Sigatoka Leaf Spot Disease on Banana Laboratory Diagnostics Manual
Edited by Dr Juliane Henderson December 2006
The PCR primers and reaction conditions for molecular diagnosis of Mycosphaerella musicola and Mycosphaerella fijiensis are pending publication and thus must be held in confidence until such time they appear in the public domain. Please also ensure these parameters do not appear in material including conference oral presentations and posters and funding body reports.
© Copyright 2006
Sigatoka Leaf Spot Disease Diagnostic Manual
Updated December 2006
Contributors
Sharon Van Brunschot André Drenth Kathy Grice Juliane Henderson Julie Pattemore Ron Peterson Susan Porchun
1 Table of Contents
Introduction...... 3 The Banana Industry in Australia ...... 3 Origin and Distribution of Black Sigatoka...... 3 Black Sigatoka in Australia...... 6 Yellow Sigatoka...... 9 Disease Symptoms...... 10 Black Sigatoka...... 10 Yellow Sigatoka...... 13 Other Leaf spots...... 17 The Pathogens ...... 18 The Causal Agents...... 18 Taxonomy of M.fijiensis and M. musicola ...... 18 Morphology ...... 18 Hosts ...... 21 Relative Distributions of M. fijiensis and M. musicola ...... 21 Life Cycle...... 21 Disease Development and Epidemiology ...... 23 Disease Cycle ...... 23 Survival of the Inoculum ...... 23 Spread of the Pathogen ...... 25 Disease Control ...... 26 Cultural Control ...... 26 Chemical Control...... 26 Microscopic Diagnosis of Disease ...... 28 The Pathogen...... 28 Molecular Diagnosis of Disease...... 29 Preliminary diagnosis and sampling...... 29 DNA extraction ...... 29 PCR Amplification ...... 32 PCR analysis using gel electrophoresis...... 40 Troubleshooting ...... 44 Troubleshooting Guide 1 - DNA Extraction...... 44 Troubleshooting Guide 2 - Gel Electrophoresis ...... 45 Troubleshooting Guide 3 – PCR Diagnostic Results ...... 47 Limitations of the Technology ...... 49 Sources of Reference Material ...... 50 Fungal cultures and DNA ...... 50 Contacts ...... 51 Further Reading ...... 52 Appendix ...... 53 Reagents for DNA extraction...... 53 Reagents for Agarose Gel Electrophoresis...... 54 References ...... 56
2 Introduction
The Banana Industry in Australia
In 2005, the wholesale value of the Australian banana industry exceeded $320 million. More than 80% of the 20.4 million, 13kg carton crop is produced in the wet tropics of North Queensland, in the region between Tully and Cooktown. The remaining 20% are grown predominantly in the subtropical regions of the eastern coast from Nambucca Heads, in New South Wales to Nambour in Queensland. There are also small producing regions in the Northern Territory and in Kununurra and Carnarvon in Western Australia.
Approximately 95% of bananas grown in Australia are of the Cavendish subgroup (including the cultivars Grand Nain and Williams) with the remaining 5% comprising mainly the Ladyfinger cultivar but also some Goldfinger, Ducasse, Sucrier, FHIA 18, Red Dacca and Plantains. The Australian market for banana is mostly that of the fresh fruit product.
Currently the only other products made from banana fruit are the small-scale production of banana puree, dried banana and banana wine. There is also a specialist manufacturer of handbags from banana fibre.
Origin and Distribution of Black Sigatoka
Black Sigatoka, which is also known as black leaf streak in some parts of the world, is caused by the fungal pathogen Mycosphaerella fijiensis (Leach 1964). Black Sigatoka affects many Musa species throughout the world including those grown on the island of New Guinea, and in the Torres Strait Islands (TSI).
M. fijiensis was first identified in the Sigatoka valley of Vita Levu in the Fiji Islands in 1963 (Leach 1964). Black leaf streak, as it was first known, quickly replaced Sigatoka, which was endemic in the region at that time. Surveys of the Oceanic region from 1964 to 1967 revealed that this new disease was already established in the Pacific and parts of the Pacific Rim. From its wide dispersal, it appeared that M. fijiensis had been present in this region for some time prior to its identification. The disease was found in the Philippines in 1970 but again it appeared that it might have been present for 15 years or more prior to this. From the examination of herbarium specimens it was concluded that M. fijiensis had been present on the island of New Guinea since 1957 and that it had been present in Taiwan in 1927 (Stover 1978). From this work Stover concluded that the centre of origin for M. fijiensis was likely to have been in PNG-Solomon Islands region.
The spread of M. fijiensis in the Latin Americas has been well documented as it appeared in this region after its initial identification in Fiji in 1963. Although the pathogen may have been
3 introduced to Honduras as early as 1969, it was only positively identified there in 1972 (Stover and Dickson 1976). It was here that the disease acquired its current name black Sigatoka. The disease slowly moved throughout Latin America and by 1981 was endemic to Central America. It has since moved south into Colombia, Ecuador, Peru and Bolivia. M. fijiensis has only recently reached the Caribbean. It was first identified in Cuba in 1992 (Vidal 1992) and more recently in Jamaica, the Dominican Republic, Trinidad, Grand Bahama Island and Haiti (Carlier et al. 2000a) (Fortune et al. 2005; Jones 2002) (Mourichon et al. 1997). The direction of the prevailing winds is believed to limit further spread by natural means of M. fijiensis within the Carribean (Carlier et al. 2000a).
Although M. fijiensis may have been present in Africa as early as 1973, it was first identified in Gabon in 1978 (Frossard 1980). It is believed that the pathogen was introduced into Africa from infected banana plants imported from Asia. From its first introduction, it has since spread to neighbouring countries and is now found in Cameroon, Cote d’Ivoire, the Democratic Republic of Congo, Nigeria, and Ghana. A separate introduction is believed to have brought the pathogen to the countries on the east coast of Africa (Carlier et al. 2000a). The disease was first identified in this region in 1987 on the island of Pemba. The disease quickly spread to the adjacent island of Zanzibar and from there to the mainland of Tanzania (Dabek and Waller 1990). By 1988 it had reached Kenya (Kung'U et al. 1992). The introduction of the disease to the East African countries of Rwanda, Burundi, (Sebasigari 1989), Uganda (Tushemereirwe and Waller 1993) and Malawi (Ploetz et al. 1992) is however thought to have been from the Democratic Republic of Congo (Carlier et al. 2000a). A map showing the worldwide distribution of M. fijiensis is presented in Figure 1.
The distribution of M. fijiensis in Asia is still unclear (see section on Other Leaf Spots) although there are several substantiated reports of this pathogen from this region. The pathogen has been confirmed as being present in Southern China (Carlier et al. 2000a; Mourichon and Fullerton 1990), Vietnam, Thailand, Taiwan and Singapore. Black Sigatoka has also been confirmed as being present in parts of Malaysia; West Malaysia, Jahore, Langkawi and East Malaysia and also as being present in parts of Indonesia, namely Halmahera, Java, Kalimantan and West Sumatra.
4 Distribution of M. musicola. xxx Distribution of M. fijiensis. ● Distribution of M. eumusae.
Figure 1: World map showing the global distribution of Mycosphaerella fijiensis, Mycosphaerella musicola and Mycosphaerella eumusae.
5 Black Sigatoka in Australia
Currently, mainland Australia is a designated ‘black Sigatoka free’ region. However, there have been nine previous incursions of the disease in North Queensland, the first detected in 1981 and the most recent incursion in 2001 in the Tully Valley. A Pest Free Area was declared on the 20th December 2004.
Although M. fijiensis was only detected on bananas on the Torres Strait Islands during a plant disease survey in 1981 its spread would indicate that it had been present in this region for a considerable period of time prior to its detection. However, as no disease surveys had been undertaken in the region prior to 1981, it cannot be certain how long the disease had been present for. The pathogen had most likely been introduced on banana plants brought over from Papua New Guinea where it is known to be endemic.
Black Sigatoka was also detected in Bamaga, on mainland Australia, during the 1981 disease survey. Although the disease was widespread through the township, only 25% of plants were infected due to the resistance of many of the banana cultivars grown. Many plants found in Bamaga belonged to the ABB genotype. The ABB genotype contains subgroups with many cultivars having higher resistance to M. fijiensis compared with other genotypes, such as the AAA subgroup of which Cavendish is a member. Another factor that impeded the spread of the disease was this region had a low density of banana plants and is situated in the dry tropics of Australia. Without additional water and nutrients, banana plants do not grow well in this region. As an initial measure to prevent the spread of black Sigatoka, the disease was declared quarantinable under the Commonwealth Quarantine Act of 1908. The region was then proclaimed a quarantine region enabling enforceable restrictions on the movement of banana material out of the area.
Due to the proximity of this region to Queensland’s major commercial banana production areas, the Queensland Department of Primary Industries and Fisheries (DPI&F) first attempted eradication in October of 1981. The Commonwealth Government and Australia’s banana growing States sponsored this eradication program. This original attempt at eradication involved the destruction of all banana plants in the Bamaga region as well as all banana plants on Thursday Island, Badu Island and Kubin Village on the western side of Moa Island. Replanting was left for a period of 6 months in order to avoid reinfection from any ascospores discharged from the banana trash. In 1984, M. fijiensis was again detected on some banana plants during routine plant disease surveillance by the DPI&F. This time the disease was restricted to the Bamaga area. It was not found on Thursday, Badu or Moa Islands (Jones 1984) indicating that the eradication program on these islands had been successful.
