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Transcript of Optimisation of Banana streak virus (BSV) diagnostic...
Optimisation of Banana streak virus (BSV) diagnostic assays
by
Pål Kristian Berntzen Bjartan
Bachelor of Chemical Engineering
Graduate Certificate of Biotechnology
A thesis submitted for the degree of
Master of Applied Science
at the
Queensland University of Technology
Centre for Tropical Crops and Biocommodities
Science and Engineering Faculty
2012
i
Abstract Bananas are one of the world’s most important crops, serving as a staple
food and an important source of income for millions of people in the
subtropics. Pests and diseases are a major constraint to banana production.
To prevent the spread of pests and disease, farmers are encouraged to use
disease‐ and insect‐free planting material obtained by micropropagation.
This option, however, does not always exclude viruses and concern remains
on the quality of planting material. Therefore, there is a demand for
effective and reliable virus indexing procedures for tissue culture (TC)
material.
Reliable diagnostic tests are currently available for all of the
economically important viruses of bananas with the exception of Banana
streak viruses (BSV, Caulimoviridae, Badnavirus). Development of a reliable
diagnostic test for BSV is complicated by the significant serological and
genetic variation reported for BSV isolates, and the presence of
endogenous BSV (eBSV). Current PCR‐ and serological‐based diagnostic
methods for BSV may not detect all species of BSV, and PCR‐based methods
may give false positives because of the presence of eBSV. Rolling circle
amplification (RCA) has been reported as a technique to detect BSV which
can also discriminate between episomal and endogenous BSV sequences.
However, the method is too expensive for large scale screening of samples
in developing countries, and little information is available regarding its
sensitivity. Therefore the development of reliable PCR‐based assays is still
considered the most appropriate option for large scale screening of banana
plants for BSV. This MSc project aimed to refine and optimise the protocols
for BSV detection, with a particular focus on developing reliable PCR‐based
diagnostics
Initially, the appropriateness and reliability of PCR and RCA as
diagnostic tests for BSV detection were assessed by testing 45 field samples
ii
of banana collected from nine districts in the Eastern region of Uganda in
February 2010. This research was also aimed at investigating the diversity
of BSV in eastern Uganda, identifying the BSV species present and
characterising any new BSV species. Out of the 45 samples tested, 38 and
40 samples were considered positive by PCR and RCA, respectively. Six
different species of BSV, namely Banana streak IM virus (BSIMV), Banana
streak MY virus (BSMYV), Banana streak OL virus (BSOLV), Banana streak
UA virus (BSUAV), Banana streak UL virus (BSULV), Banana streak UM virus
(BSUMV), were detected by PCR and confirmed by RCA and sequencing. No
new species were detected, but this was the first report of BSMYV in
Uganda. Although RCA was demonstrated to be suitable for broad‐range
detection of BSV, it proved time‐consuming and laborious for identification
in field samples.
Due to the disadvantages associated with RCA, attempts were
made to develop a reliable PCR‐based assay for the specific detection of
episomal BSOLV, Banana streak GF virus (BSGFV), BSMYV and BSIMV. For
BSOLV and BSGFV, the integrated sequences exist in rearranged, repeated
and partially inverted portions at their site of integration. Therefore, for
these two viruses, primers sets were designed by mapping previously
published sequences of their endogenous counterparts onto published
sequences of the episomal genomes. For BSOLV, two primer sets were
designed while, for BSGFV, a single primer set was designed. The episomal‐
specificity of these primer sets was assessed by testing 106 plant samples
collected during surveys in Kenya and Uganda, and 33 leaf samples from a
wide range of banana cultivars maintained in TC at the Maroochy Research
Station of the Department of Employment, Economic Development and
Innovation (DEEDI), Queensland. All of these samples had previously been
tested for episomal BSV by RCA and for both BSOLV and BSGFV by PCR
using published primer sets. The outcome from these analyses was that the
newly designed primer sets for BSOLV and BSGFV were able to distinguish
iii
between episomal BSV and eBSV in most cultivars with some B‐genome
component. In some samples, however, amplification was observed using
the putative episomal‐specific primer sets where episomal BSV was not
identified using RCA. This may reflect a difference in the sensitivity of PCR
compared to RCA, or possibly the presence of an eBSV sequence of
different conformation.
Since the sequences of the respective eBSV for BSMYV and BSIMV in
the M. balbisiana genome are not available, a series of random primer
combinations were tested in an attempt to find potential episomal‐specific
primer sets for BSMYV and BSIMV. Of an initial 20 primer combinations
screened for BSMYV detection on a small number of control samples, 11
primers sets appeared to be episomal‐specific. However, subsequent
testing of two of these primer combinations on a larger number of control
samples resulted in some inconsistent results which will require further
investigation. Testing of the 25 primer combinations for episomal‐specific
detection of BSIMV on a number of control samples showed that none
were able to discriminate between episomal and endogenous BSIMV.
The final component of this research project was the development of
an infectious clone of a BSV endemic in Australia, namely BSMYV. This was
considered important to enable the generation of large amounts of
diseased plant material needed for further research. A terminally
redundant fragment (1.3 × BSMYV genome) was cloned and transformed
into Agrobacterium tumefaciens strain AGL1, and used to inoculate 12
healthy banana plants of the cultivars Cavendish (Williams) by three
different methods. At 12 weeks post‐inoculation, (i) four of the five banana
plants inoculated by corm injection showed characteristic BSV symptoms
while the remaining plant was wilting/dying, (ii) three of the five banana
plants inoculated by needle‐pricking of the stem showed BSV symptoms,
one plant was symptomless while the remaining had died and (iii) both
banana plants inoculated by leaf infiltration were symptomless. When
iv
banana leaf samples were tested for BSMYV by PCR and RCA, BSMYV was
confirmed in all banana plants showing symptoms including those were
wilting and/or dying.
The results from this research have provided several avenues for
further research. By completely sequencing all variants of eBSOLV and
eBSGFV and fully sequencing the eBSIMV and eBSMYV regions, episomal
BSV‐specific primer sets for all eBSVs could potentially be designed that
could avoid all integrants of that particular BSV species. Furthermore, the
development of an infectious BSV clone will enable large numbers of BSV‐
infected plants to be generated for the further testing of the sensitivity of
RCA compared to other more established assays such as PCR. The
development of infectious clones also opens the possibility for virus‐
induced gene silencing studies in banana.
v
Table of Contents
Chapter 1 Introduction ......................................................................... 1
1.1 Bananas ............................................................................................ 1
1.2 Pests and diseases affecting bananas .............................................. 3
1.2.1 Pests ......................................................................................... 3
1.2.2 Diseases ................................................................................... 5
1.2.3 BSV diagnostics ...................................................................... 14
1.2.4 BSV diagnostics – issues and possible solutions .................... 16
1.3 The situation in Uganda ................................................................. 18
1.4 Project aim and objectives ............................................................. 19
Chapter 2 General materials and methods ......................................... 21
2.1 General materials ........................................................................... 21
2.1.1 Sources for specialised reagents ............................................ 21
2.1.2 Reagents and equipment for DNA amplification ................... 21
2.1.3 Oligonucleotide synthesis ...................................................... 21
2.1.4 Bacterial strains ..................................................................... 22
2.2 General methods ............................................................................ 22
2.2.1 DNA extraction from leaf tissue............................................. 22
2.2.2 PCR amplification ................................................................... 23
2.2.3 RCA ......................................................................................... 23
2.2.4 Restriction digestion .............................................................. 23
2.2.5 Agarose gel electrophoresis ................................................... 24
2.2.6 DNA extraction from agarose gel ........................................... 24
2.2.7 Dephosphorylation of 5’ ends ............................................... 24
2.2.8 DNA ligation ........................................................................... 24
2.2.9 Transformation of E. coli ........................................................ 24
2.2.10 Mini‐preparation protocol ..................................................... 25
2.2.11 Sequencing ............................................................................. 25
Chapter 3 Assessment of PCR and RCA for diagnosis of BSV in field samples..................................................................................................27
3.1 Introduction .................................................................................... 27
3.2 Methods ......................................................................................... 29
3.2.1 Samples for analysis ............................................................... 29
vi
3.2.2 PCR amplification ................................................................... 29
3.2.3 RCA amplification .................................................................. 29
3.2.4 Cloning and sequence analysis .............................................. 33
3.3 Results ............................................................................................ 35
3.3.1 Sample collection ................................................................... 35
3.3.2 PCR results ............................................................................. 35
3.3.3 RCA results ............................................................................. 40
3.4 Discussion ....................................................................................... 47
Chapter 4 Development of episomal‐specific PCR assays for the detection of banana streak viruses with endogenous counterparts ........ 51
4.1 Introduction .................................................................................... 51
4.2 Methods ......................................................................................... 53
4.2.1 Developing episomal‐specific primers for BSOLV and BSGFV 53
4.2.2 Developing episomal‐specific primers for BSMYV and BSIMV 53
4.2.3 Plant samples and nucleic acid extraction ............................. 55
4.2.4 PCR and RCA analysis ............................................................. 55
4.3 Results ............................................................................................ 59
4.3.1 Identification of potential episomal‐specific primers ........... 59
4.3.2 Primer testing ........................................................................ 65
4.3.3 Final primer assessment using the GMGC collection ............ 70
4.4 Discussion ....................................................................................... 76
Chapter 5 Development of an infectious clone of Banana streak MY virus.......................................................................................................81
5.1 Introduction .................................................................................... 81
5.2 Methods ......................................................................................... 83
5.2.1 Preparation of the infectious clone ....................................... 83
5.2.2 Plant inoculation and assessment ......................................... 86
5.3 Results ............................................................................................ 88
5.3.1 Preparation of the infectious clone ....................................... 88
5.3.2 Plant inoculation and assessment ......................................... 88
5.4 Discussion ..................................................................................... 101
Chapter 6 General discussion and conclusions..................................105
Bibliography ........................................................................................111
vii
List of Figures
Figure 1.1: (a) Map of the BSV genome
(b) Electron micrograph of BSV ................................................. 10
Figure 1.2: Unrooted tree showing the identified BSV species grouped into
three clades ............................................................................... 12
Figure 3.1: Districts in eastern Uganda where leaf samples were collected
..................................................................................................................... 36
Figure 3.2: Symptoms in leaf samples collected in eastern Uganda ........... 37
Figure 4.1: (a) Map of the integrated regions of BSOLV
(b) The integrated sequences were mapped onto the full‐length
genome ................................................................................ 60
Figure 4.2: (a) Map of the integrated regions of BSGFV
(b) The integrated sequences were mapped onto the full‐length
genome ................................................................................ 61
Figure 4.3: Genome of (a) BSMYV and (b) BSIMV with primer sequences
mapped ...................................................................................... 63
Figure 4.4: Screening of the GMGC collection ............................................. 72
Figure 5.1: Creation of the BSMYV infectious clone .................................... 84
Figure 5.2: Restriction digest of plasmid DNA ............................................. 89
Figure 5.3: Healthy banana plant ................................................................. 91
Figure 5.4: Banana plant at 12 weeks post leaf‐infiltration ......................... 92
Figure 5.5: Needle‐prick inoculated banana plant at 12 weeks post
inoculation ................................................................................. 95
Figure 5.6: Banana plant at 12 weeks post corm injection ......................... 96
Figure 5.7: Sugar cane plant at 12 weeks post needle‐prick inoculation .... 97
Figure 5.8: Screening of inoculated banana and sugarcane plants for BSMYV
by PCR (a) and RCA (b) .............................................................. 98
Figure 5.9: Needle‐prick inoculated banana plant seven months post
inoculation ................................................................................. 99
Figure 5.10: Corm injected banana plant seven months post inoculation 100
viii
List of Tables Table 3.1: Plant samples used in this study ................................................. 30
Table 3.2: Primer sequences used for PCR testing ...................................... 32
Table 3.3: Analysis of leaf samples for BSV by PCR and sequencing ........... 38
Table 3.4: Analysis of leaf samples for BSV by RCA and RFLP/sequencing .. 41
Table 4.1: Primers obtained from QUT for detection of BSIMV and BSMYV54
Table 4.2: PCR conditions for published primers used for BSV and eBSV
detection ..................................................................................... 57
Table 4.3: Potential episomal DNA‐specific primer sequences identified for
BSOLV and BSGFV ........................................................................ 62
Table 4.4: Primer combinations and results of initial screen for putative
episomal‐specific primers for BSMYV and BSIMV detection ...... 64
Table 5.1: Analysis of inoculated plants ....................................................... 90
ix
List of Abbreviations Common prefixes
Symbol Prefix Scale
k kilo 103
m milli 10−3
μ micro 10−6
n nano 10−9
p pico 10−12
Abbreviations
approximately
°C degrees Celsius
A genome component of banana derived from the Musa acuminata
species
B genome component of banana derived from the Musa balbisiana
species
BanMMV Banana mild mosaic virus
BBrMV Banana bract mosaic virus
BBTV Banana bunchy top virus
BLAST Basic Local Alignment Search Tool
bp base pair(s)
BS Black sigatoka
BSD banana streak disease
BSV Banana streak virus(es)
BSnV Banana streak n virus, where n = species' name, e.g. BSMYV
BVX Banana virus X
c complementary
CDS coding sequence
Chl:IAA chloroform‐isoamylalcohol
CIP calf intestinal phosphatase
CMV Cucumber mosaic virus
x
CTAB cetyltrimethylammonium bromide
CYMV Citrus yellow mosaic virus
D dalton(s)
DEEDI Maroochy Research Station of the Department of Employment,
Economic Development and Innovation
dH2O deionised water
DNA deoxyribonucleic acid
ds double‐stranded
e endogenous
EAHB East African Highland Banana
EDTA ethylenediaminetetraacetic acid
ELISA enzyme‐linked immunosorbent assay
EPRV endogenous pararetrovirus
g gravity
g gram(s)
gDNA genomic DNA
GM genetically modified
GMGC Global Musa Genomics Consortium
h hour(s)
IC‐PCR immunocapture‐PCR
ICTV International Committee on the Taxonomy of Viruses
IPTG isopropyl β‐D‐1‐thiogalactopyranoside
ISEM immunosorbent electron microscopy
l litre(s)
Ke Kenya (survey sample collection)
LB lysogeny broth (aka. Luria broth, Lennox broth, or Luria‐Bertani
medium)
m metre(s)
M molar concentration, mol/l
MGIS Musa Germplasm Information System
met methionine
xi
min minute(s)
mol mole(s)
MSc Master of Science
NA nucleic acid
ORF open reading frame
PCR polymerase chain reaction
pH ‐ log (proton concentration)
PTGS post‐transcriptional gene silencing
PVP polyvinylpyrrolidone
QUT Queensland University of Technology
RCA rolling circle amplification
RFLP restriction fragment length polymorphism
RNA ribonucleic acid
RNase ribonuclease
RT reverse transcriptase
RTBV Rice tungro bacilliform virus
s second(s)
siRNA small interfering RNA
SOC super optimal broth with catabolite repression
spp. species pluralis (Latin), multiple species
tRNA transfer RNA
TAE Tris‐Acetate‐EDTA buffer
TC tissue culture
tris tris (hydroxymethyl) aminomethane
U enzyme unit(s)
Ug Uganda (survey sample collection)
V volt(s)
v/v volume per volume
VIGS virus‐induced gene silencing
w/v weight per volume
X‐gal 5‐bromo‐4‐chloro‐indolyl‐galactopyranoside
xii
Statement of Original Authorship The work contained in this thesis has not been previously submitted to
meet requirements for an award at this or any other higher education
institution. To the best of my knowledge and belief, the thesis contains no
material previously published or written by another person except where
due reference is made.
Signature: Date:
Pål K. B. Bjartan
xiii
Acknowledgements I would like to thank a number of people for their guidance and support
throughout the course of my studies. First and foremost I want to thank my
supervisors, Dr Anthony James and Prof Rob Harding, for their academic
guidance, dedication and endless patience throughout the course of my
studies.
I’d also like to thank Ben Dugdale and Don Catchpoole for their
advice when I couldn’t get my cloning experiments to work. Your guidance
was crucial to the success of my experiments. Also thanks to Bojana Bokan
for providing me PCR‐primers and Karlah Norkunas for providing me the
agro‐cells. Assistance offered by the staff of CTCB is also acknowledged.
Also, so many thanks to my parents, Iver and Tove, for your financial
support when QUT decided to screw me over and raise the tuition fees.
Without your help I wouldn’t have made it.
Finally, I want to thank all the friends I’ve made throughout my stay
in here in Oz, Mayra and Sam in particular. Even though you at times have
been a great distraction from my studies, you are also the ones that have
made my stay here worthwhile. You have given me memories that I’ll never
forget and helped me unplug when I needed it sorely, which in the end
helped me get through my studies.
xiv
1
Chapter 1
Introduction
1.1 Bananas
Bananas (Musa spp.) are one of the most important, but undervalued, food
crops in the world providing sustenance to millions of people (Jones, 2000;
Karanja et al., 2008). Worldwide, bananas are cultivated on an area of some
nine million hectares; world production averaged 92 million tonnes per
annum during 1998‐2000 and was estimated at 99 million tonnes in 2001
(The World Banana Economy, 1985‐2002, 2003), of which roughly one third
was produced in the Latin American–Caribbean region, one third in Africa
and one third in the Asian–Pacific region. One of the highest consumption
rates is in the Great Lakes region of East Africa, where bananas comprise a
large proportion of the diet (AfricanCrops.net). In Uganda, the average
consumption per capita was 243 kg/year in 1996 (The World Banana
Economy, 1985‐2002, 2003).
Banana is an attractive perennial crop for developing countries due
to its ability to produce fruit all year round and, thus, provide a stable
source of income or nutritious food (The World Banana Economy, 1985‐
2002, 2003; Jones, 2000). Bananas for domestic consumption are produced
from a multitude of cultivars which are grown on various soil types in
different environments (Jones, 2000).
Bananas can be divided into two main categories, namely dessert
bananas and cooking bananas. Dessert bananas can be eaten raw when
ripe as they contain a considerable amount of sugar and are easily
digestible. There are various cultivars of dessert bananas but fruit from the
2
Cavendish cultivar subgroup is the most common (Jones, 2000). In the
period 1998‐2000, dessert bananas comprised 59% of annual world
production, of which the Cavendish subgroup accounted for 47% (The
World Banana Economy, 1985‐2002, 2003). In contrast to dessert bananas,
cooking bananas are usually starchy when ripe and need to be boiled, fried
or roasted to become palatable. They are an important part of the diet to
many people in tropical regions, and are mainly consumed locally. Of these,
the plantains are the most well known types (Jones, 2000). Cooking
bananas comprised 41% of annual world production during 1998‐2000, of
which plantain contributed 17% and the remaining part were distributed
among other cooking cultivars, among them the East African Highland
Banana (EAHB) subgroup (The World Banana Economy, 1985‐2002, 2003).
In Uganda, cultivars of the EAHB subgroup (including many different
cooking banana cultivars) and the dessert cultivar Sukali Ndizi are the most
popular (Bagamba et al., 2003).
The overwhelming majority of presently cultivated varieties of
banana are derived from the Musa acuminata (with a designated ‘A’
genome) and M. balbisiana (with a designated ‘B’ genome) species of the
Eumusa series of Musa (Price, 1995; Karanja et al., 2008). Essentially,
hybridisation between various sub‐species of the polymorphic species M.
acuminata has led to a range of diploid cultivars (designated AA in the
generally accepted classification scheme). The diploid AA cultivars have
given rise to the triploid AAA varieties through a meiosis‐associated process
termed chromosome restitution (Price, 1995). This occurs when meiosis
breaks down at the second division, and forms viable, diploid egg cells
which are then fertilised with haploid pollen (Jones, 2000). Hybridisations
between AA cultivars and M. balbisiana (designated BB) gave rise to the
various AAB and ABB types that are cultivated today. The major reason for
the preference for cultivation of triploid cultivars by man is believed to have
been parthenocarpy and the absence of seeds. Triploidy provides various
3
benefits of which the most important are that plants tend to be more
vigorous and productive than diploids. The vast majority of cultivars are
triploids; AAA cultivars provide many of the sweeter dessert varieties, while
AAB and ABB cultivars often provide the more starchy cooking varieties. M.
balbisiana is considered to be more disease resistant than M. acuminata
and these qualities are often found in the cultivars with the B component in
the genome. Only a few natural cultivars have been recognised as
tetraploids belonging to either AAAA, AAAB, AABB or ABBB genomic
groups, and these are believed to have arisen from the fertilisation of
triploid egg cells by haploid pollen. Tetraploid hybrids have been artificially
bred for disease resistance, and are becoming important in some countries
(Jones, 2000).
