Seed Health and Vigor

# Health

The health of chickpea and lentil seed greatly impacts the quality of
the crop stand and yield. Healthy seed is characterized by a high
germination rate, is whole (free of cracks or other damage), free from
foreign matter including weed seed and has no or limited seed-borne
pathogens. The health of seed often depends on the moisture the crop
receives at or near harvest. Wet conditions at or near harvest can
result in seed sprouting, greater mechanical damage to seed during
harvest and favor the development of seed-borne pathogens. Early and dry
harvests provide the greatest opportunity for healthy seed as long as
the seed is not harvested immaturely which can lead to a decrease in
germination rate as the seed ages.

Planting seed that is pathogen-free is important in preventing the
introduction of pathogens into new production areas and preventing the
build up of pathogens in the soil. Often growers will adjust the seeding
rate based on the seed germination rate. If a low germination rate is
due to disease, the grower is increasing the risk of introducing
inoculum into the field. Often diseased seed is shrunken or discolored,
so seed cleaners and color sorters can be used to sort out diseased or
damaged seed resulting in better germination rates and a healthier crop
stand. Most seed-borne pathogens of chickpeas and lentils can be managed
with a biological or chemical seed treatment.

The most common seed-borne pathogens of chickpea are *Ascochyta rabiei*
and *Botrytis cinerea*. Other seed-borne pathogens include *Sclerotinia
sclerotiorum*, *Rhizoctonia solani*, and *Fusarium* spp. In Canada,
routine seed tests for chickpea are conducted to determine the percent
presence of *Ascochyta*, *Sclerotinia* + *Botrytis*, and *Fusarium* spp.
on seed. Seed with greater than 0.3% *Ascochyta* is considered to be
high risk. *Botrytis*-infected seed is usually shrunken and discolored
and seed sorters can be used to separate diseased seed from healthy seed
due to sieve size and seed color. Up to 10% infection of both
*Sclerotinia* and *Botrytis* combined is tolerable but may result in
significant seedling blight and a seed treatment is highly recommended.
Some labs will inform growers if the level of *Fusarium* spp. on the
seed exceeds 5%. Combined percentages of *Sclerotinia*, *Botrytis*, and
*Fusarium* on seed that exceeds 10% is considered high risk seed and a
seed treatment is highly recommended. Chemical seed treatments are
available to help manage contaminated and infected seed.

In a survey conducted on lentil seed from the Palouse region of eastern
Washington and Northern Idaho, seed-borne pathogens identified were
*Ascochyta lentis*, *Botrytis cinerea*, *Fusarium acuminatum*, *F.
avenaceum*, *Macrophomina phaseolina*, *Phoma medicaginis* var.
*pinodella*, *Rhizoctonia solani*, and *Sclerotinia sclerotiorum*. *B.
cinerea* and *P. m. pinodella* were the most common in the survey. In
surveys from Canada, the main seed-borne pathogens identified on lentil
were also *Ascochyta* *lentis* and *Botrytis cinerea*. Other pathogens
that contributed to lentil seed problems were *Sclerotinia
sclerotiorum*, *Rhizoctonia solani* and *Fusarium* spp. In Canada,
contaminated lentil seed exceeding 10% *Ascochyta* is considered high
risk and should not be planted. The same seed standards for
*Sclerotinia*, *Botrytis*, and *Fusarium* spp. previously described for
chickpeas in Canada are also the same for lentils.

*Pea seedborne mosaic* virus is the only major seed-borne virus that is
transmitted in chickpea and lentil. Planting clean pea seed from
certified clean seed lots is the major means of managing this pathogen.

# Vigor

Seed vigor can be described as the ability of a seed to consistently,
rapidly and uniformly germinate at a high rate even under adverse
weather conditions. Often after planting, cool weather associated with
rainfall and cool soil temperatures can negatively impact the
germination and growth of seed. Weak seedlings are more prone to soil-
and seed-borne pathogens. Several laboratory tests that include
physical, physiological and biochemical tests have been used to
determine seed vigor. Vigor tests in lentils assessing the speed of
germination, seedling weight and electrical conductivity of seed
leachates have been evaluated but it was determined that the only
laboratory test in lentils that proved to be effective in evaluating
field emergence was the standard germination test.

Several factors such as herbicide use, release of seed leachates, and
resistance to soilborne pathogens have been associated with seed vigor
of lentils and chickpeas. Pre-harvest applications of glyphosate on
chickpeas can caused chemical-damage to seed, which upon planting have
poor root development which stresses the plants and makes them more
susceptible to soilborne pathogens. Seed leachates released from a seed
at planting have been shown to impact the activity of seed rot and
damping-off pathogens. Seed leachate levels are determined by the area
of dead tissue on the cotyledons due to ageing and imbibition cracks in
the seed coat. The greater the amount of dead tissue and imbibition
cracks, the greater the seed leachates. The higher the leachate level,
the greater the fungal activity and the higher the risk of infection by
soilborne pathogens. Therefore lentil and chickpea cultivars with low
leachate release are desired to improve the likelihood of vigorous
growth. In addition, disease resistance to soilborne diseases in
chickpea can improve vigor. Chickpeas are divided into two main types,
white-seeded Kabuli and dark-seeded Desi types. Dark-seeded Desi types
tend to be more resistant to damping-off caused by *Pythium ultimum*
than the Kabuli type.

> **Selected References**

Kaiser, W. J. 1992. Fungi associated with the seeds of commercial
lentils from the U. S. Pacific Northwest. Plant Disease 76:605-610.

Makkawi, M., El Balla, M., Bishaw, Z., and Van Gastel, A. J. G. 1999.
The relationship between seed vigour tests and field emergence in lentil
(*Lens culinaris* Medikus). Seed Science and Technology 27:657-668.

Matthews, S., Powell, A. A., and Spaeth, S. C. 1988. Seedling vigour and
susceptibility to disease and pests. Pages 619-625 in: World Crops: Cool
Season Food Legumes. R. J. Summerfield, ed. Kluwer Academic Publishers,
Dordrecht, The Netherlands.

Saskatchewan Agriculture and Food Agriculture Knowledge Centre, Regina,
Saskatchewan, Canada. 2007. Guidelines for Seed-borne Diseases of Pulse
Crops.

(Prepared by L. D. Porter)

**Seed Pathology**

**Chickpea**

Many fungi, bacteria and viruses are seed-borne in chickpea. Some of
them are important pathogens of the chickpea crop while others are weak
pathogens or saprophytes and usually have little or no effect on seed
viability. However, high infestation of seeds by pathogenic and
saprophytic fungi may adversely affect quality, vigor, and longevity of
seeds. These organisms are carried internally or externally on the seed
which may become a major inoculum source for development of epidemics
under field conditions. Infected seed has aided in the local and
international spread and survival of many of these pathogens. Different
countries have imposed quarantine restrictions on the movement of
chickpea seed to prevent the entry and spread of some seed-borne
pathogens. Movement of infected seeds may lead to the introduction of
new races or virulent pathotypes of some pathogens, or to the
distribution of compatible mating types which could lead to the
development of the teleomorph in nature. Storage of chickpea seeds in
germplasm repositories not only prolongs the longevity of the seed, but
also of many of the associated seed-borne pathogens.

**Symptoms and Causal Organisms**

Infested or infected chickpea seeds may be symptomless or exhibit
different kinds of symptoms ranging from seed discoloration and
deformation to small necrotic lesions and seed rot. Lesions on chickpea
seeds are more visible on white seeded (Kabuli) types than on dark
seeded (Desi) types.

***Major chickpea pathogens***. The most important seed-borne pathogens
of chickpea are *Ascochyta rabiei* (teleomorph: *Didymella rabiei*),
*Fusarium oxysporum* f. sp. *ciceris*, and *Botrytis cinerea*.

The Ascochyta blight pathogen infects and causes lesions on all aerial
parts of the chickpea plant. Pod infection (Fig. 1) often results in
infected seeds which may be symptomless or develop dark brown lesions of
different sizes and shapes (Fig. 2). Pycnidia (Fig. 3) may develop in
these lesions.

The Fusarium wilt pathogen infects the plant through the roots,
colonizes the xylem tissues causing a vascular wilt, and may be
transmitted in seed. Infected seeds are usually smaller, deformed and
discolored although sometimes infected seeds can be symptomless. Visible
signs of the pathogen are not observed on the seeds but the fungus forms
chlamydospore-like structures in the hilum region of the seed.

The Botrytis grey mold pathogen infects all aerial parts of the plant,
including pods and seeds. Infected pods may be empty or contain seeds
that are small, shriveled, and sometimes covered with a grayish white
mycelium. The fungus isolated from infected seeds shows a great
variability in colony morphology, sporulation and sclerotial formation
(Fig.4).

***Minor chickpea pathogens*.** Different fungi, bacteria and viruses
pathogenic to chickpea have been isolated from chickpea seeds. The fungi
include *Alternaria alternata*, *Colletotrichum dematium*, *Macrophomina
phaseolina*, *Operculella padwickii*, *Phoma medicaginis* var.
*pinodella*, *Stemphylium sarciniforme*, and *Verticillium dahliae*. One
bacterial species, *Xanthomonas campestris*, has been associated with
chickpea seeds, causing pre- and post-emergence damping-off and wilting
of older plants. Five plant viruses are reported to be seed-borne in
chickpea. Infected seeds can be smaller in size, deformed and discolored
or they may be symptomless. These viruses are *Alfalfa mosaic virus,
Broad bean mottle virus, Cucumber mosaic virus, Pea seed-borne mosaic
virus,* and *Tobacco streak virus.*

***Saprophytic fungi*.** A great number of fungi have been isolated from
chickpea seeds. Many of them occur on the seed surface but at times some
are found internally. Even though these fungi are not pathogens of the
chickpea crop, they can often adversely affect seed quality and
germination. Among the fungi found associated with the deterioration of
chickpea seed are species of *Alternaria*, *Aspergillus*,
*Cladosporium*, *Curvularia*, *Fusarium*, *Helminthosporium*, *Mucor*,
*Penicillium*, *Rhizoctonia*, *Rhizopus*, and *Stemphylium*.

**Detection**

Various techniques have been used to detect seed-borne microorganisms in
chickpea. The blotter test had been widely used for detection of *A.
rabiei* and other seed-borne fungi. However, for slow growing fungi,
such as *A. rabiei*, seeds need to be surface disinfested to avoid being
overgrown by faster growing saprophytic microorganisms which can
interfere in detection of slower growing ones. Additionally, the blotter
test is not usually able to detect small amounts of inoculum in seeds.
Plating entire seeds or pieces of seed on different agar media, mainly
potato dextrose agar (PDA), reduces the inconvenience of the blotter
test, although it requires surface disinfestation of the seed to avoid
fast growing microorganisms. On agar media many pathogenic fungi can be
identified by their colony growth and spore morphology. For example, *A.
rabiei* forms a typical colony on PDA with abundant production of
pycnidia and conidia (Fig. 5). A similar fungus, *P. medicaginis* var.
*pinodella*, can be differentiated by colony growth (Fig. 6) and
production of chlamydospores (Fig. 7). Detection of viruses in chickpea
seed is done by serological and/or molecular techniques, such as ELISA
and PCR tests, or by sap inoculation on different test plants.

**Seed to seedling transmission**

Infected seeds may produce infected seedlings which can serve as a
source of primary inoculum for disease epidemics to develop in chickpea
crops. Systemic pathogens, such as *F. oxysporum* f. sp. *ciceris* and
viruses, spread directly from the seed to the developing seedling during
the germination process. However, for other pathogens, like *A. rabiei*,
contact between lesions on the seed coat or cotyledons and the emerging
plant is necessary for transmission of the pathogen. Symptoms on
seedlings infected from seeds may appear in a short period of time or
they may be latent. In the case of *A. rabiei*, infected seedlings show
black elongated necrotic lesions on the epicotyls (Fig. 8) in 7-12 days
after germination. Within these lesions, the fungus sporulates profusely
producing pycnidia and conidia which can be disseminated by splashing
rain.

**Control**

Various methods have been used to control transmission of different
chickpea pathogens in seed. Much of the research has been done with *A.
rabiei*. To effectively control the seed-borne phase of the Ascochyta
blight pathogen, an integrated control program is required. This
consists of using a combination of practices, such as production of seed
in arid environments, crop rotation, field sanitation, chemical seed
treatments, and field inspections to produce Ascochyta blight-free seed.
These measures would also be effective in controlling other pathogens
that are transmitted in chickpea seed.

Different fungicides have been tested on chickpea seed to control the
major and minor pathogens, including benomyl, Calixin-M (tridemorph +
maneb), captan, carbendazim, Dithane M-45 (mancozeb), maneb,
propiconazole, thiabendazole, and thiram. Some of these fungicides are
also effective in controlling saprophytic fungi that infest chickpea
seed.

Disease development and spread of many chickpea pathogens are favored by
wet weather. Surface-contaminated or internally infected seed is one of
most important methods of disseminating and perpetuating these diseases.
A very effective method of controlling different chickpea diseases is to
produce pathogen-free seed for planting purposes. Dry, warm weather
impedes disease development and spread of most chickpea foliar
pathogens. When and where possible, seed production fields should be
located in arid areas where little or no rainfall occurs during the
flowering and fruiting stages, or at harvest. If irrigation is required
during the growing season, it should be done by furrow irrigation rather
than overhead sprinkling.

**Selected References**

Farr, D.F., Rossman, A.Y., Palm, M.E., and McCray, E.B. 2007. Fungal
Databases, Systematic Botany & Mycology Laboratory, ARS, USDA. Retrieved
April 29, 2007, from http://nt.ars-grin.gov/fungaldatabases/

Haware, M.P., Nene, Y.L., and Mathur, S.B. 1986. Seed-borne diseases of
chickpea. Tech. Bull. 1. Danish Gov. Inst. Seed Pathol. 32 pp.

Jones, R.A.C., and Coutts, B.A. 1996. Alfalfa mosaic and cucumber mosaic
virus infection in chickpea and lentil: incidence and seed transmission.
Ann. Appl. Biol. 129:491-506.

Kaiser, W.J. 1981. Control of Ascochyta blight of chickpea through clean
seed. Pages 117-122 in: Proc. Workshop on Ascochyta blight and winter
sowing of chickpea. M.C. Saxena and K.B. Singh, eds. ICARDA, Aleppo,
Syria.

Kimber, R.B.E., Scott, E.S., and Ramsey, M.D. 2006. Factors influencing
transmission of *Didymella rabiei* (ascochyta blight) from inoculated
seed of chickpea under controlled conditions. Eur. J. Plant Pathol.
114:175-184.

Kumar, A., Jalali, B.L., Panwar, M.S., and Sangwan, M.S. 1983. Fungi
associated with different categories of chickpea seeds and their effects
on seed germination and seedling infection. Int. Chickpea Newsl.
9:19-21.

Nene, Y.L., and Reddy, M.V. 1987. Chickpea diseases and their control.
Pages 233-270 in: The Chickpea. M.C. Saxena and K.B. Singh, eds. CAB
International, Oxford, UK.

Singh, K., Singh, A.K., and Singh, R.P. 2004. Effect of fungicidal seed
treatment on the fungi associated with chickpea. Ann. Plant Prot. Sci.
12:450-451.

Reddy, J.R., Ahmed, K.M., and Varma, B.K. 1988. Bacterial wilt of
chickpea caused by *Xanthomonas campestris* (Pam.) Dowson. Int. Chickpea
Newsl. 19: 13-15.

(Prepared by Antonio Trapero-Casas, and Walter J. Kaiser)

![](media/image1.jpeg){width="3.375in" height="2.1680555555555556in"}

Fig.1. Necrotic lesions on a chickpea pod infected by *Ascochyta
rabiei*. Concentric rings of brown pycnidia can be observed in the
lesions.

