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The formation of secondary sporangia by Pythium ultimum: The influence of organic amendments and Pythium nunn
Soil Bid. B&hem. Vol. 20. No. 2. pp. 151-156, Printed in Great Britain. All rights reserved
1988
Copyright Q
0038-0717!88 $3.00 + 0.00 1988 Pcrgamon Press pk
THE FORMATION
OF SECONDARY SPORANGIA BY PYTHIUM ULTIMUM: THE INFLUENCE OF ORGANIC AMENDMENTS AND PYTHIUM ZVUNN T. C. PAULITZ'and R.
BAKER
Department of Plant Pathology and Weed Science, Colorado State University, Fort Collins, CO 80523, U.S.A. (Accepted 20
Seprember
1987)
Summary-Sporangia of Pythium ulfimtun were placed in polycarbonate membrane sandwiches and buried in raw Nunn sandy loam amended with increasing concentrations of bean leaves, oatmeal. glucose + L-asparagine (4: I w/w) or glucose alone. After 24 h, there was a significant negative correlation between the concentration of substrates added to the soil and the formation of secondary sporangia. Glucose + t-asparagine also inhibited the formation of secondary sporangia in an axenic system. The formation of secondary sporangia in soil amended with P. nunn (300 cfu g-‘) + 0.3% (w/w). bean-leaf or oatmeal was significantly greater than in the amended treatments without P. nwtn at 7. I4 and 21 days after amendments were added to the soil. Results suggest that competitive saprophytric activity of P. nunn rapidly depletes substrates usually utilized by P. ulfimum. This results in sporangial germination then immediate formation of survival structures (in some instance of lower inoculum potential) by the latter fungus.
Fusarium sambucinum and F. solani in natural soils confirmed the principle that survival structures (chlamydospores) are formed under nutrient-limiting conditions (Griffin and Pass, 1969; Griffin, 1976). The purpose of this research was to study the influence of glucose and organic amendments on the formation of secondary sporangia of P. uftimum in natural soil conditions. Another purpose was to investigate the effects of P. nunn (Lifshitz et al., 1984c) on the formation of secondary sporangia. P. nunn is a recently described species that is mycoparasitic on P. ultimum (Lifshitz et al., 1984a, c). P. nunn reduced the population density of P. ultimum and subsequent disease on cucumbers in infested steamed soil (Paulitz and Baker, 1987a). P. nunn also effectively competed with P. uftimum for substrates that ordinarily promote saprophytic increases in the inoculum potential of the pathogen (Paulitz and Baker, 1987b). By competing for substrates and reducing the amounts of nutrients available to P. ultimum, we hypothesized that P. nunn could influence the formation of secondary sporangia of P. ultimwn.
lNTRODUCllON
uftimum Trow is a soilborne pathogen that pre- and post-emergence damping-off of seedlings and root necrosis in many economically important plants. It is considered a primary colonizing sugar fungus (Garrett, 1970) and has little saprophytic activity on previously colonized organic matter. Because of the rapid germination of sporangia (1-3 h) and the high growth rate in soil (300pm h-‘) (Stanghellini and Hancock, 1971), P. ultimum can rapidly colonize readily utilizable food sources (such as simple carbohydrates), increase its inoculum potential and subsequently induce disease. The sporangia of P. ultimum play an important role in the life cycle and ecology of this plant pathogenic fungus. Its sporangia survive adverse conditions in the soil (Stanghellini and Hancock, 1971), germinate rapidly in response to low levels of exogenous nutrients and rapidly grow through the soil to colonize fresh substrates. When nutrients become limiting, its hyphal protoplasm retracts into primary sporangia or into newly-formed secondary sporangia. Septa are formed and the empty hyphae lyse. These survival methods permit the fungus to retain and conserve its protoplasm under starvation conditions and to maintain sporangial populations in the soil. Although some work has been done on the effects of organic amendments on the germination of sporangia of P. ultimum (Agnihotri and Vaartaja, 1967), very little is known about the influence of the nutrient status of natural soils on the formation of secondary sporangia. Kleb’s Principles suggest that there is a relationship between nutrition, growth and reproduction and that reproduction begins when nutrients are exhausted (Griffin, 1981). Studies of Pythium
induces
iMATERIAlS
AND METHODS
Soil treatments Nunn sandy loam was used in all experiments. Soil was air-dried, sieved (~4 mm) and stored for 2-3 months before use. This untreated soil will be referred to as raw soil. For treatments requiring steamed soil, the soil was moistened to approx. -0.03 MPa, exposed to aerated steam at 55°C for 1 h, held under non-sterile conditions for I wk and air-dried. For autoclaved soil treatments, the soil was autoclaved for 1 h on two consecutive days, and dried in an oven at 60°C. To revive soil fungistasis, the soil was moistened to 15% w/w (-0.01 MPa) and stored for I wk before use.
*present
address: USOA-AQS, Horticultural Crops Research Laboratory, 3420 NW Orchard Ave. Corvallis, OR 97330. U.S.A. ISI
