and whitefly (Bamisia tabaci) in advanced generation like T2. The confirmed
transgenic cotton plants showing higher expression of Vip3Aa and ASAL in T2 generation
were selected for insect leaf detached bioassay. Almost 78% to 100% mortality of H.
armigera as compared to non-transgenic control, confirmed significant resistance
development in transgenic cotton plants against H. armigera expressing Vip3A toxin protein
(Figures 4.20-I A-F & 4.20-II). Besides the minor damage to the leaves of transgenic cotton
96
line L5P3, all other lines remained undamaged. All the larvae that survived after the third day
of bioassay were extremely weak and a significant delay in the larval growth was observed as
compared to the larvae fed on non-transgenic cotton plants, the results were in complete
accordance with (de Oliveira et al. 2016). The transgenic cotton lines L3P2, L5P3, L6P3B and
L6P3C showed 95%; 89%, 89%, and 72% mortality rates of whitefly in three days clip
bioassay in controlled environment grown transgenic cotton plants as compared to non-
transgenic control plants where the mortality was 10% which can be due to some natural
stress. Similarly, all the dead insects showed notable physical changes like the abnormal
stretching of the body (Figures 4.21-I A-C and 4.21-II) likely due to the expression of lectin
gene that targets the digestive tract of the insect (Javaid et al. 2016).
Conclusion
The current research work was proposed to utilize and evaluate the insecticidal
spectrum of Bt Vip3Aa and garlic lectin ASAL genes in the cotton plant to enrich the
insecticidal potential of the transgenic cotton plant against major chewing and sucking
insects. Insect bioassays using viruliferous whiteflies and 2nd
instar larvae of cotton
bollworms confirmed the efficacy of transgenes. Significant mortality of sucking and
chewing insects on transgenic cotton plants tells the success story of this study. The study is
a part of future strategy to overcome resistance build-up in insects due to extensive use of Bt
toxins against specific receptors. By using next generation Bt toxins having different target
receptors will be helpful in delaying resistance and making longer and stable insect control
which will impact on national economy and farmers.
97
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7. APPENDICES
Appendix I
Luria Bertani (LB) Medium (1 L)
Ingredients Quantity pH
Tryptone 5 g 7.5
Yeast extract 10 g
NaCl 10 g
Agar 15 g
Luria Bertani (LB) Broth (1 L)
Ingredients Quantity pH
Tryptone 5 g 7.5
Yeast extract 10 g
NaCl 10 g
Yeast Extract Peptone (YEP) Medium (1 L)
Ingredients Quantity pH
Yeast extract 10 g 7.5
Peptone 10 g
NaCl 5 g
Agar 15 g
Yeast Extract Peptone (YEP) Broth (1 L)
Ingredients Quantity pH
Yeast extract 10 g 7.5
Peptone 10 g
NaCl 5 g
124
Murashige and Skoog (MS) Medium (1 L)
Ingredients Quantity pH
