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Multipest economic thresholds on snap beans

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Multipest economic thresholds on snap beans
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Snap beans
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Gadabu, Afete Divelias, 1949-
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vii, 194 leaves : ill. ; 28 cm.

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Beans ( jstor )
Crops ( jstor )
Defoliation ( jstor )
Diseases ( jstor )
Eggs ( jstor )
Fungicides ( jstor )
Juveniles ( jstor )
Leaves ( jstor )
Root knot nematodes ( jstor )
Roundworms ( jstor )
Beans -- Diseases and pests ( lcsh )
Dissertations, Academic -- Entomology and Nematology -- UF
Entomology and Nematology thesis Ph. D
Kidney bean -- Diseases and pests ( lcsh )
City of Gainesville ( local )
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bibliography ( marcgt )
non-fiction ( marcgt )

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Thesis (Ph. D.)--University of Florida, 1986.
Bibliography:
Bibliography: leaves 176-193.
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Typescript.
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Vita.
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by Afete Divelias Gadabu.

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MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS


BY

AFETE DIVELIAS GADABU




















A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY









UNIVERSITY OF FLORIDA


1986














ACKNOWLEDGEMENTS


I wish to express my sincere gratitude to many people for their support and guidance:

Dr. V. H. Waddill, my advisor and committee chairperson, for his guidance, encouragement, suggestions, and assistance throughout my Doctor of Philosophy program at the University of Florida.

Dr. J. R. Strayer, co-chairperson, for his constructive suggestions, encouragement, and providing materials for computer work whenever I needed them.

Dr. R. McSorley, for serving on my committee, encouragement,

suggestions, and invaluable assistance with statistical data analysis.

Dr. K. Pohronezny, for serving on my committee, encouragement, suggestions, and assistance with data analysis.

Dr. S. H. Kerr, for his guidance and constructive suggestions during my Ph.D. program.

I am indebted to Dr. R. D. Berger for his advice, helpful suggestions on disease progress analysis, and allowing me to use his computer programs.

The invaluable technical assistance and friendship of Nancy Shivers, Diane Putnal, W. (Hank) Dankers, Jorge Parrado, James Reynolds, John Sarvich, and Ingeborg Stough during the long and difficult field and laboratory hours are appreciated. I am also indebted to William Meyers, Joyce Francis, Jeanette Viola, and all staff at Homestead TREC for their










help and moral support. I owe a lot to my fellow students for their suggestions and sense of humor.

Sincere thanks are extended to the Malawi government and USAID for the scholarship.

Finally, the most special and loving acknowledgement is owed to my wife Abigail, for her love, encouragement, and patience throughout the program. I would like to end with my sincere thanks to Olivia and Kondwarri, my children, for putting-up with my absence.













TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS . . . . . . . . . . . ii

ABSTRACT . . . . . . . . . . * . . vi

CHAPTER I --INTRODUCTION . . . . . . . . . 1

CHAPTER II -- LITERATURE REVIEW ON DEFOLIATION, AND THE IDENTIFICATION AND CONTROL OF ROOT-KNOT NEMATODES (Meloidogyne
spp.) AND BEAN RUST (UROMYCES PHASEOLI [PERS.] WINT.) . 7
Introduction . . . . . . . . . . 7
Simulated Leaf Damage on Crop Plants . . . . 8 Nematodes Associated with Beans . ............ 10
Occurrence and Importance of Root-knot Nematodes . 10 Epidemiology and Life Cycle of Meloidogyne spp .. 13 Control of Root-knot Nematodes . . . . . . 15
Identification of Root-knot Nematodes . . . . 16 The Importance of Bean Rust . . . . . . 18
Identification and Etiology of the Pathogen . . 19 Symptoms . . . . . . . . . . . 20
Epidemiology of the Disease . . . . . . 22
Control of the Disease . . ................ 24
Interaction of Root-knot Nematodes and other Pathogens 25

CHAPTER III -- THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN
YIELD . . . . . . . . . . . . . 27
Introduction . . . . . . . . . . 27
Materials and Methods . . . . . . . . 29
Results . . . . . . . . . . . 31
Discussion . . . . . . . . . . . 49

CHAPTER IV -- THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION
ON SNAP BEANS . . . . . . . . . . . 52
Introduction . . . . . . . . . . 52
Materials and Methods . . . . . . . . 55
Results . . . . . . . . . . . 57
Discussion . . . . . . . . . . . 73

CHAPTER V -- THE EFFECT OF BEAN RUST ON SNAP BEANS . . . 77
Introduction . . . . . . . . . . 77
Materials and Methods . . . . . . . . 80
Results . . . . . . . . . . . 82
Discussion . . . . . . . . . . . 96

CHAPTER VI -- THE EFFECT OF DEFOLIATION, VAPAM, AND BEAN RUST ON
SNAP BEANS . . . . . . . . . . . . 100
Introduction . . . . . . . . . . 100
Materials and Methods . . . . . . . . 102
Results . . . . . . . . . . . 106
Discussion . . . . . . . . . . . 139











CHAPTER VII -- THE EFFECT OF INOCULATION SYSTEM ON Meloidogyne
incognita RACE ESTABLISHMENT ON BEANS . . . . . 142
Introduction . . . . . . . . . . 142
Materials and Methods . . . . . . . . 144
Results . . . . . . . . . . . 144
Discussion . . . . . . . . . . 155

CHAPTER VIII -- SUMMARY AND CONCLUSIONS . . . . . . 160

APPENDIX -- THE EFFECT OF FUNGICIDES ON SNAP BEANS . . . 164
Introduction . . . . . . . . . . 164
Materials and Methods . . . . . . . . 165
Results . . . . . . . . . . . 166
Discussion . . . . . . . . . . 174

LITERATURE CITED . . . . . . . . . . . 176

BIOGRAPHICAL SKETCH . . . . . . . . . . . 194














Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fullfillment of the Requirements for the Degree of Doctor of Philosophy





MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS By

Afete Divelias Gadabu

May, 1986

Chairman: Dr. Van H. Waddill
Major Department: Entomology and Nematology


During 1984 and 1985, a number of greenhouse and field experiments were carried out at the Tropical Research and Education Center at Homestead in Dade County, Florida, to determine the effect of manual defoliation, Meloidogyne incognita, bean rust, and various other nematodes on 'Sprite' snap beans (Phaseolus vulgaris L.). Treatments consisted of total defoliation (100%), 0%, 25%, 50%, and 75% defoliation at various plant growth stages; 0, 10, 100, 1,000, 10,000 and 100,000 M. incognita eggs and juveniles per pot, fungicide sprays which included bitertanol at 7-day intervals, mancozeb tank-mixed with sulfur at 4-5-day, 7-day, and 14-day intervals respectively, and soil fumigation with metam-sodium at 0, 47, 94, 187, 281, and 374 L/ha and a separate one at 935 L/ha. Experiments were conducted with each series of treatments as well as with combinations of two or more types of treatments simultaneously.










Manual defoliation caused the highest snap bean yield losses at

full-bloom and pod-set both in the field and the greenhouse experiment. Snap bean yield loss was observed at the 25% defoliation level in both experiments. Total defoliation resulted in the highest yield losses.

Yield was negatively correlated to Meloidogyne incognita (Kofoid

and White) Chitwood population levels when plants were grown in pots and the nematodes were used alone. Yield was also inversely related to nematode population levels when manual defoliation occurred on nematode inoculated plants. Yield loss was observed on plants grown in soil inoculated with 10 eggs and juveniles per pot.

The bean rust disease, Uromyces phaseoli (Pers.) Wint., was

manipulated by fungicide sprays. It was observed that plants with the highest disease severity gave the lowest yield whereas plants which were virtually disease free had the highest yield. Generally, fungicide sprays increased yields. In some cases yield increases were not high enough to pay for the extra cost of fungicide sprays.

Soil funigation with metam-sodium increased yields slightly. The optimum metam-sodium application rate was 187 L/ha.

Yield was affected most by the bean rust disease when defoliation, metam-sodium and the disease were used simultaneously.


vii














CHAPTER I
INTRODUCTION



Beans, Phaseolus vugaris L., are the major protein source for many people in the world, especially in developing countries (Yamaguchi, 1978). Consequently beans are considered an important crop in the tropics, subtropics, and warm temperate areas of the world (Zaumeyer and Meiners, 1975). Zaumeyer and Meiners (1975) stated that the leading world bean producers were Brazil, Mexico, and the United States of America (U.S.A.) in descending order. Beans are grown for fresh market, processing, and dry seed. In some countries the leaves are also used as a vegetable.

Snap beans are known by various names in different areas. These names include French beans, green beans, pole beans, string beans, and wax beans (Yamaguchi, 1978). Snap beans are grown in many states in the U.S. where 132,720 metric tons (mt.) were harvested for fresh-market consumption from 35,847 hectares (ha.) in 1981 (Anon., 1981). In the same crop year, 671,640 mt. of snap beans were harvested for processing from 93,324 ha. (Anon., 1981). The gross value for snap beans in 1981 was $199,282,000 in the U.S. (Anon., 1981).

Florida is the largest producer of fresh-market snap beans in the U.S., producing nearly 40% of the crop (Anon., 1972, 1982; Rose, 1975; Ware and McCollum, 1980). In the 1981-82 production year, Florida produced 60,600 mt. of bush and pole beans from 37,206 ha. (Anon., 1982). Southeast Florida is the major producing area for snap beans in










the state with the greatest production in Dade County (Anon., 1982). Gadsden, Marion and Palm Beach counties and parts of the west central area also produce some snap beans (Anon., 1982). Rose (1975), Ware and McCollum (1980), and Anon. (1982) reported that the winter demand for fresh market snap beans in population centers to the north of Florida is usually met by supplies from the southern districts of Florida.

Whereas Florida is the largest producer of fresh market snap beans, Wisconsin is the leading producer of snap beans for processing (Kobriger and Hagedorn, 1983). Michigan, New York, and Oregon also produce more snap beans for processing than Florida (Kobriger and Hagedorn, 1983; Ware and McCollum, 1980).

Snap bean production has some inherent pest problems (Anon., 1982; Rose, 1975). Fresh market snap bean yield, however, increased from 31 cwt/acre in the 1947-1952 period to 37 cwt/acre in the 1967-1972 period, in the U.S., despite these problems (Anon., 1972). This increase in yield has been ascribed to the advent of synthetic organic pesticides during and after World War II. In Florida, snap bean yields were, however, on the decline during the same period (Rose, 1975). The decline was associated mainly with adverse weather conditions (Anon., 1982). Galvez et al. (1977) and Vargas (1980) stated that insect pests, such as leafminers, leafrollers, corn earworm, Mexican bean beetle, and others cause tremendous losses in bean yield. Many of these insects feed on the leaves reducing the photosynthetic tissue of the plant. These pests have been more-or-less controlled by insecticide sprays (Acland, 1971; Iraneta and Rodrigez, 1983).

Bean rust, Uromyces phaseoli, (Pers.) Wint., is one of the most important diseases of beans in many bean producing areas of the world










(Acland, 1971; Cook, 1978; Crispin and Dongo, 1962; Iraneta and Rodrigez, 1983; Martinez, 1983; Schwartz et al., 1979; Stoezer and Omunyin, 1983; Vargas, 1980, Zaumeyer and Thomas, 1957). Other major bean disease include anthracnose (Colletotrichum lindemuthianum Sacc. and Magn.), angular leaf spot (Isariopsis griseola Sacc.), halo blight (Pseudomonas phaseolicola (Burk.) Dows.), common blight (Xanthomonas phaseoli (E. F. Smith)), and bean common mosaic virus (Acland, 1971; Allen, 1983; Martinez, 1983; Stoetzer and Omunyin, 1983). Martinez (1983) stated that root rots caused by Macrophomina phaseolina (Tassi) Goid.,, Sclerotium rolfsii Sacc., Rhizoctonia solani Kuhn, Pythium spp. and Fusarium spp. were among the most important diseases of beans.

Fifty-seven races of the bean rust fungus, Uromyces phaseoli, have been reported in the U.S. (Laundon and Waterston, 1965). The number of races of U. phaseoli is, however, not fixed, due to controversy on what constitutes a physiologic race of a pathogen (Crispin and Dongo, 1972; Davidson and Vaughan, 1963; Groth and Shrum, 1971; McMillan, 1972).

Uromyces phaseoli has been reported to reduce the translocation of photosynthetic products from the foliage to the roots and developing seeds (Daly, 1976; Livne, 1962; Montalbini, 1973; Zaki and Durbin, 1965). The reduction of photosynthetic products translocation is exacerbated by increased water loss through the damaged leaf cuticle despite a decrease in transpiration (Vargas, 1980). Water loss increases as infection becomes more severe. Infection by U. phaseoli predisposes bean plants to other pathogens such as Pythium spp., Rhizoctonia spp. P. phaseolicola, C. lindemuthianun and many others (Vargas, 1980).










Root-knot nematodes, Meloidogyne spp., also infect bean plants. Meloidogyne spp. are most prevalent in light sandy soils with good drainage and moderately warm temperatures (25-300C) (Crispin et al., 1976). Roberts and Boothroyd (1984), however, stated that M. incognita (Kofoid and White) Chitwood is more common in southern states of the U.S. and M. hapla Chitwood is commonly found in northern states. Meloidogyne spp. reportedly limit the production of beans by interfering with nitrogen fixation by Rhizobium spp. and causing root galls (Ngundo, 1977; Sharma and Guazelli, 1982; Singh et al. 1981 a). Meloidogyne incognita has been observed to predispose beans to Fusarium wilt (Singh et al., 1981 b). Severe root-knot nematode infections may lead to 50-90% yield loss (Freire and Ferraz, 1977; Ngundo, 1977; Varon and Galvez, 1974).

Control of these pests and diseases has been based on chemical

pesticides and cultural methods (Acland, 1971; Allen, 1983; Carvalho et al., 1981; Martinez, 1983; Rhoades, 1976; Robbins et al., 1972; Shorey and Hall, 1963; Stoetzer and Omunyin, 1983; Villamonte, 1965; Yoshii, 1977; Zaumeyer and Meiners, 1975). The use of resistant varieties and flooding has been part of the management strategies for bean rust and root-knot nematodes (Crispin et al., 1976; Martinez, 1983; Ngundo, 1977; Singh et al., 1981a; Vieira, 1967).

One of the problems research scientists are confronted with, in

crop pest management, is the development of multi-pest threshold levels to be used in determining the type of management strategies and the extent to which these pest complexes have to be controlled so that maximum yield is obtained with minimum disruption of the environment. Pesticides are, however, the main means of controlling pest complexes on










many crops (Allen, 1983; Stoetzer and Omunyin, 1983). Metcalf (1975) reported that repeated use of pesticides has sometimes led to a decline in acreage or yield of the crop. As a consequence of this subtle decline in crop yield due to repeated pesticide use, an integrated pest management (IPM) approach has been advocated (Huffaker and Smith, 1980; Waddill et al., 1981). IPM aims at better understanding of the significance of biological, ecological, and economic processes in the production of the crop, and the population dynamics of the pest complex, their predators and parasites, and other factors affecting the system in the field (Huffaker and Smith, 1980).

Research was undertaken during the period 1984-1985 to investigate the effects of manual defoliation, root-knot nematode population levels, and bean rust on snap bean yield. These factors were used separately, in combinations of two or more of them together, and were studied as stated in order to determine their threshold levels singly and when they occurred simultaneously.

Specific experiments were conducted to determine

1. the effect of defoliation on yield of snap beans under field and

open greenhouse conditions,

2. the effect of Meloidogyne incognita population levels on snap bean

yield,

3. the effect of bean rust, Uromyces phaseoli on snap bean yield,

4. the effect of defoliation and M. incognita population levels on

snap bean yield,

5. the effect of defoliation and several nematode genera on snap bean

yield,






6



6. the effect of defoliation, nematodes and bean rust on snap bean

yield, and

7. the effect of inoculation system on the establishment of M.

incognita in beans (Phaseolus vulganis L.).














CHAPTER II
LITERATURE REVIEW ON DEFOLIATION AND THE IDENTIFICATION AND CONTROL OF ROOT-KNOT NEMATODES (MELOIDOGYNE SPP.) AND BEAN RUST (UROMYCES PHASEOLI [PERS.] WINT.)



Introduction



Bean insect pests such as leafminers (Liriomyza spp.), bollworms

(Heliothis armigera Hbn.), and leaf rollers (Urbanus proteus L.) as well as diseases including rust (Uromyces phaseoli [Pers.] Wint.), web blight (Thanatephones cucumeris [Frank] Donk) and angular leaf spot (Isariopsis griseola Sacc.) not only destroy plant foliage but also cause physiological damage in some cases (Acland, 1971; Galvez et al., 1977). Root-knot nematodes, Meloidogyne spp., cause prolific galls on the root system of plants which may lead to the following above-ground symptoms: incipient wilting, stunted growth, and chlorotic leaves, often with burnt out edges (Agudelo, 1980). A combination of these organisms on beans usually leads to great losses in yield. Control of these pest problems has been mainly by the use of chemical pesticides (Acland, 1971; Agudelo, 1980).

This review is a summary of the problems encountered in the identification of root-knot nematodes and bean rust and the use of simulated leaf damage on beans.

Simulated Leaf Damage on Crop Plants

Bean plants are susceptible to defoliation by insects, diseases, hail, moisture stress, and mechanical injury resulting from farm










machinery. Sometimes defoliation is initiated by chemical defoliants to facilitate harvesting (McGregor et al., 1953). In most cases defoliation occurs due to pest problems and adverse environmental conditions. To reduce defoliation by pests, preventative spray programs are usually followed (Greene and Minnick, 1967). These sprays are usually applied regardless of the anticipated crop loss. Hence, it would be desirable to determine the relationship between defoliation levels and yield losses to maximize the efficiency and rationale for spray treatments.

A wide range of yield losses due to artificial defoliation has been observed on various bean cultivars (Edje and Mughogho, 1976a, 1976b; Edje et al., 1973, 1976; Garvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho, 1983; Vieira, 1981; Waddill et al., 1984). Edje et al. (1973, 1976), Edje and Mughogho (1976a, 1976b), Vieira (1981) and Waddill et al. (1984) manually defoliated indeterminate bean cultivars. Waddill et al. (1984) reported that complete defoliation when only primary leaves were present reduced yield by about 65% and repeated weekly defoliation of 50% resulted in 34% yield loss. Vieira (1981) reported that 66% leaf area removal during the flowering and pod formation stages was detrimental to yield. Galvez et al. (1977) observed that 100% defoliation at formation of the first trifoliate leaves decreased yields of the bean cultivars ICA-Guali and Porrillo-Sentetico by 34% and 49% respectively. Greene and Minnick (1967) indicated that yield reduction in snap beans begins somewhere between 33% and 50% defoliation when defoliation occurs in the prebloom and bloom stages, respectively. Hohmann and DeCarvalho (1983) reported that removal of 25, 50, 75, and 100% of the leaf area at the pod formation stage reduced










yield by 11, 20, 20, and 70% respectively. At the same levels of leaf area reduction, defoliation at initiation of flowering decreased yield by 18, 12, 19, and 55% respectively. At the formation of the third trifoliate leaves only total defoliation affected the yield.

Kalton et al. (1945) and Weber (1955) reported that 50% and 75% leaf removal in soybeans had little effect on yield when defoliation occurred at the prebloom stage. Significant yield losses were, however, observed when plants were heavily defoliated at the bloom or pod set stages (Begum and Eden, 1963, 1964; Camery and Weber, 1953; Kalton et al., 1945; McAlister and Krober, 1958). Todd and Morgan (1972) observed significant yield reduction on soybeans with 33, 67, and 100% leaf removal at 2 wk, 4 wk, and 8 wk after first bloom. Wilkerson et al. (1984) reported that all defoliations on 'Florunner' peanuts resulted in lower stem weight to length ratios and lower pod numbers and weights. It was observed that defoliation altered the normal partitioning of photosynthates between plant parts in peanuts. Wit (1983) reported that during the most sensitive period (July) in the Netherlands, 60% defoliation induced a maximum yield reduction of 35% in Brussel sprouts. He also noted that when partial defoliation was carried out 15 wk after transplanting or later, no effect on yield was observed. Douglas et al. (1981) observed a grain yield reduction of 77% in corn when complete defoliation was carried out at silking. Grain yield losses decreased with delay in defoliation toward maturity. Less severe defoliations, however, resulted in smaller reductions in yield. Generally, grain yield was tolerant of post-silking defoliation and yield losses exceeding 20% were recorded only after 67% of the leaves were removed. Defoliation action thresholds for tomato for the prebloom and postbloom










stages have been established at 30% and 50% respectively (Keularts, 1980). Extensive research on artificial defoliation effects on tomato has been conducted by various writers (Keularts, 1980; Wolk et al., 1983). The effect of leaf removal has also been studied on cotton (Ludwig, 1926), grain sorghum (Stickler and Pauli, 1961) and wheat and oats (White, 1962; Wotmack and Thurman, 1962).

Nematodes Associated with Beans

Many nematodes have been found in and around the roots of beans (Agudelo, 1980; Allen, 1983). Among the nematodes associated with beans, root-knot nematodes, Meloidogyne spp., are the most important in tropical and subtropical regions (Agudelo, 1980; Allen, 1983). Table 1 shows the nematode species associated with beans in various bean producing areas (Agudelo, 1980; Ayala and Ramirez, 1964; Bridge, 1973; Bridge et al., 1977; Castillo and Litsinger, 1978; Caveness et al., 1975, Feakin, 1973; Hague, 1980; Sinclair and Shurtleff, 1975; Singh and Farrell, 1972).

Of the four main species of Meloidogyne, M. hapla Chitwood has a more northerly distribution than M. arenaria (Neal) Chitwood, M. incognita (Kofoid and White) Chitwood, and M. javanica (Treub) Chitwood which are cosmopolitan in warmer regions (Allen, 1983; Roberts and Boothroyd, 1984). The distribution of the other nematode genera is shown on Table 1.

Occurrence and Importance of Root-knot Nematodes

The most common species of root-knot nematodes are Meloidogyne

arenaria, M. incognita, M. hapla, and M. javanica. Meloidogyne arenaria, M. incognita, and M. javanica occur worldwide warmer regions whereas M. hapla has a more northerly distribution (Agudelo, 1980; Allen, 1983;








TABLE 1. Nematodes commonly found in association with roots of beans.


Species Distribution Reference


Meloidogyne arenaria (Neal) Chitwood Cosmopolitan, tropical to warm Agudelo, 1980; Castillo temperate regions and Litsinger, 1980 M. hapla Chitwood N. Europe, Japan, U.S.A., Agudelo, 1980; Sinclair Canada, & warmer regions of and Shurtleff, 1975 Africa and Middle East

M. incognita (Kofoid & White) Chitwood Cosmopolitan, tropical to warm Agudelo, 1980 temperate regions

M. javanica (Treub) Chitwood) Cosmopolitan, tropical to warm Agudelo, 1980; Sinclair temperate regions & Shurtleff, 1975 Pratylenchus brachyurus (Godfrey) Filipjev Cosmopolitan Agudelo, 1980; Bridge, 1973

Aphelenchoides spp. Nigeria Agudelo, 1980; Bridge et al., 1977

Rotylenchulus reniformis Linford & Oliveira W. Africa, U.S.A., Indonesia, Agudelo, 1980; Ayala & Philippines Ramirez, 1964; Singh & Farrell, 1972

Helicotylenchus spp. Cosmopolitan Agudelo, 1980; Bridge, 1973; Hague, 1980

Criconemella spp. Widespread Agudelo, 1980; Feakin, 1973








TABLE 1. Continued.


Species Distribution Reference


Belonolaimus spp. Southeastern U.S.A. Agudelo, 1980; Feakin, 1973; Sinclair & Shurtleff, 1975

Trichodorus spp. Widespread Agudelo, 1980; Feakin, 1973

Xiphinema spp. Widespread Agudelo, 1980; Caveness et al., 1975; Feakin, 1973










Roberts and Boothroyd, 1984). Meloidogyne arenaria is rarely encountered in association with beans. Meloidogyne incognita and M. javanica frequently occur simultaneously on beans (Ngundo, 1977; Saka, 1982; Santacruz, 1983; Singh et al., 1981a). The most serious threat to bean production is M. incognita (Ngundo, 1977; Singh et al., 1981a; Sharma and Guazelli, 1982). These nematodes may cause yield losses of 50 to 90% during severe infections (Freire and Ferraz, 1977; Varon and Galvez, 1974).

The limitation on bean by root-knot nematodes has been reported to be due to extensive root-galling and interference with nitrogen fixation by Rhizobium spp. (Agudelo, 1980), as well as with water and nutrient uptake. Root-knot nematode infestations often lead to abbreviated root systems (Agudelo, 1980; Franklin, 1978). Above-ground symptoms of root-knot infections include incipient wilting, chlorotic above-ground plant parts, and stunted growth (Agudelo, 1980).

Epidemiology and Life Cycle of Meloidogyne spp.

Meloidogyne spp. are most abundant in light sandy soils with

adequate drainage and temperatures of 250-30*C (Crispin et al., 1976). Root-knot nematodes are spread by irrigation and flood waters, by vegetative propagation of plant parts in soil contaminated with eggs and juveniles, which adhere to farm implements, animals, and man (Agudelo, 1980; Caveness, 1967; Crispin et al., 1976; Steadman et al., 1975; Vieira, 1967; Villamonte, 1965; Walker, 1965). The length of survival of root-knot nematodes in the soil varies with the stage of development, soil type, moisture, temperature, soil aeration, and length of the fallow period (Navarro and Barriga, 1970; Villamonte, 1965; Walker, 1965).










The life cycle of Meloidogyne spp. has several developmental stages (Taylor and Sasser, 1978). The adult female lays eggs in a gelatinous matrix. The first-stage juvenile develops and molts within the egg. What emerges from the egg is actually the second-stage juvenile, hence the general belief that root-knot juveniles grow between a series of three molts into adult males and females (Agudelo, 1980). Root-knot nematode eggs are oval or ellipsoidal and may be concave on the side. They measure 30-52 x 67-128 pm (Thorne, 1961). These eggs are usually protected from dehydration by a gelatinous matrix secreted by the female (Franklin, 1978; Taylor and Sasser, 1978).

The juvenile stages are vermiform, have a stylet about 10 pm long and have an overall length of 375-500 pm and a width of 15 pm (Robbins et al., 1972; Taylor and Sasser, 1978). Males are cylindroid and measure 0.03-0.36 x 1.2-1.5 mm. The males lack a bursa. Adult females are pyriform and usually pearly white (visible in roots without magnification). The females measure 0.27-0.75 x 0.40 x 1.30 mm and have a soft cuticle (Franklin, 1978; Taylor, 1965; Walker, 1965). The life cycle of root-knot nematodes may take 17-57 days, depending on the soil temperature and the host plant (Tyler, 1933; Taylor and Sasser, 1978).

Infection by and pathogenesis of Meloidogyne spp. are affected by plant age, plant susceptibility, population size and environmental factors (Brodie and Dukes, 1972; Gilvonio and Ravines, 1971; Nemec and Morrison, 1972; Sosa Moss and Torres, 1973). Second stage juveniles of Meloidogyne spp. enter the plant root system within 2 days after inoculation and migrate inter and/or intracellularly through the cortex into the stele (Dropkin, 1980; Ngundo and Taylor, 1975 b). The juvenile inserts its head into the vascular system of the root to obtain










nutrients from the plant. Plant cells in the vicinity of the nematode juvenile increase both in number and size (hyperplasia and hypertrophy), causing the characteristic giant cells (synctia) (Dropkin, 1980; Taylor and Sasser, 1978). The giant cells usually form near the juvenile's head by the fusion and enlargement of plant cells in response to nematode feeding. These giant cells eventually become apparent in the form of galls on the root system. Injury to plant root systems usually becomes apparent 10 days after infection. Five to six weeks after infection, epidermal cells of the roots collapse after females have deposited eggs near the outer root surface (Ngundo and Taylor, 1975a).

Control of Root-knot Nematodes, Meloidogyne spp.

The economic importance of plant-parasitic nematodes is commonly assessed by the use of soil fumigants (Mountain, 1965). Usually, an inverse relationship between yield and nematode numbers is expected (Sasser et al., 1968). The relationship between yield and nematode counts is not always inverse (Robbins et al., 1972). In many beanproducing regions, nematicides are extensively used on a preventative basis (Agudelo, 1980). The world farming community has many nematicides available depending on supply and legal registration. These nematicides include dicholoropropene-dichloropropane (DD), ethylene dibromide (EDB), phenamiphos, methyl bromide, aldicarb, metam-sodium, and DBCP (Jimenez, 1976; Parisi et al., 1972; Rhoades, 1976; Sosa Moss and Wrihs, 1973). In these operations, no attempt is made to eradicate nematodes (Thomason and McKenry, 1975). These nematicide applications are aimed at reducing the nematode populations by 80-90% in the upper 40-60 cm of the soil and are considered adequate to provide economic control (Thomason and McKenry, 1975).










Crop rotation has been used to reduce nematode numbers in bean fields (Agudelo, 1980). Beans are planted once every 2 or 4 years in rotation with a crop such as corn, which is not particularly susceptible to many nematodes parasitic on beans. Cover crops such as marigold (Tagetes minuta), rattle box (Crotalaria spectablilis), or hairy indigo (Indigofera hirsuta) have been used for this purpose (Eguiguren et al., 1975; Navarro and Barriga, 1970; Rhoades, 1976; Zaumeyer and Thomas, 1957). Other cultural practices used to reduce nematode numbers include long fallow periods, deep plowing, and flooding (Crispin et al., 1976; Vieira, 1967).

There are many bean cultivars resistant to M. incognita (Blazey et al., 1964; Christie, 1959; Fassuliotis et al., 1970; Hartman, 1968; Ngundo and Taylor, 1974; Rhoades, 1976; Varon and Galvaz, 1974; Wester et al., 1958). In some cases resistance to M. incognita is broken by simultaneous infection of M. incognita and M. javanica (Ngundo, 1977). Ngundo (1977) reported that seven bean lines were resistant to M. incognita and M. javanica when they occurred simultaneously.

