Wheat spindle streak mosaic virus DP - Plant Health Australia

DEPARTMENT OF PRIMARY INDUSTRIES

Wheat spindle streak mosaic virus

Diagnostic Manual

Wheat spindle streak mosaic virus Diagnostic Manual

2

Wheat spindle streak mosaic virus

Diagnostic Manual

Bonny Rowles-van Rijswijk

Dr James Cunnington

Dr Brendan Rodoni

Department of Primary Industries, Knoxfield

Private Mail Bag 15

Ferntree Gully Delivery Centre

Victoria 3156

Australia

Published by Primary Industries Research Victoria

Department of Primary Industries

Knoxfield

June 2005

© Copyright State of Victoria 2005

This publication is copyright. No part may be reproduced by any process except in

accordance with the provisions of the Copyright Act 1968.

Authorised by the Victorian Government, 1 Treasury Place, East Melbourne.

ISBN 1 74146 397 1


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This publication may be of assistance to you but the State of Victoria and its

employees do not guarantee that the publication is without flaw of any kind or is

wholly appropriate for your particular purposes and therefore disclaims all liability for

any error, loss or other consequence which may arise from you relying on any

information in this publication.

For more information about DPI visit the website at www.dpi.vic.gov.au

or call the Customer Service Centre on 136 186.


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Table of Contents

1. Introduction _________________________________________________ 6

2. Wheat spindle streak mosaic virus _______________________________ 7

2.1. Distribution and yield loss _______________________________________ 7

2.2. Wheat spindle streak mosaic virus description _______________________ 7

2.3. Host range of Wheat spindle streak mosaic virus ____________________ 10

2.4. Symptoms associated with Wheat spindle streak mosaic virus __________ 10

2.5. Diseases associated with causing symptoms similar to Wheat spindle streak

mosaic virus____________________________________________________ 15

2.5.1. Soil-borne wheat mosaic virus _______________________________________ 15

2.5.2. Wheat streak mosaic virus __________________________________________ 18

2.6. Management of Wheat spindle streak mosaic virus __________________ 19

3. Polymyxa graminis __________________________________________ 20

3.1. The impact of infection by the fungus, Polymyxa graminis _____________ 20

3.2. Description of Polymyxa graminis ________________________________ 21

3.3. Life cycle of Polymyxa graminis _________________________________ 24

3.3. Polymyxa graminis fungal isolation _______________________________ 30

3.4. Transmission of Wheat spindle streak mosaic virus by Polymyxa graminis _ 31

3.5. Management strategies for Polymyxa graminis ______________________ 33

4. Detection of Wheat spindle streak mosaic virus ____________________ 34

4.1. Diagnostic Flow-chart _________________________________________ 34

4.2. Sample collection ____________________________________________ 35

4.2.1. Plant ___________________________________________________________ 35

4.2.2. Soil ____________________________________________________________ 35

4.3. Total RNA extraction __________________________________________ 36

4.3.1. Equipment required _______________________________________________ 36

4.3.2. Reagents required ________________________________________________ 37

4.3.3. Method _________________________________________________________ 37

4.4. Detection of Wheat spindle streak mosaic virus in total RNA extracts using one

step RT-PCR ___________________________________________________ 39



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4.4.2. Reagents _______________________________________________________ 40

4.4.3. One-step RT-PCR detection of Wheat spindle streak mosaic virus___________ 42

4.5. DNA Sequencing of PCR Products _______________________________ 43


4.5.2. Reagents _______________________________________________________ 43

4.5.3. Method _________________________________________________________ 43

4.6. Detection of Wheat spindle streak mosaic virus with Enzyme-linked

immunosorbent assay (ELISA) _____________________________________ 44

4.6.1. Equipment_______________________________________________________ 44

4.6.2. Reagents _______________________________________________________ 44

4.6.3. Biological Reagents _______________________________________________ 45

4.6.4. Method _________________________________________________________ 45

5. Detection of Polymyxa graminis ________________________________ 47

5.1. Diagnostic Flow-chart _________________________________________ 47

5.2. Sample collection ____________________________________________ 48

5.2.1. Plant ___________________________________________________________ 48

5.2.2. Soil ____________________________________________________________ 48

5.3. Total DNA extraction __________________________________________ 49


5.3.2. Method _________________________________________________________ 49

5.4. Detection of Polymyxa graminis in total DNA extracts using PCR. _______ 50


5.4.2. Reagents _______________________________________________________ 51

5.4.3. Method _________________________________________________________ 52

5.5. DNA Sequencing of PCR Products _______________________________ 52

Acknowledgments _____________________________________________ 53

References __________________________________________________ 54

Related Articles _______________________________________________ 59

Appendix 1 – ELISA sample plans ________________________________ 61


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1. Introduction

The cost to the grains industry by incursions of exotic pests and pathogens has been

recognised by Plant Health Australia and the Department of Primary Industries,

Victoria. In preparation for potential incursions, it was decided to develop a

diagnostic capability for Wheat spindle streak mosaic virus (WSSMV), as this viral

disease has not been reported in Australia and poses a significant threat to the

grains industry. WSSMV outbreaks are favoured by cool climates (Cadle-Davidson

and Bergstrom, 2004), and it is feasible that this virus could establish under southern

Australian conditions.

WSSMV was first reported in 1957, in Ontario, Canada, and is now ranked amongst

the three most common viral diseases of wheat in this region (Tenuta and Johnson,

2003). WSSMV is transmitted by the soil-borne fungus, Polymyxa graminis

Ledingham (OMAF Staff, 2002). There is only one record of a “P. graminis-like”

fungus in Australia, that was lodged on 8 September 1959, in Murrumburrah, NSW,

on Poa annua L. (DAR 48987) (APPD, 2005). It must be noted that the identification

was not to subspecies level, and that this record has never been confirmed.

P. graminis is non-pathogenic (Kanyuka et al., 2003), but its the ability to acquire and

transmit a range of plant viruses via zoospores (swimming spores) that invade root

hairs and epidermal cells of young plants during periods of high soil moisture that

results in serious diseases in cereal crop species (Kanyuka et al., 2003; OMAF Staff,

2002). The fungus can remain in the soil for at least 8 years (OMAF Staff, 2002),

with viral particles protected from the environment within resting spores (Kanyuka et

al., 2003). The persistent, soil-borne nature of the WSSMV-vector P. graminis,

makes the use of virus-resistant crop varieties the only practical and environmentally

friendly means of control (Kanyuka et al., 2003).

As the virus and vector are so intrinsically linked, this diagnostic protocol will deal

with methods to detect both WSSMV and P. graminis. The sampling and detection

methods in this manual have not yet been trialed under Australian conditions. It is

highly recommended that the methods be validated prior to establishment as a

national standard.


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2. Wheat spindle streak mosaic virus

2.1. Distribution and yield loss

WSSMV in France, Germany, India, Italy, Japan, North America, and possibly China

(Brunt et al., 1996 onwards; Jones, 2004).

Losses from WSSMV are greatest when a sensitive wheat variety is grown, and

when fields have had several crops of winter wheat in the past 8-10 years, resulting

in the contamination of soil over the entire paddock with the fungal vector P. graminis

(Jones, 2004; OMAF Staff, 2002; Tenuta and Johnson, 2003). Wheat yield losses

are attributed to a reduction in the numbers of tillers produced (Bowden, 2005).

Losses are dependent on the wheat variety and the weather, and are difficult to

measure. A long cool spring prolongs the symptomatic period and increases losses,

as the optimum temperature for symptom development is 5°C-15°C (Bowden, 2005;

OMAF Staff, 2002; Tenuta and Johnson, 2003). Alternatively, in a warm spring,

losses may be minimal (Bowden, 2005). Data from North America shows the

variability in yield loss due to WSSMV infection; in 2004 soft white winter wheat

losses were reported to be 2-18% in Michigan state, and 7-59% in Canada. In

Pennsylvania and Georgia, losses were estimated to be 7-24% and 22-35%,

respectively (Bowden, 2005).

2.2. Wheat spindle streak mosaic virus description

WSSMV is a member of the genus Bymovirus, family Potyviridae. The virus consists

of single stranded positive sense RNA encapsulated by a protein coat (Brunt et al.,

1996 onwards). The long flexuous filamentous WSSMV particles (fig. 1) are 16 nm

wide, with no clear modal length. Measurements range from 300-2000 nm, with the

modal length probably around 700-1000 nm (Brunt et al., 1996 onwards; Jones,

2004; Lommel et al., 1986). Virions are found in the cytoplasm of leaves, roots,

mesophyll and vascular parenchyma of WSSMV-infected plants, and distinctive

pinwheel inclusion bodies are produced (Lommel et al., 1986). Infection by WSSMV

causes the chloroplasts and mitochondria to swell, resulting in disruption of these

organelles (Brunt et al., 1996).

genus007.htm


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Figure 1. Electron micrograph of Wheat spindle streak mosaic virus. Par t icles t rapped

by an t iserum f rom in f ect ed leaves. Lengt h of bar [nm ]: 100 (Adam s, 2002).

