OCCURRENCE OF MACROPHOMINAPHASEOLINA AND OTHER PATHOGENS OFEUPHORBIA LATHYRIS IN ARIZONA SOILS.

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Authors YOUNG, DEBORAH JEAN.

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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

Results . . . . . . . . . . . . . ... . Experiments with Rhizoctonia solani, Pythium

aphanidermatum, and Macrophomina phaseo1ina Phytophthora sp. Infection • .. • Root Knot Infection . . . •

Discussion .

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

Results . . . . . Soil Samples • Native Plants Pathogenicity

Discussion •

LITERATURE CITED • .

v

Page

34

34 34 36 37 37 37 38 41

42

Table

1.

2.

3.

LIST OF TABLES

Recovery of three soil-borne pathogens from representative Euphorbia 1athyris exhibiting symptoms in the field • • . . • . . . • • • .

Populations of Macrophomina phaseo1ina sclerotia in field plot A • . . • . . . • • • . . . . .

Virulence of Macrophomina phaseo1ina isolates from different vegetative communities • • • . . . • .

vi

Page

10

24

39

Figure

1.

2.

3.

LIST OF ILLUSTRATIONS

Survival of Macrophomina phaseolina sclerotia in buried Euphorbia lathyris tissue . • . • •

Survival of loose sclerotia of Macrophomina phaseolina buried in packets of soil . . .

Disease caused by Macrophomina phaseolina in three field plantings of Euphorbia lathyris

vii

Page

26

. . . . 27

.. . . . 30

ABSTRACT

Rhizoctonia solani, Pythium aphaniderrnatum, and Macrophomina

phaseolina were isolated from Euphorbia lathyris grown in fields near

Tucson, Arizona. R. solani occurred as a damping-off organism in the

fall. R. aphanidermatum infected seeds, seedlings, and mature plants

in laboratory and greenhouse tests. Although R. aphaniderrnatum was

infrequently isolated from field plants in Arizona, it was a major

pathogen of greenhouse plants growing at high temperatures in nonsterile

soil. ~. phaseolina was a major pathogen. Infection of E. lathyris

roots occurred within I mo of an October 1980 planting, but symptoms

were not significant until June. Sclerotia of this fungus ranged in

numbers from I to 246 sclerotia/g field soil. Population densities of

0.2 sc1erotium/g soil were sufficient to cause more than 90% plant

death in field plots. Some plants infected with~. phaseo1ina were

growing in an area newly cleared of native desert vegetation. Subse­

quently, M. phaseo1ina was found in uncultivated soils from four

vegetative communities in southern Arizona at elevations from 600 to

2,000 m; the fungus also was recovered from roots of several symptomless

native plants.

viii

CHAPTER 1

DISEASES OF EUPHORBIA LATHYRIS

Concern regarding the future availability of petroleum has

stimulated investigations of possible alternative or supplementary

sources of hydrocarbons. One suggested source is the gopher plant or

caper spurge, Euphorbia lathyris L. (22). Reportedly originating in

the Orient, this plant has become established as a weed in sections of

western Europe and the United States (31). It has also been cultivated

for seed in Japan and China (31).

Hydrocarbon formation by!. lathyris reportedly is increased

when plants are exposed to high levels of solar radiation (22). Also,

it has been suggested that gopher plants can be grown with less water

than many other agricultural crops (22). Therefore, E. lathyris is

being considered as a potential new crop for Arizona.

~. lathryis is not listed in the U.S.D.A. Index of Plant

Diseases in the United States (2) and, until recently, there has been

little literature on this plant. Poinsettia (E. pulcherima Willd.),

another member of this genus, is widely grown in greenhouses and is

susceptible to several soil-borne pathogens, including Rhizoctonia

solani Kuehn, Pythium spp. (37), Phytophthora sp., and Meloidogyne sp.

(2).