IS2
T. C. PAULITZand R.
Sporangia of P. ultimum
An isolate of P. ultimum (Nl) used by Lifshitz et al. (1984b) was maintained on water- agar. This pathogenic isolate, obtained originally from Nunn sandy loam, produced sporangia but not oospores. It was identified as P. ultimum based on formation of antheridia and oospores when mated with a known P. ultimum isolate (J. G. Hancock, personal communication). The fungus was grown on 2% water agar in IOO-mm-dia Petri dishes for 2 months at 26’C. The Petri dishes were kept in plastic bags to prevent the medium from drying. One-quarter of the culture was removed from the Petri dish, placed in an empty Petri dish, and flooded with distilled water. Sporangia were dislodged from the surface of the agar with a paint brush and suspended in 150 ml of distilled water. Sporangia were vacuumed into a 47-mm-dia polycarbonate membrane filter having a pore size of 0.45pm (GA-6, Gelman Sciences Inc.. Ann Arbor, Michigan) by use of a Millipore filtration apparatus. Another filter was placed over the sporangia and the membrane-sporangium sandwich was cut into 5-mm? sections. The sections were buried in the soil treatments. Tests had shown that the germination of sporangia added to raw soil, either directly or in the membrane sandwich, was not affected by soil fungistasis, even when sporangia were washed three times before being applied to the membrane. Sporangia germinated on the membrane in 1-3 h in the absence of any exogenous nutrients. However, secondary sporangia that subsequently formed were sensitive to soil fungistasis. Inoculum of P. nunn An isolate of P. nunn (N3) (Lifshitz el of., 1984b) was grown in IOOml of potato-dextrose broth for I wk. The mycelial mats from five cultures were removed and added to 6 kg of Nunn sandy loam soil that had been amended with I% (w/w) ground oatmeal and autoclaved twice. The inoculum was incubated under aseptic conditions at 26’C in 32 x 39 x 19-cm plastic tubs covered with aluminum foil. After 3 wk. over 10,000 cfu g-’ of P. nunn were detected in the inoculum by use of a Pythiumselective medium (Mircetich and Kraft, 1973). Observations of the inoculum by fluorescence microscopy (Scher and Baker, 1983) revealed a mixture of sporangia and oospores. Soil amendments
Dried green bean leaves (Phuseolus vulgaris L. “Pinto”) or rolled oats (Quaker Oats Company, Chicago) were ground in a Wiley mill and sieved (< I mm). Various concentrations of bean leaves, rolled oats, glucose + asparagine (4: 1 w/w ratio), or glucose alone were mixed with Nunn sandy loam. Treated soil was placed in eight IOO-mm-dia glass Petri dishes (40 g dish-‘). The soil was moistened to 15% (w/w) (-0.01 MPa). One section of the membrane-sporangium sandwich was buried in soil in each dish and the dishes were held at 26’C. After 24 h, the membrane-sporangium sandwiches were removed, peeled apart and stained in 0.05% aqueous cotton blue.
BAKER
Testing the effect axenic conditions
of glucose and asparagine
under
Because the addition of glucose to a raw soil induces a rapid increase in the populations of soil microbes, the following experiment was performed to separate the nutritional effects of glucose from microbial interactions that result from a glucose-induced increase in biomass. Membrane-sporangium sandwiches were prepared under axenic conditions. Sections of the sandwich were floated on sterile glucose + L-asparagine (4: I ratio) solutions of various concentrations in sterile lOO-mm-dia glass Petri dishes (20 ml dish-‘). Testing the effect of P. nunn on formation sporangia
of secondary
To determine the effect of P. nunn on secondary sporangia formation, three treatments were usedraw soil, raw soil + 0.3% (w/w) bean leaves or oatmeal, and raw soil + P. nunn (300 cfu g-‘) + 0.3% (w/w) bean leaves or oatmeal. Tests had shown that P. nunn added to raw soil without an organic amendment had no effect on the formation of secondary sporangia of P. ultimum because the population density of P. nunn did not increase saprophytically (Paulitz and Baker, 1987a). Treated soil was placed in eight 6.5-cm’ plastic pots (200g pot-‘). The soil was moistened to 15% (w/w) (-0.01 MPa) and a membrane-sporangium sandwich was buried in each pot at 0.7, 14 and 21 days. Membranes were removed 24 h later, stained and sporangia counted. Pots were held at 26’C in a growth room and watered daily to maintain the matric potential above -0.01 MPa. Analyses of data
One-hundred sporangia were counted on each membrane. Data were expressed as the ratio of secondary to primary sporangia. All experiments were performed twice. Data from the two trials were pooled, because there were no significant differences in variances between the trials, as determined by an F-test of the error mean square values. Duncan’s Multiple Range Test or t-tests were used to determine statistical differences among treatment means.
RESULTS
By staining in 0.05% aqueous cotton blue, newlyformed secondary sporangia could be distinguished from primary sporangia. Secondary sporangia stained darker and had smooth walls when compared with primary sporangia, which were usually empty and collapsed (Fig. la). Secondary sporangia most commonly formed were intercalary (Fig. 1b) or at the terminal ends of hyphae (Fig. Ic). Occasionally, secondary sporangia formed adjacent to primary sporangia (Fig. Id). When primary sporangia of P. ultimum were incubated in soil containing 0.3% bean leaves or rolled oats, significantly fewer secondary sporangia were formed when compared with raw untreated soil (Table I). Treating soil with aerated steam or by autoclaving had a similar effect. In raw soil where microbial populations were reestablished by moistening and incubating the soil for 1 wk before use,
153
sporangia of Pyrhium drimum
Sxondary
b
d
Fig. I. Primary and secondary sporsngia of P. ulrirrwrr formed on poiycarbonate membranes (0.45 ktrn pore size) buried for 24 h in r;Lw so& Soil was moistened to t 5”4 (w. w) f -0.01 MPa) immediately before adding the membrane spontngia sandwiches. After 24 h, membranes were removed and stained with 0.05% aqueous cotton blue. (a) Primary and sccondnry sponlngia; (b) newly-formed intercalary secondary sporangium. Arrows show lyscd hyphae; (c) newly-formed terminnl secondary spur@;t; td) secnnktry
sporangium (xrow) formed adjacent to primary sporangium.
significantly more secondary sporangia were formed than in soil only recently moistened. A significant negative correlation (P = 0.05) between the log,, concentration of bean leaves or rolled oats added to raw soil and the formation of secondTable
I. The
amendments
effect
of soif treatments
on the formation
and organic
ofsxond;tr~
sporan-
gia by P!vrhitmr u/ri,num Mean 2
Trcatmcnt” Raw
(unrrearcdj
1.5s B
2.52 .\
Fungistatic’ Rolled Bean
oats
(3%.