WSSMV is transmitted by the obligate fungus P. graminis (Barr, 1979). The

subspecies of P. graminis that transmits WSSMV requires 3 hours at 15oC for

attachment and host penetration to occur, and WSSMV transmission has been

reported to occur within 16 days at temperatures ranging from 7.5 to 17.5oC (Cadle-

Davidson and Bergstrom, 2004). Environmental fluctuation through thermoperiodism

between 5 to 15oC (night-day) stimulates WSSMV transmission (Cadle-Davidson and

Bergstrom, 2004). The virus is not seed transmitted (Jones, 2004), nor is it

transmitted by grafting, pollen, or contact between plants (Brunt et al., 1996

onwards).

In an infected plant, Carrol et al. (1997) detected WSSMV in roots one month prior to

its detection in leaves, suggesting that virus moves slowly from roots to leaves.

WSSMV was detected in non-symptomatic leaves from two months after sowing,

through to crop senescence, and it was concluded that WSSMV is acquired and

spread by the vector during the majority of the crop cycle (Carrol et al., 1997).

There has been much confusion surrounding the relatedness of WSSMV to Wheat

yellow mosaic virus (WYMV), and how to distinguish the two viruses. Yellow mosaic

symptoms in wheat were first recorded in Japan, in 1927, and were attributed to

WYMV (Namba et al., 1998). In Canada similar symptoms in wheat were attributed

to WSSMV, and in the USA, India, France, Italy and China the causal agent has

been labelled either WYMV or WSSMV (Chen et al., 1999; Clover and Henry, 1999;


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Namba et al., 1998). A bymovirus occurring in rye, in Germany, has been described

as WSSMV or WYMV (Chen et al., 1999).

In 1979, Usugi and Saito drew the conclusion that WSSMV was a strain of WYMV

due to similarities in morphology, virus particle length, geographic distribution,

buoyant density, stability in sap, and because the two viruses had common antigens

in serological studies. However, there were slight differences in the symptoms

produced on infected wheat varieties, and not all serological antigens were common

to both viruses (Usugi and Saito, 1979). Hariri et al. (1996) also concluded that

WSSMV and WYMV were different strains of the same virus because of their strong

serological relationship. Carroll et al. (1995) developed a polyclonal antiserum that

reacted with bymovirus isolates of WSSMV, WYMV, and Barley yellow mosaic virus.

The International Committee on Taxonomy of Viruses website

(http://image.fs.uidaho.edu/vide/descr887.htm) continues to list WYMV as a strain of

WSSMV (Brunt et al., 1996 onwards).

The discrimination between WSSMV and WYMV is important as WSSMV has a

broader host range than WYMV, infecting Triticum aestivum and T. durum, as well as

Secale cereale. WSSMV infection has also been reported to breakdown field

resistance to Soil-borne wheat mosaic virus (SBWMV) in wheat (Clover and Henry,

1999; Lommel et al., 1986).

After generating sequence data from two fragments amplified from WYMV and

WSSMV (corresponding to RNA1 (7.6 kb) and RNA2 (3.6 kb)), Namba et al. (1998)

deduced that the two viruses shared only 77% amino acid sequence identity in their

coat proteins, and 74% nucleotide sequence identity in their 3’ non-coding regions.

This result had not been expected because of their close serological relationship, but

clearly suggests that WYMV and WSSMV are distinct virus species. This

discrimination was supported by sequencing of a 1.7 kb fragment representing the 3’-

terminal sequence of RNA1 of a Canadian WSSMV isolate, as this sequence shared

98.0% homology with the French WSSMV isolate used by Namba et al. (1998) (Lu et

al., 1998). Further analysis by Chen et al. (1999) was able to divide the isolates into

two distinct groups, i) WSSMV, for North American and European isolates, and ii)

WYMV for Asian isolates; based on differences in nucleotide between the groups of

over 30%. These findings were confirmed by Clover and Henry (1999), after analysis

of a further 23 viral sequences. As there was no serological method able to

discriminate between WSSMV and WYMV and no molecular diagnostic protocols


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available for either virus, Clover and Henry (1999) developed a WSSMV-specific RT-

PCR protocol. The WSSMV-specific RT-PCRs developed by Clover and Henry

(1999) and Gitton et al. (1999) form the basis of the diagnostic protocol

recommended for the detection of WSSMV (section 4).

2.3. Host range of Wheat spindle streak mosaic

virus

WSSMV infects wheat (T. aestivum, T. durum), which is used to maintain and

propagate the virus for experimentation, and rye. WSSMV does not infect barley

(Hordeum vulgare), nor any dicotyledonous species (Brunt et al., 1996 onwards;

Jones, 2004).

2.4. Symptoms associated with Wheat spindle

streak mosaic virus

WSSMV transmission occurs primarily during autumn and to a lesser extent in spring

(Carrol et al., 1997), with significant infections taking place during cool, wet autumn

periods. Often large areas of a field may be affected (Stromberg, 2002) (fig. 2 and

3), due to the uniform distribution of P. graminis, and WSSMV infection sites tend to

follow the lower, wetter areas of the field (Bowden, 2005).

Symptoms only appear in early spring, right after green-up (Bowden, 2005),

regardless of when plants have become infected (Carrol et al., 1997). However, the

earlier in the life cycle that the wheat is infected, the more severe the symptom

expression (Stromberg, 2002). The virus requires temperatures below 17oC for

symptom development (Lommel et al., 1986). The optimum temperature for

symptom development is 10oC (Smith, 2005), but symptoms will express between

9°C and 13°C air temperature (Stromberg, 2002), and 5 to 13oC soil temperature

(Smith, 2005). If soil temperatures exceed 15oC infected plants will fail to develop

mosaic symptoms, leaving little visual evidence of the disease (Carrol et al., 1997;

Smith, 2005), although symptoms may persist on older leaves (Bowden, 2005).

In favourable conditions during early spring, the symptoms of WSSMV appear as

yellow to light green streaks or dashes on a dark green background, in young leaves.


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The discontinuous streaks are usually 3 to 6.5 mm long, and oriented parallel to leaf

veins and taper at the ends to form chlorotic spindles (Bowden, 2005; Smith, 2005;

Tenuta and Johnson, 2003) (fig. 4 to 7). WSSMV is most easily diagnosed by

symptoms when the chlorotic spindle shaped lesions appear on the upper leaves of

the plant prior to heading (Smith, 2005).

During cool spring conditions, and as plants mature, the spindle-shaped lesions and

the chlorotic streaks on older leaves coalesce and become necrotic (Smith, 2005;

Stromberg, 2002). The older leaves look mottled with yellow and green blotches. If

temperatures remain cool, spindle shaped lesions will continue to develop throughout

spring in the upper new leaves (Smith, 2005). Warmer temperatures may prevent

symptom development on newly expanded leaves, giving the appearance that plants

have outgrown the disease, with symptoms confined to older, lower leaves (Smith,

2005).

WSSMV can also cause stunting, reduced tillering, smaller heads and less seed per

head than healthy plants, although the kernels produced by infected plants are not

shrivelled or light in weight (Bowden, 2005; OMAF Staff, 2002; Smith, 2005;

Stromberg, 2002; Tenuta and Johnson, 2003) (fig. 8 and 9).


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Figure 2. Infection by Wheat spindle streak mosaic virus is usually uniform in a wheat field,

because of the uniform distribution of the fungal vector, Polymyxa graminis (Source: http://www.uky.edu/Agriculture/IPM/scoutinfo/wheat/disease/wssm/wssm.htm).

Figure 3. Wheat field showing a large central patch of the crop affected by Wheat spindle streak mosaic virus (© Erik L. Stromberg, Department of Plant Pathology, Physiology, and

Weed Science, Virginia Tech).

Figure 4. Symptoms of Wheat spindle streak mosaic virus (WSSMV) infection typically

appear in early spring right after green-up. The symptoms of WSSMV are yellow to light green streaks or dashes on a dark green background. Dashes are usually 3 mm to 6.5 mm long.

The spindle-shaped dashes and streaks are oriented parallel to the leaf veins (©Robert L. Bowden, Kansas State University).


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Figure 5. Wheat spindle streak mosaic virus produces yellow streaks running parallel to the

leaf veins (Courtesy of Gary Bergstrom, © Queen's Printer for Ontario).

Figure 6. A wheat leaf with spindle streak symptoms associated with Wheat spindle streak

mosaic virus infection (© Erik L. Stromberg, Department of Plant Pathology, Physiology, and Weed Science, Virginia Tech).

/MBS/english/common/queens.html


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Figure 7. Symptoms of advanced Wheat spindle streak mosaic virus infection, showing well-

developed short light green to yellow streaks running parallel with the veins in leaves (Source: http://www.uky.edu/Agriculture/IPM/scoutinfo/wheat/disease/wssm/wssm.htm).

Figure 8. The effect of stunting caused by Wheat spindle streak mosaic virus (WSSMV)

infection is demonstrated through the use of WSSMV-resistant (left) and WSSMV-susceptible (right) wheat varieties (© Alabama Cooperative Extension System).