African bananas are grouped into three categories, which includes
the East African bananas (AA, AAA, ABB and AB), which are mainly dessert
bananas, the African plantain bananas (AAB) grown mainly in central and
west Africa, and the EAHB (AAA) used for cooking and beer preparation.
Although bananas are not of African origin, Africa has grown into an
important zone of secondary genetic diversity (AfricanCrops.net).
1.2 Pests and diseases affecting bananas
1.2.1 Pests
1.2.1.1 Rhizome and root pests
The two major rhizome and root pests causing problems to banana plants
are the banana weevil borer and the burrowing nematode. The banana
weevil borer (Cosmopolites sordidus) is distributed widely throughout all
continents of the world where banana is grown. It attacks all Musa species
and cultivars and no resistant cultivars are known. Weevil borers are spread
from farm to farm via infested sucker planting material, with on‐farm
spread by short‐distance weevil borer movement between plants.
4
Several parasitic nematodes also cause damage to banana and
plantain. The burrowing nematode (Radopholus similis) is the most
widespread and damaging nematode attacking bananas and plantains. It is
present throughout the tropics, Australia and South Africa. R. similis is
easily spread in water run‐off, by adhering to soil particles or through de‐
suckering tools. As with the weevil borer, the main cause of spread is
infested plant material, and this is how the nematode has been introduced
into previously clean areas (Robinson, 1996).
1.2.1.2 Bunch pests
The majority of banana bunch pests cause superficial peel damage which
does not affect the eating quality of the fruit. In this respect, the
subsistence or cash cropping sector of banana production is not severely
disadvantaged by bunch pests. However, for the discerning export markets
of the world, quality standards are strict and external blemishes caused by
bunch pests are totally unacceptable.
Bunch pests include various species of thrips, such as red rust thrips
species (Chaetanaphothrips spp.), 'corky scab' thrips (caused by the Thrips
florum complex), flower thrips (Frankliniella spp.) and the banana silvering
thrips (Hercinothrips bicinctus). Bunch pests also include some species of
moths, like the banana scab moth (Nacoleia octaseama), found in Australia
and the Pacific Islands, and the banana moth (Opogona sacchari). Although
infestation of a plantation may not be widespread, the attacked bunches
can be severely damaged (Robinson, 1996).
1.2.1.3 Leaf pests
The banana aphid (Pentalonia nigronervosa) is widely distributed in all
banana‐growing areas. Its significance as a pest is due to its ability to act as
a vector for the transmission of Banana bunchy top virus (BBTV,
Nanoviridae, Babuvirus) from plant to plant (Robinson, 1996). Similarly,
mealybugs are known to transmit Banana streak virus (BSV) species (Jones
5
and Lockhart, 1993; Kubiriba et al., 2000). Various insects that visit banana
flowers, bract abscission surfaces, cut suckers and pseudostems can also
mechanically transmit disease‐causing bacteria from plant to plant
(Robinson, 1996).
1.2.2 Diseases
Bananas are susceptible to a range of serious and debilitating diseases
caused by fungi, bacteria and viruses.
1.2.2.1 Fungal diseases
The most serious fungal disease is known as black sigatoka (BS) (caused by
Mycosphaerella fijiensis), also called 'black leaf streak'. BS has caused
devastation to commercial bananas grown in all tropical localities where
banana is grown. It was first recorded from the islands of the Pacific and
has since spread to Asia, Latin America and Africa. In Africa, it has spread
throughout West, Central and East Africa and threatens the production of
bananas and plantains in the area, which are a staple food source for
millions of people (Robinson, 1996; Carlier et al., 2000).
Another important banana disease is Fusarium wilt or Panama
disease, caused by the soil‐borne pathogen Fusarium oxysporum f. sp.
cubense. This disease destroyed over 40,000 ha of Gros Michel (AAA)
export bananas in tropical America in the late 1950s and early 1960s. The
industry was only saved by the introduction of the resistant cultivars in the
Cavendish complex (Robinson, 1996; Ploetz and Pegg, 2000).
1.2.2.2 Bacterial diseases
Moko disease, caused by the bacterium Ralstonia solanacearum, is one of
the most serious diseases of bananas and plantains (Robinson, 1996). It has
caused severe crop losses in smallholder plantations in Latin America and
the Caribbean. Outside the western hemisphere, it has been reported only
in the Philippines and Indonesia. Cavendish (AAA) cultivars are known to be
susceptible as are some ABB cultivars such as 'Bluggoe'. The bacterium
6
infects the plant through wounded tissue, and can be transmitted by a
mechanical vector such as a pruning knife, but it can also be transmitted by
insects or by root‐to‐root contact (Robinson, 1996; Thwaites et al., 2000).
1.2.2.3 Viral diseases
Several viruses are known to infect banana including BBTV,
Cucumber mosaic virus (CMV), Banana bract mosaic virus (BBrMV), Banana
mild mosaic virus (BanMMV), Banana virus X (BVX) and several
Badnaviruses collectively referred to as BSV (Karanja et al., 2008; Geering,
2009).
BBTV is considered the most devastating of the viral diseases
infecting banana crops. It has been reported in several countries in Asia and
the Pacific, Australia and Central Africa, but has not been reported in Latin
America, the Canary Islands, Israel or South Africa. Disease symptoms
include the development of dark green flecks along the leaf veins which
produce a 'dot‐dash' pattern. Affected leaves are more upright than usual
and become pale yellow around the margin. Emerging leaves become
choked in the throat of the plant which creates the 'bunchy top' effect and
pronounced stunting. The insect vector of BBTV, the aphid Pentalonia
nigronervosa, is widespread. While BBTV is not mechanically transmitted,
movement of infected plant material is a concern (Robinson, 1996; Thomas
and Iskra‐Caruana, 2000). The epidemiology of BBTV is, however, very
simple and the disease should therefore be able to be effectively controlled
in developing countries. In Australia, bunchy top has been eradicated from
some regions through a strict inspection and eradication program
consisting of a combination of roguing, clean planting material schemes
and domestic quarantine (Robinson, 1996; Thomas and Iskra‐Caruana,
2000; Geering, 2009). Transgenic approaches to develop resistant cultivars
have been attempted but, despite considerable research efforts, genetically
modified (GM) banana plants with bunchy top resistance have yet to be
developed (Geering, 2009).
7
CMV infects Cavendish cultivars and 'Horn' plantain and occurs
sporadically wherever Musa is grown. The disease is characterised by
conspicuous, sharply defined interveinal chlorosis of the leaves which, in
serious outbreaks, can lead to necrosis and rotting of the heart leaf and
central cylinder. However, serious infection is seldom encountered. CMV is
transmitted from weed and vegetable host plants to banana by several
different aphid species (Robinson, 1996; Lockhart and Jones, 2000a).
BBrMV is widely distributed in the Philippines, India and Sri Lanka.
There are also records of BBrMV from Thailand, Vietnam and Western
Samoa (Rodoni et al., 1999; Thomas et al., 2000a), but the symptoms
expressed in these countries were atypical, and these records need to be
verified. Little is known about the epidemiology of BBrMV (Thomas et al.,
2000a; Geering, 2009).
BanMMV (unassigned member of Flexiviridae) has only received
attention in recent years. Following its identification during routine virus‐
indexing, this virus appears to be one of the most frequently detected
viruses in Musa germplasm (Teycheney et al., 2005a). Although BanMMV
infections are rarely associated with obvious disease symptoms (Thomas et
al., 2000b; Geering, 2009), there are indications which suggest that the
virus may interact synergistically with other banana viruses to cause more
severe disease symptoms than either one would cause alone. For example,
mixed infections of BanMMV and CMV are correlated with a necrotic
reaction (Caruana and Galzi, 1998), and banana plants in Uganda with
severe stunting and bunch malformation were shown to be infected with
an unidentified badnavirus, most likely a BSV, and a filamentous virus
believed to be BanMMV, whereas plants only infected with the badnavirus
had mild disease symptoms (Tushemereirwe et al., 1996). As yet, no vector
of BanMMV has been identified, although there is strong circumstantial
evidence for plant‐to‐plant transmission (Teycheney et al., 2005a).
8
BVX, another unassigned member of the family Flexiviridae, has not
been associated with any disease symptoms and virtually nothing is known
about the epidemiology of this virus. Although there are presently no
records of BVX outside Guadeloupe, it is suspected that its geographic
distribution may be wider (Teycheney et al., 2005b).
Banana streak virus
BSV is the most frequently observed virus affecting banana in the Americas
and most of Africa, in particular Uganda (Tushemereirwe et al., 1996;
Harper et al., 2004, 2005; Geering, 2009) and Kenya (Karanja et al. 2008).
BSD is, however, rarely encountered in either Australia or South Africa. The
reasons for this are not known, but may include epidemiological variables
such as vector type and abundance. Furthermore, legislation in Australia is
used to regulate the banana industry, including planting material, and
subsequently virus diseases are well controlled. In contrast, in Kenya and
Uganda, the high prevalence of the disease may be due to the inadvertent
use of infected planting material by farmers (Geering, 2009).
Mealybugs (Pseudococcus spp.) are the only known vector for
banana‐infecting badnaviruses (Jones and Lockhart, 1993; Kubiriba et al.,
2000), and the rate of spread by this means is believed to be very slow
(Kubiriba et al., 2000). During a 2‐year field trial in north Queensland, no
natural spread of Banana streak CA virus (BSCAV) from infected to healthy
plants was observed (Daniells et al., 2001).
Symptoms of BSD can be very severe and include stunting, bunch
abnormalities, splitting and internal necrosis of the pseudostem and fruit.
However, in some environmental conditions, infected plants may remain
symptomless. Yield losses from BSV infection vary widely (Daniells et al.,
1999; Lockhart and Jones, 2000b). In Cavendish, the yield loss has been
estimated to vary from 7% to 90% depending on the severity of the
symptoms (Lassoudière, 1974). BSV is an important limiting factor to
banana production in countries like Kenya and Uganda (Kubiriba et al.,
9
2000; Karanja et al., 2008). Additionally, BSV is recognised as a major
constraint to both the genetic improvement of banana as well as
germplasm dissemination. The reasons for this are discussed later.
BSVs are classified in the genus Badnavirus within the family
Caulimoviridae (Lockhart, 1986; Fauquet et al., 2005). They encapsidate a
non‐covalently closed, circular dsDNA genome and are known as
pararetroviruses since they replicate by reverse transcription via an RNA
intermediate (Lockhart and Jones, 2000b). Unlike true retroviruses,
however, they have no requirement for the integration of their viral
genome into the host chromosome in order to replicate and their genome
does not encode an integrase (Harper et al., 2002b). Like all members of
the Badnavirus genus, BSV have bacilliform virions of approximately 30 ×
120‐150 nm (Figure 1.1b), and a circular dsDNA genome of approximately
7.4 kbp (Harper and Hull, 1998; Lockhart and Jones, 2000b) which contains
three open reading frames (ORFs) and a tRNAmet binding site (Figure 1.1a).
The first two ORFs encode two small proteins (ORF 1 of 20.8 kD, and ORF 2
of 14.5 kD) of unknown function while the third ORF encodes a large
polyprotein (208 kD) that is proteolytically cleaved to produce the viral coat
protein, movement protein, aspartic protease, reverse transcriptase (RT)
and ribonuclease (RNase)‐H (Harper and Hull, 1998).
Replication takes place in both the cytoplasm and nucleus of host
cells. Following entry of the viral genome into the cytoplasm, the viral DNA
is transported to the nucleus where it forms super‐coiled mini‐
chromosomes. The viral DNA is transcribed into polyadenylated RNA which
is terminally redundant. Newly transcribed RNA is transported back to the
cytoplasm where it has two roles. It can either be used as a template for
viral protein synthesis, or it can undergo reverse transcription by the viral‐
encoded reverse transcriptase to synthesise the virus‐sense strand DNA.
This DNA can then re‐enter the nucleus for amplification (Harper et al.,
2002b).
10
Figure 1.1: (a) Map of the BSV genome. Conserved sequence motifs ofbadnaviruses are indicated as black segments. The positions of the threeORFs of BSV are given by filled arrows. (Modified from Harper et al., 2004)(b) Electron micrograph of BSV. The particle sizes are ca. 120 nm × 30 nm(Harper et al., 2004).
a)
b)
11
Based on current classification criteria which delimits individual
species based on nucleotide differences of greater than 20% in the
RT/RNase H‐coding region (King et al., 2011), there are presently three BSV
species recognised by the International Committee on the Taxonomy of
Viruses (ICTV), namely Banana streak GF virus (BSGFV), Banana streak MY
virus (BSMYV) and Banana streak OL virus (BSOLV), while one putative
species, Banana streak VN virus, is awaiting recognition (King et al., 2011).
Six more species, namely Banana streak CA virus (BSCAV, previously Banana
streak Cavendish virus), Banana streak IM virus (BSIMV, previously Banana
streak Imové virus), Banana streak UA virus (BSUAV, previously Banana
streak Uganda A virus), Banana streak UI virus (BSUIV, previously Banana
streak Uganda I virus), Banana streak UL virus (BSULV, previously Banana
streak Uganda L virus), Banana streak UM virus (BSUMV, previously Banana
streak Uganda M virus) have also recently been proposed based on full‐
length sequence analyses (Geering et al., 2011; James et al., 2011b). Partial
genome sequences have been reported for nine other putative virus
species, based on isolates of BSV from Uganda, named Banana streak
Uganda B–H, J and K viruses (Harper et al. 2004, 2005). However, until the
complete genomic sequences of these putative viruses have been
determined, their taxonomic status remains unresolved. Nevertheless, in
spite of the pending status with ICTV, undeniably a great amount of
diversity exists in the collection of badnaviruses, now commonly referred to
as BSV, infecting bananas (Figure 1.2).
BSV is one of the few plant viruses which have integrated sequences
present in their host plant genome. (LaFleur et al., 1996; Harper and Hull,
1998; Harper et al., 1999b; Ndowora et al., 1999; Geering et al., 2001,
2005a, b). There are two groups of integrated BSV sequences known to
occur in banana. The first group, named endogenous pararetroviruses
(EPRVs) contains the majority of integrated sequences and comprises
incomplete virus genomes (as a result of large‐scale genome
12
Figure 1.2: Unrooted tree showing the identified BSV species grouped intothree clades (shaded) based on nucleotide sequence identities of greaterthan 72%. White circles represent the different BSV species found inUganda, and for clarity only the descriptor of the Ugandan BSV species isused (Modified from Harper et al., 2005). Using the old abbreviations somecommon BSV species is denoted as following; BSOLV is abbreviated BSOlV,BSCAV as BSCavV, BSGFV as BSGfV and BSMYV as BSMysV.
13
rearrangements and gene deletions or nucleotide substitutions giving rise
to translational frame‐shifts or premature stop codons) which are incapable
of causing infection (LaFleur et al., 1996; Harper and Hull, 1998; Harper et
al., 1999b; Ndowora et al., 1999; Geering et al., 2001, 2005a, b). EPRV
sequences are believed to have been present in the Musa genome for a
very long time, almost certainly pre‐dating domestication. M. acuminata
and M. balbisiana have different populations of EPRVs, which suggest that
the integration events occurred after the divergence of these species, an
occurrence estimated to have happened approximately 4.6 million years
ago (Geering et al., 2005a; Lescot et al., 2008). Although some EPRV
sequences show homology to the genomes of known episomal BSV, many
others have no known episomal counterpart (Geering et al., 2005a). The
second type of integrated sequences are called endogenous BSV (eBSV) and
consist of the entire genome of characterised episomal BSV species which
exist as multiple non‐contiguous regions of the virus DNA combined with
host‐genomic sequences. Under certain stress conditions, particularly in
vitro propagation and hybridisation, recombination events occur in the
integrated sequences allowing the reconstituted viral genome to be
activated thereby resulting in episomal infections (Ndowora et al., 1999;
Dallot et al., 2001; Gayral et al., 2008; Côte et al., 2010; Iskra‐Caruana et
al., 2010). Whilst the incomplete EPRVs have been found in both the A‐ and
B‐genomes derived from the wild progenitors of domesticated banana,
Musa acuminata (A) and M. balbisiana (B), respectively, the eBSV have only
been detected in the B‐genome of various banana accessions (Geering et
al., 2001, 2005b; Gayral et al., 2008).
It is thought that the majority of eBSV sequences are silenced by
DNA methylation and heterochromatisation, although a few inactivateable
copies are transcribed at low levels. These non‐infectious RNA transcripts
are then thought to form a template for the synthesis of small interfering
RNAs (siRNAs), which, in turn, maintain the epigenetic control of the virus.
14
It is only when stresses such as interspecific hybridisation or tissue culture
(TC) propagation weakens the epigenetic control that activateable copies of
viral DNA are transcribed to cause infection (Dallot et al., 2001; Lheureux et
al., 2003).
A question which remains unanswered is why the rate of activation
of BSV is low in some hybrids, such as 'Goldfinger' (syn. 'FHIA‐01', AAAB
genome), whilst it is high in other cultivars. The rate of activation of eBSOLV
in 'Lady Finger' (AAB genome), a popular cultivar in subtropical Australia, is
virtually non‐existent, even though it has been propagated through TC for
many years. In stark contrast, the activation rate of eBSOLV in 'FHIA‐21'
(AAAB genome) is 58% after only six in vitro subcultures (Dallot et al.,
2001). The Goldfinger cultivar is presumed to contain an activatable allele
of eBSGFV DNA, given that this virus was originally discovered in this plant,
but the activation rate is extremely low (Gayral et al., 2008). The
commercialisation of 'Goldfinger' gives some hope to plant breeders that it
is possible with current technology to develop new, improved bananas and
plantains without eBSV being a constraint (Geering, 2009).
1.2.3 BSV diagnostics
BSV species are serologically and genomically variable (Lockhart and Jones,
2000b; Harper et al., 2002a) and this creates problems for the development
of reliable diagnostic tests. Published methods for BSV detection include
enzyme‐linked immunosorbent assay (ELISA), polymerase chain reaction
(PCR), immunosorbent electron microscopy (ISEM), immunocapture‐PCR
(IC‐PCR) and rolling circle amplification (RCA) (Dahal et al., 1998; Harper et
al., 1999a, b, 2002a, 2004, 2005; Geering et al., 2000; Le Provost et al.,
2006; Karanja et al., 2008; James et al., 2011a, b). Each of these methods
has its inherent advantages and disadvantages.
Although ELISA allows for the screening of large numbers of samples
at low cost, it suffers from a lack of sensitivity and is, therefore, unsuitable
15
for detection of BSV in asymptomatic plant tissue or where the virus occurs
in low concentration (Dahal et al., 1998; Harper et al., 2005). Further, ELISA
as well as IC‐PCR and ISEM, relies on the initial capture of virions onto a
surface using BSV‐specific antibodies. Due to the highly variable nature of
the BSV coat protein (Lockhart and Jones, 2000b; Harper et al., 2002a,
2005), it is virtually impossible to generate an antiserum that
detects/captures all isolates of BSV. This means such a diagnostic approach
has the potential to result in false negatives.
PCR is a very sensitive method for virus detection. Further, the
reagents are now becoming available at an affordable cost for routine use
in developing countries and the associated equipment has broad
application in plant biotechnology. Crucial to the successful detection of
viruses by PCR is the availability of primers that will allow the amplification
of viral genomic sequences. Primer sets for the specific detection of BSOLV,
BSCAV, BSMYV, and BSGFV have been developed (Geering et al., 2000)
along with degenerate primer sets that can detect several BSV species as
they are designed to conserved sequences in the reverse
transcriptase/RNase H‐coding region of ORF 3 of badnaviruses (Harper et
al., 2002a, 2004, 2005; Yang et al., 2003; Karanja et al., 2008). The major
problems to date with PCR is the presence of endogenous sequences which
may yield a positive result despite the absence of episomal virus (Harper et
al., 1999b, 2005; Yang et al., 2003), and the use of several primer sets
which are still considered unlikely to detect the entirety of the BSV
sequence diversity. In an attempt to circumvent the detection of integrated
DNA, immunocapture (IC)‐PCR is currently used as the “gold‐standard” for
BSV indexing. IC‐PCR using degenerate primers to amplify BSV has been
demonstrated (Harper et al. 2002a, 2004, 2005; Karanja et al., 2008).
However, this method still has serious limitations due to the inability of
antiserum to capture all BSV isolates (Harper et al., 2002a), along with the
limitations to PCR previously mentioned. Further, the presence of
16
contaminating, carry‐over nucleic acid remaining in capture tubes can also
lead to false positives by detection of EPRV and eBSV sequences in the
Musa genome (Le Provost et al., 2006; Iskra‐Caruana et al., 2009).