![](media/image2.jpeg){width="3.5in" height="1.7777777777777777in"}

Fig. 2. Kabuli chickpea seed (right) with necrotic black lesions are
infected with *Ascochyta rabiei*; healthy seeds (left).

![](media/image3.jpeg){width="3.375in" height="2.2215277777777778in"}

Fig. 3. Pycnidium of *Ascochyta rabiei* oozing conidia in a necrotic
lesion on chickpea tissue.

![](media/image4.jpeg){width="3.5in" height="2.1930555555555555in"}

Fig. 4. Isolates of *Botrytis cinerea* from chickpea seed vary greatly
in cultural characteristics, sporulation, and sclerotial formation.

![](media/image5.jpeg){width="2.625in" height="1.9652777777777777in"}

Fig. 5. Three of 10 surface disinfected desi chickpea seeds plated on
potato dextrose agar are infected with *Ascochyta rabiei*.

![](media/image6.jpeg){width="2.625in" height="1.8319444444444444in"}

Fig. 6. *Phoma medicaginis* var. *pinodella* was seed-borne in four of
10 seeds of *Cicer* *pinnatifidum*.

![](media/image7.jpeg){width="2.625in" height="1.75in"}

Fig. 7 Brown chlamydospores of *Phoma medicaginis* var. *pinodella* in
naturally infected chickpea tissue.

![](media/image8.jpeg){width="1.738888888888889in" height="2.65in"}

Fig. 8. Necrotic lesions on the stems of a chickpea seedling infected
from seed by *Ascochyta rabiei*.

**Lentil**

Several lentil pathogens may be associated with reproductive structures
and are transmitted with seed, or affect seed quality by downgrading, or
both. Nonparasitic factors such as weather and harvesting technique can
also affect seed quality and cause economic losses. On the other hand, a
high incidence of the pathogen *Ascochyta lentis* in seed may sometimes
make the seed unsuitable for planting, yet have no effect on germination
or visual appearance.

The following lentil pathogens are transported or transmitted with seed:
*Ascochyta lentis*; *Colletotrichum truncatum*, *Botrytis cinerea* and
*B. fabae*, *Erwinia rhapontici*, *Fusarium avenaceum*, *Sclerotinia
sclerotiorum*, *Stemphylium botryosum*, *Pea seed-borne mosaic
potyvirus* (PSbMV); *Bean yellow mosaic potyvirus* (BYMV); and *Broad
bean stain comovirus* (BBSV).

The incidence of *Colletotrichum truncatum* in seed is usually low, even
when seed is harvested from fields severely infested with anthracnose,
and seed-to-plant transmission has never been experimentally proven. It
is generally believed that seed infection is insignificant in the
epidemiology of anthracnose compared with soil-borne inoculum.
Discoloration of lentil seed by *C. truncatum* is uncommon. Similarly,
although *E. rhapontici* can cause pink seed in lentil, its incidence is
low and it has not been documented whether seed-to-plant transmission
occurs in the field. However, even a low incidence of pink seed can
cause economic loss because the affected seed may appear to have been
treated with a fungicide.

Species of *Ascochyta*, *Botrytis*, *Sclerotinia*, *Stemphylium*, and
*Fusarium* cause lentil diseases that are more serious in regions where
rainfall can occur in the late growing season and during harvest. These
regions include the western Great Plains of North America, parts of
Australia, the Indian subcontinent, and sometimes the Pacific Northwest
of the U.S. In contrast, in the Middle East, where lentil originated,
crops typically mature and ripen under terminal drought and depend on
residual soil moisture to fill seed. Under these conditions fungi have
little opportunity to invade and discolor maturing seed.

Pod infection by *A. lentis* leads to seed infection. The seed shrivels
and turns purplish-brown, and whitish patches of mycelium and pycnidia
may be visible on the surface. When wet weather occurs, the pathogen
spreads and can cause severe seed discoloration (Fig. 1) even after a
crop has been swathed. In Canada testing by commercial labs for internal
seed-borne *Ascochyta* is common. Annual incidence and severity of
Ascochyta blight in Saskatchewan (the main area of lentil production)
have been reported regularly in the Canadian Plant Disease Survey since
1990. In wet years values in individual samples, incidence typically
ranges up to 50%, with provincial means from 2 to 7%. Similar high
levels of seed-borne *Ascochyta* have been reported in Australia.

The frequency of seed-to-seedling transmission of *A. lentis* is only
10% even under favorable conditions of soil moisture and temperature. In
the 1980s, a four-year study in Saskatchewan designed to relate seed
infection with subsequent epidemic development showed variable results.
However, it led to the recommendation that in the semi-arid brown and
dark brown soil zones planting seed with up to 5% infection could be
tolerated, given normal weather conditions.

The cultivars used in the study were susceptible to *A. lentis*; it is
not known if different tolerances might apply with the newer
*Ascochyta*-resistant cultivars now grown in Canada. However, it is
noteworthy that recent reports in the Canadian Plant Disease Survey
indicate high levels of infection and discoloration in seed samples of
resistant cultivars. This emphasizes how strongly seed infection can be
influenced by wet weather and saprophytic growth of *Ascochyta* on
mature plant tissue.

*Botrytis cinerea* and *B*. *fabae* develop on lentil crops at different
stages throughout the growing season in different areas of the world,
but cause seed discoloration (Fig. 2) when wet weather leads to dense
growth during flowering and maturation. Under these conditions,
infection is initiated in dead petals that persist in the flower after
pollination. The fungus then spreads into the developing pod. Some seed
is so shriveled that it cannot be harvested, but levels of infection in
harvested seed may reach 30 %. Many of the infected seeds are no longer
viable.

In addition to its effect on visual appearance, *B. cinerea* can lead to
seedling blight when infected seed is planted. Viable infected seeds
produce seedlings that die before or soon after emergence. In addition,
because the fungus is fast-growing, it can spread from an infected
seedling to adjacent healthy plants. Thus, the effect of seed-borne
*Botrytis* on the crop may be numerically higher than the initial
percent seed infection. On the other hand, it is not clear whether the
level of *Botrytis* seedling blight in a crop influences the level of
botrytis gray mold that may develop later in the season in that crop.

Although sclerotia of *S. sclerotiorum* may contaminate seed harvested
from an infested crop, the pathogen is seldom found at high levels
within lentil seed. The mechanism by which it infests lentil seed, the
effect it has on seed, and the conditions conducive to an outbreak are
all similar to those for *Botrytis*. However, unlike *Botrytis* spp.,
*S. sclerotiorum* does not have a conidial stage and its potential for
secondary spread from infection foci is limited to direct mycelial
growth in dense crop canopies. Thus, fewer pods and seeds in a crop
become infected.

*Stemphylium botryosum* is frequently found in lentil seed. However, as
it is a ubiquitous organism that is also saprophytic on dead plant
material, the role of seed-borne inoculum in initiating epidemics of
Stemphylium blight is uncertain. A high level of Stemphylium blight
induced by artificial inoculation in a growth chamber has been shown to
cause seed discoloration, but whether the disease causes seed
discoloration in nature is unknown.

*Fusarium avenaceum* is a common cause of soil-borne seedling blight and
root rot in lentil and is not usually considered to be seed-borne on
lentil. However, in Canada moderate levels (≥ 5%) of infection have been
reported in harvested seed in years with persistent wet weather after
the crop matures. Transmission to the pods and seed is presumably due to
rain splash of inoculum from soil and crop residues on the soil surface.
Infested seed shows reduced germination and vigor; under wet conditions
seed is most likely discolored by a mixture of pathogens and saprophytic
fungi, such as *Alternaria alternata*.

Although three viruses are known to be transmitted in lentil seed, very
little is known about their effects on seed quality. Seeds for plants
infected with BBSV occasionally shown dark staining and plants infected
with PSbMV may produce seeds that are small and misshapen.

Nonparasitic factors that cause seed discoloration and downgrading
include hail damage and freezing injury. Hail on maturing lentil crops
leads to shelling out of seed from ripe pods and brownish bruises on
seeds retained within pods that were green when the storm occurred (Fig.
3). Because lentil has an indeterminate growth habit, late-season
rainfall and cool weather result in continued growth and flowering. In
western Canada this makes crops susceptible to early frosts. Immature
seeds that were frozen turn dark green and shrivel (Fig. 4) as they dry
down, thus contaminating healthy grain that matured earlier.

Lentil seed that is harvested mechanically or handled with augers is
susceptible to damage from chipping and splitting, especially if the
moisture content is below 16 %. This is especially the case with
cultivars that have large flat, rather than small round seeds. Physical
damage can be reduced by using slow combine cylinder and auger speeds.
Mechanical harvesting can also lead to a condition referred to as "earth
tag", especially in crops that have not matured uniformly or that are
very weedy. Green plant material passing through the combine is crushed
and releases juices on the lentil seeds, which then trap soil and dust
particles on their surface. The risk of earth tag can be reduced by
using desiccants to dry crops down before harvest. Lentil seeds with
light green seed coats also have a natural tendency to turn brown as
they age, especially at high storage temperatures. This is because of
oxidation of phenolic compounds in the seed coat. The discoloration
results in a gradual reduction in market value.

Selected References:

Bailey, K.L., Gossen, B.D., Gugel,R.K., and Morrall, R.A.A. 2002.
Diseases of Field Crops in Canada, 3^rd^ ed. Can. Phytopathol. Soc.,
Saskatoon, Canada.

Bayaa, B., and Erskine, W. 1998. Diseases of lentil. Pages 442-443 in:
The Pathology of Food and Pasture Legumes. D.J. Allen and J.M. Lenné,
eds. ICRISAT / CAB International, Wallingford, U.K.

Beniwal, S.P.S., Bayaa, B., Weigand, S., Makkouk, K.H., and Saxena, M.C.
1993. Field Guide to Lentil Diseases and Insect Pests. International
Center for Agricultural Research in the Dry Areas, Aleppo, Syria.
(<http://www.icarda.cgiar.org/Publications/Field_Guides/Lentil/Lent7.html>)

Bos, L., Hampton, R.O., and Makkouk, K.M. 1988. Viruses and virus
diseases of pea, lentil, faba bean and chickpea. Pages 591-615 in: World
Crops: Cool Season Food Legumes. R.J.

> Summerfield, ed. Kluwer Academic Publishers, Dordrecht, The
> Netherlands.

Huang, H. C., Erickson, R. S., Yanke, L. J., Hsieh, T. F., and Morrall,
R. A. A. 2003. First report of pink seed of lentil and chickpea caused
by *Erwinia rhapontici* in Canada. Plant Dis. 87: 1398.

Morrall, R.A.A. 1997. Evolution of lentil diseases over 25 years in
western Canada. Can. J. Plant Pathol. 19: 197-207.

Morrall, R.A.A., Carriere, B., Ernst, B., Pearse, C., Schmeling, D., and
Thomson, L. 2007. Seed- borne pathogens of lentil in Saskatchewan in
2006. Can. Plant Dis. Surv. 87: 120-121.

([http://www.cps-scp.ca/cpds.htm]{.underline})

Morrall, R.A.A., Carriere, B., Pearse, C., Schmeling, D., and Thomson,
L. 2003. Seed-borne pathogens of lentil in Saskatchewan in 2002. Can.
Plant Dis. Surv. 83: 121-123.

([http://www.cps-scp.ca/cpds.htm]{.underline})

Saskatchewan Pulse Growers. 2000. Pulse Production Manual. 2^nd^ ed.
Saskatchewan Pulse Growers, Saskatoon, Canada.
([www.saskpulse.com/growing/index.php](http://www.saskpulse.com/growing/index.php))

(Prepared by R. A. A. Morrall)

![](media/image9.jpeg){width="3.0in" height="2.1333333333333333in"}

Fig. 1. Comparison of healthy (left) and Ascochyta infected (right)
lentil seeds of two varieties Laird and Eston. (Courtesy R. A. A.
Morrall)

![](media/image10.jpeg){width="3.0in" height="2.0131944444444443in"}

Fig. 2. Seed discoloration of lentil caused by *Botrytis* spp. (Courtesy
R. A. A. Morrall)

![](media/image11.jpeg){width="3.0in" height="2.0930555555555554in"}

Fig. 3. Brownish bruises on lentil seeds caused by hail when pods were
still green (Courtesy R. A. A. Morrall).

![](media/image12.jpeg){width="3.0in" height="2.8930555555555557in"}

Fig. 4. Dark green discoloration and shriveling of lentil seeds caused
by frost. (Courtesy R. A. A. Morrall)

**Pythium seedling and root rot**

Pythium seedling and root rot of chickpea has been reported from India,
the US (Washington State, Idaho, California), Iran, Spain, Brazil and
Canada (Alberta). The large-seeded kabuli types are much more
susceptible than the dark-seeded, small irregularly shaped desi types
with thick seed coats. *Pythium* causes pre-emergence and post-emergence
damping-off and root rot of chickpea and can cause major yield loss
without seed treatments in some locations.

Pythium seedling and root rot of lentil has been documented in the
Pacific Northwest of the U.S., India, Iran, Syria, Turkey, Canada
(Saskatchewan), and Mexico. The light-seeded low-tannin cultivars are
much more susceptible than the dark-seeded varieties, which contain
tannins that are inhibitory to *Pythium.*

**Causal Organisms**

*Pythium* spp., *P. ultimum* Trow (vars. *sporangiferum* and *ultimum*),
and *P. irregulare* Buisman have been identified as causal agents of
root rot and damping-off of chickpea, but other *Pythium* spp. may cause
disease, given the wide host range of this genus and the difficulty of
identification to species. *Pythium* spp, *P. ultimum, P.* *irregulare*
and *P. aphandidermatum* Fitz. have all been isolated from diseased
lentil plants. All species form thick-walled oospores in infected root
tissue (Fig. 1).

**Symptoms**

In chickpeas, a major symptom is seed rot and emergence failure. Seeds
which fail to emerge frequently have soil adhering to them, giving the
appearance of small brown balls (Fig. 2). Roots are also attacked,
leading to necrotic, discolored roots with less root branching and
feeder roots (root pruning). Crown and hypocotyl tissue can be
discolored and rotted. Plants are stunted, some plants can die before
flowering, and yield is reduced.

In lentils, seeds can also be damped-off. Adult plants show stunting,
yellowing or purpling of the leaves, and discoloration of main tap root
and feeder roots, in contrast to the white color of healthy roots (Fig.
3). Lateral or feeder roots are sparse, and plants often die when the
weather becomes warm and dry. In the field, infested areas show poor
stands (Fig. 4), and parts of the plant or the entire plant will turn
light grey to light brown and die (Fig. 5).

**Disease Cycle**

*Pythium* spp. survive in the soil as oospores or thick-walled
sporangia. *Pythium* spp. are stimulated to germinate by seed or root
exudates, and can rapidly infect seeds and embryos, often in less than
48 hrs*. Pythium* can infect either directly by hyphae or from zoospores
produced in sporangia .Most of the research has been done on chickpea,
but similar principles probably apply to lentils. *Pythium* on chickpea
is more severe in wetter soils with high water potentials. Disease
severity is greater in heavier, poorly-drained soils. On many hosts,
*Pythium* is favored by cool soil temperatures, which slows seed
emergence. In Spain, *Pythium* was severe on chickpeas planted in the
fall-winter, but not as prevalent in spring plantings. However, with
chickpea, damping-off was similar across the range of 10 C to 25 C.
Resistance to damping-off increases with seedling age. Seeds
pre-germinated for 48-96 hrs were resistant to seedling rot and decay by
*P.* *ultimum* Once plants are infected, oospores and sporangia are
formed in roots, which then decay and release oospores into the soil.
The disease is monocyclic- i.e. one cycle of inoculum each year.

