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OCCURRENCE OF MACROPHOMINAPHASEOLINA AND OTHER PATHOGENS OFEUPHORBIA LATHYRIS IN ARIZONA SOILS.
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University MicrOfilms
International 300 N. Zeeb Road Ann Arbor, MI48106
8227378
Young, Deborah Jean
OCCURRENCE OF MACROPHOMINA PHASEOLINA AND OTHER PATHOGENS OF EUPHORBIA LATHYRIS IN ARIZONA SOILS
The University of Arizona PH.D. 1982
University Microfilms
International 300 N. Zeeb Road, Ann Arbor,MI 48106
OCCURRENCE OF MACROPHOMINA PHASEOLINA AND OTHER PATHOGENS OF
EUPHORBIA LATHYRIS IN ARIZONA SOILS
by
Deborah Jean Young
A Dissertation Submitted to the Faculty of the
DEPARTMENT OF PLANT PATHOLOGY
In Partial Fulfillment of the Requirements For the Degree of
DOCTOR OF PHILOSOPHY
In the Graduate College
THE UNIVERSITY OF ARIZONA
1 9 8 2
THE UNIVERSITY OF ARIZONA GRADUATE COLLEGE
As members of the Final Examination Committee, we certify that we have read
the dissertation prepared by Deborah Jean Young
entitled Occurrence of Macrophomina phaseo1ina and other pathogens of
Euphorbia 1athyris in Arizona soils
and recommend that it be accepted as fulfilling the dissertation requirement
for the Degree of Doctor of Philosophy
Date
S}v4; 2?/ (f%z Date ~ r ,
Final approval and acceptance of this dissertation is contingent upon the candidate's submission of the final copy of the dissertation to the Graduate College.
I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfill~ng the dissertation requirement.
Date'
STATEMENT BY AUTHOR
This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at The University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library.
Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgment of source is'made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the head of the major department or the Dean of the Graduate College when in his judgment the proposed use of the material is in the interests of scholarship. In all other instances, however, permission must be obtained from the author.
ACKNOWLEDGMENTS
I wish to express my appreciation to Dr. S. M. Alcorn for his
guidance, encouragement, and accessibility during my graduate studies.
I would like to thank the following persons: Drs. H. E. Bloss,
R. L. Gilbertson, R. B. Hine, and M. E. Stanghellini for their interest
and suggestions; Ms. P. Rotkis for her assistance in the field and
laboratory; Mr. D. Vannickerk for his help in the laboratory. A special
thanks is extended to my husband, D. M. Chezem, who provided support
and assistance throughout these studies.
Finally, I would like to acknowledge the Office of Arid Lands
Studies and the Diamond Shamrock Corporation for financial support and
the Department of Plant Pathology for use of their facilities throughout
the course of these studies.
iii
TABLE OF CONTENTS
LIST OF TABLES . . . .
LIST OF ILLUSTRATIONS
ABSTRACT
CHAPTER
1. DISEASES OF EUPHORBIA LATHYRIS
Materials and Methods Experiments with Rhizoctonia solani Keuhn,
Pythium aphanidermatum (Edson) Fitz., and Macrophomina phaseo1ina (Tassi) Goid.
Experiments with a Phytophthora sp. Me1oidogyne spp. Inoculations
2. LATENT INFECTION OF EUPHORBIA LATHYRIS BY MACROPHOMINA PHASEOLINA AND ITS RELATIONSHIP TO POPULATIONS OF
Page
vi
vii
viii
1
2
2 6 7 8
8 13 14 14
FUNGAL PROPAGULES • • • • . • • . • 17
Materials and Methods Sc1erotia1 Populations of M. phaseolina in Soil Survival of Sclerotia in Field Soil Disease Development in the Field . Inoculum Density-Disease Incidence •
Results . • . . . . . . . . • . . . . . . Sc1erotia1 Populations of M. phaseo1ina in Survival of Sclerotia Buried in the Field Disease Development in the Field . Inoculum Density-Disease Incidence
Discussion . • . . . . . . . . . • • .
iv
Soil
17 17 18 19 21 23 23 25 28 29 29
TABLE OF CONTENTS--Continued
3. OCCURRENCE OF MACROPHOMINA PHASEOLINA IN UNCULTIVATED ARIZONA SOILS . • . . . .
Materials and Methods Collection and Assay of Soil Samples Pathogenicity Tests
240 cm in height, some in flower) growing in the greenhouse with diurnal
temperatures of 38 and 21 C (day-night) were found to be infected with
R. aphaniderrnatum. A blue-purple color developed in stems near the soil
line; lower leaves became chlorotic. Chlorosis and bluing of the stern
progressed upward. Within 4 wk of the initial symptoms, most plants
were dead or nearly so. ~. aphanidermatum was consistently isolated
from roots and from all portions of affected stems.