6 Failure of the initial eradication program in Bamaga was thought to be due to several factors. Regrowth from eradicated plants may have provided a source of inoculum or some banana plants may have been missed during eradication despite thorough surveys of the area. Another possibility is that local residents may have hidden plants prior to the eradication program and then returned them to Bamaga after the reintroduction of clean planting material, and so providing a source of inoculum (Jones 1989). At that time it was decided not to proceed any further with eradication attempts and the disease was instead managed and contained in Bamaga. This was possible due to quarantine restrictions preventing the movement of banana material out of the Torres Strait Islands and Bamaga to other parts of Australia and also due to the remoteness and isolation of the region at the time. Trials were undertaken in the region to evaluate the resistance of several banana cultivars to black Sigatoka (Jones 1984). A program was also introduced during this time by the Queensland Banana Industry Protection Board (BIPB) to replace all the bananas in the Bamaga area with more resistant cultivars such as Bluggoe and Tu8 (Jones 1989).
In 1988, a second attempt at eradication was undertaken due to increased pressure from the banana industry. A small commercial plantation of Cavendish bananas had been planted at Bamaga and the access road had been upgraded leading to an increase in traffic to the region. The industry feared that black Sigatoka would move south into the large commercial growing regions devastating the industry. Eradication of the pathogen was effected through the destruction of all banana plants in the area, replanting with resistant lines such as Tu8, Bluggoe and Ducasse, and the favourable climate of a long, hot, dry season. Another incursion in this region detected in 1999 has been linked to regrowth of plants from the 1988 eradication program and also to the importation of plants directly from Thursday Island (Davis et al. 2000). Again, all susceptible plants were removed and buried. Black Sigatoka has not been detected in the region since the 1999 eradication.
There have been six further small incursions since 1988 (Figure 2) all in isolated, banana growing regions. On each occasion, officers from DPI&F have managed the eradication of the disease by the removal and destruction of all banana plants in a buffer zone of up to a 50 km around the site of the infection. The area was then allowed to remain free of banana plants for at least 4 months before replanting with cultivars more resistant to black Sigatoka. Eradication was also assisted by the natural climatic conditions. Many regions in Cape York experience long, hot and dry seasons interspersed with a short, wet season. Banana plants do not survive the dry season in this area without irrigation and M. fijiensis does not proliferate under these conditions (Peterson 2002).
All of these incursions have been attributed to the movement of diseased plants between sites of infection. The close proximity of the Torres Strait Islands to the mainland and the strong cultural ties between various communities residing in this region suggest a logical
7 ingress for the pathogen. The initial movement of diseased banana plant material and suckers from the Torres Strait Islands could have brought the pathogen to mainland Australia and then the subsequent movement of planting material transported it between sites of infection on the mainland.
The most recent and most serious of incursions was in Tully in 2001. This incursion differed from the previous ones in two ways: (i) this incursion was in the largest commercial banana production area in Australia and (ii) Tully has one of the highest rainfalls in Australia. The prior eradication strategy of removing all banana plants in the region was revised as this now involved 4400 ha of banana plants. An alternative strategy was developed which relied heavily on deleafing of all banana plants to achieve a zero disease level in the region. Constant surveillance also enabled scientists to monitor the disease. Diagnosis of diseased leaf material was however more difficult than usual. Heavy rains in the area at the time had washed away all of the fungal structures used to differentiate M. fijiensis from the endemic M. musicola. Molecular diagnosis was a useful tool used to differentiate between the two pathogens and was one of the many factors contributing to the success of the eradication campaign.
Figure 2: Map of Queensland showing the locations of all Mycosphaerella fijiensis incursions on mainland Australia since 1981. All incursions were eradicated by the Queensland Department of Primary Industries and Fisheries (DPI&F).
8 Yellow Sigatoka
Yellow Sigatoka is caused by the fungal pathogen Mycosphaerella musicola Leach which is closely related to M. fijiensis. The pathogen is considered to have a worldwide distribution. It has not however been reported in the banana growing regions of the Canary Islands, Egypt and Israel (Jones 2000) and its exact distribution through Asia is still unclear.
Stover (1962) hypothesised the mode of spread of M. musicola worldwide (Stover 1962). Working from disease records, Stover proposed that M. musicola was moved from Java where Zimmermann first described the anamorph of this pathogen, Pseudocercospora musae in 1902, to Fiji on banana leaf material used as packing material in shipping containers. M. musicola was first identified in Fiji in 1913 by Massee (Massee 1914). From here Stover proposed that the pathogen moved to the east coast of Australia on the prevailing winds in around 1924. At this time there was a disease epidemic in the Fijian banana plantations of the Sigatoka Valley which was causing inoculum levels to be exceptionally high. Once in Australia the disease quickly spread throughout banana plantations, many of which had been left unmanaged due to the severe banana bunchy top disease (BBTD) epidemic. Stover hypothesised that the combination of exceptionally high levels of inoculum during the epidemic years of the late 1920s and early 1930s, coupled with unusual climatic conditions and air turbulence, could have resulted in enough viable ascospores surviving the long journey on the tradewinds to cause the disease outbreaks in Africa and South America.
This hypothesis was tested using RFLP markers to study the genetic structure of the global population of M. musicola (Hayden et al. 2003). Hayden (2003) found that Stover’s hypothesis was supported by the fact that many alleles found in the Indonesian population were detected in the Australian, African and South American populations. The hypothesis however was not supported when the Australian population was compared with the African and the South American populations. Both of these populations possessed alleles not present in the Australian population. Further, Hayden concluded from the genetic differentiation data that it is likely that the African and South American M. musicola populations arose from the Indonesian population in a separate founder event from that of the Australian population.
In Australia, M. musicola was found to have spread to the banana growing regions of New South Wales by 1927 (Simmonds 1928). M. musicola is now endemic throughout all banana- growing regions in Queensland and northern New South Wales. In Western Australia it was first detected in Kununurra in 1990 although it is thought to have been present for some time before this first report (Shivas and Kesavan 1992). It is now identified as a common pathogen to the banana growing regions of the Kimberleys. The pathogen has also been detected in banana growing regions in the Northern Territory.
9 Disease Symptoms
Black Sigatoka
Black Sigatoka causes large necrotic lesions on the leaves of the banana plant and early drop (collapse) of the entire leaf (Figure 3). The resulting loss of photosynthetic capacity leads to slower filling of fingers, reduced yields and finger size and premature ripening of fingers. Field losses vary from 30-50% depending on the climatic conditions (Gauhl et al. 2000; Stover 1983) and are presently 5-10% in even well-managed plantations with good control strategies (R. Romero, pers. comm.). In subsistence crops of plantain, yield loss has been estimated to be up to 33% during the first crop cycle and up to 76% in the second (Mobambo et al. 1996).
CIRAD
(b)
(a) Photo by Juliane Henderson (c) CIRAD
Figure 3(a): Black sigatoka infection in a managed banana plantation in Costa Rica (Photo by Juliane Henderson), (b) heavily diseased leaf (Image courtesy of CIRAD) and (c) underside of a diseased leaf (Image courtesy of CIRAD).
10 There are six recognised stages in symptom development (Fouré 1987; Meredith and Lawrence 1969). A brief description of each stage follows:
Stage 1: Initially, tiny specks < 0.25 mm and white to yellowish in colour that quickly turn a reddish brown, appear on the abaxial surface (underside) of the leaf laminar. This first stage is also known as the ‘initial speck stage’ ).
Stage 2: The tiny reddish brown specks elongate and widen, becoming streaks approximately 2mm X <1 mm. This stage is also referred to as the ‘initial streak stage’. The streaks are more clearly visible on the abaxial surface of the leaf laminar than the adaxial surface (upper side) of the leaf. Conidia and conidiophores may be present (Figure 4(a)).
(a) Stage 3: The streaks continue to expand in size and change colour to a very dark brown, almost black, colour. This is also referred to as the ‘second streak stage’. Where infection is heavy, the streaks overlap to give a black appearance to large (a) areas of the leaf. The streaks are clearly visible from the adaxial side of the leaf. Conidia and conidiophores are present at this stage (Figure 4(b)).
Stage 4: The streaks continue to enlarge and become more elliptical in shape as it broadens and a water-soaked border may develop around the edges. This stage is known as the ‘first spot stage’ (Figure 4(c)).
(b) Stage 5: This stage also known as the ‘second spot stage’ is characterised by the central region of the spot becoming slightly depressed. The water soaked border may develop a yellow halo around it. Where infection is heavy, large areas of leaf tissue collapses. (Figure 4(d)).
Stage 6: The final stage, also referred to as the ‘third spot stage’, is when the centre of each spot becomes dry and pale grey to beige in colour. Perithecia and ascospores are present in stage 6 lesions. Surrounding each of the spots is a distinctive black border. Where infection is heavy the large areas of the leaf become necrotic. The spots remain visible even after the death and desiccation of the leaf due to the dark border encircling each of the individual spots (Figure 4(e)).
11 Figure 4(a): Stage 2 symptoms or ‘First Streak Stage’. Figure 4(b): Stage 3 symptoms or ‘Second Streak Stage’. Conidia and conidiophores may be present at this stage. Streaks are now almost black. Conidia and conidiophores Note that Stage 1 symtpoms (initial speck stage) are barely are present. visible at <0.25mm.
Figure 4(c): Stage 4 symptoms or ‘First Spot Stage’. The Figure 4(d): Stage 5 symptoms or ‘Second Spot Stage’. streaks are becoming more elliptical and have a water- Note the blackening in the centre of the spots. The water- soaked border. soaked border begins to develop a yellow halo.