Figure 9. A wheat field with a cultivar susceptible to Wheat spindle streak mosaic virus growing on the left, and a resistant cultivar growing on the left, showing the reduction in

tillering due to virus infection (© Erik L. Stromberg, Department of Plant Pathology, Physiology, and Weed Science, Virginia Tech).


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2.5. Diseases associated with causing symptoms

similar to Wheat spindle streak mosaic virus

WYMV has not been included in this list, as symptoms caused by both WYMV and

WSSMV are very similar, and both infect wheat (Clover and Henry, 1999). These

viruses can only be accurately differentiated using molecular tests.

2.5.1. Soil-borne wheat mosaic virus

Soil-borne wheat mosaic virus (SBWMV) (synonym Wheat soil-borne mosaic virus)

and WSSMV have similar symptoms, life cycle and field distribution patterns, and

share the same fungal vector (Bowden, 2005; OMAF Staff, 2002; Tenuta and

Johnson, 2003). In certain cases, both SBWMV and WSSMV may be present in the

same field (OMAF Staff, 2002; Tenuta and Johnson, 2003), and the situation is

further confused by the fact that many plants are found to be infected with both

viruses (Bowden, 2005).

Although both viruses are vectored by the same fungus, the similarity between

WSSMV and SBWMV is only superficial, as the viruses are unrelated (Bowden,

2005; Linker et al., 1994; Smith, 2005). WSSMV belongs to the genus Bymovirus,

and SBWMV belongs to the genus Furovirus. Also, WSSMV infects wheat and rye,

whereas SBWMV can also infect barley (Linker et al., 1994).

As with WSSMV, symptoms of SBWMV infection often appear in low, wet areas, and

may cover all or most of a field due to the distribution of the fungal vector (fig. 10)

(Linker et al., 1994). Symptoms of WSSMV are usually visible prior to symptoms

associated with SBWMV, which typically appear in early spring right after green-up

(Bowden, 2005). The symptoms of SBWMV are most obvious in the early spring

months (Linker et al., 1994), and as with WSSMV, symptoms will not be expressed in

late spring when conditions are warmer (Linker et al., 1994; Smith, 2005).

The characteristic symptoms of WSSMV are the yellow-to-light green tapered streaks

running parallel to the leaf veins (OMAF Staff, 2002). In contrast, SBWMV typically

has a mosaic of green islands or blotches occurring on the leaves, ranging from mild

green to a prominent yellow that are longer than they are wide, against a yellow


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background (fig. 11 and 12) (Bowden, 2005; Linker et al., 1994; OMAF Staff, 2002;

Smith, 2005; Tenuta and Johnson, 2003). It must be noted that plants severely

infected with WSSMV resemble infection by WSBMV as they may develop mottled

leaves (Bowden, 2005), and that both WSSMV and SBWMV cause plant stunting

(fig. 13) (Linker et al., 1994; OMAF Staff, 2002).

Figure 10. Soil-borne wheat mosaic virus (SBWMV) field disease pattern (left), and patch of

SBWMV affected winter wheat (right) (Left; Source: http://www.plantpathology.tamu.edu/appel/vwheat3.htm, right; ©Department for Environment,

Food and Rural Affairs)

Figure 11. Soil-borne wheat mosaic virus (SBWMV) symptoms on wheat (left), and close-up

of SBWMV affected plants (©Department for Environment, Food and Rural Affairs)


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Figure 12. In each photograph, a healthy plant (left) is compared to a plant showing

symptoms of Soil-borne wheat mosaic virus infection (right).

Figure 13. Severe stunting of wheat caused by Soil-borne wheat mosaic virus

(© Ohio State University).


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2.5.2. Wheat streak mosaic virus

Wheat streak mosaic virus (WSMV) belongs to the genus Rymovirus (family

Potyviridae), and is transmitted by the wheat leaf curl mite (Aceria tulipae). This virus

produces symptoms that are similar to WSSMV infection, ie., stunted plants with

mottled green and yellow streaks extending the length of the leaf (fig. 14 and 15).

Plants can also exhibit rolling or curling of the youngest leaves due to damage by the

mite vector. Symptoms of WSMV typically appear in the spring (Stromberg, 2002).

Unlike WSSMV, WSMV field infection is not evenly distributed, infection is usually

located near the field edges, and infected plants continue to show symptoms

throughout the increasingly warm spring weather (Smith, 2005; Stromberg, 2002).

Figure 14. Streak mosaic pattern of Wheat streak mosaic virus (Left; © 2000 by the

University of Nebraska, right; http://www.oznet.ksu.edu/path-ext/Alerts/2002%20alerts/alert04.asp)

Figure 15. As the wheat crop develops, plants affected with Wheat streak mosaic virus are typically severely stunted with yellow mottled and streaked leaves. (© Erik L. Stromberg,

Department of Plant Pathology, Physiology, and Weed Science, Virginia Tech).


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2.6. Management of Wheat spindle streak mosaic

virus

There is no cure for a WSSMV-infected plant, and therefore preventative measures

are employed to reduce infection rates (Anon, 2003). Crop rotation as a

management tool to reduce the incidence of disease is only moderately effective

because the virus can remain infective in the soil for many years inside spores of the

fungal vector (Bowden, 2005; Linker et al., 1994; OMAF Staff, 2002; Tenuta and

Johnson, 2003). Manipulation of planting dates, irrigation schedules and paddock

drainage by avoiding exposure of crops to high risk low-lying wet areas (Bowden,

2005) could reduce virus incidence. Anecdotal evidence has shown fields that have

had liberal amounts of poultry and livestock manure added have a reduction in

WSSMV build-up (OMAF Staff, 2002; Tenuta and Johnson, 2003).

Chemical control is ineffective, as fungicides will not control the virus or the vector

(Anon, 2003) due to the longevity of the fungal resting spores (Campbell, 1996;

Kanyuka et al., 2003). The use of fumigants is not a practical or an affordable option

for broadacre farmers (Roget, 2004), and internationally, it is not an ecologically

acceptable form of disease control (Kanyuka et al., 2003).

The current thinking is that the only way to control WSSMV is to plant resistant

varieties (Anon, 2003). Presently there are no cultivars completely resistant to

WSSMV (Bowden, 2005), although many sources of resistance to WSSMV have

been identified in commercial wheat varieties and in wild Triticum species, with the

resistance trait being highly heritable (Kanyuka et al., 2003). Other wheat cultivars

such as Sierra, Mustang, and Thunderbird are highly susceptible and should be

avoided in high risk districts (Bowden, 2005).


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3. Polymyxa graminis

3.1. The impact of infection by the fungus, Polymyxa

graminis

Polymyxa graminis is a plasmodiophorid, a weakly pathogenic eukaryotic obligate

biotrophic parasite of plant roots with a worldwide distribution (Barr, 1979; Gitton et

al., 1999; Kanyuka et al., 2003; Ward et al., 2005), which includes New Zealand

(Jones, 2004). Although P. graminis is weakly pathogenic, its ability to vector cereal

crop diseases is what causes significant yield reductions (Kanyuka et al., 2003).

As previously stated, P. graminis is the vector for WSSMV. This fungus is also a

vector for the following viral diseases (Anon, 2005; Barr, 1979; Gitton et al., 1999;

Kanyuka et al., 2003; Morales et al., 1999; Ward et al., 2005):

- Aubian wheat mosaic virus

- Barley mild mosaic virus

- Barley yellow mosaic virus

- Chinese wheat mosaic virus

- Indian peanut clump virus (IPCV)

- Oat golden stripe virus

- Oat mosaic virus

- Peanut clump virus (PCV)

- Rice necrosis mosaic virus

- Rice stripe necrosis virus (RSNV)

- Soil-borne cereal mosaic virus

- Soil-borne wheat mosaic virus

- Sorghum chlorotic spot virus

- Wheat yellow mosaic virus

As the viruses are protected from the environment within the P. graminis resting

spores, which may remain dormant but viable for decades, the diseases are very

persistent (Kanyuka et al., 2003). After fungal spore germination, the viruses are

carried within the swimming zoospores to the plant host (Ward et al., 2005). The

host specificity of P. graminis is complex. For example, a West African isolate of P.

graminis that transmits RSNV is able to complete its lifecycle within rice roots. A

second West African isolate of P. graminis that is capable of transmitting PCV can

not complete its lifecycle within rice roots (Anon, 2005). It is therefore imperative to

understand the biology of local isolates of P. graminis and their ability to transmit

plant viruses.


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3.2. Description of Polymyxa graminis

Polymyxa graminis is an obligate root-infecting organism that was originally

described from wheat by Ledingham (1939). Barr (1979) examined roots of common

weeds for the presence of plasmodiophoraceous fungi, but was unable to identify a

natural reservoir of host species. The recorded hosts for P. graminis are listed in

Table 1.

Table 1: Recorded plant hosts of Polymyxa graminis.