Identification of Root-knot Nematodes

Maggenti (1981) and Taylor and Sasser (1978) state that root-knot nematodes were first described by Berneley in England in 1855. M. incognita was studied, independently, in the U.S. by Neal and Atkinson in 1889 (Maggenti, 1981). Maggenti (1981) reported that Neal indicated that root-knot nematodes occurred in Florida before 1805. During these early studies, Meloidogyne spp. were described under the species names Heterodera marioni or H. radicicola (Maggenti, 1981). Chitwood and Chitwood (1950), as a result of their work on the taxonomy of root-knot nematodes, placed them under the genus Meloidogyne. Chitwood and Chitwood recognized five species of Meloidogyne and one subspecies.










Esser et al. (1976), however, recognized 35 species in this genus. Dickson (unpublished) reported that more than 50 species of Meloidogyne were identified. The number of species in this genus fluctuates due to various identification procedures used and discovery of new species each year.

Root-knot nematode speciation is based on perineal patterns, the distance between stylet knobs and the dorsal esophageal gland opening, the second-stage juvenile morphology, chromosome number, electrophoresis, and host range (Maggenti, 1981; Taylor and Sasser, 1978). Host differentials are also used to separate races of the same species (Taylor and Sasser, 1978).

Meloidogyne incognita is the most widely distributed species of

root-knot nematode, comprising 52% of a world collection (40'N to 330S) in areas where annual temperatures are normally within the 18-300C range (Taylor and Sasser, 1978). This species has four host races as follows: race 1 does not infect 'Deltapine' 16 cotton, 'NC95' tobacco, and 'Florunner' peanuts; race 2 does not infect 'Deltapine' cotton, and 'Florunner' peanuts; race 3 does not infect 'NC95' tobacco and 'Florunner' peanuts; and race 4 does not infect 'Florunner' peanuts only. All four races infect 'California Wonder' pepper, 'Charleston Grey' watermelon and 'Rutgers' tomato (Taylor and Sasser, 1978). Meloidogyne incognita has a very extensive host range and frequently coexists with M. javanica (Dickson, unpublished; Santacruz, 1983). Meloidogyne javanica is the second most widely distributed species, forming 31% of a world collection (Taylor and Sasser, 1978). Meloidogyne javanica has no known host races but exhibits variation in chromosome numbers. M. hapla and M. arenaria comprised 8 and 7% of a world










collection respectively (Taylor and Sasser, 1978). These two species are known to have 2 host races each (Dickson, unpublished).

The Importance of Bean Rust

Bean rust is known to occur whenever beans are grown (Vargas, 1980). Bean rust is the most important bean disease in Central and South America (Augustin et al., 1972; Crispin and Dongo, 1962; Makram et al., 1973; Zaumeyer and Meiners, 1975). Bean rust has been reported to reduce the yield of snap beans in New Zealand, Egypt, and Australia (Ballantyne, 1974; Makram et al., 1973; Yen and Brien, 1960). Yields of dry beans have been lowered by infections of bean rust in Kenya and Turkey (Mukumya, 1974; Rudolph and Baykal, 1978).

Although the occurrence of bean rust was characterized as sporadic in the U.S. (Harter et al., 1935), Vargas (1980) reported yield losses as high as 40-80% in the U.S. are caused by this disease. Brazil is reported to incur losses of 35-50% due to bean rust infections (Vargas, 1980).

Bean rust was reported to be responsible for the bulk of the yield losses in Navy beans in Michigan (Andersen, 1975). The disease was reported to be troublesome in snap bean fields of North Dakota and Minnesota (Meiners, 1977). Zaumeyer and Meiners (1975) reported that prior to 1945, bean rust was a major disease in irrigated fields in Colorado, western Nebraska, Wyoming and Montana. In their review, Zaumeyer and Meiners (1975) reported bean rust was no longer a problem in those areas, although it was still occasionally important in fall snap bean crops along the Atlantic seaboard and in winter crops grown in Florida.










Identification and Etiology of the Pathogen

Bean rust is caused by the fungus Uromyces phaseoli [Pers.] Wint. (= U. phaseoli typica (Reben) Wint. = U. appendiculatus [Pers.] Unger). The fungus was first described in Germany in 1795 (Cook, 1978). The pathogen is an autoecious macrocyclic rust fungus (Kolmer et al., 1984; Laundon and Waterston, 1965). This pathogen is parasitic on the leguminous genera Dolichos, Phaseolus, and Vigna (Laundon and Waterston, 1965). The fungus is transmitted generally through wind-borne uredospores. Uredospores are rusty orange in color and ellipsoidal to obvoidal in shape, 20-30 x 20-26 pm in measurement (Laundon and Waterston, 1965).

The aecial and pycnial stages are rare in U. phaseoli (Harter et

al., 1935). Harter et al. (1935) did not observe any aecia or pycnia of this fungus in the field. In Queensland, Ogle and Johnson (1974) did not report seeing mature aecia or pycnia of U. phaseoli. The absence of aecia on U. phaseoli under field conditions has also been reported in Maryland (Marcus, 1952). The aecial stage of this fungus has, however, been observed and reported in New York and Virginia (Fromme and Wingard, 1921; Jones, 1960). Both aecia and pycnia were reported to occur on field grown beans in North Dakota by Venette et al. (1978).

Fromme and Wingard (1918) and Harter et al. (1935) reported that telia form under unfavorable conditions for the development of the pathogen such as low temperatures, decreased host vigor, and increased host resistance. The propensity of the pathogen to form telia was suspected to be an innate character of the fungal isolate (Harter et al., 1935). It has been reported that teliospores do not occur in Florida (Townsend, 1939). Consequently, Townsend (1939) suggested










uredospores blown in from the north serve as primary inocula in Florida. Later, Kidney (1980) observed telia in Alachua and Dade counties, contrary to Townsend's findings.

Uredospores overwinter in infected crop debris and trellis poles (Davison and Vaughan, 1963). These overwintered uredospores are known to initiate the disease cycle in the next growing season in Oregon and Maryland (Davison and Vaughan, 1963; Marcus, 1952). In Florida, colder temperatures than those normal for that state are apparently required for the uredospores to be viable for relatively long periods (Davison and Vaughan, 1963).

Disease development is frequently initiated by uredospores under natural conditions. The uredospore produces a germ tube upon germination. An appressorium which molds itself into the stomatal ledge is formed when the germ tube gets in touch with the stoma (Mendgen, 1973). An infection peg develops, from the appressorium and pushes the guard cells apart until the fungal cytoplasm is transferred into the substomatal vesicle (Vargas, 1980). Enzymes, lipid bodies, and glycogen particles are contained in the vesicle (Mendgen, 1973). The fungus develops infection hyphae and haustoria as it proceeds inter-cellularly in the host tissue (Mendgen and Heitefuss, 1975; Vargas, 1980).

The bean rust fungus may complete its life cycle within 10-15 days after inoculation (Yarwood, 1961). Uredospores are released passively from pustules and disseminated by farm implements, insects, animals, and wind currents (Yarwood, 1961; Zaumeyer and Thomas, 1957).

Symptoms

Apparently, bean rust is primarily a foliar disease which occasionally occurs on pods, stems and branches (Fromme and Wingard, 1918; Laundon and Waterston, 1965; Vargas, 1980).










The uredia (uredinia) are the major diagnostic sign of the pathogen (Fromme and Wingard, 1918). In a susceptible reaction, symptoms of bean rust first appear on the lower leaf surface as minute, whitish, slightly raised spots about 5-6 days after infection. These spots enlarge to form mature reddish-brown pustules which rupture the epidermis and obtain a diameter of 1-2 mm, 10-12 days after infection (Vargas, 1980). The uredia reach a diameter of 5 mm by the 14th day after infection (Rey and Lozano, 1961). The size of the uredia varies depending on environmental conditions as well as the host. The uredia may appear powdery due to uredospores protruding from them (Fromme and Wingard, 1918). Uredia often appear on both leaf surfaces. The uredia are frequently surrounded by chlorotic halos and eventually by rings of secondary and tertiary sori (Zaumeyer and Thomas, 1957). As infection progresses, the leaf becomes debilitated and the chlorotic areas surrounding pustules coalesce, while tissue ramified by the fungus remains green, apparently, as a result of starch accumulation (Wang, 1961). Severe rust infections may cause premature abscission. Bean rust rarely causes small, circular necrotic lesions on pods (Kucharek and Simone, 1980).

Rust infection has been reported to cause increased respiratory

rates in susceptible hosts (Daly et al., 1961). Twenty-four hours after infection starch accumulation decreases sharply in the vicinity of the fleck. Accumulation of starch at the perimeter of the lesion, however, resumes 96-120 hr after infection. The quantity of starch in this area decreases at the time the fungus sporulates (Schipper and Mirocha, 1969). Rust infections cause leakage of ions, amino acids, and sugars in susceptible plant leaves (Hoppe and Heitefuss, 1974a). Hoppe and Heitefuss (1974b) presented evidence that rust infection caused damage










to chloroplast membranes. Raggi (1978) reported decreased photosynthetic rates in rust-infected plants.

Epidemiology of the Disease

Fromme and Wingard (1921) reported that since rust rarely attacks pods directly, resulting losses are insidious and difficult to assess. Yield losses are, however, more likely to be severe when plants are infected during the prebloom and flowering stages of development (Almeida et al., 1977; Costa, 1972; Crispin et al., 1976; Nasser, 1976; Wimalajeewa and Thavam, 1973; Yoshii and Galvez, 1975). Early infection of some bean varieties can lead to almost complete crop loss in some seasons (Fromme and Wingard, 1921; Howland and MaCartney, 1966; Townsend, 1939). Townsend (1939) indicated that total loss of the entire crop due to rust has occurred in Florida.

The variability in the prevalence of bean rust seasonally and

geographically is partly due to environmental conditions (Augustin et al., 1972; Gonzalez, 1976; Harter and Zaumeyer, 1941; Harter et al., 1935; Schein, 1961). Infection by U. phaseoli is favored by prolonged periods (8-18 hours) of at least 95% RH and moderate temperatures (15-270C) (Augustin et al., 1972; Gonzalez, 1976, Schein, 1961). The optimum temperature for uredospore germination was reported to be 14.5*C whereas the optimum temperature for infection was 17*C (Harter et al., 1935). Crispin et al. (1976), Schein (1961), and Zaumeyer and Thomas (1957), however, reported that any temperatures below 150 may retard fungal development. Day length and light intensity are also important factors for the development of the bean rust fungus (Harter and Zaumeyer, 1941).










Fifty-seven races of U. phaseoli have been identified in the U.S. (Stavely, 1984). Laundon and Waterston (1965) reported 35 races of U. phaseoli. Races 1 and 2 were identified from specimens obtained from Washington, D.C. and California (Harter et al., 1935). Twenty races of U. phaseoli were differentiated according to their reaction on seven bean cultivars (Barter and Zaumeyer, 1941). Fisher (1952) isolated 10 races from the Rocky Mountain states and Maryland. Race 31 was identified from New Mexico and race 32 from Maryland (Sappenfield, 1954; Zaumeyer, 1960). Hikida (1961, 1962) isolated and identified races 33 and 34 in Oregon. Race 35 was isolated by McMillan (1972) from the bean cultivar Dade, which was bred for resistance to previously known races of U. phaseoli in Florida. McMillan (1972) reported that races 1, 2, 5, 9, 10, 11, and 35 occur in Florida.

There is tremendous variability in the reaction pattern of U.

phaseoli races (Groth and Shrum, 1977). In many areas where several races occur, there is usually one race which is greatly predominating (Fisher, 1952).

Uromyces phaseoli races have also been identified outside the U.S. Thirty-one races have been identified in Mexico (Crispin and Dongo, 1962), 10 races in Colombia (Zuniga and Victoria, 1975), 46 races in Brazil (Pereira and Chaves, 1977), 12 races in Puerto Rico (Lopez, 1976), 4 races in Nicaragua, 5 races in Honduras (Vargas, 1969, 1970), 7 races in Guatemala (Vargas, 1970), 5 races in El Salvador (Vargas, 1971), 4 races in Peru (Guerra and Dongo, 1973), 11 races in Costa Rica, 11 races in Australia, and 8 races in East Africa (Ballantyne, 1974, 1975; Fisher, 1952; Ogle and Johnson, 1974).










Control of the Disease

Cultural control measures of this disease include crop rotation and removal of old plant debris (Vieira, 1967). Reduced plant density and planting date adjustment for specific production areas may reduce rust incidence (Vargas, 1980). Resistant varieties of beans have been used for the control of rust (Augustine et al., 1972; Ballantyne, 1974; Coyne and Schuster, 1975; Crispin et al., 1976; Madriz and Vargas, 1975; Meiners, 1974; Rivera, 1977; Rodriguez, 1976).

Fungicidal sprays are usually recommended to help manage bean rust. Since bean rust reduces yields more severely when infection occurs before flowering than when infection is initiated after flowering, fungicidal sprays are, therefore, more effective if applied during early plant development (Yoshii and Galvez, 1975). Of the older fungicides, sulfur dusts have given relatively good control (Ballantyne, 1975; Harter et al., 1935; Zaumeyer and Thomas, 1957). Sulfur is usually applied at the rate of 25-30 kg/ha every 7-10 days. Generally, protectant fungicides fail in areas where rainfall is frequent because deposits are washed off too soon. Other preventative chemicals applied at schedule similar to that of sulfur are chlorothalonil (225 g/ha), maneb (4-5 kg/ha), and mancozeb (3-4 kg/ha) (Costa, 1972; Crispin et al., 1976; Hilty and Mullins, 1975; Vieira, 1967; Wimalajeewa and Thavam, 1973).

Plantvax (Oxycarboxin) is somewhat therapeutic when sprayed 20 to 40 days after planting at the rate of 1.8-2.5 kg/ha (Costa, 1972; Frenhani et al., 1971; Hilty and Mullins, 1975). McMillan et al. (1982) reported effective control of bean rust when bean plants were sprayed weekly with bitertanol or triadimefon. These fungicides are not registered for use on beans at this time. While certain fungicides are










effective against bean rust, their use is regulated by their estimated cost-effectiveness. Thus, Issa and Arruda (1964) cited by Vargas (1980) concluded that chemical control of bean rust was not economically practical in Brazil. This conclusion may apply to most tropical beanproducing areas. The use of fungicides in highly mechanized agricultural systems, such as the U.S., may be economically feasible provided registration conditions are met.

Interaction of Root-knot Nematodes and Other Pathogens

Increased incidence of plant diseases has been reported to be

associated with the presence of root-knot nematodes (Brodie and Cooper, 1964; Carter, 1975a,b; Cauquil and Shepherd, 1970; Minton et al., 1975; Morrell and Bloom, 1981; Norton, 1960; Reynolds and Hanson, 1957; Schuster, 1959; Thomason et al., 1959; Van Gundy et al., 1977). Carter (1975), Cauquil and Shepherd (1970), Norton (1960), Reynolds and Hanson (1957), and White (1962) reported increase incidence of soreshin of cotton (Rhizoctonia solani Kuhn), root rot (Pythium debaryanum Hesse) and Fusarium wilt (Fusarium oxysporum Schlecht) when Meloidogyne incognita (Kofoid and White) Chitwood was present. Increased incidence of southern blight, Sclerotium rolfsii Sacc., was observed in soybeans infested with root-knot nematodes (Minton et al., 1975). The interaction of M. incognita and bacterial wilt (Corynebacterium fluccumfaciens (Hedges) Dows.) was reported on beans by Schuster (1959). Van Gundy et al. (1977) reported the enhancement of the development of R. solani in the presence of exudates from galls caused by M. incognita. Morrell and Bloom (1981) reported a significant increase in the percentage of Fusarium wilt occurrence and vessel infection at 21*C in the presence of M. incognita in tomato. Meloidogyne-Fusarium synergism was also observed










in cowpea (Thomason et al., 1959). Interaction of root-knot nematodes is not limited to nematode-fungus or nematode bacterium complexes. Meloidogyne incognita has also been reported to interact with other nematodes (Thomas and Clark, 1983). Thus, Thomas and Clark (1983) reported that early season counts of M. incognita and Rotylenchulus reniformis Linford and Oliveira were positively correlated with later counts of the same nematode, but negative correlations were found between early M. incognita and subsequent R. reniformis, and between early R. reniformis and subsequent M. incognita counts. The authors suggested that a competitive interaction existed with each species capable of inhibiting the other and becoming the dominant population.

Bookbinder and Bloom (1980) reported the interaction of Meloidogyne spp. with bean rust, Uromyces phaseoli (Pers.) Wint. They observed that U. phaseoli and M. incognita were synergistic in suppressing shoot and root weights of beans. Meloidogyne incognita infections reduced uredial diameter of U. phaseoli significantly. It was observed that simultaneous inoculations of U. phaseoli and M. incognita resulted in reduced mean numbers of galls per gram of root tissue. Similar effects were observed when U. phaseoli was inoculated first. Meloidogyne incognita numbers were consequently reduced by 62% in rusted plants. This reduction in nematode numbers was probably due to suppressed translocation of photosythates to the roots (Bookbinder and Bloom, 1980). Egg hatch was, nevertheless, not affected by the fungus.














CHAPTER III
THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN (PHASEOLUS VULGARIS L.) YIELD



Introduction



Snap beans, Phaseolus vulgaris L., are defoliated by leaf eating insects, diseases, mechanical injury, and adverse weather conditions (Agudelo, 1980; Costa and Rossetto, 1972; Ruppel and Idrobo, 1962; Schoonhoven and Cardona, 1980; Vargas, 1980). Thus, an understanding of the yield-loss relationship between pest infestations and a crop is essential for the development of an integrated pest management program. Much information on the relationship between a crop and pest infestations has been obtained by simulating pest attack through manual defoliation of plants (Edje et al., 1972, 1973, 1976; Edje and Mughogho, 1976a, b; Galvez et al., 1977; Greene and Minnick, 1967; Vieira, 1981; Waddill et al.; 1984). Manual defoliation is not a precise simulator of pest defoliation (Ruesink and Kogan, 1975); however, it provides a good estimate of the host-pest relationship. To minimize or avoid defoliation by pests, producers often resort to preventive pesticide applications on their crop (Greene and Minnick, 1967). These pesticide applications are a form of insurance on the crop when little knowledge on the pest damage-yield loss relationship is available.

Beans are defoliated by a wide range of leaf-eating insects including leafminers (Liriomyza spp.), cabbage looper (Trichoplusia ni










(Hub.)), leafroller (Urbanus proteus L.), and beetles (Systates spp.). Pohronezny et al (1978) reported that Liriomyza spp. were considered by many local farmers in Dade County, Florida, as the most serious pest on their crops. Farmers expect yield loss as long as same leaf damage is observed, but Harris (1974) showed that leaf consumption by pests does not necessarily result in yield reduction.

Greene and Minnick (1967) reported that snap bean yield was not significantly reduced until more than 33% of the leaf surface was removed during blooming. It was later observed that snap bean plants tolerated up to 66% defoliation if damage occurred before flowering (Greene, 1971). Vieira (1981) found that 66% leaf loss on an indeterminate bean cultivar reduced yield when defoliation occurred during flowering and pod formation. At the first trifoliate leaf stage, Galvez et al. (1977) found that total (100%) defoliation decreased yield of two bean cultivars by 34 and 49% respectively. Total defoliation of plants when only primary leaves were present resulted in yield reduction of 65% on pole beans in Dade County (Waddill et al. 1984).

Kalton et al. (1945) and Weber (1955) reported that up to 75% leaf removal in soybeans had little effect on yield if plants were defoliated prebloom. Defoliation in the bloom and pod formation stages, however, resulted in significant yield losses (Begum and Eden, 1963, 1964; Camery and Weber, 1953; Kalton et al., 1945; McAlister and Krober, 1958). Todd and Morgan (1972) reported significant yield reductions on soybeans with 33%, 67% and 100% defoliation at 2 wk, 4 wk, and 8 wk after first bloom. Research on the effects of defoliation on crop yield has also been conducted on tomato, cotton, corn, wheat, oats, and other grain crops (Dungan, 1930; Keularts, 1980; Ludwig, 1926; Stickler and Pauli, 1961; White, 1946; Wolk et al. 1983; Womack and Thurman, 1962).










This study on snap beans (Phaseolus vulgaris L., 'Sprite') was

conducted to determine the plant growth stage most sensitive to defoliation, and the effects of defoliation on yield.



Materials and Methods



Two defoliation experiments were conducted in the summer and fall, 1984 in the greenhouse and field, respectively. Bush snap beans (Phaseolus vulgaris L. 'Sprite') were grown at the Tropical Research and Education Center in Homestead, Dade County, Florida. The crops were grown on Rockdale soil (pH ca. 7.8). The greenhouse and field experiments were planted on 25 June 1984, and 24 October 1984, respectively. Fertilizer (8:16:16) was applied at the rate of 448 kg/ha according to the University of Florida Extension recommendations (Stall and Sherman, 1983). Benlate(R) (550g ai/ha) was applied fortnightly for control of certain diseases and sprays of Trigard(R) (150 g/ha) were applied at the same frequency for leafminer (Liriomyza spp.) control. Ambush(R) (40 g ai/ha) or Pydrin(R) (250 g ai/ha) was applied at 14-day intervals for cowpea curculio (Chalcodermus aeneus Boh.) control.

Defoliation levels investigated were total (100%), 25%, 50%, and 75%. An undefoliated control was included. Plants were defoliated at the primary leaf stage, first trifoliate leaf, third trifoliate leaf, flower bud formation, full bloom, and pod set. Beans were harvested on 8-20 August 1984 and 18-26 December 1984. The harvest was not graded since cowpea curculio feeding damage to pods was extensive. Greenhouse experiment

Rockdale soil (3030 L) was fumigated with Dowfume(R) MC2 (681 g) on a cement slab under a tightly sealed polyethylene sheet. Number two










(7.5 L) pots were filled with 6.4 L soil and placed on a corrugated bench 0.91 m high. Six seeds were planted in each pot and thinned to three after emergence. A plot consisted of three pots with three plants/pot. The crop was irrigated twice a day using an automatic time-controlled, water-mist-producing overhead system. The foliage was removed from the distal end of the petiole and the correct number of leaves removed at a particular growth stage was determined by leaf counts.

The treatments were replicated four times and randomized in a complete block. Fresh weights of pods were determined. Yield loss (percentage) was computed from the untreated control yield at each growth stage. The dollar economic value was computed by extrapolating plot data to a per hectare basis. Plot yield data were subjected to analysis of variance and regression using the general linear models procedure of SAS (Ray, 1982).

Field experiment

The herbicides Treflan(R) (841 g ai/ha) and Dual(R) (1.7 kg ai/ha) were applied to the site prior to planting. Plots were kept as weedfree as possible by mechanical cultivation. Plots were three rows wide (0.91 m row spacing) and 6 m long. Seeds were mechanically planted at 8-10 cm spacing within the row. The crop was irrigated using an overhead










sprinkler system. The foliage was removed from the distal end of the petiole and the correct proportion of the foliage removed was determined from leaf counts.

Ambush(R) (40 g ai/ha) or Pydrin(R) (250 g ai/ha) was sprayed at 14 day intervals for leafroller and cowpea curculio control. Benlate(R) (550 g ai/ha) and Trigard (150 g ai/ha) were applied fortnightly for disease and leafminer control respectively. Mesurol(R) (200 g ai/ha) was applied as needed for snail and slug control. Treatments were replicated four times in a randomized complete block. Fresh pod weights were determined and yield loss computed from the undefoliated plot data at each growth stage. Yield data were analyzed by analysis of variance and regression utilizing the general linear models procedure of SAS (Ray, 1982).



Results

Defoliation had a significant effect on yield in both the greenhouse and the field, with F-values of 50.16 and 39.95 (p < 0.001) respectively (Table 2). There was no significant interaction between time of defoliation and defoliation levels under greenhouse conditions (F = 0.79). There was significant interaction between defoliation levels and time of defoliation in the field (F = 3.83). Analysis of variance on the effects of time of defoliation showed that there were significant differences between defoliation times (F = 6.23) in the field but not in the greenhouse.

Regression analysis of the relationship between yield in g/plot and defoliation level produced models of the form: Y = a+ bx where Y = log (yield), x = defoliation level and a = intercept. The quadratic model










(a + bx = cx2) resulted in higher coefficients of determination (R2) (Table 3). Generally the fit of the quadratic models to the data was better than the linear model, although the increase in R2 was generally less than 10%. Thus the predictive powers of the linear and quadratic models were more or less similar.

Defoliation did not reduce yield proportionally to its magnitude in both the greenhouse and the field (Tables 4, 5 and Figures 1, 3). Conversely, 25% defoliation resulted in yield increases at the primary leaf stage and full bloom in the field (Table 5). Defoliating plants at the first trifoliate leaf stage at 75% level increased yield by 4% under field conditions (Table 5). No yield increases due to defoliation at pod set in the greenhouse and at full bloom and pod set in the field were observed (Table 5). Total defoliation at pod set resulted in 74% yield loss in the greenhouse but losses of 95% and 92% yield loss at full bloom and pod set in the field.

Yield loss in the greenhouse ranged from 16% to 74% whereas in the field it ranged from -4% to 95% (Table 5). The least yield reduction in the greenhouse was observed at 50% defoliation when foliage was removed at pod set. There was, however, an increase in yield in the field when 75% of the foliage was removed at the first trifoliate leaf stage (Table 5).

Gross dollar values of 'Sprite' snap beans per hectare are shown in Tables 6 and 7 and Figures 5-10. These values were computed based the following price ranges $6.00 (low), $11.00 (medium), and $20.00 (high) for 13.62 kg of snap beans. Since the undefoliated control generally gave higher yields, dollar values obtained from it were higher. In the field, however, 75% defoliation at the first trifoliate leaf stage gave











TABLE 2. F-values from analysis of variance for the effects of defoliation, time (plant growth stage) and their interaction on snap
bean yield.



Field Greenhouse



Source F Prob. F F Prob. F



Defoliation 50.16 0.0001 39.95 0.0001 Growth Stage 6.23 0.0001 0.35 0.92 Defoliation X

Growth Stage 3.83 0.001 1.04 0.79






TABLE 3. Regression equations for the relationship between yield and defoliation.


Plant Growth Stage Greenhouse Field


Linear
2 2 Primary leaf y = 127.5 76x R = 0.48 y = 1525 805x R = 0.41
0.9)a 2 2 (y = 2.11 0.39x) (R 2 = 0.49) (y = 3.18 0.35x) (R2 = 0.34) y = 108.1 4.64x R2 = 0.22 y = 1197 686x R2 = 0.16 First trifoliate leaf (y = 2.01 0.24x) (R2 = 0.22) (y = 3.08 0.62x) (R2 = 0.25) y = 119.5 57.8x R2 = 0.35 y = 1420.4 922x R2 = 0.59 Third trifoliate leaf (y = 2.08 0.31x) (R2 = 0.38) (y = 3.2 0.55x) (R2 = 0.58) y = 117 64.5x R 2 = 0.42 y = 1286.5 863x R2 = 0.61 Flower bud formation (y = 2.07 0.36x) (R2 = 0.42) (y = 3.14 0.54x) (R 2 = 0.53) y = 115.9 55.9x R2 = 0.30 y = 1266.2 999x R2 = 0.66 Full bloom (y = 2.05 0.31x) (R2 = 0.26) (y = 3.36 1.47x) (R2 = 0.46) y = 125.2 76.2x R2 = 0.51 y = 1441 1390.3x R2 = 0.73* Pod set (y = 2.13 0.48x) (R = 0.51) (y = 3.25 1.06x) (R = 0.81)

Quadratic
2 2 2 2=0.43 Primary leaf y = 129.2 89.3x + 13.3 R2 = 0.48 y = 1446 173.6x 631.x R2 = 0.43 (y = 2.07 0.14x 0.25x2) (R2 = 0.22) (y = 3.15 0.08x 0.26x ) 2 (R2 = 0.36)
y = 117 117.3x + 70.9x2 R2 = 0.27 y = 1005.6 + 845.4x 1511.4x R = 0.24 First trifoliate leaf (y = 2.05 0.54x + 0.29x2 (R = 0.25) (y = 2.85 + 1.16x 1.78x ) (R = 0.43)
2 =2 2 y = 117.3 39.9x 17.8- R2 = 0.34 y = 1269.8 + 283.7x 1295.7x R2 = 0.68*
Third trifoliate leaf (y = 2.05 0.Olx 0.23 ) (R2 = 0.41) (y = 3.07 + 0.52x 1.07x ) 2 (R2 = 0.77)
y = 115.7 53.5x 11x2 R2 = 0.43 y = 1284.7 848.7x 1 .3x R2 = 0.61
Flower bud formation (y = 2.03 0.06x 0.3 ) (R2 = 0.45) (y = 3.1 0.17x 0.37x ) 2 (R2 = 0.55)
y = 116.3 58.9x 3x 2 R2 = 0.29 y = 1129.8 + 91.1x 1099.9x R = 0.73* Full bloom (y = 2.03 0.16x 0.15x (R 2 = 0.27) (y = 2.94 + 1.86x 3.32x )2 (R = 0.67)
y = 114.4 + 10.7x 86.9 R2 = 0.55 y = 1552 -2282.6x + 892. 3x R2 = 0.87* Pod set (y = 2.02 0.38x 0.86x ) (R = 0.65) (y = 3.17 0.42x 0.64x ) (R = 0.84)

a Figures in parentheses are y = log (yield); x = defoliation as proportion.


* R significant at 0.05.











TABLE 4. Effect of defoliation and defoliation time on snap bean yield (g/plot) in the greenhouse and field. Data are means of 4 replicates.


Snap bean yield (g/plot) by plant growth stage


First Third Flower
Primary leaf trifoliate leaf trifoliate leaf bud formation Full bloom Pod set Defoliation
level (%) Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field 0 138 1361 122 1168 125 1293 128 1295 121 1138 122 1530 25 89 1403 84 835 97 1210 89 1113 94 1140 96 1085 50 92 1105 72 933 99 1140 89 673 79 690 102 495 75 84 1050 82 1215 81 820 78 620 93 813 84 457 100 46 613 65 120 54 335 46 350 52 52 32 125








TABLE 5. Relationship between defoliation, time and yield (loss (%)) of snap beans in the greenhouse
and field.


Yield (% loss by plant growth stage


First Third Flower
Primary leaf triboliate leaf trifoliate leaf bud formation Full bloom Pod set Defoliation
level (Z) Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field Greenhouse Field

0 0 0 0 0 0 0 0 0 00 0 0 25 36 -3 31 29 22 16 30 14 22 0 21 29 50 34 19 41 20 21 12 30 48 35 39 16 68 75 39 23 32 -4 35 37 39 52 23 29 31 70 100 69 55 47 74 57 74 64 73 57 95 74 92










2.25




2.0




1.75-


C
A


Defoliation Level


Effects of defoliation and time of defoliation on snap beans yield in the greenhouse (linear models).