Figure 4(e): Stage 5 & 6 ‘Third Mature Spot’ symptoms. Multiple lesion stages are present. Note the pale grey centres of the Stage 6 lesions.
Figure 4: Images of five of the six diagnostic stages of development for black Sigatoka disease (Images reproduced with permission, Department of Primary Industries and Fisheries, Queensland).
12 Yellow Sigatoka Yellow Sigatoka disease is similar to black Sigatoka (Figure 5). There are, however, some distinguishing diagnostic features for yellow Sigatoka. The disease ultimately has the same effect on yields as black Sigatoka, although yellow Sigatoka disease development is slower, enabling it to be controlled through deleafing and the use of fungicides.
(b) CIRAD
(a) (c)
Figure 5: (a) Banana plant infected with Mycosphaerella musicola. Note later stage lesions are always present in the lower leaves which are older while the newer leaves show the earlier stage symptoms. Symptom development can be used in conjunction with other tools to assist with diagnosis. The photographs in (b) and (c) show advanced lesions on leaves.
(Images (a) & (c) reproduced with permission, Department of Primary Industries & Fisheries, Queensland (b) courtesy of CIRAD & INIBAP)
13 Yellow Sigatoka can be differentiated from black Sigatoka at the early stages of lesion development (Stages 1 and 2) on visual symptoms. At later stages, examination of the conidiophores and conidia requires compound microscopy. There have been several descriptions of the development of individual lesions of Sigatoka disease over the years which are well summarised in Meredith (1970) (Meredith 1970). Brun’s description (Brun 1958) is similar to that of Leach (Leach 1946) except that Brun excludes Leach’s 5th stage (second spot stage). A brief description of each stage as per Brun follows:
Stage 1: This stage is characterised by the appearance of very small light green dots or dashes of approximately 1 mm in length. (Figure 6(a))
Stage 2: The small dot or dash of Stage 1 elongates into a light green streak several millimetres long. (Figure 6(b)&(c))
Stage 3: At this stage there is a change in the colour of the streak to a rusty brown. The streak becomes elongated and widens slightly. The border of the streak is ill defined. (Figure 6(d))
Stage 4: The streak becomes more elliptical and is a definite spot with a sunken dark brown centre. It is often surrounded by a yellow halo. At this stage the conidia and conidiophores are produced. (Figure 6(e))
Stage 5: The final stage has a grey dried out centre and an obvious black margin. This black margin can still be seen even after the leaf has dried out. (Figure 6(f))
The stages of both yellow and black Sigatoka are summarised in Table 1.
14 Figure 6(a): Stage 1 lesions of yellow Sigatoka Figure 6(b): Lesions associated with Stage 2a (early) of characterised by the light green dots and dashes which are yellow Sigatoka. Note light green streaks which are the about 1 mm in length. characteristic lesions at this stage.
Figure 6(c): Lesions associated with Stage 2b (late) of Figure 6(d): Stage 3 symptoms associated with yellow yellow Sigatoka. Note the change in colour of the streaks Sigatoka. Note that the streaks from stage 2 have now from light green to rusty brown. elongated and widened.
Figure 6(e): Stage 4 symptoms associated with yellow Figure 6(f): Stage 5 symptoms of yellow Sigatoka. Note Sigatoka. Note that the Stage 3 streaks have now become the dried out grey centre with the black ring outside. spots. Conidia and conidiophores may be present from this stage
Figure 6(a)-(f): Images of the five stages of lesion development in yellow Sigatoka. (Images reproduced with permission, Department of Primary Industries and Fisheries, Queensland).
15
Lesion Stage Yellow Sigatoka Black Sigatoka
Stage 1 Very small light green dot or dash Small pigmented spot of white or up to 1 mm long yellow, similar to yellow Sigatoka stage 1
Stage 2 Light green streak several Brown streak, visible on underside of millimetres long leaf, later visible on leaf upper surface as yellow streak; colour changes progressively to brown, then black on upper leaf surface
Stage 3 An elongated rusty brown spot Enlarged stage 2, streaks become with an poorly defined border longer
Stage 4 A mature spot with a dark brown Appears on leaf underside as brown sunken centre; often surrounded spot, as a black spot on upper leaf by a yellow halo, conidiophores surface and conidia are produced at this stage
Stage 5 Spot has developed a grey, dried Elliptical spot is totally black on the out centre and a peripheral black underside of the leaf, surrounded by ring which is evident even after a yellow halo the leaf has dried out
Stage 6 Centre of spot dries out, turns grey and is surrounded by a well-defined margin and a bright yellow halo
Table 1: Summary of the different lesion stages associated with yellow and black Sigatoka leaf spot diseases of banana.
16 Other Leaf spots A third leaf spot disease, Eumusae leaf spot (Figure 7), has recently been described. This disease is caused by the pathogen, Mycosphaerella eumusae, anamorph Pseudocercospora eumusae which is very closely related to M. fijiensis and M.musicola (Carlier et al. 2000c; Crous and Mourichon 2002). Distribution of this pathogen is still uncertain however originally it was found serendipitously in Asia in West Malaysia, Thailand, Vietnam, Southern India and Sri Lanka during a survey initiated by INIBAP to determine the distribution of M. fijiensis and M. musicola within South and South-east Asia. Samples collected as Sigatoka leaf spots were found to be associated with this new pathogen (Carlier et al. 2000b; Carlier et al. 2000c; Crous and Mourichon 2002). Little is known about M. eumusae however, one isolate was found infecting a banana known to be resistant to M. fijiensis. Morphologically it is similar to M. musicola.
Several other leaf spot diseases may produce lesions with a similar appearance to those of M. fijiensis and M. musicola however these other pathogens can be easily distinguished using light microscopy as they are morphologically quite different.
Figure 7: Banana leaves with lesions associated with Eumusae leaf spot (ELS) caused by the fungal pathogen Mycosphaerella eumusae. (Images reproduced with permission, INIBAP & CABI, UK)
17 The Pathogens
The Causal Agents
Black Sigatoka is caused by the heterothallic ascomycetous fungi Mycosphaerella fijiensis Morelet (anamorph Paracercospora fijiensis). Yellow Sigatoka is caused by another, Mycosphaerella musicola Leach ex Mulder (anamorph Pseudocercospora musae). Jones (2000) has a comprehensive chapter describing fungal leaf diseases of banana plants.
Taxonomy of M.fijiensis and M. musicola
Morphology
Although M. fijiensis and M. musicola are extremely closely related, and some stages of the disease appear similar, the morphology of the conidia and conidiophores can be routinely used to differentiate the two pathogens. Conidia produced by M. fijiensis are pale to medium olive green with paler tips, 1-10 septate, obclavate to cylindro-obclavate and straight or curved. Conidia produced by M. musicola however are a paler shade of olive green, 0-6 septate, cylindrical to obclavate-cylindrical, and also either straight or curved. The conidia of M. fijiensis are also discernable by the thickened basal hilum at the base.
Conidiophores of Paracercospora fijiensis can be either straight or bent, are pale to medium brown, 0-5 septate, often geniculate and are generally unbranched. A diagnostic characteristic are the distinctive scars present on the tip of the conidiophores. Conidiophores of P. musae differ in that they are straight, hyaline, mostly without septa, geniculation or branching scars are not present.
Another difference between the two pathogens is the location of the conidiophores. When P. fijiensis is the infecting pathogen they are mainly found on the abaxial surface of the leaf whereas conidiophores of P. musae are abundant on both surfaces. Conidiophores are also present much earlier in the lesions associated with black Sigatoka, as early as stage 2 lesions, whereas they are generally not present until the disease has progressed to stage 4 lesions for yellow Sigatoka. Images of these structures are presented in Figures 8 and 9. The structures of Mycosphaerella eumusae are also presented in Figure 10 for comparison. The morphology of the anamorphs of M. fijiensis and M. musicola is summarised in Table 2.
18 (X 10) CIRAD (X 40) CIRAD
Figure 8 (a) Fruiting bodies of Paracercospora Figure 8(b) Conidia produced by the fruiting bodies fijiensis found in lesions associated with black of Paracercospora fijiensis Note the basal Sigatoka. Note the small number of conidia thickening which is not found in Pseudocercospora produced by these in comparison with M. musae or Pseudocercospora eumusae. musicola and M. eumusae.
(Images courtesy of CIRAD & INIBAP)
(X CIRAD (X CIRAD
Figure 9 (a): Fruiting bodies of Figure 9 (b): Conidia of Pseudocercospora Pseudocercospora musae found in musae. No basal thickening is present. Up lesions associated with yellow to 50 conidia may be associated with a Sigatoka disease. single sporodochia.
(Images courtesy of CIRAD & INIBAP)
19 (X CIRAD (X CIRAD
Figure 10 (a): Fruiting bodies of Figure 10 (b): Conidia of Pseudocercospora eumusae. Pseudocercospora eumusae found in Conidia are shorter (21.2-41.6 x 2.5 µm) than those of lesions associated with Eumusae leaf P. musae (10-109 x 2-6 µm). spot. Conidiophores are found mainly on the abaxial leaf surface.
