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Copyright 2005, Board of Trustees, University
Ft. Pierce ARC Research Report RL-1978-2 May, 1978
A TECHNIQUE FOR ENUMERATING VIABLE i IBRAR
OF SCEROTIUM ROLFSII IN SO
R. SonodaR 979
Abstract n lorida
A technique was developed for determining the presence of viable
propagules of Sclnrotium rolfii and of estimating propagule populations
in soil. Soil to be assayed was amended with oven-dried, ground shoots
of Digitaria deeiumbens and rolled out on a plate to a depth of 3 mm with
a glass rod. After incubation for 48 hours under high humidity, circu-
lar patches of mycelial growth originating from propagules were easily
seen and recorded. There were no differences in the propagule density
determined by this technique and a sieving method. With the new tech-
nique, samples were processed about three times faster than with the
Methods used to determine the number of viable propagules of
Sclerotium rolfsii Sacc. in soil usually involve the transfer of
sclerotia or baits from soil to another substrate (1, 2, 3). No means
for direct determination of populations of viable propagules of the
fungus in soil have been reported. The standard washing and sieving
method of assay (2) requires that sclerotia be picked out from among
seed and other material that may resemble sclerotia in size and color.
In the following study, two characteristics of S. rolfsii were exploited.
These were: 1) the ability of its propagules to-initiate growth in the
presence of undecomposed plant residues in soil and 2) the production of
heavy mycelial growth on the surface of soil with these residues,
especially under high humidity conditions. These factors were used to
develop a quick, easy to use, direct method of assay. This technique
was compared with Leach and Davey's sieving technique (2).
Development of Technique
Sclerotia of S. rolfsii placed on the surface of moistened Oldsmar
fine sand soil amended with 1% w/w dried, ground shoots of Pangola
(Digitaria decumbens) germinated and produced abundant mycelial growth.
Mycelial growth was more profuse in high humidity chambers. Sclerotia
placed on soil not amended with plant material germinated erratically,'
and made poor growth.
Oldsmar fine sand soil was obtained from a field previously crop-
ped to tomato where S. rolfsii had infected many of the plants. The
i/ Associate Professor, University of Florida, Institute of Food and
Agricultural Sciences, Agricultural Research Center, Fort Pierce.
I thank Girl Scout Volunteer laboratory aides Catherine Driscoll,
Cecelia Driscoll, Teresa Ford, Debra Henderson and Brenda Johnson
for technical assistance.
soil was air-dried, moistened to ab crt- ng capacity and
divided into two lots. One lot of soil vas amended with 1% w/w dried
Pangola; the other was not. Soil tm both lots was placed in 150 mm
diameter petri dishes to depths zf 2.7, 1 .4, and 8.1 mm. The dishes
were placed on a wire screen support over 500 ml distilled water, in
a 31 x 23 x 10 cm plastic containe- s-nd incubated at 30 + 10 C. After
24 hours of incubation, a few discrete circular patches of mycelial
growth of S. rolfsii were visible o the soil surface. Mere circular
patches were visible at 48 hours. At T2 hours some patches of growth
coalesced, but no further increase in their numbers *as seen. Scil to
which dried Pangola was added had abotkt five ti;e as many patches of
growth as unamended soil. Soil ..4 n deep, in smn eca.se ha dtice--
as many patches of growth as soil 2.7mm deep, hoWever, often -there
were less than twice as many patches. This indicated that .4 lnm was
too deep for our purposes.
Rnm-:`T samples of dried Pangola were tested for the presence of
S. rofstii. The dried materials were added to virgin soil infested
with S. rolfsii sclerotia and to noninfested virgin soil. The soil was
plated out as described in the preceding paragraph. No patches of S.
rolfsii growth were produced on noninfested virgin soil with any ofthe
sample':. Patches of S. rolfsii mycelia grew over the infested soil.
Air.-dried infested soil amended with 1% w/w dried Pangola was
moistened at 10% increments from 30 90% water-holding capacity of the
soil and incubated in high humidity chambers for 48 hours. No diffe-
rence in numbers of circular patches of growth was noticed at various
moisture levels, although mycelia of other fungi were present on pieces
of Pangola on soils with 80 90% of water-holding capacity.