Naturally Infected Field Plants. Stems of ~. 1athyris up to
2-mo-o1d became discolored at the soil line and collapsed; black
lesions occurred on taproots. R. solani was the primary organism
associated with these infections; damping-off was most cornmon in
November and December 1979 in field plots (Table 1). R. solani also
caused discrete brown lesions, discoloration, and dieback of roots to
approximately 30-cm depths on older field-grown E. lathyris. This
occurred particularly in June 1979 and March, June, and July 1980
(Table 1). Affected plants had necrotic lower leaves, chlorotic upper
leaves, and were stunted.
During 1979 and 1980, most established plants thaL died were
infected with M. phaseolina. Some of these were from plots previously
2 fumigated with methyl bromide (0.97 kg/IO m ) and _f.rom plots newly
cleared of native desert ·vegetation. Lower leaves became chlorotic
and wilted, then necrotic; the necrosis proceeded acropetally. Brown
lesions occurred on roots and at the soil line on hypocotyls. The
cortex of the tap and lateral roots often sloughed off. Sclerotia
formed under the root cortex and in the stem pith. In one sampling of
30 plants in June 1980, 60% were infected at the soil line but 13.3%
had infections of roots at 15-20 cm depths.
Although M. phaseolina could be recovered from stems of field-
infected plants, none of the 100 seeds nor 100 seed capsules were
infected.
P. aphanidermatum was isolated during summer months from field-
grown plants that received frequent irrigations or rains (Table 1).
Such plants were flaccid and had foliar symptoms similar to those
caused by!. solani and~. phaseolina. However, none had the blue
color exhibited by the infected greenhouse-grown plants.
9
Table 1. Recovery of three soil-borne pathogens from representative EUEhorbia lathyris exhibiting symptoms in the field.
Plants 1.,...2-mo-olda Plants 3.,...4-mo-old Plants 5-9.,...mo-old No. b No. No.
Isolation dates plants Rsc Mpd Pae plants Rs Hp Pa plants
June 1979 10f og 0 0 14 7 5 0 0
July-Sept 1979 19 2 2 0 37 1 19 0 0
Oct-Dec 1979 37 8 2 0 8 1 0 0 2
Jan-Mar 1980 0 0 0 0 0 0 0 0 25
Apr-June 1980 14 0 2 0 8 0 4 0 107
July-Aug 1980 18 0 2 2h 16 1 6 4h 27
Totals 97 10 8 2 83 10 34 4 161
a determined from time of seeding; generally 2 wks later. Plant age emergence
b Total number of plants with symptoms from which isolations w'ere attempted.
cRs = Rhizoctonia solani.
~s = MacroEhomina Ehaseolina.
epa = Pythium aEhanidermatum.
Rs Mp
0 0
0 0
0 0
12 1
15 57
9 7
36 65
fDifferences between total plants and plants with the designated fungi represent plants from which Fusarium spp., CeEha1osEorium spp., AsEergillus sp., Dactylaria sp., Penicillium sp., RfiizoEuS sp., and/or Trichoderma sp. were isolated.
gNumber of plants with symptoms from which designated pathogen was isolated.
hCollected from areas with excess water.
Pa
0
0
0
0 1h
3h
4
I-' o
11
Inoculations ''lith R. solani. loJhen separate lots of 100 seeds
3 4 105 were planted in metal trays and inoculated with 0, 10 , 10 , or
hyphal fragments per ml, an average of 89, 84, 60, and 43 seedlings
emerged, respectively, after 19 days. R. solani was consistently
recovered from inoculated seeds and damped-off seedlings.
One-mo-old!. lathyris plants in infested soil were wilted
after 3 days at all temperatures tested. After 7 days, 100, 90, and 60%
of the plants at 21, 30, and 36 C, respectively, were dead. R. solani
was recovered only from affected plants .. Symptoms included black
lesions on the taproots and lower stems; hypocotyls frequently
collapsed. Similar results were obtained using 2-mo-old plants
incubated at 30 and 36 C.
No symptoms occurred on aerial parts of 6-mo-old plants,
although scattered small, brown lesions were present on lateral roots
of inoculated plants. R. solani was recovered from these lesions on 100
and 83% of the inoculated plants at 21 and 30 C, respectively.
Inoculations with M. phaseolina. One-mo-old E. lathyris plants,
inoculated by the drench procedure and incubated either in the green-
house or at 34 and 26 C (day-night) in a growth chamber, exhibited no
stem or foliar symptoms by 2 mo. However, brown lesions were scattered
on the taproots of 50 and 60% of the inoculated plants, respectively,
from which M. phaseolina was consistently isolated. Some darkening of
vascular bundles at the soil line of inoculated stems also occurred.
