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RESEARCH ARTICLE
CURRENT SCIENCE, VOL. 81, NO. 6, 25 SEPTEMBER 2001
673
*For correspondence. (e-mail: maheshwaridk@rediffmail.com)
Isolation of siderophore-producing strains of
Rhizobium meliloti and their biocontrol potential
against Macrophomina phaseolina that causes
charcoal rot of groundnut
N. K. Arora†, S. C. Kang# and D. K. Maheshwari†,*
†Department of Botany and Microbiology, Gurukul Kangri University, Hardwar 249 404, India
#Department of Biotechnology, Taegu University, Kyungbuk, Kyongsan 512-516, Korea
Of the 12 isolates of root-nodulating bacterium,
Rhizobium meliloti isolated from the medicinal plant,
Mucuna pruriens, only two were able to produce
siderophores. The two isolates (RMP3 and RMP5)
were able to inhibit a widely occurring plant patho-
gen, Macrophomina phaseolina that causes charcoal
rot in groundnut. Further, there was a marked en-
hancement in percentage seed germination, seedling
biomass, nodule number and nodule fresh weight of
M. phaseolina-infected groundnut plants inoculated
with the strains RMP3 and RMP5, compared to
uninoculated and uninfected controls. Thus plant
growth promotory nature of both the rhizobial
strains was confirmed.
EXTENSIVE use of chemicals to control plant diseases
has disturbed the delicate ecological balance of the soil,
leading to groundwater contamination, development of
resistant races of pathogen and health risks to humans1.
Reluctance of most companies to test newer chemicals
due to the financial and registration difficulties has fur-
ther aggravated the problem2. However, considerable
attention has been paid to plant growth-promoting
rhizobacteria (PGPR), as the best alternative to chemi-
cals, to facilitate eco-friendly biological control of soil
and seed-borne pathogens3.
Most of the microorganisms which exhibit PGPR ac-
tivity, belong to the Gram-negative group and among
these fluorescent Pseudomonads are the most widely
studied. The siderophores of fluorescent Pseudomonads
and their introduction into the rhizosphere have been
widely explored. Siderophore production in iron stress
conditions confers upon these organisms an added ad-
vantage, resulting in exclusion of pathogens due to iron
starvation. However, use of antagonistic rhizobia has an
added advantage in that they have also the ability to fix
nitrogen. Different strains of rhizobia have now been
reported to produce siderophores. Strains of root-
nodulating bacteria have also been reported to produce
phytohormones like indole acetic acid (IAA)4, and anti-
bacterial compounds like rhizobiotoxin5 and bacterio-
cins6. This ability also confers upon nodule bacteria a
selective advantage and may lead to both direct and
indirect control of plant pathogens.
Macrophomina phaseolina (Tassi) Goid, is one of the
most destructive plant pathogens causing charcoal rot,
dry-root rot, wilt, leaf blight, stem blight and damping-
off diseases in a wide range of host plants7,8. M. phaseo-
lina has a very wide host range and therefore, it is not
easy to attain host resistance/tolerance. Chemical man-
agement is often uneconomical and not feasible, be-
cause the pathogen is both seed- and soil-borne8.
Biocontrol can thus offer a very good alternative for
management of this pathogen.
Unfortunately, most of the biocontrol agents perform
in vitro but fail to control the pathogens in field condi-
tions. The present investigation is aimed at assessing
the antagonistic potential of Rhizobium meliloti isolates,
both in vitro and in vivo, against M. phaseolina, that
causes charcoal rot of groundnut.
The root-nodulating bacterium isolates from Mucuna
pruriens were morphologically, biochemically and
physiologically tested according to Holt et al.9 and were
identified to be Rhizobium meliloti10.
The pathogenic strain of M. phaseolina was isolated
from the diseased seeds of groundnut by the blotter
technique11. The pathogen was identified using standard
mycological literature.