(Images courtesy of CIRAD & INIBAP)
Paracercospora fijiensis Pseudocercospora musae
Conidiophores • first appear at stage 2 or • first appear at stage 4 or initial streak stage first spot stage
• can be either straight or • are straight, hyaline, bent, are pale to medium mostly without septa, brown, 0-5 septate, often geniculation or branching geniculate, generally and they do not have any unbranched with spore scaring distinctive, slightly thickened spore scars that • abundant on both surfaces are diagnostic for this pathogen
• mainly found on the abaxial surface of the leaf
• emerge singly or in small groups
Conidia • pale to medium olive • a paler shade of olive green with paler tips, 1-10 green, 0-6 septate, septate, obclavate to cylindrical to obclavate- cylindro-obclavate and cylindrical, and also either straight or curved straight or curved
• thickened basal hilum • no basal thickening present
Table 2: Comparison of the morphology of Paracercospora fijiensis and Pseudocercospora musae.
20 Hosts
Currently the only known hosts of M. fijiensis and M. musicola are Musa spp. These species and subspecies all vary in their levels of resistance to M. fijiensis and M. musicola. There is one report in the literature of M. musicola having been isolated from leaf spots on a Heliconia species in Venezuela (Madiz et al. 1991).
Relative distributions of M. fijiensis and M. musicola
M. fijiensis is found throughout the world’s tropical banana growing regions from the Tropic of Cancer to the Tropic of Capricorn with the exception of Australia. M. musicola also causes serious yield losses in regions not affected by black Sigatoka. As this pathogen can proliferate at lower temperatures and lower relative humidity, M. musicola is more widespread than M. fijiensis. Yellow Sigatoka is often the dominant disease at higher altitudes (>1200 m) although it appears that M. fijiensis is becoming more adapted to higher altitudes and is gradually replacing M. musicola in these regions (Carlier et al. 2000a). Figure 1 shows the worldwide distribution of both of these pathogens.
Life Cycle
Most infections of M. fijiensis and M. musicola begin with spores being deposited on the susceptible cigar leaf of the banana plant. Spores will germinate within 2-3 hours of being deposited on the leaf surface if there is a water film present or if the humidity is very high. The optimal temperature for germination of M. fijiensis spores is 27ºC. For M. musicola the optimal temperature for germination of conidia is between 25-29ºC and for ascospores it is between 25-26ºC. The germ tube then grows epiphytically for several days (2-3 days for M. fijiensis and 4-6 days for M. musicola) before penetrating the leaf via stomata in a hydrotropic response through the formation of appressoria or stomatopodia over the stomata (Meredith 1970; Stover 1980).
Once inside the leaf, the infection hypha forms a large substomatal vesicle. Fine hyphae then grow through the mesophyll layers into an air chamber and then into the palisade tissue. From here the hyphae grow out into other air chambers eventually emerging through stomata in the streak that has developed. Again, epiphytic growth occurs before the re-entry of the hypha into the leaf through another stomate.
Conidia are observable from stage 2 of black Sigatoka whereas they are generally only visible from stage 4 of yellow Sigatoka. Perithecia form during stages 5 and 6 of black Sigatoka and during stage 5 of yellow Sigatoka. Overall the disease cycle is much faster for M. fijiensis than for M. musicola due to shorter time required to complete the life cycle. Generally, it has been observed, the optimal conditions for M. fijiensis are those where there is, on average, higher temperatures and higher relative humidity. See Table 2 for the disease development and associated structures found during the disease cycle for each of these pathogens.
21 Comprehensive cytological studies of the interactions between M. fijiensis and three banana genotypes have been undertaken (Beveraggi et al. 1995) (Sallé et al. 1989). From these studies it was found that there is a relatively long period of biotrophy before any incompatible reactions are observed in susceptible cultivars. The pathogen colonises the leaf tissue, growing intercellularly without the production of haustoria, for almost a month. During this period, little evidence of the presence of the pathogen can be seen externally. Cytological changes are visible in the parenchyma cells after about 28 days although the cells still appear healthy. There is contact between the hyphae and the cells but no localised reaction. Externally, stage 2 or the initial streak stage symptoms are visible.
After 41 days, stage 5 or second spot stage symptoms are visible externally. In the tissue sections taken from the susceptible cultivar ‘Grande Naine’ at this time during the studies, three distinctive zones were seen. Zone I which corresponded to the cells within the necrotic spot contained plasmolysed cells. Zone II corresponded to the yellow halo and this region contained cells with large intracellular globules. At the boundary of Zones II & III an intercellular substance, later identified as polyphenol, was noted. This substance formed intercellular bridges and host cells in contact with it showed degeneration of the cell wall. Beyond this boundary the remaining host cells in Zone III appeared to be healthy. Hyphae were observed throughout the intercellular spaces in all of the zones (Sallé et al. 1989).
Importantly, haustoria were never observed during the invasion of any host by M. fijiensis. The progression of the disease was similar in the partially resistant cultivar, ‘Fougamou’, except that the growth rate of hyphae in the susceptible cultivar was much higher than in the partially resistant cultivar (Beveraggi et al. 1995; Sallé et al. 1989). In the highly resistant cultivar, ‘Yangambi Km5’, a compatible reaction was not observed. There was no biotrophic period; rather fungal growth was blocked at the site of penetration. Stomatal guard cells became necrotic and there was a deposition of polyphenolic substances around the outside of the cell walls of the host and the pathogen. This is consistent with a hypersensitive response (Beveraggi et al. 1995; Sallé et al. 1989).
22 Disease Development and Epidemiology
Disease Cycle
The disease cycle for both M. fijiensis and M. musicola is similar with only minor differences as outlined previously. As M. fijiensis produces considerably less conidia and for a shorter period of time than M. musicola, ascospores are the main dispersal agent for this pathogen (Stover 1980). Both conidia and ascospores are important for dispersal of M. musicola (Stover 1971) however for both pathogens ascospores are involved in the movement of the pathogen over longer distances rather than conidia. A distinctive line spotting pattern of infection is produced when the source of inoculum is conidia dislodged by rain splashes. These run down the inside of the cigar leaf cylinder contacting the lower point of the cylinder resulting in a line of infection. The deposition of ascospores by wind currents is generally on the terminal end of these leaves resulting in a distinctive leaf tip infection (Meredith 1970; Stover 1972).
The disease cycle is much faster for black Sigatoka than it is for yellow Sigatoka, as seen by the earlier appearance of spots. Inoculation studies conducted in Honduras demonstrated that spotting associated with M. fijiensis infections appeared 8-10 days faster than that associated with M. musicola infections. Ascospore maturation time is also shorter at 2 weeks for M. fijiensis compared with 4 weeks for M. musicola (Stover 1980). A diagrammatic representation of the disease cycle for M. musicola is presented in Figure 11.
Survival of the Inoculum
Production of perithecia and the subsequent discharge of ascospores continues for several months. Even in severely necrotic tissue, ascospore ejection can continue for more than two months, this is the case also where the leaf has been removed and placed on the ground (Carlier et al. 2000a). Ascospore release remains high for three weeks after removal of the leaf from the plant and then decreased rapidly over the next six weeks until the tenth week when the leaves themselves had disintegrated (Gauhl 1994). The survival of ascospores is directly related to the time it takes for the disintegration of the diseased leaf material (Stover 1980). Ascospores ejected are no longer viable after 6 hours of exposure to UV radiation (Parnell et al. 1998).
23
Figure 11: Life cycle of Mycosphaerella musicola, the fungal pathogen causing yellow Sigatoka (Reproduced with permission, Department of Primary Industries & Fisheries, Queensland).
24 Spread of the Pathogen
Both M. musicola and M. fijiensis are dispersed within banana blocks by rain splash of conidia. Movement between blocks is possible through the aerial spread of ascospores ejected from the perithecia. Due to the larger amount of conidia produced by M. musicola than by M. fijiensis, conidia are considered the main means of spread for M. musicola while ascospores are the main method of dispersal of M. fijiensis (Stover and Dickson 1976).
Long distance spread may also be via the wind dispersal of ascospores. The short time that ejected ascospores can survive UV irradiation suggests that the distance viable ascospores are dispersed by this method will also be affected by the amount of cloud cover and the distance travelled through the night (Parnell et al. 1998). Recent population studies of both M. fijiensis (Rivas et al. 2004) and M. musicola (Hayden et al. 2005) however, suggest limited long distance dispersal—less than 50 m—of these pathogens based on the genetic structure of the populations. In many cases long distance movement, especially intercontinental movement, of the pathogen is thought to be more likely due to the direct transportation of germplasm from an infected area to a new region (Rivas et al. 2004).
25 Disease Control
Control of black and yellow Sigatoka can be carried out in a number of ways including removal of infected leaf area, chemical control or the use of more resistant varieties.
Cultural Control
To reduce inoculum levels in plantations it is extremely important to remove any leaves with lesions from the plant. Lesions associated with M. fijiensis infections contain perithecia that continue to produce and eject ascospores for several months. This time can be decreased significantly if infected leaves are removed from the plant and placed on the ground enabling them to degenerate faster.
Another factor that can be manipulated, to some degree, is that of relative humidity. The use of an efficient drainage system within plantations can assist with reducing the relative humidity by removing excess groundwater and rainwater as quickly as possible. Where irrigation of plantations is required, the use of drippers or under plant systems prevents humidity in the canopy from rising to levels favourable for germination of ascospores. For this reason, the planting density can also affect the spread of disease.
Chemical Control
In regions where M. fijiensis is endemic, chemical control of this pathogen also effectively controls M. musicola. Unfortunately, the converse is not true.
The fungicides used in the control of black and yellow Sigatoka fall into three categories: (i) protectant fungicides, (ii) systemic fungicides, and (iii) oil.