**Control**

Pythium seed and seedling rots can be effectively controlled by chemical
seed treatments. These include mefenoxam and metalxyl, both which are
specific for Oomycetes. Both are registered for use on lentil and
chickpea in US, Canada, and Australia. Captan, a more general protectant
fungicide, is registered in the US on lentils, and thiram, another
protectant, is used in Australia and Canada. Some of the registrations
on lentils are only for low-tannin varieties. These protectant chemicals
can also be formulated with other fungicides, such as fludioxonil, for
protection against *Rhizoctonia.* However, none of these treatments are
systemic in the root and will not control root rot. There are no
effective chemical soil treatments that are registered or economical.

Numerous biological seed treatments have been tested and found
effective, including *Penicillium oxalicum, Pythium oligandrum,
Psuedomonas fluorescens,* and *Bacillus* spp. In the U.S., Kodiak
(*Bacillus* *subtilis*) and T-22 Planter Box (*Trichoderma harzianum*)
are registered on lentils and chickpea. In addition, a large amount of
research has been conducted in India on biological control using seed
treatments with PGPR (plant-growth promoting rhizobacteria), primarily
with *Pseudomonas* and *Bacillus* spp., which are used by growers.

Cultural techniques, such as later planting to avoid wet, cool soils,
can reduce disease. For example, in eastern Washington, chickpeas are
typically planted in late spring after field peas. Pythium diseases may
be less severe on well-drained sandy soils, compared to heavy clay
soils. Crop rotation is probably not effective, given the wide host
range of *Pythium* spp. and the ability to survive form more than one
year in the soil. However, recent work (Paulitz, unpublished), using DNA
quantification of 10 *Pythium* spp. across eastern Washington, suggests
that grain legumes may favor and select for different *Pythium* species
than wheat and barley.

**Selected References**

Beniwal, S. P. S., Baya'a, B., Weigand, S., Makkouk, K. and Saxena, M.
C. 1993. Field Guide to Lentil Diseases and Insect Pests. ICARDA,
Aleppo, Syria

Bhatti, M. A., and Kraft, J. M. 1992. Influence of soil moisture on root
rot and wilt of chickpea. Plant Disease 76:1259-1262.

Kaiser, W. J. and Hannan, R. M. 1983. Etiology and control of seed decay
and preemergence damping-off of chickpea by *Pythium ultimum*. Plant
Disease 67: 77-81.

Kaiser, W. J., and Hannan, R. M. 1984. Biological control of seed rot
and preemergence damping-off of chickpea with *Penicillium oxalicum*.
Plant Disease 68: 806-811.

Kaiser, W. J. and Horner, G. M. 1980. Root rot of irrigated lentils in
Iran. Can. J. Botany 58: 2549-2556.

Nene, Y. L. and Reddy, M. V. 1987. Chickpea diseases and their control.
Pages 233- In: The Chickpea. M. C. Saxena and K. B. Singh, eds. CAB
International, Wallingford, Oxon, UK.

Paulitz, T. C., Dugan, F., Chen, W., and Grünwald, N. J. 2004. First
report of *Pythium irregulare* on lentils in the United States. Plant
Disease 88: 310.

Trapero, Casas, A., Kaiser, W. J. and Ingram, D. M. 1990. Control of
Pythium seed rot and preemergence damping-off of chickpea in the U.S.
Pacific Northwest and Spain. Plant Dis. 74: 563-569.

Westerlund, F. V., Campbell, R. N. and Kimble, K. H. 1974. Fungal root
rots and wilt of chickpea in California. Phytopathology 64: 432-436.

(Prepared by T. Paulitz)

![](media/image13.jpeg){width="3.375in" height="2.702777777777778in"}

Figure 1. Oospores of *Pythium irregulare* group I (sensu Matsumoto) in
roots of lentils. Arrows indicate antheridia. (Courtesy T. Paulitz)

![](media/image14.jpeg){width="2.342361111111111in"
height="3.123611111111111in"}

Figure 2. Top: After 9 days, untreated chickpea seed (white-seeded)
planted in a field soil were rotted by *P. ultimum*. Germinated seed on
right treated with metalaxyl. Soil naturally adheres to decayed seeds,
giving them the appearance of a brown ball. Bottom: Pre- and post
emergence damping-off of chickpea caused by *P. ultimum* in field soil
collected at Central Ferry, WA. Ten seeds of a white-seeded line were
planted in the pot. (Courtesy W. Kaiser)

![](media/image15.jpeg){width="2.9319444444444445in"
height="3.498611111111111in"}

Figure 3. Roots of lentils rotted by *P. irregulare* group I. Plants
taken from a field near Garfield, WA. Note black discoloration of tap
root, and lack of lateral roots (top). Above ground, leaves of infected
plants turn beige to light-brown (bottom). (Courtesy T. Paulitz)

![](media/image16.jpeg){width="3.001388888888889in"
height="2.0006944444444446in"}

Figure 4. Field of lentils infected by *P. irregulare* (near Garfield,
WA). Note poor stand and white to grey discoloration of plants.
(Courtesy T. Paulitz)

![](media/image17.jpeg){width="3.251388888888889in"
height="2.154166666666667in"}

Figure 5. Resistance to root rot varied greatly among lentil lines grown
in irrigated trials at Karaj, Iran. Most plants in many lines were
killed by root rot. The primary pathogen was *P. ultimum.* Inset- Green
plant shows resistance to *P. aphandidermatum*, from a variety trial in
Shirez, Iran. (Courtesy W. Kaiser).

**Phytophthora Root Rot**

Soil-borne diseases of chickpea have been attributed to *Phytophthora*
species in at least five countries: Australia, Argentina, India,
Pakistan, and Spain. Names given to these diseases include root rot,
root and stem rot, foot blight, collar rot, damping off, and
pre-emergence damping off. The term "Phytophthora root rot (PRR)" can be
applied to all of these, but usually applies to post-emergent root
disease of chickpea caused by *Phytophthora medicaginis* Hansen and
Maxwell, previously identified as *P. cryptogea*, *P. megasperma* var.
*sojae*, *P. megasperma* f. sp. *medicaginis*, and *P. sojae* f. sp.
*medicaginis*. *P. medicaginis* or other *Phytophthora* species have not
been recorded as pathogens of lentil, in contrast to chickpea.

*P. medicaginis* and the legume diseases it causes have been reviewed.
Its major host is alfalfa (*Medicago sativa* L.) and it occurs worldwide
wherever alfalfa is grown, causing mild to severe root disease. This
pathogen is prevalent in, but not restricted to, soils in which alfalfa
has been grown.

In chickpea, PRR caused by *P. medicaginis* is a problem only in cool
season plantings. Crop loss assessment data have not been published.
However, PRR is a lethal disease that can affect large areas in chickpea
crops. Major losses occur in Australia. Aerial (Fig. 1) and ground
inspections indicated that about 20% of the area of chickpea crops in
northern New South Wales, Australia, died prematurely and was not
harvestable in 1988, a season with high rainfall conducive to PRR. The
occurrence of PRR in Australia is mainly in rain-fed grain production
areas of southern Queensland and northern New South Wales (latitude
26-33^º^S). Chickpeas in this region are most commonly grown in
Vertosols (moisture-retentive, neutral to alkaline cracking clay soils)
in rotation with cool season cereals. Alfalfa is grown as a forage crop
but is not often used directly in crop rotations on grain farms. Alfalfa
and annual *Medicago* spp. are naturalized and occur occasionally in
roadsides and pastures.

There is a lack of published information on PRR of chickpea from other
countries, which suggests that PRR is not regarded as a major disease
beyond Australia. It has not been recorded occurring naturally on
chickpea in North America, even though *P. medicaginis* occurs there
wherever alfalfa is grown.

**Symptoms**

Symptoms of PRR in chickpea can develop from seedling emergence to near
maturity. The disease is commonly lethal, causing wilting, chlorosis,
and rapid death of plants (Fig. 2) a week or later after heavy rain. Tap
roots of wilted plants are girdled by brown or black zones of rotting
that sometimes extend up to a centimeter above ground level. Premature
death over large areas often equates to 100% yield loss because
harvesting is not profitable.

Disease can also be sub-lethal. In this form, symptom expression is
dependent on soil moisture or possibly other factors. Lesions can be
restricted to lateral roots and/or the lower tap root, in which case
limited defoliation or foliar stress symptoms may be evident. Plants
sometimes survive an initial severe infection after a period of soil
saturation by regenerating lateral roots. Such plants have green shoot
tops but defoliated lower stems and reduced or no seed yield (Fig. 3).

PRR has a characteristic field distribution. It is most intense in or
confined to watercourses, depressions, areas of soil with heavy texture
and high clay content, and other areas where water collects and soil
saturation occurs (Figs. 1, 4). Physiological waterlogging unrelated to
*P. medicaginis* can also occur in these areas, but is distinguishable
by criteria listed in Table 1.

**Causal Organism**

*P. medicaginis* is the only species of *Phytophthora* that causes PRR
in Australia, based on extensive field surveys, taxonomic work, and
pathogenicity tests. Under synonyms it has reportedly caused PRR in
Argentina, India, and Pakistan although these identifications and/or
associations were not conclusive. A *Phytophthora* sp. closely related
to but distinct from *P. medicaginis* has been identified as the cause
of PRR in Spain and *Phytophthora citrophthora* is reportedly associated
with an occurrence of PRR in Argentina. The combined evidence indicates
that *P. medicaginis* is the most significant cause of PRR worldwide.

All isolates of *P. medicaginis*, whether from alfalfa or chickpea,
appear to be aggressive on chickpea. They show some specialization in
aggressiveness to alfalfa genotypes, but no distinct pathogenic races as
occur in some *Phytophthora* spp. such as *P. sojae*.

*P. medicaginis* is homothallic, producing oospores in single cultures.
Antheridia are either paragynous or amphigynous with a tendency toward
the latter. Oogonia are 37-39 µm diameter but occasionally 32-43 µm.
Oospores are 28-34 µm in diameter with oospore walls 2-4 µm thick.
Sporangia produced in flooded or liquid cultures are ovoid to ellipsoid
or pear-shaped and non papillate, with length 50-60 µm and width 24-30
µm. All of these morphological characteristics plus pathogenicity on
chickpea or alfalfa need to be considered for identification. In culture
growth is optimal at 25-26^º^C with little or no growth at 5^º^C and
35^º^C.

The life cycle is similar to that of other homothallic species of
*Phytophthora*. Hyphae in roots produce compatible oogonia and
antheridia which fuse, resulting in formation of thick-walled oospores.
These oospores are very resistant to the environment and can survive in
soil after roots have died. They germinate in a water-saturated
environment to form sporangia. Host exudates stimulate oospore
germination provided the pH is between 3.5 and 10.5 (optimum 6).
Sporangia produce motile zoospores which are released into the soil
solution. Zoospores are attracted chemotactically to the host roots
where they encyst. Cysts germinate to produce hyphae which then
penetrate and colonize the host. Colonization and death of roots
continue even while the host remains alive and the soil moist. Hyphae
produce new oogonia and antheridia which fuse to form the next
generation of oospores. Hyphae of some isolates produce chlamydospores
which can act as survival structures. The primary inoculum is usually
infected plant debris or soil containing oospores or chlamydospores.
These propagules and zoospores are dispersed by water during irrigation
and flood events. Zoospores can swim only a few mm in the soil water
matrix; movement over longer distances depends on water infiltrating or
moving over the soil.

**Host Range and Epidemiology**

*P. medicaginis* causes disease in a limited number of genera and
species within the family *Fabaceae*. It is recognized as a damaging
pathogen of alfalfa worldwide although resistant varieties have been
developed. Both alfalfa and chickpea can become infected and develop
symptoms at any growth stage. Field and greenhouse experiments in the
United States and Australia have shown the following species are also
susceptible: annual and perennial medics (*Medicago* spp,), sainfoin
(*Onobrychis vicifolia* Scop.), sulla (*Hedysarum* spp.), *Sesbania*
spp., and *Cicer* spp. other than chickpea. In these experiments,
infection was incited at seedling stages. Older plants were tested and
found not to be susceptible in the case of sainfoin. Prevalence of any
of the hosts listed above could lead to establishment of *P.
medicaginis* in chickpea cropping soils.

After *P. medicaginis* infects a crop of a susceptible host, it persists
for many years in the soil through a succession of non-host crops
(including cereals) and/or fallows. Survival for at least 13 years in
the absence of alfalfa has been recorded in the United States and for at
least three years in the absence of chickpea in Australia. The levels of
inoculum are reduced by crop rotation but remain sufficient to incite
disease in a season with high rainfall. Dormant oospores are reported to
be the main survival mechanism. However, *P. medicaginis* reportedly
colonizes live or dead non-host roots and is likely to survive actively
in the absence of a live host.

Development of PRR disease in chickpea requires that the soil be flooded
or saturated with water. Often, only poorly-drained parts of fields are
affected. Previously diseased fields can produce profitable crops if
soil saturation does not occur. Saturation triggers oospore germination
and production of sporangia and motile zoospores. Stomata of chickpea
seedlings at or just below the soil line are a significant infection
point for zoospores, particularly when there is water at the surface.

Soil temperatures below 22^º^C or above 28^º^C are likely to be
sub-optimal for disease development and soil temperatures below 18^º^C
or above 31^º^C inhibitory, based on *in vitro* studies on fungal
isolates and *in vivo* studies with inoculated alfalfa. However, field
or greenhouse studies of temperature effects have not been reported for
chickpea.

**Diagnostic Techniques**

Diagnosis of PRR is based on symptomatology, field distribution, soil
type, and isolation and positive identification of the pathogen. Except
for isolation, these are described above. Isolation of *P. medicaginis*
needs to be done from margins of active lesions from symptomatic living
plants. Roots must be washed thoroughly without damaging them and then,
for best results, washed for an additional 2 hr under running tap water.
Root sections 2-4 mm long with active lesions should be removed
aseptically, blotted dry on sterile tissue paper, placed on selective
agar and incubated in the dark at 25^º^C. A number of media selective
for *P. medicaginis* are available; these usually contain pimaricin and
antibacterial agents (vancomycin or rifampicin) incorporated into a base
medium such as V8 or corn meal agar. Hymexazol is partially inhibitory
to *P. medicaginis* but when used at 50 mg/L is beneficial for isolating
from soil-contaminated plants where *Pythium* spp. can be a problem.
When present, *P. medicaginis* will grow out from portions of the
infected tissue in 5-7 days. Isolation may also be done by baiting.
Sprouted alfalfa seeds or excised cotyledons, floated on sterile water
containing diseased tissue and incubated at 22^º^C, will attract
zoospores which infect the bait tissue, leading to production of
sporangia and, later, oospores. The infected cotyledons can be
transferred to corn meal agar plates with rifampicin to obtain pure
cultures. Chickpea seedlings can be used to bait *P. medicaginis* from
soil in pots.

Molecular techniques have been developed which can detect *P.
medicaginis* and distinguish it from other *Phytophthora* species. These
techniques include a real-time fluorescent PCR method that can detect
and quantify *P. medicaginis* in alfalfa; this may also be applicable to
chickpea. A preliminary baiting step would probably be required for
detection in soil.

**Management**

The first step in reducing losses from PRR is to avoid fields that are
poorly drained, prone to flooding or have a history of PRR of chickpea
or alfalfa. If practicable, grading fields to eliminate low spots will
reduce the risk of PRR. Minimizing practices that lead to soil
compaction, such as aggressive tillage, are also likely to reduce PRR.
In some countries, chickpeas are grown in no-tillage systems that can
lead to improved soil structure and hence improved drainage. Crop
rotation with wheat (a non-host) reduces incidence of diseased plants.
However, none of these practices eliminates the risk of major losses if
saturation does occur due to heavy or prolonged rainfall.