Therefore, the development of a new, more effective approach that can
overcome the dependence of antisera to capture BSV virions is needed.
A new promising method for detection of BSV is the application of
RCA combined with restriction fragment length polymorphisms (RFLPs)
(James, 2011; James et al., 2011a). RCA uses bacteriophage φ29 DNA
polymerase to amplify circular DNA molecules in a sequence‐independent
manner (Blanco et al., 1989; Dean et al., 2001). Previous studies have
shown that RCA in combination with RFLP is a highly reproducible tool for
geminivirus diagnosis, and is largely independent of viral genome
organisation, source plant type and origin, and sample preparation (Haible
et al., 2006; Schubert et al., 2007). Since the episomal DNA of BSV is
circular as opposed to the integrated sequences, RCA is able to discriminate
between episomal and integrated BSV sequences and thus overcome the
occurrence of false‐positives. An additional practical advantage of using
RCA is that it will amplify uncharacterised BSV species for which sequence
information is not available, making it useful to detect new BSV species.
One of the major disadvantages of this approach, however, is that for BSV
identification it is relatively expensive which may make it prohibitive for
widespread use in developing countries.
1.2.4 BSV diagnostics – issues and possible solutions
One of the difficulties faced by Australian researchers involved in
the development and optimisation of diagnostic assays for BSV is the
limited availability of diseased plant material. Despite being widespread in
East Africa, the disease is sporadic in Australia due largely to the
implementation of disease control programs. Further, due to the slow rate
of transmission by the mealybug vector and the inability to mechanically
transmit the virus (Jones and Lockhart, 1993; Kubiriba et al., 2000; Huang
17
and Hartung, 2001), obtaining large numbers of infected plants as controls
is difficult. One approach to generate large numbers of infected plants is to
construct an infectious clone of the virus.
An infectious clone is essentially a replication competent double‐
stranded DNA copy of the viral genome carried in a bacterial plasmid.
These transgenic plasmids can be introduced into cells by transformation to
produce infectious virus. Successful development of infectious clones of the
Tungrovirus, Rice tungro bacilliform virus (RTBV), and the Badnavirus, Citrus
yellow mosaic virus (CYMV), have been reported (Dasgupta et al., 1991;
Huang and Hartung, 2001). The existing model for production of a
successful infectious clone of Badnavirus involves the expression (following
entry to the plant cell) of a larger than unit length terminally redundant
transcript of the viral genomic DNA by host DNA‐dependent RNA
polymerase, using the cloned virus DNA as template. During a normal virus
infection, this greater than full‐length transcript is produced and works as
the template for viral minus‐strand DNA synthesis using the virus‐encoded
replicase protein, as well as a polycistronic RNA for expression of the virus
encoded proteins (Pfeiffer and Hohn, 1983; Hohn et al., 1985; Temin, 1989;
Medberry et al., 1990; Dasgupta et al., 1991; Qu et al., 1991; Bouhida et al.,
1993; Jacquot et al., 1999).
There are several ways to inoculate plants with the infectious clone.
Wounding the plant with an inoculated scalpel or shooting it into the tissue
with a biolistic particle delivery system are two options. The method that
may prove to be the most efficient is Agrobacterium‐mediated inoculation,
where the vector is transformed into competent cells of Agrobacterium
tumefaciens which in turn are used to inoculate plants. This method has
been shown to be the most efficient or, in some cases, the only way to
cause systemic infection in plants using infectious clones of badnaviruses
and caulimoviruses (Grimsley et al., 1986; Dasgupta et al., 1991; Huang and
Hartung, 2001).
18
1.3 The situation in Uganda
Banana crops in Uganda have two major problems. Firstly, the staple food
landraces of EAHB are low in two essential micronutrients namely pro‐
vitamin A and iron (Fungo et al., 2007; Honfo et al., 2007). As a result,
Ugandan populations dependent on a banana‐based diet have a high
incidence of vitamin A deficiency and iron deficiency anaemia (Bachou,
2002). Secondly, as with most other banana‐producing countries, Ugandan
bananas are affected with a wide range of devastating pests and diseases
that threaten the food supply (Robinson, 1996; Bagamba et al., 2003). Of
these, BSV is considered the most important virus. Two approaches are
currently being used in an effort to overcome these two issues.
As part of a Bill and Melinda Gates Grand Challenges in Global
Health Project, researchers within the CTCB at QUT are attempting to
develop genetically modified bananas with increased levels of pro‐vitamin
A and iron. This approach has been successfully used with other crops such
as rice, and was the only real option due to the difficulties associated with
conventional banana breeding, mainly due to the low fertility of the
common cultivars and landraces. An important parallel and complementary
project was also initiated to address the need for reliable virus indexing of
planting material prior to its distribution to growers. It is with this latter
project that my research project is concerned.
One of the widely used and accepted strategies for the production
of disease free bananas in developing countries such as Uganda is
micropropagation, often referred to as TC. Agriculture in East Africa consists
mainly of small‐scale farm activity in which the cultural practice in banana
production has been for farmers to transplant banana suckers from one
farm to another. This has, in turn, substantially contributed to the efficient
transmission of banana viruses throughout the region. As a result of this,
farmers are encouraged to use clean, disease‐free and insect‐free planting
material, which is effectively obtained by TC propagation techniques (Africa
19
Harvest, 2009). Micropropagation of bananas involves the excision of the
meristem of usually a field grown banana and its establishment in a sterile
environment on sterile media. By applying this method, fungal, bacterial or
nematode diseases can be avoided, as infected or infested meristems are
easily recognised in culture and discarded. The meristems can then be
stimulated to produce multiple plantlets through the manipulation of the
plant hormone levels in the media. Hence, a very large number of plantlets
can be generated from a single meristem, each genetically identical to the
original plant. The plantlets are then removed from the sterile
environment, acclimatised and grown in a nursery to a size appropriate for
planting in the field. A danger of micropropagation, however, is that if the
original parent plant is virus infected, all progeny (which may be in the
millions) will also be infected (Daniells et al., 2001). This not only causes
potentially devastating effect on the crop plants but also poses a threat of
international virus spread. It is imperative, therefore, that a reliable virus
indexing scheme is available to screen for viruses. Banana virus indexing
schemes, combined with micropropagation, is well established in a number
of countries including Australia. However, the major problem with these
schemes is the lack of a reliable diagnostic test for BSV.
1.4 Project aim and objectives
The current diagnostic tests for BSV have serious limitations due to the
extreme serological and genetic differences between the BSV species.
Serological methods lack the ability to detect the vast differences in the
BSV coat protein, and these methods are therefore unsuitable for reliable
detection of all BSV species. The challenge with PCR detection of BSV lies
firstly with the hypervariable genome, which makes the primer design very
difficult, and secondly with the presence of integrated sequences in the
banana genome. Although BSV sequences are integrated in both the A‐ and
B‐genomes of banana, it is with the B genome that the major challenges lie
20
as this genome contains sequences identical to four known episomal BSV
species. A novel BSV diagnostic test, based on a combination of RCA and
RFLPs, has been developed at QUT which appears promising. However, RCA
has the problem that it is relatively expensive and, therefore, may not be
affordable to developing countries where the need is highest. A PCR‐based
diagnostic test is a cost‐effective and sensitive alternative, but this
approach demands the design of primer sets that will avoid detection of
eBSV. The overall aim of this project was to improve the current strategies
for BSV detection. This aim was to be achieved through the following three
objectives:
1. Compare PCR and RCA as diagnostic assays for BSV
2. Develop episomal‐specific primer sets for BSV species with
endogenous counterparts
3. Develop an infectious clone of BSMYV to generate BSV infected
plant material for assessment of diagnostic tests.
21
Chapter 2
General materials and methods
2.1 General materials
2.1.1 Sources for specialised reagents
Laboratory reagents were acquired from scientific supply companies such
as Sigma Aldrich and Crown Scientific. Agarose for gel electrophoresis was
obtained from Roche Diagnostics and stain used was SYBR® Safe DNA Gel
Stain provided by Invitrogen Corp. Agar for bacterial culture was purchased
from Oxoid. Molecular weight markers were purchased from Bioline, while
modifying and restriction enzymes were purchased from Roche Diagnostics
or New England Biolabs.
2.1.2 Reagents and equipment for DNA amplification
Rolling circle amplification (RCA) of DNA was conducted using the Illustra
TempliPhi™ 100 Amplification Kit provided by GE Healthcare,
Buckinghamshire, United Kingdom. Polymerase chain reaction (PCR)
amplification was undertaken using GoTaq® Green Master Mix provided by
Promega. Both amplification reactions were undertaken on a Peltier
Thermal Cycler‐PTC200 from MJ Research.
2.1.3 Oligonucleotide synthesis
Oligonucleotide primers were synthesised by GeneWorks and supplied as
precipitated DNA, usually within the range 30‐50 nmol. Stock solutions
were made by dissolving primers with deionised water (dH2O) to
concentrations of 100 or 200 pmol/µl. Stock solutions were diluted with
dH2O to working concentrations of 5 pmol/µl.
22
2.1.4 Bacterial strains
Heat‐shock competent Escherichia coli (E. coli) strain XL1‐blue, used for
general plasmid cloning, and Agrobacterium tumefaciens (A. tumefaciens)
strain AGL1, used for the inoculation of Cavendish banana plants, was
kindly supplied by colleagues Dr Anthony James and Ms Karlah Norkunas.
2.2 General methods
2.2.1 DNA extraction from leaf tissue
Total NA was isolated from leaf tissue from various cultivars of Musa and
Saccharum using a cetyltrimethylammonium bromide (CTAB)‐based
method. Either fresh tissue (0.4 g) or dried tissue (0.04 g) was ground to a
fine powder with washed sand in liquid nitrogen with a mortar and pestle.
To the ground leaf tissue, 3 ml CTAB extraction buffer (0.1 M tris
(hydroxymethyl) aminomethane (Tris)‐HCl solution, pH 8.0, 0.05 M
ethylenediaminetetraacetic acid (EDTA), pH 8.0, 1.4 M sodium chloride,
0.08 M sodium sulphite, 10 g/l polyvinylpyrrolidone (PVP) and 0.05 M
CTAB) was added, and the homogenate was distributed to two 2 ml
microfuge tubes. The homogenates were incubated at 65°C for 30 min in a
water bath, inverting tubes several times. The homogenate was centrifuged
at 18,000 g for 5 min. Supernatant (750 μl) was extracted with an equal
volume of chloroform‐isoamylalcohol (Chl:IAA) solution (24:1 v/v). The
phases were vortexed thoroughly and then centrifuged at 18,000 g for 5
min. Supernatant (600 μl) was removed and extracted a second time with
an equal volume of Chl:IAA as described. The supernatant (450 μl) was
removed and nucleic acids (NA) precipitated by adding an equal volume of
isopropanol (100%). Total NA was collected by centrifugation at 18,000 g
for 5 min, and pellets washed using 500 μl of ethanol (70%) followed by
centrifugation at 18,000 g for 1 min. Pellets were air dried and resuspended
in 50 μl of dH2O at 4°C overnight. Where indicated, NA extracts were
23
analysed using the Nanodrop2000 spectrophotometer and diluted to 10
ng/μl for PCR.
2.2.2 PCR amplification
PCR amplification was undertaken using GoTaq® Green Master Mix
(Promega). Unless otherwise stated, 2× GoTaq mixture, 5 pmol each of
sense and anti‐sense primer, 10 ng CTAB NA extract and dH2O were added
up to a final volume of 20 μl and then incubated in a Peltier Thermal Cycler‐
PTC200 (MJ Research) under the following conditions: Initial denaturation
at 94 °C for 2 min, followed by 35 cycles of denaturation at 94 °C for 20 s,
annealing at the temperature specified for 20 s and extension at 72 °C for
time specified, with the subsequent final extension for a minimum of 2
min. Samples where held at 14 °C until analysis by agarose gel
electrophoresis.
2.2.3 RCA
DNA was amplified using the Illustra TempliPhi 100 Amplification kit (GE
Healthcare), essentially according to the manufacturer’s instructions but
with slight modifications as described by James et al., (2011a). Nucleic acid
extract (1 µl) was mixed with 3 µl of kit sample buffer and 1 µl of a 5 µM
stock solution (approximately 4.16 pmol/µl of each primer) of degenerate
primers. The mixture was denatured at 95°C for 3 min and cooled on ice for
another 3 min, and subsequently kit reaction buffer (5 µl) and polymerase
(0.2 µl) was added. The mixtures were incubated at 30°C for 18 h and then
the reactions were stopped by incubation at 65°C for 10 min. Samples were
held at 14 °C prior to subsequent analysis.
2.2.4 Restriction digestion
Unless stated otherwise, the entire RCA reaction volume (10 µl) or 15 µl of
miniprep DNA was digested using 2× buffer, restriction enzyme(s) (2 U) and
dH2O in a total volume of 20 µl. The samples were incubated for 1.5‐2 h at
the recommended temperature, by manufacturer, for each enzyme used.
24
2.2.5 Agarose gel electrophoresis
Agarose gels were prepared by melting 1% (w/v) solid agarose in Tris‐
Acetate‐EDTA (TAE) buffer. Samples were loaded and electrophoresed at
100 V or 120 V for 45 min for PCR, and 120 V at 60 or 90 min for RCA and
miniprep DNA restriction digests. Gels were stained with 0.25× SYBR® Safe
DNA Gel stain.
2.2.6 DNA extraction from agarose gel
DNA bands were excised from agarose gels and stored at ‐20 °C until DNA
recovery. DNA was extracted from the agarose gel using Quantum Prep™
Freeze 'N' Squeeze DNA Gel Extraction Spin Columns (Bio Rad), using their
standard protocol.
2.2.7 Dephosphorylation of 5’ ends
The phosphate group was removed from the 5’ ends of linearised plasmid
DNA by incubating with 3 U of calf intestinal alkaline phosphatase (CIP,
Roche) at 37 °C for 50 min, with the subsequent heat‐inactivation at 65 °C.
Following dephosphorylation, CIP was removed from the DNA sample using
agarose gel electrophoresis and DNA gel extraction using Quantum Prep™
Freeze 'N' Squeeze DNA Gel Extraction Spin Columns (Bio Rad).
2.2.8 DNA ligation
The specific ligation methods are described in each chapter.
2.2.9 Transformation of E. coli
Chemically competent E. coli strain XL1‐blue was transformed by heat
shock. Approximately 10 µl of ligation mix were added per 50 µl of
competent E. coli culture and incubated on ice for about 15 min. Cells were
heat shocked at 42 °C for 90 s, returned to ice, resuspended in 150 µl LB or
SOC (Sambrook and Russell, 2001) and incubated for approximately 1 h at
37 °C with shaking. Transformed E. coli were plated onto solid LB agar
25
medium containing the appropriate antibiotics (ampicillin, 50 µg/ml, unless
specified otherwise), supplemented with IPTG (0.2 g/l) and X‐gal (40 mg/l)
for white/blue selection vector systems, and incubated at 37 °C for 18‐24 h.
Colonies were selected, using white/blue selection when
possible/appropriate, and incubated in 1 ml of liquid LB with the
appropriate antibiotics at 37 °C overnight with shaking.
2.2.10 Mini‐preparation protocol
Bacterial cells from overnight cultures were pelleted in a microcentrifuge at
18,000 g for 30 s. Pellets were resuspended in 150 µl glucose‐Tris‐EDTA
buffer (50 mM glucose, 25 mM Tris‐HCl pH 8.0, 10 mM EDTA) by vortexing.
Cells were then lysed by the addition of 150 µl sodium hydroxide
(0.2M)/sodium dodecyl sulphate (1% w/v) solution, followed by gentle
inversion. Chromosomal DNA and proteins were precipitated by adding 150
µl potassium acetate solution (5 M, 200 µl) followed by 200 µl Chl:IAA
solution (24:1) and centrifugation at 18,000 g for 5 min. Supernatant (400
µl) was removed and the DNA precipitated with 900 µl ethanol (100 %),
before centrifuging at 18,000 g for another 5 min. The DNA pellets were
washed in 350 µl ethanol (70% v/v) and centrifuged again at 18,000 g for 2
min. The pellets were air or vacuum dried and resuspended in 50 µl dH2O
with 1 µl RNaseA (20 µg/ml) added.
2.2.11 Sequencing
All sequencing reactions used plasmid DNA and the ABI Big Dye Terminator
Cycle Sequencing Kit version 3.1 (Applied Biosystems). Each reaction
contained 1 µl Big Dye v3.1 mix, 2 µl plasmid DNA, 3.33 pmol of primer, 3.5
µl of sequencing buffer (400 mM Tris pH 9.0, 10 mM MgCl2) and dH2O in a
final volume of 20 µl. All sequencing cycle reactions used an initial
denaturation of 95 °C for 5 min followed by 40 cycles of denaturation at 95
°C for 30 s, annealing at 50 °C for 20 s and extension at 60 °C for 4 min.
Following this, extension products were precipitated by adding 0.1 volume
26
of sodium acetate (3 M) and EDTA (125 mM) and 2.5 volumes of 100%
ethanol, and incubating at room temperature for 15 min. Products were
pelleted by centrifuging at 18,000 g for 20 min. Pellets were washed with
250 µl ethanol (70 %) and centrifuged at 18,000 g for 5 min. The dried DNA
pellets were capillary sequenced either by Griffith University DNA
Sequencing Facility or at QUT.
27
Chapter 3
Assessment of PCR and RCA for diagnosis of BSV in field samples
3.1 Introduction
Banana streak virus (BSV), which causes banana streak disease (BSD), is the
most common virus affecting banana in the Americas and most of Africa
(Tushemereirwe et al., 1996; Harper et al., 2005). BSV is highly prevalent in
Uganda and Kenya, and has a high diversity of species (Harper et al., 2002a,
2004, 2005; Karanja et al., 2008; James, 2011; James et al., 2011b).
PCR is a very sensitive method for virus detection and primer sets
for the specific detection of nine BSV species have been reported (Geering
et al., 2000; James, 2011; James et al., 2011b). However, while the EPRVs
are of only minor concern for developing primers for reliable PCR
detection, one of the major problems to date with PCR‐based diagnostic
methods is the presence of endogenous BSV (eBSV) in the Musa B‐genome,
as they are full‐length sequences of the corresponding episomal virus
which may yield a positive result despite the absence of episomal virus
(Harper et al., 1999b, 2005; Yang et al., 2003).
Rolling circle amplification (RCA) combined with restriction fragment
length polymorphisms (RFLPs) is a relatively new and promising method for
detection of episomal BSV (James, 2011; James et al., 2011a). An additional
practical advantage of using RCA is that it will amplify uncharacterised BSV
species for which sequence information is not available, making it useful to
detect new BSV species.
28
In order to further determine the diversity of BSV in Africa, a study
was undertaken to identify and characterise the BSV species present in
eastern Uganda using both PCR‐ and RCA‐based tests. This chapter
describes the results of this study and discusses PCR and RCA as methods
for virus detection.
29
3.2 Methods
3.2.1 Samples for analysis
Forty‐five leaf samples (Table 3.1) were collected from nine districts in
eastern Uganda in February 2010. Leaf samples were dried over silica gel
and sent to QUT for analysis. Total nucleic acid was extracted from the leaf
samples as described in section 2.2.1, and nucleic acid extracts were
diluted to 10 ng/µl using deionised water.
3.2.2 PCR amplification
PCR testing was conducted for the detection of BSV species using primers
listed in Table 3.2. As an internal extraction control, extracts were initially
tested for the banana actin gene. PCRs were conducted as described in
section 2.2.2 with PCR extension times set to 30 s for all primer sets, while
specific annealing temperatures for each primer set are listed in Table 3.2.
PCRs were analysed by electrophoresis through agarose gels (section 2.2.5)
and DNA fragments visualized on a Safe imager blue‐light transilluminator.