A multidisciplinary research team initiated agronomic experi­

mentation with E. lathyris in 1979. Plants with disease symptoms

1

grown in greenhouses at the University of Arizona and the Environmental

Research Laboratory (ERL) and in field plots in Arizona, New Mexico,

Nevada, and Utah were submitted for diagnosis from March 1979 through

August 1980. The purpose of this research was (i) to isolate and

identify potential pathogens from diseased!. lathyris plants, (ii) to

determine the pathogenicity of the isolates, and (iii) to determine

whether Meloidogyne incognita (Kofoid and lVhite) Chitwood and M.

javanica (Treub.) Chitwood, which also occur in Arizona soils (21) are

pathogens of!. lathyris.

Materials and Methods

Experiments with Rhizoctonia solani Keuhn, Pythium aphanidermatum (Edson) Fitz., and Macrophomina phaseolina (Tassi) Goid.

Isolation Procedures. Diseased~. lathyris plants of varying

ages were collected from ~1arch 1979 through August 1980 from greenhouse

and field plantings. One field planting in Arizona was newly cleared

of desert vegetation.

Sections of roots or lower stems were washed in running tap

water for 3 min, 0.5% sodium hypochlorite for 30-60 sec, and distilled

water for 1 min, then placed on 2% water agar + streptomycin sulfate at

100 ppm (WAS). The tissues were incubated in the dark at 20, 30, and

34 C for 14 days. Pure cultures, obtained from hypha 1 tips, were

subsequently maintained on V-8 agar (10% V-8 juice + 2% Bacto agar),

Difco potato-dextrose agar (PDA) , or Difco Czapek's agar.

2

3

Seed capsules were collected from dead, field-grown!. lathyris

plants in August 1981. Seeds and capsules were washed and surface

sterilized as above. They were incubated for 1 wk at 34 C on a

selective medium (mPDA) for M. phaseolina, which contained PDA + 250 ppm

streptomycin sulfate, 250 ppm penicillin G, and 100 ppm chloroneb

(1,4-dichloro-2,5-dimethoxybenzene, E. I. DuPont de Nemours & Co., Inc.,

Wilmington, DE 19898).

Growing Test Plants. E. lathyris seeds (from plants grovm in

Chico, CA) were planted in metal trays (48 x 35 x 8 em) or 10- or l5-cm

diam pots, respectively containing 3,800, 260, and 300 g of a 1:1:2

mixture of perlite, peat, and Baccto potting medium (Michigan Peat Co.,

P.O. Box 66388, Houston, TX 77006). As appropriate, the planting

medium was steam-sterilized for 12 hr before use. Seeded containers

were placed on greenhouse benches or at 21 C in growth chambers with

a l2-hr photoperiod until plant emergence.

Additionally, forty 4-mo-old!. lathyris plants, planted in

nonsterile sand, were observed for disease. These plants were grown

in a greenhouse with diurnal temperatures of 38 C (high) and 21 C (low)

with high humidity.

Inoculation Procedures. The following procedures were followed

for all three fungi. Plants 1- and 2-rno-old were grown in 10-cm-diam

pots; 100 ml of inoculum were added to each pot; 10 test and 10 control

plants were used per experiment. Six-mo-old plants were grown in 15-cm­

diam pots, to each of which was added 375 ml of inoculum; six test and

six control plants were used per experiment. Isolations were made from

plants showing symptoms. Experiments with~. solani and P.

aphanidermatum were terminated after 30 days; tests with M. phaseolina

were run for 60 days. Each experiment was done once per temperature

regime per inoculum concentration noted.

Tests with R. solani. ~. solani was grown on PDA plates for

5 days at 30 C in the dark, then comminuted for 30 sec in distilled

water. The resulting suspension of hyphal fragments and agar was used

as a soil drench. A multinucleate isolate from E. lathyris roots,

identified according to Parmeter and Hhitney (27), was used throughout

this study. In preemergence tests, 100~. lathyris seeds were planted

in a metal tray, after which 2,000 ml of inoculum containing 103

, 104 , 5 .

or 10 hypha 1 fragments per ml, was poured on the surface. Trays were

incubated at 21 C. In postemergence tests, l-mo-old plants were

inoculated by adding suspensions of 104 hyphal fragments per ml to the

soil surface. Plants were then incubated at 21, 30, and 36 C. Two-mo­

old~. lathyris were inoculated with suspensions containing 105 hypha 1

fragments per ml and incubated at 21 and 30 C.

