Cereal chlorotic mottle virus
R. S. Greber
Department of Primary Industries, Indooroopilly, 4068, Australia
Described by Greber (1977
Greber & Gowanlock (1979)
A virus with bacilliform particles c. 65 x 240 nm containing RNA, protein
and, probably, lipid. Transmitted in a persistent manner by cicadellid leafhoppers but
not by inoculation of sap. Infects plants only in the family Gramineae; natural infection
occurs in maize and weed grasses and less frequently in temperate cereals. Reported only
from eastern Australia.
In maize, the virus causes fine chlorotic striations in the leaves (Fig.1
variable stunting and tassel sterility
); epiphytotics occur
sporadically, occasionally reaching 60% incidence but usually 0-10%. Naturally
infected temperate cereals, e.g.
barley (Hordeum vulgare; Fig.2
and wheat (Triticum aestivum; Fig.3
) also show striations and stunting but
epiphytotics do not occur in winter-grown crops. Species in several grass genera
provide a natural reservoir of virus (Greber, 1981
Reported only from Eastern Australia (latitude 19°-29°S) but range may be wider.
Host Range and Symptomatology
Known plant hosts are all in the family Gramineae, scattered through the
sub-families, but with fewer hosts in the Andropogonoidae. The following species are
hosts: Avena sativa, Bromus unioloides, Digitaria ciliaris, Dinebra retroflexa,
Echinochloa colona, Eleusine indica, Eragrostis cilianensis, Hordeum vulgare,
Leptochloa filiformis, Secale cereale, Setaria italica, Setaria verticillata,
x Secale cereale
), T. aestivum,
T. monococcum, T. tauschii, Urochloa panicoides, Zea mays.
Some of these species
(e.g. Dinebra retroflexa
spp.) are more susceptible than
others. No marked genotype differences have been noted within species except in
maize (Greber, 1981
This virus is not transmitted by inoculation of sap, only by the leafhopper
vectors; young seedlings are the most readily infected. Symptoms in most hosts consist
of various types of chlorotic striation with or without stunting. Striations tend to
decrease in intensity in chronic infections. Some degree of sterility is usually
caused, particularly in the male inflorescence in maize.
- Zea mays (maize). Some lines and hybrids are highly resistant
others are highly susceptible and these can be used as differential hosts. Susceptible
lines (e.g. B37, H84) produce fine short chlorotic striations (Fig.1) along
the veins over the leaf lamina, leaf sheath and ear husks. Symptoms appear
c. 14 days after leafhopper inoculation at 25°C.
- Triticum spp. (wheat) and x Triticosecale. First symptoms are
prominent chlorotic striations (Fig.3) which take up much of the area of the
first few leaves emerging after inoculation. Striations in later leaves are
usually finer and less prominent.
- Dinebra retroflexa. Readily infected and shows persisting chlorotic striations.
- Leptochloa filiformis. Severe chlorotic striations which turn to rusty
brown necrosis. The severe leaf symptoms are accompanied by deformation of the inflorescence.
- Sorghum bicolor (sorghum) and Oryza sativa (rice) are not infected.
- Maize is the best source of virus for purification. Cultures can be maintained
in Dinebra retroflexa. Vectors can be reared free of the virus on Chioris
gayana (for Nesoclutha pallida) and Sorghum bicolor (for Cicadulina bimaculata).
- Dinebra retroflexa is a good test plant in leafhopper transmission experiments.
No evidence available of strain differentiation.
Transmission by Vectors
Efficiently transmitted by at least two species of cicadellid leafhopper, Nesoclutha pallida
) and Cicadulina bimaculata
The virus was transmitted efficiently by leafhoppers that were injected with
a preparation of purified virus but not by insects that fed on the same preparation
through a membrane (R. Greber, unpublished data). For N. pallida,
minimum acquisition access period was 4 h (R. Greber, unpublished data) and the
latent period was 12-27 days for insects that acquired the virus from plants
or 8-10 days for insects injected with purified virus preparations (R. Greber,
unpublished data). The virus usually persists in the vector for life but some insects
cease to transmit up to 4 weeks before death
). The virus is acquired
both by adults and by nymphs and is transmitted through the moult
The virus infects cells of brain and salivary glands of both vector species
and apparently multiplies in the vector (Greber & Gowanlock, 1979
unpublished data). The vectors have a wide feeding range among species of Gramineae
but acquisition from and transmission to some host species is effected only by
though N. pallida
injected with infective sap from these
species will transmit to other species (Greber, 1981
Transmission through Seed
Not transmitted through seed of maize or D. retroflexa.