Large glass plates (3 mm thick) were used to increase the speed
of handling samples and the capacity of the technique. The glass
plates were cut into 20.3 x 25.5 cm sections. Glass rods (3 mm diam.)
were glued to the four edges of the plates forming a 3 mm high container.
Moistened soil amended with 1% w/w dried Pangola was placed on the
plates. The soil was slightly flattened by hand, then rolled out 3 mm
deep over the plate with a glass rod. Excess soil was discarded or
used in subsequent plates. The plates, separated with 4 mm glass rods,
were stacked over each other in plastic humidity chambers. The cham-
bers were covered and either incubated on the laboratory bench (260 +
20 C) or in 300 + 10 C incubators.
The glass plates were weighed before use. Dry weight of the soil
was subsequently determined by drying soil on the plate in an oven
after patches of mycelial growth were counted. The population of S.
rolfsii was calculated as the number of patches of growth per gm dry
weight soil. Dry weight of soil did not differ more than 4% from test
to test for individual plates, thus in most tests dry weight of soil on
a particular plate was considered to be constant and dry weight deter,
mination was made once for each plate.
Comparison of Techniques
The plate technique and the sieving technique were compared. About
1600 g air-dried infested soil was divided into two lots of 800 g each.
Eight g of dried Pangola and enough water to bring the moisture level to
60% water-holding capacity was added to one lot and the lot further
divided into four aliquots which were rolled out separately on the glass
plates. Enough soil was used to completely fill the plate. The glass
plates were then placed in the humidity containers. The other 800 g lot
of soil was also divided into four aliquots of 200 g each and sieved
through a series of 10, 20 and 40 mesh screens. Sclerotia were col-
lected from the screens and transferred to petri dishes containing soil
amended with 1% dried Pangola.
The time required to process four replicates of each sample using
the plate technique was 12 minutes. This included adding dried Pangola
to soil, adding water, and rolling the soil out on the glass plates and
putting the plates into plastic containers. The average time required
for four replicates in the sieving technique was 32 minutes. This in-
cluded weighing the sample, washing soil through the sieve (including
rewashing to uncover sclerotia covered by soil and debris) and trans-
ferring sclerotia to dried Pangola-amended soil. The time required
to prepare the humidity chambers or to prepare petri dishes with
amended soil was not determined. The mean density of propagules
detected by the two techniques did'not differ at the 5% level of sign-
ificance within the two lots of each sample.
The patches of circular growth of S. rolfsii on the surface of in-
fested soil amended with 1% w/w dried Pangola were easy to see. Labora-
tory helpers can be readily trained to identify mycelia of S. rolfsii
as well as prepare the plates. In the dieving technique proficiency
in recognizing sclerotia is required.
Samples assayed by the plate technique can be processed about Trt- -
times faster than by the sieving technique. Furthermore, a minimum of
equipment is required for the plate technique, and dried, ground tissue
of other plants can be substituted for Pangola shoots. When ambient
laboratory temperatures are 250 C or more an incubator is not necessary.
In addition, because soil moisture requirement for the technique is
broad, soil brought in from the field generally does not have to be
moistened when adding plant residue.
The technique has been used routinely for determining the presence
and the population of S. rolfsii at the Agricultural Research Center,
Fort Pierce for the past three years. All samples have been from sandy
soil. The suitability of the technique for other types of soils has
not been investigated. Toribio (4) recently reported a similar techni-
que. In his technique, water was added to air-dried soil. No plant
residues were used.
1. Avizohar-Hershenzon, A. and P. Shacked. 1968. A baiting methods
for estimating the saprophytic activity of Sclerotium rolfsii
in soil. Phytopath. 58: 410-413.
2. Leach, L. D., and A. E. Davey. 1938. Determining the sclerotium
population of Sclerotium rolfsii by soil analysis and pre-
dicting losses of sugar beets on the basis of these analyses.
J. Agr. Res. 56: 619-631.
3. Rodriquez-Kabana, R., P. A. Backman, and E. A. Wiggins. 1974.
Determination of sclerotial populations of Sclerotium rolfsii
in soil by a rapid flotation-sieving technique. Phytopath.
4. Toribio, J. A. 1977. Une technique simple de denombrement direct
des sclerotes viables de Sclerotium rolfsii dans de sol. Ann.
Phytopath. 9: 177-182.