Siderophore production by the isolates was tested by
chrome azurol S (CAS) assay12. Siderophore production
was also checked by the top layer method. The strains
were spread over YEM agar and incubated for 48 h at
30°C. After incubation, a thin layer of CAS reagent12 in
0.7% agar was spread on the bacterial growth and plates
were again incubated for 24 h at 30°C. Formation of
yellow-orange halo around the colonies indicates
siderophore production13. To determine the type of
siderophore, culture supernatants were used. The pres-
ence of catechol-phenolic-type siderophores was tested
according to Arnow14 and Rioux et al.15, taking 2,3-
RESEARCH ARTICLE
CURRENT SCIENCE, VOL. 81, NO. 6, 25 SEPTEMBER 2001 674
dihydroxybenzoic acid as the standard. The presence of
hydroxamate siderophores in the supernatant was
checked according to Gibson and Magrath16, taking the
absorption spectrum of supernatant in the visible range
in a Shimadzu UV-VIS spectrophotometer (model UV-
1601).
Rhizobial isolates were checked for the ability to
produce hydrocyanic acid (HCN) and IAA. HCN pro-
duction was determined on tryptic soya agar17. Produc-
tion of IAA was determined according to the modified
method of Gupta et al.18.
The bacterial isolates were screened for ability to
inhibit M. phaseolina on agar plates. Rhizobial strains
were cultured in YEM broth. M. phaseolina was also
raised on YEM agar. A 6 mm mycelial disc of M.
phaseolina was centrally placed on YEM agar plates
and exponentially grown rhizobial strain (24 h) was
spotted on two sides of the mycelial disc. The plates
were incubated at 30°C for 5 days, to measure inhibition
of radial fungal growth as a clear zone between fungal and
bacterial colonies. Per cent inhibition was determined by
the reduction in fungal growth compared to control.
Earthen pots (24 × 12 × 12 cm) were filled with gar-
den soil. Mycelia of M. phaseolina were inoculated in
pre-sterilized oat grains in one-litre capacity flasks and
incubated for 5 days at 30°C, to prepare the culture for
soil infestation. The grain-based culture of M. phaseo-
lina was mixed in both the sterile and non-sterile soil,
so as to make the inoculum level of approximately
105 cfu g–1 soil. Rhizobial strains RMP3 and RMP5 were
grown in YEM broth up to log phase (108 cells ml–1).
The cells were harvested by centrifugation and coated
(approximately 108 cfu per seed) on the surface-
sterilized groundnut seeds by the slurry method19. After
drying for 3–4 h, the seeds were sown in the pots. The
experiment was designed with the following treatments
with groundnut: (i) soil (control); (ii) soil + M. phaseo-
lina; (iii) soil + RMP3; (iv) soil + RMP5; (v) soil +
RMP3 + M. phaseolina; (vi) soil + RMP5 + M. phaseo-
lina. Five replicates of each treatment were taken. The
plants were watered with tap water whenever required.
The disease incidence was noted as percentage of the
plants showing charcoal rot after 60 days. Plants were
harvested to measure seedling biomass and nodule fresh
weight of a local variety of Arachis hypogaea, and
compared with controls.
Root colonization study was carried out in earthen
pots filled with garden soil. The soil was infested with
M. phaseolina and surface-sterilized seeds, bacterized
with the isolates RMP3 and RMP5, resistant to ampicil-
lin (10 µg) were sown in respective pots. Controls were
also maintained as described earlier. Plants were up-
rooted carefully along with the adhering rhizosphere
soil. The soil was removed by shaking the roots gently.
One gram of the rhizosphere soil was serially diluted in
sterile distilled water for determining cfu g–1 soil. Cfu
counts of both rhizobia and M. phaseolina were deter-
mined on YEM agar by the spread plate technique. The
plates were incubated at 30°C for 48 h and 5 days for
rhizobia and fungi, respectively. For isolating and iden-
tifying rhizobial colonies, YEM agar was supplemented
with 10 µg ampicillin and actidione (to control fungal
growth).
Of the 12 rhizobial isolates screened for siderophore
production on CAS agar, only 02, RMP3 and RMP5
showed orange colour production and yellow-orange
coloured halo around the colonies, on CAS reagent
overlaid on YEM agar. Larger halo was formed around
the colonies of strain RMP5 in comparison to those of
strain RMP3, after 24 h of incubation. The 48-h-old su-
pernatant of culture broth of both the strains showed a
major peak at 400 nm, which corresponds to hydrox-
amate-type of siderophore. The culture supernatant of
none of the strains produced phenolate–catechol
siderophore. In rhizobia, the ability to synthesize
siderophore is restricted to a limited range of strains,
rather than a wide distribution13. There are now, how-
ever, a wide range and type of siderophores reported in
rhizobia20. Hydroxymate-type of siderophores have also
been reported in different rhizobial strains13,21.