3.2.3 RCA amplification
RCA testing of nucleic acid extracts was undertaken as described in section
2.2.3. RCA reactions were restriction digested separately using either StuI
or PstI as described in section 2.2.4. All samples that yielded no digest
products using these two enzymes were RCA amplified and digested again
using EcoRI, BamHI, SacI, XbaI, SalI and HindIII in separate reactions. Digest
products were electrophoresed through agarose gels (section 2.2.5) and
DNA fragments visualized on a Safe imager blue‐light transilluminator. For
samples with full‐length restriction fragments or RFLPs distinct from those
predicted from characterised full‐length BSV species, bands were excised
(section 2.2.6) for analysis by cloning and sequencing. Where the RFLP
matched to predicted profiles for BSV species with published full‐length
sequences, these were used to identify the species present. For samples
30
Table 3.1: Plant samples used in
this study
Sample
number
District
Cultivar
Genome
type
Symptoms
188
Luwero
Yangambi Km5
AAA
chlorotic streak, necrotic flecks
189
Luwero
Nfuka
EAH‐AAA
mild
chlorotic flecking
190
Luwero
Nfuka
EAH‐AAA
strong chlorotic flecks
191
Luwero
Lusumba
EAH‐AAA
strong chlorotic/necrotic flecks
192
Luwero
Mpologoma
EAH‐AAA
mild
chlorotic streaking
193
Luwero
unsure
EAH‐AAA
yellow flecking
194
Luwero
Mpologoma
EAH‐AAA
chlorotic flecking
195
Luwero
Mpologoma
EAH‐AAA
chlorotic flecking
196
Luwero
Mpologoma
EAH‐AAA
strong chlorotic flecking
197
Luwero
Kisansa
EAH‐AAA
mild
yellow flecking
198
Luwero
Kisansa
EAH‐AAA
mild
flecking
199
Kam
uli
Mpologoma
EAH‐AAA
mild
chlorotic streaking
200
Kam
uli
Mpologoma
EAH‐AAA
mild
chlorotic flecks
201
Kam
uli
Mpologoma
EAH‐AAA
mild
chlorotic flecks
202
Kam
uli
Mpologoma
EAH‐AAA
mild
chlorotic flecks
203
Kam
uli
Malila
EAH‐AAA
strong chlorotic streaking
204
Kam
uli
Enzirabushera
EAH‐AAA
strong chlorotic streaking
205
Jinja
Mbazirume
EAH‐AAA
strong chlorotic streaking
206
Jinja
Ntinka
EAH‐AAA
strong chlorotic streaking
207
Mbale
Nam
unga
EAH‐AAA
mild
yellow‐green
flecks
208
Mbale
Embululu
EAH‐AAA
mild
yellow flecks
31
209
Mbale
Embululu
EAH‐AAA
strong chlorotic /necrotic flecks
210
Mbale
Dihakho
EAH‐AAA
strong chlorosis, some necrosis
211
Mbale
unknown
EAH‐AAA
moderate streaking
212
Mbale
unknown
EAH‐AAA
very m
ild flecking
213
Manafwa
Mbwazirume
EAH‐AAA
chlorotic flecking
214
Manafwa
Embululu
EAH‐AAA
yellow blotches
215
Manafwa
Murrumbidgee
EAH‐AAA
no sym
ptoms
216
Bududa
Pisang ceylan
AAB
chlorosis, necrosis
217
Bududa
Pisang ceylan
AAB
yellow/green
chlorosis
218
Bududa
Khyebusi
EAH‐AAA
mild
flecking
219
Bududa
Lisitalo
EAH‐AAA
mild
yellow flecking
220
Manafa
Kulone
EAH‐AAA
severe chlorosis
221
Manafa
Mulure
EAH‐AAA
yellow blotches
222
Manafa
Embululu
EAH‐AAA
mild
yellow flecking
223
Manafa
Embululu
EAH‐AAA
yellow flecks, necrosis
224
Manafa
Gros Michel (Bogoya)
AAA
chlorotic flecking
225
Manafa
Embululu
EAH‐AAA
strong flecking
226
Sironko
Mulure
EAH‐AAA
chlorotic/necrotic streaks
227
Sironko
Lusindalo
EAH‐AAA
chlorotic streaks
228
Sironko
Em
bululu
EAH‐AAA
chlorotic streaks
229
Sironko
Giant Caven
dish
AAA
chlorotic streaks
230
Sironko
Kayinja (Pisang aw
ak)
ABB
chlorotic/necrotic streaks
231
Kapuchorw
a Mpologoma
EAH‐AAA
chlorotic/necrotic streaks
232
Kapuchorw
a Gros Michel (Bogoya)
AAA
chlorotic streaks
32
Table 3.2: Primer seq
uen
ces used for PCR testing
Target
BSV
Primer nam
e Annealing
temp. (°C)
Sequence (5'‐3')
Product length
(bp)
Region of genome
amplified
Reference
BSM
yV
Mys‐F1
57
TAAAAGCACAGCTC
AGAACAAACC
589
6481–7069
Geering et al.
(2000)
Mys‐R1 (c)
CTC
CGTG
ATTTCTTCGTG
GTC
BSG
FV
GF‐F1
64
ACGAACTA
TCACGACTTGTTCAAGC
476
6354–6829
GF‐R1 (c)
TCGGTG
GAATA
GTC
CTG
AGTC
TTC
BSCAV
Cav‐F1
57
AGGATTGGATG
TGAAGTTTG
AGC
782
6425–7207
Cav‐R1 (c)
ACCAATA
ATG
CAAGGGACGC
BSO
LV
RD‐F1
57
ATC
TGAAGGTG
TGTTGATC
AATG
C522
6499–7020
RD‐R1 (c)
GCTC
ACTC
CGCATC
TTATCAGTC
BSIMV
B107seq1
50
GCTA
GGAAGAAAAGTCTG
GG
475
7417–122
James et al.
(2011b)
B109seq11 (c)
TGCAAGTCTA
CTTACACAGC
BSU
AV
B105seq2
60
CTC
AGCGGCAAGATTAGGAAGG
517
6513–7029
B105seq15 (c)
TCCCCATTGGTC
GTC
ATTGC
BSU
IV
B89seq2
50
GAATC
CTC
AAAGGTA
CCCC
619
435–1053
B89seq10 (c)
CATG
AGGTCAAGCATA
TGC
BSU
LV
B113seq16
60
GAACTG
ACAGTA
GCGCAATC
G943
6282–7224
B113seq17 (c)
GACTTGGCTTGCCTG
AGTA
TCG
BSU
M
B94seq3
50
GACGAGCTG
CAAGCTC
TCAGG
467
973–1439
B93seq5 (c)
TGTG
CCTA
TTCTG
AGGTTGG
Musa
actin
ActinDNAf
50
CTG
GTG
ATG
GTG
TGAGCCAC
664
ActinDNAr (c)
CATG
AAATA
GCTG
CGAAACG
(c) den
otes primers that anneals to the complementary strand.
33
where a consistent, novel RFLP was observed, representative samples were
selected for cloning and sequencing.
3.2.4 Cloning and sequence analysis
PCR/RCA products were extracted from agarose gels as previously
described (section 2.2.6). A 4.5 µl aliquot of gel‐purified PCR product was
ligated into pGEM®‐T Easy vector (Promega) at 4 °C overnight as
recommended by the manufacturer. RCA fragments were ligated into 3 µl
of appropriately digested (section 2.2.4), alkaline phosphatase‐treated
pUC19 (section 2.2.7), using 0.5 U T4 DNA ligase (Promega) and 10× ligation
buffer in 20 µl volumes, and incubated at 4 °C overnight. Ligation reactions
were transformed into competent Escherichia coli as described in section
2.2.9 and plasmid DNA was isolated (section 2.2.10) and subsequently
restriction digested (section 2.2.4), to confirm the presence of the desired
insert.
DNA sequencing was carried out as described in section 2.2.11. In
the case of PCR amplicons, three samples which tested positive for a
specific BSV were cloned and sequenced to confirm amplification of the
expected virus. For each sample (PCR or RCA derived), inserts from at least
three independent clones were sequenced. For PCR products, universal
m13 primers were used to sequence plasmid inserts, whereas for RCA
products either universal primers or the degenerate badnavirus primers
(Badna‐FP/RP; Yang et al., 2003) which anneal in the RT/RNaseH‐coding
region were used.
Sequences were analysed using the Vector NTI Advance suite
v.10.3.0 (Invitrogen Corp.). ContigExpress was used to trim vector
sequences and assemble sequences from clones of each sample. The
identity of cloned fragments was determined by comparison to published
sequence information in National Centre for Biotechnology Information
database (http://www.ncbi.nlm.nih.gov) using the Basic local alignment
34
search tool (BLAST) programs. For RCA fragments, classification of
sequences as BSV species was based on sequence comparisons using a
region of the RT/RNaseH‐coding region, delimited by either the Badna‐FP
or Badna‐RP primers (Yang et al., 2003) where possible. Badnavirus
sequence differences within the RT/RNaseH‐coding region of more than
20% are considered to be distinct Badnavirus species (King et al., 2011),
and this was used for identification by matches to published sequences. In
samples where the sequence obtained was outside the RT/RNaseH‐coding
region, identification was made by sequence identity to published full‐
length BSV sequences where similarity was greater than 80%. In the case of
PCR products, the sequences were compared against the specific region of
the BSV genome that PCR primers were known to amplify (Table 3.2).
35
3.3 Results
3.3.1 Sample collection
Leaf samples were collected from 45 plants from nine districts in Eastern
Uganda during February, 2010 (Table 3.1, Figure 3.1). Of the 45 samples, 44
were recorded as showing some symptoms associated with BSV infection
such as chlorotic and necrotic flecks or yellow blotches, while one sample
was asymptomatic (Figure 3.2). Symptom expression ranged from very mild
flecking to strong chlorotic and necrotic flecking. Of the 45 samples, three
samples were from triploid cultivars derived of both A‐ and B‐genome
origin, specifically AAB (samples 216 and 217) and ABB (230) while the
remainder were triploid cultivars of pure A‐genome origin (AAA).
3.3.2 PCR results
Using primers specific for nine BSV species (BSCAV, BSGFV, BSIMV, BSMYV,
BSOLV, BSUAV, BSUIV, BSULV and BSUMV), 38 of the 45 samples tested
positive for one or more BSV (Table 3.3). The extraction control actin PCR
test was positive for all samples. Of the 38 PCR positive samples, five
samples (189, 219, 224, 230 and 231) tested positive for more than one
BSV species. Sample 230 (ABB genome) tested positive for all four BSV with
known integrated counterparts, while dual infections were detected in the
other four samples (all of which had A‐only genomes). Seven samples (195,
196, 200, 204, 215, 218 and 227) did not test positive despite only one
sample (215) originating from symptomless leaves.
PCR screening for BSMYV resulted in three positive samples (216,
217 and 230), all of which have a B‐genome component. BSOLV was
detected in five samples (188, 189, 190, 219 and 230), while BSGFV was
detected in one sample (230) and BSIMV was detected in three samples
(219, 230 and 231). These four BSV, all with a known integrated
counterpart, were all found in sample 230, which has an ABB genotype,
however the remaining samples in which these viruses were detected
36
Figure 3.1: Districts in eastern Uganda where leaf samples were collected:(A) Luweero, (B) Kamuli, (C) Jinja, (D) Mbale, (E) Manafwa, (F) Manafa, (G)Bududa, (H) Sironko and (I) Kapchorwa.
37
Figure 3.2: Symptoms in leaf samples collected
in eastern Uganda. (a) Sam
ple 205 displayed m
ild chlorotic flecks, (b) sample 215 w
assymptomless, (c) sam
ple 221 displayed yellow blotches and (d) sample 226 displayed chlorotic and necrotic flecks.
a)
b)
c)
d)
38
Table 3.3: A
nalysis of leaf samples for BSV
by PCR and seq
uen
cing
Sample
number
PCR results
Sequence ID
BSCAV
BSG
FV
BSIMV
BSM
YV
BSO
LV
BSU
AV
BSU
IV
BSU
LV
BSU
MV
Species
% identity
188
‐ ‐
‐ ‐
+ ‐
‐ ‐
‐ BSO
LV
98%
189
‐ ‐
‐ ‐
+ ‐
+ ‐
‐ BSO
LV
BSU
IV
95%
95%
190
‐ ‐
‐ ‐
+ ‐
‐ ‐
‐ BSO
LV
91%
191
‐ ‐
‐ ‐
‐ ‐
‐ ‐
+ BSU
MV
91%
192
‐ ‐
‐ ‐
‐ ‐
‐ +
‐ BSU
LV
94%
193
‐ ‐
‐ ‐
‐ ‐
‐ +
‐ BSU
LV
94%
194
‐ ‐
‐ ‐
‐ ‐
‐ +
‐ BSU
LV
88%
195
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
196
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
197
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
198
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
199
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
200
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
201
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
202
‐
‐ ‐
‐ ‐
‐ ‐
+ ‐
203
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
204
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
205
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
206
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
207
‐ ‐
‐ ‐
‐ ‐
+ ‐
‐ BSU
IV
94%
208
‐ ‐
‐ ‐
‐ +
‐ ‐
‐ BSU
AV
91%
209
‐ ‐
‐ ‐
‐ +
‐ ‐
‐ BSU
AV
93%
39
Sample
number
PCR results
Sequence ID
BSCAV
BSG
FV
BSIMV
BSM
YV
BSO
LV
BSU
AV
BSU
IV
BSU
LV
BSU
MV
Species
% identity
210
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
211
‐ ‐
‐ ‐
‐ +
‐ ‐
‐ BSU
AV
86%
212
‐ ‐
‐ ‐
‐ +
‐ ‐
‐ BSU
AV
88%
213
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
214
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
215
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
216
‐ ‐
‐ +
‐ ‐
‐ ‐
‐
217
‐ ‐
‐ +
‐ ‐
‐ ‐
‐
218
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
219
‐ ‐
+ ‐
+ ‐
‐ ‐
‐ BSIMV
100%
220
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
221
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
222
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
223
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
224
‐ ‐
‐ ‐
‐ +
+ ‐
‐ BSU
IV
95%
225
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
226
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
227
‐ ‐
‐ ‐
‐ ‐
‐ ‐
‐
228
‐ ‐
‐ ‐
‐ +
‐ ‐
‐
229
‐ ‐
‐ ‐
‐ ‐
‐ +
‐
230
‐ +
+ +
+ ‐
‐ ‐
‐
231
‐ ‐
+ ‐
‐ ‐
‐ ‐
+ BSIMV
BSU
MV
98%
98%
232
‐ ‐
‐ ‐
‐ ‐
‐ ‐
+ BSU
MV
91%
Total
0
1
3
3
5
15
3
12
3
40
contain only A‐genomes. Four BSVs were only detected in samples from
cultivars with AAA genotypes. BSUIV was detected in three samples (189,
207 and 224), as was BSUMV (191, 231 and 232). BSUAV was detected in
15 samples (203, 208–214, 220–225 and 228) and BSULV in 12 samples
(192–194, 197–199, 201, 202, 205, 206, 226 and 229). The remaining BSV
species, BSCAV, was not detected in any of the leaf samples tested.
To confirm the specific detection of each BSV using PCR, the
amplicons from selected samples were cloned and sequenced. PCR
amplicons from samples positive for BSMYV and BSGFV were not cloned, as
these two BSV were only detected in cultivars which contain a B‐genome.
For the remaining six BSV species detected using PCR, three to four samples
were sequenced (Table 3.3). BLASTN searches performed using the
sequences from each PCR product confirmed the specific detection of each
BSV, with a sequence identity of greater than 86% to the reference
sequence in all cases (Table 3.3).
3.3.3 RCA results
Of the 45 samples analysed, 40 samples were considered positive for BSV
infection using RCA (Table 3.4). In total, 25 of these 40 samples were
analysed by cloning and sequencing of RCA products. BLAST searches of the
sequences resulted in a sequence identity of at least 91% for all samples,
except those in which BSULV was identified, for which sequence identity
was in the order of 82–92% (Table 3.4). Interestingly, all cloned fragments
matched significantly (i.e. >80%) to a published BSV sequence.
Five samples (200, 215, 218, 228 and 230) were considered RCA
negative due to the absence of restriction fragments following digestion
with PstI, StuI, EcoRI, XbaI, BamHI, HindIII, SacI and SalI. Of these five
samples, one sample (215) was from an asymptomatic plant with the
remaining four samples derived from leaves showing characteristic BSV
symptoms. Samples 200, 215 and 218 also tested negative for nine BSV
41
Table 3.4: A
nalysis of leaf samples for BSV
by RCA and RFLP/sequen
cing
Sample
number
RE digest pattern (kb
p)
Fragments sequenced
RFLP
match to
sample
Sequence ID
StuI
PstI
RE used
Ban
d lengths
(kbp)
Species
%
identity
StuI
PstI
188
FLa , 6, 5, 4.2, 3.1, 2.5, 1.8, 1.6
4.5, 3, 3×
2.5
+
4.5
3
2.5
BSU
MV
BSO
LV
94%
92%
98%
189
‐ FL, 5, 2.5
+
5
BSO
LV
96%
190
FL, 6, 5, 4.2, 3.9, 3.1, 2.9, 2.5, 2.4, 1.8, 1.6,
1.5
4.5, 3, 3×
2.5
188
BSU
MV
BSO
LV
191
6.5, 4.5, 2.5,1.7
‐ +
2.5
BSU
MV
93%
192
‐ 6
193
BSU
LV
193
‐ 6
+
6
BSU
LV
89%
194
FL, 5
FL
N/A
b
195
FL, 3
FL
+
FL
BSU
LV
83%
196
FL
FL
N/A
197
FL, 5
FL
N/A
198
FL
6
193
BSU
LV
199
FL
6
193
BSU
LV
200
‐ ‐
201
3, 2.5, 2
6, 5, 2.8, 1.8
+
6
BSU
LV
92%
202
‐ 6, 2.8, 1.2
+
6
BSU
LV
89%
203
FL
‐ +
FL
BSU
AV
94%
42
Sample
number
RE digest pattern (kb
p)
Fragments sequenced
RFLP
match to
sample
Sequence ID
StuI
PstI
RE used
Ban
d lengths
(kbp)
Species
%
identity
StuI
PstI
204
6, 2
FL
+
FL
–c
205
FL
‐
N/A
206
FL
FL, 5
+ FL
BSU
LV
87%
207
FL
4, 3.5
N/A
208
FL
4.5, 3
+
FL
BSU
AV
93%
209
FL
2×
3
+
3 3
BSU
AV
91%
93%
210
FL
2×
3
209
BSU
AV
211
5.2, 2.4,
FL
+
FL
BSU
AV
94%
212
5, 2.4
FL
213
BSU
AV
213
5, 2.4
FL
+
5
BSU
AV
94%
214
5, 2.4
FL
213
BSU
AV
215
‐ ‐
216
FL
FL
+
FL
BSM
YV
99%
217
FL
FL
+
FL
BSM
YV
98%
218
‐ ‐
219
4.5, 2.5, 2, 1.2, 0.9
FL, 5, 2.7
+
FL
BSIMV
98%
220
5, 2.4
4.5, 3.2
+
4.5
3.2
BSU
AV
96%
93%
221
5, 2.4
4.5, 3
+ 4.5
BSU
AV
94%
222
5, 2.4
FL
213
BSU
AV
43
Sample
number
RE digest pattern (kb
p)
Fragments sequenced
RFLP
match to
sample
Sequence ID
StuI
PstI
RE used
Ban
d lengths
(kbp)
Species
%
identity
StuI
PstI
223
FL
FL
+
FL
BSU
AV
94%
224
5, 2.4
FL
213
BSU
AV
225
5, 2.4
FL
+
FL
BSU
AV
93%
226
FL
7, 1
+ 7
BSU
LV
92%
227
‐ 5, 2.7
+
5
BSU
LV
85%
228
‐ ‐
229
FL
FL
+
FL
BSU
LV
82%
230
‐ ‐
231
6, 5, 2.4, 1
FL, 4.5, 3.5
232
BSU
MV
232
6, 2
x 3, 1.5
FL, 4.5, 3.5
+
FL
4.5
3.5
BSU
MV
91%
93%
91%
Identified
35
a
FL represents single‐cut, full‐length BSV
bands of 7.5–8
kbp.
b
“N/A” indicates samples not analysed
by BLAST or where RFLP m
atch were not possible.
c Not virus DNA
44
species by PCR, while sample 230 (ABB genome) only tested positive for the
four BSV species with known eBSVs and was therefore likely to be a false
positive due to amplification of integrated sequences. Only sample 228,
which was PCR positive for BSUAV, could be considered PCR positive and
RCA negative.
BSOLV was identified in three samples (188, 189 and 190; all PCR
positive for BSOLV). For samples 188 and 190, identification was based on
predicted RFLPs from published BSV sequences (2635, 2442 and 2312 bp
digested with PstI; 3111, 2436 and 1842 bp digested with StuI) and was
confirmed by sequencing one of the 2.5 kbp BSOLV fragments created by
PstI digestion from sample 188. Although a characteristic RFLP was not
observed with sample 189, BSOLV was identified in this sample following
sequencing of a 5 kbp PstI fragment, which showed 96% similarity to the
published BSOLV sequence.