Common name Botanical name Source

Barley Hordeum vulgare Barr, 1979; Rush, 2003

Couch grass, Quackgrass Agropyron repens Barr, 1979

Millet Panicum miliaceum Rush, 2003

Rye Secale cereale Barr, 1979; Rush, 2003

Sorghum Sorghum vulgare Rush, 2003

Wheat Triticum aestivum, T. durum Barr, 1979; Rush, 2003

Polymyxa graminis belongs to the order Plasmodiophorales, family

Plasmodiophoraceae (Kanyuka et al., 2003; Ward et al., 2005). In total, ten genera

are recognised in the Plasmodiophorales, all of which are intracellular parasites of

higher plants or fungi (Ward and Adams, 1998): Polymyxa, Spongospora,

Plasmodiophora, Ligniera, Membranosorus, Octomyxa, Sorodiscus, Sorosphaera,

Tetramyxa and Woronina (Kanyuka et al., 2003). Several species of Polymyxa,

Spongospora and Plasmodiophora genera, including P. graminis, are of significant

agronomic importance (Kanyuka et al., 2003).

There are several distinctive characteristics common amongst plasmodiophorids.

They are obligate intracellular parasites (Kanyuka et al., 2003; Rush, 2003), replicate

through cruciform nuclear division (Rush, 2003), they form multinucleated plasmodia

(Kanyuka et al., 2003; Rush, 2003), and their zoospores have two, anterior whiplash

flagella of unequal length (Kanyuka et al., 2003; Rush, 2003) (fig. 16). All produce

environmentally resistant, long-lived resting spores (cysts) that are often clustered

together to form a sporosorus (cystosorus) (Braselton, 1995; Rush, 2003). The

production of zoospores and long-lived sporosori are significant features in the

epidemiology of the diseases caused by plasmodiophorids, and the viruses they

vector (Rush, 2003).

javascript:popup('citart1','b18','10.1046/j.1364-3703.2003.00177.x','bj13643703200300177x','')


22

Figure 16. Scanning electron micrograph of a single biflagellate Polymyxa graminis zoospore,

showing the classic plasmodiophorid anterior whiplash flagella of unequal length (Kanyuka et al., 2003).

The genus Polymyxa, has two recognised species, P. graminis and P. betae. Both

species are morphologically indistinguishable (Barr, 1979; Legreve et al., 2002; Ward

and Adams, 1998), but were separated due to host range (Barr, 1979; Legreve et al.,

2002; Rush, 2003), and differences in cyst walls and zoosporangial plasmodia (Barr,

1979). P. graminis parasitises mostly monocotyledons, in the family Poaceae,

whereas P. betae favours dicotyledons within the Chenopodiaceae and related plant

families, such as Amaranthaceae, Portulaceae and Caryophyllaceae (Legreve et al.,

2002; Ward and Adams, 1998; Ward et al., 2005). The cyst walls in P. graminis are

fused, whereas in P. betae they are separated by a cementing substance, and the

zoosporangial plasmodia of P. graminis have longer discharge tubes than those of P.

betae (Barr, 1979). In recent times, ribosomal DNA ITS sequences have separated

P. graminis from P. betae, with isolates of P. graminis exhibiting a wider diversity in

ITS sequence when compared to P. betae (Rush, 2003).

The diversity within ITS sequences of P. graminis isolates may be a reflection of their

differences in temperature and host range. Indian isolates have a significantly higher

temperature optimum (27-30oC) than isolates from Europe and Canada (15-20oC,

Rush, 2003). A reduction in temperature to 15oC disables the Indian isolates, making

them incapable of vectoring PCV, although at that temperature the virus can replicate

and cause symptoms of infection (Rush, 2003).

The diversity in P. graminis isolates is reflected in host range (Legreve et al., 2002),

with tropical and subtropical isolates exhibiting a continuum in host specificity, as

opposed to an absolute, discrete host range favoured by isolates from temperate

regions (Rush, 2003). P. graminis normally infects graminaceous species, but

isolates from India are capable of infecting the dicotyledon, peanut (Arachis

hypogaea) (Rush, 2003). From an epidemiological standpoint, this distinction is


23

important, as in vector-virus-host relationships, tropical isolates of P. graminis do not

need to complete their life-cycle within the virus susceptible plant (Rush, 2003). An

example is the P. graminis capable of transmitting PCV and IPCV. Both viruses

infect peanut, but peanut is a poor host for the fungal vector (Rush, 2003). Due to

the aggressiveness of this P. graminis isolate, its inability to complete its life cycle in

the peanut plant has little effect on disease severity of PCV and IPCV (Rush, 2003).

Legreve et al. (2002) proposed dividing P. graminis into five taxa based on the

combination of host range, temperature requirements, and genome characteristics,

as listed below.

1. P. graminis f. sp. temperata Legreve, Delfosse and Maraite

P. graminis on graminaceous species, particularly barley or wheat. P. graminis has

been detected once infecting sugar beet after heavy inoculation, favoured by

temperatures of 15-20oC, from temperate regions (Belgium, Canada, China, France,

Germany, UK), ITS1-5.8S gene-ITS2 sequence type Pg-I.

2. Polymyxa graminis f. sp. tepida Legreve, Delfosse and Maraite

P. graminis on graminaceous species, particularly barley, oat and wheat, favoured by

temperatures of 15-20oC, from temperate regions (Canada, UK) and ITS1-5.8S gene-

ITS2 sequence type Pg-II.

3. Polymyxa graminis f. sp. tropicalis Legreve, Delfosse and Maraite

P. graminis on graminaceous species, particularly sorghum, pearl millet and maize,

occasionally on wheat and barley, rarely on groundnut or sugar beet, favoured by

temperatures above 23oC, from tropical regions (India, Senegal), ITS1-5.8S gene-

ITS2 sequence type Pg-IIIa or b.

4. Polymyxa graminis f. sp. subtropicalis Legreve, Delfosse and Maraite

P. graminis on graminaceous species, particularly sorghum and pearl millet but also

wheat and barley, occasionally groundnut and sugar beet, favoured by temperatures

above 23oC, from sub-tropical regions (India, Pakistan) and ITS1-5.8S gene-ITS2

sequence type Pg-IVa or b.

5. Polymyxa graminis f. sp. colombiana Legreve, Delfosse and Maraite

P. graminis on rice from Colombia, ITS1-5.8S gene-ITS2 sequence type Pg-V.


24

Of the five P. graminis taxa, two are known to occur in temperate regions (Ward et

al., 2005). The biological significance of the two temperate ribotypes is not clear, but

the trend found by Ward et al. (2005) was that ribotype I was isolated from barley,

and ribotype II from wheat and other cereals.

3.3. Life cycle of Polymyxa graminis

The lifecycle of P. graminis is separated into two phases, 1) the production of

zoospores, and 2) the production of resting spores, with each phase initiated by the

attachment and penetration of the plant host epidermal or root hair cells by

zoospores (Jones, 2004; Kanyuka et al., 2003). The zoospores form within

zoosporangia located within root hairs or epidermal cells (Kanyuka et al., 2003), and

the zoosporangia (resting spores) burst open releasing the short-lived, motile

zoospores into the soil around the roots (Kanyuka et al., 2003) (fig. 17 and 18).

Figure 17. Diagram of the lifecycle of Polymyxa graminis (Kanyuka et al., 2003)

The zoospores, with the aid of their two flagella, swim short distances through the

soil moisture film (moisture between soil particles) to infect roots (Jones, 2004). The


25

fungus invades the root hairs of young susceptible plants in the autumn during

periods of high soil moisture (Bowden, 2005) (fig. 19). The zoospore contents

enlarge within the invaded host cell and undergo several cycles of synchronous

mitotic 'cruciform' nuclear divisions and reduction in nuclear size (Kanyuka et al.,

2003). The Polymyxa cytoplasm becomes an irregularly shaped multinucleate sac

(plasmodium), which develops into a zoosporangium (Ward et al., 2005), separated

from the host cytoplasm by a distinct cell wall (Kanyuka et al., 2003). Septa form

within the zoosporangium, dividing it into lobes that expand in volume followed by

several cycles of 'non-cruciform' mitotic nuclear divisions, and many form exit tubes

extending to the host cell wall (Kanyuka et al., 2003). The secondary zoospores are

cleaved apart, become rounded when they mature, and the septa between

zoosporangial segments disintegrate (Kanyuka et al., 2003), later developing into

resting spore clusters (Ward et al., 2005).

Figure 18. Life cycle of Polymyxa graminis: (a) mature sporangia with exit tubes passing through cells. Motile zoospores emerging formed from one sporangium; (b) biflagellate zoospores; (c) living amoeboid zoospores; 9d) amoeboid spores during period of active growth; (e) individual plasmodia (meronts); (f) same cell as d after the plasmodia have coalesced; (g) plasmodia just prior to cleavage into incipient cystosori; (h) cleavage of

plasmodia into polygonal resting spores (cystosori); (i) wheat cell completely filled with resting spores. (Adapted from Ledingham) (Integrated Pest Management Copyright © 2002

University of Illinios at Urbana-Champaign).

http://www.uiuc.edu/


26

Figure 19. Diagram of Polymyxa graminis zoospore encystment and penetration of root cells

(S: Stachel, R: Rohr: adhesium, N: nucleus, V: vacuole, L: lipid droplet) (Kanyuka et al., 2003).