Letters represent the plant growth stage defoliation occurred. A = primary leaf, B = first trifoliate leaf, C = third trifoliate leaf, D = flower bud formation, E = full bloom, and F = pod set.


Figure 1.













Primary leaf stage First trifoliate leaf Third trifoliate leaf Flower bud formation Full bloom Pod formation


1


0.5


0.75


1.0


Defoliation Level


Effects of defoliation and time of defoliation on snap bean yield in the greenhouse (quadratic models).


Letters represent the plant growth stage defoliation occurred.


2.5




2.0-


1.5




1.0-


0.5




0
0


_I
0.25


Figure 2.






























































Figure 3.


0.25 0.5 0.75 1.0
Defoliation Level
Effects of defoliation and time of defoliation on snap bean yield under field conditions (linear models).


Letters represent the plant growth stage defoliation occurred. A = primary leaf, B = first trifoliate leaf, C = thrid trifoliate leaf, D = flower bud formation, E = full bloom, and F = pod set.












3.25


Figure 4.


0 0.25 0.5 0.75 1.0

Defoliation Level
Effects of defoliation and time of defoliation on snap bean yield under field conditions (quadratic models).


Letters represent the plant growth stage defoliation occurred. A = primary leaf, B = first trifoliate leaf, C = third trifoliate leaf, D = flower bud formation, E = full bloom, and F = pod set.







TABLE 6. Effects of defoliation and defoliation time on gross dollar values per hectare of 'Sprite' snap beans
grown in the greenhouse.


Gross dollar values per hectare
Time of defoliation (growth stage)

Defoliation First trifoliate Third trifoliate Flower bud level (%) Price rangea Primary leaf leaf leaf formation Full bloom Pod set


0 low 292 258 265 270 256 258
medium 535 472 485 495 468 472 high 974 859 885 900 852 859

25 low 187 178 207 189 199 204
medium 343 326 379 347 365 373 high 623 593 689 630 664 679

50 low 193 152 209 189 167 216
medium 353 279 384 347 307 397 high 643 507 698 630 558 772

75 low 178 175 172 165 197 178
medium 327 321 316 302 361 326 high 594 584 574 549 656 593

100 low 96 137 115 59 110 67
medium 177 250 211 109 201 123 high 332 455 384 198 366 223


a low = $7/13.62 kg; medium = $11.20/13.62 kg; and high


= $20/13.62 kr of snap beans.







TABLE 7. Effects of defoliation and defoliation time (plant growth stage) on gross dollar
Sprite snap beans grown in the field.


values per hectare of


Gross dollar values per hectare
Time of defoliation (plant growth stage)

Defoliation Primary First trifoliate Third trifoliate Flower bud level (%) Price range leaf leaf leaf formation Full bloom Pod set


0 low 333 285 316 317 278 374
medium 611 523 580 582 510 686 high 1110 952 1054 1057 928 1248

25 low 343 176 266 273 279 266
medium 629 323 487 500 511 487 high 1143 588 886 909 930 886

50 low 270 228 278 165 170 157
medium 495 419 510 302 311 288 high 869 761 928 549 566 524

75 low 256 297 200 152 197 112
medium 470 544 367 279 361 206 high 855 990 666 507 656 374

100 low 150 74 82 86 110 30
medium 275 136 151 157 201 55 high 500 247 274 285 366 100

a Low = $7/13.62 kg; medium $11.20/13.62 kg; and high $20/13.62 kg of snap beans.
Low = $7/13.62 kg; medium = $11.20/ 13.62 kg; and high = $20/13.62 kg of snap beans.

























1000 -Price range medium low


800





> 600'



O 0
S400





200







0 25 50 75 100
Defoliation Level


Figure 5. Influence of defoliation on gross dollar value per hactare
of 'Sprite' snap beans defoliated at the primary leaf
stage in the greenhouse.





















1000 Price range medium low 800




w
600


r-4 0

m 400
0





200






0 25 50 75 100 Defoliation Level Figure 6. Influence of defoliation on gross dollar value per
hectare of 'Sprite' snap beans defoliated at the
third trifoliate leaf stage in the greenhouse.



















1000
Price range medium low


800





600





400





200





0
0 25 50 75 100 Defoliation Level Figure 7. Influence of defoliation on gross dollar value per hectare
of 'Sprite' snap beans defoliated at pod set in the
greenhouse.


















Price range


High medium low


25 50 Defoliation Level


Influence of defoliation on gross values per hectare of 'Sprite' snap beans defoliated at the primary leaf stage in the field.


1200 1000 800


600


400. 200-


Figure 8.




















Price range


E high E medium
1 low


Defoliation Level (%)


Figure 9. Influence of defoliation
hectare of 'Sprite' snap trifoliate leaf stage in


on gross dollar values per beans defoliated at the third the field.


1200 1000


800 -


400


200





0


75
75


100





48








1300 Price range 3 high

120 medium 1200 o
low




1000





800





600





400





200 -0
0 25 50 75 100 Defoliation Level (%) Figure 10.. Influence of defoliation on gross dollar values per hectare
of 'Sprite' snap beans defoliated at pod set in the field.










a higher dollar value than the undefoliated control. Removal of 25% of the foliage at the primary leaf stage and full bloom also resulted in slightly higher dollar values than the undefoliated control. Generally, increased defoliation resulted in lower dollar values per hectare.



Discussion

Total defoliation when only primary leaves were present resulted in 69% and 55% yield loss in the greenhouse and field respectively. This level of defoliation resulted in 57%, 95%, 74% and 92% yield reduction in the greenhouse and field when plants were defoliated at full bloom and pod set, respectively. The lower yield reduction in the greenhouse may be due to the better controlled environmental conditions. The only growth stage at which total defoliation resulted in less yield loss in the field was at the primary leaf stage. This may have been due to better recovery of plants from the total defoliation in the field.

Removing 25% of the foliage resulted in yield loss of at least 20% in the greenhouse at all growth stages. This is a substantial loss in terms of dollar values. Thus, it appears that the economic threshold under greenhouse conditions was between 0 and 25% defoliation. In the field the economic threshold level varied with the plant growth stage (Table 4). The increase in yield in the field may have been due to compensatory reactions of the plant. The compensation may have resulted from increased photosynthesis due to increased exposure of the remaining photosynthetically active foliage to light. The same may apply to the increase in yield of plants with 75% defoliation at the first trifoliate leaf stage. Reducing foliage may have increased air circulation among










plants which, indirectly, may have increased carbon dioxide uptake and hence photosynthesis. Increased air circulation may also have resulted in reduced leaf surface humidity which may have reduced subtle fungal diseases from being established on the crop.

Removing all leaves from plants at pod set and full bloom resulted in substantial loss under both conditions probably because at these stages the developing pods were deprived of photosynthates normally manufactured in these leaves. The pods which developed probably utilized reserved photosynthates initially and thereafter photosynthates which were produced from the few leaves which were formed after defoliation. At the primary leaf stage, total defoliation slowed down the growth rate of the plants. Under greenhouse conditions, recovery may have been slow and plants may have been etiolated due to insufficient light, hence the higher yield loss. In the field, total defoliation at the first trifoliate leaf stage through flower bud formation resulted in 74 and 73% yield loss. This loss in yield is essentially similar in magnitude indicating that the sensitivity of plants at these stages was more or less the same.

Results obtained in these experiments seem to show yield increases due to defoliation. This may have been due to chance effects resulting from many factors including plant characteristics and the environment. Generally, data show tendency to lower production and hence dollar values with increased defoliation. The decrease in production due to defoliation may have been to enhancement of pathogen entry through wounds made during manual defoliation. Defoliating with scissors also led to water loss through direct evaporation. Defoliation also reduced the leaf area for photosynthesis. One can only suspect that the










inconsistency was due to the imprecise nature of manual defoliation in simulating insect damage. It is possible that removing whole fractions of leaf surfaces had a different effect on plants from damage done by leaf feeding pests which usually occurs at random.

Although the influence of defoliation on yield was not consistent

at all plant growth stages, plants showed more sensitivity at full bloom and pod set. This was an indication that leaf damaging pests should be managed before these plant growth stages. If left unchecked and if plants become heavily defoliated, substantial loss in yield would be expected. Insecticides are generally applied as soon as insect pest infestations are detected. Insecticide application normally starts before the pest populations exceed threshold levels. Disease control chemicals are primarily preventatives applied well before the diseases are observed. Since pesticide sprays against diseases and insect pests were the same at all defoliation levels and plant growth stages, the grower would incur loss in gross dollar values proportional to loss in yield. In this study the threshold level for defoliation was below 25%. At all plant growth stages.














CHAPTER IV
THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION ON SNAP BEANS



Introduction



Root-knot nematodes, especially Meloidogyne incognita (Kofoid and White) Chitwood, are a serious threat to bean (Phaseolus vulgaris L.) production in many bean-producing areas of the world (Agudelo, 1980; Allen, 1983; Ngundo, 1977; Sharma and Guazelli, 1982; Singh et al. 1981a). M. incognita has been reported to cause extensive root-galling on bean plants, which interferes with nitrogen fixation by Rhizobium spp. as well as nutrient uptake by the root system. Moreover, M. incognita has been reported to increase the severity of other pathogens through predisposition of host plant root tissues (Carter, 1975a,b; Golden and Van Gundy, 1975; Powell, 1971; Powell and Nusbaum, 1960; Porter and Powell, 1967; Sasser et al., 1955). Thus, yield loss caused by M. incognita and related species is not always a unitary effect, but often a result of interaction of these nematodes with other plantparasitic organisms.

Yield losses of 50 to 90% in field beans have been reported due to severe root-knot nematode infections (Agudelo, 1980; Freire and Ferraz, 1977; Ngundo, 1977; Varon and Galvez, 1974). Yield decreases caused by M. incognita are also well known in other crops (Allen, 1983; Lamberti, 1979). Crop yield is usually expected to be inversely related to nematode counts (Sasser et al., 1968). In view of this theory, Barker










et al. (1976), Di Vito et al. (1981), and Di Vito and Ekanayake (1983) reported the relationship between initial M. incognita densities and plant growth or yield of tomato and sugar beet. Barker et al. (1976) showed that M. incognita suppressed yields of tomato in North Carolina by up to 85% in the coastal plains and 20-30% in mountain locations. Meloidogyne incognita has been observed to cause yield losses of 30-60% and up to 15% in eggplant and pepper (Capsicum frutescens L.) respectively (Lamberti, 1975). Yield losses due to M. incognita infections have been reported on okra (Hibiscus esculentus L.), sweet potato (Ipomoea batatas (L.) Lam.), celery (Apium graveolens L.), and carrot (Daucus carota L.) (Lamberti, 1971).

Root-knot nematodes rarely occur alone on any crop (Powell, 1971). Thus, nematodes may occur together with other plant pests and diseases. McSorley and Waddill (1982) reported yield loss partitioning on yellow squash (Cucurbita pepo L.) into nematode and insect components by using multiple regression. The partitioning of yield loss was facilitated by the use of selective pesticides. Consequently, McSorley and Waddill (1982) suggested that it may be imperative to separate pests into nematode and insect components when complexes of several pests were present. This separation of yield loss components would be facilitated by monitoring field pest populations during the growing season, at specific intervals, to detect population changes (McSorley and Waddill, 1982).

Beans are susceptible to defoliation by insects, adverse environmental conditions, diseases, and mechanical injury (Agudelo, 1980). Hence an understanding of the relationship between crop yield and pest infestations is essential for the development of sound pest management










strategies. One way of elucidating this relationship has been manual defoliation of plants to simulate pest damage (Douglas et al., 1981; Galvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho, 1983; Kalton et al., 1945; Keularts, 1980; Wit, 1983, Wolk et al., 1983). Hohmann and De Carvalho (1983) reported that leaf area reduction of 25, 50, 75, and 100% on the bean cultivar Carioca reduced yield by 11, 20, 20, and 70% respectively when defoliation was done at the pod formation stage. At the same percentage of leaf area reduction, defoliation at initiation of flowering decreased yield by 18, 12, 19 and 55% respectively. Greene and Minnick (1967) indicated that yield reduction in 'Harvester' snap beans due to leaf removal began somewhere between 33% and 50% defoliation when plants were defoliated in the bloom or pre-bloom stages. Working on inderterminate snap beans, Waddill et al. (1984) noted that the removal of both primary leaves when only primary leaves were present resulted in yield reduction of up to 65%. Vieira (1981) reported that 66% defoliation of an indeterminate bean cultivar at the flowering and pod formation stages was detrimental to yield. Galvez et al. (1977) observed that total defoliation at the formation of the first trifoliate leaves reduced yields of the bean cultivars ICAGuali and Porrillo-Sentetico by 34% and 49% respectively. These observations indicate that the magnitude of yield loss due to defoliation depends not only on the severity of defoliation but also on the growth stage (time) the defoliation takes place and the cultivar of beans grown. Thus, manual defoliation provides a useful estimate of the host-pest relationship despite its imprecision in simulating pest damage (Ruesink and Kogan, 1975).










This study was conducted to investigate the relationship between M. incognita population levels, manual defoliation, and their interaction to snap bean yield.



Materials and Methods



Two studies were conducted in a greenhouse at the Tropical Research and Education Center in Homestead, Dade County, Florida, in the summer and fall 1984. One experiment was an inoculation experiment with M. incognita designed to determine the effect of this pest alone on yield, and the other experiment examined the simultaneous effect of M. incognita inoculation and manual defoliation. The first experiment (M. incognita alone) was designed as a randomized complete-block replicated four times, involving six different nematode population levels. The other experiment was a 5x4x6 factorial replicated four times and included the following treatments: 5 defoliation levels of 0, 25, 50, 75, and 100%; 4 nematode population levels of 0, 1,000, 10,000, and 100,000 eggs and juveniles per pot; and defoliation at the following 6 plant growth stages: primary leaf, first trifoliate leaf, third trifoliate leaf, flower bud formation, full bloom, and pod set.

Preparations for both experiments were made by covering Rockdale soil (3030 L) with a polyethylene sheet and fumigating it with Dowfume(R) MC2 (681g). Two -gallon, side-drain, plastic pots were filled with 6.4 L soil and placed on corrugated greenhouse benches 0.91 m high. Fertilizer (8:16:16) was applied before planting at 3 g/pot (448 kg/ha). Plants were top-dressed with the same fertilizer at 1.5 g/pot four weeks after germination. The M. incognita inoculation test










was planted on 25 June 1984 and harvested on 28 August 1984. The M. incognita x defoliation study was planted on 24 October 1984 and harvested on 26 December 1984 to 3 January 1985. In each experiment, a plot comprised of 2 pots, each containing 3 plants. Irrigation was provided by an automatic time-controlled, overhead, water-mist-producing system twice a day.

The Meloidogyne incognita population used in each experiment was originally obtained from Hausa potato (Coleus parviflorous Benth.) and was maintained on greenhouse-grown tomato (Lycopersicon esculentum Mill.) 'FloraDade' plants. Meloidogyne incognita eggs were extracted by the sodium hypochlorite (NaOCl) method of Hussey and Barker (1973). A

0.525% NaOCl solution was made from Thrift King(R) commercial bleach (5.25% NaOCl) by dilution with cold tap water (25*C). Tomato roots were thoroughly washed of soil with running tap water. The clean roots were cut into 2-3 cm long pieces and 120 g of the cut root material was manually shaken in 200 ml of the NaOCl solution for 3.5 minutes. The shaken material was serially filtered through 100-mesh, 230-mesh and 500-mesh sieves. The number of eggs and juveniles of M. incognita per ml was determined by counting in a watch glass under a dissecting microscope (20X). Appropriate dilutions of the nematode eggs and juveniles were made according to the population levels used.

Plants were inoculated 10 days after planting by drenching their bases with the nematode egg and juvenile suspension. The nematode population levels of 0, 10, 100, 1,000, 10,000, and 100,000 per pot were equivalent to 0, 0.16, 1.6, 15.6, 156 and 1562 eggs and juveniles per 100 ml soil, respectively.










In the experiment involving simultaneous nematode inoculation and defoliation, plants were defoliated manually with a pair of scissors. The correct number or proportion of leaves to be removed was determined by leaf counts at each plant growth stage. To eliminate additional uncontrolled defoliation, plants were sprayed with Ambush(R) (40 g ai/ha) for bean leaf roller (Urbanus proteus L.) and cowpea curculio (Chalcodermus aeneus Boh.) control; Trigard(R) (150 g ai/ha) for leafminer (Liriomyza spp.) control, and Benlate(R) (550 g ai/ha) for disease control. These pesticides applied at 14-day intervals. Yield was taken from all six plants. Pods less than 7 cm in length and diseased or damaged ones were discarded.

Yield data were subjected to analysis of variance and regression analysis using the general linear models procedure of SAS (Ray, 1982). Nematode inoculation data were also analyzed using Seinhorst's models (Ekanayake and Di Vito, 1984; Ferris, 1984; Ferris et al., 1981; Seinhorst, 1965).



Results



Meloidogyne incognita alone

Analysis of variance on the effect of Meloidogyne incognita population levels on yield showed a significant relationship with F = 26.2*** (Table 8). Regression analysis of the data produced models of the form Y = a + bx or Y = a + bx + cx2 where Y = yield (g/plot) or log (yield), x = log (M. incognita population + 1) (Table 9). Quadratic models consistently gave somewhat higher coefficients of determination (R2 values. The predictive ability of the quadratic models was, however,









TABLE 8. Effect of M. incognita on snap bean yield. Data are means of
4 replicates.



No. of Log (M. incognita Yield (g/plot)a Yield loss M. incognita/Pot Population + 1) (%)



0 0 128 0 10 1.0 103 19

100 2.0 79 38

1000 3.0 70 45

10000 4.0 68 47

100000 5.0 53 59


a Data rounded off to the nearest whole number. F = 26.2*** for M. incognita populations.










not greatly superior to that of linear models (Figure 11). Yield reduction was initiated even by the lowest nematode population.

The data did not fit the Seinhorst (1965) model, which is of the
= m+ (-rn z[P-TI
form Y = m + (1-m) Z where Y = ratio between yield at nematode population level p and in the absence of nematodes, m = minimum yield at very high nematode population levels, T = tolerance limit (the nematode population level below which yield reduction does not occur), and Z = the proportion of the plant undamaged in the presence of parasitism or infection by one nematode (Ferris et al., 1981; Ferris, 1984; Seinhorst, 1965 and 1972). The Seinhorst model produced a R2-value of only 0.00661, an indication that the data had a poor fit to the model. Data were, moreover, linear in trend (Figure 11).

Table 10 shows the gross dollar values per hectare of snap beans. These values were computed from the price ranges of $6.00 (low), $11.00 (medium), and $20.00 (high) per bushel (13.62 kg) of beans. These were derived from gross yield and prices given above. They are what the grower would get without subtracting the cost of production which includes labor, pesticides, rent, farm machinery depreciation, and/or interest in loans. The nematode-free plots produced at least twice as much money as plots with 10,000 or 100,000 eggs and juveniles of M. incognita per pot which were equivalent to 156 and 1562 eggs and juveniles per 100 ml soil. Nematode-free plants produced $404, $740 and $1345 per hectare at the low, medium, and high prices respectively compared to $326, $598, and $1087 when pots were inoculated with 10 eggs and juveniles (= 0.16/100 ml soil). This level of M. incognita population resulted in a 19% loss in gross dollar value per hectare ($77, $142, and $258 loss at low, medium and high prices, respectively).










Meloidogyne incognita inoculation x defoliation

Analysis of variance on the relationship between yield and M.

incognita defoliation, and their interaction showed that there was no significant interaction between defoliation and M. incognita at all plant growth stages during which defoliation was done (Table 11). There were, however, significant differences among M. incognita population levels, and there were also significant differences in yield with defoliation level at all plant growth stages (Table 11). Regression analysis produced models of the form Y = a + bx + cn, where Y = yield (g/plot, x = defoliation level (as a fraction), and n = log (M. incognita population + 1) (Table 12). Quadratic models of the form Y = a + blx + b2x2 + cln + C2n2 gave somewhat higher coefficients of determination (R2) (Table 12), but are more difficult to visualize graphically. In general, yield was reduced much faster by nematodes than when defoliation was held constant then when nematode populations were held constant and defoliation levels were changed (Figure 13).

Table 13 shows data on the effects of defoliation, and M.

incognita, when they occurred simultaneously, on snap beans. The lowest yields were obtained when 100% of the leaf area was removed at the first trifoliate leaf stage or at flower bud formation and plants were inoculated with 100,000 eggs and juveniles/pot (Table 13). Total defoliation at any time from the third trifoliate leaf stage through to pod set reduced yield by at least 93% in the presence of an initial nematode population density of 100,000 eggs and juveniles/pot. There was, generally, no apparent synergistic effect when defoliation and M. incognita occurred simultaneously (Tables 13 and 14), which was evidenced by the lack of significance for the interaction term in the analysis of variance.










TABLE 9. Regression equations for the relationship
population levels (x) and yield component


between M. incognita
(y).


Yield component Model Linear Quadratic



Yield Y = 118.6 -13.98x Y = 126.7 26x + 2.4x2 R2 = 0.81* R2 = 0.87*



Log (Yield) Y = 2.1 0.1x Y = 2.1-0.1x + 0.005x2 R2 = 0.84** R2 = 0.85*


* R significant at 0.05

** R significant at 0.01











160



140 120


mUU- Linear
2 Quadratic
,a

80

3 60



40



20 .
0 1 2 3 4 5 Log (M. incognita Population + 1)


Effects of M. incognita on snap bean yield.


Figure 11.










TABLE 10.


The influence of M. incognita on the gross dollar value per hectare of snap beans grown in a greenhouse.


Log (M. incognita Price rangea Gross dollar values

population +1) per hectare



0 low 404 medium 740 high 1345

1.04 low 326 medium 598 high 1087

2.0 low 248 medium 456 high 828

3.0 low 222 medium 407 high 740

4.0 low 215 medium 395 high 718

5.0 low 167 medium 307 high 558

a Low = $6; medium $11; high = $20/13.62 kg of snap beans. Low = $6; medium = $11; high = $20/13.62 kg of snap beans.







TABLE 11. F-values and probability levels from analysis of variance for the effects of defoliation and M.
incognita and their interaction on snap bean yield.


Plant Growth Stage

Primary First tri- Third tri- Flower bud Full Pod Source leaf foliate leaf foliate leaf formation bloom set


Prob. Prob. Prob. Prob. Prob. Prob. F F F F F F F F F F F F Defoliation 17.67 0.0001 67.84 0.0001 66 0.0001 41.6 0.0001 100 0.0001 140 0.0001 Log (M. incognita
population +1) 23 0.0001 23.5 0.0001 35 0.0001 38 0.0001 28.95 0.0001 56.15 0.0001 Defoliation x
Log (M. incognita
population +1) 0.56 0.8871 0.54 0.8395 1.5 0.0962 0.72 0.7396 1 0.4601 0.38 0.9575





TABLE 12. Regressions equations for the relationship between M. incognita defoliation and yield.


Growth stage plants were defoliated


Linear equations

Primary leaf stage First trifoliate leaf stage Third trifoliate leaf stage Flower bud formation stage Full bloom stage Pod set stage


Quadratic equations

Primary leaf stage First trifoliate leaf stage Third trifoliate leaf stage Flower bud formation stage Full bloom stage Pod set stage


aRegression equatona Regression equation


259-22.12N-25.1x (R2 = 0.62**) 256.51-21.04N-32.3x (R2 = 0.59**)
2
267.04-21.51N-33x (R = 0.72**)
2
258.15-24.17N-30.19x (R = 0.64**)
2
237.26-17.75N-31.34x (R = 0.58**)
2
25176-21.44N-29.77x (R = 0.58**)




241.97 + 2.07N-5.13N2-7.9x-4.3x2 (R = 0.66**) 227.11 + 5.39N-5.61N2+8.09x-10.1x2 (R2 = 0.67**) 239.73 2.42N-4.03N2+7.42x-10.1x2 (R2 = 0.79**) 243.49 10.31N-2.94N2-10.53x-4.94x2 (R2 = 0.66**) 204.27-0.57-3.64N2+22.68x-13.5x2 (R = 0.79**) 218.66-0.24N-4.5N2+21.65x-12.86x2 (R2 = 6.68**)


aY = Yield (g/plot). N = Log (M. incognita population + 1). X = Defoliation level (fraction).
** = R significant at 0.01.







Effects of M. incognita and defoliation on snap bean yield (g/plot). Data are means of 4


replicates.


Snap bean yield (g/plot) by plant growth stage


First Third Flower
Defoliation Log (M. incognita Primary trifoliate trifoliate bud Full Pod
level population +1) leaf leaf leaf formation bloom set


0 0 261 278 274 288 238 247 0 3.0 219 247 230 207 188 193 0 4.0 173 163 150 155 122 143 0 5.0 112 96 102 87 87 89 0.25 0 213 166 225 215 224 220 0.25 3.0 186 138 173 148 182 175 0.25 4.0 139 118 141 129 141 135 0.25 5.0 84 60 121 95 110 106 0.50 0 217 235 204 196 187 203 0.50 3.0 172 183 170 145 157 154 0.50 4.0 147 140 153 94 126 111 0.50 5.0 87 107 114 71 98 74 0.75 0 185 188 193 180 145 216 0.75 3.0 145 156 156 144 135 170 0.75 4.0 106 114 87 95 109 136 0.75 5.0 92 80 66 80 69 66 1.0 0 126 46 80 83 48 38 1.0 3.0 87 30 50 50 28 35 1.0 4.0 58 21 29 27 17 23 1.0 5.0 41 12 19 16 17 15


TABLE 13.











Price range


High Medium low


.5 = ~ .~ I - a a m I I ~ - I - I I -


0 1 2 Log (M. incognita


3 4 5 population +1)


Figure 12.


The influence of M incognita on gross dollar value per hectare of 'Sprite' snap beans.


1400 1200 1000


800 600


400 200





0





68





255
















00
> 23 218








'75 155




















Figure 13: Effects of defoliation and M. incognita on snap
bean yield.





The influence of M. incognita and defoliation on snap bean yield loss (%).


Yield loss (%) by plant growth stage


First Third Flower
Defoliation Log (M. incognita Primary trifoliate trifoliate bud Full Pod
level population +1) leaf leaf leaf formation bloom set


0 0 0 0 0 0 0 0 0 3.0 16 11 16 28 21 22 0 4.0 34 41 45 46 49 42 0 5.0 57 65 63 70 62 60 0.25 0 19 40 18 25 6 11 0.25 3.0 29 50 37 49 24 29 0.25 4.0 47 58 49 55 41 45 0.25 5.0 68 78 56 67 53 57 0.50 0 17 15 26 32 21 18 0.50 3.0 34 34 38 50 34 38 0.50 4.0 44 50 44 67 47 55 0.50 5.0 67 62 58 75 59 70 0.75 0 29 32 30 38 39 12 0.75 3.0 45 44 43 50 43 31 0.75 4.0 60 59 68 67 54 45 0.75 5.0 65 71 76 72 71 73 1.0 0 52 83 71 71 80 84 1.0 3.0 67 89 82 83 88 86 1.0 4.0 78 93 90 91 93 91 1.0 5.0 84 96 93 94 93 94


TABLE 14.







TABLE 15. Effects of M. incognita and defoliation on the gross dollar values per hectare of snap
beans.


Gross dollar values by plant growth stage


First Third
Defoliation Log (M. incognita Price Primary trifoliate trifoliate Flower bud Full
level population +1) range leaf leaf leaf formation bloom Pod set


0



3.0 4.0 5.0



0



3.0 4.0


low medium high

low medium high

low medium high

low medium high

low medium high

low medium high

low medium high


830 1522 2768

697 1278 2325

548 1005 1827

357 655 1191

673 1233 2242

589
1081 1965

440 807 1467


882 1617 2940

785 1439 2617

520 954 1734

309 566 1029

529 970 1764

441 808
1470

370 679 1235


870 1595 2900

731 1340 2436

478 877 1595

322 590 1073

714 1308 2379

548 1005 1827

444 813 1479


916 1679 3053

659 1209 2198

495 907 1649

275 504 916

687 1259 2289

467 856 1557

412 756 1374


755 1384 2517

596 1094 1988

385 706 1283


783 1435 2609

611 1119 2035

454 833 1514


284 313 521 574 956 1044


710 1301 2366

574 1052 1913

445 817 1485


697 1278 2323

556 1019 1853

430 789 1435


0.25


0.25


0.25







TABLE 15. continued


Gross dollar values by plant growth stage


First Third
Defoliation Log (M. incognita Price Primary trifoliate trifoliate Flower bud Full
level population +1) range leaf leaf leaf formation bloom Pod set


0.25


5.0


0.5


3.0


4.0


0.5


0.75


0.75


low
medium high

low medium high

low medium high

low
medium high

low medium high

low medium high

low
medium high


266 487 885

689
1264 2298

548 1005 1827

465 853
1551

274 502 912

589 1081
1965

457 838
1523


194 355 641

750 1375 2499

582 1067
1940

441 808
1470

335 615 1118

600 1099 1999

494 905 1646


383
702 1276

644 1181 2147

540 989 1799

482 893 1624

366 670 1219

522 957 1740

496 909 1653


302
554 1007

586
1075 1954

458 840 1527

302
554 1003

229 420 764

568
1041 1893

458 840 1527


355 651
1184

596
1094 1988

586 1075
1954

398 730 1327

310 568
1032

461 845 1536

430 789 1435


337 617
1122

642 1177 2140

485 890
1618

352
646 1175

235 430 783

689 1263 2297

540 990 1800







TABLE 15. continued.