None of the isolates produced hydrocyanic acid. Ear-
lier studies have also reported a very low incidence of
cyanogens in rhizobia and in other PGPR22,23. In fact, it
has been reported that production of HCN proved to be
deleterious to the plant22. However, all the twelve
strains produced IAA. Production of IAA is reported to
be more common in rhizobia. Prevost et al.24 reported
that 96% of the rhizobial isolates produced IAA,
whereas Antoun et al.22 observed IAA production by
56% strains of R. meliloti.
Of all the isolates only the two siderophore-producing
strains, RMP3 and RMP5, showed strong antagonism
against M. phaseolina (Figure 1). Strains RMP3 and
RMP5 showed 72% and 77% inhibition of fungal
growth, respectively after 5 days of incubation, com-
pared to control. There was a gradual increase in fungal
inhibition with incubation time and strain RMP5 was
found to be more effective than RMP3 (Figure 2). The
Figure 1. Antagonistic effect of strain RMP5 on M. phaseolina on
YEM agar.
RESEARCH ARTICLE
CURRENT SCIENCE, VOL. 81, NO. 6, 25 SEPTEMBER 2001
675
Figure 2. Inhibition of M. phaseolina by strain RMP5.
Table 1. Effect of seed bacterization with rhizobial strains on percentage germination, seedling biomass and nodule fresh
weight of groundnut in M. phaseolina-infested soil
Seedling biomass (g/plant) Nodule fresh weight (mg/plant)
Treatment Seed germination (%) 30 days 60 days 30 days 60 days
Control* 78.0 4.24 9.68 14.8 61.4
M. phaseolina• 58.0 2.94 3.24 9.6 24.2
RMP3♦ 88.8**,b 4.48 12.72**,b 20.2 88.7**,b
RMP5♦ 90.9**,b 4.72 13.21**,b 24.7 96.1**,a,b
RMP3 + M. phaseolina 85.1**,a 4.31 11.28**,a 13.7 71.6**,a
RMP5 + M. phaseolina 88.6**,b 4.45 11.78**,a 17.1 75.6**,a
∗With no pathogen and rhizobia; •With no rhizobia; ♦Without M. phaseolina; Results are mean of five replicates ± SD.
**Highly significant at P > 0.01; a,bMeans in the column followed by same letter are not significantly different.
formation of inhibition zone by the rhizobial antagonis-
tic strains RMP3 and RMP5 against M. phaseolina in
vitro, indicated secretion of certain metabolites by both
the strains. Increase in inhibition percentage with incu-
bation period, suggested release of metabolites with
incubation.
Rhizobial isolates RMP3 and RMP5 did not show
nodulation in groundnut plants, as evidenced by the
absence of nodulation in sterile soil. Seed germination
percentage of groundnut increased considerably in M.
phaseolina-infested soil by seed bacterization with both
the strains (Table 1). Strain RMP5 showed 52.8% and
RMP3 46.7% increase in germination percentage over
pathogen control (Table 1). The per cent germination of
seeds coated with rhizobial cells was even better than
the control. Bacterization with strains RMP3 and RMP5
increased the plant biomass and nodule fresh weight in
comparison to pathogen-infested control. Seedlings
(unbacterized) in M. phaseolina-infested soil developed
symptoms of charcoal rot. The black spots were clearly
visible on the stem of wilted plants. These plants
showed poor nodulation as evidenced by about 61%
decline in fresh nodule weight in comparison to control.