Six-mo-old plants similarly inoculated and incubated at 34 and 26 C
for 2 mo also did not show foliar symptoms. However, 50 and 40% of
12
these plants had root lesions, as described, from which H. phaseo1ina
was isolated. Control plants remained symptomless.
Separate lots of 4-mo-o1d!. 1athyris plants were inoculated by
the toothpick and soil-drench methods and then incubated at 34 and 26 C.
Symptoms developed within 7 days on all toothpick-inoculated plants,
but only after 17 days on drench-inoculated plants. Early symptoms
included wilting and chlorosis of leaves, beginning with the tips of
the lower leaves. Stems turned dark at the soil line. In the affected
stem region, vascular bundles were dark, and sclerotia formed in the
hollow center. In advanced stages of the disease, cortical tissues
were sloughed from roots; sclerotia also had formed on the inside of
these tissues.
Inoculations with P. aphanidermatum. In tray tests, seedling
emergence at 21 C
inoculations with
after 19 days was 96, 56, 23, and 7% following
345 0, 10 , 10 , and 10 propagu1es per rol. Seedling
emergence from similarly inoculated trays at greenhouse temperatures
after 19 days was 85, 44, 16, and 0%, respectively.
Plants 1-, 2-, and 6-mo-old became infected under all experi-
mental conditions. Symptoms occurred on younger plants as early as
3 days following inoculation. Tap and lateral roots darkened; black
lesions occurred on the stems at the soil line; and yellowing, then
dying of leaves proceeded acropeta11y. The apical portions of stems
became flaccid. Unlike plants inoculated with!. solani or M.
phaseolina, however, stems of greenhouse plants inoculated with P.
aphanidermatum developed a blue color at the soil line, which in some
instances progressed to the shoot tip. At 30 C, 6-mo-o1d plants died
within 14 days.
Phytophthora sp. Infection
13
Periodically, a Phytophthora sp., which readily produced
sporangia and hypha1 swellings but no oospores, was isolated from
greenhouse-grown~. 1athyris plants with stem and root rot. This fungus
formed oospores when mated with the A1 isolate of Phytophthora
parasitica Tucker (done by Zeinab E1pHama1awi, University of California,
Riverside), and therefore is considered to be R. parasitica.
R. parasitica caused preemergence damping-off; seedling
emergence was 52 and 98% from inoculated and uninocu1ated seeds
respectively. After emergence, 10 (38%) seedlings from inoculated
seeds had black lesions on ther tap roots, from which R. parasitica was
reiso1ated. This fungus was not recovered from symptomless control
plants.
Postemergence infection occurred with plants at all ages tested.
One- and 2-mo-o1d plants showed above-ground symptoms 5-10 days after
inoculation; black lesions occurred on tap roots and the base of stems;
lower leaves were chlorotic. Three- and 4-mo-o1d plants were wilted
and chlorotic in 7-14 days. Infection proceeded up the stem tissue,
from which the fungus was readily isolated.
While infection was noted at all three temperatures and in the
greenhouse, reiso1ation from plants showing symptoms was more
successful with plants grown at 20 and 25 C than with plants grown
at 30 C.
14
Root Knot Infection
All M. incognita-inoculated plants formed root galls. However,
younger plants were more severely affected, as indicated by ratings of
4, 1, and 2 for plants 2-, 4-, and 6-mo of age, respectively. Plants
inoculated at l-mo were smaller than uninoculated controls; older
inoculated plants were not stunted. Chlorosis did not appear to be an
indication of infection.
Root galls also occurred on all M. javanica-inoculated plants:
l-mo-old plants had more extensive galling (mean score of 3.4) than
did 3-mo-old plants (mean score of 2). These plants did not become
stunted or chlorotic.
Discussion
R. solani, M. phaseolina, P. aphanidermatum, K. parasitica, M.
incognita, and M. javanica were pathogenic to E. lathyris. In the
greenhouse, infections by R. solani were favored by air temperatures
of 21 C; in the field, this fungus was associated particularly with
losses of young plants (under 2 mo) following October plantings. R.
solani generally did not kill older !. lathyris plants; however, it
caused root lesions to soil depths of at least 30 cm.
M. phaseolina is a serious pathogen of established!. lathyris
plants in the vicinity of Tucson during the hot season. The recovery
of M. phaseolina from representative, naturally-infected!. lathyris
grown in the summer in western Arizona and Utah indicates that this
fungus is also a pathogen in these areas. Infection by M. phaseolina
is favored by high temperature and stress.
