NC State and USDA Cucumber Disease Handbook
Belly Rot (Rhizoctonia solani Kuhn)
Belly rot is caused by the fungus Rhizoctonia solani Kuhn
(anastomosis group 4). This fungus can cause fruit rot or damping-off
Occurrence and Importance:
Belly rot (soil rot, Rhizoctonia rot) is one of the most common
fruit rots of cucumbers in North Carolina. Losses vary from year
to year with a loss of 5 to 10 percent not unusual. The condition
is more prevalent in warm, humid weather. Fruit become infected
where they contact the soil in the field. Apparently healthy fruit
which have already been infected can develop severe symptoms within
Very small cucumbers show yellowish brown superficial discoloration.
Large fruit have a dark brown water soaked decay most often on
the side of the fruit in contact with the soil. The fruit may
have small cracks in the rotten area. Under humid conditions a
dense brown mold covers the rotted area.
The fruit can develop symptoms 24 hours after coming in contact
with the fungus and the entire fruit can rot in 72 hours. Temperatures
below 50°F retard disease development during transit and
Rhizoctonia solani Kuhn anamorph of Thanatephorus cucumeris
(Frank) Donk, anastomosis group 4, is the organism which causes
belly rot in cucumber.
Very rapid growth, color varies among isolates from nearly white
to gray and brown. Occasionally produces brown or black sclerotia,
especially if cultures are maintained for a few weeks.
Rhizoctonia solani is a multinucleate organism. If the
fungus has only one or two nuclei per cell it is not R. solani.
Many isolates have been collected from both North Carolina and
Mississippi, most from cucumber. For detailed information on a
particular isolate, see the isolate database.
Isolate Rs-143N was the isolate most widely used in 1992. It
was isolated from North Carolina soil in 1986. (crop and specific
Always store isolates in at least two different locations and
by two different methods.
1. Frozen on oat grains:
Autoclave equal amounts of oats and water in a 250ml flask for
1 1/2 hours on two consecutive days. Add two or three PDA disks
of Rhizoc when cool. Allow to incubate in the dark for 7-10 days.
If the flask is shaken daily it may help keep the fungus from
cementing all of the oats together. Under the hood, pour some
oat grains into a petri dish and allow them to dry for 12-24 hours.
When dry, place oat grains in sterile test tubes, seal, and put
in the freezer. The isolates can be kept this way for at least
a year. This is the preferred method of long term storage.
2. Frozen in glycerol:
Sterilize glycerol by autoclaving it for 15 minutes. Pour glycerol
into labeled vials, and in each vial place PDA disks or strips
of Rhizoc. Freeze vials. This is a published method, but there
has been some difficulty in keeping the fungus alive with this
3. Storage under water:
Make slant tubes of PDA and allow Rhizoc to colonize (3-5 days).
Cover with not more than one inch of sterile water. Store on the
shelf in the lab.
4. Storage under oil:
Same as storage under water except with mineral oil used instead
Generally the level of inoculum is reported as number of colonized
oat grains per square inch.
Source of Resistance:
PI 165509, PI 19708-, Marketmore 76
Differentials - Controls:
Supergreen, Coolgreen, PI 432855 are susceptible
Rhizoctonia sclerotia production medium:
Defrost frozen green beans and drain off the liquid. Autoclave
100 g of beans per flask for 30 minutes. Infest each flask with
three disks (2 mm in diameter) then incubate at 27°C for
3-4 weeks. Air-dry on paper towels and grind in a Waring blender
for about 1 minute. Sieving on 25- and 50- mesh sieves result
in 300-710 µm diameter sclerotia.
Van Bruggen, A.H.C. and Arneson, P.A. 1985. A Quantifiable Type
of Inoculum of Rhizoctonia solani. Plant Disease 69:966-969.
Anastomosis Group Methods for Rhizoctonia solani:
Observation by light microscopy:
1) The isolates for test are incubated for 2 to 5 days at 25°C
under darkness on PDA (Figure 1, a, b).
2) Inocula [agar disks (about 5mm in diameter) which hold
young mycelia] (Figure 1, c) are placed
on sterilized cellophane film which is mounted on medium for test
(water agar) (Figure 1, d), and incubated
for 24 to 72 hours at 25°C under darkness.
3) The hyphae which grow from the inocula contact each other
at the area where the two colonies overlapped (Figure
4) The cellophane film (about 15x15mm2) which holds the fusing
hyphae is cut off with a razor blade (Figure
1, e) and mounted on a micro slide with a forceps (Figure
5) This film is stained with a dye (e.g. 0.05% toluidine blue
0) and covered with a cover glass for light microscopic observation
(Figure 1, f, g).
6) One drop of the stain solution is put into underside of the
cover glass, and the glass is mounted on the film very carefully
to prevent destruction of the structure of fusing point (Figure
Observation of nuclei of fusing hyphae by light microscopy:
1) The preparation and transfer of inocula, and conditions for
incubation are the same as the previous method.
2) The cellophane film which is cut off from agar plate is reversed
(Figure 2, a) and mounted on the micro
slide (Figure 2, b), and the slide is coated
with melted water agar (45-50°C) to prevent the damage of
fusing hyphae (Figure 2, c).
3) The slide holding cellophane film with hyphae is fixed and
stained according to the procedure of Herr (1979).*
4) The film is removed from the slide spontaneously during hydrolyzation
with HC1. The film is reversed again in the stain solution in
a small Petri plate, and stained (Figure 2,
5) The fusing hyphae are observed after decoloring for suitable
period to observe the nuclei (Figure 2,
Hyphal fusion frequency (FF, %):
1) FF = probability of fusion between a hypha and another hypha
under a certain condition. The ability of hyphal fusion of one
isolate is, therefore, expressed as that frequency.
2) A formula to calculate the FF is as follows:
the sum of fusion points in 15 fields x 100
FF(%) = _____________________________________________
the sum of contact points in the same fields
3) The contact points include the fusing, crossing, and tip to
side and side by side hyphal adhering points. The hyphae are fused
in 3 plates in each test. One field magnified 100 time covers
2.7 mm2. Five fields per plate are observed. The test is reciprocated
at least two times.
4) Herr, L. J. 1979. Phytopathology 69: 958-961.