Gross dollar values by plant growth stage

First Third
Defoliation Log (M. incognita Price Primary trifoliate trifoliate Flower bud Full
level population +1) range leaf leaf leaf formation bloom Pod set


0.75 4.0 low 332 362 278 302 347 430 medium 609 663 510 554 636 789 high 1107 1206 928 1007 1157 1435

0.75 5.0 low 291 256 209 256 219 133 medium 533 469 383 470 401 244 high 969 853 696 855 730 443

1.0 0 low 399 150 252 266 151 125 medium 731 275 463 488 277 229 high 1329 499 841 885 504 417

1.0 3.0 low 274 97 153 155 91 110 medium 502 178 288 285 167 201 high 912 323 523 518 303 366

1.0 4.0 low 183 62 87 82 53 70 medium 335 113 160 151 97 129 high 609 206 291 275 176 235

1.0 5.0 low 133 35 61 55 53 47 medium 244 65 111 101 97 86 high 443 117 203 184 176 157










Gross dollar values are shown in Table 15 and Figure 12. These

values were computed from gross yield per hectare based on the following price ranges $6.00 (low), $11.00 (medium), and $20.00 (high). Hence these are gross values without deducting production costs. The gross dollar values (Table 15) show that there was a wide range at each plant growth stage. Generally highest dollar values were obtained when the plants were nematode-free and when no defoliation occurred. The combination of nematodes and defoliation had inconsistent effects on gross dollar values. If defoliation were held constant at any level, gross dollar values decreased as the nematode populations were increased (Table 15). If nematode populations were held constant the decrease in gross dollar values was not always consistent with the levels of defoliation. This is shown clearly at the primary leaf, first trifoliate leaf, full bloom and pod set stages of plant growth. At these stages some lower levels of defoliation have smaller dollar values than higher defoliation levels. As expected, loss in dollar values was similar to yield loss since the former was obtained from yield (g/plot), but varies greatly depending on the current market price which can fluctuate widely.



Discussion



There were significant differences in yield when M. incognita inoculated to snap beans. In this test 10 eggs and juveniles/pot reduced yield by at least 19%. In the defoliation M. incognita interaction test, 1,000 eggs and juveniles/pot depressed yield by only 11-28% when plants were not defoliated (Table 14). Similar trends










occurred when 10,000 or 100,000 eggs and juveniles/pot were used. This discrepancy in M. incognita effects on snap bean yield may be due to the difference in the seasons in which the two trials were conducted. Tyler (1933) reported that at temperatures ranging from 27.50C to 300C, females of Meloidogyne spp. developed from infective juveniles to the egg-laying stage in 17 days; at 24.50C in 21 to 30 days; at 200C, in 31 days; at 15.40C in 57 days; and at temperatures above 33.5%C or below 15.40C, females failed to reach maturity on tomato plants. Decker and Casamayor-Garcia (1966) stated that one generation of M. incognita developed on lettuce within 26 days at a mean temperature of 23.30C. They further stated that from the time of larval invasion up to the commencement of egg-laying required at least 19 days. Lamberti (1979) observed that M. incognita rarely started to invade root tissues when the soil temperature was below 18*C. Since the two tests being discussed here were conducted at different times of the year, it is likely that in the summer test soil temperatures were generally higher than in the fall trial. Thus, the life cycle of the nematode may have been completed in a shorter period of time in the summer than in the fall. Consequently, more M. incognita generations (at least 3) may have been completed during this season. There is also the possibility that the quality of the inoculum was different in the two tests since the source of the eggs and juveniles for the fail test was also exposed to relatively lower temperatures than the summer inoculum. The verification of this phenomenon can only be obtained by conducting further tests. The higher yields in the fall test may also be due to the fact that the nematodes were not able to invade the root tissues of the plants as fast as they could under optimum summer conditions. During










these tests soil temperatures were not taken which in fact precludes the comparison of edaphic temperatures during the time the two tests were conducted. Soil temperatures are generally warmer when air temperatures are higher.

The results obtained in the test where M. incognita was used alone indicate that the threshold population level was between 0 and 10 eggs and juveniles/pot whereas in the fall experiment the threshold population level was between 0 and 1,000 eggs and juveniles/pot. During the fall, 10 and 100 eggs and juveniles were not used hence the threshold for this test could have possibly, been similar to the summer threshold level. Generally, defoliation increased yield loss when combined with nematodes (Table 14). The influence of defoliation level on yield was not as drastic as expected. It is not apparent why defoliation had this slight effect on yield. This is not, however, in agreement with the general principle that nematodes predispose plants to diseases and other pests. Statistical analysis showed that defoliation and M. incognita acted independently in influencing yield.

The gross dollar values had a trend similar to that of yield since they were computed from gross yield. These values are gross figures from which one has to deduct production costs which include pesticides, land rent, labor, interest on loans (if any), and farm machinery depreciation. Thus, net income would depend on the cost of production and current market prices of snap beans. Snap bean production is costly (Taylor and Wilkowske, 1984). Loss in gross income ranges compared to nematode free plants were $470 to $1443; $441 to $2441; $521 to $2059; $764 to $2167; $151 to $2201; and $286 to $2192 when plants were defoliated at various levels at the primary leaf, first trifoliate leaf,










third trifoliate leaf, flower bud formation, full bloom, and pod set stages respectively. If plants were not defoliated but were inoculated with nematodes, loss in income ranged from $443 to $1577; $323 to $1911; $464 to $1827; $855 to $2137; $529 to $1561; and $674 to $1565 when plants were meant to be defoliated at the primary leaf, first trifoliate leaf, third trifoliate leaf, flower bud formation, full bloom, and pod set stages respectively. In each case the lower loss in income is for the 1,000 eggs and juveniles/pot population level and the higher value in loss was for the 100,000 eggs and juveniles/pot (Table 15). These losses are based on yield loss disregarding production costs. The loss in income has been computed using the high market price of snap beans. The yields of snap beans in both studies were low, and thus, the grower would have had a loss in income in both cases.














CHAPTER V
THE EFFECT OF BEAN RUST, UROMYCES PHASEOLI (PERS.) WINT.,
ON SNAP BEANS, PHASEOLUS VULGARIS L. 'Sprite'



Introduction



Bean rust, caused by the fungus Uromyces phaseoli (Pers.) Wint., is a serious disease of beans, Phaseolus vulgaris L. (Agudelo, 1980; Allen, 1983, McMillan et al., 1982, Pohronezny et al., 1984). The disease causes severe damage on winter and spring grown snap beans in south Florida (McMillan, 1982; Pohronezny et al., 1984). Usually, U. phaseoli first appears in January and becomes progressively more severe February through May (Pohronezy et al., 1984). Initial inoculum is believed to come from infected bean plant debris in abandoned fields. Losses of up to 78% in pinto beans, 74.2% and 18.4% in 'Ex Rico 23' and 'Bat 308' field beans, respectively, have been reported from severely infected crops in the United States and Latin America (CIAT, 1983; Kelly, 1982).

Bean rust, U. phaseoli, is an autoecious polycyclic disease whose rates of increase are affected by timing, amount of sporulation, light intensity, relative humidity, and relative cultivar susceptibility (Cohen and Rotem, 1970; Cook, 1978; Imhoff et al., 1982 a,b). Rotem et al. (1973) reported that, in an automatic humidity chamber study, humidity was inversely related to the sporulation of U. phaseoli. Infection by U. phaseoli has, however, been reported to be favored by prolonged periods of at least 95% relative humidity and moderate temperatures (15-270C) (Augustin et al., 1972; Gonzalez, 1976; Schein, 1961).

77










Uromyces phaseoli progress on artificially inoculated beans, P. vulgaris 'Bountiful', depended more on length and frequency of wetting periods than on temperature (Imhoff et al., 1982a).

Yield loss due to disease has been observed to be proportional to the area under the disease progress curve or proportional to disease severity at some critical stage of host growth (Madden et al., 1981; Raymundo and Hooker, 1981; Romig and Calpouzos, 1970; Shaner and Finney, 1977; Teng et al. 1979). In many of these studies, area under the disease progress curve satisfactorily explained the relationship between diseases and yield losses. Disease severity at one or more points in time and rate of increase of the disease were also satisfactory disease parameters employed to explain the relationship between disease and yield loss (James, 1974; James and Teng, 1979; Main, 1977).

Berger (1981) compared the logistic and Gompertz models for disease progress curve fitting. It was observed that the Gompertz model consistently gave better fit to the data examined than the logistic model for disease severity values outside the 0.05 < y < 0.6 range (Figure 14). The Gompertz model was superior to the logistic model in linearizing 113 selected disease progress curves (Berger, 1981).

Growers often resort to routine fungicide sprays for disease

control. Currently, weekly sprays with mancozeb are applied for disease control on beans (McMillan et al., 1982; Pohronezny et al. 1984). The effectiveness of these sprays depends on spray coverage and disease severity but in many cases disease control is less than satisfactory (McMillan et al. 1982). Usually, sprays are initiated before disease signs and/or symptoms are observed on the crop.

The present studies were conducted to determine the effect of bean rust on 'Sprite' snap beans under field conditions.


















































Time


Figure 14. General progress curve of a plant disease.










Materials and Methods



Two trials were conducted at the Tropical Research and Education

Center in Homestead, Dade County, Florida, on Rockdale soil (pH ca.7.8). The first trial was planted on 27 February 1985 and the second on 21 March 1985. Beans were harvested on 25 April 1985 and 13 May 1985 respectively.

In both trials plots were 3 rows wide (0.91 m row spacing) and 3 m long. Beans were planted 7-10 cm apart within the row. Prior to planting the herbicides Treflan(R) (841 g ai/ha) and Dual(R) (1.7 kg ai/ha) were applied to the site. Fertilizer (8:16:16) was applied at 448 kg/ha before planting. Plants were top dressed at 224 kg/ha just before flower bud formation. The crops were sprayed with Ambush(R) (40 g ai/ha) fortnightly for cowpea curculio (Chalcodermus aeneus Boh.) control. Slugs and snails were controlled by Mesurol(R) (200 g ai/ha) pellets. Plants were irrigated using an overhead sprinkler system.

In both trials five treatments were arranged in a randomized

complete block design with four replications. Fungicides were used as a tool to manipulate disease levels. Treatments used were (a) no fungicide; (b) bitertanol (57 g ai/ha) at 7-day intervals; (c) mancozeb (0.7 kg ai/ha) tank-mixed with sulfur (4.5 kg ai/ha) at 4-5-day intervals;

(d) same as (c) but at 7-day intervals; and (e) same as (c) but at 14-day intervals. All sprays were applied with Helena(R) sticker or Nu Film-17(R) as a spreader/sticker. Bitertanol plots were virtually disease free.

Plants were inoculated at the primary leaf growth stage using infected pole bean leaves collected from abandoned bean fields. The










infected leaves were clipped on to wire stakes 25 cm above ground level. Two stakes were placed in each plot, one on each end, depending on general wind direction. Disease progress was monitored by taking trifoliate leaves at random from each plot once a week. At each sampling occasion leaves were taken from the same relative level within the canopy. Disease severity (proportion of leaf area infected by the a-b
disease) was determined using the mathematical model y = a-b, where y =
a
disease severity, a = area of leaf before cutting out diseased tissue, and b = area of leaf after cutting out diseased tissue. The mean of the

5 trifoliate leaves was the measure used in the final data. Leaf area was determined by a LiCor(R) portable area meter (Model LI-300, Lombdar Instruments Corp).

Disease progress curves are generally sigmoid in shape (Imhoff et al., 1982a). The generalized progress curve is shown in Figure 14. Progress curves of bean rust in these studies were obtained by determining disease severity at weekly intervals as indicated above. Area under the disease progress curve was computed using the general
n
model: y = 1 [(X.+ni + Xi)/2] [ti + 1 ti] in which y = area under th
the disease progress curve, x = disease severity at the i observation, ti = time (days) at the ith observation, and n = total number of observations. The computations were facilitated by the use of a computer program provided by Dr. R. D. Berger. The computer program employed the following model: y = (((n (x) + n (x + 1))/2) t (x)) where y = area under the disease progress curve, x = disease severity, n = number of disease severity values, and t = time (days) at which observation is made. The rate of disease progress was determined by using the Gompertz model which consistently gave better fit to the data (Berger, 1981).










Gross dollar values were obtained by multiplying yield (kg/ha) by current market prices of 13.62 kg of snap beans which were $6 (low), $11 (medium), and $20 (high), respectively. Net returns from investment were derived by subtracting the gross dollar values of unsprayed plants and the cost of mancozeb from the values realized from sprayed plants. No net returns for bitertanol spray were computed because this fungicide was experimental and its price was not available.

Data were analyzed by the analysis of variance and regression analysis using the general linear procedure of SAS (Ray, 1982).



Results



Figures 15 and 16 show disease progress in trials 1 and 2 respectively. Progress patterns were similar in both trials although disease severity values were higher for trial 2.

Analysis of variance of yield data by treatments gave F values of 5.37 and 10.77 with probabilities of 0.01 and 0.005 for trials 1 and 2, respectively. This showed that there was a significant relationship between yield and disease severity. The disease free plants produced higher yield in trial 2 than in trial 1 (Table 16). In trial 1 disease severity ranged from 0.098 to 0.76 and yield was from 1102 g to 2723 g/plot whereas in trial 2 disease severity ranged from 0.4 to 0.86 and yield was 276 g to 3214 g (Table 16). Generally, where the disease occurred, yield was lower in trial 2 than in trial 1.

Figure 17 shows the relationship between yield loss and maximum proportion of foliage infected (disease severity) for both trials. In trial 1, 0.098, 0.46, 0.65, and 0.76 disease severity resulted in 21%,










17%, 35%, and 60% yield loss respectively whereas in trial 2 disease severity of 0.4, 0.47, 0.71, and 0.86 lead to 55%, 56%, 80%, and 91% yield loss respectively. Figure 18 is a representation of the relationship between area under the disease progress curve (AUDC) and yield for trials 1 and 2. Both disease severity and AUDC were positively correlated with yield loss which showed that these disease measures were inversely related to yield. Figures 17 and 18 are similar in shape. Regression equations between disease severity and snap bean yield shown in Tables 17 and 18 are similar in nature. Regression analysis of the data produced the model y = a + bx, y = yield (g/plot), and x = disease parameters. Both disease severity and AUDC were significantly correlated with yield at full bloom and pod set in trial 1 (Table 17). When pods were fully developed only disease severity was significantly correlated with yield. In trial 2, disease severity and AUDC were significantly correlated with yield at pod set through the stage when pods were fully formed (Table 18). In trial 1 the coefficient of determination (R2) at full bloom or later range from 0.46 to 0.93 (Table 17) while in trial 2 the coefficient of determination (R2) at pod set or later ranged from 0.89 to 0.99 (Table 18). In trial 1 bean rust severity was not significantly correlated with yield at the stage when pods were half developed whereas the in trial 2 the disease was not significantly related to yield at flower bud formation and full bloom (Tables 17 and 18).

The gross dollar values per hectare of snap beans infected by bean rust are shown in Table 19 and Figures 19 and 20. The virtually disease free plants gave the highest dollar values per hectare in both trials. These plants were sprayed with bitertanol an experiment fungicide, which is currently not registered for rust control on beans. Therefore, no










Therefore, no price information on this product is given. In trial 1, plants with a 0.46 disease severity gave a higher dollar value than plants with a 0.098 disease severity. These dollar values corresponded to spray intervals of mancozeb and sulfur of 7-days and 4-5 days (Table 19). Plants with a disease severity of 0.4 and 0.47 gave dollar values of $2367 and $2327 respectively in trial 2. These disease severity values corresponded with 4-5-day and 7-day spray schedules of mancozeb and sulfur (Table 19). In trial 2, gross dollar values were consistently inversely related to both disease parameters (Table 10). The plants with the highest disease parameter produced the lowest gross dollar value.

The relationship between disease severity and net returns from investment per hectare of snap beans is shown in Table 20. No net returns are shown for the virtually disease free plants because an experimental fungicide with no price tag was used on them. In trial 1 there was no improvement on net returns by spraying beans at 4-5 day intervals from the 7-day intervals. Actually, there was a loss in net income by spraying plants more often (Table 20). In trial 1, there were substantial increases in net returns when plants were sprayed at 7-day intervals compared to the 14-day spray schedule. The increases in returns were $1069, $578, and $306 at the high, medium and low prices at the 7-day spray schedule from the 14-day schedule in trial 1. From the 14-day spray schedule to the 4-5-day spray interval there were increases in net returns of $901, $475, and $239 at the high, medium, and low prices respectively in trial 1. By increasing spray frequency from 7-day to 4-5-day intervals there were net losses of $168, $103, and $67 at the high medium, and low prices respectively. Thus, in trial 1 there


































.0


,14 19 24 29 34 39 4
14 19 24 29 34 39 4


Days After Inoculation


Figure 15.


Disease progress curves for Uromyces phaseoli on snap beans sprayed with mancozeb at various frequencies (Trial 1).


A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals, D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur at 14-day intervals.


















































0.1


5 10 15 20 25 30 35 40 45 50


Days After Inoculation


Figure 16.


Disease progress curves for Uromyces phaseoli on snap beans sprayed with mancozeb at various frequencies (Trial 1).


A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals, D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur at 14-day intervals.


..................










TABLE 16. Effects of Uromyces phaseoli on snap bean yield.


Disease Parameters

Maximum Area under disease

Proportion of progress curve

foliage infested (sq. units)

Trial 1 Trial 2 Trial 1 Trial 2


0.76 0.65 0.46

0.098

0.0


0.86 0.71 0.47 0.40 0.0


5.64 5.93 4.11 0.98

0.014


13.90 12.85 9.28 7.33 0.0


Yield (g/plot)


Trial 1


1102 1778 2248 2158 2723


Trial 2


276 642 1426 1452 3214










TABLE 17. Regression equations for the relationship between level bean
rust disease and snap bean yield (Trial 1).


Plant growth stage


Regression equation


Third trifoliate leaf Flower bud formation Full bloom


Pod set


Disease incidence negligible

" "


= 2666.71 = 2428.64 = 2718.09 = 2812.42 = 2622.1 = 2441.97 = 2618.35 = 2627.26


Pods half developed


Pods fully developed


- 29738.55x

- 5481.99d

- 3407.12x

- 181.8d

- 605.23x

- 1617.92d

- 196.94x

- 1689.48d


(R2

(R2 (R2

(R2 (R2 (R2

(R2 (R2


= 0.9**) = 0.83*) = 0.93**) = 0.84*) = 0.69NS) = 0.46NS) = 0.71NS) = 0.8*)


R significant at P < 0.05 R significant at P ( 0.01 = yield (g/plot) = area under the disease progress curve = proportion of foliage infected = not significant at 0.05









TABLE 18. Regression equations for the relationship between bean rust
and snap bean yield (Trial 2).


Plant growth stage


First and Third trifoliate


Flower bud formation


Full bloom Pod set


Pods half formed


Pods fully formed


Regression equation


Disease incidence negligible


y = 2492.5 y = 2443.73 y = 1940.83 y = 1734.26 y = 3113.46 y = 2886.63 y = 3099.74 y = 3220.42 y = 3165.01 y = 3288.9


- 7790.18x (R2


- 23678.58d

- 1141.86x

- 1384.95x

- 893.33x

- 5340.87d

- 369.12x

- 3224.52d

- 203.23x

- 2795.75d


(R2 (R2 (R2 (R2 (R2 (R2 (R2 (R2 (R2


= 0.39NS) = 0.36NS) = 0.16NS) = 0.21NS) = 0.89*) = 0.92**) = 0.98**) = 0.99**) = 0.98**) = 0.98**)


R Significant at 0.05 R Significant at 0.01 = yield (g/plot) = area under the disease progress curve = proportion of foliage infected = not significant

























*- Trial 1

-.Trial 2


Maximum Proportion of Foliage Infected


Figure 17.


The influence of the maximum proportion of foliage infected by Uromyces phaseoli on snap bean yield.


1.0





.8-


/


0 j


~ff/



0















- Trial 1

- Trial 2


Figure 18.


Area Under Disease Progress Curve

The influence of the area under disease progress curve on snap bean yield.


1.0




.8




.6




.4




.2




0
0






TABLE 19.


The influence of Uromyces phaseoli on the gross dollar value per hectare of snap beans (Phaseolus vulgaris 'Sprite').


Disease Parameters at harvest Spray Maximum Proportion of Area under disease
frequency foliage infected progress curve Gross dollar values (days) Price range

Trial 1 Trial 2 Trial 1 Trial 2 Trial 1 Trial 2


0 0.76 0.86 5.64 13.9 high 1797 449 medium 987 247 low 539 135 14 0.65 0.71 5.93 12.85 high 2574 1047 medium 1416 576 low 772 314

7 0.46 0.47 4.11 9.28 high 3665 2327 medium 2016 1280 low 1100 698 4-5 0.098 0.4 0.98 7.33 high 3520 2367 medium 1936 1302 low 1056 710 7a 0.0 0.0 0.014 0.0 high 4446 5244 medium 2446 2884 low 1334 1573

a Sprayed with bitertanol.






TABLE 20.


The relationship between disease severity and net return per hectare of snap beans sprayed with mancozeb and sulfur.


Maximum
Fungicide Proportion of Net return ($) No. of spray frequency foliage infected in investment sprays Price range

Trial 1 Trial 2 Trial 1 Trial 2 Trial 1 Trial 2


0 0 0 0.76 0.86 high 0 0 medium 0 0 low 0 0 3 14-day 14 0.65 0.71 high 743 564 medium 395 295 low 199 145 5(6)a 7-day 7 0.46 0.47 high 1812 1810 medium 973 965 low 505 495 7(8) 4-5-day 4-5 0.098 0.4 high 1644 1828 medium 870 965 low 438 485

a Number of sprays in trial 2 are in parentheses.




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112
TABLE 24. Regression equations for the relationship between various
nematode populations, during the growing season, and snap
bean yield. Nematode populations were transformed to log
(population +1).
Time
Regression Equation
a
Coefficient of
determination (R )
Preplant
y
=
497.3-21x,+7.lx.
1 4
0.025
NS
y
=
538.4-20.3Xj
0.018
NS
Midseason
y
=
894.6-191.lx,-11.lx-25.4x-66.6x.-8.2x_
1 2 3 4 5
0.17
NS
y
=
902.7-200.6x -12.3x -29.6x -67.2x.
12 3 4
0.17
NS
y
=
841.7-220.7xj-24.8x2~20.3x3
0.14
NS
y
=
767.7-187.2Xj-30.4x2
0.13
NS
y
=
770.4-251.61Xj
0.11
NS
Harvest
y
=
797.3-157.6x.-35.5x0-28.6x+9.4x,
12 3 4
0.17
NS
y
=
783.5-142.8Xj-27.lx2-25.8x3
0.17
NS
y
=
731.7-132xj-27.3x2
0.15
NS
y
=
682.8-137Xj
0.13
NS
a
= Criconemella
= Helicotylenchus
= Meloidogyne
= Rotylenchulus
= Tylenchorhynchus
NS = Not significant at 0.05


169
TABLE 43. Mean snap bean yield from plants defoliated at various levels
and various fungicide spray frequencies.
Defoliation
levels (%)
Plant
Fungicide Primary leaf
Growth
Flower
Stage
bud formation
0
No fungicide
143
152
0
Bitertanol (7-day)a
212
232
0
Mancozeb (4-5-day)
185
157
0
Mancozeb (7-day)
144
155
0
Mancozeb (14-day)
126
131
25
No fungicide
112
126
25
Bitertanol (7-day)
181
172
25
Mancozeb (4-5-day)
162
152
25
Mancozeb (7-day)
147
149
25
Mancozeb (14-day)
116
73
50
No fungicide
99
95
50
Bitertanol (7-day)
180
168
50
Mancozeb (4-5-day)
154
153
50
Mancozeb (14-day)
106
128
75
No fungicide
104
98
75
Bitertanol (7-day)
135
134
75
Mancozeb (4-5-day)
121
100
75
Mancozeb (7-day)
108
106
75
Mancozeb (14-day)
109
59
100
No fungicide
100
74
100
Bitertanol (7-day)
93
106
100
Mancozeb (4-5-day)
81
113
100
Mancozeb (7-day)
60
77
100
Mancozeb (14-day)
76
56
a
Figures in parentheses are spray intervals.


Number of juveniles
150
Figure 30
No. juveniles hatched from eggs obtained from bean plants.


Yield (g/plot)
136
igure 27.
The effect of metam-sodium on nsap bean yield when plants
were not defoliated.


88
TABLE 17. Regression equations for the relationship between level bean
rust disease and snap bean yield (Trial 1).
Plant growth stage
Regression equation
Third trifoliate leaf
Flower bud formation
Full bloom
Pod set
Pods half developed
Pods fully developed
Disease incidence negligible
It II II
y =
2666.71
- 29738.55x
(R2
=
0.9**)
y =
2428.64
- 5481.99d
(R2
=
0.83*)
y =
2718.09
- 3407.12x
(R2
=
0.93**)
y =
2812.42
- 181.8d
(R2
=
0.84*)
y =
2622.1
- 605.23x
(R2
=
0.69NS)
y =
2441.97
- 1617.92d
(R2
=
0.46NS)
y =
2618.35
- 196.94x
(R2
=
0.71NS)
y =
2627.26
- 1689.48d
(R2
=
0.8*)
* R significant at P ^ 0.05
** R significant at P < 0.01
y = yield (g/plot)
x = area under the disease progress curve
d = proportion of foliage infected
NS = not significant at 0.05


Number of juveniles
151
Figure 31. No. juveniles hatched from eggs obtained from tomato plants.


V£>


2
the state with the greatest production in Dade County (Anon., 1982).
Gadsden, Marion and Palm Beach counties and parts of the west central
area also produce some snap beans (Anon., 1982). Rose (1975), Ware and
McCollum (1980), and Anon. (1982) reported that the winter demand for
fresh market snap beans in population centers to the north of Florida is
usually met by supplies from the southern districts of Florida.
Whereas Florida is the largest producer of fresh market snap beans,
Wisconsin is the leading producer of snap beans for processing (Kobriger
and Hagedorn, 1983). Michigan, New York, and Oregon also produce more
snap beans for processing than Florida (Kobriger and Hagedorn, 1983;
Ware and McCollum, 1980).
Snap bean production has some inherent pest problems (Anon., 1982;
Rose, 1975). Fresh market snap bean yield, however, increased from 31
cwt/acre in the 1947-1952 period to 37 cwt/acre in the 1967-1972 period,
in the U.S., despite these problems (Anon., 1972). This increase in
yield has been ascribed to the advent of synthetic organic pesticides
during and after World War II. In Florida, snap bean yields were,
however, on the decline during the same period (Rose, 1975). The
decline was associated mainly with adverse weather conditions (Anon.,
1982). Galvez et al. (1977) and Vargas (1980) stated that insect pests,
such as leafminers, leafrollers, corn earworm, Mexican bean beetle, and
others cause tremendous losses in bean yield. Many of these insects
feed on the leaves reducing the photosynthetic tissue of the plant.
These pests have been more-or-less controlled by insecticide sprays
(Acland, 1971; Iraneta and Rodrigez, 1983).
Bean rust, Uromyces phaseoli, (Pers.) Wint., is one of the most
important diseases of beans in many bean producing areas of the world


102
observed that the prediction of yield loss was more accurate when both
insect and nematode pests were present. Bookbinder and Bloom (1980)
reported that Meloidogyne spp. interacted with bean rust, Uromyces
phaseoli, on beans. The root-knot nematodes and the disease had an
additive effect on the suppression of shoot and root weights of bean
plants. Meloidogyne incognita infections reduced uredial diameter of _U.
phaseoli. Similar effects were observed if IJ. phaseoli was inoculated
first. Bookbinder and Bloom (1980) observed that rusted plants had 62%
less M. incognita than uninfected plants. They suggested this was due
to suppressed translocation of photosynthates to the roots. _U. phaseoli
infection did not affect M. incognita egg hatch.
This study was conducted to determine the relationship between
defoliation, nematodes and bean rust and yield of snap beans.
Materials and Methods
General
Three trials were conducted at the Tropical Research and Education
Center in Homestead, Dade County, Florida. Experimental sites were on
Rockdale soil (pH ca. 7.8) planted in fall 1984 and early spring 1985.
The fields had been previously cropped in tomato (Lycopersicon
esculentum Mill.). In all three trials snap beans (Phaseolus vulgaris
L. 'Sprite') were used. Prior to planting, the herbicides Treflan^^
(R)
(841 ai/ha) and Dual" (1.7 kg ai/ha) were applied to the site.
Fertilizer (8:16:16) was applied preplant at 448 kg/ha according to the
University of Florida Extension recommendation (Stall and Sherman,
1983). The plants were topdressed with 224 kg/ha fertilizer at flower


TABLE 7. Effects of defoliation and defoliation time (plant growth stage) on gross dollar values per hectare of
Sprite snap beans grown in the field.
Gross dollar values per hectare
Time of
defoliation (plant growth stage)
Defoliation
level (%)
Primary
Price range3 leaf
First trifoliate
leaf
Third trifoliate Flower bud
leaf formation Full bloom Pod set
0
low
333
285
316
317
278
374
medium
611
523
580
582
510
686
high
1110
952
1054
1057
928
1248
25
low
343
176
266
273
279
266
medium
629
323
487
500
511
487
high
1143
588
886
909
930
886
50
low
270
228
278
165
170
157
medium
495
419
510
302
311
288
high
869
761
928
549
566
524
75
low
256
297
200
152
197
112
medium
470
544
367
279
361
206
high
855
990
666
507
656
374
100
low
150
74
82
86
110
30
medium
275
136
151
157
201
55
high
500
247
274
285
366
100
a
Low = $7/13.62 kg; medium = $11.20/13.62 kg; and high = $20/13.62 kg of snap beans


130
Days After Inoculation
Figure 24. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at 14-day intervals, defoliated at
various levels and soil fumigated with metam-sodium.
19 = no defoliation, no metam-sodium and mancozeb + sulfur at 14-day
intervals, 20 = no defoliation, 935 L/ha metam-sodium and mancozeb +
sulfur at 14-day intervals, 21 = 25% defoliation, no metam-sodium and
mancozeb + sulfur at 14-day intervals, 22 = 35% defoliation, 935 L/ha
metam-sodium and mancozeb + sulfur at 14-day intervals, 23 = 50% de
foliation, no metam-sodium and mancozeb + sulfur at 14-day intervals,
and 24 = 50% defoliation, 935 L/ha metam-sodium and mancozeb + sulfur
at 14-day intervals.