Tests with M. phaseolina. M. phaseolina, isolated from E.

lathyris roots and identified according to Dhingra and Sinclair (13),

was used in these experiments. Pathogenicity of ~. phaseolina was

determined by two methods. Wooden toothpicks were autoclaved for 1 hr

in distilled water, then placed in a broth containing 0.5% tryptone,

0.25% yeast extract, 0.1% dextrose, and 2 parts soil extract (prepared

by autoclaving 1,000 g of field soil with 1,500 ml of distilled water

for 1 hr and then filtering the suspension into sterile flasks through

4

5

Whatman No.1 filter paper). The broth with toothpicks was autoclave-

sterilized for 20 min, after which it was inoculated with~. phaseolina

and incubated at 34 C, in the dark, for 21 days. By this time,

sclerotia had formed on the toothpicks. Stems (10-15 mID diam) of

plants to be inoculated were wiped with 2% amphyl (Sterling Drug Inc.,

Lehn and Fink Div., Montvale, NJ 07645) and pierced with a sterile

needle 3 cm above the soil line. Toothpicks with sclerotia were

inserted into the holes. Sterile toothpicks were similarly inserted

into control plants. Plants were placed in growth chambers with 14 hr

of light at 34 C and 10 hr of dark at 26 C (33) and watered only when

the top 8 cm of soil was dry.

For the second procedure, ~. phaseolina was grown for 7 days on

PDA at 34 C in the dark. A suspension of hypha! fragments and sclerotia

5 (10 Iml) was prepared as described for R. solani. Two-5 cm of the tap

and lateral roots were pruned from plants to be inoculated, after which

the remaining roots were soaked for 1-2 min in 100 ml of inoculum per

plant. Inoculated plants were then placed singly in sterile soil in

lo-c~diam pots; the remaining inoculum was poured on the soil. Tap

water was substituted for the inoculum in treating control plants.

Tests with P. aphanidermatum. An isolate of R. aphanidermatum

(25), recovered from!. lathyris roots, was grown 3 days at 34 C in the

dark on V-8 agar plates, then comminuted for 30 sec on distilled water.

This suspension contained hyphal fragments, oospores, and sporangia,

each of which was counted as one propagule. Seeds in metal trays were

exposed to the same inoculum concentrations and procedures as for R.

solani. Trays were incubated at 21 C or in the greenhouse. Plants 1-

and 2-mo-old were inoculated by drenching the soil surface with

5 suspensions ranging from 10 to 1 x 10 propagules per mI. Plants were

then placed in growth chambers at 30-33 or at 35-36 C. Six-mo-old

plants were similarly inoculated with suspensions containing 104

propagules per ml and were incubated at 37 C.

Experiments with a Phytophthora sp.

In September .1979, isolations were attempted (as described

above) from greenhouse-grown plants with stem and root rot. To

determine the pathogenicity of the Phytophthora sp. isolated, test

plants were grown as previously described for R. solani, one plant per

10-cnrdiam pot, which contained 260 g soil. Inoculum was prepared by

growing cultures on Difco cornmeal agar (CMA) for 7 days in the dark

at 27 C. Sufficient cultures, including agar, were then comminuted for

30 sec in distilled water to provide a suspension of hyphal fragments

and sporangia containing 104 propagules per mI. One hundred ml of

inoculum was used per compartment of a Speedling tray, which contained

40 g soil.

In the preemergence damping-off trial, 50 control and 50 test

seeds were planted, one per compartment of a Speedling tray; after

which the soil was drenched with inoculum. The tray was incubated in

a 20 C growth chamber with l2-hr photoperiod for 1 mo.