WW)
leaves (3%.
w’w)
I.lIC 042C
0.72 c
Steamed”
O.lTD
Autoclavcde ratio
‘Sporangium membrane-nporsngium to the soil. are
not
Means Mean
wiches. moistened I wk
%OII
incubarcd
data
were added
thesame
letter
according
Test.
to
Means
of IWO triuls.
are
n = 16.
to 15% (W w) immedrately of membrane-sporangium
to 15%
before
treated
by
different,
Separation
the pooled
“All soil moistened to the addition ‘Soil
sandwiches followed
after
1-i h
drtermlned
significantly
Duncan’s from
ratio’
I qxangia
prior sand-
(W w) .md held dt 26’C
for
to use.
w-ith
aerated
ztc;Lm at 55 C for
at 76 C
for I wk and air-dried. ‘So11 auroxtaved for I h on IWO consecutive
days and
at 60 C.
non-sterile
I h.
conditions
dried
under
ary sporangia was found (Fig. Lb). These amcndmcnts almost completely inhibited the formation of secondary sporangia. A similar trend was found with glucose + L-asparaginc (Fig. Zc) and with glucose alone (Fig. 2d). The formation of secondary sporangia of P. uftimum on membranes incubated in sterile solutions also was inhibited by increasing concentrations of glucose and L-asparagine (4: I) (Fig. 3). At the beginning of the cxpcriments. the formation of secondary sporangia in raw soil amended with 0.3”/b bean leaves or OLltITltX~l w3s not significantly difkent (P = 0.05) from the corresponding treatments with P. nunn added at 300 cfu g-’ (Fig. 3). The formation of secondary sporangia in all amended treatments was significantly less (P = 0.05) than in the untreated soil at time 0. At days 7, 14 and 21 the sporangia ratio was greater in the amended treatments where P. nunn was added as compared with the amended trcatmcnts without P. nun. DlSCUSSlON The data in Figs 2-3 suggest that adding organic matter to soil may affect the activity of P. uirimum in at least two ways. When availabk nutrient levels are low, sporangium germination may be followed immediately by secondary spotangium formation. We observed this in treatments with no or smaller amounts of added nutrients; thcsc had higher ratios of second-
T. C. PALZITZand R. BAKER (bf
2.0t
0
Log % bean
leaf
-2.0
(w/w)
-1.5
-1.0
-0.5
0
0.5
Log % oatmeoltw/w)
2.or
2.0- (cl
(dl Glucose
Glucoseand asparogine r -0.93
--i 0
0
-2.5
-2.0
-1.5
Log % glucose
-1.0
-0.5
0
(w/w 1
Fig. 2. Effect of bean leaves, oatmeal, glucose + asparagine. or glucose alone on formation of secondary ulrimum in raw soil after 24 h. Sporangia were incubated in moist raw soil (-0.01 MPa) in polycarbonate membranes (0.45 pm pore size). Points are the average of pooled data from two trials (n = 16). One-hundred sporangia from each membrane were counted. Sporangia ratio (2,/I ‘) = No. of secondary sporangia to No. of primary sporangia.
sporangia by P.
ary sporangia formation than treatments with higher amounts of glucose, bean leaves or oatmeal. At higher nutrient concentrations, extended vegetative growth and utilization of substrates may occur with resultant increases in population density (Hancock, 1977; Johnson et al., 1981; Watson, 1970; Yarwood, 1966).
0
-2.5
-2.0
-1.5
Log % glucose
-1.0
-0.5
0
1w/w 1
Fig. 3. Effect of glucose + asparagine on formation of secondary sporangia of P. ulfimum under axenic conditions. Primary sporangia were incubated on polycarbonate membranes (0.45pm pore size) floated on sterile glucoseasparagine (1: I w,‘w) for 24 h.
Results in Fig. 4 suggest that P. nunn may compete with P. ufrimum for readily available substrates that would ordinarily promote saprophytic increases in the inoculum potential of the pathogen. Such competition may induce P. ulfimum to cease vegetative growth and convert its protoplasm into secondary sporangia which function as survival structures. Thus, when bean leaf or oatmeal were added to soil with P. IIMR, germinating sporangia did not continue vegetative growth but converted to secondary sporangia in soil that was incubated for 7 days. Without P. nunn, 14 days were required for secondary sporangium formation to reach the same level as in the raw. nonamended soil. Competition between P. nunn and P. ultimum for available substrates could be a mechanism involved in the biological control of Pythium damping-off observed by Paulitz and Baker (1987a, b). Other primary colonizing fungi influencing the colonization of organic matter by P. t&mum have been suggested as candidate antagonists. Bouhot (1981) controlled damping-off of cucumber in naturally-infested soil by adding fermented organic matter. He suggested that suppression was due to saprophytic competition by species of Mucorales. Martin and Hancock (1986) found several soils that did not support the sapro-
Secondary sporangia of Pyrhium ultimum l.Sr
0
I
I
I
I
I
7
14
21
28
Days
Fig. 4. The effect of P. nunn on formation of secondary sporangia of P. ulfimum in raw soil amended with bean leaves or oatmeal. Bean leaves or oatmeal added at 0.3% (w/w) to raw soil at start of experiment. P. nunn added at 3OOcfug-‘. Soil was moistened to 15% (-0.01 MPa) and polycarbonate membrane (0.45pm pore size) sandwiches containing sporangia of P. ultimum were buried in the soil at 0.7, I4 and 21 days. Sporangia ratio was determined after 24 h. Duncan’s mean separation test was used on the data at each sample time.