On the other hand, plants raised from bacterized seeds
were healthy and showed no signs of charcoal rot dis-
ease in M. phaseolina-infested soil. After 60 days, the
disease incidence reached 84%. Disease incidence re-
duced to a mere 3.8% in RMP5 and 7.9% in RMP3-
treated plants. Seed bacterization with RMP5 and RMP3
increased seedling biomass by 22% and 16.5% respec-
tively, of control, in infested soil. Nodule fresh weight
was also enhanced by seed bacterization with rhizobial
strains (Table 1). The increment in seedling biomass
and nodule fresh weight, by seed coating, was even
more in non-infested soil. There was a 36% increment
in seedling biomass and 56.5% in nodule fresh weight
RESEARCH ARTICLE
CURRENT SCIENCE, VOL. 81, NO. 6, 25 SEPTEMBER 2001 676
Table 2. In vivo population density of isolates RMP3 and RMP5 and fungal pathogen M. phaseolina in presence of each other
Log cfu g–1 soil after time (days)
7 15 30 45 60
RMP3* 7.16 ± 0.02 6.98 ± 0.019 6.48 ± 0.015 5.61 ± 0.018 5.00 ± 0.013
RMP5* 7.38 ± 0.021 7.20 ± 0.021 6.77 ± 0.019 5.89 ± 0.017 5.37 ± 0.015
M. phasolina+ 5.23 ± 0.013 5.52 ± 0.014 6.49 ± 0.012 6.56 ± 0.016 6.04 ± 0.014
RMP3 + (M. phaseolina) 6.90 ± 0.024 6.67 ± 0.018 5.74 ± 0.017 5.07 ± 0.018 4.55 ± 0.015
RMP5 + (M. phaseolina) 7.11 ± 0.019 6.86 ± 0.019 6.10 ± 0.021 5.42 ± 0.017 4.88 ± 0.011
M. phaseolina + (RMP3) 5.08 ± 0.019 4.53 ± 0.018 3.31 ± 0.014 2.33 ± 0.014 1.18 ± 0.011
M. phaseolina + (RMP5) 5.01 ± 0.017 4.41 ± 0.020 3.18 ± 0.022 2.11 ± 0.013 1.03 ± 0.009
*Without M. phaseolina; +Without rhizobia.
Results are mean of five replicates ± SD.
by strain RMP5 in non-infested soil compared to con-
trol, which was slightly more than that of strain RMP3
(Table 1).
Both RMP3 and RMP5 maintained high cfu g–1 up to
60 days in the presence of M. phaseolina, which was
marginally lower than the population of both the iso-
lates in soil without fungal infestation. The population
density of RMP5 was slightly higher than that of RMP3
both in infested and non-infested soils. Both the strains
strongly inhibited the M. phaseolina population in the
rhizosphere. The population of the pathogenic fungus
declined from 105 cfu g–1 to 67 and 62 cfu g–1 after 60
days by RMP3 and RMP5 bacterization, respectively
(Table 2).
There was a considerable decrease in disease inci-
dence, improvement in seedling biomass and fresh nod-
ule weight by seed bacterization with the strains RMP3
and RMP5, in the presence of pathogen. Thus rhizobial
strains not only acted as biocontrol agents against M.
phaseolina, but also proved to be plant growth promo-
tory in nature as evidenced by the increase in seedling
biomass and fresh nodule weight over uninoculated con-
trols. Siderophore and IAA have been reported to be
responsible for biocontrol and plant growth promotory
nature of various PGPR strains22,25. The root coloniza-
tion data also supported the biocontrol ability and
PGPR ability of both the rhizobial isolates. A steep de-
cline in the rhizosphere population of M. phaseolina in
the presence of rhizobial strains and a high population
density of both the rhizobial strains in rhizosphere, in-
dicated their association with groundnut roots. It has
also been reported earlier that Rhizobium reduced char-
coal rot disease caused by Macrophomina spp26. Strains
of R. japonicum have been reported to protect soybean
from M. phaseolina infection5. Rhizobia have earlier
also been reported for their ability to act as PGPR for
both legumes27 and non legumes22,28.
The isolates of R. meliloti, RMP5 and RMP3 have
proved to be effective in promoting the growth of a
non-host, besides inhibiting M. phaseolina causing
charcoal rot disease in groundnut. Use of rhizobia has
an added advantage over other organisms like
Pseudomonads, in that they have the ability to fix
nitrogen symbiotically with legumes and are eco-
friendly and non-pathogenic to humans. Another benefit
is that there is a better technical knowledge of inoculant
production and application for rhizobia. Thus two
isolates of R. meliloti (RMP3 and RMP5) appear to have
a great potential in controlling soil- and seed-borne
diseases caused by M. phaseolina.