133
Tables 28, 29, and 30 show nematode genera detected in the soil at
preplant, midseason, and at harvest. Metam-sodium rate had no signifi
cant effects on nematode numbers at all sampling times.
Total yield (g/plot) and yield loss are shown in Table 31. The
lowest yield was obtained from plots fumigated at 47 L/ha metam-sodium
and plants defoliated at the 0.75 level. Gross dollar values are given
in Table 32 and net income is shown in Table 33. These gross dollar
values were computed from total yield (per hectare) multiplied by the
following price ranges: $6.00 (low), $11.00 (medium), and $20.00 (high)
per bushel (13.62 kg). Net dollar values were obtained by subtracting
the cost of metam-sodium per hectare from the gross dollar values.
Gross dollar values generally decreased with the increase in defoliation
level. There were net returns on investment when plants were not
defoliated (Table 33). There were no net returns on investment when
plants were defoliated at any level (Table 33).
The effect of defoliation, metam-sodium, bean rust, and their interaction
on snap bean yield
Analysis of variance showed that there were no significant
differences among defoliation levels and metam-sodium rates at the 0.05
level (Table 34-1). There was no significant interaction among
defoliation levels, metam-sodium rates, and fungicides at the 0.05 level
(Table 34-1). There were, however, significant differences among
fungicide sprays (F = 7.62**). When analysis of variance was used for
yield data analysis with disease severity as one of the independent
factors, there were no significant differences in yield based on disease
severity (maximum proportion of foliage infected) (F < 1). There was no
significant interaction among defoliation levels, metam-sodium rates,


32
2 2
(a + bx = ex ) resulted in higher coefficients of determination (R )
(Table 3). Generally the fit of the quadratic models to the data was
2
better than the linear model, although the increase in R was generally
less than 10%. Thus the predictive powers of the linear and quadratic
models were more or less similar.
Defoliation did not reduce yield proportionally to its magnitude in
both the greenhouse and the field (Tables 4, 5 and Figures 1, 3).
Conversely, 25% defoliation resulted in yield increases at the primary
leaf stage and full bloom in the field (Table 5). Defoliating plants at
the first trifoliate leaf stage at 75% level increased yield by 4% under
field conditions (Table 5). No yield increases due to defoliation at
pod set in the greenhouse and at full bloom and pod set in the field
were observed (Table 5). Total defoliation at pod set resulted in 74%
yield loss in the greenhouse but losses of 95% and 92% yield loss at
full bloom and pod set in the field.
Yield loss in the greenhouse ranged from 16% to 74% whereas in the
field it ranged from -4% to 95% (Table 5). The least yield reduction in
the greenhouse was observed at 50% defoliation when foliage was removed
at pod set. There was, however, an increase in yield in the field when
75% of the foliage was removed at the first trifoliate leaf stage (Table
5).
Gross dollar values of 'Sprite' snap beans per hectare are shown in
Tables 6 and 7 and Figures 5-10. These values were computed based the
following price ranges $6.00 (low), $11.00 (medium), and $20.00 (high)
for 13.62 kg of snap beans. Since the undefoliated control generally
gave higher yields, dollar values obtained from it were higher. In the
field, however, 75% defoliation at the first trifoliate leaf stage gave


TABLE 38. Effect of defoliation, metam-sodium and bean rust on yield of gross dollar value per hectare of
snap beans. Data are means of 3 replicates.
Disease severity Gross dollar values
Defoliation level
(proportion of foliage)
Metam-sodium
(Liters/ha)
(proportion of
foliage infested)
Yield
(g/plot)
high
Price Range
medium
low
0
0
0.87
184
300
165
90
0
935
0.83
365
596
328
179
0.25
0
0.64
260
424
233
127
0.25
935
0.71
345
563
310
169
0.5
0
0.77
220
359
197
108
0.5
935
0.74
338
551
303
165
0
0
0.02
807
1316
724
395
0
935
0
827
1349
742
405
0.25
0
0
570
930
512
279
0.25
935
0.01
499
814
448
244
0.5
0
0
446
727
400
219
0.5
935
0
548
895
492
268
0
0
0.56
349
569
313
171
0
935
0.59
573
935
514
281
0.25
0
0.70
520
848
467
255
0.25
935
0.66
276
450
247
135
0.5
0
0.68
258
420
231
126
0.5
935
0.71
143
233
128
70
0
0
0.71
363
593
326
178
0
935
0.74
406
662
364
199
0.25
0
0.75
190
310
171
93
0.25
935
0.83
359
585
322
176
0.5
0
0.72
327
533
293
160
0.5
935
0.81
397
648
357
196
131


123
TABLE 34-2. Mean snap bean yield per plot sprayed with fungicides.
Fungicide
Spray
Frequency
Q
Mean yield (g/plot)
No fungicide
0
284a
Mancozeb
14-day
396a
Mancozeb
7-day
342a
Bitertanol
7-day
616b
Means followed by the same letter are not significantly different at
P 0.05 (Duncan's multiple range test).


182
Harris, P. 1974. A possible explanation of plant yield increases
following insect damage. Agro-Ecosystems 1: 219-225.
Harter, L. L., C. F. Andrus and W. J. Zaumeyer. 1935. Studies on bean
rust caused by Urmyces phaseoli typica on bean. J. Agrie. Res. 50:
737-759.
Harter, L. L. and W. J. Zaumeyer. 1941. Differentiation of physiologic
races of Uromyces phaseoli typica on beans. J. Agrie. Res. 62:
717-730.
Hartmann, R. W. 1968. Manoa Wonder, a new root-knot nematode resistant
pole bean. Hawaii Agrie. Expt. Sta. Univ. Hawaii, Circ. 67. 12 pp.
Hikida, H. R. 1961. Race 33 of Uromyces phaseoli var. typica Arth., A
Distinct Physiologic Race of Bean Rust from Oregon. Plant Dis.
Rep. 45: 388.
Hikida, H. R. 1962. Races of bean rust, Uromyces phaseoli in
Willamette Valley Dis. Abstr. 22: 3341-3342.
Hilty, J. W. and C. A. Mullins. 1975. Chemical control of snap bean
rust. Tennessee Farm and Home Sci. 93: 4-5.
Hohmann, C. L. and S. M. De Carvalho. 1983. Effect of defoliation on
yield of beans. An. Soc. Entomol. Bras. 12: 3-10.
Hoppe, H. and R. Heitefuss. 1974a. Permeability and membrane lipid
metabolism of vulgaris infected with U. phaseoli. I. Changes in
the efflux of cell constituents. Physiol. Plant Pathol. 4: 5-9.
Hoppe, H. and R. Heitefuss. 1974b. Permeability and membrane lipid
metabolism of 1?. vulgarlyinfected with U. phaseoli II. Changes in
lipid concentration and P incorporation into phospholipids.
Physiol. Plant Pathol. 4: 11-23.
Howland, A. K. and J. C. MaCartney. 1966. East African bean rust
studies. East Afr. Agrie. For. J. 32: 208-210.
Huffaker, C. B. and R. F. Smith. 1980. Rationale, organization and
development of a national IPM. pp. 1-24 In: New Technology of
Pest Control. C. B. Huffaker (ed.). Wiley, New York.
Hussey, R. S. and K. R. Barker. 1973. A comparison of methods of
collecting inocula of Meloidogyne spp. including a new technique.
Plant Dis. Rep. 57: 1025-1028.
Imhoff, M. W., K. J. Leonard, and C. E. Main. 1982a. Analysis of
disease progress curves, gradients, and incidence-severity
relationships for field and phytotron bean rust epidemics.
Phytopathology 72: 72-80.
Imhoff, M. W., K. J. Leonard, and C. E. Main. 1982b. Patterns of bean
rust lesion size increase and spore production. Phytopathology 72:
441-446.


175
largest dollar values. Four to five-day sprays of mancozeb were
generally better than 7-day spray schedules on yield. Bitertanol gave
the highest yield and highest dollar values. Four to five-day sprays of
mancozeb and sulfur were generally better than 7-day sprays. This may
have been due to the presence of zinc, an element required for plant
growth. May be the more frequent sprays provided more of this element
than the 7-day spray schedule of mancozeb and sulfur rather than the
efficacy of the fungicide since there was virtually no apparent disease
build up on the plants. It is possible that the more frequent sprays
controlled other diseases resulting in higher yield. This same fact may
be true for the fortnightly spray schedule which generally gave the
lowest yield among the fungicide sprays.
The net income value picture is not clear with regard to bitertanol
since its market price was not known. Moreover, the net income depends
on snap bean prices which fluctuate widely. Furthermore, these values
do not include other production costs such as farm machinery, labor and
interest on loans. There was no apparent benefit from the shorter (4-5
day) spray schedule. The extra sprays did not produce enough increase
in yield to justify added cost. From these results the ideal mancozeb
spray schedule was the 7-day interval. Biteranol increased snap bean
yield substantially probably because this chemical controlled some other
subtle disease and it may have plant growth regulating properties. This
has been suggested by teh non-establishment of bean rust disease in this
study. Bitertanol has no known micronutrients which ruled out the
micronutrient effect.


172
TABLE 44. Continued
Defoliation
level (%)
Fungicide
Price
range
Plant
of
Primary
leaf
Growth Stage
defoliation
Flower bud
formation
low
636
887
100
Mancozeb
medium
1165
1625
(4-5-day)
high
2118
2955
low
471
604
100
Mancozeb
medium
863
1108
(7-day)
high
1569
2014
low
596
439
100
Mancozeb
medium
1093
806
(14-day)
high
1988
1465


Log (Yield)
40
Figure 4. Effects of defoliation and time of defoliation on snap
bean yield under field conditions (quadratic models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = third trifoliate
leaf, D = flower bud formation, E = full bloom, and F = pod set.


94
ai
3
CO
>
)-i
CO
O
a
en
en
o
u
O
5000
4000
3000
2000
1000
0
0 0.098 0.46 0.65 0.76
Disease Severity
Figure 19. Influence of disease severity on gross dollar value per
hectare of 'Sprite' snap beans in trial 1.


144
(Briggs, 1946; Feder and Feldmesser, 1955). Sodium hypochlorite has
also been used for sterilizing processing substrates and equipment in
diagnostic nematology laboratory work (Esser, 1972), so its adverse
effects on nematodes at higher concentrations is well known.
The main objectives of this study were twofold (1) to determine the
influence of the initial M. incognita population density on yield of
beans and effects of inoculation method on the establishment of M.
incognita on beans, and (2) to determine the effect of sodium hypochlo
rite (NaOCl) concentration on the number of M. incognita eggs and
juveniles extracted from infected bean and tomato plants, and the
influence of NaOCl concentration on egg hatch.
Materials and Methods
Meloidogyne incognita (Kofoid and White) Chitwood, obtained from
Hausa potato (Coleus parviflorous Benth.), was maintained on greenhouse-
grown tomato (Lycopersicon esculentum Mill Floradade). Infected bean
roots were obtained from an earlier experiment conducted in a greenhouse
at the Tropical Research and Education Center in Homestead, Dade County,
Florida. Sodium hypochlorite (NaOCl) solutions (0.13, 0.26, 0.525, 1.3
and 2.6%) were made from Thrift King^ commercial bleach (5.25% NaOCl)
diluted serially with cold tap water (25C). Roots were cleaned of
soil, cut into 2-3 cm pieces, mixed, and 120 g of the mixture was used
for egg and juvenile extraction by the sodium hypochlorite method
(Hussey and Barker, 1973), except that a 230-mesh sieve was used instead
of the 200-mesh. Comparisons of egg and juvenile numbers extracted by
the various concentrations of NaOCl were made.


Log (Yield)
37
Figure 1. Effects of defoliation and time of defoliation on snap beans
yield in the greenhouse (linear models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = third trifoliate leaf,
D = flower bud formation, E = full bloom, and F = pod set.


114
TABLE 26. Net returns on investment per hectare of snap beans.
Metam-sodium
(Liters/ha)
Price Range
low
medium
high
0
47
94
187
281
0
0
0
41
137
311
- 7
112
325
-141
- 10
210
-338
-247
- 85
-425
-283
- 29
374


14
The life cycle of Meloidogyne spp. has several developmental stages
(Taylor and Sasser, 1978). The adult female lays eggs in a gelatinous
matrix. The first-stage juvenile develops and molts within the egg.
What emerges from the egg is actually the second-stage juvenile, hence
the general belief that root-knot juveniles grow between a series of
three molts into adult males and females (Agudelo, 1980). Root-knot
nematode eggs are oval or ellipsoidal and may be concave on the side.
They measure 30-52 x 67-128 pm (Thorne, 1961). These eggs are usually
protected from dehydration by a gelatinous matrix secreted by the female
(Franklin, 1978; Taylor and Sasser, 1978).
The juvenile stages are vermiform, have a stylet about 10 jum long
and have an overall length of 375-500 pm and a width of 15 pm (Robbins
et al., 1972; Taylor and Sasser, 1978). Males are cylindroid and
measure 0.03-0.36 x 1.2-1.5 mm. The males lack a bursa. Adult females
are pyriform and usually pearly white (visible in roots without magni
fication) The females measure 0.27-0.75 x 0.40 x 1.30 mm and have a
soft cuticle (Franklin, 1978; Taylor, 1965; Walker, 1965). The life
cycle of root-knot nematodes may take 17-57 days, depending on the soil
temperature and the host plant (Tyler, 1933; Taylor and Sasser, 1978).
Infection by and pathogenesis of Meloidogyne spp. are affected by
plant age, plant susceptibility, population size and environmental
factors (Brodie and Dukes, 1972; Gilvonio and Ravines, 1971; Nemec and
Morrison, 1972; Sosa Moss and Torres, 1973). Second stage juveniles of
Meloidogyne spp. enter the plant root system within 2 days after inocula
tion and migrate inter and/or intracellularly through the cortex into
the stele (Dropkin, 1980; Ngundo and Taylor, 1975 b). The juvenile
inserts its head into the vascular system of the root to obtain


53
et al. (1976), Di Vito et al. (1981), and Di Vito and Ekanayake (1983)
reported the relationship between initial M. incognita densities and
plant growth or yield of tomato and sugar beet. Barker et al. (1976)
showed that M. incognita suppressed yields of tomato in North Carolina
by up to 85% in the coastal plains and 20-30% in mountain locations.
Meloidogyne incognita has been observed to cause yield losses of 30-60%
and up to 15% in eggplant and pepper (Capsicum frutescens L.) respec
tively (Lamberti, 1975). Yield losses due to M. incognita infections
have been reported on okra (Hibiscus esculentus L.), sweet potato
(Ipomoea batatas (L.) Lam.), celery (Apium graveolens L.), and carrot
(Daucus carota L.) (Lamberti, 1971).
Root-knot nematodes rarely occur alone on any crop (Powell, 1971).
Thus, nematodes may occur together with other plant pests and diseases.
McSorley and Waddill (1982) reported yield loss partitioning on yellow
squash (Cucrbita pepo L.) into nematode and insect components by using
multiple regression. The partitioning of yield loss was facilitated by
the use of selective pesticides. Consequently, McSorley and Waddill
(1982) suggested that it may be imperative to separate pests into
nematode and insect components when complexes of several pests were
present. This separation of yield loss components would be facilitated
by monitoring field pest populations during the growing season, at
specific intervals, to detect population changes (McSorley and Waddill,
1982).
Beans are susceptible to defoliation by insects, adverse environ
mental conditions, diseases, and mechanical injury (Agudelo, 1980).
Hence an understanding of the relationship between crop yield and pest
infestations is essential for the development of sound pest management


60
Meloidogyne incognita inoculation x defoliation
Analysis of variance on the relationship between yield and M.
incognita defoliation, and their interaction showed that there was no
significant interaction between defoliation and M. incognita at all
plant growth stages during which defoliation was done (Table 11). There
were, however, significant differences among M. incognita population
levels, and there were also significant differences in yield with defo
liation level at all plant growth stages (Table 11). Regression analy
sis produced models of the form Y = a + bx + cn, where Y = yield
(g/plot, x = defoliation level (as a fraction), and n = log (M.
incognita population + 1) (Table 12). Quadratic models of the form Y =
2 2
a + b^x + b^x + c^n + C£n gave somewhat higher coefficients of
2
determination (R ) (Table 12), but are more difficult to visualize
graphically. In general, yield was reduced much faster by nematodes
than when defoliation was held constant then when nematode populations
were held constant and defoliation levels were changed (Figure 13).
Table 13 shows data on the effects of defoliation, and M.
incognita, when they occurred simultaneously, on snap beans. The lowest
yields were obtained when 100% of the leaf area was removed at the first
trifoliate leaf stage or at flower bud formation and plants were inocu
lated with 100,000 eggs and juveniles/pot (Table 13). Total defoliation
at any time from the third trifoliate leaf stage through to pod set
reduced yield by at least 93% in the presence of an initial nematode
population density of 100,000 eggs and juveniles/pot. There was, gen
erally, no apparent synergistic effect when defoliation and M. incognita
occurred simultaneously (Tables 13 and 14), which was evidenced by the
lack of significance for the interaction term in the analysis of
variance.


107
2
+ 1). The linear models gave low R at all soil sampling times (Table
24).
Gross dollar values per hectare are shown in Table 25. The highest
dollar value was obtained with the 374 L/ha soil fumigation, as
expected, since this was the rate at which the highest yield was
achieved. The gross dollar values were based on yield per hectare and
the following prices $6.00 (low), $11.00 (medium), and $20.00 (high) per
bushel of snap beans (13.62 kg). Net income is shown in Table 26.
Yields obtained in this study were generally low. These yields were low
probably because of the less ideal temperatures at the time the study
was conducted. Under these conditions the grower would have made a
profit at 47 L/ha, and 94 L/ha at the low, medium, and high snap bean
prices respectively (Table 26). The dollar values were based on the
metam-sodium cost of $1.59/L (McSorley and Pohronezny, 1984); no other
expenses have been used in economic analyses.
The effect of metam-sodium and defoliation on snap beans
Analysis of variance on the influence of metam-sodium, defoliation,
and their interaction on snap beans showed that there was no significant
interaction between metam-sodium and manual defoliation (Table 27).
There were also no significant differences among metam-sodium rates (F =
1.5 NS) on yield. There were, however, significant differences among
defoliation levels (F = 20.22**) (Table 27). Regression analysis of
2 2
yield data on metam-sodium rates produced the model y = a + bx cx (R
= 0.34) where y = yield (g/plot), x = metam-sodium rate (gal/acre)
(Figure 21). The cubic model of the form y = a + bx + cx + dx (R =
0.46) was obtained when yield data were analyzed by regression against
manual defoliation (Figure 28).


50
plants which, indirectly, may have increased carbon dioxide uptake and
hence photosynthesis. Increased air circulation may also have resulted
in reduced leaf surface humidity which may have reduced subtle fungal
diseases from being established on the crop.
Removing all leaves from plants at pod set and full bloom resulted
in substantial loss under both conditions probably because at these
stages the developing pods were deprived of photosynthates normally
manufactured in these leaves. The pods which developed probably
utilized reserved photosynthates initially and thereafter photosynthates
which were produced from the few leaves which were formed after
defoliation. At the primary leaf stage, total defoliation slowed down
the growth rate of the plants. Under greenhouse conditions, recovery
may have been slow and plants may have been etiolated due to
insufficient light, hence the higher yield loss. In the field, total
defoliation at the first trifoliate leaf stage through flower bud
formation resulted in 74 and 73% yield loss. This loss in yield is
essentially similar in magnitude indicating that the sensitivity of
plants at these stages was more or less the same.
Results obtained in these experiments seem to show yield increases
due to defoliation. This may have been due to chance effects resulting
from many factors including plant characteristics and the environment.
Generally, data show tendency to lower production and hence dollar
values with increased defoliation. The decrease in production due to
defoliation may have been to enhancement of pathogen entry through
wounds made during manual defoliation. Defoliating with scissors also
led to water loss through direct evaporation. Defoliation also reduced
the leaf area for photosynthesis. One can only suspect that the


TABLE 36. Nematode genera and disease severity values found in plots fumigated with metam-sodium.
Figures are means of 3 replicates (Preplant soil samples).
Defoliation
(proportion of
level
foliage)
Disease severity
(maximum
Metam-sodium proportion of
(Liters/ha) foliage infected)
Helicotylenchus
(No./lOO ml)
Meloidogyne
(no./100 ml)
Rotylenchulus
(No./lOO ml)
0
0
0.02
0
0
19
0
935
0.05
0
0
16
0.25
0
0.04
5
0
5
0.25
935
0.04
0
0
2
0.50
0
0.03
0
0
3
0.50
935
0.03
5
0
2
0
0
0.01
5
0
36
0
935
0.01
0
0
15
0.25
0
0
5
0
37
0.25
935
0.03
0
0
15
0.50
0
0.0
5
0
37
0.50
935
0.0
0
0
15
0
0
0.02
0
0
13
0
935
0.04
5
0
27
0.25
0
0.02
0
0
9
0.25
935
0.02
0
0
9
0.50
0
0.02
0
5
2
0.50
935
0.5
0
0
12
0
0
0.04
10
0
12
0
935
0.02
10
0
45
0.25
0
0.03
0
0
6
0.25
935
0.02
0
0
3
0.50
0
0.03
10
0
30
0.50
935
0.01
0
0
40
126


190
Shorey, H. H. and I. M. Hall. 1963. Toxicity of chemical and microbial
insecticides to pest and beneficial insects on poled tomatoes. J.
Econ. Entomol. 56: 813-817.
Shurtleff, M. C. 1966. How to control plant diseases in the home and
garden. Iowa State Univ. Press, Ames, Iowa. 529 pp.
Sinclair, J. B. and M. C. Shurtleff (eds.). 1975. Compendium of
Soybean Diseases. Amer. Phytopath. Soc., St. Paul. Minnesota. 69
pp.
Singh, D. B., P. P. Reddy, V. R. Rao, and R. Rajendram. 1981a. Culti-
vars of French beans resistant to root-knot nematode, M. incognita.
Trop. Pest Management 27: 29-31.
Singh, D. B., P. P. Reddy, and S. R. Sharma. 1981b. Effect of root-
knot nematode, M. incognita on Fusarium wilt of French beans.
Indian J. Nematol. 11: 84-85.
Singh, N. D. and K. M. Farrel. 1972. Occurrence of Rotylenchulus
reniformis in Trinidad, West Indies. Plant Dis. Rep. 56: 551.
Sosa Moss, C. and J. M. Torres. 1973. Respuesta de frijol ejotero a 7
niveres de poblacin de M. incognita. Mematropica 3: 17 (Abstr.).
Sosa Moss, C. and H. Wrish. 1973. Uso de melaza de cana en fijol
ejutero para combatir M. incognita. Nematropica 3: 18 (Abstr.)
Stall, W. M. and M. Sherman. 1983. Snap bean production in Florida.
Fla. Coop. Ext. Service Circ. No. 100. 4 pp.
Stavely, J. R. 1984. Pathogenic specialization in Uromyces phaseoli in
the United States and rust resistance in beans. Plant Dis. 68:
95-99.
Steadman, J. R., C. R. Maier, H. F. Schwartz, and E. D. Kerr. 1975.
Pollution of surface irrigation waters by plant pathogenic
organisms. Water Res. Bull. 11: 796-804.
Stickler, F. C. and A. W. Pauli. 1961. Leaf removal in grain sorghum.
1. Effects of certain defoliation treatments on yield and compon
ents of yield. Agron. J. 53: 99-102.
Stoetzer, H. A. I. and M. E. Omunyin. 1983. Controlling bean pests and
diseases, pp. 22-24. In: Kenya Farmer (August Special Issue:
Food beans in Kenya).
Taylor, A. L. 1965. Los pequeos pero destructores nematodes, pp.
929. In: Enfermedades de las plantas, USDA Ed., Herrero, Mexico.
Taylor, A. L. and J. N. Sasser. 1978. Biology, Identification and
Control of Root-knot Nematodes. North Carolina Graphics, Raleigh,
111 pp.


0
help and moral support. I owe a lot to my fellow students for their
suggestions and sense of humor.
Sincere thanks are extended to the Malawi government and USAID for
the scholarship.
Finally, the most special and loving acknowledgement is owed to my
wife Abigail, for her love, encouragement, and patience throughout the
program. I would like to end with my sincere thanks to Olivia and
Kondwarri, my children, for putting-up with my absence.
iii


54
strategies. One way of elucidating this relationship has been manual
defoliation of plants to simulate pest damage (Douglas et al., 1981;
Galvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho,
1983; Kalton et al., 1945; Keularts, 1980; Wit, 1983, Wolk et al.,
1983). Hohmann and De Carvalho (1983) reported that leaf area reduction
of 25, 50, 75, and 100% on the bean cultivar Carioca reduced yield by
11, 20, 20, and 70% respectively when defoliation was done at the pod
formation stage. At the same percentage of leaf area reduction, defoli
ation at initiation of flowering decreased yield by 18, 12, 19 and 55%
respectively. Greene and Minnick (1967) indicated that yield reduction
in 'Harvester' snap beans due to leaf removal began somewhere between
33% and 50% defoliation when plants were defoliated in the bloom or
pre-bloom stages. Working on inderterminate snap beans, Waddill et al.
(1984) noted that the removal of both primary leaves when only primary
leaves were present resulted in yield reduction of up to 65%. Vieira
(1981) reported that 66% defoliation of an indeterminate bean cultivar
at the flowering and pod formation stages was detrimental to yield.
Galvez et al. (1977) observed that total defoliation at the formation of
the first trifoliate leaves reduced yields of the bean cultivars ICA-
Guali and Porrillo-Sentetico by 34% and 49% respectively. These obser
vations indicate that the magnitude of yield loss due to defoliation
depends not only on the severity of defoliation but also on the growth
stage (time) the defoliation takes place and the cultivar of beans
grown. Thus, manual defoliation provides a useful estimate of the
host-pest relationship despite its imprecision in simulating pest damage
(Ruesink and Kogan, 1975).


25
effective against bean rust, their use is regulated by their estimated
cost-effectiveness. Thus, Issa and Arruda (1964) cited by Vargas (1980)
concluded that chemical control of bean rust was not economically
practical in Brazil. This conclusion may apply to most tropical bean-
producing areas. The use of fungicides in highly mechanized
agricultural systems, such as the U.S., may be economically feasible
provided registration conditions are met.
Interaction of Root-knot Nematodes and Other Pathogens
Increased incidence of plant diseases has been reported to be
associated with the presence of root-knot nematodes (Brodie and Cooper,
1964; Carter, 1975a,b; Cauquil and Shepherd, 1970; Minton et al., 1975;
Morrell and Bloom, 1981; Norton, 1960; Reynolds and Hanson, 1957;
Schuster, 1959; Thomason et al., 1959; Van Gundy et al., 1977). Carter
(1975), Cauquil and Shepherd (1970), Norton (1960), Reynolds and Hanson
(1957), and White (1962) reported increase incidence of soreshin of
cotton (Rhizoctonia solani Kuhn), root rot (Pythium debaryanum Hesse)
and Fusarium wilt (Fusarium oxysporum Schlecht) when Meloidogyne incognita
(Kofoid and White) Chitwood was present. Increased incidence of southern
blight, Sclerotium rolfsii Sacc., was observed in soybeans infested with
root-knot nematodes (Minton et al., 1975). The interaction of M.
incognita and bacterial wilt (Corynebacterium fluccumfaciens (Hedges)
Dows.) was reported on beans by Schuster (1959). Van Gundy et al.
(1977) reported the enhancement of the development of R. solani in the
presence of exudates from galls caused by M. incognita. Morrell and
Bloom (1981) reported a significant increase in the percentage of
Fusarium wilt occurrence and vessel infection at 21C in the presence of
M. incognita in tomato. Meloidogyne-Fusarium synergism was also observed


TABLE 12. Regressions equations for the relationship between M. incognita defoliation and yield.
Growth stage plants
were defoliated
Regression equation
a
Linear equations
Primary leaf stage Y =
First trifoliate leaf stage Y =
Third trifoliate leaf stage Y =
Flower bud formation stage Y =
Full bloom stage Y =
Pod set stage Y =
259-22.12N-25.lx (R2 = 0.62**)
256.51-21.04N-32.3x (R2 = 0.59**)
267.04-21.51N-33x (R2 = 0.72**)
258.15-24.17N-30.19x (R2 = 0.64**)
237.26-17.75N-31.34x (R2 = 0.58**)
25176-21.44N-29.77x (R2 = 0.58**)
Quadratic equations
Primary leaf stage Y
First trifoliate leaf stage Y
Third trifoliate leaf stage Y
Flower bud formation stage Y
Full bloom stage Y
Pod set stage Y
241.97 + 2.07N-5.13N2-7.9x-4.3x2 (R2 = 0.66**)
227.11 + 5.39N-5.61N2+8.09x-10.1x2 (R2 = 0.67**)
239.73 2.42N-4.03N2+7.42x-10.1x2 (R2 = 0.79**)
243.49 10.31N-2.94N2-10.53x-4.94x2 (R2 = 0.66**)
204.27-0.57-3.64N2+22.68x-13.5x2 (R2 = 0.79**)
218.66-0.24N-4.5N2+21.65x-12.86x2 (R2 = 6.68**)
aY = Yield (g/plot).
N = Log (M. incognita population + 1).
X = Defoliation level (fraction).
** = R significant at 0.01.


48
O 25 50 75 100
Defoliation Level (%)
Figure 10.. Influence of defoliation on gross dollar values per hectare
of 'Sprite' snap beans defoliated at pod set in the field.


T
H*
GQ
C
T
D
Gross dollar value
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fD
H
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22
to chloroplast membranes. Raggi (1978) reported decreased photosynthe
tic rates in rust-infected plants.
Epidemiology of the Disease
Fromme and Wingard (1921) reported that since rust rarely attacks
pods directly, resulting losses are insidious and difficult to assess.
Yield losses are, however, more likely to be severe when plants are
infected during the prebloom and flowering stages of development (Almeida
et al., 1977; Costa, 1972; Crispin et al., 1976; Nasser, 1976; Wimala-
jeewa and Thavam, 1973; Yoshii and Galvez, 1975). Early infection of
some bean varieties can lead to almost complete crop loss in some
seasons (Fromme and Wingard, 1921; Howland and MaCartney, 1966; Townsend,
1939). Townsend (1939) indicated that total loss of the entire crop due
to rust has occurred in Florida.
The variability in the prevalence of bean rust seasonally and
geographically is partly due to environmental conditions (Augustin et
al., 1972; Gonzalez, 1976; Harter and Zaumeyer, 1941; Harter et al.,
1935; Schein, 1961). Infection by U. phaseoli is favored by prolonged
periods (8-18 hours) of at least 95% RH and moderate temperatures
(15-27C) (Augustin et al., 1972; Gonzalez, 1976, Schein, 1961). The
optimum temperature for uredospore germination was reported to be 14.5C
whereas the optimum temperature for infection was 17C (Harter et al.,
1935). Crispin et al. (1976), Schein (1961), and Zaumeyer and Thomas
(1957), however, reported that any temperatures below 15 may retard
fungal development. Day length and light intensity are also important
factors for the development of the bean rust fungus (Harter and Zaumeyer,
1941).