BSIMV was only identified in one sample (219; also PCR positive for
BSIMV) by sequence analysis of a full‐length PstI fragment, while BSMYV
was only identified in two samples (216 and 217; also PCR positive for
BSMYV) by sequence analysis of full‐length StuI fragments from both
samples.
BSUMV was identified in three samples (188, 191 and 232; 191 and
232 were PCR positive for BSUMV) based on sequence analysis of 4.5 and
a 3 kbp PstI fragments for sample 188, a 2.5 kbp StuI fragment for
sample 191 and FL, 4.5 and 3.5 kbp PstI fragments for sample 232.
BSUMV was also suspected in two others (190 and 231) based on RFLP
matches (231 was PCR positive for BSUMV). Based on the presence of 4.5
and 3 kbp PstI fragments in sample 190 (in addition to BSOLV‐specific
fragments therein) which are identical to the two BSUMV fragments
sequenced in sample 188, in addition to a number of matching StuI
fragments in both samples, this sample probably also contains BSUMV.
45
Similarly the presence of FL, 4.5 and 3.5 kbp PstI fragments in sample
231, confirmed by sequencing in sample 232 to be BSUMV, suggests the
presence of BSUMV in this sample.
BSUAV was identified in nine samples (203, 208, 209, 211, 213, 220,
221, 223 and 225; all of which were PCR positive for BSUAV) based on
sequence analysis of the full‐length StuI fragments from samples 203, 208
and 223, the two 3 kbp PstI fragments from sample 209, the full‐length
PstI fragment from 211, the 5 kbp StuI fragment from sample 213, the 4.5
kbp and 3.2 kbp PstI fragments from sample 220, the 4.5 kbp PstI fragment
from sample 221 and the full‐length PstI fragment for sample 225.
Additionally, BSUAV was suspected in five other samples (210, 212, 214,
222 and 224; all of which were PCR positive for BSUAV) based on RFLPs.
Sample 210 possessed identical StuI/PstI RFLPs to sample 209 while
samples 212, 214, 222 and 224 displayed identical StuI/PstI RFLPs to
samples 211 and 213. Of the 15 BSUAV PCR positive samples, only sample
228 was not considered positive using RCA.
BSULV was identified in eight samples (193, 195, 201, 202, 206, 226,
227 and 229; all except 195 and 227 were PCR positive for BSULV) based on
sequencing of PstI digest fragments (Table 3.4) and was also suspected in
three samples (192, 198 and 199; all of which were PCR positive) based on
the presence of a 6 kbp PstI fragment which is identical to the fragment
observed in samples 193, 201 and 202 (Table 3.4). All sequences showed
more than 82% similarity to the published sequence. However, in contrast
to BLAST sequence analysis to all other BSVs, samples 193, 202 and 206
displayed a large insertion of 100–200 bp where the insertion sequences
matched no sequences in GenBank when conducting a BlastN search. Of
the 12 BSULV PCR positive samples, three samples (194, 197 and 206) with
full‐length digest fragments, consistent with samples confirmed to be
infected with BSULV, were not analysed by cloning and sequencing.
46
In the remaining samples (194, 196, 197, 204, 205 and 207), all of
which were considered RCA positive for BSV infection, either full‐length
digest fragments or distinct profiles precluded identification based on
RFLPs. Cloning was either unsuccessful in these samples or, in the case of
sample 204, the sequence obtained was not of badnavirus origin.
47
3.4 Discussion
In this chapter, banana leaf samples from eastern Uganda were analysed to
identify the BSV species present, and to potentially detect new species of
BSV. Eight species of BSV were detected by PCR using species‐specific
primers. BSCAV was not detected in any of the 45 samples, while BSGFV
and BSMYV were only detected in cultivars with a B‐genome component.
BSULV and BSUAV were the most prevalent BSV species detected with 15
and 12 PCR positive samples each, respectively. When tested by
RCA/sequencing, six known species of BSV were identified while no novel
BSV sequences were identified. This is the first report of episomal detection
of these six species from eastern Uganda, and is the first report of the
detection of BSMYV in field samples from Uganda.
Detection of BSV using RCA relies on either detection of predicted
RFLPs, or cloning and sequencing of RCA fragments. As demonstrated with
samples of both BSUAV and BSULV studied in this project, a reliance on
distinct RFLPS for identification can be difficult due to sequence variability
and thus, in many cases, sequencing is necessary for definitive
identification. The broad sequence diversity is further illustrated in samples
where RCA/sequencing identified a BSV species, but PCR using species‐
specific primer did not (e.g. BSUMV in samples 188 and 190). Whether the
failure of PCR reflects sequence diversity in the primer binding sites is not
certain, but users of PCR‐based diagnostics must be mindful of this.
In three samples (194, 197 and 206) identified as BSULV, an
insertion of approximately 100–200 bp was found which showed no
homology to published sequences. This finding is not unexpected
considering the highly variable nature of badnavirus genomes, particularly
since their location was outside the coding regions. These three isolates
may represent a novel isolate and this warrants further investigation.
48
Several discrepancies were found when comparing the results from
PCR and RCA analyses. In two cases, a PCR product was amplified from
samples but the samples were considered negative by RCA due to the
absence of restriction fragments with a suite of restriction enzymes.
Conversely, in six samples testing positive by RCA, PCR either didn’t
generate an amplicon or a different BSV species was detected. The exact
reasons for this are not known. At present, the sensitivity of RCA as a
diagnostic method has not been demonstrated, particularly in comparison
to PCR. That RCA has less sensitivity for detection of low levels of virus
could help explain the failure of RCA to detect all samples detected using
PCR. A loss of restriction sites due to sequence variability is another
possibility, while limited cloning/sequencing of samples could explain why
PCR detections were not all confirmed in RCA analysis. On the other hand,
three samples which were considered positive for BSV infection using RCA
were not PCR positive using any of the nine primer sets. The reasons for
this could include sequence variability affecting primer binding or the
presence of novel BSV species for which primers are not available.
Both RCA and PCR have inherent advantages and disadvantages.
PCR is an inexpensive, robust, highly specific and sensitive method for
detecting BSV, and is suitable for diagnosing known BSV species for which
primers have been made. However, it faces problems with the generation
of false positives derived from sequences integrated in the Musa B‐genome
and with detection of uncharacterised species. RCA on the other hand has
the advantages of being able to target a broad spectrum of viruses,
independent of whether the sequence is known beforehand. RCA has
issues, however, with cost and with inconclusive species determination.
RFLP profiles for characterised species of BSV found in Uganda, along with
BSMYV, have been made by James et al. (2011a, b). Whereas these
previous studies were focussed on the characterisation of BSV species in
Western Uganda, the present study focussed on BSV isolates from Eastern
49
Uganda. The experience from the present work, however, is that the
restriction profiles for some BSV species are quite variable, likely due to the
hypervariable BSV genomic sequences. RCA is therefore more suitable for
non‐specific broad‐spectrum detection (a simple yes/no assay) and
investigation of potential new species.
One option to improve PCR‐based diagnostics for BSV would be to
develop primers capable of detecting episomal BSMYV, BSIMV, BSOLV and
BSGFV in banana cultivars with a B‐genome component. DNA sequence
information is now available for the loci where endogenous BSOLV and
BSGFV are present within the M. balbisiana genome (Geering et al., 2005a,
b; Gayral et al., 2008), thus opening the possibility of designing PCR primers
which do not detect the eBSV sequence, and thereby specifically detect the
episomal forms of these BSV species. Additionally, since little information
exists on the sensitivity of RCA for detection of BSV infection, screening of a
large number of infected plants by both PCR and RCA should be undertaken
to assess/compare the reliability and sensitivity of both methods for BSV
diagnosis.
50
51
Chapter 4
Development of episomal‐specific PCR assays for the detection of banana streak viruses with
endogenous counterparts
4.1 Introduction
Banana streak virus (BSV) species are serologically and genomically
variable (Lockhart and Jones, 2000b; Harper et al., 2002a) and this creates
problems for the development of reliable diagnostic tests. Several BSV
species are known to have some or all of their genome integrated into the
Musa balbisiana genome. Integrated DNA of three species, namely Banana
streak OL virus (BSOLV), Banana streak GF virus (BSGFV) and Banana streak
IM virus (BSIMV) have been shown to be activated under stress conditions
to cause episomal infection (Harper et al., 1999b; Ndowora et al., 1999;
Lheureux et al., 2003; Geering et al., 2005a, b; Gayral et al., 2008; Côte et
al., 2010; Iskra‐Caruana, 2010). A fourth BSV species, Banana streak MY
virus, is known to have at least part of its sequence integrated into the M.
balbisiana genome (Geering et al., 2005a, b), and there is speculation this
BSV species can also be activated from integrated sequences to cause
infection. Although PCR primers for the detection of these four BSV species
have been published (Harper et al., 1999b; Geering et al., 2000; James et
al., 2011b), they are not able to discriminate between episomal and
endogenous sequences present in the M. balbisiana genome, and so their
use can result in false positives by detecting eBSV.
The development of a diagnostic method which can overcome the
dependence on antisera‐based approaches and is cost‐favourable for use in
developing countries would be useful. In a recent study (James, 2011)
52
comparing PCR and RCA for the detection of field isolates of BSV, PCR was
demonstrated to be a suitable method for the detection of all characterised
species in bananas with A only genomes. In contrast, in bananas with some
B genome component, PCR was only suitable for detecting species without
endogenous counterparts. DNA sequence information is now available for
the loci where endogenous BSOLV and BSGFV are present within the M.
balbisiana genome. These integrated sequences contain multiple partial
fragments in repeated units with discontinuous and inverted portions
amongst banana genomic sequences at the integrated sites (Geering et al.,
2005a, b; Gayral et al., 2008). Based on the discontiguous nature of these
integrated BSV sequences it may be possible to design PCR primers which
do not detect the eBSV sequence, and thereby specifically detect the
episomal form of BSV. Therefore, the objective of work described in this
chapter was to attempt to develop PCR‐based diagnostic assays for the
specific detection of episomal DNA for BSV species with integrated
counterparts.
53
4.2 Methods
4.2.1 Developing episomal‐specific primers for BSOLV and BSGFV
BAC clone sequences of the M. balbisiana genome which contain eBSOLV
and eBSGFV are publicly available (GenBank accession numbers AF106946
and AP009334 for eBSOLV, and AP009325 and AP009326 for eBSGFV). The
published eBSV sequences were mapped onto published episomal virus
sequences of BSOLV and BSGFV (GenBank accession numbers AJ002234
and AY493509, respectively) using VectorNTI suite v.10.3.0, and analysed
for potential primer sites which may specifically amplify episomal DNA of
each BSV. Putative primer binding sites were assessed visually following
mapping of the integrated DNA sequences onto the episomal form of each
BSV species. By comparing the annealing sites on episomal DNA and
binding sites on the sequence from the integrated locus, primers were
selected which would specifically amplify episomal DNA, or would give a
distinct product size difference when detecting the two templates.
4.2.2 Developing episomal‐specific primers for BSMYV and BSIMV
For BSMYV and BSIMV, sequences of the respective eBSV in the M.
balbisiana genome are not available. Therefore, a selection of primers
available from previous studies in the Centre for Tropical Crops and
Biocommodities (CTCB), QUT (kindly provided by Dr Anthony James and Ms
Bojana Bokan) (Table 4.1) with annealing sites distributed across the virus
genomes, were assessed in different combinations. Primers were mapped
to the full‐length BSMYV and BSIMV genomes (GenBank accession numbers
AY805074 and HQ593112, respectively) to allow combinations of primers to
be selected.
54
Table 4.1: Primers obtained from QUT for detection of BSIMV and BSMYV Target virus Primer name Sequence (5'–3') Reference
BSIMV
B107seq2 GTCCTTGAATCCTGCAATCC (c)
Anthony James, unpublished
B108seq1 TTCCTCTGAAGCAATTATCC (c)
B108seq2 CCAGAACATACGTTATGCCB108seq3 CAATCCAGATTCTTCCTTC (c)B108seq4 GAATGCACCTGCCATCTTCCB108seq5 CTTGATGTCATTGAGTTCTTCC (c)B108seq6 CCAGCGGCAGTTACAATCCGB109seq1 CTTTGTTAACTCTGGATTCCB109seq2 ATGGAAGTAGTCATTGAGG (c)B109seq3 GAATTCATCATCATCTTCGAG (c)B109seq4 CCATCAAAGTCTTTGAGGACCCB109seq5 GCCCGATGAGCCAGTGAGC (c)B109seq6 GGCTTCCAGTACAACATCCB109seq7 CCAGAAGCTTCCTGTTCTGGB109seq8 CCCCAAGACAAGCTTACATB109seq9 ATGCAAGTCTACTTACACAGC (c)B109seq10 GTTCAAACATGAAGATTTCGGB109seq12 TGAAGGGGAAGATAACTACC
BSMYV
Mys‐F3 GAAACTCCAAGATCAAGCTTCAGCC Anthony James, unpublished
BSVIII_SalI_F_ GTCGACACAGGACCACTGCCATGATTGTTG
Bojana Bokan, unpublished
BSVIII_SalI_R_ GTCGACTGGATGAGGGTCTGATCACTTTCA (c)
BSVIII_SphI_F_ GCATGCACCTACTATATCACCAGCCAGCCG
BSVIII_SphI_R_ GCATGCTCTGCATCTTTCGACAACTTTAGT (c)
BSVIII_BstXI_F_ CCATGGAAGTGGACCTAGCTGAAGGCAATC
BSVIII_BstXI_R_ CCACTTCCATGGCTGATATGATAGACCGAT (c)
BSV_ORF1_F GAATTC ATGGATACTTACTGGGATAAAACC
BSV_ORF1_R CTCGAG TCATCCAATAATAACTTTCTCCAG (c)
BSV_ORF2_F CTCGAG TCATTGTAGGGATCTTAGAATTTC
BSV_ORF2_R GAATTC ATGAGTCTAGCCAACACCAAGGCT (c)
BSV_ORF3_F CAATTG ATGACAACTCGAAGGTCCAGTCTA
BSV_ORF3_R CTCGAG CTAACCGGGATCTCCTCCGTCAGG (c)
(c) denotes primers that anneals to complementary strands.
55
4.2.3 Plant samples and nucleic acid extraction
Plant samples used in this study were obtained from several different
sources:
1. Leaf samples of bananas that had been collected during field surveys
in Uganda (Ug) in 2010, and Kenya (Ke) in 2009 and comprised 65
and 41 samples, respectively.
2. A collection of 33 leaf samples from a wide range of banana cultivars
maintained in tissue culture (TC) were obtained from the Maroochy
Research Station of the Department of Employment, Economic
Development and Innovation (DEEDI), Queensland.
3. Fifty‐two nucleic acid extracts prepared from leaf samples were
obtained from the Global Musa Genomics Consortium (GMGC;
www.musagenomics.org).
For the samples collected in Kenya in 2009, the 33 TC samples obtained
from DEEDI and 20 of the 65 samples from Uganda collected in 2010, DNA
extracts (CTAB method) were already available and had been analysed for
BSV by RCA and PCR using all the available published primer sets by
Anthony James (CTCB, QUT). The remaining 45 leaf samples were collected
from Uganda in 2010 (Chapter 3). For the GMGC collection, samples were
provided by CIRAD and a quality control PCR for the actin house‐keeping
gene had been previously carried out (Anthony James, CTCB).
4.2.4 PCR and RCA analysis
Generally, PCR analysis was undertaken as described in section 2.2.2. For
each set of putative episomal‐specific primers designed for BSOLV and
BSGFV, the annealing temperature was optimised by a gradient PCR
conducted on an episomal BSV‐positive extract using an annealing
temperature range of 50 to 70 °C and an extension time of two min (BSOLV)
or one minute (BSGFV). Testing of BSOLV‐ and BSGFV‐specific primer sets
56
was then conducted by screening several collections of plant samples by
PCR using the optimised parameters.
As an initial quick screen to assess the suitability of BSMYV‐ and
BSIMV‐specific primer combinations, four samples were used. These four
samples comprised (i) a positive control sample previously shown using
RCA to be infected with the test BSV species (to test for episomal BSV); (ii)
two episomal BSV‐negative samples from cultivars with some B‐genome
component shown to test positive for the respective eBSV using PCR with
published primers and (iii) a known BSV‐negative healthy sample from an
A‐only genome banana.
For BSMYV, 20 primer combinations were tested while for BSIMV, 25
primer combinations were tested. PCR was carried out using an annealing
temperature of 50 °C and extension time of 4 min (BSMYV) or 3 min
(BSIMV). Following initial primer testing, potential episomal‐specific primer
sets which amplified episomal BSMYV only were chosen and used to screen
selected samples from Ke2009, Ug2010, DEEDI and GMGC sample
collections using PCR with the annealing temperatures of 65 °C or 68 °C and
an extension time of 2 min. Optimised annealing temperatures were
determined for the primer sets selected for BSMYV by gradient PCR, using
an annealing temperature range of 50 to 70 °C with an extension time of 2
min. Further testing of potential episomal‐specific BSIMV primers were
undertaken using the conditions listed above.
To screen for the presence of BSV, and as a comparison to detection
with putative episomal‐specific primer sets, primers RD‐F1/R1, Mys‐F1/R1
and GF‐F1/R1, which specifically detect BSOLV, BSMYV and BSGFV,
respectively (Table 4.2; Geering et al. 2000) were used to test the GMGC
sample collection. This collection was considered to be free of episomal
BSV due to the presumptive virus‐indexed status of the collection and also
to contain accurate data on sample genotypes. Primer annealing
57
Table 4.2: PCR conditions for published
primers used for BSV
and eBSV
detection
Target
Primer
nam
e
Sequence (5'–3')
Amplicon size
on episomal
target (bp)
Region
amplified
(bp)
Amplicon
size on eBSV
(bp)
Annealing
temperature
(°C)
Extension
time
Reference
BSO
LV RD‐F1
ATCTG
AAGGTG
TGTTG
ATCAATG
C
522
6499–7020
522
57
30
s Geering et
al. (2000)
RD‐R1
GCTCACTCCGCATCTTATCAGTC
(c)
eBSO
LVMusa T3‐2
GGCTTATG
ATG
CTG
ACCACAT
Not am
plified
n/a
1931
50
2min
Lhereu
x et
al. (2003)
BSV
510
TTCTCGACCATAAATTG
TAT (c)
BSG
FV GF‐F1
ACGAACTA
TCACGACTTGTTCAAGC
476
6354–6829
476
64
30
s
Geering et
al. (2000)
GF‐R1
TCGGTG
GAATA
GTC
CTG
AGTC
TTC (c)
eBSG
FVVM1‐F
TTG
TCCAAAATCTG
CTCGTG
Not am
plified
n/a
481
50
(Côte et al.,
2010)
VM1‐R
TGTAATTCCTG
CTCCTG
CAA (c)
BSM
YV
Mys‐F1
TAAAAGCACAGCTC
AGAACAAACC
589
6481–7069
589
57
30
s Geering et
al., (2000)
Mys‐R1
CTC
CGTG
ATTTC
TTCGTG
GTC
(c)
BSIMV B107seq1
GCTA
GGAAGAAAAGTC
TGGG
475
7417–122
475
50
30
s James et al.,
(2011b)
B109seq11 TG
CAAGTC
TACTTACACAGC (c)
(c)
den
otes primers that anneals to complementary strands.
58
temperatures and extension times for these two primers sets are listed in
Table 4.2.
For the detection of eBSV loci in plant samples primer sets that
were designed to amplify the border region between the integrated BSOLV
and BSGFV and banana chromosomal DNA (Ndowora, 1998; Lheureux et
al., 2003; Côte et al., 2010) were used. PCR was conducted on the GMGC
collection samples using annealing temperatures and extension times listed
in Table 4.2.
RCA was conducted on the nucleic acid samples obtained from
GMGC to screen for the presence of episomal BSV. RCA was undertaken as
described in section 2.2.3, with RCA products digested using StuI.
PCR and RCA reactions were analysed by agarose gel electrophoresis
as described in section 2.2.5.
59
4.3 Results
4.3.1 Identification of potential episomal‐specific primers
4.3.1.1 BSOLV and BSGFV
A comparison of eBSOLV and eBSGFV sequences with their respective
episomal sequences (Figure 4.1 and Figure 4.2) allowed several potential
episomal DNA‐specific primer sets to be identified (Table 4.3).
For BSOLV, two primer sets were designed to amplify fragments of
1489 and 1249 bp from an episomal template. The sense‐strand primer
annealing sites were chosen at 5240–5261 and 5480–5503 bp for BSOLV‐F3
and ‐F2, respectively. The anti‐sense primer annealing site was located at
6705–6728 for BSOLV‐R2.