Weakly to moderately immunogenic. Antisera prepared by intramuscular injection of
purified virus had titres of 1/32 to 1/128 in gel-diffusion tests. Two, sometimes
three, precipitin lines are formed (Fig.10
) and the relative position and intensity
of these lines varies in different tests. Addition of 1% Triton X-100 to the antigen
well usually enhances the reaction. Virus also reacts in ring-precipitin tests, and
the gel-diffusion reaction with infective sap is useful in diagnosis. All major
antigens are sedimented from fresh infective sap (100,000 g
, 110 min)
but one, presumably the G-protein, is released as a soluble antigen after treatment
with 1% Triton X-100 (Fig.10
) or by degradation of stored preparations.
Particle morphology and virus/vector relations place the virus in the
plant rhabdovirus group
). It is classified within Subgroup II
because of the substantial association of the particles with the nuclei of
infected plant and insect cells
(Greber & Gowanlock, 1979
). The rhabdoviruses
infecting Gramineae are transmitted either by delphacid or cicadellid leafhoppers
but never by vectors of both types. This allows differentiation on both the type
of vector and association of the particles with the nucleus. The virus resembles
wheat striate mosaic virus in its cellular distribution and site of maturation and
in having a cicadellid vector; however, no serological relationship has been
demonstrated (Greber & Gowanlock, 1979
R. G. Timian, personal communication).
No serological reaction was obtained with antiserum to maize mosaic virus
(Greber & Gowanlock, 1979
a virus which also differs in having a delphacid vector
and in not being reported to infect festucoid grasses.
Barley yellow striate mosaic virus
northern cereal mosaic virus
are delphacid-transmitted and have
characteristics of Subgroup I (Peters, 1981
Stability in Sap
Virus in Dinebra retroflexa
sap, assayed by injection into
had a thermal inactivation point (10 min) between 45 and 50°C.
Purified virus, assayed in the same way was infective after 7 days at 4°C
(R. Greber, unpublished data).
A modification of the method of Jackson & Christie (1977)
Tissue (40-80 g) with good symptoms is checked for high particle concentration by
negative stain electron microscopy and then ground in a cold mortar, squeezed
through cloth and re-extracted using 0.1 M Tris-HCl buffer, pH 8.4, containing
0.04 M sodium sulphite. The metal salts used by Jackson & Christie (1977)
omitted. After a low speed centrifugation and adjusting to pH 7.4 the extract is
centrifuged in tubes containing solutions of 300 mg/ml sucrose layered over 600
mg/ml sucrose; the virus is collected from the interface and filtered through
Celite (10 g in a 9 cm funnel). The filtrate and washings are then passed through
two successive gradients of 100-400 mg/ml sucrose and 300-600 mg/ml sucrose and
pelleted in the manner described by Jackson & Christie (1977)
preparations have a milky appearance and gradient bands are very easily located.
Most particles from purified preparations appear relatively intact when fixed
with glutaraldehyde before electron microscope examination.
Properties of Particles
The particles sediment as a single band with a buoyant density in sucrose
solution of 1.165 g/cm3
. The particle mass (from density and dimensions)
is about 5.7 x 108
daltons (R. Greber, unpublished data). Light-scattering
is substantial, but the uncorrected A260
ratio is 1.17.
There is a gradual increase of uncorrected absorbance from A280
Normally bacilliform, 65 nm x 240 nm in thin section and 75 nm x 250 nm when fixed
in glutaraldehyde before negative staining (Fig.5
, top particle). Unfixed particles
disrupt in phosphotungstate negative stain to yield bullet-shaped structures, the
larger particles measuring 75 nm x 215 nm (Fig.5
, second particle). A prominent
net-like surface pattern (Fig.6
) is evident both in negative stain and in tangential
thin sections (Greber, 1979
Greber & Gowanlock, 1979
). This pattern correlates
with the number of projections seen around the perimeter (52 in longitudinal sections
and 17 in cross sections; Fig.7
). The helix, which is infrequently penetrated by
negative stain (Fig.5
, third particle), has c.
48 turns, approximately two
turns for each row of projections. In cross section (Fig.7
) the densely stained,
spiked outer zone is clearly separated from an inner stained circle; the central
core (35 nm) is largely unstained, with no central dot. The envelope projections
and net-pattern continue round the hemispherical ends, but at one end of each
particle (at the point of detachment from the nuclear membrane) the inner structure
is less densely stained within the hemisphere (Fig.5
, bottom particle).
Particle CompositionNucleic acid:
RNA (R. Greber, unpublished data), probably single-stranded
negative sense, as in other rhabdoviruses
), and arranged in a helix
5.5 µm in total length.
Protein: Five polypeptides are resolved by electrophoresis in
polyacrylamide/SDS gels (J. Dale & R. Greber, unpublished data) with M. Wt.