The susceptibility of plants 1, 2, 3, and 4 mo after the time

of seeding also was determined. ~venty-five inoculated and 25 control

plants were used of each age under greenhouse conditions. In addition,

6

10 inoculated and 10 control plants of each age were incubated at 20,

25, and 30 C in growth chambers with l2-hr photoperiods. Each test

was terminated after 1 mo.

Isolations were attempted from all inoculated plants as

previously described. Plant tissue was placed either on WAS or CMA and

incubated 4-7 days in the dark at 27 C. Pure cultures, from hyphal

tips, we~e subsequently maintained on CMA. The isolate was sent to the

University of California, Riverside for species designation.

Meloidogyne spp. Inoculations

Meloidogyne incognita (Kogoid and White) Chitwood and M.

javanica (Treub) Chitwood occur in southern Arizona soils (21).

Although root k~ot was not reported in any of the field plantings, the

susceptibility of ~. lathyris to these nematodes was studied in

greenhouse experiments.

7

E. lathxris plants, 1-, 2-, 4-, and 6-mo-old (from time of

seeding) were grown in l5-cm-diam pots in a 1:2 (v/v) mixture of

pasteurized field soil:30 mesh silica sand on greenhouse benches for

experiments with~. incognita. Second stage larvae were extracted from

Chile pepper roots (Capsicum fructescens L. cv. Anahe±m) (21). Fifty ml

of inoculum containing 5,000 larvae in a water suspension was poured on

the soil surface of each of seven pots per age group. Tomato plants

(Lycopersicum esculentum Mill. cv. Bonny Best) were similarly inoculated.

Four E. lathyris plants and two tomato plants in each age group were

used as uninoculated controls.

8

!. 1athryis plants (1- and 3-mo-o1d) were used for inoculations

with M. javanica. Ten thousand larvae (in a 50 m1 water suspension)

were added to each of 14 pots as above. Inoculated tomato plants and

uninocu1ated tomato and!. 1athyris plants were again used as controls.

To determine whether infection by Me1oidogyne spp. occurred,

plants were removed from the soil after 3 mo and observed for stunting,

chlorosis, and root galling. The amount of galling on individual plants

in a test was scored on a scale of 0 to 5 (0 = no galls, 5 = 80-100%

roots galled). Average scores were then calculated.

Results

Experiments with Rhizoctonia solani, Pythium aphaniderrnatum, and Macrophomina phaseo1ina

Naturally Infected Greenhouse Plants. Four-mo-o1d plants (180-

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.

Survival of Sclerotia Buried in the Field

Initially (January 1981) 2,000 (± 240) sc1erotia/g plant tissue

were buried in packets in plot A. Numbers declined then increased to

75,000 (± 7,000)/g colonized tissue in May. Numerous sclerotia were

observed macroscopically; plant tissues were black and partially

decomposed. During the next 8 mo, populations fluctuated but there was

an overall decline in numbers (Fig. 1). There was little change in

sc1erotia1 numbers of infested stem tissue stored in the laboratory.

After 1 yr, 1 g of tissue contained 2,200 (± 400) sclerotia.

Approximately 600 sc1erotia/g soil were recovered from the

buried packets containing loose sclerotia in soil each mo until May.

Numbers increased to 3,200 in September t~en stabilized at approxi­

mately 1,000 sc1erotia/g soil during the remainder of the test period

(Fig. 2).

90-

75-

60-

45-

30-

15 -

-..

-I'"

26

-r-

-r-

-r-

-r-

... ~; ~.i.i. o~-~~::~: ~~~ ... ~~~ ... ~ .. ~.~ ... ~~t=~~~~ ... __ ~rm:~::~ .. ~ .. ~. ___

o ..... c

Q) ~ >'c >-0'

~ ~ ~ ~ ~ ....: 0-Q)

CJ)

SAMPLING DATES

..... u o

::0 o Z

<i c Q) 0

Q J

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

coulteria (Hook. f.) Wats., Echinocloa colonum (L.) Link., Hymenothrix

wislizeni Gray, Leptochloa filiformis (Lam.) Beauv., Physalis wrightii

Gray, and Solanum elaegnifolium Cav.