For BSGFV, a single primer set was designed which was predicted to
amplify a 729 bp fragment from episomal BSGFV DNA. When designing
primers for BSGFV, the sense primer annealing site was chosen at 6075–
6096 bp while the anti‐sense primer annealing site was located at 6781–
6803 bp.
4.3.1.2 BSMYV and BSIMV
Because the published sequence of integrated BSMYV and BSIMV are not
publicly available, a more random approach was taken for these two BSV.
Primers which were previously available were obtained and mapped onto
their respective BSV episomal genome sequences (Figure 4.3) and then
combinations selected (Table 4.4). Combinations were made to ensure that
all regions of the genome of both viruses were included in the assessment,
with amplicons ranging from 925 to 3937 bp produced in PCRs for BSMYV
and ranging from 837 to 3144 bp for BSIMV (Table 4.4).
60
Figure 4.1: (a) Map of the integrated regions of BSOLV. The integratedBSOLV segments are illustrated by green arrows or green lines for smallfragments. (b) The integrated sequences were mapped onto the full‐lengthgenome. The primer sites are marked as blue lines. As illustrated in a), theydo not bind to a single continuous fragment but are located on fragmentsthat are far apart on the M. balbisiana genome. Primers that anneal to thecomplementary strand are marked “(c)”.
a)
b)
61
Figure 4.2: (a) Map of the integrated regions of BSGFV. The integratedBSGFV segments are illustrated by green arrows or as a green line forsegment #6. (b) The integrated sequences were mapped onto the full‐length genome. The primer sites are marked as blue lines. As illustrated ina), they do not bind to a single continuous fragment but are located onfragments that are far apart on the M. balbisiana genome and bothprimers bind to the complementary strand. Primers that anneal to thecomplementary strand are marked “(c)”.
a)
b)
62
Table 4.3: Potential episomal D
NA‐specific primer seq
uen
ces identified
for BSO
LV and BSG
FV
Target
virus
Primer
nam
e Sequence (5'–3')
Amplicon size
on episomal
target (bp)
Region
amplified
(bp)
Amplicon size
on eBSV
(bp)
Annealing
temperature
(°C)
Extension
time
BSO
LV BSO
LV‐F2
CAAGTA
GCAATG
GACCCAGAGTC
T 1249
1489
5480–6728
5240–6728
2412
2898
61
57
1 2 min
min
BSO
LV‐R2 CCCTG
TCAGGATTTC
TGGATG
TTC (c)
BSO
LV‐F3
GTG
ATC
AGGCCTTCAAGCTC
AA
BSG
FV BSG
FV‐F1 TC
TGTG
CTTATG
CAAGTG
GACG
729
6075–6803
Not am
plified
60
30 s
BSG
FV‐R1 GAGGGTC
GTC
TATG
TCGGAGTTG (c)
(c) den
otes primers that anneals to complemen
tary strands
63
Figure 4.3: Genome of (a) BSMYV and (b) BSIMV with primer sequencesmapped. Primers that anneal to the complementary strand are denoted ‘c’.
a)
b)
64
Table 4.4: Primer combinations and results of initial screen for putative episomal‐specific primers for BSMYV and BSIMV detection
PCR product
length (bp)
Primer selection PCR test results
Primer set #
Forward primer
Reverse primer
Positive control
B‐genome
1a
B‐genome
2
A‐genome
b
BSMYV 1 BSV_ORF1_F BSV_ORF2_R 925 ‐ ‐ ‐ ‐ 2 BSVIII_BstXI_R_ 1486 + ‐ ‐ ‐ 3 BSVIII_SphI_R_ 3581 ‐ ‐ ‐ ‐ 4 BSV_ORF2_F BSVIII_BstXI_R_ 960 + ‐ ‐ ‐ 5 BSVIII_SphI_R_ 3050 ‐ ‐ ‐ ‐ 6 BSV_ORF3_F BSVIII_SphI_R_ 2655 ‐ ‐ ‐ ‐ 7 BSVIII_SalI_R_ 3937 ‐ ‐ ‐ ‐ 8 BSVIII_BstXI_F_ BSVIII_SphI_R_ 2102 + ‐ ‐ ‐ 9 BSVIII_SalI_R_ 3384 + ‐ ‐ ‐ 10 BSVIII_SphI_F_ BSVIII_SalI_R_ 1288 + + + ‐ 11 BSV_ORF3_R 2961 + ‐ ‐ ‐ 12 BSVIII_SalI_F_ BSV_ORF3_R 1679 + ‐ ‐ ‐ 13 BSV_ORF1_R 3274 ‐ ‐ ‐ ‐ 14 Mys‐F1 BSV_ORF1_R 2220 ‐ ‐ ‐ ‐ 15 BSV_ORF2_R 2615 ‐ ‐ ‐ ‐ 16 BSVIII_BstXI_R_ 3170 + ‐ ‐ ‐ 17 Mys‐F3 BSV_ORF3_R 1139 + ‐ ‐ ‐ 18 BSV_ORF1_R 2739 ‐ ‐ ‐ ‐ 19 BSV_ORF2_R 3134 + ‐ ‐ ‐ 20 BSVIII_BstXI_R_ 3689 + ‐ ‐ ‐
BSIMV 1 B107seq1 B109seq5 2151 + ‐ + ‐ 2 B109seq1 B109seq5 1181 ‐ ‐ ‐ ‐ 3 B109seq1 B109seq3 1995 ‐ ‐ ‐ ‐ 4 B109seq7 B109seq5 964 ‐ ‐ ‐ ‐ 5 B109seq7 B109seq3 1778 ‐ ‐ ‐ ‐ 6 B109seq4 B109seq3 1111 + ‐ + ‐ 7 B109seq4 B109seq2 1927 + ‐ + ‐ 8 B109seq1 B109seq2 2811 ‐ ‐ ‐ ‐ 9 B109seq4 B108seq5 2899 + ‐ + ‐ 10 B109seq6 B108seq5 2112 ‐ ‐ ‐ ‐ 11 B109seq6 B108seq3 2677 ‐ ‐ ‐ ‐ 12 B109seq8 B108seq3 1828 + ‐ + ‐ 13 B109seq1 B108seq1 2686 ‐ ‐ ‐ ‐ 14 B109seq12 B108seq3 992 + ‐ + ‐ 15 B109seq12 B108seq1 1850 ‐ ‐ ‐ ‐ 16 B109seq12 B107seq2 2848 + ‐ + ‐ 17 B108seq2 B108seq1 837 ‐ ‐ ‐ ‐ 18 B108seq2 B107seq2 1835 ‐ ‐ ‐ ‐ 19 B108seq2 B109seq11 2827 ‐ ‐ ‐ ‐ 20 B108seq4 B107seq2 1048 + ‐ + ‐ 21 B108seq4 B109seq11 2039 + ‐ + ‐ 22 B108seq6 B109seq11 1428 + ‐ + ‐ 23 B108seq6 B109seq5 3144 + ‐ + ‐ 24 B107seq1 B109seq5 2151 + ‐ + ‐ 25 B107seq1 B109seq3 2990 ‐ ‐ ‐ ‐
a ‘B‐genome 1’ and ‘B‐genome 2’ are samples from cultivars with some B‐genome component previously tested negative for the listed BSV by RCA and positive by PCR using published primers for the listed BSV.
b ‘A‐genome’ is a healthy banana cultivar with an A‐only genome.
65
4.3.2 Primer testing
4.3.2.1 BSOLV
PCR annealing temperatures were initially optimised for the two BSOLV
primer sets (F2/R2 and F3/R2) using gradient PCR on a known episomal
BSOLV‐infected sample as template. The results from this pilot study
showed that the optimal annealing temperatures for amplification of the
expected size amplicons from a known episomal BSOLV template using
primer pairs F2/R2 and F3/R2 were 61°C and 57°C, respectively (Table 4.3).
Following optimisation, primers BSOLV‐F2/R2 were used to test a
total of 106 samples, derived from Ke2009 and Ug2010 survey collections,
which had previously been tested for episomal BSV by RCA and for BSOLV
by PCR using published primers RD‐F1/R1. Of these 106 samples, 17 were
previously shown to be infected with episomal BSOLV using RCA (and all 17
were also PCR positive using published primers RD‐F1/R1). When the 106
samples were tested using BSOLV‐F2/R2, the expected size product of 1.2
kbp was amplified from 16 of the 17 samples previously shown to contain
episomal BSOLV (with only sample Ug190 testing negative). Additionally,
two further samples, Ug230 (from a cultivar with an ABB genome), which
had tested PCR positive using primers RD‐F1/R1, and Ug201 (from a cultivar
of pure A‐genome origin), which had tested PCR negative using primers RD‐
F1/R1, were also positive using BSOLV‐F2/R2. Of the remaining 87 samples
in these two collections (22 of which contained some B‐genome
component), three samples (Ke156, Ke176 and Ug219, all from cultivars of
pure A‐genome origin), which had previously tested BSOLV positive by PCR
using primers RD‐F1/R1, tested negative by PCR using BSOLV‐F2/R2. These
three samples, along with Ug201, were all RCA positive for badnavirus
infection; however, BSOLV was not confirmed in these samples.
Furthermore, Ug230, which tested positive using both RD‐F1/R1 and
BSOLV‐F2/R2, was negative by RCA.
66
To further assess the ability of primers BSOLV‐F2/R2 for specific
detection of episomal BSOLV, the primers were used to screen the DEEDI
collection. Previous RCA analysis had shown that sample 5 contained
episomal BSOLV based on RFLP identification. Additionally, when the DEEDI
collection was screened using published primers RD‐F1/R1, 14 of the 33
samples (all from cultivars with some B‐genome component), including
sample 5, tested positive for BSOLV. PCR screening of the DEEDI collection
using BSOLV‐F2/R2 resulted in four positive samples (5, 10, 12 and 26) all
from cultivars with some B‐genome component, all of which were positive
using primers RD‐F1/R1.
The second potential episomal‐specific primer set, BSOLV F3/R2,
was also tested for their ability to detect BSOLV infection by screening a
subset of 20 samples (233–252) from the Ug2010 survey sample collection.
For these 20 samples, primer set BSOLV F3/R2 gave the same results as
primer set BSOLV F2/R2. Therefore, no further testing of BSOLV F3/R2 was
undertaken. PCR screening resulted in nine positive samples (Ug2010: 237–
240, 242, 243, 245, 247 and 248), comprising two samples from cultivars of
pure A‐genome and seven with some B‐genome component. All nine
samples were considered positive for BSOLV when previously screened by
RCA. Of the remaining 11 samples which tested negative using BSOLV
F3/R2, five of which contain some B‐genome component, none were
previously considered positive for episomal BSOLV using RCA.
PCR screening of the DEEDI samples using BSOLV F3/R2 resulted in
the same four positive samples as primers BSOLV F2/R2, only one of which
was considered positive when screened by RCA.
4.3.2.2 BSGFV
PCR annealing temperatures were initially optimised for the BSGFV
primer set (BSGFV‐F1/R1) using gradient PCR on a known episomal BSGFV‐
infected sample as template. The results from this pilot study showed that
67
the optimal annealing temperatures for amplification of the expected 729
bp amplicons from a known episomal BSGFV template was 60°C (Table 4.3).
Following optimisation, primers BSGFV‐F1/R1 were used to test a
total of 106 samples, derived from Ke2009 and Ug2010 survey collections,
which had previously been tested for episomal BSV by RCA and for BSGFV
by PCR using published primers GF‐F1/R1. Of these 106 samples, 13
samples were known to be infected with episomal BSGFV using RCA (and all
13 were also PCR positive using the published primers GF‐F1/R1). When the
106 samples were tested using the newly developed primers BSGFV‐F1/R1,
the expected size product of 700 bp was amplified from all of the 13
samples previously shown to contain episomal BSGFV (Ke2009: 146, 148,
176, 180 and 181; Ug2010: 239, 240, 242, 244, 246–248 and 250).
Interestingly, a 700 bp product was also amplified from two additional
samples (Ug243 and 251, also positive using primers GF‐F1/R1) from
cultivars with some B‐genome component. When previously screened by
RCA, sample 243 had been considered positive for badnavirus (although
BSGFV had not been identified) while sample 251 had tested negative for
badnavirus. Of the remaining 91 samples previously tested in these two
collections, 22 of which contained some B‐genome component, none were
found to contain episomal BSGFV using primers BSGFV‐F1/R1. The strong
correlation between the results obtained using primers BSGFV‐F1/R1 and
those obtained using RCA suggested that primers BSGFV‐F1/R1 were
promising candidates for PCR‐based detection of episomal BSGFV.
To further test the ability of primers BSGFV‐F1/R1 to specifically
detect episomal BSGFV, the primers were used to test the DEEDI collection.
Previous RCA analysis had shown that none of these 33 samples contained
episomal BSGFV, although 15 samples (all from cultivars with some B‐
genome component) had tested PCR positive for BSGFV using published
primers GF‐F1/R1 suggesting the presence of eBSGFV. When these 33
samples were tested using primers BSGFV‐F1/R1, all samples tested
68
negative, consistent with previous work using RCA and, in contrast to
screening with published primers GF‐F1/R1, no episomal BSGFV was found.
This result further demonstrated the ability of primers BSGFV‐F1/R1 to
avoid detection of eBSGFV in B‐genome banana cultivars.
4.3.2.3 BSMYV
PCR was undertaken with 20 BSMYV‐specific primer combinations to assess
their ability to specifically detect episomal BSMYV (Table 4.4). For initial
screening, four DNA extracts were selected which included one known
BSMYV‐infected Cavendish (AAA‐genome), two from banana cultivars with
some B‐genome component (Ug239 and Ug240) which were negative for
episomal BSMYV but positive for eBSMYV and one from a healthy banana
cultivar with an A‐only genome (Ug236). Importantly, the sample
containing BSMYV, as well as samples Ug239 and Ug240 containing
eBSMYV, all tested positive using the previously published BSMYV
diagnostic primers Mys‐F1/R1.
Of the 20 primer sets selected, 11 of these (numbered 2, 4, 8, 9, 10,
11, 12, 16, 17, 19 and 20; Table 4.4) amplified the expected product from
the known episomal BSMYV‐infected sample. Of these 11, however, one
primer set (number 10) also amplified the expected size product from the
two samples, Ug239 and Ug240, which contained eBSMYV. No products
were amplified from any of the samples using the remaining nine primer
sets (Table 4.4). Primer sets 12 and 17 were selected for further testing
based on the amplification of strong bands of a relatively small size (1703
bp and 1109 bp, respectively). To further optimise these primer sets, the
primers were used to test two known episomal BSMYV positive samples
(Ke149 and Ug249), as well as two RCA‐negative samples with some B‐
genome component (Ug239 and 240). Whereas primer set 17 amplified the
expected 1.1 kbp amplicon in all four samples, the use of primer set 12
resulted in the amplification of the expected 1.7 kbp amplicon only in the
69
two episomal BSMYV‐positive samples, although a weak amplicon of 600
bp was present in sample 239.
Based on the promising results obtained using primer set 12, this
set of primers was subsequently used to screen 41 samples from the
Ke2009 survey sample collection. Of these 41 samples, eight were
previously shown to be infected with episomal BSMYV using RCA.
Additionally, all eight RCA positive samples were PCR positive when tested
with published primers Mys‐F1/R1. PCR screening using primer set 12
resulted in seven positive samples (Ke149, 150, 173, 175, 176, 182 and
184), all of which were previously shown to be infected with episomal
BSMYV using RCA. Of the remaining 34 samples (11 from cultivars with
some B‐genome component), none were considered positive when tested
with primer set 12. The 600 bp band previously observed in the two RCA‐
negative samples from cultivars of some B‐genome component was not
detected in any of the samples.
4.3.2.4 BSIMV
The method for assessing the specificity of primers for detecting episomal
BSIMV was essentially the same as for BSMYV. For initial screening, 25
BSIMV‐specific primer combinations were tested for potential episomal‐
specificity on four DNA extracts including one known episomal BSIMV‐
infected, two from banana cultivars with some B‐genome component
(Ug241 and Ug244) which were negative for episomal BSIMV but positive
for eBSIMV and one from a healthy banana cultivar with an A‐only genome
(Ug236). Samples Ug236, 241 and 244 were all shown to test positive using
the previously published BSIMV diagnostic primers (Table 4.2).
Of the 25 primer sets selected, 12 of these (numbered 1, 6, 7, 9, 12,
14, 16, 20, 21, 22, 23 and 24) amplified the expected products (Table 4.4)
from the known episomal BSIMV‐infected sample. However, all of these 12
primer sets also amplified the same sized products from Ug241, but not
70
Ug244, both of which were known to be negative for episomal BSIMV but
positive for eBSIMV. None of the remaining 13 primer sets amplified
products from any samples (Table 4.4). Based on these results, none of the
25 primers sets were considered specific for the detection of episomal
BSIMV.
To further investigate the reasons for the amplification of the
products produced from sample Ug241 and not Ug244 by the 12 primer
sets listed above, four of the primer sets (1, 6, 14 and 20) were chosen for
further analysis. The primer sets were selected based on either intensity of
amplicon (set 1), small size of amplicons (set 6) or both these criteria (sets
14 and 20), and also because they amplified different regions on the BSIMV
genome. All four primer sets were used to screen 10 samples from the
DEEDI collection (DEEDI: 4, 5, 10, 12, 13, 16, 23, 27, 28 and 31), all of which
were derived from cultivars with some B‐genome component and were
considered negative for BSIMV when tested previously by RCA and/or PCR.
Amplicons of the expected size were detected in eight, 10, nine and six of
the 10 samples using primer sets 1, 6, 14 and 20, respectively. The
detection of the expected product in samples previously demonstrated to
be negative by RCA was highly suggestive of non‐specific amplification,
possibly of eBSIMV, using these four primer combinations. As such further
testing was discontinued.
4.3.3 Final primer assessment using the GMGC collection
To further assess the ability of the putative episomal‐specific primer
pairs BSOLV‐F2/R2, BSGFV‐F1/R1 and BSMYV set 12, all three primer pairs
were used to screen the GMGC collection. This collection was chosen as it
was believed to include a virus‐indexed, true‐to‐type collection comprising
a broad genetic base of Musa germplasm. As such, it was expected to
provide a reliable sample set with which to assess the episomal‐specificity
of the primers.
71
When the 52 samples from the GMGC collection were screened by
PCR using primers BSGFV‐F1/R1, 11 positive samples (GMGC: 1–4, 9, 10,
27, 32, 34, 45 and 46) comprising one sample (32) from a cultivar of pure A‐
genome and ten with some B‐genome component, were PCR positive.
Using primer set BSOLV‐F2/R2, three positive samples (12, 18 and 20;
Figure 4.4a) were detected, all of which contained some B‐genome
component. Using primer set 12 for BSMYV detection, the expected size
1.7 kbp amplicon was generated in 21 samples (GMGC: 1–5, 9–12, 17, 18,
20, 21, 34, 36, 39–41, 45, 46 and 48), all of which possessed some B‐
genome component. The large number of positive samples was
unexpected, as this collection was considered to be free of episomal BSV
infection.
In order to clarify the episomal virus status of the GMGC collection,
a subset of samples was tested using RCA. The ten B‐genome samples
which were PCR positive using BSGFV‐F1/R1 as well as the three samples
which were PCR positive using BSOLV‐F2/R2, were RCA‐amplified and the
reactions digested using StuI to detect episomal BSV DNA. Of the 10 BSGFV
PCR‐positive samples tested, four (2, 3, 34 and 45) produced digest profiles
indicative of BSGFV infection (6034 and 1229 bp), while the remaining six
samples did not produce any digest products. Of the three samples which
tested positive using BSOLV‐F2/R2, no positive identification of BSOLV could
be made using RCA, based on known RFLPs for this virus. However, clear
bands of approximately 3 kbp in samples 12 and 18 and 5 kbp in sample 20,
along with several smaller bands in these samples were detected, which
could indicate that BSV infection associated with a distinct RFLP profile may
be present. This finding indicates that, although the GMGC sample
collection was originally believed to be free of episomal BSV infection, this
was clearly not the case. Therefore, the detection of many PCR positive
samples using the primers developed in this study could be considered to
72
Figure 4.4: Screen
ing of the GMGC collection using (a) the episomal specific primer sets, BSO
LV F2–R
2, and (b) the published
primer sets,
RD‐F1–F2. “M” den
otes marker lanes, “+” is positive control and “NTC
” is a negative template control.
73
be a reflection of episomal BSV infection in some samples, despite not all
testing positive for episomal BSV in this instance.