The fungus forms dark clusters of resting spores in the cortical tissues within the

infected root (fig. 20 to 25). These spore balls are released into the soil when the

roots decay (Bowden, 2005), and the fungus survive as clusters of thick-walled

resting spores within the soil (Ward et al., 2005) for more than 15 years in the

absence of access to susceptible hosts (Jones, 2004). The cycle is restarted when

the resting spores germinate to each produce one biflagellate zoospore (Kanyuka et

al., 2003), which, in turn, will infect the root epidermal cells of a suitable plant host

(Ward et al., 2005), and behaving in the same manner as zoospores derived from

zoosporangia (Kanyuka et al., 2003).

Figure 20. Transmission electron micrograph of resting spores of Polymyxa graminis in root cell of Triticum monococcum (Source: http://oak.cats.ohiou.edu/~braselto/plasmos/poly.html)


27

Figure 21. Clusters of Polymyxa graminis resting spores inside root cells of wheat (X 205)

(Source: http://www.bsu.edu/classes/ruch/msa/barr.html).


28

Figure 22. Grass roots cleared of protoplasm to reveal the presence of clusters of small

resting spores of Polymyxa graminis. Top: low-power view of a root. Bottom: two clusters of resting spores in a single root cortical cell

(Source: http://helios.bto.ed.ac.uk/bto/microbes/chytrid.htm).

Figure 23. Images of Polymyxa graminis in roots. A, B and C: P. graminis zoosporangia

(zsp). Some zoosporangial exit tubes (et) are indicated and function for release of zoospores to the extracellular environment. D: P. graminis sporogenic plasmodium (spl) in root cells E and F: P. graminis sporosori (sp). Some individual resting spores (rs) are indicated. Bars =

100μm (© American Phytopathological Society 2004)


29

Figure 24. Resting spores of Polymyxa graminis in Sorghum arundinaceum roots

(Source: http://www.dpvweb.net/dpv/showfig.php?dpvno=235&figno=05

Figure 25. Resting spores of the fungus Polymyxa graminis, which delivers several viruses to

plants (Source: http://www.acad.swarthmore.edu/bio5/maldonado/risks.htm).


30

3.3. Polymyxa graminis fungal isolation

Polymyxa graminis is an obligate biotroph and can therefore only be maintained in

the roots of host plants (Kanyuka et al., 2003; Subr et al., 2002). As P. graminis can

not be grown in culture, all purified preparations could potentially be contaminated

with soil microorganisms and plant debris (Subr et al., 2002). It is generally difficult

to obtain good quality P. graminis DNA for molecular studies that is free from the

contaminating DNA of the host plant or other organisms (Ward et al. 1994),

especially when samples are collected from the field (Kanyuka et al., 2003).

Adams et al. (1986), Mutasa et al. (1993), Subr et al. (2002) and Ward et al. (1994)

have isolated P. graminis spores for molecular biological experiments by propagating

the fungus on host plant roots in semi-sterile sand cultures. Subr et al. (2002) used

subtractive hybridisation to further isolate P. graminis-specific DNA from plant roots.

However, even in experimental glasshouse conditions, P. graminis multiplication is

slow, taking 34 weeks to produce zoospores and approximately 23 months to

produce resting spores (Kanyuka et al., 2003).





31

3.4. Transmission of Wheat spindle streak mosaic

virus by Polymyxa graminis

WSSMV particles are thought to be present in viruliferous P. graminis resting spores

and zoospores (Kanyuka et al., 2003; Ward et al., 2005). Driskel et al. (2004)

produced direct evidence that WSSMV is internalised by P. graminis, as WSSMV

coat protein was detected in the resting spores using immunofluorescence (fig. 26).

Figure 26. Immunofluorescence labelling of Wheat spindle streak mosaic virus (WSSMV) coat protein (CP) in cross sections of barley roots. A, B, C and D: Transmitted light and

fluorescence images of Polymyxa graminis resting spores or sporosori in root cortical cells. A and C show sporosori that were labelled with WSSMV CP antiserum (B and D respectively). E: Resting spores, F: treated with buffer and secondary antiserum. Bar = 20μm. A, C and E:

Transmitted light images of resting spores or sporosori in root cortical cells. B and D: WSSMV CP was detected in resting spores by immunofluorescence labelling. Sections A and B, C and D, E and F are the same, but images were taken using either a transmitted light detector or a

Kr/Arg laser (Driskel et al., 2004 © American Phytopathological Society).


32

WSSMV cannot be removed from zoospores by washing, or inactivated by

application of antiserum. P. graminis resting spores remain viruliferous after

treatments with diluted NaOH and HCl (Kanyuka et al., 2003). The precise

mechanism of virus uptake and transfer is unknown (Rush, 2003), and it is also not

known whether WSSMV is able to replicate within P. graminis, although indirect

evidence suggests that it does not (Kanyuka et al., 2003). Zoospores released from

viruliferous P. graminis isolates grown in virus-resistant host plants no longer contain

WSSMV, or loose the ability to transmit the virus (Kanyuka et al., 2003).

Polymyxa graminis acquires WSSMV when it multiplies inside virus-infected plant

cells as the virus is incorporated in the resulting plasmodium (Rush, 2003). If this

plasmodium develops into a zoosporangium, the secondary zoospores will be

viruliferous, and if it develops into a sporosorus, the virus particles will survive inside

the resting spores for years (Rush, 2003). When the plant host cell dies and

deteriorates, the infected resting spores are released into the surrounding soil, and

upon germination, will release motile viruliferous primary zoospores in search of a

plant host (Rush, 2003) (fig. 17).

Motile P. graminis zoospores are released from resting spores or zoosporangia, and

infects the plant via root hairs or epidermal cells (Rush, 2003). If the zoospore is

viruliferous, virus particles are introduced into the plant cytoplasm soon after contents

of the zoospore are injected into the cell (Rush, 2003). The virus replicates and

initiates disease, becoming systemic, causing symptoms in leaves, and affecting

plant growth and yield (Rush, 2003; Ward et al., 2005).


33

3.5. Management strategies for Polymyxa graminis

Polymyxa graminis is a ubiquitous soil inhabitant wherever susceptible crops are

produced (Rush, 2003). Once the fungus is introduced to new areas of cereal

production, eradication is difficult as the dormant resting spores are able to persist for

many years in soil in the absence of a plant host (Jones, 2004). Therefore, strict

farm hygiene must be implemented to prevent spread, and techniques such as

minimum or zero tillage can be utilised to reduce soil movement via cultivation

(Jones, 2004).

This fungus survives in a variety of soil types that vary greatly in pH, texture, and

structure (Rush, 2003). For example, P. graminis can function as a vector in soil pH

values 4 to 8 with minimal effect on its ability to vector Rice stripe necrosis virus

(Rush, 2003). Infection of plant roots by P. graminis is favoured by high soil moisture

conditions, so well drained soils typically reduce infection (Rush, 2003). However,

heavy soils with poor structure and compaction layers, which slow drainage and

retard root growth, enhance root infection (Rush, 2003).

Due to the longevity and hardiness of P. graminis resting spores, host resistance to

the fungus by cereal cultivars would be a very desirable trait. Unfortunately, at

present there are no sources of natural P. graminis resistance. The European barley

cultivars tested so far have been highly susceptible to P. graminis and there were

some differences between wheat cultivars in their degree of susceptibility (Ward et

al., 2005).


34

4. Detection of Wheat spindle streak mosaic

virus

Please note, that the WSSMV detection system outlined in this manual is based on

the RT-PCR protocols published by Gitton et al. (1999) and Clover and Henry (1999),

and that these protocols have not been validated under Australian conditions.

4.1. Diagnostic Flow-chart

If POSITIVE: Sequence PCR product for

verification of WSSMV

infection

If NEGATIVE:

Stop

WSSMV not detected

RNA Extraction

Housekeeping

RT-PCR Primer pair (MDH-H968/MDH-C1163)

(Nassuth et al., 2000)

+

RT-PCR

Primer pair 2 (WSSMVF/Oligo-d(T)Not1) (Clover and Henry, 1999)

RT-PCR

Primer pair 1 (Fw4/Rw4)

(Gitton et al.,

1999)

Examine symptoms Tissue Sampling

-

Repeat RNA extraction,

and re-test with the Housekeeping

RT-PCR

+

-

+

-

Plant Sample

If WSSMV is detected, ELISA can be utilised to screen

large numbers of plant samples

Sap

Extraction

ELISA

(After WSSMV has been confirmed by

RT-PCR)


35

4.2. Sample collection

4.2.1. Plant

Record plant symptoms. Collect leaves and roots from plants suspected of WSSMV

infection. Wash roots clean of soil under fast running tap water. Store plant material

at 4oC or at –20oC (Gitton et al., 1999; Kingsnorth et al., 2003).