15
Of considerable interest and practical importance is the fact
that a number of~. lathyris plants growing in newly cleared desert
were infected with this fungus in the first season. While this fungus
has been recovered from desert soils during qualitative surveys of soil
microorganisms (14, 30, 36) to my knowledge these findings are the first
to suggest that the population of M. phaseolina in virgin soil can be
sufficient to cause disease. K. aphanidermatum also has been recovered
from nonagricultural soils in Arizona (34) and!. solani is thought to
occur in arable land worldwide (4). That these fungi are indigenous in
uncultivated lands is important because of their disease-causing
potential on "new" crops (e.g., jojoba, guayule (22), and buffalo
gourd (5), in addition to~. lathyris (22) and because such land may
need to be brought under cultivation to replace that lost to urban
sprawl.
P. aphanidermatum was isolated from occasional field plants
exposed to excess water during the hot summer. However, based on
observations and isolations from naturally-infected and inoculated
~. lathyris growing in the greenhouse, K. aphanidermatum can decay seeds
and kill plants through at least the flowering stage when environmental
conditions are optimal.
The most favorable time for planting~. lathyris in the field in
the vicinity of Tucson is either fall or spring (28). Because
chemicals may provide protection against damping-off, fall planting Of
~. lathyris seems most appropriate to achieve the most biomass by the
following season. To avoid infection by ~. phaseolina and by K.
aphanidermatum, and to conserve the amount of water needed to grow the
crop, harvest in early summer should be contemplated.
~. incognita, M. javanica, and R. parasitica have not been
recovered from diseased, field-grown!. lathyris plants in the south
western United States. Although!. lathyris appears to tolerate M.
javanica, these three pathogens are known to occur in the southwest
and have the potential to cause disease problems.
16
CHAPTER 2
LATENT INFECTION OF EUPHORBIA LATHYRIS BY MACROPHOMINA PHASEOLINA AND ITS RELATIONSHIP TO
POPULATIONS OF FUNGAL PROPAGll.ES
Macrophomina phaseo1ina was the most prevalent pathogen of
established field plantings of E. 1athyris in southern Arizona. In order
to determine methods of control, it is important to know (i) when
infection occurs and (ii) the number of propagu1es required to cause
infection. This study concerns these parameters and their re1ation-
shipes) to M. phaseo1ina infection of E. 1athyris.
Materials and Methods
Sc1erotia1 Populations of M. phaseo1ina in Soil
The number of viable sc1erotia/g of air-dried field soil (Gila
silt loam) was determined (32) in five field plots (four of which had
different cropping histories) using a composite soil sample from each
plot. Composites consisted of 10 cores, 28 cm x 30 rom each, taken at
equal intervals across each plot (described below). The composites
were air-dried for 48-72 hr at 20 C, sifted through a 2 mm sieve, and
then mixed thoroughly by transferring the soil repeatedly between two
beakers. Dried soils were stored at room temperature in lightly capped
jars. Three 1 g replications were assayed from each composite. Means
and their standard deviations were determined. To verify the
reliability of this technique, a soil sample collected and assayed on
17
January 13, 1981 by these procedures was reassayed after 1 yr of
storage.
Fluctuations in the sclerotial populations in plot A (180 x
18
13 m), previously planted to safflower, was determined by taking one
composite sample monthly (February 1981-January 1982). The sclerotial
populations of ~. phaseolina in plots B, C, and D, each of which
measured 180 x 5 m, were similarly assayed monthly from April (prior to
a spring planting of !. lathyris) through August 1981. These plots had
previously been planted with safflower, cotton, and soybeans,
respectively.
To determine the horizontal distribution of sclerotial popula
tions, 10 individual soil cores, taken along diagonal paths in February
1981 in plots A and E; were assayed separately. Plot E (180 x 25 m)
had been cropped with E. lathyris for three consecutive plantings.
Survival of Sclerotia in Field Soil
Naturally infected stems and roots of !. 1athyris were
collected from dead field plants in October 1980. The material was
air-dried and then cut into 5 cm segments. Five g of infested tissue
was placed on a 15 x 15 cm nylon screen (14 x 18 mesh) and stapled into
a rectangular packet. Each packet was attached to a wooden stake with
nylon line and buried along one edge of plot A, 15 cm below the soil
surface, in January 1981. One packet was retrieved monthly (February
1981-February 1982). The only water received by the site during this
test period was from rain. The number of viable sclerotia/g infected
tissue was determined by grinding the tissue in a Wiley mill (40 mesh)
and assaying it with the soil assay technique. Additionally, stem
tissue collected in October 1980 was stored in lightly capped glass
jars at 20 C. The population density of this material was determined
after 1 yr of storage.