phytic growth of P. ultimum on crop residues. In these soils, P. oligundrum, another primary colonizing fungus, had greater saprophytic activity than in conducive soils. The addition of chloride increased the colonization of organic matter by P. oligandrum, but reduced the colonization and subsequent population increases of P. ultimum. The formation of secondary sporangia of P. ultimum in soil was influenced by the amount of readily available substrates (Figs 2-3). The high negative correlation between the quantity of substrates and the formation of secondary sporangia in the soil might be used as a bioassay to measure the conduciveness of a soil to the saprophytic growth of P. ultimum. Other bioassays have been developed that involve the addition of organic amendments. Lifshitz and Hancock (1981) added cotton leaves and observed an increase in population density of P. uffimum. They found that the increase in population densities was correlated with disease severity in untreated soil. Bouhot (1975a-c) devised an assay where oat flakes were mixed with the sample soil that was applied to the base of cucumber seedlings. By mixing the amended sample soil with sterile soil in various concentrations, the inoculum potential of Pythium spp in natural soils was quantified. In our system, further testing is needed to determine how well formation of secondary sporangia is correlated with disease incidence in natural soils. Finally, these experiments might offer another explanation for the observations of Lifshitz er of. (1984b). They amended a raw soil containing a low indigenous population of P. nunn with six weekly additions of ground bean leaves. After 6 wk. the population density of P. nunn increased to 4000 cfu g-t. Sporangia of P. ultimum were placed on Nucleopore membranes, buried in the soil and induced to germinate by adding 0.3% bean leaves. After 48 h, the membranes were removed and examined. In the amended treatments with P. nunn, the
155
formation of small (l&l5 pm) secondary sporangia was greater than in the raw soil; however, the formation of large (20-24 pm) secondary sporangia was inhibited. They hypothesized that some diffusible substance (enzymatic or toxic) was involved which inhibited the formation of secondary sporangia. Our results suggest that the high population density and activity of P. nunn induced by the addition of organic matter during the 6 wk so reduced the nutrient status of the soil that P. ultimum, when induced to germinate, was not able to successfully occupy substrates, reverted to the formation of small secondary sporangia, or became susceptible to germination-lysis phenomena. This shift in the frequency of size distribution of secondary sporangia, from large to small, might be explained by a reduction in the nutrient status of the soil. REFERENCES Agnihotri V. P. and Vaartaja 0. (1967) Effects of amendments, soil moisture contents. and temperatures on germination of Pyrhium sporangia under the influence of soil mycostasis. Phylopafhology 57. I 116-l 120. Bouhot D. (1975a) Recherches sur l’ecologie des champignons parasites dans le soil. V. Une technique selective d’estimation du potentiel infectieux des sol, terreaux et substrats infestes par Pyrhium sp., etudes qualitatives Annales de Phyropatholgie 7, 9-18.
Bouhot D. (1975b) Recherches sur I’ecologie des champignons parasites dans le soil. VII. Quantification de la technique d’estimation du potentiel infectieux des sols, terreaux et substrats infest&s par Pyfhium sp. Annales de Phytoparhologie 7, 147-154.
Bouhot D. (1975~) Technique selective et quantitative d’estimation du potentiel infectieux des sob, terreaux et substrats infest& par Pyfhium sp. Mode d’emploi. Annales de Phytopathologie 7. 155-158.
Bouhot D. (1981) Induction dune resistance biologique aux Pyrhium dans les sol par I’appport dune matiere organique. Soil Biology & Biochemisrry 13, 269-274. Garrett S. D. (1970) Pathogenic Root Infecting _ I Fungi. - Cambridge University Press.Griffin D. H. (1981) Fungal Physiology. Wiley, New York. Griffin G. J. (1976) Roles of low pH, carbon and inorganic nitrogen source use in chlamydospore formation by Fusarium solani. 1381-1389.
Canadian Journal
of Microbiology
22,
Griffin G. J. and Pass T. (1969) Behaviour of Fusarium roseum “Sambucinum” under carbon starvation conditions in relation to survival in soil. Canadian Journal of Microbiology 15, I l7- 126. Hancock J. G. (1977) Factors affecting soil populations of Pylhium ulrbnum in the San Joaquin Valley of California. Hilgardia 45, I07-122.
Johnson L. F.. Hsieh C. C. and Sutherland E. D. (1981) Effects of exogenous nutrients and inoculum qua&y on the virulence of Pyrhium ulrimum to cotton hypocotyls. Phyropathology 71, 629-632.
Lifshitz R. and Hancock J. G. (1981) An enrichment method to estimate potential seedling disease caused by low densities of Pyrhium ulfimum inocula in soils. Planr Disease 64, 828-829.
Lifshitz R., Duper M.. Elad Y. and Baker R. (1984a) Hyphal interactions between Pythium nunn and several soil fungi. Canadian Journal of Microbiology 30, 1482-1487.
Lifshitz R., Sneh B. and Baker R. (1984b) Soil suppressiveness to a plant pathogenic Pylhium species. Phytoparhology 74, 105&1061. Lifshitx R., Stanghellini M. and Baker R. (1984c) A new
156
T. C. PAULITZand R. BIKER
species of Pyfhium isolated from soil in Colorado. MyCofaxon 20, 373-379. Martin F. N. and Hancock J. G. (1986) Association of chemical and biological factors in soils suppressive to Pythium ultimum. Phyropathology 76, 122 I-123
I.
Mircetich S. M. and Kraft I. M. (1973) Efficiency of various selective media in determining Pyrhium population in soil. Mycopathologia et Mycologia Applicara SO, 15 I-161.
Paulitz T. C. and Baker R. (1987a) Biological control of Pythium damping-off of cucumbers with Pythium mum: population dynamics and disease suppression. Phyropathology 77, 335-340.
Paulitz T. C. and Baker R. (1987b) Biological control of Pythium damping-off of cucumbers with Pythium mmn:
influence of soil environment and organic amendments. Phytopathology 77, 341-346.
Scher F. M. and Baker R. (1983) Fluorescent microscope technique for viewing fungi in soil and its application to studies of a Fusarium suppressive soil. Soil Biology & Biochemistry IS. 7 15-7 18.
Stanghellini M. E. and Hancock J. G. (1971) The sporangium of Pyrhium ulrimum as a survival structure in soil. Phytopathology 61, 157-164.