4.2.2. Soil

Collect soil samples in a “W” pattern across the field. Take a 1-cup sample of soil

every ten metres. Soil samples can be pooled. To increase the likelihood of

detecting virulent P. graminis, include soil samples collected from the wettest area of

the paddock.

To recover WSSMV from the soil samples, rear T. aestivum seedlings in the suspect-

virus-infected soil (Clover and Henry, 1999). Incubate two-week old seedlings of T.

aestivum in a slurry of 40 g of infected soil at 20oC for 10-14 days. After incubation,

transplant into sterile sand to give a 1:9 dilution of infected soil:sand. Grow the

plants for a further 3 weeks at 20oC, then decrease the temperature to 10oC. Record

plant symptoms. Take leaf samples monthly. Store plant material at 4oC or at –

20oC.

Currently, there are no soil extraction protocols for extracting WSSMV or P. graminis

directly from soil.


36

4.3. Total RNA extraction

Wear disposable gloves and a lab coat at all times. Have an autoclave bag ready to

dispose of all plant material, tips, tubes, gloves, and paper towel that has come into

contact with any suspect plant material. A footbath containing disinfectant located at

the doorway of the laboratory must be used when exiting the lab.

4.3.1. Equipment required

1. 2-20 L, 20-200 L, and 200-1000 L pipettes and sterile tips

2. Autoclave

3. Autoclave bags

4. Balance (at least 2 decimal places)

5. Disposable gloves

6. Microcentrifuge

7. Sterile microcentrifuge tubes

8. Paper towel

9. RNeasy® Plant Mini Kit (QiagenTM) (Kingsnorth et al., 2003)

10. Sharps container

11. Sterile scalpel blades and scalpel blade handle

12. Waterbath or heatblock set at 70oC

13. Weighboats

ALSO OR OR

1a. 1b. 1c.

Autoclaved mortar Qiagen Tissue Lyser Homex tissue macerater

and pestle (QiagenTM) Homex bags

Fume hood 2 ml snap-lock tubes (Bioreba AG / BioSys)

Sterile sand Liquid Nitrogen Plastic disposable

Stainless steel beads pasteur pipettes


37

4.3.2. Reagents required

1. MacKenzie buffer (MacKenzie et al., 1997)

Chemical Amount Final Concentration

Guanidine thiocyanate (CH5N3·CHNS) 23.64 g 4 M

3M Sodium acetate (C2H3NaO2) 3.33 ml 0.2 M

0.5M EDTA (C10H16N2O8) 2.5 ml 25 mM

PVP-40 (Polyvinylpyrrolidone) 1.25 g 2.5% (w/v)

Add sterile distilled water to final volume of 50 ml

Store at room temperature

Please note, fresh MacKenzie buffer should be prepared every 3-6 months

2. β-mercaptoethanol (C2H6OS)

3. 20% N-Lauroylsarcosine solution (w/v)

4. 100% Ethanol

4.3.3. Method

The RNA extraction method is based on that described by MacKenzie et al. (1997).

All steps are carried out at room temperature and is as follows:

1a. If using a mortar and pestle to homogenise samples:

1a-1. Determine the number of samples and label plastic tubes accordingly.

1a-2. Use new clean gloves and scalpel blades for each sample.

1a-3. Cut each new sample on fresh paper towel on the bench.

1a-4. Weigh out 400 mg of plant sample.

1a-5. Place sample in mortar.

1a-6. Add 1980 l of MacKenzie buffer.

1a-7. Add 20 l of β-mercaptoethanol in the fumehood.

1a-8. Homogenise in fume hood.

1a-9. Pipette 1.0 ml of the mixture into a labelled microcentrifuge tube (you

may need to cut the end of the pipette tip if the slurry is too thick).

1a-10. Continue to step 2.


38

1b. If using the QiagenTM Tissue Lyser:

1b-1. Determine the number of samples and label the 2 ml snap-lock tubes

accordingly.

1b-2. Use new clean gloves and scalpel blades for each sample.

1b-3. Cut each new sample on fresh paper towel on the bench.

1b-4. Weigh out 100 mg of plant sample and place sample in the

appropriate tube.

1b-5. Add 990 l of MacKenzie buffer.

1b-6. Add 10 l of β-mercaptoethanol in the fumehood.

1b-7. Close tubes.

1b-8. Place tubes in the Adaptor Set, in the QiagenTM Tissue Lyser. Grind

for 1 min at 30 Hz.

1b-9. Rotate tubes within the Adaptor Set, so that tubes in the centre are

moved to the outside.

1b-10. Continue to step 2.

1c. If using the Homex tissue macerater:

1c-1. Determine the number of samples and label plastic tubes accordingly.

1c-2. Use new clean gloves and scalpel blades for each sample.

1c-3. Cut each new sample on fresh paper towel on the bench.

1c-4. Weigh out 200 mg of plant sample.

1c-5. Place sample in Homex bag.

1c-6. Add 1980 l of MacKenzie buffer.

1c-7. Add 20 l of β-mercaptoethanol in the fumehood.

1c-8. Macerate tissue with the Homex.

1c-9. With a plastic disposable pasteur pipette, transfer 1.0 ml of the mixture

into a labelled microcentrifuge tube.

1c-10. Continue to step 2.

2. Carefully read the RNeasy Mini Handbook.

3. Add 100 l of 20% Sarkosyl to each tube and mix.

4. Incubate tubes at 70oC for 10 minutes.

5. Spin tubes in microcentrifuge for 1 minute at 13,000 rpm.

6. Continue with step 4 of the “RNeasy® Plant Mini Protocol for Isolation of Total

RNA from Plant Cells and Tissues and Filamentous Fungi” on page 75 of the

RNeasy Plant Mini Handbook and follow as per manufacturer's instructions.


39

4.4. Detection of Wheat spindle streak mosaic virus

in total RNA extracts using one step RT-PCR

For the reliable detection of WSSMV, total RNA extracts are subjected to three RT-

PCR tests, as outlined below, with primer sequences and annealing temperatures

listed in Table 2.

1. Primer pair Fw4 (F) / Rw4 (R). This primer pair amplifies a 457 bp region of the

RNA1 component of the WSSMV genome, from bases 730 to 1186 on the

reference WSSMV nucleotide sequence X73883. Please note that this primer

pair does not amplify isolates from Okalahoma or China, but that it will faintly

detect Barley yellow mosaic virus (Gitton et al., 1999).

2. Primer pair WSSMVF (F) / Oligo-d(T)Not1 (R). This primer pair amplifies a

region on the WSSMV coat protein gene on RNA1, generating a 982 bp fragment

(Clover and Henry, 1999).

3. House-keeping gene (Primer pair MDH-H968 (F) / MDH-C1163 (R). The MDH-

H968/MDH-C1163 primer pair are designed to amplify a 196 bp region of the

plant mRNA encoding malate dehydrogenase (MDH) gene (Nassuth et al., 2000).

This gene is highly conserved among plants and therefore RT-PCR amplification

of the MDH mRNA is used as an internal RT-PCR control to, a) determine the

quality of the RNA extract, and b) determine whether the RNA extract contains

inhibitors that will interfere with the activity of the reverse transcriptase and Taq

DNA polymerase enzymes. This RT-PCR is particularly important when

confirming the absence of WSSMV in the test sample.


1. 0-2 l, 2-20 l, 20-200 l, and 200-1000 l pipettes and sterile tips

2. 0.2 or 0.5 ml sterile PCR tubes

3. Bulb spinner or centrifuge


5. Freezer

6. Gel electrophoresis tanks, rigs and racks

7. DNA Molecular Weight markers

8. Ice

9. Leather gardening gloves


40

10. Sterile microcentrifuge tubes to store reagents

11. Microwave

12. Power pack

13. Thermocycler

14. UV transilluminator with camera

4.4.2. Reagents

1. Primers

For the detection of WSSMV use the three primers sets listed in Table 2. Each

primer is used as a stock solution at a concentration of 10 M.

Table 2: Primers required for the detection of Wheat spindle streak mosaic virus

Primer Sequence (5'-3') PCR Cycling Conditions

Fw41 AAG GAA ATA AAT ACC

GCC CCA G

1 cycle [48oC 30 min],


25 cycles [94oC 1 min, 56oC 1 min, 72oC 1.5

min],

1 cycle [72oC 10 min]

Rw41 TCA TAC CCG ACT CTT

CCA GCA C

WSSMVF2 CAG CAA CCA AAG TYR

CAG CAA C



10 cycles [94oC 30 s, 56oC 30 s, 68oC 1 min],

25 cycles [94oC 30 s, 56oC 30 s, 68oC 2 min],

1 cycle [68oC 7 min]

Oligo-d(T)No

t12

CAA TTC GCG GCC GCT

MDH- H9683 GCA TCT GTG GTT CTT

GCA GG

1 cycle [48 C 45 min],

1 cycle [94 C 2 min],

35 cycles [92 C 30 secs, 54oC 30 secs, 72 C

1 min],

1 cycle [72 C 5 mins]

MDH-C11633 CCT TTG AGT CCA CAA

GCC AA

1 (Gitton et al., 1999), 2 (Clover and Henry, 1999), 3 (Nassuth et al., 2000)

2. PCR Controls

1. Positive control - RNA extract from plant tissue infected with WSSMV.


41

- Alternatively a “plasmid control” that has the target WSSMV

sequence cloned into the plasmid

2. Negative plant control - RNA extract from uninfected plant tissue of the same

species as that used for the positive control.