The survival potential of loose sclerotia in soil was compared
to that of the sclerotia in plant residue. Ground plant residue con
taining approximately 400 (± 48) sclerotia (determined as above) was
mixed with 2.0 g of air-dried field soil from plot A, which contained
2.0 (± 1.0) sclerotia/g soil. This mixture was placed on a 10 x 10 em
square (100-200 mesh) of nylon screen, which was then placed on a
10 x 10 nylon screen (14 x 18 mesh) and stapled into a rectangular
packet. Packets were buried in plot A and retrieved as above. Soil
from the packets was assayed by the above described technique.
Disease Development in the Field
19
Seed germination of !. lathyris is optimal when soil tempera
tures are between 16 and 26 C (28). Therefore, two planting dates,
October and May, were used. Three thousand seeds were planted in plot A
in October 1980 (soil temperature ca. 26 C at planting depth). The
plot was irrigated at the time of planting, received 11.05 em of rain
during the winter months, and was furrow irrigated every other week
from May through July. There also was 13 em of rain from May through
July. Sixteen healthy-appearing plants were removed from the field in
April and in June 1981 to determine whether tap, lateral, and feeder
roots were infected at depths from 1-30 em below the soil surface.
Roots were divided into four 7-8 em segments, which were washed in
running tap water for 2 min, stirred in a solution of 0.5% sodium
hypochlorite for 5 min, and rinsed in distilled water for 2 min.
Isolations from tap, lateral, and feeder roots in each root section
were made on mPDA as described in Chapter 1. All samples were
incubated 1 wk in the dark at 34 C and observed for the presence of
M. phaseo1ina growing from the root segments.
By August 1981, about 90% of the!. 1athyris in this plot were
dead. Healthy-appearing weeds (14 species, 3 plants/species) were
removed from the surrounding area at this time and segments of their
roots plated on mPDA as above. Subsequently,!. 1athyris plants were
mowed and disked into the soil.
!. 1athyris was again seeded into plot A on 20 October 1981.
The irrigation schedule was the same as the previous year. This
planting was observe.d biweekly for root infection by!:!. phaseo1ina.
Twenty healthy-appearing plants were removed from the field at the
time of emergence (3 November) and biweekly until February 1982.
Isolations on mPDA were made as previously described.
20
Disease development also was observed in plots B, C, and D of
E. 1aLhyris. One thousand seeds were planted in each plot, which
consisted of two 180 m rows (100 cm centers), in May 1981. Seeds were
planted 5 cm deep and 8 cm apart. The plots were irrigated at the time
of planting, then at 2 wk intervals. Healthy and diseased plants were
counted at emergence, twice a month during June and July, and at the
end of August. All infected plants were removed from the plot at the
time when data were recorded. Isolations from roots of representative
infected plants (approximately 20 plants/sampling date) were made
on WAS.
Inoculum Density-Disease Incidence
Euphorbia 1athyris was seeded into flats of pasteurized soil
(1 field soi1:1 peat:1 sand, v/v) and germinated on benches in the
greenhouse. Seedlings were transplanted, one per pot (10-cm diam x
10-cm depth) 1 mo after seeding. Plants were inoculated 3 mo later.
In all procedures, test plants (approximately 40 em in height) were
first uprooted and rinsed well in tap water to remove adhering soil
particles.
Four inoculation procedures were used:
21
(i) An isolate of M. phaseo1ina from field-grown E. 1athyris
plants was cultured at 34 C in the dark for 14 days in flasks containing
50 m1 of Difco potato-dextrose broth (PDA) and then comminuted for
30 sec in distilled water. The resulting suspension of sclerotia and
hypha1 fragments was used as stock inoculum. Ptopagu1e concentrations
were determined using a Levy counting chamber (Arthur S. Thomas Co.,
Philadelphia, PA 19100).
Roots of each test plant were soaked in 100 m1 of a dilution of
this suspension for 1-2 min while control plant roots were soaked in
distilled water. Treated plants were placed singly in a mixture of
steam-sterilized field soi1:vermicu1ite (2:1, v/v) in lS-cm diam x lS-cm
depth pots. The remainder of each 100 m1 suspension was poured into
the pot containing the corresponding plant. Five plants were inoculated
with each concentration; five control plants also were used. Plants
22
were incubated for 4 wk in a growth chamber with 14 hr of light at 34 C
and 20 hr of darkness at 26 C.
In this and the following three experiments, test plants were
watered approximately every other day. Each procedure was done once.