Watson A. G. (1970) The effect of cover crops incorporated into field soil on Pythium ulrimum populations and inoculum potentials. Phytoparhology 69, 1537 (Abstr.). Yarwood C. E. (1966) Detection of Pyrhium in soil. Planr Disease Reporter SO, 791-792.
1988
Copyright Q
0038-0717!88 $3.00 + 0.00 1988 Pcrgamon Press pk
THE FORMATION
OF SECONDARY SPORANGIA BY PYTHIUM ULTIMUM: THE INFLUENCE OF ORGANIC AMENDMENTS AND PYTHIUM ZVUNN T. C. PAULITZ'and R.
BAKER
Department of Plant Pathology and Weed Science, Colorado State University, Fort Collins, CO 80523, U.S.A. (Accepted 20
Seprember
1987)
Summary-Sporangia of Pythium ulfimtun were placed in polycarbonate membrane sandwiches and buried in raw Nunn sandy loam amended with increasing concentrations of bean leaves, oatmeal. glucose + L-asparagine (4: I w/w) or glucose alone. After 24 h, there was a significant negative correlation between the concentration of substrates added to the soil and the formation of secondary sporangia. Glucose + t-asparagine also inhibited the formation of secondary sporangia in an axenic system. The formation of secondary sporangia in soil amended with P. nunn (300 cfu g-‘) + 0.3% (w/w). bean-leaf or oatmeal was significantly greater than in the amended treatments without P. nwtn at 7. I4 and 21 days after amendments were added to the soil. Results suggest that competitive saprophytric activity of P. nunn rapidly depletes substrates usually utilized by P. ulfimum. This results in sporangial germination then immediate formation of survival structures (in some instance of lower inoculum potential) by the latter fungus.
Fusarium sambucinum and F. solani in natural soils confirmed the principle that survival structures (chlamydospores) are formed under nutrient-limiting conditions (Griffin and Pass, 1969; Griffin, 1976). The purpose of this research was to study the influence of glucose and organic amendments on the formation of secondary sporangia of P. uftimum in natural soil conditions. Another purpose was to investigate the effects of P. nunn (Lifshitz et al., 1984c) on the formation of secondary sporangia. P. nunn is a recently described species that is mycoparasitic on P. ultimum (Lifshitz et al., 1984a, c). P. nunn reduced the population density of P. ultimum and subsequent disease on cucumbers in infested steamed soil (Paulitz and Baker, 1987a). P. nunn also effectively competed with P. uftimum for substrates that ordinarily promote saprophytic increases in the inoculum potential of the pathogen (Paulitz and Baker, 1987b). By competing for substrates and reducing the amounts of nutrients available to P. ultimum, we hypothesized that P. nunn could influence the formation of secondary sporangia of P. ultimwn.
lNTRODUCllON
uftimum Trow is a soilborne pathogen that pre- and post-emergence damping-off of seedlings and root necrosis in many economically important plants. It is considered a primary colonizing sugar fungus (Garrett, 1970) and has little saprophytic activity on previously colonized organic matter. Because of the rapid germination of sporangia (1-3 h) and the high growth rate in soil (300pm h-‘) (Stanghellini and Hancock, 1971), P. ultimum can rapidly colonize readily utilizable food sources (such as simple carbohydrates), increase its inoculum potential and subsequently induce disease. The sporangia of P. ultimum play an important role in the life cycle and ecology of this plant pathogenic fungus. Its sporangia survive adverse conditions in the soil (Stanghellini and Hancock, 1971), germinate rapidly in response to low levels of exogenous nutrients and rapidly grow through the soil to colonize fresh substrates. When nutrients become limiting, its hyphal protoplasm retracts into primary sporangia or into newly-formed secondary sporangia. Septa are formed and the empty hyphae lyse. These survival methods permit the fungus to retain and conserve its protoplasm under starvation conditions and to maintain sporangial populations in the soil. Although some work has been done on the effects of organic amendments on the germination of sporangia of P. ultimum (Agnihotri and Vaartaja, 1967), very little is known about the influence of the nutrient status of natural soils on the formation of secondary sporangia. Kleb’s Principles suggest that there is a relationship between nutrition, growth and reproduction and that reproduction begins when nutrients are exhausted (Griffin, 1981). Studies of Pythium
induces
iMATERIAlS
AND METHODS
Soil treatments Nunn sandy loam was used in all experiments. Soil was air-dried, sieved (~4 mm) and stored for 2-3 months before use. This untreated soil will be referred to as raw soil. For treatments requiring steamed soil, the soil was moistened to approx. -0.03 MPa, exposed to aerated steam at 55°C for 1 h, held under non-sterile conditions for I wk and air-dried. For autoclaved soil treatments, the soil was autoclaved for 1 h on two consecutive days, and dried in an oven at 60°C. To revive soil fungistasis, the soil was moistened to 15% w/w (-0.01 MPa) and stored for I wk before use.
*present
address: USOA-AQS, Horticultural Crops Research Laboratory, 3420 NW Orchard Ave. Corvallis, OR 97330. U.S.A. ISI