1. Loper, J. E. and Ishimaru, C. A., in The Rhizosphere and Plant
Growth (eds Keister, D. L. and Cregan, P. B.), Kluwer Aca-
demic Publishers, 1991, pp. 253–261.
2. Heydasi, A. and Misaghi, J. J., Plant Soil, 1998, 202, 109–
116.
3. Weller, D. M., Chandler, J. L. and Feldman, A. W., Annu. Rev.
Phytopathol., 1988, 26, 379–407.
4. Wang, T. L., Wood, E. A. and Brewin, N. J., Planta, 1982, 155,
343–349.
5. Chakraborty, U. and Purkayastha, R. P., Can. J. Microbiol.,
1984, 30, 285–289.
6. Wilson, R. A., Handley, B. A. and Beringer, J. E., Soil Biol.
Biochem., 1998, 30, 413–417.
7. Young, P. A., Tex. Agric. Exp. Stn. Bull., 1949, 712, 1–33.
8. Singh, S. K., Nene, Y. L. and Reddy, M. V., Plant Dis., 1990,
74, 812–814.
9. Holt, J. G., Krieg, N. R., Sneath, P. H. A., Staley, J. T. and Wil-
liams, S. T., in Bergey’s Manual of Determinative Bacteriology,
Williams and Wilkins Press, Baltimore, USA, 1994.
10. Arora, N. K., Kumar, V. and Maheshwari, D. K., Symbiosis,
2000, 29, 121–137.
11. De Tempe, J., in Proceedings of International Seed Testing As-
sociation, 1963, vol. 28, 1933.
12. Schwyn, B. and Neilands, J. B., Anal. Biochem., 1987, 160, 47–
56.
13. Carson, K. C., Holliday, S., Glenn, A. R. and Dilworth, M. J.,
Arch. Microbiol., 1992, 157, 264–271.
14. Arnow, L. E., J. Biol. Chem., 1937, 118, 531–537.
15. Rioux, C. R., Jordan, D. C. and Rattray, J. B. M., Anal. Bio-
chem., 1983, 133, 163–169.
16. Gibson, F. and Magrath, D. I., Biochim. Biophys. Acta, 1969,
192, 175–184.
17. Miller, R. L. and Higgins, E. J., Phytopathology, 1970, 60, 104–
110.
RESEARCH ARTICLE
CURRENT SCIENCE, VOL. 81, NO. 6, 25 SEPTEMBER 2001
677
18. Gupta, C. P., Sharma, A., Dubey, R. C. and Maheshwari, D. K.,
Cytobios, 1999, 99, 183–189.
19. Somasegaran, P. and Hoben, H. J., in Handbook for Rhizobia:
Methods in Legume-Rhizobium Technology (ed. Garber, R. C.),
Springer Verlag, New York, 1994.
20. Guerinot, M. L., Plant Soil, 1991, 130, 199–209.
21. Pessmark, M., Pittman, P., Buyer, J. S., Schwyn, B., Gill, P. R.
and Neilands J. B., J. Am. Chem. Soc., 1993, 115, 3950–3956.
22. Antoun, H., Beauchamp, C. J., Goussard, N., Chabot, R. and La-
lande, R., Plant Soil, 1998, 204, 57–67.
23. De Britto Alvarez, M. A., Gagne, S. and Antoun, H., Environ.
Microbiol., 1995, 61, 194–199.
24. Prevost, D., Bordeleau, L. M., Caudry-Reznick, S., Schulman,
H. M. and Antoun, H., Plant Soil, 1987, 98, 313–324.
25. Noel, T. C., Sheng, C., Yost, C. K., Pharis, R. P. and Hynes,
M. F., Can. J. Microbiol., 1996, 42, 279–283.
26. Purkayastha, R. P., Menon, U. and Chakraborty, B. N., Indian J.
Exp. Biol., 1981, 19, 462–465.
27. Werner, D., Symbiosis of Plants and Microbes, Chapman and
Hall Publication, London, 1992.
28. Trinick, M. J., Nature, 1973, 244, 459–460.
ACKNOWLEDGEMENTS. Thanks are due to Council of Scientific
and Industrial Research and Technical Mission on Oilseeds and
Pulses for financial help, and Dr. Dharm Pal for encouragement.
Received 25 April 2001; revised accepted 25 July 2001