Test plants were inoculated with approximate concentrations of 0, 1, 5,
25, and 50 sclerotia/g soil. In all experiments, isolations from
roots on both mPDA and WAS were attempted from all plants with symptoms
and from representative control plants. Roots were treated as
described previously.
(ii) This procedure was the same as the first, except that pots
were suspended in a wooden box, internally heated to ca. 34 C, which
was located in a greenhouse.
(iii) Residue from M. phaseolina-infected field plants was
ground through a 40 mesh screen in a Wiley mill. The ground material
then was added to a steam-sterilized mixture of field soil:vermiculite:
sand (3:1:1, v/v) to supply desired sclerotial concentrations, which
were determined using the soil assay technique (32). Uprooted plants
were placed singly in pots (15~cm diarn x l5-cm depth) containing a
given infested soil mixture. Five plants were inoculated at each
concentration and incubated for 6 wk in the described soil box (34 C).
Equal numbers of control plants were grown in a noninfested soil mix.
(iv) Naturally-infested field soil containing a high concentra
tion (ca. 50 sclerotia/g soil) of M. phaseolina was diluted with
pasteurized soil (1 field soil:l peat:l sand, v/v) to achieve appro
priate concentrations. All concentrations were determined using the
soil assay technique. Ten plants were inoculated, placed singly in
23
pots (lO-cm diam x 10-em depth) containing soil with a given population
of sclerotia, and incubated in the above described 34-26 C growth
chamber for 8 wk.
Results
Sclerotial Populations of M. phaseolina in Soil
The composite soil sample from plot A, collected on 13 January
1981, contained 0.7 (± 0.6) sclerotia/g soil. One year later, the same
soil (stored at room temperature in the laboratory) contained 0.6
(± 0.6) sclerotia/g soil.
The detectable number of viable sclerotia in plot A (Table 2)
generally remained low from February through September 1981, although
25% of the plants showed disease symptoms by July and there was 90%
death by the end of August. The population density increased sub
stantially after the plant residue was plowed into the soil (October)
and continued to increase in November and December, but decreased
during January 1982.
Assays of 10 individual cores in February 1981 from plot A
(previously planted to safflower) and from plot E (cropped consecu~
tively to!. lathyris) showed that sclerotia were not evenly distributed
throughout the respective fields. On a diagonal path across the field,
plot A contained 3, 0, 0, 1, 1, 0, 1, 5, 0, and 0 sclerotia/g soil in
each core. Plot E contained 10, 11, 53, 53, 88, 35, 3, 3, 6, and 1
sclerotia/g soil in each core.
The vertical distribution of sclerotia in plot A also varied.
Samples taken in April 1981 had 0.6, 2, 4.6, and 2 sclerotia/g soil at
Table 2. Populations of Macrophomina phaseo1ina sc1erotiaa in field plot Ab.
Date Means Range Comments
24
2/26/81
3/26/81
4/24/81
5/19/81
6/19/81
7/14/81
7/31/81
8/13/81
9/8/81
1.1 1.6 0-5 Planted 10/80 to E. 1athyris
10/23/81
11/3/81
12/4/81
1/9/82
1/27/82
29.5
0.5
5.0
1.3
2.6
0.3
1.0
2.3
47.0
80.0
246.0
55.3
69.6
6.3
0.4
2.6
2.3
0.5
0.5
1.0
3.2
7.2
31. 7
44.0
22.3
16.0
25-34
0-0.8
2-7
0-4
2-3
0-1
0-2
0-6
39-53
45-107
197-281
40-81
54-86
3% d o ° °d d o ~sease ~nc~ ence
14% disease incidence
25% disease incidence
90% disease incidence; plow under residue
Replanted to E. 1athyris
aNumbers of viable sc1erotia/g soil based on an assay of a composite sample (10 soil cores) taken on each date. Means are from three readings per sample.
b Plot A planted to safflower in 1979.
CStandard deviations. d Numbers of plants with disease symptoms/numbers of plants emerged x 100.
25
depths of 0-7, 8-14, 15-21, and 22-28 cm. Samples taken in the same
field, 6 wk later, had 0.7, 0.2, 0.2, and 0.1 sc1erotia/g soil at these
depths.
The inoculum densities in plots B, C, and D prior to the spring
1981 planting of E. 1athyris were 0.7 (± 0.5), 0.2 (± 0.2), and 11.5
(± 2.9) sc1erotia/g soil, respectively. A1.though a previous crop of
soybeans (plot D) resulted in more than ten times the initial sc1erotia1
population density than did crops of cotton or safflower, there were no
significant differences between the monthly population densities
throughout the growing season in each plot. Plot E (cropped consecu
tively with!. 1athyris) contained 87 (± 2) sclerotia at this time.