IS2
T. C. PAULITZand R.
Sporangia of P. ultimum
An isolate of P. ultimum (Nl) used by Lifshitz et al. (1984b) was maintained on water- agar. This pathogenic isolate, obtained originally from Nunn sandy loam, produced sporangia but not oospores. It was identified as P. ultimum based on formation of antheridia and oospores when mated with a known P. ultimum isolate (J. G. Hancock, personal communication). The fungus was grown on 2% water agar in IOO-mm-dia Petri dishes for 2 months at 26’C. The Petri dishes were kept in plastic bags to prevent the medium from drying. One-quarter of the culture was removed from the Petri dish, placed in an empty Petri dish, and flooded with distilled water. Sporangia were dislodged from the surface of the agar with a paint brush and suspended in 150 ml of distilled water. Sporangia were vacuumed into a 47-mm-dia polycarbonate membrane filter having a pore size of 0.45pm (GA-6, Gelman Sciences Inc.. Ann Arbor, Michigan) by use of a Millipore filtration apparatus. Another filter was placed over the sporangia and the membrane-sporangium sandwich was cut into 5-mm? sections. The sections were buried in the soil treatments. Tests had shown that the germination of sporangia added to raw soil, either directly or in the membrane sandwich, was not affected by soil fungistasis, even when sporangia were washed three times before being applied to the membrane. Sporangia germinated on the membrane in 1-3 h in the absence of any exogenous nutrients. However, secondary sporangia that subsequently formed were sensitive to soil fungistasis. Inoculum of P. nunn An isolate of P. nunn (N3) (Lifshitz el of., 1984b) was grown in IOOml of potato-dextrose broth for I wk. The mycelial mats from five cultures were removed and added to 6 kg of Nunn sandy loam soil that had been amended with I% (w/w) ground oatmeal and autoclaved twice. The inoculum was incubated under aseptic conditions at 26’C in 32 x 39 x 19-cm plastic tubs covered with aluminum foil. After 3 wk. over 10,000 cfu g-’ of P. nunn were detected in the inoculum by use of a Pythiumselective medium (Mircetich and Kraft, 1973). Observations of the inoculum by fluorescence microscopy (Scher and Baker, 1983) revealed a mixture of sporangia and oospores. Soil amendments
Dried green bean leaves (Phuseolus vulgaris L. “Pinto”) or rolled oats (Quaker Oats Company, Chicago) were ground in a Wiley mill and sieved (< I mm). Various concentrations of bean leaves, rolled oats, glucose + asparagine (4: 1 w/w ratio), or glucose alone were mixed with Nunn sandy loam. Treated soil was placed in eight IOO-mm-dia glass Petri dishes (40 g dish-‘). The soil was moistened to 15% (w/w) (-0.01 MPa). One section of the membrane-sporangium sandwich was buried in soil in each dish and the dishes were held at 26’C. After 24 h, the membrane-sporangium sandwiches were removed, peeled apart and stained in 0.05% aqueous cotton blue.
BAKER
Testing the effect axenic conditions
of glucose and asparagine
under
Because the addition of glucose to a raw soil induces a rapid increase in the populations of soil microbes, the following experiment was performed to separate the nutritional effects of glucose from microbial interactions that result from a glucose-induced increase in biomass. Membrane-sporangium sandwiches were prepared under axenic conditions. Sections of the sandwich were floated on sterile glucose + L-asparagine (4: I ratio) solutions of various concentrations in sterile lOO-mm-dia glass Petri dishes (20 ml dish-‘). Testing the effect of P. nunn on formation sporangia
of secondary
To determine the effect of P. nunn on secondary sporangia formation, three treatments were usedraw soil, raw soil + 0.3% (w/w) bean leaves or oatmeal, and raw soil + P. nunn (300 cfu g-‘) + 0.3% (w/w) bean leaves or oatmeal. Tests had shown that P. nunn added to raw soil without an organic amendment had no effect on the formation of secondary sporangia of P. ultimum because the population density of P. nunn did not increase saprophytically (Paulitz and Baker, 1987a). Treated soil was placed in eight 6.5-cm’ plastic pots (200g pot-‘). The soil was moistened to 15% (w/w) (-0.01 MPa) and a membrane-sporangium sandwich was buried in each pot at 0.7, 14 and 21 days. Membranes were removed 24 h later, stained and sporangia counted. Pots were held at 26’C in a growth room and watered daily to maintain the matric potential above -0.01 MPa. Analyses of data
One-hundred sporangia were counted on each membrane. Data were expressed as the ratio of secondary to primary sporangia. All experiments were performed twice. Data from the two trials were pooled, because there were no significant differences in variances between the trials, as determined by an F-test of the error mean square values. Duncan’s Multiple Range Test or t-tests were used to determine statistical differences among treatment means.
RESULTS
By staining in 0.05% aqueous cotton blue, newlyformed secondary sporangia could be distinguished from primary sporangia. Secondary sporangia stained darker and had smooth walls when compared with primary sporangia, which were usually empty and collapsed (Fig. la). Secondary sporangia most commonly formed were intercalary (Fig. 1b) or at the terminal ends of hyphae (Fig. Ic). Occasionally, secondary sporangia formed adjacent to primary sporangia (Fig. Id). When primary sporangia of P. ultimum were incubated in soil containing 0.3% bean leaves or rolled oats, significantly fewer secondary sporangia were formed when compared with raw untreated soil (Table I). Treating soil with aerated steam or by autoclaving had a similar effect. In raw soil where microbial populations were reestablished by moistening and incubating the soil for 1 wk before use,
153
sporangia of Pyrhium drimum
Sxondary
b
d
Fig. I. Primary and secondary sporsngia of P. ulrirrwrr formed on poiycarbonate membranes (0.45 ktrn pore size) buried for 24 h in r;Lw so& Soil was moistened to t 5”4 (w. w) f -0.01 MPa) immediately before adding the membrane spontngia sandwiches. After 24 h, membranes were removed and stained with 0.05% aqueous cotton blue. (a) Primary and sccondnry sponlngia; (b) newly-formed intercalary secondary sporangium. Arrows show lyscd hyphae; (c) newly-formed terminnl secondary spur@;t; td) secnnktry
sporangium (xrow) formed adjacent to primary sporangium.
significantly more secondary sporangia were formed than in soil only recently moistened. A significant negative correlation (P = 0.05) between the log,, concentration of bean leaves or rolled oats added to raw soil and the formation of secondTable
I. The
amendments
effect
of soif treatments
on the formation
and organic
ofsxond;tr~
sporan-
gia by P!vrhitmr u/ri,num Mean 2
Trcatmcnt” Raw
(unrrearcdj
1.5s B
2.52 .\
Fungistatic’ Rolled Bean
oats
(3%.