Protectant fungicides do not penetrate the leaf surface and are broad-spectrum fungicides. The fungicidal action occurs on the leaf surface and so is used at early stages of infection or as a preventative. It is the result of a reaction of the fungicide with essential enzymes containing thiol groups which produces a non-selective toxicity. The protectant fungicides used to control yellow Sigatoka in Australia are dithiocarbamates and in particular are formulations of mancozeb a compound made of zinc and magnesium salts and ethylene bisdithiocarbamate. In Australia these fungicides are generally applied with the addition of oil.
Systemic fungicides do penetrate the leaf surface and so they are used after infections have taken place—up to stage 3 lesions. Systemic fungicides used in Australia are from the triazole group. These compounds inhibit a cytochrome P-450 mono-oxygenase enzyme. This enzyme catalyses the C-14 demethylation reaction which is part of the ergosterol biosynthesis
26 pathway. This group of fungicides are generally referred to as demethylation inhibitors (DMI). The fungicides Bumper® 250 EC, and Tilt® 250 EC (both propiconazoles); Folicur® 430 EC (a tebuconazole); Opus® 75 (epoxiconazole) and Score® (difenoconazole) are registered for use on bananas in Australia.
The third group, oils, are also widely used fungicides and are generally applied with both protectant and systemic fungicides. It is still unclear how petroleum oils work as fungicides. In vitro experiments on M. musicola showed that the oil had no effect on the growth of the fungus. It is thought that the oil may act by stimulating the plants own defence responses. Care has to be taken though when using oil in spraying programs as it can build up on the leaf surface resulting in reduced gaseous exchange and photosynthesis leading to a potential decrease in yield.
In some regions of the world a disease forecasting system is used to assist with managing the effective use of fungicides. The system is based on the monitoring of plantation disease levels together with rainfall, temperature and relative humidity. The rationale for the use of this system is that it enables plantation managers to time fungicide applications by predicting the optimal time for spraying in an effort to reduce the amount of chemicals required to control the disease. This is important in regions where the crop is grown as a staple as it reduces the cost of production. However it has also been adopted in some banana growing regions in an effort to prevent fungicide resistance developing in local populations of M. fijiensis. This method has been used successfully in French Antilles for the management of Sigatoka disease and in Cameroon for black Sigatoka. It has not been as successful though in Central America possibly due to the more favourable climatic conditions which enable M.fijiensis populations to proliferate at a much faster rate and thus increasing the number of sexual cycles attained over a given period of time. This leads to an increase in the amount of recombination in the population providing a higher potential for resistance to develop.
27 Microscopic Diagnosis of Disease
It is difficult for the less experienced plant pathologist to confidently identify black or yellow Sigatoka based on leaf spot symptoms alone. In fact there are several diseases that can, at different stages and under certain conditions, appear to be identical. The development and appearance of symptoms can differ for each of the Sigatoka diseases as a consequence of various biotic and abiotic factors such as the prevailing weather conditions, nutritional state of the plant and inoculum levels present. They will also vary due to the different levels of resistance among different cultivars of Musa spp. These differences are used to classify banana cultivars into resistance groupings.
A preliminary diagnosis can sometimes be made based on the leaf spot symptoms. When used in conjunction with light microscopy a definitive diagnosis can generally be made. Microscopy is therefore an indispensable tool in the diagnosis of these pathogens. Although morphologically these two pathogens are very similar, there are some small but significant differences between them. Generally, these differences can be observed on microscope slides prepared directly from the diseased leaf tissue.
Leaf tissue with suspected early stage Sigatoka leaf spot lesions should be incubated overnight at 100% relative humidity at around 25ºC. Generally the samples will already be contained inside a sealed plastic bag from collection which achieves this high humidity without additional incubation. The high humidity will ensure a profusion of conidia and conidiophores for identification. As noted previously, conidia are present much earlier in M. fijiensis infections and can be observed as early as stage 2 lesions whereas conidia can only be seen from stage 4 lesions in M. musicola infections.
The Pathogen
Images of the identifying structures (conidiophores and sporodochia) are presented in Figures 8 and 9. Note the scaring and basal thickening in the conidiophores of M. fjiensis. Images of the closely related M. eumusae are presented as well for comparison (Figure 10).
28 Molecular Diagnosis of Disease
Preliminary diagnosis and sampling
Sections of banana leaves showing disease are inspected for symptoms and a preliminary diagnosis made where possible based on lesion appearance. When conidia are present, morphological identifications are made. In the absence of conidia, ascospores may be used to produce conidia in culture. If a positive diagnosis cannot be made using symptomology and conidial morphology, molecular diagnosis using gel-based PCR, is carried out as below.
DNA extraction
DNA is extracted directly from leaf tissue using a modified method of Stewart, C. N. and Via, L. E. (1993) “A rapid CTAB DNA isolation technique useful for RAPD fingerprinting and other PCR applications”. BioTechniques 14(5): 748-750
Consumables and equipment
Item Ordering Information
Pipettes: P20, P200, P1000 Pipetman P20, P100, P200 John Morris Scientific www.johnmorris.com.au
Sterile 1.5mL microfuge tubes Quantum 1.5mL Microtubes (500) Quantum Scientific P/L www.quantum-scientific.com.au Catalogue No. QSP505
Sterile plastic micropestles Kontes 1.5mL Pellet Pestles (100) Quantum Scientific P/L www.quantum-scientific.com.au Catalogue No. 749521-1500
Aerosol Resistant (plugged) tips Axygen Maximum Recovery, racked and pre-sterilised Quantum Scientific P/L www.quantum-scientific.com.au TF-20-L-R-S (suits P20) TF-200-L-R-S (suits P200) TF-1000-L-R-S (suits P1000)
Microcentrifuge (24-well)
29 Cork Borer (5mm, ethanol sterilisable)
Gas flame
Heating waterbath (set to 70ºC)
Small liquid nitrogen dewar/flask
Bench top shaking platform or similar
Reagents
Liquid nitrogen
CTAB Extraction Buffer (2% w/v N-cetyl-NNN-trimethyl ammonium bromide, 1.42M NaCl, 20mM EDTA, 100mM Tris-HCl (pH8.0), 2% polyvinylpyrrolidone (PVP-40), 5mM ascorbic acid, 4mM diethyldithiocarbamic acid (DIECA) See appendix for extraction buffer preparation details
β-mercaptoethanol
Chloroform-isoamyl alcohol (24:1)
Isopropanol (100%)
Sterile dH2O
Sterile, acid-washed sand
Notes
• DNA extraction should be carried out in an area which is physically separated from areas used for PCR preparation and PCR analysis, using equipment dedicated to extraction only
• Pre-warm extraction buffer at 70ºC
• Pipetting for steps 4-7 should be carried out in a fume cupboard
30 Procedure
1. Using a 5mm cork borer, excise 8-15 discs (approximately 50mg) from fungal leaf spot lesions on banana leaves
2. Place the excised leaf discs into a sterile microfuge tube. Samples may be stored at 4ºC or –20ºC until ready to extract
3. Dip microfuge tube in liquid nitrogen and then homogenise using a micropestle (Alternative method available, see note below ♦). A small amount of sterile, acid- washed sand may be used to aid grinding. Complete homogenisation is not necessary
4. Add 600µL of pre-warmed N-cetyl-NNN-trimethyl ammonium bromide (CTAB) extraction buffer (70ºC) and 3µL β-mercaptoethanol to the microfuge tube and continue homogenisation to emulsify
5. Add 500µL chloroform-isoamyl alcohol (24:1) and shake homogenate at room temperature for 5 min on a shaking platform set to 500rpm
6. Centrifuge at 1000g for 5 min at room temperature
7. Transfer the aqueous phase to a fresh microfuge tube and add 0.7 X volume of 100% isopropanol (approximately 350µL)
8. Mix by inversion and incubate at room temperature for 5 min
9. Centrifuge at 14000g in a benchtop microfuge for 20 min at room temperature
10. Draw off the supernatant using a pipette tip and discard. Air dry the pellet for approximately 15-20 min, taking care not to over dry. When no traces of isopropanol can be detected, resuspend in 100µL sterile water
♦ Alternative homogenisation method
1. If available, a Fastprep FP120 machine (Bio101) or similar apparatus, can be used to homogenise the sample. This is extremely beneficial if large numbers of samples are being extracted. For this method the leaf discs are placed in a 2mL screw top tube with O-ring together with approximately 250mg of 1.5–2.5 mm glass beads
2. Add 600µL of pre-warmed CTAB extraction buffer (70ºC) and 3µL β-mercaptoethanol to the tube
3. Process at 4 m/sec (speed 4) for 45 seconds, using the Fastprep FP120 machine (Bio101)
4. Continue as above from step 5
31 PCR Amplification
Background: The Sigatoka PCR diagnostic assay design
The Sigatoka PCR assay is designed to amplify specific sections of the ribosomal DNA of M. fijiensis and M. musicola. Using this region allows good species differentiation as well as good sensitivity as ribosomal DNA is present in high copy number in the pathogen genome.
Specificity of the PCR assays have been achieved using sequence polymorphisms in the internal transcribed spacer (ITS) regions; the black Sigatoka diagnostic primer (MFfor) is designed to the ITS1 region, while the yellow Sigatoka diagnostic primer (MMfor) is designed to the ITS2 region (Figure 12). Using different regions of the ITS for detection of yellow and black Sigatoka also introduces a size differential between the PCR products. This allows quicker identification of the result and serves as an assay control in the laboratory.