MS Basal Salt 4.43 g 5.8
Sucrose 30 g
Phytagel 3 g
Murashige and Skoog (MS) Broth (1 L)
Ingredients Quantity pH
MS Basal Salt 4.43 g 5.8
Sucrose 30 g
B5 Vitamin Complex (50 mL)
Ingredients Quantity
10 mM Nicotinic Acid 40.6 µL
10 mM Pyridoxine HCL 24.3 µL
10 mM Thymine HCL 148.25 µL
10 mM Myoinositol 280 µL
125
Appendix II
50X Tris-Acetate (TAE) Buffer (pH 8.6)
Ingredients Quantity
Tris Base 242 g
Glacial Acetic Acid 57.1 mL
Disodium EDTA 18.61 g
0.8% Agarose gel
Ingredients Quantity
1X TAE 100 mL
Agarose 0.8 g
Ethidium Bromide
Ingredients Quantity
Ethidium Bromide Tablet 0.1 g
Distilled H2O 100 mL
6X DNA Loading Dye
Ingredients Quantity
Ficoll 20%
EDTA 0.1 M
SDS 1%
Bromophenol Blue 0.25%
Xylene Cyanol 0.25%
126
Appendix III
Rifampicin
Ingredients Quantity
Rifampicin powder 12.5 mg
70% Ethanol 10 mL
Ampicillin
Ingredients Quantity
Ampicillin powder 1 g
Distilled H2O 10 mL
Tetracycline
Ingredients Quantity
Tetracycline powder 12.5 mg
70% Ethanol 10 mL
Kanamycin
Ingredients Quantity
Kanamycin powder 1 g
Distilled H2O 20 mL
Cefotaxime
Ingredients Quantity
Cefotaxime Powder 0.25 g
Distilled H2O 5 mL
127
Appendix IV
CTAB DNA Extraction Buffer
Stock Working Volume (50 mL)
10 % CTAB 2% 10 mL
5M NaCl 1.4 M 14 mL
0.5 M EDTA (pH 8.0) 0.02 M 2 mL
1 M Tris-Cl (pH 8.0) 0.1 M 5 mL
10 % PVP 2 % 10 mL
Autoclaved H2O 8.5 mL
β-Mercepto Ethanol 2% 0.5 mL
RNA Extraction Buffer
Stock Working Volume (50 mL)
10% CTAB 2% 2.5 mL
5M NaCl 1.4 M 10 mL
0.5 M EDTA (pH 8.0) 0.02 M 1.25 mL
1 M Tris-Cl (pH 8.0) 0.1 M 2.5 mL
10% PVP 2% 2.5 mL
Autoclaved H2O 30.25 mL
β-Mercepto Ethanol 2% 1 mL
SSTE Buffer
Stock Working Volume (50 mL)
5% SDS 0.5% 1 mL
1.5 M NaCl 1 M 2.85 mL
0.5 M EDTA (pH 8.0) 1 mM 20 µL
1M Tris-Cl (pH 8.0) 10 mM 100 µL
128
Protein Extraction Buffer
Stock Volume (45 mL)
5M NaCl 900 µL
0.5 M EDTA 900 µL
1M Sucrose 9 mL
10% SDS 450 µL
1M Tris (pH 8.0) 9 mL
100 mM PMSF 900 µL
MDTT 4.5 mL
Tween 20 3 µL
Distilled H2O 19.275 mL
129
Appendix V
20X SSC Stock, pH 7.0 (1L)
NaCl 175 g
Trisodium citrate hydrate 88.2 g
Double distilled H2O 800 mL
2X SSC Solution
20X SSC 28.5 mL
Double distilled H2O 256.5 mL
4X SSC/0.1% Tween 20 (70 mL)
20X SSC 14 mL
Double distilled H2O 56 mL
Tween 20 70 µL
RNase A Digestion Solution
1 mg/mL RNase 60 µL
20X SSC 60 µL
Double distilled H2O 480 µL
1X TBS Buffer, pH 7.5 (1 L)
Tris base 6.5 g
NaCl 8.76 g
Double distilled H2O 800 mL
Denaturing Solution (70 mL)
Deionized Formamide 49 mL
20X SSC 7 mL
Double distilled H2O 1 mL
pH adjusted to 7.0-7.5 with 1M HCL and volume raised up to 70 mL
130
1X PBS Buffer, pH 7.4 (1 L)
NaCl 8 g
KCl 0.2 g
Na2HPO4 1.44 g
KH2PO4 0.24 g
Double distilled H2O 800 mL
Mellavaine Buffer, pH 7.0 (20 mL)
0.2 M Na2HPO4 16.47 mL
0.1 M Citric Acid 3.53 mL
Hybridization Solution
Reagents Quantity
dFA (50-70%) 60 µL
20X SSC 12 µL
Dextran Sulphate (50%) 24 µL
Carrier DNA (10µg/µl) 4 µL