WW)
leaves (3%.
w’w)
I.lIC 042C
0.72 c
Steamed”
O.lTD
Autoclavcde ratio
‘Sporangium membrane-nporsngium to the soil. are
not
Means Mean
wiches. moistened I wk
%OII
incubarcd
data
were added
thesame
letter
according
Test.
to
Means
of IWO triuls.
are
n = 16.
to 15% (W w) immedrately of membrane-sporangium
to 15%
before
treated
by
different,
Separation
the pooled
“All soil moistened to the addition ‘Soil
sandwiches followed
after
1-i h
drtermlned
significantly
Duncan’s from
ratio’
I qxangia
prior sand-
(W w) .md held dt 26’C
for
to use.
w-ith
aerated
ztc;Lm at 55 C for
at 76 C
for I wk and air-dried. ‘So11 auroxtaved for I h on IWO consecutive
days and
at 60 C.
non-sterile
I h.
conditions
dried
under
ary sporangia was found (Fig. Lb). These amcndmcnts almost completely inhibited the formation of secondary sporangia. A similar trend was found with glucose + L-asparaginc (Fig. Zc) and with glucose alone (Fig. 2d). The formation of secondary sporangia of P. uftimum on membranes incubated in sterile solutions also was inhibited by increasing concentrations of glucose and L-asparagine (4: I) (Fig. 3). At the beginning of the cxpcriments. the formation of secondary sporangia in raw soil amended with 0.3”/b bean leaves or OLltITltX~l w3s not significantly difkent (P = 0.05) from the corresponding treatments with P. nunn added at 300 cfu g-’ (Fig. 3). The formation of secondary sporangia in all amended treatments was significantly less (P = 0.05) than in the untreated soil at time 0. At days 7, 14 and 21 the sporangia ratio was greater in the amended treatments where P. nunn was added as compared with the amended trcatmcnts without P. nun. DlSCUSSlON The data in Figs 2-3 suggest that adding organic matter to soil may affect the activity of P. uirimum in at least two ways. When availabk nutrient levels are low, sporangium germination may be followed immediately by secondary spotangium formation. We observed this in treatments with no or smaller amounts of added nutrients; thcsc had higher ratios of second-
T. C. PALZITZand R. BAKER (bf
2.0t
0
Log % bean
leaf
-2.0
(w/w)
-1.5
-1.0
-0.5
0
0.5
Log % oatmeoltw/w)
2.or
2.0- (cl
(dl Glucose
Glucoseand asparogine r -0.93
--i 0
0
-2.5
-2.0
-1.5
Log % glucose
-1.0
-0.5
0
(w/w 1
Fig. 2. Effect of bean leaves, oatmeal, glucose + asparagine. or glucose alone on formation of secondary ulrimum in raw soil after 24 h. Sporangia were incubated in moist raw soil (-0.01 MPa) in polycarbonate membranes (0.45 pm pore size). Points are the average of pooled data from two trials (n = 16). One-hundred sporangia from each membrane were counted. Sporangia ratio (2,/I ‘) = No. of secondary sporangia to No. of primary sporangia.
sporangia by P.
ary sporangia formation than treatments with higher amounts of glucose, bean leaves or oatmeal. At higher nutrient concentrations, extended vegetative growth and utilization of substrates may occur with resultant increases in population density (Hancock, 1977; Johnson et al., 1981; Watson, 1970; Yarwood, 1966).
0
-2.5
-2.0
-1.5
Log % glucose
-1.0
-0.5
0
1w/w 1
Fig. 3. Effect of glucose + asparagine on formation of secondary sporangia of P. ulfimum under axenic conditions. Primary sporangia were incubated on polycarbonate membranes (0.45pm pore size) floated on sterile glucoseasparagine (1: I w,‘w) for 24 h.
Results in Fig. 4 suggest that P. nunn may compete with P. ufrimum for readily available substrates that would ordinarily promote saprophytic increases in the inoculum potential of the pathogen. Such competition may induce P. ulfimum to cease vegetative growth and convert its protoplasm into secondary sporangia which function as survival structures. Thus, when bean leaf or oatmeal were added to soil with P. IIMR, germinating sporangia did not continue vegetative growth but converted to secondary sporangia in soil that was incubated for 7 days. Without P. nunn, 14 days were required for secondary sporangium formation to reach the same level as in the raw. nonamended soil. Competition between P. nunn and P. ultimum for available substrates could be a mechanism involved in the biological control of Pythium damping-off observed by Paulitz and Baker (1987a, b). Other primary colonizing fungi influencing the colonization of organic matter by P. t&mum have been suggested as candidate antagonists. Bouhot (1981) controlled damping-off of cucumber in naturally-infested soil by adding fermented organic matter. He suggested that suppression was due to saprophytic competition by species of Mucorales. Martin and Hancock (1986) found several soils that did not support the sapro-
Secondary sporangia of Pyrhium ultimum l.Sr
0
I
I
I
I
I
7
14
21
28
Days
Fig. 4. The effect of P. nunn on formation of secondary sporangia of P. ulfimum in raw soil amended with bean leaves or oatmeal. Bean leaves or oatmeal added at 0.3% (w/w) to raw soil at start of experiment. P. nunn added at 3OOcfug-‘. Soil was moistened to 15% (-0.01 MPa) and polycarbonate membrane (0.45pm pore size) sandwiches containing sporangia of P. ultimum were buried in the soil at 0.7, I4 and 21 days. Sporangia ratio was determined after 24 h. Duncan’s mean separation test was used on the data at each sample time.