The reverse primers for both the yellow and black Sigatoka assays are located in the 26S ribosomal gene region. For the yellow Sigatoka assay, the forward primer (MMfor) is paired with the universal reverse primer R635 (Liu et al., 1991) to amplify a PCR product of size approximately 650 bp. In the black Sigatoka assay, the forward primer (MFfor) is paired with a modified version of the universal reverse primer R635 to amplify a PCR product of size approximately 1000bp. The modified reverse primer (R635-mod) was produced by shortening R635 by two bases at the 3’end. This allows more efficient amplification of black Sigatoka when used in a PCR with primer MFfor.
Yellow Sigatoka ∼650bp
MMfor R635 ITS1 ITS2 18S (partial) 5.8S 26S (partial)
MFfor R635mod
Black Sigatoka ∼1000bp
Figure 12: Primer map showing the location of Sigatoka diagnostic primers in the ITS regions and the expected sizes of the PCR products.
32 The Banana Internal Control Assay
To avoid potential false negative results, an internal control PCR assay has been developed to the banana DNA in the extract. A primer pair (MWPL1-110 and MWPL1-197) has been designed to amplify a 180 base pair fragment of the putative banana pectate lyase encoding gene (MWPL1) isolated from Musa acuminata cv Williams. This is a single copy gene which is differentially expressed in ripening fruit. Our research has demonstrated that this band can be amplified in all tissue types from a wide range of cultivars within the genotypes AA, AB, AAB, ABB, AAA, AAAA and BB.
The internal control reactions should be set up in parallel with the diagnostic assays. For each sample to be tested, an internal control assay should be included.
PCR Primer Sequences
Target Primer Name (direction) Sequence (5’→3’)
M. fijiensis MFFor (forward) CGGAGGCCGTCTAAACACT
M. fijiensis R635-mod (reverse) GGTCCGTGTTTCAAGAC
M. musicola MMFor (forward) CCGCGGCCGTTAAATCTTCAAA
M. musicola R635 (Liu et al., 1991) GGTCCGTGTTTCAAGACGG
Banana Internal MWPL1-110 (forward) GTCCGCGACCCTGAATTAGTAGT Control
Banana Internal MWPL1-197 (reverse) CGGTGCCGCATGACAAG Control
Consumables and equipment
Item Ordering Information
Pipettes: P2, P20, P200, P1000 Pipetman P2, P20, P100, P200 John Morris Scientific www.johnmorris.com.au
Aerosol Resistant (plugged) tips Axygen Maximum Recovery, racked and pre- sterilised Quantum Scientific P/L www.quantum-scientific.com.au TF-300-L-R-S (suits P2)
33 TF-20-L-R-S (suits P20) TF-200-L-R-S (suits P200) TF-1000-L-R-S (suits P1000)
Sterile 1.5mL microfuge tubes Quantum 1.5mL Microtubes (500) Quantum Scientific P/L www.quantum-scientific.com.au Catalogue No. QSP505
PCR Strip tubes with attached lids PCR 8-strip tubes (120 strips) www.eppendorf.com.au Catalogue No. 0030 124.359
PCR rack
Microcentrifuge
Capsulefuge (suit 8-strip PCR tubes)
96-well, 0.5mL plate format PCR Thermal MJ Research PTC-100 Cycler. The Sigatoka assay was optimised on Geneworks a MJ Research PTC-100 but most brands should be suitable www.geneworks.com.au Call for specs and quote
Reagents
Item Ordering Information
Taq DNA polymerase, enzyme buffer and AmpliTaq Gold DNA Polymerase with Buffer MgCl2 II and MgCl2 (250 U) Applied Biosystems www.appliedbiosystems.com.au Catalogue No. N8080241 dNTPs (set of dATP, dCTP, dGTP, dTTP) dNTP Set (40µmol or 100mM each) Promega www.promega.com.au Catalogue No. U1240
Sterile water Millipore® or injectable grade
PCR primers 40nmol scale, HPLC-purified grade www.geneworks.com.au
34 Preparation and storage of PCR stocks
Reagent Details
Sterile water • Obtain injectable water • Store in sterile vials at room temp
10X Buffer II • Supplied with enzyme • Aliquot on first thaw and be sure to mix well before using • Store –20ºC
MgCl2 • Supplied with enzyme • Aliquot on first thaw and mix well before using • Store –20ºC dNTP mix • Purchased as individual stocks of dATP, dCTP, dGTP and dTTP at 100mM • Take 10µL of each into 460µL sterile water to give a final working solution of 2mM each dNTP • Store aliquotted at –20ºC and avoid multiple freeze/thaws where possible
Primer stocks • Resuspend lyophilised pellet to 100µM • To prepare a 20µM working solution, dilute 1:5 in sterile water and store aliquotted at –20ºC • Do not dilute all stock as primers are more stable at higher concentrations
AmpliTaq Gold DNA Polymerase (250U) • Supplied at 5 U/µL, use 0.2µL (1U) per reaction • Store at –20ºC in a cyclic defrost freezer
35 Notes
• PCR set-up should be carried out in a designated, “PCR-clean” area, using dedicated equipment. A PCR-clean laboratory coat should be worn. No movement of equipment or reagents should occur between set-up and processing of PCR products, excepting prepared reactions which may travel one- way to the template addition laboratory. Backwards movement of racks between the template addition area to the PCR preparation area must be avoided.
• Internal control PCRs must be set up in parallel with diagnostic samples for quality control.
• It is recommended to only set-up one PCR experiment per day. PCR set-up should never directly follow handling of PCR products.
• Reagents, except Taq DNA polymerase, should be thoroughly thawed, mixed and pulse-centrifuged to collect contents at the bottom of the tube.
• Preparation of mastermixes is recommended when multiple samples are to be analysed; this permits standardisation of reagents across the tubes and minimises pipetting errors. When preparing mastermixes, include two extra tubes for positive and negative PCR controls. Also, allow extra tubes (usually 10% of the number being tested, ie. 20 samples + 2 extra) to account for losses due to pipetting errors caused by tip retention during aliquotting.
36 PCR set-up Procedure
1. In a sterile microfuge tube, prepare mastermixes for each of the black and yellow Sigatoka assays by adding the reagents in the order they appear in the table below. Reagent amounts are for a single reaction of final volume 25µL, allowing for addition of 1µL DNA template or 1µL sterile water as a negative PCR control. At the same time as you are setting up the diagnostic assays, ensure that a separate internal control assay is set up for each sample being screened
Sigatoka Diagnostic PCR Set-up
Reagent Final M. musicola M. fijiensis Concentration (Yellow Sigatoka) (Black Sigatoka)
Sterile water 15.55µL 15.55µL
10X PCR Buffer II 1X 2.5µL 2.5µL
MgCl2 (25mM) 3mM 3µL 3µL dNTPs (2mM) 100µM 1.25µL 1.25µL
Primer MMfor (20µM) 0.6µM 0.75µL -
Primer R635 (20µM) 0.6µM 0.75µL -
Primer MFFor (20µM) 0.6µM - 0.75µL
Primer R635-mod (20uM) 0.6µM - 0.75µL
TaqGold DNA Polymerase 1U 0.2µL 0.2µL (5U/µL)
TOTAL 24µL 24µL
37 Banana Internal Control PCR Set-up
Reagent Final Concentration Banana Internal Control
Sterile water 15µL
10X PCR Buffer II 1X 2.5µL
MgCl2 (25mM) 3mM 3µL dNTPs (2mM) 200µM 2.5µL
Primer MWPL1-110 (20µM) 0.3µM 0.4µL
Primer MWPL1-197 (20µM) 0.3µM 0.4µL
TaqGold DNA Polymerase 1U 0.2µL (5U/µL)
TOTAL 24µL
2. Aliquot 24µL to PCR 8-strip tubes. Add 1µL of sterile water to the negative control tube. To ensure a reliable control against contamination during mastermix preparation, the negative control tube must remain closed from this point until post- PCR analysis.
3. Transfer the tubes to the template addition section. Where possible, racks should not be removed from the PCR clean section. If this cannot be avoided, the tubes should be transferred to another rack before template addition and the “template-free” rack returned to the PCR clean laboratory immediately.
4. Using filter tips, add 1µL template DNA (from a 1:100 dilution of extract) to individual PCR tubes. The use of plugged tips at template addition ensures the source of the DNA being tested and prevents cross-contamination from pipette barrels. Open only one PCR tube at a time and close the lid immediately following template addition. Add 1µL of positive control DNA to the last tube.
Important: take care to prevent cross-contamination between template tubes and PCR tubes. Opening template tubes with the left-hand and opening PCR tubes with the right hand is good practice in preventing cross-contamination of samples.
5. Pulse-spin the PCR strip-tubes using a capsule microcentrifuge to collect the DNA template at the bottom of the tube with the PCR reagents.
38 6. Proceed to PCR thermal cycling or store reactions at 4ºC until ready. Following storage at 4ºC, condensation will form on the lid of the PCR tubes and another pulse spin will be necessary.
PCR thermal cycling Procedure
1. Programming should be carried out in a 96-well plate thermal cycler with a hot-lid attachment. If a hot-lid cycler is not available, add 20µL of sterile paraffin oil to each reaction (in the PCR clean room, before proceeding to template addition) to prevent evaporation of reagents during cycling.