Seed dressing with metalaxyl controls pre-emergent and early
post-emergent disease caused by *P. medicaginis*, but is expensive and
does not prevent late season PRR. For these reasons metalaxyl is not
used by growers. While repeated soil drenches with metalaxyl (Fig. 2) or
phosphorous acid or its derivatives have been shown to control chickpea
PRR and other Phytophthora diseases in horticultural and forestry
situations, drenching chickpea crops is neither practicable nor
cost-effective. There has been no commercial application of biological
control products in chickpea.

Australian chickpea breeding programs have incorporated resistance from
ICC11870 (CPI56564), a desi type chickpea landrace from Tajikistan, into
several Australian varieties such as Yorker (Fig. 5). This resistance
can confer a yield advantage of more than 100% over highly susceptible
varieties under conditions that are moderately conducive to disease.
Much higher levels of resistance occurring in *C. echinospermum* (Fig.
6), a closely related species in the primary gene pool, have been
incorporated into interspecific hybrids and lines derived from these are
in the advanced stages in breeding programs. The mechanism of resistance
in *Cicer* species including chickpea is not understood, but it is of
the rate-limiting type rather than immunity.

**References**

Brinsmead, R. B., Rettke, M. L., Irwin, J. A. G., Ryley, M. J., and
Langdon, P. W. 1981. Resistance in chickpea to *Phytophthora megasperma*
f. sp. *medicaginis.* Plant Dis. 69:504-506.

CAB International, 2006. *Phytophthora medicaginis* \[text revised 2006
by R. Southwell\]. In: Crop Protection Compendium. Wallingford, UK: CAB
International. http://www.cabicompendium.org/cpc/home.asp.

Dale, M. L., and Irwin, J. A. G. 1991. Stomata as an infection court for
*Phytophthora megasperma* f. sp. *medicaginis* in chickpea and a
histological study of infection. Phytopathology 81:375-379.

Erwin, D. C., and Ribeiro O. K, 1996. Phytophthora Diseases Worldwide.
American Phytopathological Society, St. Paul, MN.

Frezzi, M. J. 1950. Las especies de "Phytophthora" en la Argentina.
Revista de investgaciones agricolas (Ministerio de Agricultura y
Gansderia, Argentina) 4:47-133.

Hansen, E. M., and Maxwell, D. P. 1991. Species of the *Phytophthora
megasperma* complex. Mycologia 83:376-381.

Knights, E.J., Southwell, R.J., Schwinghamer, M.W., and Harden, S. 2008.
Resistance to *Phytophthora medicaginis* Hansen and Maxwell in wild
*Cicer* species and its use in breeding root rot resistant chickpea
(*Cicer arietinum* L.). Australian Journal of Agricultural Research 59:
383-387. doi:10.1071/AR07175

Liew, E. C. Y., and Irwin, J. A. G. 1994. Comparative studies on
*Phytophthora megasperma* isolates from chickpea in Australia and Spain.
Mycol. Res. 98:1284-1290.

Stack, J. P., and Millar, R. L. 1985. Relative survival potential of
propagules of *Phytophthora megasperma* f. sp. *medicaginis*.
Phytopathology 75:1025-1031.

Suryanarayana, D., and Pathak, S. R. 1968. Foot blight--a new disease of
gram. FAO Plant Prot. Bull. 16:71-73.

(Prepared by M. W. Schwinghamer, R. J. Southwell, K. J. Moore, and E. J.
Knights)

Table 1. Distinctions between Phytophthora root rot and waterlogging in
chickpea

+-----------------------------------+----------------------------------+
| > Phytophthora root rot           | > Waterlogging                   |
+-----------------------------------+----------------------------------+
| > *P. medicaginis* present based  | > *P. medicaginis* absent (low   |
| > on baiting or isolation from    | > oxygen kills roots)            |
| > roots, or oospores in roots     |                                  |
+-----------------------------------+----------------------------------+
| > Occurs at any growth stage with | > Most likely to occur near      |
| > onset at temperatures up to     | > maturity at high temperatures  |
| > 31^0^ C                         |                                  |
+-----------------------------------+----------------------------------+
| > Onset a week or later after     | > Onset within two days after    |
| > flooding                        | > flooding                       |
+-----------------------------------+----------------------------------+
| > Lower leaves fall off           | > Plants die too fast to lose    |
|                                   | > leaves                         |
+-----------------------------------+----------------------------------+
| > Roots rotted                    | > Initially roots not rotted     |
+-----------------------------------+----------------------------------+
| > Plants easily pulled out        | > Initially plants not easily    |
|                                   | > pulled out                     |
+-----------------------------------+----------------------------------+
| > Controllable by metalaxyl       | > Insensitive to metalaxyl       |
| > drenches (not a cost effective  |                                  |
| > control measure)                |                                  |
+-----------------------------------+----------------------------------+
| > Partially controllable by crop  | > PRR resistance and seed        |
| > rotation, choice of variety,    | > treatment ineffective,         |
| > and seed treatment              | > manageable by field selection, |
|                                   | > withholding irrigation at      |
|                                   | > podding, choice of variety     |
+-----------------------------------+----------------------------------+

![](media/image18.png){width="3.3756944444444446in"
height="2.238888888888889in"}

**Fig. 1.** Aerial view of premature death of chickpeas (yellow areas)
from Phytophthora root rot compared with survival in elevated contour
banks. (Courtesy M. W. Schwinghamer)

![](media/image19.jpeg){width="3.2506944444444446in"
height="2.2368055555555557in"}

**Fig. 2.** Chlorosis and rapid death of chickpeas from Phytophthora
root rot in experimental plots. Surviving plots at center and left and
patches affected by runoff were protected by drench treatments of
metalaxyl through a period of soil saturation, thus distinguishing
*Phytophthora* rather than waterlogging as the cause of symptoms.
(Courtesy M. W. Schwinghamer)

![](media/image20.jpeg){width="1.9701388888888889in" height="2.875in"}

**Fig. 3**. Sub-lethal infection of chickpea by Phytophthora root rot,
with symptoms of browning and rotting of tap and lateral roots,
generation of new lateral roots, defoliation extending to shoot tops,
reduced podding. (Courtesy M. W. Schwinghamer)

![](media/image21.png){width="3.3756944444444446in"
height="2.2729166666666667in"}

**Plate 4**. Phytophthora root rot of chickpea, confined to an area that
was flooded temporarily. (Courtesy M. W. Schwinghamer)

![](media/image22.jpeg){width="3.254166666666667in"
height="2.1590277777777778in"}

**Fig. 5**. Resistance derived from a chickpea landrace in commercial
variety Yorker (right foreground). Trial plots were in a commercial
chickpea crop in a field infested naturally with Phytophthora root rot.
(Courtesy K. J. Moore)

![](media/image23.jpeg){width="4.411111111111111in"
height="2.9993055555555554in"}

**Fig. 6**. Resistant (R) and susceptible (S) reactions of chickpea (Ch)
and *Cicer echinospermum* (Ce) to Phytophthora root rot in a field
disease nursery. *C. echinospermum*, a slow-growing species with a
relatively prostrate growth habit, has been crossed with chickpea in
Australia to improve resistance. (Courtesy M. W. Schwinghamer)

Rhizoctonia Seed and Seedling Rot (Wet root rot)

*Rhizoctonia solani* can cause seed and seedling rot of both lentil and
chickpea as well as many other agricultural crops worldwide. The
pathogen is highly favored under no-till or reduced-till soil conditions
when organic matter is high. Sandy soils are particularly favorable to
this pathogen. The pathogen can infect at a wide range of soil
temperatures but is favored by cool (11 to 18ºC), wet soil conditions.

# Symptoms and Signs

Reddish to dark brown root lesions can develop on epicotyls and
hypocotyls at any plant stage from emergence to maturity. Upon
infection, seed rot or pre- and post-emergence damping-off of the
seedlings can result. Root lesions usually have a sunken wet appearance
and can severely constrict individual roots or root tips (Fig. 1). Dark
brown lesion appears above the main stem region on the main stem and may
expand to lower branches in order plants (Fig. 2). Often root tissue
below the constriction point will continue to survive and be healthy at
least initially (Fig. 3), but a lesion or collection of lesions can be
severe enough to pinch-off roots resulting in pointed brown tips at the
point of infection which is commonly referred to as root pruning. Small
black sclerotia that are irregular to spherical in shape and size may
form on old lesions. Infected plants become stunted and leaves turn
yellow beginning with lower leaves and progressing to the upper leaves
over time (Fig. 4). Often, circular patches of stunted plants can be
observed in fields. Due to the lack of a vigorous root system, infected
plants can be easily pulled from the soil.

## Causal organisms

Wet root rot of chickpea and lentil is caused by *Rhizoctonia solani*
Kühn (teleomorph *Thanatephorus cucumeris* (A. B. Frank) Donk). The
mycelium is described as being initially hyaline in color but turning
yellowish to brown with increasing age. The mycelial growth is
characterized by branches growing at right angles from each other with
small constrictions of the mycelium (3.5-7.2 µm wide) at the point of
branching. Septae are generally formed close to the base of each branch.
*R. solani* is divided into different anastamosis groups based on the
ability of hyphae to fuse with one another. Reported anastamosis groups
of *R. solani* affecting chickpea include AG-1, AG-4, AG-7 and AG-8, and
only AG-8 is reported to infect lentil. Survival spores are dark brown
to black sclerotia consisting of a mass of moniliform cells.
Thick-celled chlamydospores may also develop from the hyphae and act as
long term survival structures. The teleomorph often found on soil or
plant parts near the soil surface is not considered to be a major factor
in the pathology of this pathogen and is characterized by blackish,
barrel-shaped basidia that form under conditions of high humidity.

## Disease Cycle and Epidemiology

*R. solani* is both a soil and seed-borne pathogen of lentil and
chickpea but soilborne inoculum is considered the primary inoculum
source. Wind, rain/irrigation water, and farm machinery are the key
means of dissemination of infection propagules. Both the hypocotyls and
epicotyls of developing plants are the primary infection points. The
pathogen is capable of producing its own infection court through the use
of infection pegs produced from infection cushions that penetrate the
root cuticle and epidermis, but infection can also take place through
natural openings and wounds. *R. solani* can move both intracellular and
intercellular through the plant tissue and can survive in the soil for
extended periods of time as sclerotia (six years), as thick-walled
mycelium or as a saprophyte on organic debris.

# Management

Good seedbed preparation, crop rotation, disease-free seed, well-drained
soils, and tillage are the principal means of limiting root diseases in
lentils and chickpeas to this pathogen. Herbicide carryover, cold or
compacted soils and conservation tillage practices can increase the
potential for root rot and seedling blights caused by *R. solani*.
Chickpea and lentil cultivars with thick epicotyls have a less chance of
pre- and post-emergence damping-off by *R. solani* than those with thin
epicotyls, since large epicotyls are less likely to be completely
girdled upon infection. Chickpea and lentil cultivars with reduced seed
exudates of carbohydrates and amino acids at planting also tend to have
reduced disease problems. Since *R. solani* can remain viable in the
soil for several years, crop rotations with corn, small grains and other
non-host crops will limit the build up of soil inoculum. Since faba
bean, field bean, field pea, mustard, canola, rapeseed, soybean and
sunflower are hosts of *R. solani*, caution should be used when these
crops are grown in rotation with chickpeas and lentils to avoid the
build up of soil inoculum. A crop rotation including lentils and
chickpeas every 3 years or longer is recommended to reduce the severity
of this disease. Commercial biological and chemical seed treatments that
reduce the incidence of *Rhizoctonia* seed rot are available.

> **Selected References**

Cook, R. J., Schillinger, W. F., and Christensen, N. W. 2002.
Rhizoctonia root rot and

take-all of wheat in diverse direct-seed spring cropping systems. Can.
J.

Plant Pathol. 24: 349-358.

Shukla, T. N. Ahmad, Z. U., and Garg, S. K. 1972. Root rot of lentil.
Indian

Phytopathology 25: 584-586.

Sneh, B., Burpee, L., and Ogoshi, A. 1991. Identification of
*Rhizoctonia* species.

American Phytopathological Society Press, St. Paul, MN.

Sunders, S., Kataria, H. R., Satyavir, Sheoran, O. P. 2003.
Characterization of

*Rhizoctonia solani* associated with root/collar rots and blights.
Indian Phytopath.

56: 27-33.

Westerlund, F.V., Campbell, R. N., and Kimble, K.A. 1974. Fungal root
rot and wilt of chickpea in California. Phytopathology 64:432-436.

(Prepared by L. D. Porter)

![](media/image24.png){width="2.0548611111111112in"
height="2.2263888888888888in"}

Fig. 1. Root symptoms of chickpea infected by *Rhizoctonia solani.*
(Courtesy W. Chen and T. Paulitz)

![](media/image25.png){width="2.2493055555555554in"
height="1.8791666666666667in"}

Fig. 2. Advanced symptoms of wet root rot caused by Rhizoctonia solani
on old chickpea plant showing lesions on lower portion of stem and
bracnches. (Courtesy ICRISAT)

![](media/image26.jpeg){width="1.9736111111111112in"
height="3.079861111111111in"}

Fig. 3. Lentil plants infected by *Rhizoctonia solani* showing that the
main root below the infection (constriction) may remain healthy and
intact at least initially, but may become affected as the disease
progresses. (Courtesy ICARDA)

![](media/image27.png){width="2.7506944444444446in"
height="2.0659722222222223in"}![](media/image28.jpeg){width="1.9333333333333333in"
height="2.7069444444444444in"}

Fig. 4. A. Field symptoms (yellowing and stunting) of chickpea plants
infected by *Rhizoctonia solani.* (Courtesy W. Chen). B. A lentil plant
infected by *R. solani* showing yellowing of leaves progressing from
lower to upper plant parts. (Courtesy ICARDA)

**Dry Root Rot**

Chickpea

Dry root rot is a serious disease of chickpea in warm regions with high
temperature and dry soil condition particularly at reproductive stage.
The disease was first reported from India in 1931. Dry root rot is a
serious disease problem, particularly in semi-arid tropics Ethiopia, and
central and southern part of India, where the crop is grown under
rainfed and hot summer conditions. It has been reported from Australia,
Bangladesh, Ethiopia, India, Iran, Kenya, Lebanon, Myanmar, Mexico,
Nepal, Pakistan, Spain, Sudan, Turkey, and USA. The disease also occurs
on lentil.

Symptoms

Dry root rot usually occurs during flowering and podding stages, and
diseased plants are usually scattered in the field. Symptoms include
drooping of petioles and leaflets of very top part of the plant, not the
lower part of the plants, differentiating the disease from Fusarium
wilt. Sometimes the topmost leaves are chlorotic while the rest of the
plant is dry. The leaves and stems of affected plants are straw colored,
but in some cases the lower leaves and stems show brown color. The most
conspicuous symptom is sudden appearance of drying of the whole plants
that are scattered in the field (Fig. 1). When plants are uprooted, the
lower portion of the tap root usually remains in the soil. Tap root is
dark, shows signs of rotting and devoid of any lateral and fine roots
(Fig. 2). The root is dry (unless there is recent rains or irrigation).
Sometimes a grayish mycelial growth can be seen near the tip of the tap
root. The dead root is very brittle, shows shredding of the bark , and
root tips are easy to break. Dark and minute (80 to 175 um) sclerotia
may be seen with a hand lens on and in the inner side of the bark (Fig.
3). The stem pith at the collar region may contain minute sclerotia or
sparse thread-like mycelium (Fig. 4). *The pathogen* also infects
chickpea seedlings only under high temperature conditions.