3. Negative buffer control - an aliquot of the RT-PCR “Master Mix” without template.

4. The house keeping RT-PCR, using primers MDH-H968/MDH-C1163, reduces the

risk of false negative results. The generation of a band confirms the presence of

RNA in the extract, and that the RNA extract does not contain inhibitors. Failure

to produce an amplicon of expected size (196 bp) indicates that either dilution of

the RNA extract or re-extraction of RNA from the sample is required.

3. RT-PCR reagents

1. One-step RT-PCR kit (Invitrogen® SuperScriptTM One-Step RT-PCR with

Platinum® Taq, Catalogue No. 12574-026, is recommended)

2. Nuclease-free water

4. 5x TBE Buffer

Per 1 litre

Tris (C4H11NO3) 54 g

Boric acid (H3BO3) 27.5 g

0.5M EDTA ([CH2.N(CH2.COOH).CH2COONa]2.2H2O) pH 8.0 20 ml

Store at room temperature.

5. 1% Agarose gel with ethidium bromide

Use a 1% DNA grade agarose gel made with 0.5x TBE solution, and stained with

0.03 g/ml Ethidium bromide.

6. 1x TE Buffer

Per 100 ml

1 M Tris-HCl (pH 8.0) 1 ml

0.5 M EDTA 200 μl

Adjust pH to 8.0± 0.2. Store at room temperature.

7. 6x loading dye

Per 100 ml


42

1 x TE 10 ml

Glycerol (Sigma 200-289-5) 50 ml

Bromophenol blue (Sigma 263-653-2) trace (0.2%)

Store at room temperature.

4.4.3. One-step RT-PCR detection of Wheat spindle streak mosaic virus

This method is to be repeated for each set of the three primer pairs listed in Table 2.

Use one-step RT-PCR reagents as specified by the manufacturer. Some volumes

outlined below may vary depending on the buffer and enzyme concentrations

specified by the manufacturer. Ensure that the final volume of the RT-PCR reaction

is 25 µl by altering the volume of nuclease-free water accordingly.

1. Label sterile PCR tubes

2. Prepare "Master Mix" on ice in a sterile microcentrifuge tube.

The “Master Mix” usually contains buffer, forward and reverse primers, RT/Taq

and nuclease-free water.

Prepare the “Master Mix” according to the RT/Taq manufacturer’s

recommendations.

Ensure that the final volume for each reaction is 24 l.

Add 24 l of “Master Mix” to each PCR tube.

3. Add 1 l of each template (total RNA extract) to each corresponding PCR tube.

4. Cycle the tubes using the RT-PCR conditions listed in Table 2.

5. At completion of the RT-PCR, mix 10 l of each reaction with 2 l of 6x gel

loading dye, and load samples onto a 2% agarose gel with ethidium bromide.

6. Electrophorese in 0.5 x TBE at 100V for 45 minutes or until the bromophenol blue

front has migrated half way down the length of the gel.

7. Visualise and photograph gel on UV transilluminator.


43

4.5. DNA Sequencing of PCR Products


1. 0-2 l, 2-20 l, 20-200 l, and 200-1000 l pipettes and tips

2. 0.2 or 0.5 ml PCR tubes

3. 1.5 or 2 ml centrifuge tubes to store reagents


5. Freezer

6. Ice machine

7. Latex gloves

8. PC with Internet access

9. QIAQuick® PCR Purification Kit (QiagenTM)

10. Thermocycler

11. UV illuminator

4.5.2. Reagents

1. ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction Kits (Applied

Biosystems (www.appliedbiosystems.com))

2. Forward and Reverse primers (As per section 4.4.2).

3. Sterile dH2O.

4.5.3. Method

PCR products are cleaned using the QIAquick Spin kit (Qiagen) as per

manufacturer’s instructions. The purified PCR products are prepared for sequencing

with ABI Big Dye (Roche), as per the manufacturer’s instructions. Sequencing is out-

sourced to Monash University, Melbourne or similar provider. The DNA sequences

are compared against sequences on the GenBank database using the program

BlastN (Altschul et al., 1997), to confirm if the positive PCR product sequence is from

WSSMV.

http://www.appliedbiosystems.com/


44

4.6. Detection of Wheat spindle streak mosaic virus

with Enzyme-linked immunosorbent assay (ELISA)

4.6.1. Equipment

1. Wheat spindle streak mosaic virus coating antibody (Agdia; CAB 43001/0500)

(Store at 4oC)

2. Wheat spindle streak mosaic virus enzyme conjugated antibody (Agdia, ECA

43001/0500) (Store at 4oC)

3. Balance

4. Homex tissue macerater (Bioreba AG / BioSys)

5. Homex bags (Bioreba AG / BioSys)

6. Multiskan MS micro-titre plate reader (Labsystems)

7. Micro-titre plate (Nalgene)

8. Incubator

9. Fridge

10. 20-200 l pipette tips

11. Plastic sample cups

12. Plastic zip-lock bags

4.6.2. Reagents

Make all reagents with distilled water only.

1. Coating buffer

Per 1 litre

Sodium carbonate (anhydrous) 1.59 g

Sodium bicarbonate 2.93 g

Sodium azide 0.2 g

Adjust pH to 9.6. Store at 4oC.

2. ELISA extraction buffer

Per 1 litre

Sodium sulfite (anhydrous) 1.3 g

Polyvinylpyrrolidine (PVP) MW 24-40,000 MW 20.0g

Sodium azide 0.2 g


45

Powdered egg (chicken) albumin, Grade II 2.0 g

Tween-20 20.0 g


3. PBST buffer (Wash buffer)

Per 1 litre

Sodium chloride 8.0 g

Sodium phosphate, dibasic (anhydrous) 1.15 g

Potassium phosphate, monobasic (anhydrous) 0.2 g

Potassium chloride 0.2 g

Tween-20 0.5 g


4. ECI buffer

Per 1 litre

Bovine serum albumin (BSA) 2.0 g

Polyvinylpyrrolidine (PVP) MW 24-40,000 MW 20.0g

Sodium azide 0.2 g


5. Substrate visualising buffer

Dissolve 5 mg P-nitrophenyl phosphate in 5 ml substrate buffer (9.7 % (v/v)

Dethanolamine, pH 9.8)

4.6.3. Biological Reagents

Due to quarantine restrictions, there is no positive control available for WSSMV.

For a negative control use tissue from a non-infected plant of the same species as

that being tested.

4.6.4. Method

1. Dilute the concentrated WSSMV coating antibody into coating buffer at the

dilution recommended by the manufacturer. Make 1 ml per 8 wells. Use

immediately.

2. Add 100 μl of prepared coating buffer to each well. Incubate for 4 hours at 37oC.

(see Appendix 1 for layout with or without using the outer wells).


46

3. Homogenise plant tissue: for negative controls and each sample, homogenise 1

gram of leaf or root tissue 10 ml extraction buffer.

4. After incubation of coating antibody, empty wells into a sink. Fill wells to

overflowing with PBST. Quickly empty. Repeat 4 to 8 times. Final rinse with

distilled water.

5. After washing, hold the plate upside down and tap firmly on a folded paper towel

to dry the wells.

6. To each micro-titre plate load 100 μl each of negative and buffer controls in

duplicate wells.

7. Add 100 μl of each sample homogenate into duplicate wells. Ensure that you

have a record of sample location (well grid reference) for each sample (Appendix

1).

8. Incubate plate overnight at 4oC.

9. Ten minutes prior to washing make enzyme antibody conjugate. Dilute the

concentrated WSSMV enzyme antibody conjugate into ECI buffer at the dilution

recommended by the manufacturer. Make 1 ml per 8 wells. Mix thoroughly.

10. After incubation of plant sap extract, empty wells into a sink with a quick flipping

motion. Fill wells to overflowing with PBST buffer. Quickly empty. Repeat 4 to 8

times and include several brief distilled water washes.


to dry the wells.

12. Add 100 μl of prepared enzyme antibody conjugate per well.

13. Incubate plate at 37oC for 4 hours.

14. Wash plate three times with PBST buffer and three times with distilled water.


to dry the wells.

16. Add 100 μl of substrate visualising buffer to each well. Incubate at room

temperature for 30 to 60 minutes.

17. Measure the micro-titre plate with a plate reader (Labsystems) at an optical

density of 405 nm.

18. Samples are considered positive if the OD readings are 2.5 times the (corrected)

healthy control OD reading.


47

5. Detection of Polymyxa graminis

The P. graminis detection system outlined in this manual is based on the PCR

protocols published by Ward and Adams (1998) and Ward et al. (2005). The

protocols have not been validated for Australian conditions.