As a comparison to the results obtained using the putative
episomal‐specific primers, published primers for the detection of BSOLV,
BSGFV and BSMYV, which detect both episomal and eBSV sequences, were
used to screen the 52 samples in the GMGC collection. Using primer set
RD‐F1/R1 (for detection of BSOLV), a total of 30 samples (1–5, 9–13, 15,
17–21, 23, 27, 28, 30, 36, 39–41 and 43–48) tested positive (Figure 4.4b),
comprising five samples from A‐only genotype cultivars (13, 15, 19, 43 and
44), 24 samples from cultivars with some B‐genome component and
sample 30 with an AS genotype. However, the positive A‐genome cultivar
samples produced weak amplicons compared to samples with some B‐
genome component. These primers, not intentionally designed to avoid
amplifying eBSOLV, thus amplified 24 out of the 26 samples from cultivars
having some B‐genome component in this collection. In contrast, only three
of these 24 B‐genome samples were amplified using the primer sets BSOLV‐
F2/R2 (12, 18 and 20).
Using primer set GF‐F1/R1 (for detection of BSGFV), a total of 28
samples (1–5, 9–13, 18, 20, 21, 23, 27, 28, 30, 32, 34, 36, 39–41 and 43–47)
tested positive, comprising four samples from A‐only genotype cultivars
(13, 32, 43 and 44), 23 samples from cultivars with some B‐genome
component and again sample 30 with an AS genotype. However, the
positive A‐only cultivar samples again displayed weak amplicons compared
to samples with some B‐genome component. In contrast, 11 of these 28
positive samples were positive when tested using primer set BSGFV‐F1/R2,
including sample 32 with a AAA genome.
Using primer set Mys‐F1/R1 (for detection of BSMYV), a total of 27
samples (1–5, 9–13, 17–21, 23, 27, 28, 34, 36, 39–41, 44–46 and 48) tested
positive, comprising three samples from A‐only genotype cultivars (13, 19
74
and 44) and 24 samples from cultivars with some B‐genome component.
However, the positive A‐only cultivar samples again displayed weak
amplicons compared to samples with some B‐genome component. In
comparison, 21 samples from cultivars with some B‐genome component
were amplified using the putative episomal‐specific primer set 12, all of
which were positive using Mys‐F1/R1.
As these three published primer sets were not designed to be
selective between episomal and integrated BSV sequences, it was expected
that all samples which contain a B‐genome could test positive for BSV using
these primer sets, as a result of amplification of eBSV sequences. However,
the detection of BSV in A‐only genome samples was of concern. Since
activation of BSV from eBSV sequences is only known to occur in B‐genome
banana cultivars, the detection of BSV in A‐only genome samples using the
published primers, but not using the putative episomal‐specific primer sets
developed herein, suggested that the newly designed primers were not
detecting episomal infection in some samples.
To further resolve this issue, primer sets were obtained which were
specific to the integrated loci of both BSGFV and BSOLV and these were
also used to test the 52 samples in the GMGC collection. The occurrence of
positive results in DNA samples using these primer sets should be indicative
of the presence of B‐genome DNA, as eBSV loci are only known to occur in
bananas which contain some B‐genome component. PCR was carried out
using two published primer sets specific to the junction of eBSV/Musa
genomic sequences of the integrated loci of eBSOLV and eBSGFV.
PCR screening using the eBSOLV‐specific primer set, Musa‐T3‐
2/BSV510, resulted in a total of 28 positive samples, comprising four
samples (6, 7, 13 and 19) of A‐only genomes and 24 samples from cultivars
with some B‐genome component (1–5, 9–12, 17, 18, 20, 21, 23, 27, 34, 36,
39–41, and 45–48). The amplicons produced from the A‐only genome
samples were very faint, while the eBSOLV primer set failed to amplify two
75
B‐genome samples (28 and 31). This result suggested that Musa B‐genome
DNA was present in at least four samples considered to contain A‐only
genome DNA, while in two samples either B‐genome DNA was not present,
or these distinct cultivars do not contain eBSOLV.
PCR screening of the GMGC collection using the eBSGFV‐specific
primer set, VM1‐F/R, also resulted in a total of 28 positive samples,
comprising three samples of A‐only genomes (33, 43 and 44), 24 samples
from cultivars with some B‐genome component (1–5, 9–12, 18, 20, 21, 23,
27, 28, 34, 36, 39–41, and 45–48), and sample 30 (AS‐genome). The
amplicons for the A‐genome cultivars were again very faint. The eBSGFV
primer set failed to amplify two B‐genome samples (17 and 31), and in one
sample (48), a distinct fragment of approximately 1.2 kbp was amplified
instead of the expected 481 bp amplicon. Consistent with previous findings
using primers for eBSOLV, PCR using primers for eBSGFV indicated that B‐
genome DNA was present in several samples considered to have A‐only
genomes, while several samples with B‐genomes were not positive.
In summary, testing of the GMGC collection using primers which
were specific for eBSV loci indicated that the true‐to‐type status of this
collection, which originally made this sample set attractive for screening for
episomal BSV specificity, was not assured. Further, the demonstration that
this collection contained episomal BSV in several samples tested
confounded efforts to make an accurate assessment of the specificity of the
putative episomal‐specific primers developed in this study.
76
4.4 Discussion
The aim of this chapter was to design primers which specifically
amplify episomal virus DNA for the four BSV species which have integrated
counterparts within the M. balbisiana genome. This was theoretically
possible because, at least for two BSV species (BSOLV and BSGFV), the
endogenous sequences exist as multiple discontinuous partial fragments
for which BAC clone sequences were available (Geering et al., 2005a, b;
Gayral et al., 2008). However, for BSMYV and BSIMV, information on the
structure of the integrated sequences was not available and primer design
in this case was essentially random.
BAC clone sequences of the integrated versions of BSOLV and BSGFV
were mapped onto the genomes of their corresponding episomal viruses,
and potentially episomal‐specific primer sets were identified. The initial
screening for BSOLV in previously‐indexed field samples using BSOLV‐F2/R2
generated promising results with 16 out of 17 samples known to contain
episomal BSOLV detected, and a large number of B‐genome samples not
giving rise to the expected PCR product. Several additional samples were
also positive where BSV infection was present but BSOLV had not been
confirmed by RCA analysis. Similar results were obtained when screening
the DEEDI collection, with a known episomal BSOLV‐infected sample testing
positive and the majority of B‐genome samples not. However, the
detection of a positive result in several samples is of interest as this may
indicate either episomal BSOLV infection in these samples, or the presence
of an alternate, uncharacterised eBSOLV locus which amplified. If episomal
BSOLV is present, then PCR using BSOLV‐F2/R2 may be more sensitive for
BSV detection than RCA.
Similar promising results were obtained when primer pair BSGFV‐
F1/R1 was used to test previously‐indexed field samples, with 13 out of 15
PCR‐positive samples previously identified as infected with episomal
BSGFV. Of the two additional field samples which tested positive, one
77
sample was RCA positive, but a different BSV was identified (BSOLV), and
one sample from a cultivar of some B‐genome component was RCA
negative. Of the remaining 91 samples, none were previously shown to be
positive for episomal BSGFV. When the BSGFV‐F1/R1 primer set was used
to screen the DEEDI collection no samples tested positive, which was
consistent with previous work to detect episomal BSGFV using RCA. These
results indicated some success in avoiding detection of eBSGFV in B‐
genome samples using these primers, since both the field collections and
DEEDI collection contained a range of banana cultivars with some B‐
genome component where no eBSGFV was detected.
Of an initial 20 primer combinations screened for BSMYV detection,
11 appeared promising with BSMYV‐free B‐genome samples not amplified
in the initial test, despite amplification in the positive control sample. Of
these, two primer sets, 12 and 17, which seemed the most ideal with
respect to amplicon strength and size, were investigated further. Since set
17 was subsequently found to amplify eBSMYV in the following PCR
optimisation step, only set 12 was used further to screen field samples.
When testing the Ke2009 collection (41 samples), seven out of eight
samples with episomal BSMYV were detected although only weak
amplicons were generated. While the primers selected showed promise,
further work would be required to give acceptable PCR‐based detection.
Initial tests using the available BSIMV primer sets resulted in
amplification from one of the two episomal BSIMV negative samples used
as negative controls, implying amplification of eBSIMV. To investigate the
possibility that the positive B‐genome sample could prove to be an
exception, 10 BSIMV RCA‐negative samples with some B‐genome
component were used to test four of the 25 primer sets. The primer sets
each amplified between six and ten of these samples which suggested that
the primers were not suitable for discrimination of episomal and
endogenous BSIMV.
78
To further assess the episomal‐specificity of BSOLV, BSGFV and
BSMYV primers, a reference collection (GMGC) of 52 banana samples from
a diverse range of cultivars (including 26 cultivars with some B‐genome
component) was analysed. The high confidence in cultivar identification in
this sample set, along with the BSV‐indexed status, should have allowed a
straightforward assessment of primer specificity for episomal targets.
Using BSOLV F2/F3–R2, three of the 52 samples were positive (from
cultivars with some B‐genome component) while all remaining samples
tested negative. These three samples were later screened by RCA and all
three samples were considered potentially BSV positive. Screening of the
GMGC collection using BSGFV‐F1/R1 resulted in 11 positive samples (10
from cultivars of some B‐genome component). The 10 samples from B‐
genome cultivars were screened by RCA using a single restriction enzyme
and four were considered BSGFV positive. These results were unexpected
since all samples were presumed to be BSV free, as international collections
are virus‐indexed. However, the maintenance of bananas as TC accessions
can lead to the activation of eBSV sequences and episomal infections. With
13 PCR positive samples from B‐genome cultivars it is not unreasonable to
postulate that BSV infection in these samples arose from activation of eBSV.
Although episomal BSV infection was not confirmed in all 14 PCR‐positive
samples, only a single RCA reaction and digest were attempted and further
work may identify episomal BSV in other samples.
Twelve of the 13 GMGC samples screened using RCA were also PCR
positive for BSMYV using primer set 12. None of these 12 samples
produced the expected approximately full‐length band previously
associated with a BSMYV infection in RCA analysis. While this could mean a
loss of the StuI restriction site due to mutation of the BSMYV genome or a
difference in sensitivity between RCA and PCR, it is also possible that these
positive results using primer set 12 were caused by the amplification of
eBSMYV sequences.
79
The detection of eBSV sequences in A‐genome samples (using
primers for eBSOLV and eBSGFV), as well as detection of BSV sequences in
A‐genome samples, complicated the analysis. The presence of BSV
sequences in A‐genome samples suggests that either (i) contamination with
B‐genome cultivar’s DNA has occurred, (ii) previous virus indexing failed or
(iii) cultivar identification should be called into question. These issues could
be addressed by (i) obtaining samples from the same cultivars but from an
alternative source and/or (ii) conduct further testing using the putative
eBSV‐specific primers to confirm their specificity. Further, the virus status of
the source material should be independently validated.
RCA is a relatively new diagnostic method for badnavirus detection
and no information exists on its sensitivity, for example where virus
infection is at a low concentration. It must therefore be considered that
PCR results in RCA‐negative samples may reflect a difference in sensitivity
of PCR and RCA for BSV detection. A comprehensive assessment of the
sensitivity of RCA compared to PCR would shed light on this aspect of
episomal BSV diagnosis. Designing PCR primer sets that can distinguish
between integrated and episomal BSV, by mapping the integrated
sequences onto their episomal counterpart appears possible. The potential
existence of eBSVs of different conformity from the published sequences
(for BSGFV and BSOLV) may be an issue and further genomic sequencing of
Musa germplasm would shed light on this, as well as allow characterisation
of the eBSMYV and eBSIMV loci.
80
81
Chapter 5
Development of an infectious clone of Banana streak MY virus
5.1 Introduction
An important issue faced by Australian‐based BSV researchers is the limited
availability of diseased plant material. While BSV is widespread in East
Africa, the disease is sporadic in Australia due largely to the
implementation of strict quarantine and disease control programs. Further,
due to the slow rate of transmission by the mealybug vector, the inability to
mechanically transmit the virus (Jones and Lockhart, 1993; Kubiriba et al.,
2000; Huang and Hartung, 2001) and difficulties in importing diseased
material due to quarantine restrictions, obtaining large numbers of infected
plants is time consuming and laborious. The development of an infectious
BSV clone would overcome these difficulties.
Successful development of infectious clones of the tungrovirus, Rice
tungro bacilliform virus (RTBV), and the badnaviruses, Citrus yellow mosaic
virus (CYMV) and Sugarcane bacilliform virus (SCBV), have been reported
(Dasgupta et al., 1991; Bouhida et al., 1993; Huang and Hartung, 2001). The
model proposed for production of a successful infectious clone (for
Badnavirus) involves the expression (following entry to the plant cell) of a
larger‐than‐unit‐length, terminally redundant transcript of the viral
genomic DNA by host DNA‐dependent RNA polymerase, using the cloned
virus DNA as template. This greater‐than‐full‐length transcript is produced
during normal virus infection and is the template for viral minus‐strand
DNA synthesis using the virus‐encoded replicase protein as well as a
polycistronic RNA for expression of the virus encoded proteins (Pfeiffer and
Hohn, 1983; Hohn et al., 1985; Temin, 1989; Medberry et al., 1990;
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Dasgupta et al., 1991; Qu et al., 1991; Bouhida et al., 1993; Jacquot et al.,
1999). Delivery methods for the infectious DNA molecule into intact plant
cells can involve particle bombardment or Agrobacterium‐mediated
transformation.
Researchers within the CTCB at QUT have recently developed rolling
circle amplification (RCA) as an assay for the detection of a wide range of
episomal BSV species in infected plants (James, 2011; James et al., 2011a,
b). However, a regular supply of infected material is now required in order
to optimise the assay and allow comparative analyses to other assay
formats such as polymerase chain reaction (PCR). The research described in
this chapter was aimed at developing an infectious clone of a BSV species
endemic in Australia, namely Banana streak MY virus (BSMYV), to enable
the generation of large numbers of infected plants for use in future
diagnostic assay optimisation and validation. Further, since the badnavirus
SCBV has been shown to infect banana (Bouhida et al. 1993), the ability of
the BSV clone to infect sugarcane was also investigated.
83
5.2 Methods
5.2.1 Preparation of the infectious clone
The strategy for the construction of an infectious clone of BSMYV was
based on previously reported strategies for other members of the
Caulimoviridae family, with minor modifications. This strategy utilises a
terminally redundant cloned DNA molecule which contains the intergenic
region (which possesses the regulatory sequences for genome expression)
at both ends of the molecule. The terminally redundant virus sequence was
amplified in two pieces by RCA, with a common site selected within the
coding portion of the genome allowing the two molecules to be joined with
an uninterrupted reading frame. The two BSMYV fragments were amplified
by RCA (Section 2.2.3) using gDNA extracted from a BSMYV‐infected
banana leaf sample (Section 2.2.1) as template. Restriction sites for
digesting the RCA‐amplified DNA were determined based on the published
sequence of the BSMYV genome (GenBank No. AY805074; Figure 5.1).
Following amplification, the products from the two reactions were double‐
digested either with XmaI and SalI or with SalI and StuI. The pOpt‐EBX
vector (kindly supplied by Don Catchpoole) was chosen as the binary vector
as it contains suitable restriction sites in the multiple cloning site. Vector
DNA was digested with XmaI and StuI as described in section 2.2.4, except
that the digest was incubated overnight. Following digestion, the vector
was analysed using agarose gel electrophoresis (section 2.2.5), gel‐purified
(section 2.2.6) and de‐phosphorylated (section 2.2.7).
The two virus‐derived fragments were ligated into linearised pOPT‐
EBX vector DNA by mixing pOpt‐EBX DNA (10 µl, 14 ng/µl), linearised
XmaI and SalI‐fragment DNA (23 µl, 6.9 ng/µl) and SalI and StuI‐fragment
DNA (13 µl, 6.4 ng/µl), 10× ligation buffer (6 µl), T4 DNA ligase (3 µl, 1
U/µl, Promega) and adding distilled water to a total volume of 60 µl. The
ligation mixture was incubated at 4 °C overnight. The reaction was
84
Figure 5.1: Creation of the BSMYV infectious clone. Two fragments wereproduced by digestion of RCA‐amplified BSMYV DNA (green). Thesefragments were ligated together into the pOpt‐EBX binary vector, creating aterminally redundant BSMYV insert in the vector. Functional codingsequences (CDS) are coloured in orange, while the flanking, inactive CDSfragments at each end of the insert are cross‐hatched.
85
transformed into 300 μL of competent E. coli cells and the transformed
cells selected on LB with kanamycin (50 µg/ml; Section 2.2.9). Plasmid DNA
was isolated and screened for the BSMYV insert DNA using StuI and SmaI
digestion (Sections 2.2.10 and 2.2.4). Additionally, to confirm the predicted
ligation across the SalI site at position 5422 in the BSMYV genome, PCR was
done using primers BSVIII_Sph1_F and BSV_ORF3_R (Table 4.1) at an
annealing temperature of 55 °C and an extension time of 3 min. Selected
clones were sequenced across all three restriction sites to confirm insert
presence and orientation, and (for the SalI ligation site) to confirm the
integrity of the ORF3 reading frame. Sequencing was conducted using
primers NPTII‐Int‐R (5'‐GGTCACGACGAGATCCTCGCCGTCGG‐3') over the
XmaI‐site, RB (5'‐GCTGCACTGAACGTCAGAAGC‐3') over the StuI‐site and
Mys‐F2 (5'‐CAGGGGTATATCGGGGAAGAG‐3') over the SalI‐site as described
in section 2.2.11.
After confirming the integrity of the infectious clone in pOPT‐EBX,
the cloned DNA was transformed in Agrobacterium. Miniprep DNA (30 µl)
was precipitated as described in the mini‐preparation protocol (Section
2.2.10) and dissolved in deionised water (10 µl). The DNA was then added
to competent Agrobacterium tumefaciens strain AGL1 cells (50 µl). The
mixture was transferred to a pre‐chilled electroporator cuvette (path length
= 2 mm) (BioRad) and pulsed in an EC100 electroporator (Sigma‐Aldrich) at
2.8 kV (14 kV/cm) for 5 ms then immediately suspended in 1 ml of SOC. The
culture was then incubated at 28 °C for approximately 2 h with shaking.
Transformed cells were selected on LB agar with kanamycin (50 µg/ml) and
rifampicin (25 µg/ml), and incubated for two days at 28 °C. One colony
from each plate was selected and incubated in 1 ml of liquid LB with
kanamycin (50 µg/ml) and rifampicin (25 µg/ml) at 28 °C for two days and
plasmid DNA was isolated as described in section 2.2.10. As described
above, PCR was used to confirm the presence of the recombinant vector
86
DNA (Section 2.2.2) using the primers Mys‐F2 and BSV_ORF3_R (Table 4.1)
with an annealing temperature of 50 °C and an extension time of 2 min.
5.2.2 Plant inoculation and assessment
Banana and sugar cane plants were inoculated using recombinant
Agrobacterium harbouring the cloned BSMYV DNA. Tissue cultured banana
plants (cultivar Williams‐Cavendish) and setts from field grown sugar cane
plants (cultivar Q155) were planted in premium grade potting mix
(Searles®, Australia) and grown to the 3rd–4th leaf stage. A mixed inoculum
was prepared by combining three individual Agrobacterium cultures
(originating from three distinct colonies) into liquid LB (50 ml)
supplemented with kanamycin (50 µg/ml) and rifampicin (25 µg/ml) which
was incubated for two days at 28 °C. The culture was centrifuged after
incubation to precipitate cells, and the media was removed. The cell pellet
was re‐suspended in MMA‐solution (10 ml; 10 mM MES, 10 mM MgCl2, 200
µM acetosyringone) and incubated at room temperature for 2‐3 h before
inoculation. The plants were inoculated by three methods:
1. Corm inoculation – 200‐500 µl of inoculum was injected into the
corm of five banana plants.
2. Leaf infiltration – 200‐500 µl of inoculum were pressed into the
abaxial surface of the leaf using a 1 ml syringe on two banana plants.
3. Small needle‐pricks (approximately 20) were made along the
pseudostem of five banana plants and into the approximate
meristematic region between the third and fourth fully‐expanded
leaves of four sugar cane plants, and 10‐15 µl of inoculum injected
at each site.
The plants were grown in a plant growth room at a temperature of
28 °C, a relative humidity of 70% and illumination at 350 µmol
photons/(m2∙s) with a 16 h photoperiod. The plants were observed at 5 and
87
12 weeks post‐inoculation, and leaf samples collected 12 weeks post‐
inoculation and analysed by PCR and RCA.