13
Roberts and Boothroyd, 1984). Meloidogyne arenaria is rarely
encountered in association with beans. Meloidogyne incognita and M.
javanica frequently occur simultaneously on beans (Ngundo, 1977; Saka,
1982; Santacruz, 1983; Singh et al., 1981a). The most serious threat to
bean production is M. incognita (Ngundo, 1977; Singh et al., 1981a;
Sharma and Guazelli, 1982). These nematodes may cause yield losses of
50 to 90% during severe infections (Freire and Ferraz, 1977; Varn and
Galvez, 1974).
The limitation on bean by root-knot nematodes has been reported to
4
be due to extensive root-galling and interference with nitrogen fixation
by Rhizobium spp. (Agudelo, 1980), as well as with water and nutrient
uptake. Root-knot nematode infestations often lead to abbreviated root
systems (Agudelo, 1980; Franklin, 1978). Above-ground symptoms of
root-knot infections include incipient wilting, chlorotic above-ground
plant parts, and stunted growth (Agudelo, 1980).
Epidemiology and Life Cycle of Meloidogyne spp.
Meloidogyne spp. are most abundant in light sandy soils with
adequate drainage and temperatures of 25-30C (Crispin et al., 1976).
Root-knot nematodes are spread by irrigation and flood waters, by
vegetative propagation of plant parts in soil contaminated with eggs and
juveniles, which adhere to farm implements, animals, and man (Agudelo,
1980; Caveness, 1967; Crispin et al., 1976; Steadman et al., 1975;
Vieira, 1967; Villamonte, 1965; Walker, 1965). The length of survival
of root-knot nematodes in the soil varies with the stage of development,
soil type, moisture, temperature, soil aeration, and length of the
fallow period (Navarro and Barriga, 1970; Villamonte, 1965; Walker,
1965).


APPENDIX
EFFECTS OF DEFOLIATION AND FUNGICIDES ON SNAP BEANS
Introduction
Bean rust, caused by Uromyces phaseoli (Pers.) Wint., is one of the
most prevalent diseases on beans (Phaseolus vulgaris L.) grown in
Florida and other bean producing areas of the world (Agudelo, 1980;
Allen, 1983; Cook, 1978; Kidney, 1980). Bean rust is common in warm,
(15-27C) moist weather (Augustin et al., 1972; Gonzalez, 1976;
Shurtleff, 1966). Uromyces phaseoli is an autoecious/monoecious fungus
which infects mainly plants in the legume family (Cook, 1978; Shurtleff,
1966). Symptoms of the disease are confined largely to leaves, but
young stems and branches are occasionally infected. On severe infection
symptoms or signs may appear on pods (Cook, 1978; Vargas, 1980).
Lesions first appear as small slightly raised spots that are light in
color and more often on the lower leaf surfaces. These pustules gra
dually enlarge to 1-2 mm in size and may rupture the leaf epidermis 8
days after infection (Allen, 1983). A ring of secondary sori usually
develops around the original infection locus on susceptible varieties.
Bean rust is commonly controlled by weekly sprays of mancozeb
tank-mixed with sulfur or other of the ethylene bis-dithiocarbamate
fungicides (Pohronezny et al., 1984). A few experimental fungicides are
available for bean rust control. One of these more potent fungicides
for bean rust control is bitertanol. Thus, this study was conducted to
164


BIOGRAPHICAL SCKETCH
Afete Divelias Gadabu was born on 19 March 1949, in Lilongwe,
Malawi. He attended primary school at Kamzimbi Primary School and went
to Bwaila Secondary School where he got his Junior Certificate of
education in 1968. He went to Blantyre Secondary School in 1969 and
obtained a Cambridge School Certificate in 1970. In 1971, he received a
scholarship to the University of Malawi from which he graduated with a
Bachelor of Science in biology and chemistry in 1975.
In the same year he joined the Civil Service as a research officer
in the Department of Agricultural Research of the Ministry of Agricul
ture. In 1976 he received a scholarship to the University of Newcastle
upon Tyne, U.K. He graduated with a Master of Science in applied
entomology in 1978 and returned to Malawi. He worked for the Department
of Agricultural Research as a research entomologist. In 1982, he
received a scholarship to the University of Florida where he has been a
graduate student in the Department of Entomology and Nematology from
1983 to 1986. Upon fulfilling the requirements for the degree of Doctor
of Philosophy, he is returning to Malawi to continue working as a
research entomologist.
194


158
comparable in all three inoculation methods. The extent of galling was,
however, not reflected in yield in the soil-mix inoculated plants.
There were significant differences in snap bean yield among
nematode egg and juvenile densities in the seed-drench method (F =
14.34, P 0.01). This may be due to some juveniles ecloding from the
eggs in the vicinity of the germinating seeds since the nematode eggs
hatched over a period of up to 14 days. Hence many of them may have
been able to penetrate bean radicles soon after hatching. There were no
significant differences in snap bean yield (F = 3.09, P ^ 0.1) among
nematode egg and juvenile levels in the soil-mix inoculation method
probably because some of the eggs and juveniles leached out with irriga
tion water. Moreover, juveniles may not have been able to swim up to
the vicinity of germinating seeds against the downward flow of water.
The seed drench method apparently concentrated nematodes, more, within
the vicinity of the plant root system so the local population density
was higher than if much of the inoculum was scattered through the entire
volume of soil. Gall-inoculated plants produced the lowest yield and
differences in yield among gall treatments were significant at P = 0.01
(F = 26.18). This may be due to the natural environment in the galls
within which juveniles ecloded over a long period.
There was lack of association between yields obtained from
2
seed-drench and soil-mix inoculated plants (X = 7.74, 0.10 < P < 0.20).
This lack of association does not necessarily prove that the yields were
different from each other but there was a chance that they were
different.
The data did not fit the Seinhorst model probably because the
nematode population levels were in a logarithmic progression. In most


TABLE 13. Effects of M. incognita and defoliation on snap bean yield (g/plot). Data are means of 4
replicates.
Snap bean yield
(g/plot) by plant growth
stage
Defoliation
level
Log (M. incognita
population +1)
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower
bud
formation
Full
bloom
Pod
set
0
0
261
278
274
288
238
247
0
3.0
219
247
230
207
188
193
0
4.0
173
163
150
155
122
143
0
5.0
112
96
102
87
87
89
0.25
0
213
166
225
215
224
220
0.25
3.0
186
138
173
148
182
175
0.25
4.0
139
118
141
129
141
135
0.25
5.0
84
60
121
95
110
106
0.50
0
217
235
204
196
187
203
0.50
3.0
172
183
170
145
157
154
0.50
4.0
147
140
153
94
126
111
0.50
5.0
87
107
114
71
98
74
0.75
0
185
188
193
180
145
216
0.75
3.0
145
156
156
144
135
170
0.75
4.0
106
114
87
95
109
136
0.75
5.0
92
80
66
80
69
66
1.0
0
126
46
80
83
48
38
1.0
3.0
87
30
50
50
28
35
1.0
4.0
58
21
29
27
17
23
1.0
5.0
41
12
19
16
17
15
on
ON


59
not greatly superior to that of linear models (Figure 11). Yield
reduction was initiated even by the lowest nematode population.
The data did not fit the Seinhorst (1965) model, which is of the
r p-Tl
form Y = m + (1-m) Z where Y = ratio between yield at nematode
population level p and in the absence of nematodes, m = minimum yield at
very high nematode population levels, T = tolerance limit (the nematode
population level below which yield reduction does not occur), and Z =
the proportion of the plant undamaged in the presence of parasitism or
infection by one nematode (Ferris et al., 1981; Ferris, 1984; Seinhorst,
2
1965 and 1972). The Seinhorst model produced a R -value of only 0.00661,
an indication that the data had a poor fit to the model. Data were,
moreover, linear in trend (Figure 11).
Table 10 shows the gross dollar values per hectare of snap beans.
These values were computed from the price ranges of $6.00 (low), $11.00
(medium), and $20.00 (high) per bushel (13.62 kg) of beans. These were
derived from gross yield and prices given above. They are what the
grower would get without subtracting the cost of production which
includes labor, pesticides, rent, farm machinery depreciation, and/or
interest in loans. The nematode-free plots produced at least twice as
much money as plots with 10,000 or 100,000 eggs and juveniles of M.
incognita per pot which were equivalent to 156 and 1562 eggs and juve
niles per 100 ml soil. Nematode-free plants produced $404, $740 and
$1345 per hectare at the low, medium, and high prices respectively
compared to $326, $598, and $1087 when pots were inoculated with 10 eggs
and juveniles (= 0.16/100 ml soil). This level of M. incognita popula
tion resulted in a 19% loss in gross dollar value per hectare ($77,
$142, and $258 loss at low, medium and high prices, respectively).


143
environmental factors on crop damage functions of root-knot nematodes
has shown that these factors significantly affect the establishment of
the nematodes and the growth of the crop and yield (Ferris, 1980;
McKenry, 1983; Noe and Barker, 1983; Roberts, 1983).
The derivation of mathematical models relating nematode densities
to crop damage has been discussed by Ferris (1980, 1984) and Seinhorst
(1965, 1972). In these models, the relationship between the initial
density of root-infesting nematodes and yield or other growth parameters
of infected plants are expressed the assumptions that (i) up to a
certain density the yield is not affected and (ii) a certain minimum
yield remains unaffected by the nematodes even at the highest densities
(Seinhorst, 1965).
The relationship between M. incognita and other Meloidogyne spp.
initial population densities and plant growth and/or yield has been
reported on tobacco (Barker et al., 1981, Ekanayake and Di Vito, 1984),
beans (Melakberhan et al, 1983), tomato (Barker et al., 1976) and pepper
(Di Vito et al., 1982). In these studies, inoculum consisted of eggs of
M. incognita or other Meloidogyne spp. eggs extracted by the sodium
hypochlorite method (Hussey and Barker, 1973) a factor which may be
critical to the development of damage functions in these studies. Vrain
(1977) evaluated the infectivity of three types of inocula which consis
ted of intact egg masses, eggs extracted with 0.53% NaOCl, and larvae
hatched from NaOCl-treated eggs. The data obtained showed the limita
tion of egg masses, low infectivity from NaOCl-extracted eggs and
sensitivity of larvae to relatively high temperatures.
Sodium hypochlorite dilutions have been used for the sterilization
of the surface of nematodes and their eggs in laboratory studies


173
TABLE 45. Net income (dollars) per hectare of snap beans defoliated at
the primary leaf and flower bud formation stages and sprayed
with various fungicides
Plant growth
Plants were
Defoliation
level(%)
stage
defoliated
Fungicide
Primary leaf
Flower bud formation
0
No fungicide
0
0
0
Bitertanol (7-day)
1805
2092
0
Mancozeb (4-5-day)
929
- 39
0
Mancozeb (7-day)
- 87
- 34
0
Mancozeb (14-day)
- 501
- 662
25
No fungicide
- 811
- 732
25
Bitertanol (7-day)
694
523
25
Mancozeb (4-5-day)
328
- 169
25
Mancozeb (7-day)
8
191
25
Mancozeb (14-day)
- 370
- 789
50
No fungicide
-1151
-1491
50
Bitertanol (7-day)
968
418
50
Mancozeb (4-5-day)
118
- 143
50
Mancozeb (7-day)
-1046
- 741
50
Mancozeb (14-day)
- 763
-2123
75
No fungicide
-1020
-1412
75
Bitertanol (7-day)
- 209
- 471
75
Mancozeb (4-5-day)
- 745
-1529
75
Mancozeb (7-day)
-1028
-1316
75
Mancozeb (14-day)
- 946
-2489
100
No fungicide
-1125
-2040
100
Bitertanol (7-day)
-1308
-1203
100
Mancozeb (4-5-day)
-1791
-1189
100
Mancozeb (7-day)
-2284
-2074
100
Mancozeb (14-day)
-1809
-2567


145
For the egg-hatch test, 1 ml of M. Incognita eggs and juveniles
suspended in tap water was put in a watch glass and 2 ml of tap water
was added to the suspension. The initial number of eggs and juveniles
was determined by counting them under a dissecting microscope. The
watch glasses were kept at room temperature (24-30C). Subsequent egg
hatch in each treatment were assessed every 2 days until hatching
levelled off (Vrain (1977).
The inoculation method studies were established on 31 August 1984,
in 1-quart side-drain black plastic pots filled with 1 L of soil (1 part
(R)
sand to 3 parts Palmetto Rich Earth' The soil was inoculated with
eggs and juveniles extracted with the 0.525% NaOCl solution. One series
of pots was inoculated by thoroughly mixing the inoculum with the soil.
The other series was inoculated by drenching the seeds with the
inoculum. A third series was inoculated by placing the appropriate
number of galls in the pot. Each gall contained an average of 246 eggs
and juveniles.
The egg and juvenile populations investigated were 0, 10, 100,
1,000, 10,000, and 100,000 per pot and the numbers of galls were 0, 1,
10, 100, and 500 galls per pot. Since each gall contained an average of
246 eggs and juveniles the gall inoculum was, therefore, equivalent to
0, 246, 2,460, 24,600 and 123,000 eggs and juveniles per pot. Three
seeds were planted in each pot and plants were thinned to one plant/pot
after germination. Treatments were replicated four times in a randomized
complete block. Pots were placed on corrugated benches 0.91 m high in
an open greenhouse and watered twice daily using an automatic time-
controlled water mist-forming system. Beans were harvested on 25


TABLE 10. The influence of M. incognita on the gross dollar value per
hectare of snap beans grown in a greenhouse.
Log (M. incognita
population +1)
Price range3
Gross dollar values
per hectare
0
low
404
medium
740
high
1345
1.04
low
326
medium
598
high
1087
2.0
low
248
medium
456
high
828
3.0
low
222
medium
407
high
740
4.0
low
215
medium
395
high
718
5.0
low
167
medium
307
high
558
a
Low = $6; medium = $11; high = $20/13.62 kg of snap beans.


129
Figure 23. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at 7 day intervals, defoliated at
various levels and soil fumigated with metam-sodium.
13 = no defoliation, no metam-sodium and mancozeb + sulfur at 7-day inter
vals, 14 = no defoliation, 935 L/ha metam-sodium and mancozeb + sulfur at
7-day intervals, 15 = 25% defoliation, no metam-sodium and mancozeb 4-
sulfur at 7-day intervals, 16 = 25% defoliation, 935 L/ha metam-sodium
and mancozeb + sulfur at 7-day intervals, 17 = 50% defoliation, no metam-
sodium and mancozeb + sulfur at 7-day intervals, and 18 = 50% defoliation,
935 L/ha metam sodium and mancozeb + sulfur at 7-day intervals.


930
880-
780-
680
580
480
380
280
180
-gure
137
y = 650.4-772.4x-45x2+462.4x3
R2=0.46
0.25 0.5
Defoliation Level
0.75
The general relationship between defoliation and
snap bean yield at all metam-sodium rates.


Proportion of Leaf Infected
86
Figure 16. Disease progress curves for Uromyces phaseoli on snap
beans sprayed with mancozeb at various frequencies
(Trial 1).
A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals,
D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur
at 14-day intervals.


113
TABLE 25. The effect of metam-sodium on gross dollar values per hectare
of beans.
Price Range
Vapam
(liters/ha)
low
medium
high
0
176
323
587
47
292
535
972
94
318
583
1061
187
333
610
1094
281
285
522
948
374
346
634
1153


120
TABLE 32. The effect of defoliation and metam-sodium
value per hectare of snap beans.
on the
gross
dollar
Gross
dollar
values
Defoliation
level
Metam-sodium
by
price
range
(proportion of
foliage)
(Liters/ha)
Low
Medium
High
0
0
267
489
889
0
47
269
494
898
0
94
361
662
1204
0
187
416
763
1388
0
374
278
510
927
0.25
0
230
421
765
0.25
47
204
374
681
0.25
94
292
535
972
0.25
187
197
362
658
0.25
374
207
279
690
0.5
0
129
237
431
0.5
47
124
227
412
0.5
94
182
333
605
0.5
187
162
296
539
0.5
374
164
301
547
0.75
0
129
236
429
0.75
47
91
167
303
0.75
94
135
247
450
0.75
187
95
174
315
0.75
374
164
255
463


156
There were more eggs and juveniles extracted from tomato than from
bean roots (Table 40). This may be an indication that tomato is a more
comparitible root-knot nematode host than beans. Consequently the
nematodes may have reproduced at a faster rate on tomato than on beans.
It is possible that this difference in extracted eggs and juveniles may
be a reflection of age differences. Eggs and juveniles were extracted
63 days after inoculation from bean roots whereas eggs and juveniles
were extracted from tomato plants 50 days after inoculation. Thus,
there might have been more juveniles which had already emerged from bean
root galls than on tomato root galls. Consequently, more juveniles may
have been washed off along with soil from bean roots which may have lead
to the low numbers of eggs and juveniles extracted. Moreover, weighing
did not imply the same number of galls or eggs on the root material
used. Weighing may thus have biased the number of egg masses used
towards the tomato source. There is little information in the
literature pertaining to variation in the number of eggs per gall in
various hosts. This suggests that variation in egg numbers per gall on
various hosts be investigated.
More juveniles of M. incognita may have emerged on the day of
extraction from NaOCl-treated eggs than from water-treated eggs, probab
ly because the NaOCl reacted with the egg shell, thereby inducing early
hatch in eggs with fully developed juveniles. This effect of NaOCl on
egg hatch had, however, some negative effects when the concentration was
above 0.525%. The negative effect of NaOCl on eggs was particularly
evident with the 2.6% NaOCl treated eggs. This high concentration of
NaOCl gave rise to numbers of juveniles comparable to those obtained
from water-treated eggs, despite the 2.6% solution having extracted the


MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS
BY
AFETE DIVELIAS GADABU
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1986

ACKNOWLEDGEMENTS
I wish to express my sincere gratitude to many people for their
support and guidance:
Dr. V. H. Waddill, my advisor and committee chairperson, for his
guidance, encouragement, suggestions, and assistance throughout my
Doctor of Philosophy program at the University of Florida.
Dr. J. R. Strayer, co-chairperson, for his constructive suggestions,
encouragement, and providing materials for computer work whenever I
needed them.
Dr. R. McSorley, for serving on my committee, encouragement,
suggestions, and invaluable assistance with statistical data analysis.
Dr. K. Pohronezny, for serving on my committee, encouragement,
suggestions, and assistance with data analysis.
Dr. S. H. Kerr, for his guidance and constructive suggestions
during my Ph.D. program.
I am indebted to Dr. R. D. Berger for his advice, helpful sugges
tions on disease progress analysis, and allowing me to use his computer
programs.
The invaluable technical assistance and friendship of Nancy Shivers,
Diane Putnal, W. (Hank) Dankers, Jorge Parrado, James Reynolds, John
Sarvich, and Ingeborg Stough during the long and difficult field and
laboratory hours are appreciated. I am also indebted to William Meyers,
Joyce Francis, Jeanette Viola, and all staff at Homestead TREC for their
ii

0
help and moral support. I owe a lot to my fellow students for their
suggestions and sense of humor.
Sincere thanks are extended to the Malawi government and USAID for
the scholarship.
Finally, the most special and loving acknowledgement is owed to my
wife Abigail, for her love, encouragement, and patience throughout the
program. I would like to end with my sincere thanks to Olivia and
Kondwarri, my children, for putting-up with my absence.
iii

TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS
ABSTRACT vi
CHAPTER I INTRODUCTION 1
CHAPTER II LITERATURE REVIEW ON DEFOLIATION, AND THE IDENTI
FICATION AND CONTROL OF ROOT-KNOT NEMATODES (Meloidogyne
spp.) AND BEAN RUST (UROMYCES PHASEOLI [PERS.] WINT.) ... 7
Introduction 7
Simulated Leaf Damage on Crop Plants 8
Nematodes Associated with Beans 10
Occurrence and Importance of Root-knot Nematodes ... 10
Epidemiology and Life Cycle of Meloidogyne spp. ... 13
Control of Root-knot Nematodes 15
Identification of Root-knot Nematodes 16
The Importance of Bean Rust 18
Identification and Etiology of the Pathogen 19
Symptoms 20
Epidemiology of the Disease 22
Control of the Disease 2A
Interaction of Root-knot Nematodes and other Pathogens 25
CHAPTER III THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN
YIELD 27
Introduction 27
Materials and Methods 29
Results 31
Discussion 49
CHAPTER IV THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION
ON SNAP BEANS 52
Introduction 52
Materials and Methods 55
Results 57
Discussion 73
CHAPTER V THE EFFECT OF BEAN RUST ON SNAP BEANS 77
Introduction 77
Materials and Methods 80
Results 82
Discussion 96
CHAPTER VI THE EFFECT OF DEFOLIATION, VAPAM, AND BEAN RUST ON
SNAP BEANS 100
Introduction 100
Materials and Methods 102
Results 106
Discussion 139
iv

CHAPTER VII THE EFFECT OF INOCULATION SYSTEM ON Meloidogyne
incognita RACE ESTABLISHMENT ON BEANS 142
Introduction 142
Materials and Methods 144
Results 144
Discussion 155
CHAPTER VIII SUMMARY AND CONCLUSIONS 160
APPENDIX THE EFFECT OF FUNGICIDES ON SNAP BEANS 164
Introduction 164
Materials and Methods 165
Results 166
Discussion 174
LITERATURE CITED 176
BIOGRAPHICAL SKETCH 194
v

Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fullfillment
of the Requirements for the Degree of
Doctor of Philosophy
MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS
By
Afete Divelias Gadabu
May, 1986
Chairman: Dr. Van H. Waddill
Major Department: Entomology and Nematology
During 1984 and 1985, a number of greenhouse and field experiments
were carried out at the Tropical Research and Education Center at
Homestead in Dade County, Florida, to determine the effect of manual
defoliation, Meloidogyne incognita, bean rust, and various other nema
todes on Sprite' snap beans (Phaseolus vulgaris L.). Treatments
consisted of total defoliation (100%), 0%, 25%, 50%, and 75% defoliation
at various plant growth stages; 0, 10, 100, 1,000, 10,000 and 100,000 M.
incognita eggs and juveniles per pot, fungicide sprays which included
bitertanol at 7-day intervals, mancozeb tank-mixed with sulfur at
4-5-day, 7-day, and 14-day intervals respectively, and soil fumigation
with metam-sodium at 0, 47, 94, 187, 281, and 374 L/ha and a separate
one at 935 L/ha. Experiments were conducted with each series of
treatments as well as with combinations of two or more types of
treatments simultaneously.
vi

Manual defoliation caused the highest snap bean yield losses at
full-bloom and pod-set both in the field and the greenhouse experiment.
Snap bean yield loss was observed at the 25% defoliation level in both
experiments. Total defoliation resulted in the highest yield losses.
Yield was negatively correlated to Meloidogyne incognita (Kofoid
and White) Chitwood population levels when plants were grown in pots and
the nematodes were used alone. Yield was also inversely related to
nematode population levels when manual defoliation occurred on nematode
inoculated plants. Yield loss was observed on plants grown in soil
inoculated with 10 eggs and juveniles per pot.
The bean rust disease, Uromyces phaseoli (Pers.) Wint., was
manipulated by fungicide sprays. It was observed that plants with the
highest disease severity gave the lowest yield whereas plants which were
virtually disease free had the highest yield. Generally, fungicide
sprays increased yields. In some cases yield increases were not high
enough to pay for the extra cost of fungicide sprays.
Soil funigation with metam-sodium increased yields slightly. The
optimum metam-sodium application rate was 187 L/ha.
Yield was affected most by the bean rust disease when defoliation,
metam-sodium and the disease were used simultaneously.
vii

CHAPTER I
INTRODUCTION
Beans, Phaseolus vugaris L., are the major protein source for many
people in the world, especially in developing countries (Yamaguchi,
1978). Consequently beans are considered an important crop in the
tropics, subtropics, and warm temperate areas of the world (Zaumeyer and
Meiners, 1975). Zaumeyer and Meiners (1975) stated that the leading
world bean producers were Brazil, Mexico, and the United States of
America (U.S.A.) in descending order. Beans are grown for fresh market,
processing, and dry seed. In some countries the leaves are also used as
a vegetable.
Snap beans are known by various names in different areas. These
names include French beans, green beans, pole beans, string beans, and
wax beans (Yamaguchi, 1978). Snap beans are grown in many states in the
U.S. where 132,720 metric tons (mt.) were harvested for fresh-market
consumption from 35,847 hectares (ha.) in 1981 (Anon., 1981). In the
same crop year, 671,640 mt. of snap beans were harvested for processing
from 93,324 ha. (Anon., 1981). The gross value for snap beans in 1981
was $199,282,000 in the U.S. (Anon., 1981).
Florida is the largest producer of fresh-market snap beans in the
U.S., producing nearly 40% of the crop (Anon., 1972, 1982; Rose, 1975;
Ware and McCollum, 1980). In the 1981-82 production year, Florida
produced 60,600 mt. of bush and pole beans from 37,206 ha. (Anon.,
1982). Southeast Florida is the major producing area for snap beans in
1

2
the state with the greatest production in Dade County (Anon., 1982).
Gadsden, Marion and Palm Beach counties and parts of the west central
area also produce some snap beans (Anon., 1982). Rose (1975), Ware and
McCollum (1980), and Anon. (1982) reported that the winter demand for
fresh market snap beans in population centers to the north of Florida is
usually met by supplies from the southern districts of Florida.
Whereas Florida is the largest producer of fresh market snap beans,
Wisconsin is the leading producer of snap beans for processing (Kobriger
and Hagedorn, 1983). Michigan, New York, and Oregon also produce more
snap beans for processing than Florida (Kobriger and Hagedorn, 1983;
Ware and McCollum, 1980).
Snap bean production has some inherent pest problems (Anon., 1982;
Rose, 1975). Fresh market snap bean yield, however, increased from 31
cwt/acre in the 1947-1952 period to 37 cwt/acre in the 1967-1972 period,
in the U.S., despite these problems (Anon., 1972). This increase in
yield has been ascribed to the advent of synthetic organic pesticides
during and after World War II. In Florida, snap bean yields were,
however, on the decline during the same period (Rose, 1975). The
decline was associated mainly with adverse weather conditions (Anon.,
1982). Galvez et al. (1977) and Vargas (1980) stated that insect pests,
such as leafminers, leafrollers, corn earworm, Mexican bean beetle, and
others cause tremendous losses in bean yield. Many of these insects
feed on the leaves reducing the photosynthetic tissue of the plant.
These pests have been more-or-less controlled by insecticide sprays
(Acland, 1971; Iraneta and Rodrigez, 1983).
Bean rust, Uromyces phaseoli, (Pers.) Wint., is one of the most
important diseases of beans in many bean producing areas of the world

3
(Acland, 1971; Cook, 1978; Crispin and Dongo, 1962; Iraneta and
Rodrigez, 1983; Martinez, 1983; Schwartz et al., 1979; Stoezer and
Omunyin, 1983; Vargas, 1980, Zaumeyer and Thomas, 1957). Other major
bean disease include anthracnose (Colletotrichum lindemuthianum Sacc.
and Magn.), angular leaf spot (Isariopsis griseola Sacc.), halo blight
(Pseudomonas phaseolicola (Burk.) Dows.), common blight (Xanthomonas
phaseoli (E. F. Smith)), and bean common mosaic virus (Acland, 1971;
Allen, 1983; Martinez, 1983; Stoetzer and Omunyin, 1983). Martinez
(1983) stated that root rots caused by Macrophomina phaseolina (Tassi)
Gold.,, Sclerotium rolfsii Sacc., Rhizoctonia solani Kuhn, Pythium spp.
and Fusarium spp. were among the most important diseases of beans.
Fifty-seven races of the bean rust fungus, Uromyces phaseoli, have
been reported in the U.S. (Laundon and Waterston, 1965). The number of
races of _U. phaseoli is, however, not fixed, due to controversy on what
constitutes a physiologic race of a pathogen (Crispin and Dongo, 1972;
Davidson and Vaughan, 1963; Groth and Shrum, 1971; McMillan, 1972).
Uromyces phaseoli has been reported to reduce the translocation of
photosynthetic products from the foliage to the roots and developing
seeds (Daly, 1976; Livne, 1962; Montalbini, 1973; Zaki and Durbin,
1965). The reduction of photosynthetic products translocation is
exacerbated by increased water loss through the damaged leaf cuticle
despite a decrease in transpiration (Vargas, 1980). Water loss in
creases as infection becomes more severe. Infection by IJ. phaseoli
predisposes bean plants to other pathogens such as Pythium spp.,
Rhizoctonia spp. F_. phaseolicola, £. lindemuthianun and many others
(Vargas, 1980).