Causal organism

Dry root rot is caused by the fungus *Macrophomina phaseolina* (Tassi)
Goid. Most of the literatures on dry root rot used the name *Rhizoctonia
bataticola* (Taubenh.) E.J. Butler, which is no longer valid. Detailed
morphological studies of chickpea isolates of *M. phaseolina* are not
available, and the pycnidial form has not been described on chickpea. A
polyclone antibody detection of the disease is available

*Macrophomina phaseolina* infects more than 500 plant species, including
many cultivated plants particularly belonging to the legume family, and
is widely distributed in warm climates. Although significant
phenotypical and pathogenic variations have been reported in *M.
phaseolina*, there is no evidence of host specialization. Cross
inoculation studies showed that isolates from chickpea and other crops
(pigeonpea, peanut and sorghum) are all pathogenic on safflower,
sorghum, bajra (*Pennisetum americanum* L. Leeke*)*, pigeonpea and
chickpea. Chickpea isolates were more pathogenic to other crops than to
chickpea. *Macrophomina phaseolina* can destroy epidermal cells of
chickpea roots without penetration, possibly by action of secreted
macerating enzymes.

Disease cycle and epidemiology

*Macrophomina phaseolina* is a facultative saprophyte and is both seed-
and soil-borne. It survives between crop seasons as mycelium and
sclerotia in infected chickpea debris. Its sclerotia are viable after
more than 12 months in soil. Dry root rot is a monocyclic disease. There
is no secondary spread of the disease from plant to plant in the same
season. Inoculum in soil is the source of primary inoculum. The inoculum
density and environmental condition determine the disease incidence.

Dry root rot is a high temperature disease, and hot weather and dry
condition are dependent factors for disease development. It occurs when
daily minimum temperature is above 20 C and maximum above 30 C. The
disease is favored by moisture stress (dry conditions) in sandy and poor
soils. Seedling infection cannot occur if the temperature is below 18 C.
Sandy loam soils are more favorable to the disease development than clay
loam soils. Nitrogen fertilizer enhances the disease, while phosphorus
fertilizer reduces disease severity.

*Macrophomina phaseolina* synergistically interact with Fusarium wilt
and with root knot nematodes (*Meloidogyne* spp.) Co-infection of
chickpea by *M. phaseolina* with either *Fusarium oxysporum* f. sp.
*ciceris* or with *Meloidogyne* sp. causes higher levels of disease than
inoculations with the pathogens alone.

Management

Use of resistant cultivars is the best option in managing dry root rot.
A number of cultivars and germplasm lines are identified as resistant
either to dry root rot or to both dry root rot and Fusarium wilt. A
study using one cross between resistant and susceptible lines revealed
that the resistance to dry root rot is monogenic, showing 3R:1S
segregation ratio.

Fungicides as seed treatment, drenching or spray can be used to
effectively manage the disease. Effective fungicides include captan,
chlorothalonil, carboxin, and carbendazim plus thiram.

Deep plowing and removal of infected host debris from the soil reduce
the disease severity. Moisture stress conditions should be avoided.
Early planting to help avoid later dry and hot conditions. Timely sowing
of early maturing varieties is good option to escape the hot weather
conditions near the maturity. Soil amendments with organic matter help
manipulate the inoculum density and inoculum potential. Amendments with
straw of wheat, corn, sorghum or oat reduced pathogen population and
disease incidence. However, amendment with straw of chickpea, lentil or
mustard had little effect on dry root rot, and amendment with straw of
urd (*Vigna mungo* L. Hepper), mungbean (*Vigna radiata* L. R. Wilczek)
or *Trifolium* increased the disease. It seems that organic amendments
with high a C/N ratio are detrimental to dry root rot of chickpea. A
number of biocontrol agents (*Trichoderma* spp.) have also been tested
as effective control measures.

If feasible, inducing soil moisture such as irrigation can control the
disease.

**Selected references**

Bhatt, J., and Vadhera, I. 1997. Histopathological studies on
cohabitation of *Pratylenchus thornei* and *Rhizoctonia bataticola* on
chickpea (*Cicer arietinum* L.). Advances in Plant Sciences 10:33-37.

Charu Jain, and Trivedi, P. C. 1998. Effect of the interaction between
*Meloidogyne incognita*, *Fusarium oxysporum* and *Rhizoctonia
bataticola* on chickpea. Journal of Phytological Research 11:95-97.

Dhrub-Singh, and Nema, K. G. 1987. Effect of soil amendment on
*Rhizoctonia bataticola* causing dry root rot of chickpea. International
Chickpea Newsletter 17:23-25.

Gangwar, R. K., Prajapati, R. K., Kamalwanshi, R. S., and Singh, S. R.
2007. Association of *Meloidogyne incognita* and *Rhizoctonia
bataticola* in chickpea. Progressive Agriculture 7: 183-184.

Nene, Y. L., Haware, M. P., and Reddy, M. V. 1978. Diagnosis of some
wilt-like disorders of chickpea (*Cicer arietinum* L.). Information
Bulletin No. 3, International Crops Research Institute for the Semi-Arid
Tropics, Hyderabad, India. 44 pp.

Pande, S., Kishore, G. K., and Rao, J. N. 2004 Evaluation of chickpea
lines for resistance to dry root rot caused by *Rhizoctonia bataticola*.
International Chickpea and Pigeonpea Newsletter 11:37-38.

Prajapati, R. K., Srivastava, S. S. L., and Chaudhary, R. G. 2003.
Incidence of chickpea dry root rot in Uttar Pradesh and Uttaranchal and
efficacy of seed dressing fungicides on seed germination and seedling
infection. Farm Science Journal 12:170-171.

Rajinder Singh, and Sindhan, G. S. 1998. Effect of fungicides on the
incidence of dry root rot and biochemical status of chickpea plants.
Plant Disease Research 13:14-17.

Singh, P. J., and Mehrotra, R. S. 1982. Influence of soil moisture and
temperature on *Rhizoctonia bataticola* infection of gram. Indian
Phytopathology 35: 327-329.

Singh, P. J., Nagra, P., and Mehrotra, R. S. 1981. Effect of organic
amendments on root rot of gram (*Cicer arietinum* L.) and their
influence on plant growth. Plant and Soil 63:199-208.

Singh, S. K., and Nene, Y. L. 1990. Cross inoculation studies on
*Rhizoctonia bataticola* isolates from different crops. Indian
Phytopathology 43:446-448.

Singh, S. K., Nene, Y. L., and Reddy, M. V. 1990. Effect of age on
susceptibility of chickpea to *Rhizoctonia bataticola*. International
Chickpea Newsletter 23:25-26.

![](media/image29.jpeg){width="2.8743055555555554in" height="1.7625in"}

Fig. 1. Dry root rot of chickpea caused by *Macrophomina phaseolina*
occurring in patches in the field. (Courtesy ICRISAT)

![](media/image30.jpeg){width="1.4631944444444445in" height="2.3875in"}

Fig. 2. Dry root rot of chickpea showing rotting root system devoid of
lateral and fine roots. (Courtesy ICRISAT)

![](media/image31.jpeg){width="1.4402777777777778in"
height="2.966666666666667in"}

Fig. 3. Minute sclerotia of *Macrophomina phaseolina* on infected
chickpea stem on and in insider of the bark. (Courtesy ICRISAT)

![](media/image32.jpeg){width="2.3465277777777778in"
height="3.1958333333333333in"}

Fig. 4. Minute sclerotia of *Macrophomina phaseolina* in the pith tissue
at collar region of chickpea stem. (Courtesy ICRISAT)

**Fusarium Wilt**

**Chickpea**

Fusarium wilt of chickpea was first reported in India by Butler in 1918
but its etiology was not sorted out until 1940 by Padwick. The disease
now occurs widespread in most chickpea growing areas in Asia, Africa,
southern Europe and the Americas. A rough estimate of a yearly 10 to 15%
yield loss has been reported from India and Spain, but 70% to total loss
of the crop can occur in years of severe attacks by the disease.
Fusarium wilt reduces chickpea production by decreasing both seed yield
and seed weight.

**Symptoms**

The disease can appear at any growth stage. Affected plants may be
grouped in patches or appear spread across a field (Fig. 1). Symptoms
can develop in highly susceptible cultivars within 25 days after sowing
('early wilt'), but usually are more conspicuous at the onset of
flowering, by 6 to 8 weeks after sowing, and can also occur up to the
pod stage ('late wilt'). Early wilting causes more yield loss than late
wilting. Affected seedlings show flaccidity of individual leaves
followed by a dull-green discoloration, desiccation and collapse of the
entire plant (Figs. 2 and 3). Late wilted plants exhibit drooping of the
petioles, rachis and leaflets, followed by yellowing and necrosis of
foliage. When uprooted before being severely affected or dried, affected
seedlings and plants show no external root discoloration. However,
dark-brown discoloration of xylem tissues can be seen when roots and
stem of a plant are split vertically or cross-sectioned (Fig. 4).
Sometimes only a few branches of a plant are affected resulting partial
wilt (Fig. 5).

**Causal organism**

Fusarium wilt of chickpea is caused by the monophyletic forma specialis
*Fusarium oxysporum* (Schlechtend.:Fr.) f. sp. *ciceris* (Padwick) Matuo
& K. Sato. The fungus was first named *Fusarium orthoceras* Appel &
Wollenw. var. *ciceri* by Padwick, and later Chattopadhyay and Sen Gupta
renamed the pathogen *Fusarium oxysporum* Schl. f. sp. *ciceri*
(Padwick) Snyder & Hansen.

On potato-dextrose agar (PDA) and potato-sucrose agar (PSA) media, the
aerial mycelium is at first white and cottony but later it may become
cream or salmon in color or remain white, when cultures are incubated
under near-UV light (NUV). Hyphae are septate and profusely branched.
*F. oxysporum* f. sp. *ciceris* can grow within a temperature range of
7.5 to 35ºC and pH range of 4 to 9.4. Optimum conditions for mycelial
growth are 25 to 27.5ºC and pH 5.1 to 5.9, depending upon strains.
Optimum pH for sporulation is 7.1 to 7.9.

Microconidia form abundantly on short, unbranched phialides arising
laterally on the hyphae; they are never in chains but grouped in false
heads. Microconidia are oval to ellipsoid, mono- or bi-cellular and
measure 2.5-3.5 x 5-11 µm. Macroconidia are sparsely produced and less
abundant than microconidia; they are thin-walled, usually three- to
five-septate, fusoid, slightly curved, pointed at both ends with a foot
shaped basal cell, and measure 3.5-4.5 x 25-65 µm. Macroconidia form on
individual phialides in aerial mycelium and on branched conidiophores in
sporodochia (Fig. 6). Chlamydospores are formed in old mycelia and
affected chickpea tissues; they are smooth or rough walled, terminal or
intercalary in hyphae, and may be formed single, in pairs, or in short
chains. Pionnotal colonies (slimy or gelatin-like) may also form after
frequent subculturing, which may be associated to loss of virulence of
the pathogen.

Two pathotypes (yellowing and wilting) and eight pathogenic races (races
0, 1A, 1B/C, 2, 3, 4, 5, and 6) have been identified in *F. oxysporum*
f. sp. *ciceris*. Pathotypes cause distinct yellowing (slow, progressive
and acropetal foliar yellowing) or wilting (fast, severe chlorosis and
flaccidity) syndromes (Fig. 7). The pathotypes and races can be
differentiated by pathogenicity assays. Races 0 and 1B/C are of the
yellowing pathotype whereas races 1A through 6 belong to the wilting
one. Pathogenic races can be identified by the severity of disease
reactions on a set of 10 differential chickpea cultivars (Table 1).
Races 1A through race 5 can be distinguished on a more concise set of
eight differentials based on 100% wilt (Table 2).

The eight pathogenic races also differ in their geographic distribution.
Races 2, 3, and 4 have only been reported in India, whereas races 0,
1B/C, 5, and 6 are found mainly in the Mediterranean region and the
United States (California). Race 1A is more widespread and has been
reported in India, California, and the Mediterranean region. An
intraspecific phylogeny of races was inferred in *F. oxysporum* f. sp.
*ciceris* whereby each of the eight races forms a monophyletic lineage.
Also, a simple stepwise pattern of evolution of the races was
demonstrated whereby virulence has been acquired according to two
simplest scenarios of few parallel gains or losses. In these scenarios,
race 0 (the least virulent), or race 1A (the most widely widespread),
would be ancestral to other races, respectively.

**Disease cycle and epidemiology**

Infested soil and infected seeds are sources of primary inoculum. It can
survive in soil and plant debris as chlamydospores for at least 6 years.
The pathogen is also internally seed-borne and it has been observed as
chlamydospore-like structures in the hilum region of the seed. Infected
seeds play an important role in the long-distance dispersal of the
pathogen and in its introduction into new areas.

*F. oxysporum* f. sp. *ciceris* is pathogenic to *Cicer* species only.
However, the fungus can also invade root tissues of other grain legumes
such as bean (*Phaseolus vulgaris*), faba bean (*Vicia faba*), lentil
(*Lens culinaris*), pea (*Pisum sativum*), and pigeon pea (*Cajanus
cajans*), as well as of many dicotyledonous weeds, which thus serve as
symptomless carriers of the pathogen. Soilborne chlamydospores are
primary inoculum for the disease. Germination of chlamydospores in soil
is stimulated by seed and root exudates of hosts and nonhosts. The
pathogen invades germinating seeds and seedlings directly without
wounds. After reaching the xylem tissue, the fungus spreads up in the
xylem vessels by means of hyphal growth and microconidia (Fig. 8), and
to adjacent vessels by lateral mycelial growth from infected ones. The
systemic colonization along the plant axis proceeds slowly since it
takes some 15 to 20 days for fungal structures to form in the stem xylem
vessels. This determinative phase of pathogenesis is then followed by
development of symptoms (i.e., the expressive phase) once intense
colonization of xylem vessels in root and lower stem has occurred.
Histological modifications in infected stems are associated with the
development of morphological symptoms, including formation of cavities
between phloem and xylem and xylem and medulla, as well as between
phloem and cortical parenchyma.

Abundant chlamydospores form in severely infected plants that eventually
are released into the soil as infested debris decomposes. Chlamydospores
may undergo cycles of renewal by limited saprophytic growth supported by
organic debris and root exudates, as well as by transient infections of
hosts and nonhosts. Short-distance spread of *F. oxysporum* f. sp.
*ciceris* can occur through the dispersal of infested soil or plant
fragments by water, wind, machinery and human activity.

Development of Fusarium wilt of chickpea can be influenced by the
virulence (defined as the amount of disease caused by a pathogen
genotype on a given host genotype) of pathogenic races, inoculum density
of the pathogen in soil, environmental conditions (e.g., air and soil
temperature, soil moisture, soil pH, etc) and cultivar susceptibility.

High temperature is critical for development of Fusarium wilt. An
unidentified race of *F. oxysporum* f. sp. *ciceris* caused increased
wilt with decreasing soil matrix potential at 25 and 30ºC, but
moderately at 15 and 20ºC, with an inoculum density of 500 and 1,000
propagules g^-1^ soil. No disease developed at 10ºC even with 5,000
propagules g^-1^ soil. Differences in virulence to a chickpea cultivar
also occur between races within a pathotype. For example, 5,000
chlamydospores g^-1^ soil of race 1B/C were needed to cause same amount
of disease that 1,000 chlamydospores g^-1^ soil of race 1A in cv. PV61,
and the amount of disease developed with 5,000 chlamydospores g^-1^ soil
of race 1A was equal to that developed with 1,000 chlamydospores g^-1^
soil of race 5.