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

(35), Sclerotium rolfsii (6), Sclerotium cepivorum (1, 12),

Cylindrocladium crotolariae (19, 20), and Rhizoctonia solani (8).

The clustered nature of M. phaseolina in soil may cause large variations

32

in sc1erotia1 numbers between individual samples. Therefore the use of

composite samples is important.

Dhingra and Sinclair (13) suggest that increases in sc1erotia1

populations of M. phaseo1ina occur as a consequent of root and stem

colonization. Sclerotia are formed only on colonized organic matter

(26). In my studies, the initial number of viable sclerotia in E.

1athyris stems was 2,000/g tissue. The detectable sc1erotia1 popula­

tion increased 37 fold after 5 mo of burial. The fact that, macro­

scopically, the infested stems were ,black with 'the numerous sclerotia

after being retrieved from the soil implies that an increase in actual

numbers of sclerotia occurred and that the observed population increase

was not due to enhanced germination of sclerotia. Also, increases in

the number of sclerotia in field plots particularly occurred after

infected plants were plowed under. Sclerotia are released into the

soil as tissues degrade (9). Fluctuations in sc1erotia1 populations

in both plant debris and soil were also observed by Short, Wyllie, and

Bristow (32). Smaller increases in populations of loose sclerotia in

soil may have resulted from the presence of a more limited food base.

Edmunds (15) has suggested that latent infection occurs in

charcoal rot of sorghum. M. phaseo1ina also can infect!. 1athyris

without immediately inducing symptoms. Thus, control measures aimed

at preventing infection must be applied prior to seed germination.

Since infection can occur within a few weeks following a fall planting

of E. 1athyris, chemical protection would have to be afforded from this

time until harvest the following spring-summer. With!. 1athyris,

initial infection frequently occurs within the top 7 cm of soil as the

fungus invades small roots. However, sclerotia were found throughout

the plow depth and plant infection may occur at depths of 30 cm below

the soil surface. This implies that nonsystemic chemicals would have

to be placed at these depths.

33

Asymptomatic root infection of several weeds occurred in fields

known to contain M. phaseo1ina in the soil. Therefore, weed hosts may

serve to increase the sc1erotia1 population, as suggested by Bruton

(7), who fou1d M. phaseo1ina-infecLed horse purslane (Trianthema

portu1acastrum L.) in canta10up fields. Eradications of weed hosts is

important in rotation programs designed to reduce field populations of

M. phaseo1ina.

Ghaffar and Erwin (18) found that cotton plants infected with

M. phaseo1ina did not wilt or have root rot when they were watered

regularly. Similarly, M. phaseo1ina infection of sorghum (15) and

Fusarium roseum f. sp. cu1morum infection of wheat (10) cause disease

symptoms only when the plants are stressed for water. Therefore, it

is possible that symptoms on E. 1athyris could be avoided if the plants

are irrigated frequently. Nevertheless, this is not a practical

solution to the control of charcoal rot of !. 1athyris in semi-arid

regions, such as southern Arizona, where large water requirements

would obviate production of !. 1athyris.

CHAPTER 3

OCCURRENCE OF MACROPHOMINA PHASEOLINA IN UNCULTIVATED ARIZONA SOILS

A stand of E. lathyris growing in a field newly cleared of

desert vegetation was seriously reduced by M. phaseolina (Chapter 1).

Although this fungus may be seed-borne (12), I have not been able to

recover it from seeds or capsules of~. lathyris (Chapter 1). However,

qualitative surveys indicate that M. phaseolina can occur in desert

soils in Arizona (30), Nevada (14), and New Mexico (36). Since these

studies were nonquantitative, the numbers of sclerotia of M. phaseolina

that naturally occur in selected, uncultivated soils in Arizona were

investigated.