4
Root-knot nematodes, Meloidogyne spp., also infect bean plants.
Meloidogyne spp. are most prevalent in light sandy soils with good
drainage and moderately warm temperatures (25-30C) (Crispin et al.,
1976). Roberts and Boothroyd (1984), however, stated that M. incognita
(Kofoid and White) Chitwood is more common in southern states of the
U.S. and M. hapla Chitwood is commonly found in northern states.
Meloidogyne spp. reportedly limit the production of beans by interfering
with nitrogen fixation by Rhizobium spp. and causing root galls (Ngundo,
1977; Sharma and Guazelli, 1982; Singh et al. 1981 a). Meloidogyne
incognita has been observed to predispose beans to Fusarium wilt (Singh
et al., 1981 b). Severe root-knot nematode infections may lead to
50-90% yield loss (Freire and Ferraz, 1977; Ngundo, 1977; Varn and
Galvez, 1974).
Control of these pests and diseases has been based on chemical
pesticides and cultural methods (Acland, 1971; Allen, 1983; Carvalho et
al., 1981; Martinez, 1983; Rhoades, 1976; Robbins et al., 1972; Shorey
and Hall, 1963; Stoetzer and Omunyin, 1983; Villamonte, 1965; Yoshii,
1977; Zaumeyer and Meiners, 1975). The use of resistant varieties and
flooding has been part of the management strategies for bean rust and
root-knot nematodes (Crispin et al., 1976; Martinez, 1983; Ngundo, 1977;
Singh et al., 1981a; Vieira, 1967).
One of the problems research scientists are confronted with, in
crop pest management, is the development of multi-pest threshold levels
to be used in determining the type of management strategies and the
extent to which these pest complexes have to be controlled so that
maximum yield is obtained with minimum disruption of the environment.
Pesticides are, however, the main means of controlling pest complexes on

5
many crops (Allen, 1983; Stoetzer and Omunyin, 1983). Metcalf (1975)
reported that repeated use of pesticides has sometimes led to a decline
in acreage or yield of the crop. As a consequence of this subtle
decline in crop yield due to repeated pesticide use, an integrated pest
management (IPM) approach has been advocated (Huffaker and Smith, 1980;
Waddill et al., 1981). IPM aims at better understanding of the signifi
cance of biological, ecological, and economic processes in the production
of the crop, and the population dynamics of the pest complex, their
predators and parasites, and other factors affecting the system in the
field (Huffaker and Smith, 1980).
Research was undertaken during the period 1984-1985 to investigate
the effects of manual defoliation, root-knot nematode population levels,
and bean rust on snap bean yield. These factors were used separately,
in combinations of two or more of them together, and were studied as
stated in order to determine their threshold levels singly and when they
occurred simultaneously.
Specific experiments were conducted to determine
1. the effect of defoliation on yield of snap beans under field and
open greenhouse conditions,
2. the effect of Meloidogyne incognita population levels on snap bean
yield,
3. the effect of bean rust, Uromyces phaseoli on snap bean yield,
4. the effect of defoliation and M. incognita population levels on
snap bean yield,
5. the effect of defoliation and several nematode genera on snap bean
yield,

6
6. the effect of defoliation, nematodes and bean rust on snap bean
yield, and
7. the effect of inoculation system on the establishment of M.
incognita in beans (Phaseolus vulganis L.).

CHAPTER II
LITERATURE REVIEW ON DEFOLIATION AND THE IDENTIFICATION AND CONTROL
OF ROOT-KNOT NEMATODES (MELOIDOGYNE SPP.) AND BEAN RUST
(UROMYCES PHASEOLI [PERS.] WINT.)
Introduction
Bean insect pests such as leafminers (Liriomyza spp.), bollworms
(Heliothis armigera Hbn.) and leaf rollers (Urbanus proteus L.) as well
as diseases including rust (Uromyces phaseoli [Pers.] Wint.)> web blight
(Thanatephones cucumeris [Frank] Donk) and angular leaf spot (Isariopsis
griseola Sacc.) not only destroy plant foliage but also cause physio
logical damage in some cases (Acland, 1971; Galvez et al., 1977).
Root-knot nematodes, Meloidogyne spp., cause prolific galls on the root
system of plants which may lead to the following above-ground symptoms:
incipient wilting, stunted growth, and chlorotic leaves, often with
burnt out edges (Agudelo, 1980). A combination of these organisms on
beans usually leads to great losses in yield. Control of these pest
problems has been mainly by the use of chemical pesticides (Acland,
1971; Agudelo, 1980).
This review is a summary of the problems encountered in the identi
fication of root-knot nematodes and bean rust and the use of simulated
leaf damage on beans.
Simulated Leaf Damage on Crop Plants
Bean plants are susceptible to defoliation by insects, diseases,
hail, moisture stress, and mechanical injury resulting from farm
7

8
machinery. Sometimes defoliation is initiated by chemical defoliants to
facilitate harvesting (McGregor et al., 1953). In most cases defolia
tion occurs due to pest problems and adverse environmental conditions.
To reduce defoliation by pests, preventative spray programs are usually
followed (Greene and Minnick, 1967). These sprays are usually applied
regardless of the anticipated crop loss. Hence, it would be desirable
to determine the relationship between defoliation levels and yield
losses to maximize the efficiency and rationale for spray treatments.
A wide range of yield losses due to artificial defoliation has been
observed on various bean cultivars (Edje and Mughogho, 1976a, 1976b;
Edje et al., 1973, 1976; Garvez et al., 1977; Greene and Minnick, 1967;
Hohmann and De Carvalho, 1983; Vieira, 1981; Waddill et al., 1984).
Edje et al. (1973, 1976), Edje and Mughogho (1976a, 1976b), Vieira
(1981) and Waddill et al. (1984) manually defoliated indeterminate bean
cultivars. Waddill et al. (1984) reported that complete defoliation
when only primary leaves were present reduced yield by about 65% and
repeated weekly defoliation of 50% resulted in 34% yield loss. Vieira
(1981) reported that 66% leaf area removal during the flowering and pod
formation stages was detrimental to yield. Galvez et al. (1977) ob
served that 100% defoliation at formation of the first trifoliate leaves
decreased yields of the bean cultivars ICA-Guali and Porrillo-Sentetico
by 34% and 49% respectively. Greene and Minnick (1967) indicated that
yield reduction in snap beans begins somewhere between 33% and 50%
defoliation when defoliation occurs in the prebloom and bloom stages,
respectively. Hohmann and DeCarvalho (1983) reported that removal of
25, 50, 75, and 100% of the leaf area at the pod formation stage reduced

9
yield by 11, 20, 20, and 70% respectively. At the same levels of leaf
area reduction, defoliation at initiation of flowering decreased yield
by 18, 12, 19, and 55% respectively. At the formation of the third
trifoliate leaves only total defoliation affected the yield.
Kalton et al. (1945) and Weber (1955) reported that 50% and 75%
leaf removal in soybeans had little effect on yield when defoliation
occurred at the prebloom stage. Significant yield losses were, however,
observed when plants were heavily defoliated at the bloom or pod set
stages (Begum and Eden, 1963, 1964; Camery and Weber, 1953; Kalton et
al., 1945; McAlister and Krober, 1958). Todd and Morgan (1972) observed
significant yield reduction on soybeans with 33, 67, and 100% leaf
removal at 2 wk, 4 wk, and 8 wk after first bloom. Wilkerson et al.
(1984) reported that all defoliations on Florunner' peanuts resulted in
lower stem weight to length ratios and lower pod numbers and weights.
It was observed that defoliation altered the normal partitioning of
photosynthates between plant parts in peanuts. Wit (1983) reported that
during the most sensitive period (July) in the Netherlands, 60% defolia
tion induced a maximum yield reduction of 35% in Brussel sprouts. He
also noted that when partial defoliation was carried out 15 wk after
transplanting or later, no effect on yield was observed. Douglas et al.
(1981) observed a grain yield reduction of 77% in corn when complete
defoliation was carried out at silking. Grain yield losses decreased
with delay in defoliation toward maturity. Less severe defoliations,
however, resulted in smaller reductions in yield. Generally, grain
yield was tolerant of post-silking defoliation and yield losses
exceeding 20% were recorded only after 67% of the leaves were removed.
Defoliation action thresholds for tomato for the prebloom and postbloom

10
stages have been established at 30% and 50% respectively (Keularts,
1980). Extensive research on artificial defoliation effects on tomato
has been conducted by various writers (Keularts, 1980; Wolk et al.,
1983). The effect of leaf removal has also been studied on cotton
(Ludwig, 1926), grain sorghum (Stickler and Pauli, 1961) and wheat and
oats (White, 1962; Wotmack and Thurman, 1962).
Nematodes Associated with Beans
Many nematodes have been found in and around the roots of beans
(Agudelo, 1980; Allen, 1983). Among the nematodes associated with
beans, root-knot nematodes, Meloidogyne spp., are the most important in
tropical and subtropical regions (Agudelo, 1980; Allen, 1983). Table 1
shows the nematode species associated with beans in various bean produc
ing areas (Agudelo, 1980; Ayala and Ramirez, 1964; Bridge, 1973; Bridge
et al., 1977; Castillo and Litsinger, 1978; Caveness et al., 1975,
Feakin, 1973; Hague, 1980; Sinclair and Shurtleff, 1975; Singh and
Farrell, 1972).
Of the four main species of Meloidogyne, M. hapla Chitwood has a
more northerly distribution than M. arenaria (Neal) Chitwood, M.
incognita (Kofoid and White) Chitwood, and M. javanica (Treub) Chitwood
which are cosmopolitan in warmer regions (Allen, 1983; Roberts and
Boothroyd, 1984). The distribution of the other nematode genera is
shown on Table 1.
Occurrence and Importance of Root-knot Nematodes
The most common species of root-knot nematodes are Meloidogyne
arenaria, M. incognita, M. hapla, and M. javanica. Meloidogyne arenaria,
M. incognita, and M. javanica occur worldwide warmer regions whereas M.
hapla has a more northerly distribution (Agudelo, 1980; Allen, 1983;

TABLE 1. Nematodes commonly found In association with roots of beans.
Species
Distribution
Reference
Meloidogyne arenaria (Neal) Chitwood
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980; Castillo
and Litsinger, 1980
M. hapla Chitwood
N. Europe, Japan, U.S.A., Agudelo, 1980; Sinclair
Canada, & warmer regions of and Shurtleff, 1975
Africa and Middle East
M. incognita (Kofoid & White) Chitwood
M. javanica (Treub) Chitwood)
Pratylenchus brachyurus (Godfrey) Filipjev
Aphelenchoides spp.
Rotylenchulus reniformis Linford & Oliveira
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980;
& Shurtleff,
Sinclair
1975
Cosmopolitan
Agudelo, 1980;
1973
Bridge,
Nigeria
Agudelo, 1980;
et al., 1977
Bridge
W. Africa, U.S.A., Indonesia,
Agudelo, 1980;
Ayala &
Philippines Ramirez, 1964; Singh
& Farrell, 1972
Helicotylenchus spp.
Cosmopolitan
Agudelo, 1980; Bridge,
1973; Hague, 1980
Criconemella spp.
Widespread
Agudelo, 1980; Feakin,
1973

TABLE 1. Continued.
Species
Distribution
Reference
Belonolaimus spp.
Southeastern U.S.A.
Agudelo, 1980; Feakin,
1973; Sinclair &
Shurtleff, 1975
Trichodorus spp.
Widespread
Agudelo, 1980; Feakin,
1973
Xiphinema spp.
Widespread
Agudelo, 1980; Caveness
et al., 1975; Feakin,
1973

13
Roberts and Boothroyd, 1984). Meloidogyne arenaria is rarely
encountered in association with beans. Meloidogyne incognita and M.
javanica frequently occur simultaneously on beans (Ngundo, 1977; Saka,
1982; Santacruz, 1983; Singh et al., 1981a). The most serious threat to
bean production is M. incognita (Ngundo, 1977; Singh et al., 1981a;
Sharma and Guazelli, 1982). These nematodes may cause yield losses of
50 to 90% during severe infections (Freire and Ferraz, 1977; Varn and
Galvez, 1974).
The limitation on bean by root-knot nematodes has been reported to
4
be due to extensive root-galling and interference with nitrogen fixation
by Rhizobium spp. (Agudelo, 1980), as well as with water and nutrient
uptake. Root-knot nematode infestations often lead to abbreviated root
systems (Agudelo, 1980; Franklin, 1978). Above-ground symptoms of
root-knot infections include incipient wilting, chlorotic above-ground
plant parts, and stunted growth (Agudelo, 1980).
Epidemiology and Life Cycle of Meloidogyne spp.
Meloidogyne spp. are most abundant in light sandy soils with
adequate drainage and temperatures of 25-30C (Crispin et al., 1976).
Root-knot nematodes are spread by irrigation and flood waters, by
vegetative propagation of plant parts in soil contaminated with eggs and
juveniles, which adhere to farm implements, animals, and man (Agudelo,
1980; Caveness, 1967; Crispin et al., 1976; Steadman et al., 1975;
Vieira, 1967; Villamonte, 1965; Walker, 1965). The length of survival
of root-knot nematodes in the soil varies with the stage of development,
soil type, moisture, temperature, soil aeration, and length of the
fallow period (Navarro and Barriga, 1970; Villamonte, 1965; Walker,
1965).

14
The life cycle of Meloidogyne spp. has several developmental stages
(Taylor and Sasser, 1978). The adult female lays eggs in a gelatinous
matrix. The first-stage juvenile develops and molts within the egg.
What emerges from the egg is actually the second-stage juvenile, hence
the general belief that root-knot juveniles grow between a series of
three molts into adult males and females (Agudelo, 1980). Root-knot
nematode eggs are oval or ellipsoidal and may be concave on the side.
They measure 30-52 x 67-128 pm (Thorne, 1961). These eggs are usually
protected from dehydration by a gelatinous matrix secreted by the female
(Franklin, 1978; Taylor and Sasser, 1978).
The juvenile stages are vermiform, have a stylet about 10 jum long
and have an overall length of 375-500 pm and a width of 15 pm (Robbins
et al., 1972; Taylor and Sasser, 1978). Males are cylindroid and
measure 0.03-0.36 x 1.2-1.5 mm. The males lack a bursa. Adult females
are pyriform and usually pearly white (visible in roots without magni
fication) The females measure 0.27-0.75 x 0.40 x 1.30 mm and have a
soft cuticle (Franklin, 1978; Taylor, 1965; Walker, 1965). The life
cycle of root-knot nematodes may take 17-57 days, depending on the soil
temperature and the host plant (Tyler, 1933; Taylor and Sasser, 1978).
Infection by and pathogenesis of Meloidogyne spp. are affected by
plant age, plant susceptibility, population size and environmental
factors (Brodie and Dukes, 1972; Gilvonio and Ravines, 1971; Nemec and
Morrison, 1972; Sosa Moss and Torres, 1973). Second stage juveniles of
Meloidogyne spp. enter the plant root system within 2 days after inocula
tion and migrate inter and/or intracellularly through the cortex into
the stele (Dropkin, 1980; Ngundo and Taylor, 1975 b). The juvenile
inserts its head into the vascular system of the root to obtain

15
nutrients from the plant. Plant cells in the vicinity of the nematode
juvenile increase both in number and size (hyperplasia and hypertrophy),
causing the characteristic giant cells (synctia) (Dropkin, 1980; Taylor
and Sasser, 1978). The giant cells usually form near the juvenile's
head by the fusion and enlargement of plant cells in response to
nematode feeding. These giant cells eventually become apparent in the
form of galls on the root system. Injury to plant root systems usually
becomes apparent 10 days after infection. Five to six weeks after
infection, epidermal cells of the roots collapse after females have
deposited eggs near the outer root surface (Ngundo and Taylor, 1975a).
Control of Root-knot Nematodes, Meloidogyne spp.
The economic importance of plant-parasitic nematodes is commonly
assessed by the use of soil fumigants (Mountain, 1965). Usually, an
inverse relationship between yield and nematode numbers is expected
(Sasser et al., 1968). The relationship between yield and nematode
counts is not always inverse (Robbins et al., 1972). In many bean-
producing regions, nematicides are extensively used on a preventative
basis (Agudelo, 1980). The world farming community has many nematicides
available depending on supply and legal registration. These nematicides
include dicholoropropene-dichloropropane (DD), ethylene dibromide (EDB),
phenamiphos, methyl bromide, aldicarb, metam-sodium, and DBCP (Jimenez,
1976; Parisi et al., 1972; Rhoades, 1976; Sosa Moss and Wrihs, 1973).
In these operations, no attempt is made to eradicate nematodes (Thomason
and McKenry, 1975). These nematicide applications are aimed at reducing
the nematode populations by 80-90% in the upper 40-60 cm of the soil and
are considered adequate to provide economic control (Thomason and
McKenry, 1975).

16
Crop rotation has been used to reduce nematode numbers in bean
fields (Agudelo, 1980). Beans are planted once every 2 or 4 years in
rotation with a crop such as corn, which is not particularly susceptible
to many nematodes parasitic on beans. Cover crops such as marigold
(Tagetes minuta), rattle box (Crotalaria spectablilis), or hairy indigo
(Indigofera hirsuta) have been used for this purpose (Eguiguren et al.,
1975; Navarro and Barriga, 1970; Rhoades, 1976; Zaumeyer and Thomas,
1957). Other cultural practices used to reduce nematode numbers include
long fallow periods, deep plowing, and flooding (Crispin et al., 1976;
Vieira, 1967).
There are many bean cultivars resistant to M. incognita (Blazey et
al., 1964; Christie, 1959; Fassuliotis et al., 1970; Hartman, 1968;
Ngundo and Taylor, 1974; Rhoades, 1976; Varn and Galvaz, 1974; Wester
et al., 1958). In some cases resistance to M. incognita is broken by
simultaneous infection of M. incognita and M. javanica (Ngundo, 1977).
Ngundo (1977) reported that seven bean lines were resistant to M.
incognita and M. j avanica when they occurred simultaneously.
Identification of Root-knot Nematodes
Maggenti (1981) and Taylor and Sasser (1978) state that root-knot
nematodes were first described by Berneley in England in 1855. M.
incognita was studied, independently, in the U.S. by Neal and Atkinson
in 1889 (Maggenti, 1981). Maggenti (1981) reported that Neal indicated
that root-knot nematodes occurred in Florida before 1805. During these
early studies, Meloidogyne spp. were described under the species names
Heterodera marioni or H. radicicola (Maggenti, 1981). Chitwood and
Chitwood (1950), as a result of their work on the taxonomy of root-knot
nematodes, placed them under the genus Meloidogyne. Chitwood and
Chitwood recognized five species of Meloidogyne and one subspecies.

17
Esser et al. (1976), however, recognized 35 species in this genus.
Dickson (unpublished) reported that more than 50 species of Meloidogyne
were identified. The number of species in this genus fluctuates due to
various identification procedures used and discovery of new species each
year.
Root-knot nematode speciation is based on perineal patterns, the
distance between stylet knobs and the dorsal esophageal gland opening,
the second-stage juvenile morphology, chromosome number, electrophoresis,
and host range (Maggenti, 1981; Taylor and Sasser, 1978). Host differen
tials are also used to separate races of the same species (Taylor and
Sasser, 1978).
Meloidogyne incognita is the most widely distributed species of
root-knot nematode, comprising 52% of a world collection (40N to 33S)
in areas where annual temperatures are normally within the 18-30C range
(Taylor and Sasser, 1978). This species has four host races as follows:
race 1 does not infect 'Deltapine' 16 cotton, 'NC95' tobacco, and
'Florunner' peanuts; race 2 does not infect 'Deltapine' cotton, and
'Florunner' peanuts; race 3 does not infect 'NC95' tobacco and
'Florunner' peanuts; and race 4 does not infect 'Florunner' peanuts
only. All four races infect 'California Wonder' pepper, 'Charleston
Grey' watermelon and 'Rutgers' tomato (Taylor and Sasser, 1978).
Meloidogyne incognita has a very extensive host range and frequently
coexists with M. javanica (Dickson, unpublished; Santacruz, 1983).
Meloidogyne javanica is the second most widely distributed species,
forming 31% of a world collection (Taylor and Sasser, 1978). Meloidogyne
javanica has no known host races but exhibits variation in chromosome
numbers. M. hapla and M. arenaria comprised 8 and 7% of a world

18
collection respectively (Taylor and Sasser, 1978). These two species
are known to have 2 host races each (Dickson, unpublished).
The Importance of Bean Rust
Bean rust is known to occur whenever beans are grown (Vargas,
1980). Bean rust is the most important bean disease in Central and
South America (Augustin et al., 1972; Crispin and Dongo, 1962; Makram et
al., 1973; Zaumeyer and Meiners, 1975). Bean rust has been reported to
reduce the yield of snap beans in New Zealand, Egypt, and Australia
(Ballantyne, 1974; Makram et al., 1973; Yen and Brien, 1960). Yields of
dry beans have been lowered by infections of bean rust in Kenya and
Turkey (Mukumya, 1974; Rudolph and Baykal, 1978).
Although the occurrence of bean rust was characterized as sporadic
in the U.S. (Harter et al., 1935), Vargas (1980) reported yield losses
as high as 40-80% in the U.S. are caused by this disease. Brazil is
reported to incur losses of 35-50% due to bean rust infections (Vargas,
1980).
Bean rust was reported to be responsible for the bulk of the yield
losses in Navy beans in Michigan (Andersen, 1975). The disease was
reported to be troublesome in snap bean fields of North Dakota and
Minnesota (Meiners, 1977). Zaumeyer and Meiners (1975) reported that
prior to 1945, bean rust was a major disease in irrigated fields in
Colorado, western Nebraska, Wyoming and Montana. In their review,
Zaumeyer and Meiners (1975) reported bean rust was no longer a problem
in those areas, although it was still occasionally important in fall
snap bean crops along the Atlantic seaboard and in winter crops grown in
Florida.

19
Identification and Etiology of the Pathogen
Bean rust is caused by the fungus Uromyces phaseoli [Pers.] Wint.
(= U. phaseoli typica (Reben) Wint. = U. appendiculatus [Pers.] Unger).
The fungus was first described in Germany in 1795 (Cook, 1978). The
pathogen is an autoecious macrocyclic rust fungus (Kolmer et al., 1984;
Laundon and Waterston, 1965). This pathogen is parasitic on the legumi
nous genera Dolichos, Phaseolus, and Vigna (Laundon and Waterston,
1965). The fungus is transmitted generally through wind-borne uredo-
spores. Uredospores are rusty orange in color and ellipsoidal to
obvoidal in shape, 20-30 x 20-26 yum in measurement (Laundon and Waters
ton, 1965).
The aecial and pycnial stages are rare in U. phaseoli (Harter et
al., 1935). Harter et al. (1935) did not observe any aecia or pycnia of
this fungus in the field. In Queensland, Ogle and Johnson (1974) did
not report seeing mature aecia or pycnia of IJ. phaseoli. The absence of
aecia on U. phaseoli under field conditions has also been reported in
Maryland (Marcus, 1952). The aecial stage of this fungus has, however,
been observed and reported in New York and Virginia (Fromme and Wingard,
1921; Jones, 1960). Both aecia and pycnia were reported to occur on
field grown beans in North Dakota by Venette et al. (1978).
Fromme and Wingard (1918) and Harter et al. (1935) reported that
telia form under unfavorable conditions for the development of the
pathogen such as low temperatures, decreased host vigor, and increased
host resistance. The propensity of the pathogen to form telia was
suspected to be an innate character of the fungal isolate (Harter et
al., 1935). It has been reported that teliospores do not occur in
Florida (Townsend, 1939). Consequently, Townsend (1939) suggested

20
uredospores blown in from the north serve as primary inocula in Florida.
Later, Kidney (1980) observed telia in Alachua and Dade counties,
contrary to Townsend's findings.
Uredospores overwinter in infected crop debris and trellis poles
(Davison and Vaughan, 1963). These overwintered uredospores are known
to initiate the disease cycle in the next growing season in Oregon and
Maryland (Davison and Vaughan, 1963; Marcus, 1952). In Florida, colder
temperatures than those normal for that state are apparently required
for the uredospores to be viable for relatively long periods (Davison
and Vaughan, 1963).
Disease development is frequently initiated by uredospores under
natural conditions. The uredospore produces a germ tube upon germina
tion. An appressorium which molds itself into the stomatal ledge is
formed when the germ tube gets in touch with the stoma (Mendgen, 1973).
An infection peg develops, from the appressorium and pushes the guard
cells apart until the fungal cytoplasm is transferred into the substoma-
tal vesicle (Vargas, 1980). Enzymes, lipid bodies, and glycogen parti
cles are contained in the vesicle (Mendgen, 1973). The fungus develops
infection hyphae and haustoria as it proceeds inter-cellularly in the
host tissue (Mendgen and Heitefuss, 1975; Vargas, 1980).
The bean rust fungus may complete its life cycle within 10-15 days
after inoculation (Yarwood, 1961). Uredospores are released passively
from pustules and disseminated by farm implements, insects, animals, and
wind currents (Yarwood, 1961; Zaumeyer and Thomas, 1957).
Symptoms
Apparently, bean rust is primarily a foliar disease which occasion
ally occurs on pods, stems and branches (Fromme and Wingard, 1918;
Laundon and Waterston, 1965; Vargas, 1980).

21
The uredia (uredinia) are the major diagnostic sign of the pathogen
(Fromme and Wingard, 1918). In a susceptible reaction, symptoms of bean
rust first appear on the lower leaf surface as minute, whitish, slightly
raised spots about 5-6 days after infection. These spots enlarge to
form mature reddish-brown pustules which rupture the epidermis and
obtain a diameter of 1-2 mm, 10-12 days after infection (Vargas, 1980).
The uredia reach a diameter of 5 mm by the 14th day after infection (Rey
and Lozano, 1961). The size of the uredia varies depending on environ
mental conditions as well as the host. The uredia may appear powdery
due to uredospores protruding from them (Fromme and Wingard, 1918).
Uredia often appear on both leaf surfaces. The uredia are frequently
surrounded by chlorotic halos and eventually by rings of secondary and
tertiary sori (Zaumeyer and Thomas, 1957). As infection progresses, the
leaf becomes debilitated and the chlorotic areas surrounding pustules
coalesce, while tissue ramified by the fungus remains green, apparently,
as a result of starch accumulation (Wang, 1961). Severe rust infections
may cause premature abscission. Bean rust rarely causes small, circular
necrotic lesions on pods (Kucharek and Simone, 1980).
Rust infection has been reported to cause increased respiratory
rates in susceptible hosts (Daly et al., 1961). Twenty-four hours after
infection starch accumulation decreases sharply in the vicinity of the
fleck. Accumulation of starch at the perimeter of the lesion, however,
resumes 96-120 hr after infection. The quantity of starch in this area
decreases at the time the fungus sporulates (Schipper and Mirocha,
1969). Rust infections cause leakage of ions, amino acids, and sugars
in susceptible plant leaves (Hoppe and Heitefuss, 1974a). Hoppe and
Heitefuss (1974b) presented evidence that rust infection caused damage

22
to chloroplast membranes. Raggi (1978) reported decreased photosynthe
tic rates in rust-infected plants.
Epidemiology of the Disease
Fromme and Wingard (1921) reported that since rust rarely attacks
pods directly, resulting losses are insidious and difficult to assess.
Yield losses are, however, more likely to be severe when plants are
infected during the prebloom and flowering stages of development (Almeida
et al., 1977; Costa, 1972; Crispin et al., 1976; Nasser, 1976; Wimala-
jeewa and Thavam, 1973; Yoshii and Galvez, 1975). Early infection of
some bean varieties can lead to almost complete crop loss in some
seasons (Fromme and Wingard, 1921; Howland and MaCartney, 1966; Townsend,
1939). Townsend (1939) indicated that total loss of the entire crop due
to rust has occurred in Florida.
The variability in the prevalence of bean rust seasonally and
geographically is partly due to environmental conditions (Augustin et
al., 1972; Gonzalez, 1976; Harter and Zaumeyer, 1941; Harter et al.,
1935; Schein, 1961). Infection by U. phaseoli is favored by prolonged
periods (8-18 hours) of at least 95% RH and moderate temperatures
(15-27C) (Augustin et al., 1972; Gonzalez, 1976, Schein, 1961). The
optimum temperature for uredospore germination was reported to be 14.5C
whereas the optimum temperature for infection was 17C (Harter et al.,
1935). Crispin et al. (1976), Schein (1961), and Zaumeyer and Thomas
(1957), however, reported that any temperatures below 15 may retard
fungal development. Day length and light intensity are also important
factors for the development of the bean rust fungus (Harter and Zaumeyer,
1941).

23
Fifty-seven races of U. phaseoli have been identified in the U.S.
(Stavely, 1984). Laundon and Waterston (1965) reported 35 races of U.
phaseoli. Races 1 and 2 were identified from specimens obtained from
Washington, D.C. and California (Harter et al., 1935). Twenty races of
_U. phaseoli were differentiated according to their reaction on seven
bean cultivars (Harter and Zaumeyer, 1941). Fisher (1952) isolated 10
races from the Rocky Mountain states and Maryland. Race 31 was identi
fied from New Mexico and race 32 from Maryland (Sappenfield, 1954;
Zaumeyer, 1960). Hikida (1961, 1962) isolated and identified races 33
and 34 in Oregon. Race 35 was isolated by McMillan (1972) from the bean
cultivar Dade, which was bred for resistance to previously known races
of £. phaseoli in Florida. McMillan (1972) reported that races 1, 2, 5,
9, 10, 11, and 35 occur in Florida.
There is tremendous variability in the reaction pattern of U.
phaseoli races (Groth and Shrum, 1977). In many areas where several
races occur, there is usually one race which is greatly predominating
(Fisher, 1952).
Uromyces phaseoli races have also been identified outside the U.S.
Thirty-one races have been identified in Mexico (Crispin and Dongo,
1962), 10 races in Colombia (Zuniga and Victoria, 1975), 46 races in
Brazil (Pereira and Chaves, 1977), 12 races in Puerto Rico (Lopez,
1976), 4 races in Nicaragua, 5 races in Honduras (Vargas, 1969, 1970), 7
races in Guatemala (Vargas, 1970), 5 races in El Salvador (Vargas,
1971), 4 races in Peru (Guerra and Dongo, 1973), 11 races in Costa Rica,
11 races in Australia, and 8 races in East Africa (Ballantyne, 1974,
1975; Fisher, 1952; Ogle and Johnson, 1974).