Fig. 1. Survival of Macrophomina phaseo1ina sclerotia in buried Euphorbia 1athyris tissue -- Columns represent .the mean number of sc1erotia/g tissue, from three replications. Bars represent standard deviations.
Fig. 2.
50- - ....
(!)
0:: W a..
(\J
o X--1 «0 -(/)
ro Ow 0::_ wo:: --10 (.)1 (/)0:: lL.-0« d z z « w ~
40-
30-
20-
10-
0 rm t 0
.ci - IV c \J..
-,...
~ 0 ~
ai~ iIi ill ~ >Q. 0
<t ~
--
IV C :::3
J
--
- ....
--
~ (.)
o (.) Q)
o
-,...
·c o
J
SAMPLING DATES
Survival of loose sclerotia of Macrophomina in packets of soil -- Columns represent the sclerotia/g soil, from three replications. standard deviations.
phaseolina buried mean number of Bars represent
27
28
Disease Development in the Field
Roots of healthy-appearing 7- and 9-mo-old~. lathyris plants
(collected from plot A in May and June 1981) were infected by H.
phaseolina. Thirty-four percent of the successful isolations were from
roots occurring 0-7 cm below the soil surface. M. phaseolina-infected
roots were found at approximately equal proportions (20%) in the soils
depths of 8-14, 15-21, and 22-28 cm. Isolations from lateral and
feeder roots frequently yielded M. phaseolina, but tap roots rarely were
found to be infected.
Asymptomatic weed hosts from which M. phaseolina was isolated
included Amaranthus palmeri Wats., Euphorbia hyssopifolia L., Euphorbia
prostrata Aiton., Ipomea coccinea L., Sonchus oleraceus L., and
Tidestromia languinosa (Nutt.) StandI. M. phaseolina was not isolated
from Amaranthus graecizans L., Ambrosia confertiflora DC., Boerhaavia
In the October 1981 planting of ~. lathyris (plot A),
progressively more roots were found to be infected with M. phaseolina
from plant emergence in November through February. By December 90% or
more of the healthy-appearing plants were infected with this fungus.
Above-ground symptoms, however, were essentially suppressed until
temperatures increased in May and June.
Infections of E. lathyris in plots B, C, and D (previously
planted to safflower, cotton, and soybean, respectively) during the
May-August 1981 growing season were attributed mainly to M. phaseolina,
based on isolations from plants showing symptoms. Although plot D
initially had a higher sc1erotia1 concentration, this did not
substantially increase the number of plants which had succumbed to
M. phaseo1ina (Fig. 3). The final incidence of disease was 90% or
greater in all three plots, regardless of the initial numbers of
sclerotia.
Inoculum Density-Disease Incidence
Occasionally, ~. 1athyris could be infected when as few as
one sc1erotium/g soil occurred. However, 100, 20, 100, and 70% of
the roots were infected when there were five sc1erotia/g soil, using
procedures i, ii, iii, and iv, respectively. At concentrations of
29
25 sr.1erotia/g soil, M. phaseo1ina was recovered from 100, 100, 70, and
80% of the inoculated plants in experiments i, ii, iii, and iv,
respectively. M. phaseo1ina successfully infected all of the plants
in all procedures when the concentration of sclerotia in the soil was
50/g. Under all experimental conditions, at least 21 days of incuba
tion were required before above-ground symptoms occurred.
Discussion
E. 1athyris is highly susceptible to ~. phaseo1ina. All field
soils tested contained sufficient sclerotia of ~. phaseo1ina to infect
~. 1athyris. Greenhouse experiments overall indicate that at least
five sc1erotia/g soil are required for a majority of the plants to be
infected. In field plots, however, an initial population of 0.2
(± 0.2) sc1erotia/g soil (plot C) caused 90% death of ~. 1athyris.
Although initial populations of M. phaseo1ina in soil varied following
90
80
70
W 60 en <l: w en 50 o
cf2. 40
30
20
10
•• ---e. plot B
.---. plot C •......• plot 0
I . I • •
.. I : I
: I •• . , .' / ./
:/
~.
..... I
I I
I I
..... ..... .....
.......... .....
• I , ,. ,: ,: ,: ,..
i j
·i :, :', :/
: / : ,
: , : ,
: / .• I ,
/ I
I .....
.. / I OL-·~==~-~~~-L-L-L~~~~~~
4 5 6 7 8 9 10 II 12 13 14 15 16 17 PLANT AGE (WKS)
Fig. 3. Disease caused by Macrophomina phaseo1ina in three field plantings of Euphorbia 1athyris -~ Fields were planted on 5/5/81. Prior to planting, plots B, C, and D contained 0.7 (± 0.5), 0.2 (± 0.2), and 11.5 (± 2.9) sc1erotia/g of soil, respectively.