Studies have modeled complex interactions among soil temperature, race
of the pathogen, inoculum density and host cultivar. For instance,
optimal temperatures vary for different race and cultivar combinations.
At optimum soil temperature, different inoculum densities are required
to develop maximum disease in different race/cultivar conbination. Risk
threshold charts indicated that limitation in disease by a deficient
factor is compensated by another factor. These charts can be applied to
predict the potential threat of Fusarium wilt in a geographic area based
on soil temperature, the race and inoculum density in soil, and
susceptibility of cultivars.

The effects of air and soil temperature on Fusarium wilt in chickpea
result in the development of disease epidemics being strongly related to
the date of sowing. Thus, under Mediterranean type of climate, advancing
chickpea sowing from early spring to early winter significantly delays
epidemic onset, slow down epidemic development, and reduces the final
amount of disease. The delay in epidemic onset decreases yield loss in a
linear relationship. Conversely, yield loss is increased exponentially
with the rate of disease progression favored by the delay of sowing.

**Diagnostic techniques**

Careful examination of uprooted, affected plants for the absence of
external root symptoms and presence of dark-brown discoloration in xylem
tissues of roots and stem can help in the diagnosis of Fusarium wilt in
chickpeas. Care should be taken not to confuse Fusarium wilt symptoms
with yellowing, wilting, and phloem discoloration that are exhibited by
chickpea infected by some plant viruses (e.g., pea streak carlavirus,
PSV). Similarly, when isolating the fungus from diseased plants in pure
culture, care should be taken with non-pathogenic *F. oxysporum* that
usually grows out of root and lower stem tissues, particularly if plants
are severely affected.

Morphological identification of *F. oxysporum* f. sp. *ciceris* must be
done on nutrient-poor media such as carnation leaf agar. Macroconidial
morphology should preferably be based on sporodochial macroconidia with
exposure of cultures to fluorescent light and preferably also to NUV
light. Repeated culture of the fungus on PDA or PSA should be avoided as
they tend to promote mutations leading to degeneration of strains.
Degenerated strains may show slower growth rates, loss of virulence,
abundant aerial mycelium, reduced pigmentation, or paucity of
macroconidia. Isolates can be stored in soil tubes to minimize loss of
pathogen traits. Conidia from liquid culture are transferred to sterile
soil (2 ml conidial suspension in 10 g of autoclaved soil in a test
tube) and after few days of incubation the infested soil is let to dry
and stored at 3 to 5ºC. Dormant cultures can also be stored in silica
gel, sterile filter paper, or as lyophilized cultures. Identification of
the pathogen can alternatively be made by means of a specific-PCR assay
using primers specific for the forma specialis as well as for races 0,
1A, 5, and 6. Also PCR primers are available for the identification of
the wilting pathotype both in planta and in soil.

**Management**

Fusarium wilt of chickpea is a monocyclic disease. Therefore, management
of the disease would be best achieved by exclusion and eradication of
the pathogen, as well as reducing the efficiency of primary inoculum.
Healthy seed should be produced in pathogen-free areas to avoid
seed-borne dissemination of the pathogen. Seed-borne inoculum can be
eradicated by seed dressing with commercial rates of Benlate T (30%
benomyl + 30% thiram).

Inoculum in soil can be reduced or eradicated by removal of host debris
and soil solarization, respectively. However, the most practical and
cost-efficient individual measure for management of Fusarium wilt of
chickpea is the use of resistant cultivars. Resistance to *F. oxysporum*
f. sp. *ciceris* races was identified mainly in desi germplasm as well
as in wild *Cicer* spp. Resistance against both races 0 and 5 was
identified in entries of *C. bijugum*, *C. cuneatum*, *C. judaicum*,
whereas entries of *C. canariense* and *C. chorassanicum* were resistant
to race 0 but susceptible to race 5. Valuable resistance was also
identified in kabuli germplasm. Over 160 resistant desi and kabuli
germplasm lines have been identified at ICRISAT and ICARDA, some of
which carry broad-based resistance. Kabuli lines and cultivars with
resistance against one or several races are available: line ILC 9784
(races 0, 1A, and 5); lines ILC 9785, ILC 9786, FLIP 86-93C, FLIP 87-33C
and FLIP 87-38C (races 0 and 1A); lines CA334.20.4, CA336.14.3.0 and
ICC14216K (race 5); cvs. ICCV2 through ICCV6 (race 1A), Sonora80 and
Surutato77 (several races), and UC15 and UC27 (races 0, 1A, 3, 4, 5, and
6).

Resistance in chickpea against specific races of *F. oxysporum* f. sp.
*ciceris* is governed by major resistance genes. Differentiation between
early wilting and late wilting phenotypes among segregating genotypes
led to hypothesis that resistance to race 1A is controlled by at least
three independent loci, of which two confer resistance in the recessive
form and a third is operative as a dominant allele. Either of the genes
in homozygous recessive form, and the dominant allele in the third
locus, independently confers late wilting resistance, but any of these
two resistance genes together confer complete resistance. The
complete-resistant phenotype of 'WR315' to races 0 and 5 and 'JG62' to
race 0 is characterized by inability of the pathogen to colonize the
vascular tissue following penetration. Rather, penetration of resistant
roots and growth of the pathogen within the cortical tissues of roots
and lower stem occurs without development of localized cell death. While
genotypes with complete resistance show no disease under natural
infections in the field, late-wilting genotypes may develop disease few
weeks before harvesting but show low incidence or no disease at
flowering time.

Subsequent work, mainly at the USDA-ARS Grain Legume Genetics and
Physiology Unit, WSU, Pullman, have revealed that resistance against
races 1A, 2, 3, 4 and 5 in 'WR315' is governed each by a single
recessive gene, namely *foc-1, -2, -3, -4*, and *-5*, respectively.
These genes are located in two clusters on linkage group 2: one that
contains the closely linked *foc-1* and *foc-4* resistance genes and the
other that comprises *foc-3* and *foc-5* ones. A second recessive gene
for resistance to race 4 was identified in 'Surutato77'. Also a
recessive gene conferring resistance against race 0 was identified in
desi line ICC 4958 (which also carries *foc-5*), but *foc-0* and *foc-5*
are not linked. Accumulation of phytoanticipins maackiain and medicarpin
in root tissues and exudates was proposed as a mechanism of resistance
in 'CPS1', 'ICCV2' and 'WR315'. Later work concluded that resistance to
the pathogen may not require salicylic- and methyl jasmonate -regulation
of defence-related genes.

Observed durability of the resistant cultivars used suggests that there
may be little or no selection for resistance-breaking races in this
pathosystem. For example, a high diversity of *F. oxysporum* f. sp.
*ciceris* races exists in the Mediterranean Basin, where resistant
cultivars generally have not been used. Conversely, widespread use of
race 1A-resistant cultivars in India has not led yet to reports of race
6, which specifically overcomes that resistance; rather, races 2, 3, and
4 which are virulent to race 1A-resistant cultivars, were reported in
India long before the release of these cultivars.

The resistant response of chickpea cultivars to infection by *F.
oxysporum* f. sp. *ciceris* races can be significantly influenced by
both temperature and co-infection with root-knot nematodes. Thus,
'Ayala' and 'PV1' chickpeas are moderately and highly resistant,
respectively, to race 1A at 24ºC, but highly susceptible at 27ºC.
Infection of chickpea by *Meloidogyne artiellia*, *M. incognita* or *M.
javanica* can breakdown resistance to *F. oxysporum* f. sp. *ciceris*.
Infection of the plant by *M. artiellia* overcame the complete
resistance to race 5 in lines CA334.20.4 and CA336.14.3.0, but not in
ICC14216K at any inoculum density, and the resistance of 'UC27' to the
same race was overcome with 30,000 chlamydospores g^-1^ soil but not
with 3,000 chlamydospores g^-1^ soil.

Planting resistant cultivars should be used in combination with good
cultural practices for the management, e.g., long crop rotations when
possible and early sowing. In Mediterranean environments, advancing the
sowing date from early spring to early winter can contribute to control
of the disease and, in addition, increase yield. However, these benefits
can be overridden if the cultivar sown is highly susceptible to wilt, a
highly virulent race of the pathogen prevails in soil, or both. This
emphasizes the usefulness of chickpea cultivars with a late-wilting
disease reaction.

**Selected references**

Castillo, P., Navas-Cortés, J.A., Gomar-Tinoco, D., Di Vito, M., and
Jiménez-Díaz, R.M. 2003. Interactions between *Meloidogyne artiellia*,
the cereal and legume root-knot nematode, and *Fusarium oxysporum* f.
sp. *ciceris* race 5 in chickpea. Phytopathology 93: 1513-1523.

Jiménez-Gasco, M.M., and Jiménez-Díaz, R.M. 2003. Development of a
specific polymerase chain reaction-based assay for the identification of
*Fusarium oxysporum* f. sp. *ciceris* and its pathogenic races 0, 1A, 5,
and 6. Phytopathology 93: 200-209.

Jiménez-Gasco, M.M., Navas-Cortés, J.A., and Jiménez-Díaz, R.M. 2004.
The *Fusarium oxysporum* f. sp. *ciceris* / *Cicer arietinum*
pathosystem: a case study of the evolution of plant-pathogenic fungi
into races and pathotypes. Internat. Microbiol. 7: 95-104.

Landa, B.B., Navas-Cortés, J.A., Jiménez-Gasco, M.M., Katan, J., Retig,
B., and Jiménez-Díaz, R.M. 2006. Temperature response of chickpea
cultivars to races of *Fusarium oxysporum* f. sp. *ciceris*, the causal
agent of Fusarium wilt. Plant Dis. 90: 365-374.

Navas-Cortés, J.A., Hau, B., and Jiménez-Díaz, R.M. 1998. Effect of
sowing date, host cultivar, and race of *Fusarium oxysporum* f. sp.
*ciceris* on development of Fusarium wilt of chickpea. Phytopathology
88: 1338-1346.

Navas-Cortés, J.A., Landa, B.B., Méndez-Rodríguez, M.A., and
Jiménez-Díaz, R.M. 2007. Quantitative modeling of the effects of
temperature and inoculum density of *Fusarium oxysporum* f. sp.
*ciceris* races 0 and 5 on the development of Fusarium wilt in chickpea
cultivars. Phytopathology 97: 564-573.

Sharma, K.D., Chen, W., and Muehlbauer, F.J. 2005. Genetics of chickpea
resistance to five races of Fusarium wilt and a concise set of race
differentials for *Fusarium oxysporum* f. sp. *ciceris*. Plant Dis. 89:
385-390.

Sharma, K.D. and Muehlbauer, F.J. 2007. Fusarium wilt of chickpea:
physiological specialization, genetics of resistance and resistance
genes. Euphytica. DOI 10.1007/s10681-007-9401-y.

(Prepared by R. M. Jiménez-Díaz, M. del Mar Jiménez-Gasco, B. B. Landa
del Castillo, P. Castillo, and J. A. Navas-Cortés)

**Table 1.** Disease reaction of differential chickpea lines to the
inoculation with pathogenic races of *Fusarium oxysporum* f. sp.
*ciceris* ^a^

  -------------- -- ------------ ------- ------- ------ ------- ------ ----- -----
  Differential      Pathogenic                                               
  chickpea line     race                                                     

                    0            1A      1B/C    2      3       4      5     6

                                                                             

  12-071/10054      S            M       S       R      R       R      R     M

  JG-62             R            S       S       S      S       S      S     S

  C-104             M            M       R/M     S      S       S      S     M

  JG-74             R            R       R       S      R       R      M     R

  CPS-1             R            R       R       S      M       M      M     R

  BG-212            R            R       R       S      M       M      R     R

  WR-315            R            R       R       R      S       R      R     R

  ICCV-2            R            R       R       S      S       S      S     M

  ICCV-4            R            R       R       S      S       S      S     M

  P-2245            S            S       S       S      S       S      S     S

                                                                             
  -------------- -- ------------ ------- ------- ------ ------- ------ ----- -----

^a^ Disease evaluated on a 0-4 severity scale depending on the
percentage of affected foliar tissue (0=0%, 1=1-33%, 2=24-66, 3=67-100,
4=dead plant) at 40 days after sowing in a infested soil (ref). Average
disease reactions of \<1 and \>3 were considered resistant (R) and
susceptible (S), respectively. Intermediate disease reactions were
considered moderate susceptible (M).

**Table 2.** Disease reaction of selected chickpea differentials to five
races of *Fusarium oxysporum* f. sp. *ciceris* (Sharma et al. 2005.
Plant Dis. 89: 385-390)

  -------------- -------------- ------------ -------- -- -------- ------- -------- -------- --
  Germplasm      Differential   Pathogenic                                                  
  accesion       line           race                                                        

                                             1A^a^       2        3       4        5        

                                                                                            

  W6-24867       JG-62^b^                    S (100)     S (94.3) S (100) S(100)   S (100)  

  W6-24868       P-2245^b^                   S (100)     S (100)  S (100) S(100)   S (100)  

  W6-24869       Sanford                     R (0)       S (100)  S (100) S (100)  S (95.0) 

  W6-24870       CRIL-1-53                   S (100)     R (0)    R (0)   R (0)    R (0)    

  W6-24871       CRIL-1-94                   R (0)       S (100)  R (0)   I (36.4) I (30.0) 

  W6-24872       CRIL-1-17                   R (0)       R (0)    R (0)   S (100)  R (0)    

  W6-24874       CRIL-1-36                   I (33.3)    S (100)  S (100) S (100)  R (0)    

  W6-24876       WR-315                      R (0)       R (0)    R (0)   R (0)    R (0)    

                                                                                            
  -------------- -------------- ------------ -------- -- -------- ------- -------- -------- --

^a^ S = susceptible, R = resistant, I = intermediate, disease incidence
(%) in parentheses.

^b^ JG-62 and P-2245 are resistant and susceptible, respectively, to
race 0

**Lentil**

Fusarium wilt is a serious soil-borne disease of lentil. Yield losses
can be significant in highly infected areas, particularly under dry and
warm conditions that are conducive to disease development. The disease
occurs in most lentil production regions and has become a limiting
factor of lentil production in certain areas. The disease has been
reported to occur in many countries including Algeria, Argentina,
Bangladesh, Brazil, Bulgaria, Canada, Chile, Colombia, Czechoslovakia,
Egypt, Ethiopia, France, Hungary, India, Jordan, Myanmar, Morocco,
Nepal, Pakistan, Sudan, Syria, Turkey, Tunisia, Uruguay, USA and the
former USSR.

**Disease symptoms**

Fusarium wilt of lentil may occur at two plant growth stages: seedling
stage and during reproductive stage, in patches at both seedling and
adult stages (Fig. 9). It usually occurs from flowering to pod-filling
stages (Fig. 10). Infected plants are stunted, leaves turn yellow and
curl from upper part of the plant and move downward. Other symptoms
include wilting and drooping of top leaves resemble water deficiency,
shrinking and curling of leaves from the upper part of the plants and
move downwards without premature shedding (Fig. 11). Plants finally
become yellow and die. Root systems show reduced growth with marked
yellowish brown discoloration, poorly developed nodules, show damage at
tap root tips and with proliferation of secondary roots above the area
of tap root injury. No external mycelial growth occurs on the roots.
Discoloration of vascular tissue in the lower stem is not always
visible. However, the disease also occurs in seedling stage in India.
General symptoms at seedling stage include seed rot and sudden drooping
more like wilting and damping-off (Fig. 12). The fungus could also cause
pre-emergence damping-off and seed rot.