2. Thermal cycling:
Yellow Sigatoka Diagnostic PCR Step Temperature Time (minutes:seconds) 1 94°C 10:00 2 94°C 0:20 3 65°C 0:30 72ºC 0:30 4 Go to step 2, 39 more times 5 72°C 3:00 6 4°C 5:00 7 End
Black Sigatoka Diagnostic PCR Step Temperature Time (minutes:seconds) 1 94°C 10:00 2 94°C 0:20 3 64°C 0:30 72ºC 1:00 4 Go to step 2, 39 more times 5 72°C 3:00 6 4°C 5:00 7 End
Banana Internal Control PCR Step Temperature Time (minutes:seconds) 1 94°C 10:00 2 94°C 0:20 3 60°C 0:30 72ºC 0:30 4 Go to step 2, 39 more times 5 72°C 2:00 6 4°C 5:00 7 End
3. Reactions may be stored at 4ºC until ready to be analysed by gel electrophoresis
39 PCR analysis using gel electrophoresis
Consumables and equipment
Item Ordering Information
Pipette: P20 Pipetman 20 John Morris Scientific www.johnmorris.com.au
Pipette tips Gel loading tip Quantum Scientific P/L www.quantum-scientific.com.au Catalogue No. QSP-010 (1000, bagged, unsterilised) Catalogue No. QSP-010-R204 (1020, racked, unsterilised)
Nunc Microwell Minitrays HLA Plate 60-well Medos www.medos.com.au Catalogue No. NUN4-52256
Capsulefuge (suit 8-strip PCR tubes)
Gel-electrophoresis set-up (eg. BioRad Sub- Cell GT) 20 well combs (up to 4 combs depending on throughput required)
Reagents
Item Ordering Information
Agarose Agarose I (Biotechnology Grade; DNase, RNase, Protease Free) 500 g Astral Scientific P/L www.astralscientific.com.au Catalogue No. AM0710
DNA marker and 6X Gel loading buffer GenerulerTM 100bp Ladder Plus (0.05mg) supplied with 1mL 6X Loading Dye Solution MBI Fermentas www.fermentas.com
40 Catalogue No. SM0321
Tris-Borate EDTA Running Buffer (10X) TBE Buffer 10X Liquid (4L) Astral Scientific P/L www.astralscientific.com.au Catalogue No. AM0658
Ethidium bromide Ethidium Bromide (10mg/mL) Astral Scientific P/L www.astralscientific.com.au Catalogue No. AME406
Notes • A set of pipettes must be dedicated only to gel analysis. Amplified (PCR) product poses the highest risk of contamination of any template
• Gel-electrophoresis and preparation of PCR products for analysis must be kept separate at all times from areas used for DNA extraction and PCR set-up
• Visualisation of PCR products using ethidium bromide (EtBr) may be carried out by post-staining or by running EtBr in the gel. Protocols for both options are presented here
• CAUTION: Ethidium Bromide is a powerful mutagen, and possible carcinogen and teratogen. Refer to MSDS data sheets and safety information on the correct handling of EtBr before proceeding
Gel electrophoresis Procedure
1. Prepare a 1.5% agarose gel in 0.5X TBE. A gel with 20-wells is sufficient to run 16 samples, positive and negative PCR reactions and two molecular weight markers. If required, EtBr can be included in the gel; add 5 µL of a 10mg/mL solution to 100mL (final concentration of 0.5µg/mL).
2. While the gel is setting, prepare the PCR products for analysis. To 5µL of PCR product add 1µL of 6X gel loading buffer. Prepare enough molecular weight marker for two wells by adding 2µL (500ng) of GenerulerTM 100 bp Ladder Plus, 8µL sterile distilled water and 2µL 6X gel loading buffer.
3. Place the set gel into the buffer tank and cover with 0.5X TBE running buffer before removing the gel comb.
41 4. Load the samples (6µL) onto the gel in the following order: DNA marker, negative PCR control, test samples, positive PCR control, DNA marker.
5. Run the gel at 6V/cm until the bromophenol blue dye is three quarters along the length of the gel.
6. If EtBr has not been run in the gel, post-stain in water containing EtBr at (0.5µg/mL) for 20 min. If necessary, destain in water for 10-15 min.
Results Interpretation
a. Sigatoka Diagnostic PCR
A sample is positive for yellow Sigatoka if a PCR product of approximately 650bp is visible. Size can be estimated by comparison to bands of similar size in the DNA molecular marker lane. To control against cross-contamination and false positive results, the negative PCR control should not have amplified (no band present). As a control for the efficiency of the yellow Sigatoka PCR, a band of approximately 650 bp should be present in the positive control lane.
A sample is positive for black Sigatoka if a PCR product of approximately 1000bp is visible. Size can be estimated by comparison to bands of similar size in the DNA molecular marker lane. To control against cross-contamination and false positive results, the negative PCR control should not have amplified (no band present). As a control for the efficiency of the black Sigatoka PCR, a band of 1000bp should be present in the positive control lane.
b. Banana Internal Control PCR
The results of the Sigatoka assays can be accepted as correct if a 180bp band is visualised for each sample tested. If no band is present from the internal control PCR reactions then refer to the trouble shooting guide and reprocess the sample. Figure 13 illustrates typical amplification results for the yellow and black Sigatoka PCR assays, and the banana internal control. Extraction buffer controls are useful when a new batch of buffer has been prepared to demonstrate the batch as template-free.
42 YS BS BIC M M 1 2 3 4 5 6 7 8 9 10 11 12 13 14
1000bp 650bp
180bp
Figure 13. Typical amplification products for the yellow and black Sigatoka diagnostic PCR assays, including banana internal controls. Lane 1: 500 ng GeneRulerTM 100 bp Ladder Plus; Lanes 2-5: Yellow Sigatoka diagnostic assay (–‘ve PCR control; extraction buffer control; banana with severe mixed yellow & black Sigatoka infection; banana with early black Sigatoka infection); Lanes 6-9: Black Sigatoka diagnostic assay (- ‘ve PCR control; extraction buffer control; banana with severe mixed yellow and black Sigatoka infection, banana with early black Sigatoka infection); Lanes 10-13: Banana internal control assay (-‘ve PCR control; extraction buffer control; banana with severe mixed yellow and black Sigatoka infection, banana with early black Sigatoka infection); Lane 14: 500ng MassRulerTM DNA Ladder.
43 Troubleshooting
Troubleshooting Guide 1 - DNA Extraction
Problem Possible Causes Suggestions
No DNA pellet at extraction end • Not enough sample tissue • Use 50mg plant material • Sample has not been macerated enough • Add a small amount of sterile sand to facilitate • Extraction buffer has not been made up homogenisation correctly • Check buffer preparation and/or remake • Pellet is lost after decanting off liquids after • Use a pipette to remove supernatant centrifugation steps • Failure to add isopropanol
Gelatinous-like material present in the DNA pellet • Excess polysaccharide in the sample • A 1:100 dilution of the resuspended sample should dilute out possible PCR inhibitors
Coloured DNA pellet • Tannins present in sample • A 1:100 dilution usually amplifies well in PCR • Commonly found when extracting from despite the extract colour necrotic lesions (usually of no consequence)
Difficulty resuspending DNA pellet • Accumulation of salts, polysaccharides and • Avoid using a desiccator; 15-30 min drying on other insoluble contaminants in the plant the bench top is often sufficient tissue • Complete resuspension is not always • Overdrying of pellet necessary for diagnostic screening • Incubating the sample at 55ºC for 60 min may improve solubility
44 Troubleshooting Guide 2 - Gel Electrophoresis
Problem Possible Causes Suggestions
Sample escapes from bottom of the wells • Gel comb height set too low on adjustable • Check comb height is 2-3mm above tray comb holders before pouring gel • Bottom of the well pierced with a tip during • Do not place tip low into well when loading loading • Wait until gel is set before removing comb (15- • Gel not set when comb removed 20 min)
Samples have run backwards • Electrodes or gel tank lid has been fitted • DNA samples are –‘ve charged and so must backwards (possible on older gel rigs) always run from the black (-‘ve) cathode to the red (+’ve) anode
Samples have not run out of well • Power pack failure • Always check for voltage by looking for • Incorrect fitting of electrodes or tank lid bubbles rising from cathode • Check current (should be 40-50 mA at 100V) • Ensure correct dilution of TBE is used in the agarose gel and tank
• Ensure correct dilution of TBE is used in the Gel bands wavy, fuzzy or absent • Incorrect concentration of TBE buffer in gel or agarose gel and tank tank • Water used instead of TBE (common error) • Take care to remove air bubbles before gel • Air bubble in agarose gel begins to set • Incomplete dissolving of agarose before • Ensure all agarose is dissolved before cooling pouring gel and pouring gel Sample flows up and out of well • Insufficient glycerol or sucrose in gel loading • Check concentration of sucrose/glycerol in buffer (GLB) loading buffer or use commercially prepared
45 • Too little GLB added to sample loading buffer • Air bubble in gel loading tip • Add GLB: sample in 1:6 ratio • Sample loaded too fast • Check pipetting technique • Sample well too shallow for volume added • Check height that gel comb is set at
No bands visible on agarose gel when viewed • Incorrect concentration of EtBr added to gel • Check concentration of EtBr added to gel under UV light (including molecular weight • Post-staining EtBr solution old • Post-stain in newly prepared EtBr solution markers)
Uneven EtBr staining of gel (seen as patchiness or • EtBr added to agarose was not mixed properly • Swirl EtBr through cooled agarose (60ºC) regions of intense fluorescence) • Post-staining with multiple gels in the tank before pouring gel does not allow good contact with the solution • Do not overload post-stain tank with many gels • Destain with water and restain with fresh EtBr
46 Troubleshooting Guide 3 – PCR Diagnostic Results
Problem Possible Causes Suggestions
Failed positive control reaction • Degraded DNA template • Repeat PCR with new DNA template • Inhibitors in DNA template (preferably extracted from single-spored • Too high or too low DNA template culture to avoid PCR inhibition) concentration • Use 1-10ng DNA as control • PCR Primers degraded • Repeat PCR with new aliquots of primers, • dNTPs too old dNTPs, buffers, MgCl2 making sure to mix • PCR primer or dNTP preparation errors components thoroughly before addition • Taq DNA polymerase inactive
• Enzyme buffer or MgCl2 not completely mixed • PCR thermal cycler error or power outage
Failed negative control reaction • Contamination of one or more PCR reagents • Do not spend time trying to determine the exact source of the contamination • PCR reagents should be aliquotted such that when PCR contamination occurs, the aliquot in use at the time can be discarded • Repeat experiment with new aliquots of reagents Unexpected black Sigatoka positive result: • Cross-contamination of DNA templates • Ensure DNA +’ve control is after test samples • The black Sigatoka PCR diagnostic assay is • Check extraction buffer and negative PCR • Symptoms not expected for black Sigatoka both highly specific and sensitive: the result is controls • Unexpected incursion of the disease most likely real • Repeat DNA extraction of leaf sample and • First record of black Sigatoka in a region repeat PCR
• To further confirm the result, the diagnostic PCR product may be cloned and sequenced
47 or the ITS regions of the pathogen may be PCR-cloned and sequenced using universal ITS primers (White et al., 1990)
Unexpected yellow Sigatoka positive result: • Cross-contamination of DNA templates • Ensure DNA +’ve control is after test samples • The yellow Sigatoka PCR diagnostic assay is • Check extraction buffer and negative PCR • Symptoms not expected for yellow Sigatoka both highly specific and sensitive: the result is controls • Yellow Sigatoka not expected in region eg. most likely real • Repeat DNA extraction of leaf sample and some parts of south-east Asia and Oceania • In Australia, yellow Sigatoka is endemic and is repeat PCR
expected to amplify from DNA extracted from • To further confirm the result, the PCR product most leaf spot lesions. In addition, black may be cloned and sequenced or the ITS Sigatoka and yellow Sigatoka are often found regions may be PCR-cloned and sequenced together during a black Sigatoka incursion using universal ITS primers (White et al. 1990)
48 Limitations of the Technology
Due to the nature of PCR-based protocols, specific issues arise because of the high level of sensitivity and amplification of millions of copies of the target sequence. A few molecules of PCR-generated fragments can contaminate samples of subsequent PCR runs and result in false positives. On the other hand, a low copy number of initial target DNA sequences makes the first amplification cycles critical e.g. PCR inhibition can result in false negatives.