(x 10-3) of 77.5, 52.5, 46.5, 32 and 29; these probably correspond to
proteins G, N, NS, M1 and M2 of other rhabdoviruses
Relations with Cells and Tissues
Virus particles are present in most tissues of infected maize plants and in
brain and salivary gland tissue of infective leafhoppers. The particles appear to
receive their outer envelope at the inner nuclear membrane and are frequently
arranged as a palisade of bullet-shaped structures projecting into the perinuclear
space; they accumulate as bacilliform particles within the perinuclear space and
in cytoplasmic vesicles (Fig.4
). In insect tissue, particles may also be embedded
in granular viroplasm-like structures, within and around the nucleus. In some plant
and vector cells, large accumulations of particles may form and may appear to be
scattered throughout the nucleus. Unenveloped particles have not been observed,
but isolated complete particles are often found surrounded by individual vesicle
The main economic host is maize. Cereal chlorotic mottle virus is
cicadellid-transmitted, unlike maize mosaic virus
), a member of
maize sterile stunt virus (Greber, 1982
a member of plant rhabdovirus Subgroup I, both of which have
delphacid planthopper vectors. Particles of maize sterile stunt virus are
readily distinguished from those of cereal chlorotic mottle virus in negatively
stained leaf extracts (Greber, 1979
), because they lack the prominent surface
pattern and are longer and narrower (Fig.6
), and the stain penetration of the
core is only half as wide.
Cereal chlorotic mottle virus resembles
American wheat striate mosaic virus
(Sinha & Behki, 1972;
Jackson, Milbrath & Jedlinski, 1981) in several
respects. Particles of both viruses accumulate mainly in the perinuclear space,
both have cicadellid leafhopper vectors and the host range has a substantial
coincidence. However, no serological relationship has been detected between
the two viruses, and wheat striate mosaic virus differs from cereal chlorotic
mottle virus in having a higher buoyant density in sucrose (1.22 g/cm3:
Sinha, Harwalker & Behki, 1976),
in having only one M-protein
(Trefzer-Stevens & Lee, 1977),
in not showing a prominent net-like surface pattern in negative
stain, and in yielding a large amount of soluble antigen
(Thottappily & Sinha, 1973).
- Greber, Newsl. Aust. Pl. Path. Soc. 6: 17, 1977.
- Greber, Aust. J. agric. Res. 30: 433, 1979.
- Greber, Australas. Pl. Path. 10: 29, 1981.
- Greber, Aust. J. agric. Res. 33: 13, 1982.
- Greber & Gowanlock, Aust. J. biol. Sci. 32: 399, 1979.
- Herold, CMI/AAB Descr. Pl. Viruses 94, 4 pp., 1972.
- Jackson & Christie, Virology 77: 344, 1977.
- Jackson, Milbrath & Jedlinski, in Virus and Viruslike Diseases of Maize in the United States, pp. 51-76, ed. D. Gordon, J. Knoke & G. Scott, Ohio Agric. Res. Dev. Cent., Wooster, Ohio, 210 pp., 1981.
- Matthews, Intervirology 12: 132, 1979.
- Peters, CMI/AAB Descr. Pl. Viruses 244, 6 pp., 1981.
- Sinha & Behki, CMI/AAB Descr. Pl. Viruses 99, 4 pp., 1972.
- Sinha, Harwalkar & Behki, Phytopath. Z. 87: 314, 1976.
- Thottappilly & Sinha, Virology 53: 312, 1973.
- Trefzer-Stevens & Lee, Virology 78: 144, 1977.
Chlorotic striations caused by natural infection in maize.
Striation symptoms in naturally infected barley.
(From top): symptoms in wheat, Setaria italica and Eleusine coracana.
Thin section of infected maize cell showing perinuclear distribution of
virus particles (V). Bar represents 1 µm.
Virus particles (from top): bacilliform, glutaraldehyde-fixed in
negative stain; bullet-shaped, unfixed, in negative stain; helix penetration;
longitudinal thin section showing projections and one end less densely stained
(fixed with glutaraldehyde then osmium tetroxide, embedded in Spurrs medium,
stained with uranyl acetate and lead citrate). Bar represents 100 nm.
Prominent surface pattern and wider core penetration of cereal chlorotic
mottle virus (three upper particles) compared in negative stain to maize sterile
stunt virus (lower particle).
Transverse thin section of virus particles, showing protrusions.
Bar represents 100 nm.
Thin section of infected salivary gland cell from Cicadulina bimaculata.
Virus particles (V); nuclear membrane (NM). Bar represents 1 µm.
Thin section of infected brain cell from Nesoclutha pallida.
Virus particles (V); nuclear membrane (NM). Bar represents 500 nm.
Gel-diffusion serological reactions with cereal chlorotic mottle virus
antiserum (As). Top-purified virus (V), purified virus treated with Triton X-100 (VX),
and infective sap (IX). Below-separation of antigens
(p and s) by detergent treatment and differential centrifugation. Infective sap (IS).
All other antigen wells filled with preparations treated with 1% Triton X-100: pellet
100,000 g, 110 min (P); supernatant fluid 100,000 g,
110 min (5); supernatant fluid 100,000 g, 20 min (S'); infective sap (IX).