Probe DNA (20µg/ µL) 16 µL
H2O 24 µl
Total Volume 140 µL
131
Appendix VI
GUS Solution (10 mL)
Ingredients Quantity
X-Gluc 0.008 g
Chloramphenicol 0.001 g
1M Na2HPO4 1 mL
10% Triton X-100 200 µL
Methanol 2 mL
1M HEPES, pH 7.4 (1L)
Ingredients Quantity
HEPES 238.30 g
Distilled H2O 750 mL
10% Glycerol
Ingredients Quantity
99.9% Glycerol 10 mL
Distilled H2O 90 mL
Kinetin (1mg/mL)
Ingredients Quantity
Kinetin Powder 1 g
1 N KOH 2 mL
Raise volume up to 1 liter to make the final concentration of 1mg/mL
Indole Butyric Acid (IBA) 1mg/mL
Ingredients Quantity
IBA powder 1 g
1N NaOH 5 mL
Raise volume up to 100 mL and filter through 0.2 µm syringe filter
132
1M CaCl2 (1L)
Ingredients Quantity
CaCl2.2H2O 147 g
Distilled H2O 1000 mL
133
8. LIST OF PUBLICATIONS
1. Ahmed, M., Shahid, A.A., Akhtar, S., Latif, A., Din, S., Fanglu, M., Rao, A.Q.,
Sarwar, M.B., Husnain, T. and Xuede, W. (2018). Sucrose synthase genes: a way
forward for cotton fiber improvement. Biologia. doi.org/10.2478/s11756-018-
0078-6.
2. Ahmed, M., Shahid, A.A., Din, S., Akhtar, S., Ahad, A., Rao, A.Q., Bajwa, K.S.,
Khan, M.A., Sarwar, M.B. and Husnain, T. (2018). An overview of genetic and
hormonal control of cotton fiber development. Pakistan Journal Botany. 50 (1):
433-443.
3. Palma, L., Scott, D.J., Harris, G., Din, S., Williams, T.L., Roberts, O.J., Young, M.T.,
Caballero, P. and Berry, C. (2017). The Vip3Ag4 insecticidal protoxin from
Bacillus thuringiensis adopts a tetrameric configuration that is maintained on
proteolysis. Toxins 9: 165.
4. Rao, A.Q., Latif, A., Azam, S., Gul, A., Yaqoob, A., Din, S., Akhtar, S., Shahid, N.,
Shahid, A.A., Nasir, I.A. and Husnain, T. (2017). Management of biotic stress in
cotton. International Cotton Advisory Committee (ICAC) Recorder 35(4): 19-26.
5. Ahmad, A., Javed, M.R., Rao, A.Q., Khan, M.A., Ahad, A., Din, S., Shahid, A.A. and
Husnain, T. (2015). In-Silico determination of insecticidal potential of Vip3Aa-
Cry1Ac fusion protein against lepidopteran targets using molecular docking.
Frontier in Plant Science 6: 1081.
6. Ahad, A., Ahmad, A., Din, S., Rao, A.Q., Shahid, A.A. and Husnain, T. (2015). In
Silico study for diversing the molecular pathway of pigment formation: an
alternative to manual coloring in cotton fibers. Frontier in Plant Science 6: 751.
7. Rao, A.Q., Khan, M.A., Shahid, N., Din, S., Gul, A., Muzaffar, A., Azam, S.,
Samiullah, T.R., Batool, F., Shahid, A.A., Nasir, I.A. and Husnain, T. (2015). An
overview of Phytochrome: An important light switch and photo-sensory antenna
for regulation of vital functioning of plants. Biologia 70 (10): 1273-1283.
BOOK CHAPTER PUBLISHED
o Rao, A.Q., Din, S., Akhtar, S., Sarwar, M.B., Ahmed, M., Rashid, B., Khan, M.A.,
Qaisar, U., Shahid, A.A., Nasir, I.A. and Husnain, T. (2016). Genomics of Salinity
Tolerance in Plants. In: Plant Genomics. (pp. 273-299) Intech publishers.