phytic growth of P. ultimum on crop residues. In these soils, P. oligundrum, another primary colonizing fungus, had greater saprophytic activity than in conducive soils. The addition of chloride increased the colonization of organic matter by P. oligandrum, but reduced the colonization and subsequent population increases of P. ultimum. The formation of secondary sporangia of P. ultimum in soil was influenced by the amount of readily available substrates (Figs 2-3). The high negative correlation between the quantity of substrates and the formation of secondary sporangia in the soil might be used as a bioassay to measure the conduciveness of a soil to the saprophytic growth of P. ultimum. Other bioassays have been developed that involve the addition of organic amendments. Lifshitz and Hancock (1981) added cotton leaves and observed an increase in population density of P. uffimum. They found that the increase in population densities was correlated with disease severity in untreated soil. Bouhot (1975a-c) devised an assay where oat flakes were mixed with the sample soil that was applied to the base of cucumber seedlings. By mixing the amended sample soil with sterile soil in various concentrations, the inoculum potential of Pythium spp in natural soils was quantified. In our system, further testing is needed to determine how well formation of secondary sporangia is correlated with disease incidence in natural soils. Finally, these experiments might offer another explanation for the observations of Lifshitz er of. (1984b). They amended a raw soil containing a low indigenous population of P. nunn with six weekly additions of ground bean leaves. After 6 wk. the population density of P. nunn increased to 4000 cfu g-t. Sporangia of P. ultimum were placed on Nucleopore membranes, buried in the soil and induced to germinate by adding 0.3% bean leaves. After 48 h, the membranes were removed and examined. In the amended treatments with P. nunn, the
155
formation of small (l&l5 pm) secondary sporangia was greater than in the raw soil; however, the formation of large (20-24 pm) secondary sporangia was inhibited. They hypothesized that some diffusible substance (enzymatic or toxic) was involved which inhibited the formation of secondary sporangia. Our results suggest that the high population density and activity of P. nunn induced by the addition of organic matter during the 6 wk so reduced the nutrient status of the soil that P. ultimum, when induced to germinate, was not able to successfully occupy substrates, reverted to the formation of small secondary sporangia, or became susceptible to germination-lysis phenomena. This shift in the frequency of size distribution of secondary sporangia, from large to small, might be explained by a reduction in the nutrient status of the soil. REFERENCES Agnihotri V. P. and Vaartaja 0. (1967) Effects of amendments, soil moisture contents. and temperatures on germination of Pyrhium sporangia under the influence of soil mycostasis. Phylopafhology 57. I 116-l 120. Bouhot D. (1975a) Recherches sur l’ecologie des champignons parasites dans le soil. V. Une technique selective d’estimation du potentiel infectieux des sol, terreaux et substrats infestes par Pyrhium sp., etudes qualitatives Annales de Phyropatholgie 7, 9-18.
Bouhot D. (1975b) Recherches sur I’ecologie des champignons parasites dans le soil. VII. Quantification de la technique d’estimation du potentiel infectieux des sols, terreaux et substrats infest&s par Pyfhium sp. Annales de Phytoparhologie 7, 147-154.
Bouhot D. (1975~) Technique selective et quantitative d’estimation du potentiel infectieux des sob, terreaux et substrats infest& par Pyfhium sp. Mode d’emploi. Annales de Phytopathologie 7. 155-158.
Bouhot D. (1981) Induction dune resistance biologique aux Pyrhium dans les sol par I’appport dune matiere organique. Soil Biology & Biochemisrry 13, 269-274. Garrett S. D. (1970) Pathogenic Root Infecting _ I Fungi. - Cambridge University Press.Griffin D. H. (1981) Fungal Physiology. Wiley, New York. Griffin G. J. (1976) Roles of low pH, carbon and inorganic nitrogen source use in chlamydospore formation by Fusarium solani. 1381-1389.
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Griffin G. J. and Pass T. (1969) Behaviour of Fusarium roseum “Sambucinum” under carbon starvation conditions in relation to survival in soil. Canadian Journal of Microbiology 15, I l7- 126. Hancock J. G. (1977) Factors affecting soil populations of Pylhium ulrbnum in the San Joaquin Valley of California. Hilgardia 45, I07-122.
Johnson L. F.. Hsieh C. C. and Sutherland E. D. (1981) Effects of exogenous nutrients and inoculum qua&y on the virulence of Pyrhium ulrimum to cotton hypocotyls. Phyropathology 71, 629-632.
Lifshitz R. and Hancock J. G. (1981) An enrichment method to estimate potential seedling disease caused by low densities of Pyrhium ulfimum inocula in soils. Planr Disease 64, 828-829.
Lifshitz R., Duper M.. Elad Y. and Baker R. (1984a) Hyphal interactions between Pythium nunn and several soil fungi. Canadian Journal of Microbiology 30, 1482-1487.
Lifshitz R., Sneh B. and Baker R. (1984b) Soil suppressiveness to a plant pathogenic Pylhium species. Phytoparhology 74, 105&1061. Lifshitx R., Stanghellini M. and Baker R. (1984c) A new
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T. C. PAULITZand R. BIKER
species of Pyfhium isolated from soil in Colorado. MyCofaxon 20, 373-379. Martin F. N. and Hancock J. G. (1986) Association of chemical and biological factors in soils suppressive to Pythium ultimum. Phyropathology 76, 122 I-123
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Paulitz T. C. and Baker R. (1987b) Biological control of Pythium damping-off of cucumbers with Pythium mmn:
influence of soil environment and organic amendments. Phytopathology 77, 341-346.
Scher F. M. and Baker R. (1983) Fluorescent microscope technique for viewing fungi in soil and its application to studies of a Fusarium suppressive soil. Soil Biology & Biochemistry IS. 7 15-7 18.
Stanghellini M. E. and Hancock J. G. (1971) The sporangium of Pyrhium ulrimum as a survival structure in soil. Phytopathology 61, 157-164.
Watson A. G. (1970) The effect of cover crops incorporated into field soil on Pythium ulrimum populations and inoculum potentials. Phytoparhology 69, 1537 (Abstr.). Yarwood C. E. (1966) Detection of Pyrhium in soil. Planr Disease Reporter SO, 791-792.