PCR screening was conducted using BSMYV‐specific primers Mys‐
F1/R1 (Table 4.2) as described in section 2.2.2. RCA was conducted as
described in section 2.2.3 and reaction products were digested using
SmaI/StuI as described in section 2.2.4. To test for the presence of residual
A. tumefaciens, a PCR test was performed using the primers VCF‐AGL1 (5’–
GCCTTAAAATCATTTGTAGCGACTTCG–3’) and VCR‐AGL1 (5’–
TCATCGCTAGCTCAAACCTGCTTCTG–3’) as described in section 2.2.2 with an
annealing temperature of 63 °C and extension time of 30 s.
88
5.3 Results
5.3.1 Preparation of the infectious clone
RCA‐amplified BSMYV DNA was digested in two separate reactions using (i)
StuI/SalI and (ii) XmaI/SalI to produce fragments of 6564 and 3247 bp,
respectively. The fragments were gel purified and ligated into XmaI/StuI‐
digested pOpt‐EBX vector DNA, yielding a terminally redundant BSV
fragment of 9811 bp (or 1.3 × BSMYV genome) in the binary vector (a
total recombinant plasmid length of 18040 bp). Recombinant plasmid DNA
was subsequently transformed into E. coli and colonies screened for the
presence of both inserts by restriction analysis using StuI and SmaI to
produce fragments of 5489 bp and 2161 bp (Figure 5.2). The presence of
the cloned inserts was verified by PCR amplification across the SalI ligation
site and the correct alignment confirmed by sequence analysis over all
three ligation sites. DNA from three colonies confirmed to contain the
complete terminally redundant BSV molecule was then transformed into
Agrobacterium tumefaciens strain AGL1 for plant infection studies.
5.3.2 Plant inoculation and assessment
Banana plants were inoculated using three different methods while sugar
cane plants were inoculated using the needle‐prick method only (Table
5.1). None of the three uninoculated banana plants developed any
symptoms (Figure 5.3). At five weeks post‐inoculation, two out of five
banana plants inoculated using the corm‐injection method showed mild
leaf streaking, while the remaining banana and sugar cane plants were
symptomless.
At 12 weeks post‐inoculation (Table 5.1), the two banana plants
inoculated using the leaf infiltration method remained free from symptoms
of BSV infection, although the infiltrated leaves were senescing and dying
(Figure 5.4). Three of the five needle prick‐inoculated banana plants
showed mild leaf streaking on the newest leaves, while one plant remained
89
Figure 5.2: Restriction digest of plasmid DNA isolated from 13 E. colicolonies. Lanes 1, 5 and 8 show the presence of the predicted 5489 bp and2161 bp inserts. The 8 kbp band represents the pOpt‐EBX vector. “M”denotes molecular weight marker lanes.
a)
90
Table 5.1: A
nalysis of inoculated plants
Species
Cultivar
Inoculation
method
Plant no.
Symptoms (12 weeks)
Analysis
BSM
YV
(PCR)
Agrobacterium
(PCR)
Bad
navirus
(RCA)
Musa
hybrid
Cavendish
(Williams)
Corm
injection
1
Strong leaf streaking, stunting
+ ‐
+
2
Wilted
/dying
+ ‐
Failed
3
Strong leaf streaking, stunting
+ ‐
+
4
Strong leaf streaking, stunting
+ ‐
+
5
Strong leaf streaking, stunting
+ ‐
+
Needle
pricking
1
Weak leaf streaking
+ ‐
+
2
No sym
ptoms
‐ ‐
‐
3
Dead
+ ‐
+
4
Weak leaf streaking
+ ‐
+
5
Very weak leaf streaking
+ ‐
+
Leaf
infiltration
1
No sym
ptoms
‐ ‐
‐
2
No sym
ptoms
‐ ‐
‐
Saccharum
hybrid
Q155
Needle
pricking
1
No sym
ptoms
a (+)
a ‐
‐
2
No sym
ptoms
(+)
‐ ‐
3
No sym
ptoms
(+)
‐ ‐
4
No sym
ptoms
‐ ‐
‐
a Brackets indicates very weak band
91
Figure 5.3: (a) Healthy banana plant with (b) green leaves.
92
Figure 5.4: Banana plant at 12 weeks post leaf‐infiltration with the BSM
YV infectious clone showing no disease sym
ptoms. The infiltrated
leaf (circled) is sen
escing.
93
symptomless and the remaining plant was dead (Figure 5.5). All of the five
corm‐injected banana plants were smaller than the healthy control plants.
Of these five plants, four showed strong leaf streaking symptoms (Figure
5.6), while the remaining plant was stunted and dying. Of the four
inoculated sugar cane plants, none showed any symptoms of BSV infection
at 12 weeks post‐inoculation (Figure 5.7).
At 12 weeks post‐inoculation, a leaf sample was taken from each
inoculated plant and tested for the presence of BSV by PCR and RCA. A leaf
sample was also taken from a healthy banana plant (Williams‐Cavendish
cultivar) to use as a control for PCR and RCA screening (Figure 5.8). Using
PCR, a band of the size expected for BSMYV (589 bp) was amplified from
extracts derived from all seven banana displaying typical BSV symptoms.
Further, the dead plant inoculated using the needle prick method and one
wilted/dying plant inoculated by corm injection also tested positive (Table
5.1, Figure 5.8). When samples from the inoculated sugar cane plants were
tested by PCR, very weak bands of the expected size were amplified from
three of the four plants. To confirm that the results were due to the
presence of BSV and not due to residual A. tumefaciens, extracts were also
tested for the presence of Agrobacterium by PCR. These were compared to
a positive control using gDNA extracted from untransformed A. tumefaciens
(strain AGL1) as template. While a band of the expected size (approximately
800 bp) was amplified from the positive control, all plants tested negative,
indicating it is unlikely that residual agrobacteria were present.
Extracts from all plant samples were also tested by RCA to confirm
the PCR results. Bands of the sizes expected for BSMYV (2159 bp and 5491
bp) were obtained from extracts derived from all seven banana displaying
typical BSV symptoms. Further, one dead plant inoculated using the needle
prick method, also tested positive. The RCA reaction failed for the
wilted/dying plant inoculated by corm injection. No product of the
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expected band sizes was obtained for the four inoculated sugar cane plants
when tested by RCA (Figure 5.8).
Three symptomatic corm injected plants and the three symptomatic
needle pricked plants, all shown to be BSMYV infected by PCR and RCA, were
kept in the growth room for further observation. Seven months post‐
inoculation they all showed strong leaf streaking (Figures 5.9 & 5.10), with the
three corm‐injected plants still expressing the strongest symptoms.
95
b)
Figure 5.5: Needle‐prick inoculated banana plant at 12 weeks postinoculation (a) showing mild leaf streaking on the newest leaves (b).
a)
b)
96
Figure 5.6: B
anana plant at 12 weeks post corm
injection (a) showing strong leaf streaking on m
ost leaves (b).
b)
a)
97
Figure 5.7: Sugar cane plant at 12 weeks post needle‐prick inoculation (a).None of the leaves expressed any symptoms of disease (b).
a) b)
98
Figure 5.8: Screen
ing of inoculated banana and sugarcane plants for BSM
YV by PCR (a) and RCA (b) . “M
” den
otes marker lanes, “H
S” is
sample from a healthy banana plant, “+” is positive control and “NTC
” is a negative template control.
99
Figure 5.9: N
eedle‐prick inoculated banana plant seven m
onths post inoculation. The plant expressed
clear leaf streaking symptoms.
100
Figure 5.10: Corm
injected
banana plant seven m
onths post in
oculation. This plant expressed
the strongest leaf streaking symptoms, and
was stunted compared to the non‐inoculated controls.
101
5.4 Discussion
In this chapter, an infectious clone of BSMYV was generated using an RCA‐
based cloning strategy. Unlike previously described strategies, which have
necessitated the purification of virions in order to extract genomic DNA for
manipulation, in this study the BSV genomic DNA was amplified using RCA
prior to further manipulation thus avoiding the difficulties often associated
with virus purification. BSMYV was chosen because this BSV species is
endemic to Australia.
Since banana is not a natural host for A. tumefaciens (De Cleene and
Ley, 1976; Escobar and Dandekar, 2003), it was considered that the most
aggressive Agrobacterium strain available would increase the likelihood of
successful infection. As such, Agrobacterium tumefaciens strain AGL1 was
used as the host strain for plant inoculation in this study as this strain is
routinely used in our laboratory for banana transformation (Hoekema et al.,
1983, Lazo et al., 1991, Khanna et al., 2004). The infectivity of the putative
infectious clone was tested on both banana and sugarcane. The AAA‐
genome Cavendish cultivar was chosen as the test banana cultivar because
(i) BSMYV is known to infect this cultivar and produce characteristic
symptoms (Geering et al., 2000) and (ii) the absence of integrated copies of
the BSMYV genome in banana cultivars derived from Musa acuminata
facilitates the subsequent detection of BSMYV using PCR‐based testing. The
infectivity on sugarcane was also examined since successful infection of
banana by SCBV has been previously reported (Bouhida et al. 1993).
Three different inoculation methods were used based on those used
previously for other members of Caulimoviridae. The corm injection
method was based on the method described by Boulton et al. (1989),
which was used for successful agroinoculation of rice and banana plants
with RTBV (Dasgupta et al. 1991) and SCBV (Bouhida et al. 1993),
respectively. The needle pricking and leaf infiltration methods were
102
modified from the stem slash and needle press inoculation methods
described in Huang and Hartung (2001) for successful agroinoculation of
sweet orange with CYMV.
Twelve weeks post‐inoculation, four of the five banana plants
inoculated by corm injection expressed characteristic leaf streaking, as well
as stunting, while one plant had almost died. The presence of BSMYV was
confirmed in all five plants using PCR and these results were verified using
RCA with the exception of one plant for which RCA failed. Four out of five
needle‐prick inoculated bananas showed some leaf streaking on the
newest leaves at 12 weeks post‐inoculation and again infection was
confirmed in the symptomatic plants. Bananas inoculated using leaf
infiltration did not show symptoms of BSV infection.
Of the three inoculation methods used, corm injection was the most
effective with a 100% infection rate. Banana plants inoculated using this
method were also the first plants to express symptoms of BSV infection.
Although the reason for the high infectivity using corm injection is
unknown, the fact that the corm contains a large amount of rapidly dividing
meristematic tissue which is conducive to viral replication is a possible
explanation. The use of needle prick inoculation was also reasonably
efficient with four out of five inoculated plants becoming infected.
However, compared to corm‐inoculated plants, the symptoms were slower
to develop and no stunting was observed. Due to the limited availability of
plants, only two plants were inoculated using the leaf infiltration method.
Although leaf infiltration yielded no infection, the fact that only two plants
were tested precludes any meaningful comparisons to be made. However,
banana leaves were difficult to infiltrate due to the brittleness of the leaf
tissue which resulted in damage to the leaves and this may account for the
lack of infectivity. Given the results from this study, it is clear that corm
injection was the most reliable approach for infecting banana plants, and
this method is recommended when conducting further experiments.
103
For sugar cane, only a single inoculation method, needle pricking of
the meristem, was attempted due to limited availability of plants at the
time of the experiment. No disease symptoms were observed on any
sugarcane plants and all plants tested negative for BSMYV by RCA at 12
weeks post‐infection. However, very weak bands of the size expected for
BSMYV were amplified from three samples using PCR, suggesting very low
levels of infection with BSMYV. These results clearly indicate a need for
further investigation. Previous work has demonstrated that, whereas noble
sugar canes (Saccharum officinarum) can show symptoms of badnavirus
infection (eg. SCBV) commercial hybrids usually do not (Lockhart and
Autrey, 2000). This might explain why symptoms were not observed on the
commercial hybrid, Q155. The differences observed in the results of PCR
and RCA testing may simply reflect differences in the sensitivity of the two
methods or might suggest non‐target amplification in PCR. Despite previous
studies demonstrating badnavirus transmission across species (between
sugar cane and banana) (Bouhida et al., 1993; Reichel et al., 1997) and
recent studies which show that certain species of SCBV and BSV are so
closely related that they barely can be considered separate species (Muller
et al., 2011), cross‐species infection of plants using BSMYV could not be
conclusively demonstrated in this study.
To establish greater knowledge on the efficiency of the inoculation
methods and propensity for infection, a range of different banana and
sugarcane cultivars could be included in follow‐up studies. Further, electron
microscopic examination of sap or partially purified extracts from
inoculated plants is needed to demonstrate the presence of virions.
In summary, this study demonstrates that agro‐inoculation is an
effective approach for delivery of an infectious clone to banana.
Importantly, this method now provides a fast and efficient means of
generating large numbers of BSV‐infected plants for comparative testing of
RCA and PCR as diagnostic methods.
104
105
Chapter 6
General discussion and conclusions
Banana is an important part of the diet for millions of people in Sub‐
Saharan Africa, in particular Kenya and Uganda (Jones, 2000; Karanja et al.,
2008). However, pests and diseases are an enormous constraint to banana
production in these countries. The best strategy for preventing pests and
diseases of banana in the East African region is considered to be
distribution of disease‐ and insect‐free planting material obtained by tissue
culture (TC) techniques. However, TC faces issues with viruses, which exist
intracellularly and could, in cases where a virus infection are present in low
concentrations, not be detected during TC. In theory, a single plant can be
the progenitor to offspring in the millions using TC. Because viruses can be
retained in plants during TC, infected plants can be accidentally distributed.
Banana streak disease, caused by BSV, is widespread in Kenya
(Karanja et al., 2008) and Uganda (Tushemereirwe et al., 1996; Harper et
al., 2002a, 2004, 2005). Additionally, BSV species are a serious constraint to
the international distribution of Musa germplasm due to poor diagnostic
protocols and the presence of full‐length integrated sequences which can
become activated during TC to cause episomal virus infection. Diagnosing
BSV is difficult due to considerable nucleotide sequence variability in
characterised BSV isolates and the presence of endogenous BSV (eBSV)
sequences present in the Musa genome.
To assess the appropriateness and reliability of PCR and RCA as
diagnostic tests for BSV detection, 45 field samples of banana collected
from nine districts in the Eastern region of Uganda in February 2010 were
tested using RCA and PCR. In total, 40 samples were considered positive for
106
BSV infection by RCA and 38 by PCR, with episomal infection of six BSV
species confirmed. No new species were found and, based on partial
sequences found in Uganda (Harper et al. 2004, 2005), nine previously
reported putative BSV species were not detected. Further studies on
banana in East Africa would help confirm the existence of these putative
BSV species. Interestingly, Banana streak MY virus (BSMYV) was identified
in two samples of the cultivar Pisang ceylan (AAB genome) from Bududa.
This is an important result as BSMYV has not previously been reported in
field samples from Uganda.
When screening field samples for BSV infection, PCR proved to be a
better method for detection of the known BSV species, as this assay format
is highly specific to its target sequence and therefore allows identification
of the BSV species without further work. Constraints for PCR‐based
diagnostics remain however, with primers not available for putative BSV
species which have not been fully characterised, and because of the
presence of four endogenous BSV (eBSV) in the Musa B‐genome. To
address the constraints of eBSV, PCR primers that could discriminate
between integrated and episomal sequences of four BSV species were
investigated. Promising results were obtained for Banana streak OL virus
(BSOLV) and Banana streak GF virus (BSGFV), with primer design based on
the known sequence of eBSOLV and eBSGFV sequences. However, for
BSMYV and Banana streak IM virus (BSIMV), where primers were randomly
selected, results were not as encouraging. Characterising the eBSIMV and
eBSMYV within the M. balbisiana genome may enable the design of primer
sets specific to the episomal counterparts of these two BSVs. Additionally,
further efforts to refine and optimise the use of putative episomal‐specific
primers for detection of BSGFV and BSOLV would assist to alleviate the
constraints to PCR created by eBSVs.
In contrast to PCR, RCA proved time‐consuming and laborious for
detection of BSV in field samples. RFLPs for a range of previously published
107
BSV (James, 2011; James et al., 2011a, b) provided little aid to the
characterisation of the BSV isolates studied, as few samples yielded RFLPs
which were previously characterised. Instead, many samples had
approximately full‐length restriction fragments or, in some cases, various
different RFLPs for the same BSV species. In effect, the only way to identify
the BSV species following RCA was to clone and sequence the restriction
fragments. This time‐ and labour‐intensive process makes RCA a vastly
more costly method for diagnosis in field samples compared to PCR, where
specific identification of BSV species is the objective, despite needing to do
many individual PCR assays for the known species of BSV. However, RCA still
presents certain advantages over PCR including the non‐sequence specific
nature of amplification, which makes it suitable for detecting and
identifying new BSV species and mutated versions of known BSVs.
Additionally, for screening of field collected plant material for future
micropropagation it would be more time and cost effective to use RCA
because, from a practical perspective, it is not relevant to know which
species of BSV infects the plant material, only to know whether it is
infected or not, so that infected plant material can be discarded.
For plants in TC systems that have been screened initially with a
non‐specific broad‐range assay such as RCA, PCR could still serve a purpose.
Only four eBSV species are believed to cause episomal infection as a result
of activation during TC. If episomal‐specific primer sets for all four BSVs
with an integrated counterpart were developed, PCR would provide a time
and cost effective way of conducting routine screening of plant material
prior to distribution from TC, since only specific detection of these four
species, which can be activated during TC, is warranted.
While RCA has been demonstrated to detect a wide range of BSV
species and to specifically detect the episomal form of BSV DNA in infected
plants (James, 2011; James et al., 2011a, b), this method is a recently
utilised assay format and there is no published information regarding the
108
sensitivity of RCA compared to other assay formats. Furthermore, a central
issue faced by researchers in Australia when developing and optimising
newly developed assays (e.g. RCA) for BSV detection is the limited
availability of diseased plant material due to the implementation of strict
quarantine and disease control programs. Mealybugs are the only known
vector for BSV, but transmission is very slow (Jones and Lockhart, 1993;
Kubiriba et al., 2000; Huang and Hartung, 2001), which makes obtaining
large numbers of infected plants time consuming and laborious. Thus, in
order to provide a method for the rapid production of large numbers of
BSV‐infected plants as a means to assist further research and optimisation
on diagnostic methods for BSV detection, an infectious clone of a BSV
species endemic in Australia, namely BSMYV, was successfully generated.
Agro‐inoculation of the corm was found to be the most effective
way of inoculating banana with the BSMYV infectious clone, although
needle‐prick inoculation of the stem was also successful. These initial test
results potentially allow for the effective infection of a wide range of
banana cultivars for further studies on the sensitivity of new diagnostics
assays for BSV detection. Importantly, the development of a BSMYV
infectious clone opens the possibility for using virus‐induced gene silencing
(VIGS) as a tool for studying gene function in this important crop plant.
VIGS offers an attractive and quick alternative for disrupting the
expression of a gene, without the need to genetically transform the plant
(Baulcombe, 1999). Using VIGS, a recombinant virus carrying a sequence of
a host gene is used to infect the plant. When the virus infects and spreads
systemically the host gene is expressed by the infecting virus. The host
plant responds by degrading the expressed sequence (and its endogenous
counterpart) in a process termed post‐transcriptional gene silencing (PTGS)
(Baulcombe, 1999). The result of PTGS is to effectively ‘silence’ the
endogenous gene expression and so studies of gene function can be made
by assessing the loss of function as a result of PTGS (Ratcliff et al., 1997;
109
Schöb et al., 1997; Baulcombe, 1999; Liu et al., 2002; Verchot‐Lubicz, 2002;
Zhang et al., 2009; Purkayastha et al., 2010). Successful studies utilising
VIGS have been conducted using Rice tungro bacilliform virus (RTBV)
(Purkayastha et al., 2010), a rice‐infecting pararetrovirus from the family
Caulimoviridae, demonstrating the general utility for these large double‐
stranded DNA viruses for VIGS studies (Hay et al., 1991; Hull, 1996; King et
al., 2011).
In conclusion, the outcome of this project has been the further
investigation of PCR‐ and RCA‐based diagnostic assays for BSV, with specific
emphasis on the development of PCR‐based assays that can distinguish
between episomal and integrated BSV sequences for BSVs with known
eBSVs. These assays are ideally suited for application in East Africa where
the rapid development of the TC nursery industry has generated concern
over the multiplication and distribution of TC material which has not
undergone virus indexing. Combined with diagnostics for other important
banana viruses, such as Banana bunchy top virus (BBTV), BSV diagnostics
will improve the quality of TC material which is distributed to farmers
throughout this region.
110
111
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