Materials and Methods

Collection and Assay of Soil Samples

Soil samples were taken (August 1981-January 1982) from three

vegetative communities (24) which were chosen for their future

likelihood of cultivation. The communities, their elevation ranges,

and major plant associations were (i) Lower Colorado Subdivision of

the Sonoran Desert; 30-900 m; creosote (Larrea tridentata (DC.)

Coville), saltbush (Atriplex polycarpa (Torr.) Wats.), bursage

(Ambrosia aptera DC.), tamarix (Tamarix pentranda Pall), desert thorn

(Lycium macrodon Gray), mesquite (Prosopis juliflora (Swartz) DC. and

P. pubescens Benth), catclaw (Acacia greggi Gray), nolina (Nolina

34

35

bigelovii (Torr.) Wats); (ii) Arizona Upland Subdivision of the Sonoran

Desert; 150-1200 m; saguaro (Carnegiea gigantea (Engelm.) Britt. &

Rose), palo verde (Cercidium microphyllum (Torr.) Rose & Johnston),

brittlebush (Encelia farinosa Gray), ironwood (Olneya tesota Gray),

jojoba (Simrnondsia chinensis (Link) Schneid.), ocotillo (Fouquieria

splendens Engelm.); (iii) Desert Grasslands; 1000-1500 m; grama grass

(Bouteloua spp.), yucca (Yucca elata Engelm.), sotol (Dasylirion

wheeleri Wats.), bear grass (Nolina microcarpa Wats.), cholla and

prickly pear (Opuntia spp.). Approximately 40 individual soil cores

(30 cm x 30 mm) were taken from representative plant species' root

zones from each community.

Soils from Mt. W~ightson (elevation 2,800 m in the Santa Rita

Mountains, Santa Cruz Co.), Mt. Graham (elevation 3,200 m in the

Pinaleno Mountains, Graham Co.), and Mt. Lemmon (elevation 2,700 m in

the Santa Catalina Mountains, Pima Co.) also were assayed to determine

whether the distribution of M. phaseolina is limited by elevation. Each

mountain was sampled by taking one soil core approximately every 300 m

along the roads to each summit.

One site, Molino Basin (elevation 1,300 m), in the Santa

Catalina Mountains was sampled extensively. The predominant vegetation

at this oak woodland location consisted of Quercus oblongifolia Torr.,

Q. emoryi Torr., and g. arizonica Sarg. A single core was removed to

2 a depth of 30 cm from a 0.3 m grassy (Bromus spp.) bank on the north

side of a wash in July, August, October, November 1981, and January

1982. The soil in each core was thoroughly mixed, then processed and

subsampled as described in Chapter 2. In November, five separate cores

36

were taken at depths of 0-1, 8-14, 15-21, and 22-28 cm and individually

assayed. Soils in the surrounding area (1 km radius) were assayed in

January 1982.

To determine whether native plants in the wild are infected

with M. phaseo1ina the roots of 15 species (2 plants/species) were

collected from a Desert Grasslands habitat near Catalina, Arizona in

March 1982. Additionally, 10 species were collected from Molino Basin.

All plants, chosen on a random basis, appeared healthy. Both locations

had previously assayed sc1erotia1 populations (determined within 2 mo'

of plant collections) of approximately 20 sc1erotia/g soil. Root

segments were washed, disinfected, and incubated on selective medium

as described for E. 1athyris in Chapter 1.

Pathogenicity Tests

The virulence of a minimum of five isolates from soils from

each of the Lower Colorado and Arizona Upland Subdivisions, the Desert

Grasslands, and the Molino Basin site were compared to the virulence

of 10 isolates from soil in plot A at Marana (Chapter 2). The latter

site had a population of 50 sc1erotia/g soil and was planted to ~.