30
crops of safflower, cotton, and soybean, populations were never less
than 0.2 sclerotia/g soil. The differences in sclerotial populations
after various crops may reflect the ability of ~. phaseolina to infect
and multiply on these crops. For example, the highest population
density found in agricultural soil in southern Arizona was 246
sclerotia/g soil, which occurred after planting E. lathyris for one
year (plot A). Continuous cropping of E. lathyris (plot E) also
allowed large populations of M. phaseolina to form. The soil popula
tion of M. phaseolina also was elevated following a crop of soybeans
(plot D), which are quite susceptible to this fungus (32). Maximum
populations (per g field soil) in other agricultural soils include 60
in Missouri (32) and 168 in South Carolina (11). Besides prior
cropping history, time of sampling, and isolation technique may
influence the number of sclerotia detected in soil.
The assay technique used in this study gave equivalent
numbers of sclerotia/g soil when the same soil was examined at
intervals separated by 1 yr. Thus, the technique is reproducible
and sclerotia in air-dried soil apparently do not increase or decrease
in numbers under these conditions.
31
The horizontal distribution of ~. phaseolina in the field shows
that pockets of sclerotia can occur. Several other soil-borne
pathogens have a similar distribution--e.g., Pythium aphanidermatum
Quijotoa Lower Colorado 3 Santa Rosa Lower Colorado 3 Childsc Lower Colorado 5 Puerto Blanco Lower Colorado 5 Childs Lower Colorado Ad
Pima Canyon Arizona Upland 2 Bellota Arizona Upland. 3 Kuakatch Arizona Upland 4 Gates Pass Arizona Upland A Kuakatch Arizona Upland A Bates Well Arizona Upland A Bellota Arizona Upland A San Pedro Arizona Upland A Saguaro West Arizona Upland A
Las Guijas Desert Grasslands 2 Sasabe Desert Grasslands 4 Las Guijas Desert Grasslands 5 Las Guijas Desert Grasslands 5 Sonoita Desert Grasslands 5 Santa Margarida Desert Grasslands 5 Sasabe Desert Grasslands 7 Sasabe Desert Grasslands A Rincon Desert Grasslands A Sonoita Desert Grasslands A
40
Table 3.--Continued
Isolates,a by location Vegetative community Pathogenicityb
Molino Basin 1 Oak Woodland 2 Molino Basin 2 Oak Woodland 4 Molino Basin 3 Oak Woodland 4 Molino Basin 4 Oak Woodland 4 Molino Basin 5 Oak Woodland 5 Molino Basin 6 Oak Woodland 7
aAll isolates were from soil; vascular discoloration occurred in all inoculated plants; isolates were recovered from all inoculated plants. Isolations were made between 12/81 and 3/82.
bLeast number of weeks in which either of two Euphorbia lathyris plants inoculated with~. phaseolina succumbed to disease.
cIsolates from the same location are from different soil cores. d A = plants were still alive 60 days ofter inoculation.
41
Discussion
M. phaseo1ina was recovered from virgin soils in four vegetative
communities--Lower Colorado and Arizona Upland Subdivisions of the
Sonoran Desert, Desert Grasslands, and Oak Woodland. Generally,
sc1erotial concentrations were sufficient to cause disease in E.
1athyris (Chapter 2). That inoculum in fact can be sufficient to cause
disease was demonstrated in a planting of Pinus eldarica Medw. in newly
cleared Desert Grasslands near Catalina, Arizona. In the first year of
cultivation, approximately 10% of the pines were infected with M.
phaseo1ina (Young, D. J., and Alcorn, S. M., unpublished results).
Several of the areas assayed may be used to cultivate diverse
agricultural crops including jojoba (Simmondsia chinensis (Link)
Schneid.) and guayu1e (Parthenium argentatum Gray). M. phaseo1ina can
infect more than 300 plant species (17) including guayule (29) and
jojoba (16). Some isolates from virgin soil were as pathogenic to E.
1athyris as are isolates from an agricultural soil. Thus, this fungus
might cause substantial losses if infested soils are brought into
cultivation.
On the other hand, losses might be significantly reduced or
avoided if uncultivated soils are quantitatively surveyed first for
the presence of ~. phaseo1ina. Sclerotia1 populations could be used to
predict the potential for disease in candidate crops, Such disease
forecasting has been developed for such pathogens as Sclerotium
cepivorum (1), Sclerotium rolfsii (23), and Vertici11ium dah1iae (3).
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42
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43
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44
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