**Causal organisms**

Vascular wilt of lentil is caused by the fungal pathogen *Fusarium
oxysporum* Schlecht. emend. Snyder & Hansen f. sp. *lentis* (Vasudeva
and Srinivasan) Gorden. The fungus produces abundant aerial mycelium in
culture initially for 3 to 4 weeks, white to orange or salmon in color,
then becoming gelatinous, plectenchymatous stroma. The fungus produces
three kinds of asexual spores: microconidia, macroconidia and
chlamydospores. Conidia are formed on mycelium at the ends of free
conidiophores (phialides). Microconidia are single celled, kidney
shaped, ellipsoid to ovoid, cylindrical, oblong or slightly curved, and
hyaline, often agglutinated into false heads. Macroconidia are 1 -- 5
septate, nearly straight to fusiform, falcate, slender and thin-walled.
Terminal or intercalary chlamydospores are formed on mycelium, single
celled, rarely 2-celled, spherical to pyriform, smooth and hyaline. The
sexual state of the fungus has not been reported. Recently *F. redolens*
is reported to cause lentil wilt in Italy. *F. redolens* is
morphologically very similar to, but phylogenetically distinct from *F.
orysporum*.

**Disease cycle and epidemiology**

The pathogen *F. oxysporum* f. sp*. lentis* infects only lentil under
natural conditions, however, it can infect *Vicia montbretti* under
artificial inoculation. Although the pathogen has a limited host range,
it can survive in soil for several years as dormant spores such as
clamydospores or saprophytically on root exudates of non-host plants.
Just like in other formae speciaes of *F. oxysporum*, it exhibits great
range of morphological variability, nutritional requirements, fungicide
sensitivity, in response to temperature, and in virulence.

Fusarium wilt is a monocyclic disease, so there is no secondary spread
in the same growing season. The initial inoculum level is very important
for determining the incidence of the disease. The pathogen survives over
the season as spores and dormant mycelium in infected plant debris. The
spores are released into the soil when the infected debris decomposes.
The fungus can also be seed-borne, and the seed-borne inoculum could be
either systemic infection inside the seeds or as concomitant
contamination with the seeds. Seed-borne inoculum is an important source
for the introduction of the disease into new production areas where the
disease is previously absent.

Soil temperature and soil moisture are the two most important
environmental conditions that affect the development of Fusarium wilt of
lentil. Warm soil temperature (20 -- 30° C) and relatively dry soil (25%
water holding capacity) conditions favor the disease development. Other
environmental conditions that may also affect the disease development is
soil type/aeration, soil pH and fertility, as well as soil microflora.
High levels of general saprophytic microflora adversely affect the
disease and reduce disease severity. However, soil borne
lentil-infecting nematodes, such as *Ditylenchus* spp., *Meloidogyne*
spp. and *Pratylenchus* spp., synergistically interact with *F.
oxysporum* f. sp. *lentis* and increase disease severity, presumably by
inflicting injuries to the root and facilitating fungal invasion.

Resistance sources are available in lentil and have been incorporated
into lentil cultivars. Resistance to lentil wilt is conditioned by major
(1 to 3) resistance genes. Resistance is usually more prevalent in
small-seeded lentils. Recently, however, resistance was also found in
large-seeded lentil.

**Diagnostic techniques**

Knowing the cropping history of the field is important in making field
diagnosis of Fusarium wilt of lentil. Recent lentil production with
known history of lentil wilt is an indication of potential wilt problem.
Wilted plants can be dug up to check for reduced root system with
discoloration but without external fungal growth. External fungal growth
indicates another warm weather disease, the collar rot caused by
*Sclerotium rolfsii*. Discoloration may or may not occur in the vascular
system. At field scale, Fusarium wilt appears as isolated circular
patches (foci) that enlarge as season advances in fields with broadcast
crop, whereas the disease advances along the rows in row-sown field.
However, these field symptoms are not exclusively indicative of Fusarium
wilt.

Laboratory diagnosis should be done with caution because of potential of
other saprophytic *Fusarium* spp. that morphologically resemble the
lentil pathogen. Positive isolation from the stem, above the collar
region, is indicative of systemic infection that increases the certitude
of the pathogen identity. Definitive identification of the pathogen
requires pathogenicity tests on susceptible lentil plants.

**Management**

Host plant resistance is the most economical means for managing Fusarium
wilt of lentil. Resistance sources have been identified and incorporated
into lentil cultivars and resistant cultivars are available in several
countries. Some high yielding lentil lines (like Plant L4) are reported
to be resistant to multiple diseases including rust, Ascochyta blight
and Fusarium wilt. However, in many lentil production regions,
particularly in industrialized counties, selection of lentil cultivars
depends on market demands. Therefore, the most suitable cultivars for
the market class may not have desirable resistance. Even if resistant
cultivars are available, relying on resistance alone is not sufficient
for satisfactory management of the disease. A number of cultural
practices should be used in conjunction to adequately manage Fusarium
wilt.

Select and plant seeds that are free of infection and concomitant
contamination. This is particularly important when expanding lentil
production in new areas or in areas where lentil wilt is not previously
reported, since contaminated or infected seeds are important means of
introducing the pathogen into new areas. If disease-free and clean seeds
are not available, fungicide treatments must be implemented. Seed
treatment with vitavax, thiram, pentachloronitrobenzene, Bavistin
(methyl N-(1H-benzoimidazol-2-yl)carbamate), benlate, or with boric acid
plus KMnO~4~ can eliminate or reduce seed contamination. However, direct
application of fungicides to soil after planting for the control of
Fusarium wilt is not an economical practice due to the high cost and
technical difficulty in incorporating fungicides into soil during the
growing season, despite the fact several fungicide were effective in
controlling lentil wilt in *in vitro* and under greenhouse conditions.

Select varieties that are early maturing or adjust planting date so that
the plants can escape hot weather conditions during the susceptible
growing stages. However, the most suitable planting date varies
depending on the climatic conditions of the production regions. The
principle is to reduce the overlap of susceptible growth stages with hot
weather conditions. Select planting fields where no prior history of
lentil wilt and/or implement a 3 to 5 year rotation since, the pathogen
has a restricted host range. Numerous reports, over three decades, have
shown microbial agents can reduce Fusarium wilt of lentil. It may be
feasible on subsistence or small farms to implement cultural practices
like applying manure or composts that enhance populations of general
microflora or supplement with selected antagonistic biocontrol agents.
However, this approach is not currently practical for large-scale
applications in industrialized countries.

Selected references

Agrawal, S. C., and Prasad, K. V. V. 1997. Diseases of Lentil. Science
Publisher, Inc. Enfield, New Hampshire, 155 pages.

Bayaa, B., and Erskine, W. 1998. Diseases of Lentil. Pages 423-471 in:
The Pathology of Food and Pasture Legumes, D. J. Allen, and J. M. Lenne,
eds. CAB International, Wallingford, Oxon, OX10 8DE, United Kingdom.

Bayaa, B., Erskine, W., and Singh, M. 1997. Screening lentil for
resistance to Fuarium wilt: methodology and sources of resistance.
Euphytica 98:69-74.

Beniwal, S. P. S., Bayaa, B., Weigand, S., Makkouk, Kh., and Saxena, M.
C. 1993. Field guide to lentil diseases and insect pests. International
Center for Agricultural Research in the Dry Areas (ICARDA), Aleppo,
Syria. 107 pages.

Chaudhary, R. G., and Amarjit, K. 2002. Wilt disease as a cause of shift
from lentil cultivation in Sangod Tehsil of Kota, Rajasthan. Indian
Journal of Pulses Research 15: 193-194.

De, R. K., Ali, S. S., and Dwivedi, R. P. 2001. Effect of interaction
between *Fusarium oxysporum* f. sp. *lentis* and *Meloidogyne javanica*
on lentil. Indian Journal of Pulses Research 14: 71-73.

El-Hassan, S. A., and Gowen, S. R. 2006. Formulation and delivery of the
bacterial antagonist *Bacillus subtilis* for management of lentil
vascular wilt caused by *Fusarium oxysporum* f. sp. *lentis*. J.
Phytopath. 154: 148-155.

Eujayl, I., Erskine, W., Bayaa, B., Baum, M., and Pehu, E. 1998.
Fusarium vascular wilt in lentil: Inheritance and identification of DNA
markers for resistance. Plant Breeding 117: 497-499.

Khare, M. N. 1980. Wilt of Lentil. Jawaharlal Nehru Krishi Vishwa
Vidyalaya, Jabalpur, M.P India 155 pp.

Khare, M. N. 1981. Diseases of lentils. Pages 163-172 in: Lentils, Webb,
C., and Hawtin, G. (eds.). Commonwealth Agricultural Bureaux, Farnham
Royal, UK.

Riccioni, L., Haegi, A., and Valvassori, M. 2008. First report of
vascular wilt caused by Fusarium redolens on lentil in Italy. Plant Dis.
92:1132.

Stoilova, T., and Chavdarov, P. 2006. Evaluation of lentil germplasm for
disease resistance to Fusarium wilt (*Fusarium oxysporum* f. sp.
*lentis*). Journal of Central European Agriculture 7:121-126.

(Prepared by W. Chen and B. Bayaa)

**Chickpea**

![](media/image33.jpeg){width="2.3784722222222223in"
height="1.5590277777777777in"}

Fig. 1. Patches of Fusarium wilt-affected, spring-sown kabuli chickpeas
cv. PV-61 in southern Spain. (Courtesy R. M. Jiménez-Díaz)

![](media/image34.jpeg){width="2.5027777777777778in"
height="1.6402777777777777in"}

Fig. 2. Leaf flaccidity and dull-green discoloration of kabuli chickpeas
infected by *Fusarium oxysporum* f. sp. *ciceris.* (Courtesy R. M.
Jiménez-Díaz)

![](media/image35.jpeg){width="2.6277777777777778in"
height="1.6861111111111111in"}

Fig. 3. Kabuli chickpeas severely affected by Fusarium wilt in field
microplots artificially infested with *Fusarium oxysporum* f. sp.
*ciceris* race 5. (Courtesy R. M. Jiménez-Díaz)

![](media/image36.jpeg){width="1.4333333333333333in"
height="3.1798611111111112in"}![](media/image37.jpeg){width="3.033333333333333in"
height="1.3798611111111112in"}

Fig. 4. Internal dark-brown discoloration in tap root and main stem of
chickpea infected by *Fusarium oxysporum* f. sp. *ciceris* when affected
plants are either split vertically (A) or cross-sections (B), (Courtesy
ICRISAT)

![](media/image38.jpeg){width="2.973611111111111in"
height="3.279861111111111in"}

Fig. 5. Partial wilting caused by *Fusarium oxysporum* f. sp. *ciceris*
when only a few branches of a plant are affected. (Courtesy ICRISAT)

![](media/image39.jpeg){width="3.0027777777777778in"
height="2.063888888888889in"}

Fig. 6. Branched conidiophore and macroconidia of *Fusarium oxysporum*
f. sp. *ciceris* from a sporodochium formed on carnation leaf agar
(x100). (Courtesy R. M. Jiménez-Díaz)

![](media/image40.jpeg){width="1.3333333333333333in"
height="0.9in"}![](media/image41.jpeg){width="1.3333333333333333in"
height="0.9in"}

Fig. 7. Disease syndromes caused by *Fusarium oxysporum* f. sp.
*ciceris* pathotypes in chickpeas grown in artificially-infested soil.
**A**. Yellowing caused by race 0 (yellowing pathotype) on 'P-2245'
plants after 24 days incubation at 25ºC; **B**. Wilting caused by race 5
(wilting pathotype) on 'PV-60' plants after 24 days incubation at 25ºC.
(Courtesy R. M. Jiménez-Díaz)

![](media/image42.jpeg){width="0.8708333333333333in"
height="1.3333333333333333in"}![](media/image43.jpeg){width="1.3263888888888888in"
height="0.8798611111111111in"}

Fig. 8. Growth of *Fusarium oxysporum* f. sp. *ciceris* race 5 within
xylem vessels of susceptible chickpea cv. JG-62. **A**. Phialides and
microconidia (x550); **B.** Mycelia in mature and immature vessels
(x550). (Courtesy R. M. Jiménez-Díaz)

**Lentil**

![](media/image44.jpeg){width="3.0069444444444446in"
height="1.926388888888889in"}

Fig. 9. Fusarium wilt of lentil occurs in patches in the field.
(Courtesy ICARDA)

![](media/image45.jpeg){width="1.7465277777777777in"
height="2.5597222222222222in"}

Fig. 10. Sudden dropping of lentil plant infected by Fusarium wilt at
flowering stage. (Courtesy ICARDA)

![](media/image46.jpeg){width="1.5402777777777779in"
height="2.553472222222222in"}

Fig. 11. Fusarium wilt of lentil showing wilting and curling of leaves
without premature shedding. (Courtesy ICARDA)

![](media/image47.jpeg){width="3.020138888888889in"
height="1.7666666666666666in"}

Fig. 12. Seedling stage infection of lentil by *Fusarium oxysporum* f.
sp. *lentis* showing drooping and drying of leaves and whole seedlings.
(Courtesy ICARDA)

**Black Streak Root Rot (*Thielaviopsis* root rot*)***

**Lentil**

Black streak root rot of lentil was first observed in USA in 1985. The
pathogen is a soilborne fungus that causes black root rot on a wide
range of plants. The common name "black streak root rot" is based on
darkly pigmented chlamydospores that form black streaks on roots of
infected plants. Although the disease black streak of lentil has been
reported from USA only, the pathogen is wide spread and is reported from
more than 40 countries. The disease also occurs on chickpea.

**Symptoms**

General symptoms include stunted plants, purplish stems due to stress
response, and roots with coalescing short black streaks formed by black
chlamydospores. The roots eventually become black entirely. Lentil roots
are more susceptible to the disease than hypocotyls.

**Causal organism**

Black streak root rot of lentil is caused by *Thielaviopsis basicola*
(Berk. & Br.) Ferraris (synonym: *Chalara elegans*). *Thielaviopsis
basicola* produces two types of asexual spores: microconidia
(endoconidia) and chlamydospores. Cylindrical microconidia are produced
endogenously within a phialide and have truncate ends. Chlamydospores
are black-brown, subrectangular aleuriospores that easily break apart at
maturity (Fig. 1). Its perfect (sexual) state is unknown.

**Disease cycle and epidemiology**

*Thielaviopsis basicola* has a wide host range including about 150 plant
species, especially members of the Fabaceae, Solanaceae, and
Cucurbitaceae families. All cool season food legumes (chickpea, lentil,
pea and faba bean) are susceptible to the disease. In some crops like
chickpea, both roots and hypocotyls are equally susceptible to the
disease, however, lentil roots are more susceptible than hypocotyls to
infection by *T. basicola*.

The fungus survives for long periods as black chlamydospores in lentil
residue and in soil, and as a saprophyte growing on organic matter in
soil. Soil factors that favor the diseases are cool temperature (17 to
25 C), and high level of moisture. Chlamydospores germinate and produce
multiple germ tubes, which elongate and grow toward root surfaces.
Chlamydospores are produced abundantly on root surfaces and gave the
black appearance and characteristic feature of the disease.

**Diagnostic techniques**

A characteristic feature of the disease in the field is initial black
streaks on the root surface, then the entire root becomes black. In the
laboratory, black subrectangular chlamydospores are definitively
diagnostic of black streak root rot (Fig. 2).

**Management**

The best control method is to avoid fields with high levels of
infestation and high disease potential for black streak root rot.
Rotations with less susceptible host crops such as cereals may help
reduce or delay build up of *Thielaviopsis basicola* populations.
Fungicides are not recommended for control of black streak rot of lentil
due to lack of economical feasibility. Resistance to black streak rot in
lentil is not known.

**Selected references**

Anonymous. 2006. *Thielaviopsis basicola* \[Distribution map\].
Distribution Maps of Plant Diseases. Map 218 (Edition 4). Wallingford,
UK: CABI.