1athyris. Ten-mo-o1d E. 1athyris plants (2 plants/isolate) were

inoculated in the greenhouse by inserting toothpicks bearing sclerotia

of ~. phaseo1ina into their stems (see Chapter 1). Sterile toothpicks

were used on control plants. Treated plants were maintained in a

greenhouse (mean daytime temperature of 37 C) for 60 days.

Results

Soil Samples

M. phaseo1ina was recovered with the greatest frequency (34

of 37 samples) from the Desert Grasslands soils. Populations ranged

from 0.3 to 17 sc1erotia/g soil; the average was 4 sc1erotia/g soil.

37

The Lower Colorado and Arizona Upland Subdivisions yielded M. phaseo1ina

in 50% of the soil sampled. Populations also were lower, varying from

0.3 to 2.5 sc1erotia/g soil.

The Molino Basin site (0.3 m2) had 18, 20, 1, 14, and 35

sc1erotia/g soil in July, August, October, November (1981) and January

(1982), respectively. Other sites along the same channel contained

from 4 to 14 sc1erotia/g soil. M. P~~~~?~~?~ was not detected in soils

from adjacent hillsides. Each of the five soil cores (sampled within

the 0.3 m2 Molino Basin site in November) from each of the four depths

contained 4 to 35 sc1erotia/g soil.

The uppermost elevation at which M. phaseo1ina was found was

2,000 m (Mt. Wrightson and Mt. Graham). M. phaseo1ina occurred at

1,500 m on Mt. Lemmon.

Native Plants

Of the 25 plant species collected, ~. phaseo1ina was recovered

only from Lupinis sparsif10rus Benth. and Erigeron divergens Torr. &

Gray from the Desert Grasslands site and from Ipomea barbatisepa1a Gray,

Gossypium thurberi Todaro, and a Portulaca sp. from Molino Basin. Both

above and below ground, these plants had no disease symptoms;

sclerotia were not seen on any plant parts. M. phaseolina was not

38

recovered from an Ambrosia sp., Amsinckia tessellata Gray, Baileya

multiradiata Harv. & Gray, Bromus rubens L., Cryptantha decipiens

(Jones) Heller, Eriastrum diffusum (Gray) Mason, Euphorbia albomarginata

Torr. & Gray, Geranium carolineanum L., Haplopappus spinulosus

(Pursh.) DC., Haplopappus tenuisectus (Greene) Blake, Plantago

virginica L., Platystemon californicus Benth. at the Desert Grasslands

site or from an Amaranthus sp., Ambrosia confertiflora DC., Boerhaavia

erecta L., Datura meteloides DC., Euphorbia hyssopifolia L., Probuscidea

parviflora (Wooten) Woot. & StandI., and Tradescantia occidentalis

(Britton) Smyth. at the Molino Basin site.

Pathogenicity

All isolates from the uncultivated soils caused vascular

discoloration in the stems of E. lathyris plants; ~. phaseolina was

recovered from infected tissues from each plant. However, there was a

wide range in the virulence of the isolates tested. For example, plants

inoculated with Arizona Upland 1 were dead within 11 days (Table 3);

numerous sclerotia formed in root and stem tissues. Symptoms, including

blackening of stems at the point of inoculation, wilt, and leaf tip

necrosis were indistinguishable from those observed in plants inoculated

with isolates from E. lathyris or in naturally-infected field plants.

Ten isolates, however, did not cause plant death within the 60 days.

In such cases, 2-10 ern of vascular discoloration occurred both above

and below the point of inoculation. All ten isolates from field soil

caused plant death within 18 days.

39

Table 3. Virulence of Macrophomina phaseolina isolates from different vegetative communities.

a Isolates, by location Vegetative community Ph' . b at ogenJ.cJ.ty

Marana 1 agricultural 3 Marana 2 agricultural 3 Marana 3 agricultural 3 Marana 4 agricultural 3 Marana 5 agricultural 3 Marana 6 agricultural 3 Marana 7 agricultural 3 Marana 8 agricultural 3 Marana 9 agricultural 3 Marana 10 agricultural 3

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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