5.1. Diagnostic Flow-chart

If POSITIVE: Sequence PCR product for verification of P. graminis

infection

If NEGATIVE: Stop

P. graminis not detected

DNA Extraction

Housekeeping PCR

Primer pair (ITS1/ITS4)(Mello et al.,

1999)

+

Multiplex PCR Primer pairs (Pg.F1/Pg.R1

and Pg.F2/PgR2) (Ward et al., 2005)

PCR Primer pair 1

(Pgfw2/Pxrev7) (Ward and

Adams, 1998)

Examine symptoms Tissue Sampling

-

Repeat DNA extraction, and re-test with the

Housekeeping PCR

+

-

+

-

Plant Root Sample


48

5.2. Sample collection

5.2.1. Plant

Collect plants suspected of P. graminis infection and wash roots clean of soil under

fast running tap water. Allow to air dry, and store in a cool dry place (Barr, 1979).

5.2.2. Soil

Collect soil samples in a “W” pattern across the field. Take a 1-cup sample of soil

every ten metres. Soil samples can be pooled. To increase the likelihood of

detecting P. graminis, include soil samples collected from the wettest area of the

paddock.

To recover P. graminis from the soil samples, grow T. aestivum seedlings in the

suspect fungus-infected soil (Clover and Henry, 1999). Incubate two-week old

seedlings of T. aestivum in a slurry of 40 g of infected soil at 20oC for 10-14 days.

After incubation, transplant into sterile sand to give a 1:9 dilution of infected soil:sand.

Grow the plants for a further 3 weeks at 20oC, then decrease the temperature to

10oC. Grow the plants for a further 9-12 months. Check roots periodically for P.

graminis infection. Treat collected roots as per section 5.2.1.


49

5.3. Total DNA extraction


5. 2-20 l, 20-200 l, and 200-1000 l pipettes and sterile tips

6. Autoclave

7. Autoclave bags

8. Balance (at least 2 decimal places)


10. Microcentrifuge

11. Sterile microcentrifuge tubes

12. Paper towel

13. DNeasy® Plant Mini Kit (QiagenTM)

14. Sharps container

15. Sterile scalpel blades

16. Waterbath or heatblock

17. Weighboats

5.3.2. Method

Follow the protocol of the DNeasy® Plant Mini Kit (QiagenTM), as per the

manufacturer’s instructions.


50

5.4. Detection of Polymyxa graminis in total DNA

extracts using PCR.

For the reliable detection of P. graminis, total DNA extracts are subjected to three

PCR tests, as outlined below, with primer sequences and PCR protocols listed in

Table 2.

1. Primer pair Pgfwd2 (F) / Pxrev7 (R). This primer pair detects isolates of P.

graminis, with the exception of isolates originating from India, generating a band

of 280 bp (P. graminis type I), and 320 bp (type II) (Ward and Adams, 1998).

2. Multiplex with all Pg primer pairs: Pg.F1 and Pg.F2 (F) / Pg.R1 and Pg.R2

(R). The forward primers are in ITS1 region, and the reverse primers in the ITS2

region. Primer pair Pg.F1/Pg.R1 amplifies a 292 bp fragment only from P.

graminis ribotype I, and primer pair Pg.F2/Pg.R2 a 430 bp fragment only from

ribotype II (Ward et al., 2005).

3. House-keeping gene (Primer pair ITS1 (F) / ITS4 (R). The ITS1/ITS4 primer

pair is designed to amplify a fragment between 500-1000 bp from the nuclear

ribosomal ITS region (Mello et al., 1999). This region is highly conserved

amongst fungi. Consequently, PCR amplification of the ITS region is used as an

internal PCR control to a) determine the quality of the DNA extract, and b)

determine whether the DNA extract contains inhibitors that will interfere with the

activity of the Taq DNA polymerase enzyme. This PCR is particularly important

when confirming the absence of P. graminis in the test sample.


1. 0-2 L, 2-20 l, 20-200 l, and 200-1000 l pipettes and tips

2. 0.2 or 0.5 ml PCR tubes and 1.5 or 2 ml centrifuge tubes


4. Freezer

5. Gel tanks, rigs, racks and power pack

6. Ice machine

7. Latex gloves

8. Microwave

9. Thermocycler

10. UV transilluminator with camera


51

5.4.2. Reagents

1. Primers

For the detection of P. graminis, use the four primers sets listed in Table 2.

Each primer is used as a stock solution at a concentration of 10 M.

Table 2. Primers used in the detection of Polymyxa graminis.

Primer Sequence (5’-3’) PCR Protocol

Pgfwd21 GGA AGG ATC ATT AGC GTT GAA T 25 cycles [94oC 30 s, 56oC 1

min, 72oC 2 min] Pxrev71 GAG GCA TGC TTC CGA GGG CTC T

Pg.F12 AAC ATG TGG ATT GTG GGC TAT GTG 1 cycle 95°C 2 min

40 cycles [95°C 15 s, 60°C 20 s,

72°C 30 s]

1 cycle 72°C 5 min

Pg.F22 ATG TGG ATC GTC TCT GTT GCT GGA

Pg.R12 AAC TCC CAT TCT CCA CAA CGC AA

Pg.R22 CCT CAT CTG AGA TCT TGC CAA GT

ITS13 TCC GTA GGT GAA CCT GCG G 35 cycles [ 94oC 1 min, 50oC 1

min, 72oC 2 min]

1 cycle 72°C 5 min

ITS43 TCC TCC GCT TAT TGA TAT GC

1. Ward and Adams (1998), 2. Ward et al. (2005), 3. Mello et al. (1999).

2. PCR Controls

1. Positive control - DNA extract from plant tissue infected with P. graminis.

- Alternatively a “plasmid control” that has the target P.

graminis sequence cloned into the plasmid

2. Negative plant control - DNA extract from uninfected plant tissue.

3. Negative buffer control - an aliquot of the PCR “Master Mix” without template.

4. The house keeping PCR, using primers ITS1/ITS4 to reduce the risk of false

negative results. The generation of a band confirms the presence of DNA in the

extract, and that the DNA extract does not contain inhibitors. Failure to produce

an amplicon of expected size (500-1000 bp) indicates that either dilution of the

DNA extract or re-extraction of DNA from the sample is required.


52

3. PCR reagents

- Sterile dH2O

- 1 mM dNTPs

- 10 x concentration buffer

- 25 mM MgCl2

- Taq DNA polymerase

Use all reagents at concentrations recommended by the manufacturer of the Taq

DNA polymerase.

Please refer to section 4.4.2 for the following;

- 5x TBE buffer

- 1x TE solution

- 1% Agarose gel with ethidium bromide

- 6x loading dye

5.4.3. Method

8. Label sterile 0.2 ml centrifuge tubes

9. Prepare PCR “Master Mix” in sterile 1 ml microcentrifuge tube

10. Add 2 l sdH2O to the negative control tube, 2 l test template to each tube, and

DNA extracted from plants suspected to be infected with P. graminis into positive

control tube.

11. Place tubes in thermocycler, and use PCR conditions as listed in Table 2.

12. At completion of PCR, mix 10 l each PCR sample with 5 l loading dye

13. Load samples onto a 1% agarose gel containing ethidium bromide

14. Electrophorese in 0.5 X TBE at 100V

15. Visualise and photograph gel on UV transilluminator.

5.5. DNA Sequencing of PCR Products

As per section 4.5.


53

Acknowledgments

Plant Health Australia, for providing the funding for this project as part of their

National Diagnostic Protocols Initiative.

The Department of Primary Industries, Victoria, for allocating resources to this

project.

Dr Joanne Luck and Dr Fiona Constable for giving their time to review this document.


54

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60

Mutasa ES, Chwarszczynska DM, Adams MJ, Ward E and Asher MJC (1995)

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61

Appendix 1 – ELISA sample plans

1-45 Samples

1 2 3 4 5 6 7 8 9 10 11 12

A ● 2 6 10 14 18 22 26 30 34 38 42

B ● 2 6 10 14 18 22 26 30 34 38 42

C ● 3 7 11 15 19 23 27 31 35 39 43

D ● 3 7 11 15 19 23 27 31 35 39 43

E ● 4 8 12 16 20 24 28 32 36 40 44

F ● 4 8 12 16 20 24 28 32 36 40 44

G 1 5 9 13 17 21 25 29 33 37 41 45

H 1 5 9 13 17 21 25 29 33 37 41 45

● Positive control ● Buffer control ● Healthy control

1- 27 Samples

1 2 3 4 5 6 7 8 9 10 11 12

A - - - - - - - - - - - -

B - ● 1 4 7 10 13 16 19 22 25 -

C - ● 1 4 7 10 13 16 19 22 25 -

D - ● 2 5 8 11 14 17 20 23 26 -

E - ● 2 5 8 11 14 17 20 23 26 -

F - ● 3 6 9 12 15 18 21 24 27 -

G - ● 3 6 9 12 15 18 21 24 27 -

H - - - - - - - - - - - -

● Positive control ● Buffer control ● Healthy control

Documents

Wheat spindle streak mosaic virus DP - Plant Health Australia