Boerema, G. H., Pieters, R., and Hamers, M. E. C. 1993. Check-list for
scientific names of common parasitic fungi. Supplement Series 2c,d
(additions and corrections): Fungi on field crops: pulse (legumes),
forage crops (herbage legumes), vegetables and cruciferous crops. Neth.
J. Plant Pathol. 99 Suppl.: 1-32.

Bowden, R. L., Wiese, M. V., Crock, J. E., and Auld, D. L. 1985. Root
rot of chickpeas and lentils caused by *Thielaviopsis basicola*. Plant
Dis. 69:1089-1091.

Lloyd, A. B., and Lockwood, J. L. 1963. Effect of soil temperature, host
variety, and fungus strain on Thielaviopsis root rot of pea.
Phytopathology 53:329-331.

Subramanian, C. V. 1968. *Thielaviopsis basicola*. C.M.I. Descriptions
of Pathogenic Fungi and Bacteria No 170.

(Prepared by W. Chen and T. Paulitz)

![](media/image48.png){width="2.432638888888889in"
height="2.432638888888889in"}

Fig. 1. Chlamydospores and endoconidia of *Thielaviopsis basicola*
(Courtesy of W. Chen)

![](media/image49.jpeg){width="2.877083333333333in"
height="2.298611111111111in"}

Fig. 2. Chlamydospores of *Thielaviopsis basicola* on a
naturally-infected lentil root (Courtesy of T. Paulitz)

**Collar Rot**

Collar rot of chickpea and lentil is caused by *Sclerotium* *rolfsii*
Sacc. It occurs almost in every region where chickpea and lentil are
cultivated and is especially prevalent in areas with high soil moisture
and high temperature in the seedling stage of the crops. The disease is
frequently reported from and of economical importance in parts of
Bangladesh, Ethiopia, Pakistan and India, particularly when it crops are
is grown after rice.

**Symptoms**

The pathogen infects the collar region of the plant, causing a
yellowish-brown discoloration and rotting of the collar region (Fig. 1).
Infected young seedlings show damping-off symptoms. Plants infected at
an advanced stage gradually turn pale, droop and dry. White fluffy
mycelial growth of the fungus, generally associated with dirty white to
brown colored, mustard-seed sized sclerotia, can be seen in the infected
plant parts (Fig. 2 and 3). Sometimes the fungus may proceed downwards
causing root rot. Infected plants are easily pulled up as the root
system is poorly developed and side roots are destroyed (Fig. 1).

**Causal organism**

Collar rot of chickpea and lentils is caused by the fungus *Sclerotium*
*rolfsii* Sacc. The perfect stage of the fungus, although not reported
on chickpea and lentil, is *Corticium rolfsii* Curzi (Synonyms: *Athelia
rolfsii* (Curzi) Tu and Kimbrough). The basidial stage is rarely found
in nature.

The mycelium is septate and hyaline, branching at acute angles. Well
developed mycelium is in cord-like strands. The hypae have clamps in
form of forks and hooks or H-like connections. Newly developed mycelium
is slender, lacking clamp connections, snow white in color with silky
luster. Hyphal cells are binucleate to multinucleate. Basidia are
obovoid, 7-9 x 4-5 μm, each bearing 2-4 sterigmata that bear
basidiospores. Basidiospores are hyaline, smooth elliptical, apiculate
and 3.5-5.0 x 6-7 μm. Sclerotia are formed laterally from main hypal
strands, initially white becoming light brown to dark brown at maturity.
They are subspherical, 0.05-1.0 mm in diameter, with their surface being
finely wrinkled or pitted.

**Disease cycle and epidemiology**

The fungus has an extensive host range that comprises nearly 500 plant
species in 100 families. The most susceptible families are the
Compositae and Leguminosae families. Graminaceous species are less
susceptible. The fungus persists in many cultivated plants as well as
wild hosts. It commonly occurs in the tropics, subtropics, and other
warm temperate regions. The pathogen rarely occurs where average winter
temperatures fall below 0°C.

The fungus survives in the soil as sclerotia that remain viable for 2-3
years, but occasionally it persists as mycelium in infected tissues or
plant debris. They serve as a primary source of inoculum and are capable
of initiating infection with or without additional food source.
Sclerotia germinate in two ways: hyphal and eruptive. Hyphal germination
is characterized by the growth of individual strands of hyphae from the
sclerotial surface while eruptive germination is characterized by plugs
or aggregates of mycelium bursting through the sclerotial surface.
Eruptive germination in soil is greatest at 21-30°C and is less common
below 15°C or above 36°C. Germination is greatest at soil surface and
decreases with depth. Volatile compounds from re-moistened, undecomposed
plant tissue stimulate germination and growth of *S. rolfsii*.

*S. rolfsii* prefers low pH (\<8) environment, and grows at or near the
soil surface. Before the pathogen penetrates host tissue it produces a
considerable mass of mycelium on the plant surface, a process that can
take 2 to 10 days. The fungus produces a number of extracellular enzymes
including pectin methylesterase, cutinase, phosphatidase, galactinase,
mannase, xylanase and β-glucosidases. The production of cell
wall-degrading enzymes in conjunction with oxalic acid accounts in part
for the extensive host range of the pathogen. Action of these enzymes
results in tissue decay, further production of mycelium and the
formation of sclerotia. Sclerotia are disseminated by cultural practices
(infested soil and contaminated tools), infested transplant seedlings,
water (especially through irrigation), wind, and possibly on seeds. In
addition, a small percentage of sclerotia may survive passage through
sheep and cattle, and thus, could be spread through animal manure as
fertilizers. Sclerotia then germinate to produce mycelium which causes
new infections.

*Sclerotium* *rolfsii* is a variable fungus. Single-basidiospore strains
from the same field isolates show pronounced variability in
morphological characteristics suggesting field isolates might be
heterokaryotic.

Collar rot incidence may increase following periods of temperature and
moisture fluctuation; cycles of drying and wetting have been reported to
stimulated germination of sclerotia. The presence of an organic
substrate for mycelial growth may enhance disease severity. Factors that
increase nutrient leakage or predispose sclerotia to antagonism may also
accelerate their death. These include drying, heating, deep burial,
exposing sclerotia to chemicals and inducing changes in the integrity of
sclerotial rind. The disease becomes severe if the stubble of the
previous crop, such as paddy or sorghum, is left in the field.

**Diagnostic techniques**

The general symptoms of collar rot may look similar to those of crown
rot caused by *Sclerotinia* spp., but Sclerotium collar rot can be
differentiated from Sclerotinia crown rot by cord-like mycelial strands
(Fig. 3) and smaller (mustard seed size) sclerotia than those of
*Sclerotinia* spp (even those of *S. minor*). Another important
indication is that collar rot is favored by warm weather conditions,
whereas *Sclerotinia* crown infection occurs at cool conditions. Also
collar rot-affected seedlings are generally found in small patches in
the field and lie flat on the ground. Laboratory diagnosis involves
isolating the fungus from infected stems or sclerotia in either potato
dextrose agar (PDA) culture medium or on potato glucose agar medium
(PGA, 1.5%), pH 6. and incubated at 22°C for 48 h. After that period,
growth of an abundant whitish mycelium is observed, with clamp
connections in each septum of the mycelium, in addition to aerial
mycelium with a great number of whitish, spherical structures, the
initials of sclerotia.

**Management**

Control of collar rot in chickpea and lentil involves a combination of
cultural, biological and chemical methods. A number of cultural methods
are used to manage collar rot. The disease is greatly reduced by
manipulating the sowing date so that the seedling stage does not
coincide with the high soil moisture and temperature above 25°C.

Wide row spacing delay canopy closure and minimize microenvironment
conducive for sclerotial germination. However, whether wide row spacing
will increase yield is yet to be determined. Soil organic amendments
such as oat or maize straw have been found to limit disease incidence on
lentil. Soil solarization or solar heating is a relatively recent method
for controlling *S. rolfsii* inoculum. Sclerotia grown in vitro are
still viable after 12 hours at 45 °C, but are killed in 4-6 hours at 50
°C and in 3 hours at 55 °C. Covering soil with transparent polyethylene
sheets during the hot season increases soil temperatures and kills
sclerotia when the temperature under the sheets get hot enough for an
appropriate length of time.

The disease is reduced greatly by application of calcium fertilizers.
Increase in calcium levels in the tissues may partly offset the effect
of oxalic acid and cell wall degrading enzymes. Seed treatment
fungicides such as captan, thiram, triadimenol and carboxin are
partially effective. Other effective compounds include captan, mancozeb
and metalaxyl. Early seedling mortality is often reduced by application
of PCNB + thiram or thiram + carbendazim.

A number of antagonistic fungi have been shown to provide control
against *S. rolfsii* in controlled experiments, though field trial
results vary. Some of the commonly used organisms are: *Trichoderma
harzianum, T. viride, Bacillus subtilis, Penicillium* spp., and
*Gliocladium virens.*

Use of resistant cultivars provides good control of this soilborne
disease. Several resistant lines have been identified in Bangladesh,
India and Pakistan.

**REFERENCES**

Beniwal, S. P. S., Bayaa, B., Weigand, S., Makkouk, K., and Saxena, M.
C. 1993. Field guide to lentil diseases and insects pests. International
Center for Agricultural Research in Dry areas, Aleppo, Syria, 106pp.

Foster, N. R. 1999. Calcium nitrate to control collar rot during
chickpea germination. Seed Technology 21: 41-43.

Khare, M. N. 1981. Diseases of Lentil. Pages 163-172 in: Lentils. C.
Webb, and G. Hawtin, eds. Commonwealth Agricultural Bureaux, Farbham
Royal, UK,

Lifshitz, R., Tabachink, M., Katan, J., and Chet, I. 1983. The effect of
sublethal heating on sclerotia of *Sclerotium rolfsii.* Can. J.
Microbiol. 29: 1607-1610.

Nene, Y. L., and Reddy, M. V. 1987. Chickpea diseases and their control.
Pages 233-370 in The Chickpea. Saxena, M. C., and Singh, K. B. eds. CAB
International and International Center for Agricultural Research in the
Dry Area.Wallingford, Oxon, UK.

Peeples, J. L., Curl, E. A., and Rodriguez-Kabana, R. 1976. Effect of
the herbicide EPTC on the biocontrol activity of *Trichoderma viride*
against *Sclerotium rolfsii.* Plant Dis. 60:1050-1054.

Punja, Z. K. 1985. The biology, ecology, and control of *Sclerotium
rolfsii.* Annu. Rev. Phytopathol. 23:97-127.

Punja, Z.K., Grogan, R.G., and Unruh, T. 1982. Comparative control of
*Sclerotium rolfsii* on golf greens in northern California with
fungicides, inorganic salts, and *Trichoderma* spp. Plant Dis.
66:1125-1128.

(Prepared by E. Njambere and W. Chen)

![](media/image50.jpeg){width="1.2770833333333333in"
height="2.373611111111111in"}

Fig. 1. Collar rot of chickpea caused by *Sclerotium rolfsii* showing
yellowing of the leaves and constriction at the collar region. (Courtesy
ICRISAT)

![](media/image51.jpeg){width="2.7506944444444446in" height="1.71875in"}

Fig. 2. Sclerotia of *Sclerotium rolfsii* in soil and on collar region
of an infected chickpea plant. (Courtesy ICRISAT)

![](media/image52.jpeg){width="1.3159722222222223in"
height="2.625in"}![](media/image53.jpeg){width="1.7243055555555555in"
height="2.622916666666667in"}

Fig. 3. White mycelial strands of *Sclerotium rolfsii* on an infected
plants. A. chickpea (Courtesy ICRISAT), B. lentil (Courtesy ICARDA)

**Verticillium Wilt**

Verticillium wilt of chickpea was first reported in the United States
(California), and has also been reported in Italy, Pakistan and Tunisia,
and recently observed in Syria. The disease also affects lentil.

**Symptoms**

Symptoms of Verticillium wilt can appear on foliage and xylem tissue.
Leaves of affected plants may turn yellow prior to wilting. Xylem tissue
of plants may have light brown discoloration (Fig. 1). This brown
discoloration has been described as being lighter in color than the
xylem discoloration caused by *Fusarium oxysporum* f. sp. *ciceri*;
however, the symptoms caused by both fungi are very similar. In
controlled studies, *V. albo-atrum* did not cause rot of chickpea seeds,
as did *Fusarium* spp., *Rhizoctonia solani*, and *Macrophomina
phaseolina*. On lentil, *Verticillium* spp. were associated with root
rot of seedlings.

**Causal Organisms**

Verticillium wilt of chickpea and lentil is caused by the fungal
pathogens *V. albo-atrum* Reinke & Berth, or *V. dahliae* Kleb. Their
host ranges include both woody and herbaceous plants. Hyphal cells
contain only one nucleus, but hyphal tips may have multiple nuclei.
Conidiophores are branched and bear phialides that are arranged in
whorls, which bear conidia. Conidia are usually one-celled and hyaline.
*Verticillium* *dahliae* produces microsclerotia as resistant and
survival structures. Whereas *V. albo-atrum* does not produce
microsclerotia, but develops dark, thick-walled cells as resting
structures. However, in early literature, these two species names may
have been used interchangeably.

**Disease Cycle and Epidemiology**

The disease cycle of Verticillium wilt of chickpea and lentil has not
been described; however, it is likely similar to that previously
described on other crops. *Verticillium* spp. survive in the soil and
dead stems as microsclerotia or as dark, thick-walled cells. These
resting structures have been reported to survive anywhere from 9 months
to 4 years. *Verticillium* spp. penetrates roots directly or through
wounds, and the fungi colonize the xylem vessels. *Verticillium* spp.
can be dispersed by root-to-root contact, aerially as conidia or conidia
in blowing dust, and on seed infested with resting structures or
infected debris.

**Management**

Due to symptoms similar to Fusarium wilt, identifying that the causal
agent of the disease problem is *Verticillium* spp. is the first step in
management. Planting seed free of *Verticillium* spp. and rotation with
non-host crops are primary control measures for Verticillium wilt. If
available, plant resistant cultivars; chickpea lines resistant to
*Verticillium* spp. have been identified.

**Selected References**

Bhatti, M. A., Ali, S., and Khan, I. U. 1985. Comparative pathogenicity
of fungi associated with wilt of chickpea, with special reference to
*Verticillium albo-atrum* R & B. Trop. Pest Manag. 31:324-326.

Engelhard, A. W. 1957. Host index of *Verticillium albo-atrum* Reinke &
Berth.

(including *Verticillium dahliae* Kleb.). Plant Dis. Rep. Suppl.
244:23-49.

Erwin, D. C. 1958. Verticillium wilt of *Cicer arietinum* in southern
California. Plant Dis.

Rep. 42:1111.

Halila, H. M., and Harrabi, M. M. 1987. Wilt of chickpea in Tunisia
caused by *Verticillium albo-atrum*. Plant Dis. 71:101.

Harrabi, M. M., Halila, M. H., and Ben Brahim, M. 1985. Evaluating
sources of

> resistance to Fusarium and Verticillium wilts in chickpea. Arab J.
> Plant Prot. 3:91-93.

Ligoxigakis, E. K., Vakalounakis, D. J., and Thanassoulopoulos, C. C.
2002. Host range of *Verticillium dahliae* in cultivated species in
Crete. Phytoparasitica 30: 141-146.

Pegg, G. F., and Brady, B. L. 2002. Verticillium wilts. CABI Publishing,
New York, NY.

(Prepared by C. A. Bradley and K. A. Wise)

![](media/image54.jpeg){width="3.0131944444444443in"
height="1.5333333333333334in"}

Fig. 1. Cross-section of chickpea stems showing brown discoloration
caused by Verticillium wilt (Courtesy D. C. Erwin of University of
California via ICRISAT)