False positives can result from cross-amplification of non-target DNA, exogenous DNA from cells/cultures or aerosols, or from contaminating DNA originating from carry-over of previous experiments. A negative control (that contains no template DNA) should be included in all PCR diagnostic tests to identify false positive results. Aerosol-resistant, or filter-plugged tips ensure DNA template added to the reaction tube, is not contaminated by templates present in the barrel of the pipettor.
False negatives can arise for many reasons, including the presence of compounds derived from extracted substrates that inhibit Taq polymerase, degradation of the DNA target sequence, or reagent problems. Including a positive control for DNA (a known positive DNA sample) and the presence of an amplified internal standard for each sample can protect against false negatives.
PCR reaction inhibitors
Plant-derived compounds that inhibit the PCR reaction have been well documented e.g. phenolics in banana tissue and chemical sprays on leaf surfaces. This problem is usually easily solved by the further dilution of samples to 1:1000 in water.
49 Sources of Reference Material
Fungal cultures and DNA
Please contact the Queensland Department of Primary Industries & Fisheries Plant Pathology Herbarium (BRIP) for isolates of M. musicola and M. fijiensis to be used as positive controls in these assays.
Please note that only cultures of M. musicola, and heat-treated or γ-irradiated plant tissue infected with M. fijiensis and M. musicola, are held by the Plant Pathology Herbarium.
Plant Pathology Herbarium, Qld Dept of Primary Industries & Fisheries, Plant Pathology Building, 80 Meiers Road, Indooroopilly, Qld, 4068
Ph: +61 7 3896 9598 Fax: +61 7 3896 9533
To enquire about availability of M. fijiensis and M. musicola DNA contact:
Dr Juliane Henderson Research Fellow (Banana Diagnostics) Tree Pathology Centre University of Queensland
Located at:
Qld Dept of Primary Industries & Fisheries, Plant Pathology Building, 80 Meiers Road, Indooroopilly, Qld, 4068
Ph: +61 7 3896 9348 Fax: +61 7 3896 9533
50 Contacts
Dr Juliane Henderson Research Fellow (Banana Diagnostics) Tree Pathology Centre University of Queensland
Located at:
Qld Dept of Primary Industries & Fisheries, Plant Pathology Building, 80 Meiers Road, Indooroopilly, Qld, 4068
Ph: +61 7 3896 9348 Fax: +61 7 3896 9533
Kathy Grice Experimentalist Horticulture and Forestry Science, Delivery
Located at:
Centre for Tropical Agriculture (CTA) Qld Dept of Primary Industries & Fisheries, 28 Peters Street, Mareeba, Qld, 4880
Ph: +61 7 4048 4675 Fax: +61 7 4092 3593
51 Further Reading
Carlier, J., De Waele, D. & Escalant, J. (2003) Global evaluation of Musa germplasm for resistance to Fusarium wilt, Mycosphaerella leaf spot diseases and nematodes. INIBAP Technical Guidelines No. 6
Available online from www.inibap.org
D.R. Jones (ed) (2000) Diseases of Banana, Abacá and Enset. CABI Publishing, New York.
Henderson, J., Pattemore, J. A., Porchun, S. C., Hayden, H. L., Van Brunschot, S., Grice, K. R. E., Peterson, R. A., Thomas-Hall, S. R. and Aitken, E. A. B. (2006) Black Sigatoka disease: new technologies to strengthen eradication strategies in Australia. Australasian Plant Pathology 35, 181-193.
52 Appendix
Reagents for DNA extraction
CTAB extraction buffer (100 mL)
Stock Component Volume Added Final Concentration
CTAB 2g 2% (w/v)
5M NaCl 28.4mL 1.42M
0.5M EDTA 4mL 20mM
1M Tris-HCl (ph8.0) 10mL 100mM
PVP-40 2g 2% (w/v)
Ascorbic Acid (MW = 176.12) 88mg 5mM
DIECA (MW = 171.3) 68mg 4mM
Dissolve components completely in MilliQ water, warming up to 70ºC if required. Do not heat above 70ºC, as some components are heat labile. Divide into smaller aliquots and store out of the light. Discard unused buffer after 6 weeks.
**DO NOT AUTOCLAVE THIS BUFFER – HEAT LABILE**
5M NaCl Stock Solution
• Place 146.1g NaCl in 1 litre beaker and make up to 500mL with MilliQ water • Stir until dissolved • Divide into smaller aliquots and autoclave (121°C/15 mins)
0.5M EDTA (pH 8.0)
• Place 143.1g of EDTA in a 1 litre beaker and add 400mL of MilliQ water • Stir vigorously whilst adjusting the pH of the solution to 8.0 with NaOH (approximately 10g of solid NaOH)** • Make up to 500mL with MilliQ water • Divide into smaller aliquots and autoclave (121°C/ 15 mins)
** The disodium salt of EDTA will only go into solution when the pH has been adjusted to 8.0 with NaOH. **
53 1M Tris-HCl (ph8.0)
• Place 60.55g Tris Base in a 1 litre beaker and add 400mL of MilliQ water • Stir whilst adjusting the pH of the solution to 8.0 with concentrated HCl (42mL/L final solution)** • After solution has returned to room temperature make up to 500mL and make final pH adjustment • Divide into smaller aliquots and autoclave (121°C/ 15 mins)
** This solution will become very warm after the addition of the concentrated HCl but this is normal. **
Chloroform-Isoamyl Alcohol (24:1) To make up 100mL • mix 96mL (24 parts) chloroform with 4mL (1 part) isoamyl alcohol.
** This solution must be made up and used inside a fume cabinet. ** Chloroform is toxic and a possible carcinogen.
Reagents for Agarose Gel Electrophoresis
0.5 X Tris-borate (TBE) Running Buffer
If not purchasing the concentrated TBE make a 5 X stock solution as follows:
54g Tris base 27.5g boric acid 20mL 0.5M EDTA (pH 8.0)
• Add the above to 800 mL of distilled water and stir until dissolved • Make up to a final volume of 1.0L
Store at room temperature and discard if precipitation occurs. To make a working solution of 0.5 X TBE dilute 1:10 with MilliQ water.
54 6 X Gel-Loading Buffer
Four different gel-loading buffers are provided below. Personal preference and availability of individual components will dictate choice.
1. 0.25% bromophenol blue 0.25% xylene cyanol FF 40.0% (w/v) sucrose Dissolve above components in MilliQ water and store at 4° C.
2. 0.25% bromophenol blue 0.25% xylene cyanol FF 15% Ficoll 400 Dissolve above components in MilliQ water and store at room temperature.
3. 0.25% bromophenol blue 0.25% xylene cyanol FF 30.0% (w/v) glycerol Dissolve above components in MilliQ water and store at 4° C.
4. 0.25% bromophenol blue 40.0% (w/v) sucrose Dissolve above components in MilliQ water and store at 4° C.
55 References
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59