24
Control of the Disease
Cultural control measures of this disease include crop rotation and
removal of old plant debris (Vieira, 1967). Reduced plant density and
planting date adjustment for specific production areas may reduce rust
incidence (Vargas, 1980). Resistant varieties of beans have been used
for the control of rust (Augustine et al., 1972; Ballantyne, 1974; Coyne
and Schuster, 1975; Crispin et al., 1976; Madriz and Vargas, 1975;
Meiners, 1974; Rivera, 1977; Rodriguez, 1976).
Fungicidal sprays are usually recommended to help manage bean rust.
Since bean rust reduces yields more severely when infection occurs
before flowering than when infection is initiated after flowering,
fungicidal sprays are, therefore, more effective if applied during early
plant development (Yoshii and Galvez, 1975). Of the older fungicides,
sulfur dusts have given relatively good control (Ballantyne, 1975;
Harter et al., 1935; Zaumeyer and Thomas, 1957). Sulfur is usually
applied at the rate of 25-30 kg/ha every 7-10 days. Generally, protec
tant fungicides fail in areas where rainfall is frequent because
deposits are washed off too soon. Other preventative chemicals applied
at schedule similar to that of sulfur are chlorothalonil (225 g/ha),
maneb (4-5 kg/ha), and mancozeb (3-4 kg/ha) (Costa, 1972; Crispin et
al., 1976; Hilty and Mullins, 1975; Vieira, 1967; Wimalajeewa and
Thavam, 1973).
Plantvax (Oxycarboxin) is somewhat therapeutic when sprayed 20 to
40 days after planting at the rate of 1.8-2.5 kg/ha (Costa, 1972;
Frenhani et al., 1971; Hilty and Mullins, 1975). McMillan et al. (1982)
reported effective control of bean rust when bean plants were sprayed
weekly with bitertanol or triadimefon. These fungicides are not regis
tered for use on beans at this time. While certain fungicides are

25
effective against bean rust, their use is regulated by their estimated
cost-effectiveness. Thus, Issa and Arruda (1964) cited by Vargas (1980)
concluded that chemical control of bean rust was not economically
practical in Brazil. This conclusion may apply to most tropical bean-
producing areas. The use of fungicides in highly mechanized
agricultural systems, such as the U.S., may be economically feasible
provided registration conditions are met.
Interaction of Root-knot Nematodes and Other Pathogens
Increased incidence of plant diseases has been reported to be
associated with the presence of root-knot nematodes (Brodie and Cooper,
1964; Carter, 1975a,b; Cauquil and Shepherd, 1970; Minton et al., 1975;
Morrell and Bloom, 1981; Norton, 1960; Reynolds and Hanson, 1957;
Schuster, 1959; Thomason et al., 1959; Van Gundy et al., 1977). Carter
(1975), Cauquil and Shepherd (1970), Norton (1960), Reynolds and Hanson
(1957), and White (1962) reported increase incidence of soreshin of
cotton (Rhizoctonia solani Kuhn), root rot (Pythium debaryanum Hesse)
and Fusarium wilt (Fusarium oxysporum Schlecht) when Meloidogyne incognita
(Kofoid and White) Chitwood was present. Increased incidence of southern
blight, Sclerotium rolfsii Sacc., was observed in soybeans infested with
root-knot nematodes (Minton et al., 1975). The interaction of M.
incognita and bacterial wilt (Corynebacterium fluccumfaciens (Hedges)
Dows.) was reported on beans by Schuster (1959). Van Gundy et al.
(1977) reported the enhancement of the development of R. solani in the
presence of exudates from galls caused by M. incognita. Morrell and
Bloom (1981) reported a significant increase in the percentage of
Fusarium wilt occurrence and vessel infection at 21C in the presence of
M. incognita in tomato. Meloidogyne-Fusarium synergism was also observed

26
in cowpea (Thomason et al., 1959). Interaction of root-knot nematodes
is not limited to nematode-fungus or nematode bacterium complexes.
Meloidogyne incognita has also been reported to interact with other
nematodes (Thomas and Clark, 1983). Thus, Thomas and Clark (1983)
reported that early season counts of M. incognita and Rotylenchulus
reniformis Linford and Oliveira were positively correlated with later
counts of the same nematode, but negative correlations were found
between early M. incognita and subsequent R_. reniformis, and between
early R. reniformis and subsequent M. incognita counts. The authors
suggested that a competitive interaction existed with each species
capable of inhibiting the other and becoming the dominant population.
Bookbinder and Bloom (1980) reported the interaction of Meloidogyne
spp. with bean rust, Uromyces phaseoli (Pers.) Wint. They observed that
IJ. phaseoli and M. incognita were synergistic in suppressing shoot and
root weights of beans. Meloidogyne incognita infections reduced uredial
diameter of U. phaseoli significantly. It was observed that
simultaneous inoculations of _U. phaseoli and M. incognita resulted in
reduced mean numbers of galls per gram of root tissue. Similar effects
were observed when U. phaseoli was inoculated first. Meloidogyne
incognita numbers were consequently reduced by 62% in rusted plants.
This reduction in nematode numbers was probably due to suppressed
translocation of photosythates to the roots (Bookbinder and Bloom,
1980). Egg hatch was, nevertheless, not affected by the fungus.

CHAPTER III
THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN
(PHASEOLUS VULGARIS L.) YIELD
Introduction
Snap beans, Phaseolus vulgaris L., are defoliated by leaf eating
insects, diseases, mechanical injury, and adverse weather conditions
(Agudelo, 1980; Costa and Rossetto, 1972; Ruppel and Idrobo, 1962;
Schoonhoven and Cardona, 1980; Vargas, 1980). Thus, an understanding of
the yield-loss relationship between pest infestations and a crop is
essential for the development of an integrated pest management program.
Much information on the relationship between a crop and pest
infestations has been obtained by simulating pest attack through manual
defoliation of plants (Edje et al., 1972, 1973, 1976; Edje and Mughogho,
1976a, b; Galvez et al., 1977; Greene and Minnick, 1967; Vieira, 1981;
Waddill et al.; 1984). Manual defoliation is not a precise simulator of
pest defoliation (Ruesink and Kogan, 1975); however, it provides a good
estimate of the host-pest relationship. To minimize or avoid
defoliation by pests, producers often resort to preventive pesticide
applications on their crop (Greene and Minnick, 1967). These pesticide
applications are a form of insurance on the crop when little knowledge
on the pest damage-yield loss relationship is available.
Beans are defoliated by a wide range of leaf-eating insects includ
ing leafminers (Liriomyza spp.), cabbage looper (Trichoplusia ni
27

28
(Hub.)) leafroller (Urbanus proteus L.), and beetles (Systates spp.).
Pohronezny et al (1978) reported that Liriomyza spp. were considered by
many local farmers in Dade County, Florida, as the most serious pest on
their crops. Farmers expect yield loss as long as same leaf damage is
observed, but Harris (1974) showed that leaf consumption by pests does
not necessarily result in yield reduction.
Greene and Minnick (1967) reported that snap bean yield was not
significantly reduced until more than 33% of the leaf surface was
removed during blooming. It was later observed that snap bean plants
tolerated up to 66% defoliation if damage occurred before flowering
(Greene, 1971). Vieira (1981) found that 66% leaf loss on an indeter
minate bean cultivar reduced yield when defoliation occurred during
flowering and pod formation. At the first trifoliate leaf stage, Galvez
et al. (1977) found that total (100%) defoliation decreased yield of two
bean cultivars by 34 and 49% respectively. Total defoliation of plants
when only primary leaves were present resulted in yield reduction of 65%
on pole beans in Dade County (Waddill et al. 1984).
Kalton et al. (1945) and Weber (1955) reported that up to 75% leaf
removal in soybeans had little effect on yield if plants were defoliated
prebloom. Defoliation in the bloom and pod formation stages, however,
resulted in significant yield losses (Begum and Eden, 1963, 1964; Camery
and Weber, 1953; Kalton et al., 1945; McAlister and Krober, 1958). Todd
and Morgan (1972) reported significant yield reductions on soybeans with
33%, 67% and 100% defoliation at 2 wk, 4 wk, and 8 wk after first bloom.
Research on the effects of defoliation on crop yield has also been
conducted on tomato, cotton, corn, wheat, oats, and other grain crops
(Dungan, 1930; Keularts, 1980; Ludwig, 1926; Stickler and Pauli, 1961;
White, 1946; Wolk et al. 1983; Womack and Thurman, 1962).

29
This study on snap beans (Phaseolus vulgaris L., 'Sprite') was
conducted to determine the plant growth stage most sensitive to defolia
tion, and the effects of defoliation on yield.
Materials and Methods
Two defoliation experiments were conducted in the summer and fall,
1984 in the greenhouse and field, respectively. Bush snap beans
(Phaseolus vulgaris L. 'Sprite') were grown at the Tropical Research and
Education Center in Homestead, Dade County, Florida. The crops were
grown on Rockdale soil (pH ca. 7.8). The greenhouse and field experi
ments were planted on 25 June 1984, and 24 October 1984, respectively.
Fertilizer (8:16:16) was applied at the rate of 448 kg/ha according to
the University of Florida Extension recommendations (Stall and Sherman,
1983). Benlate^ (550g ai/ha) was applied fortnightly for control of
(R)
certain diseases and sprays of Trigardv (150 g/ha) were applied at the
(R)
same frequency for leafminer (Liriomyza spp.) control. Ambushv (40 g
(R)
ai/ha) or Pydrinv (250 g ai/ha) was applied at 14-day intervals for
cowpea curculio (Chalcodermus aeneus Boh.) control.
Defoliation levels investigated were total (100%), 25%, 50%, and
75%. An undefoliated control was included. Plants were defoliated at
the primary leaf stage, first trifoliate leaf, third trifoliate leaf,
flower bud formation, full bloom, and pod set. Beans were harvested on
8-20 August 1984 and 18-26 December 1984. The harvest was not graded
since cowpea curculio feeding damage to pods was extensive.
Greenhouse experiment
Rockdale soil (3030 L) was fumigated with Dowfume^ MC2 (681 g) on
a cement slab under a tightly sealed polyethylene sheet. Number two

30
(7.5 L) pots were filled with 6.4 L soil and placed on a corrugated
bench 0.91 m high. Six seeds were planted in each pot and thinned to
three after emergence. A plot consisted of three pots with three
plants/pot. The crop was irrigated twice a day using an automatic
time-controlled, water-mist-producing overhead system. The foliage was
removed from the distal end of the petiole and the correct number of
leaves removed at a particular growth stage was determined by leaf
counts.
The treatments were replicated four times and randomized in a
complete block. Fresh weights of pods were determined. Yield loss
(percentage) was computed from the untreated control yield at each
growth stage. The dollar economic value was computed by extrapolating
plot data to a per hectare basis. Plot yield data were subjected to
analysis of variance and regression using the general linear models
procedure of SAS (Ray, 1982).
Field experiment
The herbicides Treflan^ (841 g ai/ha) and Dual^ (1.7 kg ai/ha)
were applied to the site prior to planting. Plots were kept as weed-
free as possible by mechanical cultivation. Plots were three rows wide
(0.91 m row spacing) and 6 m long. Seeds were mechanically planted at
8-10 cm spacing within the row. The crop was irrigated using an overhead

31
sprinkler system. The foliage was removed from the distal end of the
petiole and the correct proportion of the foliage removed was determined
from leaf counts.
Ambush^ (40 g ai/ha) or Pydrin^ (250 g ai/ha) was sprayed at 14
(R)
day intervals for leafroller and cowpea curculio control. Benlatev
(550 g ai/ha) and Trigard (150 g ai/ha) were applied fortnightly for
disease and leafminer control respectively. Mesurolv (200 g ai/ha)
was applied as needed for snail and slug control. Treatments were
replicated four times in a randomized complete block. Fresh pod weights
were determined and yield loss computed from the undefoliated plot data
at each growth stage. Yield data were analyzed by analysis of variance
and regression utilizing the general linear models procedure of SAS
(Ray, 1982).
Results
Defoliation had a significant effect on yield in both the green
house and the field, with F-values of 50.16 and 39.95 (p < 0.001)
respectively (Table 2). There was no significant interaction between
time of defoliation and defoliation levels under greenhouse conditions
(F = 0.79). There was significant interaction between defoliation
levels and time of defoliation in the field (F = 3.83). Analysis of
variance on the effects of time of defoliation showed that there were
significant differences between defoliation times (F = 6.23) in the
field but not in the greenhouse.
Regression analysis of the relationship between yield in g/plot and
defoliation level produced models of the form: Y = a+ bx where Y = log
(yield), x = defoliation level and a = intercept. The quadratic model

32
2 2
(a + bx = ex ) resulted in higher coefficients of determination (R )
(Table 3). Generally the fit of the quadratic models to the data was
2
better than the linear model, although the increase in R was generally
less than 10%. Thus the predictive powers of the linear and quadratic
models were more or less similar.
Defoliation did not reduce yield proportionally to its magnitude in
both the greenhouse and the field (Tables 4, 5 and Figures 1, 3).
Conversely, 25% defoliation resulted in yield increases at the primary
leaf stage and full bloom in the field (Table 5). Defoliating plants at
the first trifoliate leaf stage at 75% level increased yield by 4% under
field conditions (Table 5). No yield increases due to defoliation at
pod set in the greenhouse and at full bloom and pod set in the field
were observed (Table 5). Total defoliation at pod set resulted in 74%
yield loss in the greenhouse but losses of 95% and 92% yield loss at
full bloom and pod set in the field.
Yield loss in the greenhouse ranged from 16% to 74% whereas in the
field it ranged from -4% to 95% (Table 5). The least yield reduction in
the greenhouse was observed at 50% defoliation when foliage was removed
at pod set. There was, however, an increase in yield in the field when
75% of the foliage was removed at the first trifoliate leaf stage (Table
5).
Gross dollar values of 'Sprite' snap beans per hectare are shown in
Tables 6 and 7 and Figures 5-10. These values were computed based the
following price ranges $6.00 (low), $11.00 (medium), and $20.00 (high)
for 13.62 kg of snap beans. Since the undefoliated control generally
gave higher yields, dollar values obtained from it were higher. In the
field, however, 75% defoliation at the first trifoliate leaf stage gave

33
TABLE 2. F-values from analysis of variance for the effects of defoli
ation, time (plant growth stage) and their interaction on snap
bean yield.
Field
Greenhouse
Source
F
Prob. F
F
Prob. F
Defoliation
50.16
0.0001
39.95
0.0001
Growth Stage
6.23
0.0001
0.35
0.92
Defoliation X
Growth Stage
3.83
0.001
1.04
0.79

TABLE 3. Regression equations for the relationship between yield and defoliation
Plant Growth Stage Greenhouse Field
Linear
Primary leaf
y
=
127.5
- 76x
R2 =
0.48
y
=
1525 -
805x
R2 = 0.41
(y
=
2.11 -
0.39x)a
(R2 =
0.49)
(y
=
3.18 -
0.35x)
(R2 = 0.34)
y
=
108.1
- 4.64x
R2 =
0.22
y
=
1197 -
686x
RZ = 0.16
First trifoliate
leaf
(y
=
2.01 -
0.24x)
(R2 =
0.22)
(y
=
3.08 -
0.62x)
(R2 = 0.25)
y
=
119.5
- 57.8x
R2 =
0.35
y
=
1420.4
- 922x
RZ = 0.59
Third trifoliate
leaf
(y
=
2.08 -
0.31x)
(R9 =
0.38)
(y
=
3.2 0.55x)
(R2 = 0.58)
y
=
117 -
64.5x
R2 -
0.42
y
=
1286.5
- 863x
RZ = 0.61
Flower bud formation
(y
=
2.07 -
0.36x)
(R, =
0.42)
(y
=
3.14 -
0.54x)
(R2 = 0.53)
y
=
115.9
- 55.9x
E2 *
0.30
y
=
1266.2
- 999x
RZ = 0.66
Full bloom
(y
=
2.05 -
0.31x)
(R2 "
0.26)
(y
=
3.36 -
1.47x)
(R2 = 0.46)
y
=
125.2
- 76.2x
R2 =
0.51
y
=
1441 -
1390.3x
R2 = 0.73*
Pod set
(y
=
2.13 -
0.48x)
(R2 =
0.51)
(y
=
3.25 -
1.06x)
(R = 0.81)
Quadratic
Primary leaf
y
=
129.2
- 89.3x
<- 13.3x2
2
R2
=
0.48
y
=
1446 -
173.6x -
631.4x2
R2
=
0.43
(y
=
2.07 -
0.14x -
0.25xZ)
cr;
=
0.22)
(y
=
3.15 -
0.08x 0
.26x2)
(r;
=
0.36)
y
=
117 -
117.3x +
70.9x2
K
=
0.27
y
=
1005.6
+ 845.4x
- 1531.4x
R2
=
0.24
First trifoliate
leaf
(y
=
2.05 -
0.54x +
0.29x2)
(Ro
=
0.25)
(y
=
2.85 +
1.16x 1
. 78x2)
(r;
=
0.43)
y
=
117.3
- 39.9x
- 17.8xZ
R2
=
0.34
y
=
1269.8
+ 283.7x
- 1205.7x
R2
=
0.68*
Third trifoliate
leaf
(y
=
2.05 -
O.Olx -
0.23xZ)
(Ro
=
0.41)
(y
=
3.07 +
0.52x 1
.07x2)
CR,
=
0.77)
y
=
115.7
- 53.5x
- llxZ
R2
=
0.43
y
=
1284.7
- 848.7x
- 14.3xZ
r7
=
0.61
Flower bud formation
(y
=
2.03 -
0.06x -
0.3xZ)
(R,
=
0.45)
(y
=
3.1 (
D.17x 0.
37x2)
(R
=
0.55)
y
=
116.3
- 58.9x
- 3x2
R2
=
0.29
y
=
1129.8
+ 91.lx -
1090.9x
R2
=
0.73*
Full bloom
(y
=
2.03 -
0.16x -
0.15x1
(R2
=
0.27)
(y
=
2.94 +
1.86x 3
32x2)
(Ro
=
0.67)
y
=
114.A
+ 10.7x
- 86.9x2
R2
=
0.55
V
=
1552
2282.6x +
892.3x
R2
=
0.87*
Pod set
(y
=
2.02 -
0.38x -
0.86x )
(R2
=
0.65)
(y
=
3.17 -
0.42x 0
.64x )
(R2
=
0.84)
Figures in parentheses are y = log (yield); x = defoliation as proportion.
* R significant at 0.05

TABLE 4. Effect of defoliation and defoliation time on snap bean yield (g/plot) in the greenhouse and
field. Data are means of 4 replicates.
Snap
bean yield
(g/plot)
by plant growth
stage
Primary
leaf
First
trifoliate
leaf
Third
trifoliate leaf
Flower
bud formation
Full bloom
Pod set
Defoliation
level (I)
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
0
138
1361
122
1168
125
1293
128
1295
121
1138
122
1530
25
89
1403
84
835
97
1210
89
1113
94
1140
96
1085
50
92
1105
72
933
99
1140
89
673
79
690
102
495
75
84
1050
82
1215
81
820
78
620
93
813
84
457
100
46
613
65
120
54
335
46
350
52
52
32
125

TABLE 5. Relationship between defoliation, time and yield (loss (%)) of snap beans in the greenhouse
and field.
Yield (Z loss by plant growth stage
Defoliation
level (Z)
Primary
leaf
First
triboliate
leaf
Third
trifoliate
leaf
Flower
bud formation
Full bloom
Pod set
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
0
0
0
0
0
0
0
0
0
0
0
0
0
25
36
-3
31
29
22
16
30
14
22
0
21
29
50
34
19
41
20
21
12
30
48
35
39
16
68
75
39
23
32
-4
35
37
39
52
23
29
31
70
100
69
55
47
74
57
74
64
73
57
95
74
92

Log (Yield)
37
Figure 1. Effects of defoliation and time of defoliation on snap beans
yield in the greenhouse (linear models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = third trifoliate leaf,
D = flower bud formation, E = full bloom, and F = pod set.

Log (Yield + 1)
38
2.5
2.0
1.5
1.0
0.5
0
Figure 2.
A Primary leaf stage
B First trifoliate leaf
C Third trifoliate leaf
Effects of defoliation and time of defoliation on snap
bean yield in the greenhouse (quadratic models).
Letters represent the plant growth stage defoliation occurred.

Log (Yield)
39
Figure 3. Effects of defoliation and time of defoliation on snap
bean yield under field conditions (linear models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = thrid trifoliate
leaf, D = flower bud formation, E = full bloom, and F = pod set.

Log (Yield)
40
Figure 4. Effects of defoliation and time of defoliation on snap
bean yield under field conditions (quadratic models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = third trifoliate
leaf, D = flower bud formation, E = full bloom, and F = pod set.

TABLE 6. Effects of defoliation and defoliation time on gross dollar values per hectare of 'Sprite' snap beans
grown in the greenhouse.
Defoliation
level (%)
Price range3
Gross
Time of
dollar values
defoliation
per hectare
(growth stage)
Primary leaf
First trifoliate
leaf
Third
trifoliate
leaf
Flower bud
formation
Full bloom
Pod set
0
low
292
258
265
270
256
258
medium
535
472
485
495
468
472
high
974
859
885
900
852
859
25
low
187
178
207
189
199
204
medium
343
326
379
347
365
373
high
623
593
689
630
664
679
50
low
193
152
209
189
167
216
medium
353
279
384
347
307
397
high
643
507
698
630
558
772
75
low
178
175
172
165
197
178
medium
327
321
316
302
361
326
high
594
584
574
549
656
593
100
low
96
137
115
59
110
67
medium
177
250
211
109
201
123
high
332
455
384
198
366
223
a
low $7/13.62 kg; medium = $11.20/13.62 kg; and high = $20/13.62 kr of snap beans

TABLE 7. Effects of defoliation and defoliation time (plant growth stage) on gross dollar values per hectare of
Sprite snap beans grown in the field.
Gross dollar values per hectare
Time of
defoliation (plant growth stage)
Defoliation
level (%)
Primary
Price range3 leaf
First trifoliate
leaf
Third trifoliate Flower bud
leaf formation Full bloom Pod set
0
low
333
285
316
317
278
374
medium
611
523
580
582
510
686
high
1110
952
1054
1057
928
1248
25
low
343
176
266
273
279
266
medium
629
323
487
500
511
487
high
1143
588
886
909
930
886
50
low
270
228
278
165
170
157
medium
495
419
510
302
311
288
high
869
761
928
549
566
524
75
low
256
297
200
152
197
112
medium
470
544
367
279
361
206
high
855
990
666
507
656
374
100
low
150
74
82
86
110
30
medium
275
136
151
157
201
55
high
500
247
274
285
366
100
a
Low = $7/13.62 kg; medium = $11.20/13.62 kg; and high = $20/13.62 kg of snap beans

43
O 25 50 75 100
Defoliation Level
Figure 5. Influence of defoliation on gross dollar value per hactare
of 'Sprite' snap beans defoliated at the primary leaf
stage in the greenhouse.

Gross dollar value
44
Figure 6
1000
800
600
400
200
high
Price range medium
low
25 50 75
Defoliation Level
Influence of defoliation on gross dollar value per
hectare of 'Sprite' snap beans defoliated at the
third trifoliate leaf stage in the greenhouse.

45
O 25 50 75 100
Defoliation Level
Figure 7. Influence of defoliation on gross dollar value per hectare
of 'Sprite' snap beans defoliated at pod set in the
greenhouse.

Gross dollar value
46
0 25 50 75 100
Defoliation Level (%)
Figure 8. Influence of defoliation on gross values per' hectare of
'Sprite' snap beans defoliated at the primary leaf stage
in the field.

Gross dollar value
1200
1000
800
600
400
200
0
0 25 50 75 100
Defoliation Level (%)
Figure 9. Influence of defoliation on gross dollar values per
hectare of 'Sprite' snap beans defoliated at the third
trifoliate leaf stage in the field.

48
O 25 50 75 100
Defoliation Level (%)
Figure 10.. Influence of defoliation on gross dollar values per hectare
of 'Sprite' snap beans defoliated at pod set in the field.

49
a higher dollar value than the undefoliated control. Removal of 25% of
the foliage at the primary leaf stage and full bloom also resulted in
slightly higher dollar values than the undefoliated control. Generally,
increased defoliation resulted in lower dollar values per hectare.
Discussion
Total defoliation when only primary leaves were present resulted in
69% and 55% yield loss in the greenhouse and field respectively. This
level of defoliation resulted in 57%, 95%, 74% and 92% yield reduction
in the greenhouse and field when plants were defoliated at full bloom
and pod set, respectively. The lower yield reduction in the greenhouse
may be due to the better controlled environmental conditions. The only
growth stage at which total defoliation resulted in less yield loss in
the field was at the primary leaf stage. This may have been due to
better recovery of plants from the total defoliation in the field.
Removing 25% of the foliage resulted in yield loss of at least 20%
in the greenhouse at all growth stages. This is a substantial loss in
terms of dollar values. Thus, it appears that the economic threshold
under greenhouse conditions was between 0 and 25% defoliation. In the
field the economic threshold level varied with the plant growth stage
(Table 4). The increase in yield in the field may have been due to
compensatory reactions of the plant. The compensation may have resulted
from increased photosynthesis due to increased exposure of the remaining
photosynthetically active foliage to light. The same may apply to the
increase in yield of plants with 75% defoliation at the first trifoliate
leaf stage. Reducing foliage may have increased air circulation among

50
plants which, indirectly, may have increased carbon dioxide uptake and
hence photosynthesis. Increased air circulation may also have resulted
in reduced leaf surface humidity which may have reduced subtle fungal
diseases from being established on the crop.
Removing all leaves from plants at pod set and full bloom resulted
in substantial loss under both conditions probably because at these
stages the developing pods were deprived of photosynthates normally
manufactured in these leaves. The pods which developed probably
utilized reserved photosynthates initially and thereafter photosynthates
which were produced from the few leaves which were formed after
defoliation. At the primary leaf stage, total defoliation slowed down
the growth rate of the plants. Under greenhouse conditions, recovery
may have been slow and plants may have been etiolated due to
insufficient light, hence the higher yield loss. In the field, total
defoliation at the first trifoliate leaf stage through flower bud
formation resulted in 74 and 73% yield loss. This loss in yield is
essentially similar in magnitude indicating that the sensitivity of
plants at these stages was more or less the same.
Results obtained in these experiments seem to show yield increases
due to defoliation. This may have been due to chance effects resulting
from many factors including plant characteristics and the environment.
Generally, data show tendency to lower production and hence dollar
values with increased defoliation. The decrease in production due to
defoliation may have been to enhancement of pathogen entry through
wounds made during manual defoliation. Defoliating with scissors also
led to water loss through direct evaporation. Defoliation also reduced
the leaf area for photosynthesis. One can only suspect that the

51
inconsistency was due to the imprecise nature of manual defoliation in
simulating insect damage. It is possible that removing whole fractions
of leaf surfaces had a different effect on plants from damage done by
leaf feeding pests which usually occurs at random.
Although the influence of defoliation on yield was not consistent
at all plant growth stages, plants showed more sensitivity at full bloom
and pod set. This was an indication that leaf damaging pests should be
managed before these plant growth stages. If left unchecked and if
plants become heavily defoliated, substantial loss in yield would be
expected. Insecticides are generally applied as soon as insect pest
infestations are detected. Insecticide application normally starts
before the pest populations exceed threshold levels. Disease control
chemicals are primarily preventatives applied well before the diseases
are observed. Since pesticide sprays against diseases and insect pests
were the same at all defoliation levels and plant growth stages, the
grower would incur loss in gross dollar values proportional to loss in
yield. In this study the threshold level for defoliation was below 25%.
At all plant growth stages.

CHAPTER IV
THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION ON SNAP BEANS
Introduction
Root-knot nematodes, especially Meloidogyne incognita (Kofoid and
White) Chitwood, are a serious threat to bean (Phaseolus vulgaris L.)
production in many bean-producing areas of the world (Agudelo, 1980;
Allen, 1983; Ngundo, 1977; Sharma and Guazelli, 1982; Singh et al.
1981a). M. incognita has been reported to cause extensive root-galling
on bean plants, which interferes with nitrogen fixation by Rhizobium
spp. as well as nutrient uptake by the root system. Moreover, M.
incognita has been reported to increase the severity of other pathogens
through predisposition of host plant root tissues (Carter, 1975a,b;
Golden and Van Gundy, 1975; Powell, 1971; Powell and Nusbaum, 1960;
Porter and Powell, 1967; Sasser et al., 1955). Thus, yield loss caused
by M. incognita and related species is not always a unitary effect, but
often a result of interaction of these nematodes with other plant-
parasitic organisms.
Yield losses of 50 to 90% in field beans have been reported due to
severe root-knot nematode infections (Agudelo, 1980; Freire and Ferraz,
1977; Ngundo, 1977; Varn and Galvez, 1974). Yield decreases caused by
M. incognita are also well known in other crops (Allen, 1983; Lambert!,
1979). Crop yield is usually expected to be inversely related to
nematode counts (Sasser et al., 1968). In view of this theory, Barker
52

53
et al. (1976), Di Vito et al. (1981), and Di Vito and Ekanayake (1983)
reported the relationship between initial M. incognita densities and
plant growth or yield of tomato and sugar beet. Barker et al. (1976)
showed that M. incognita suppressed yields of tomato in North Carolina
by up to 85% in the coastal plains and 20-30% in mountain locations.
Meloidogyne incognita has been observed to cause yield losses of 30-60%
and up to 15% in eggplant and pepper (Capsicum frutescens L.) respec
tively (Lamberti, 1975). Yield losses due to M. incognita infections
have been reported on okra (Hibiscus esculentus L.), sweet potato
(Ipomoea batatas (L.) Lam.), celery (Apium graveolens L.), and carrot
(Daucus carota L.) (Lamberti, 1971).
Root-knot nematodes rarely occur alone on any crop (Powell, 1971).
Thus, nematodes may occur together with other plant pests and diseases.
McSorley and Waddill (1982) reported yield loss partitioning on yellow
squash (Cucrbita pepo L.) into nematode and insect components by using
multiple regression. The partitioning of yield loss was facilitated by
the use of selective pesticides. Consequently, McSorley and Waddill
(1982) suggested that it may be imperative to separate pests into
nematode and insect components when complexes of several pests were
present. This separation of yield loss components would be facilitated
by monitoring field pest populations during the growing season, at
specific intervals, to detect population changes (McSorley and Waddill,
1982).
Beans are susceptible to defoliation by insects, adverse environ
mental conditions, diseases, and mechanical injury (Agudelo, 1980).
Hence an understanding of the relationship between crop yield and pest
infestations is essential for the development of sound pest management

54
strategies. One way of elucidating this relationship has been manual
defoliation of plants to simulate pest damage (Douglas et al., 1981;
Galvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho,
1983; Kalton et al., 1945; Keularts, 1980; Wit, 1983, Wolk et al.,
1983). Hohmann and De Carvalho (1983) reported that leaf area reduction
of 25, 50, 75, and 100% on the bean cultivar Carioca reduced yield by
11, 20, 20, and 70% respectively when defoliation was done at the pod
formation stage. At the same percentage of leaf area reduction, defoli
ation at initiation of flowering decreased yield by 18, 12, 19 and 55%
respectively. Greene and Minnick (1967) indicated that yield reduction
in 'Harvester' snap beans due to leaf removal began somewhere between
33% and 50% defoliation when plants were defoliated in the bloom or
pre-bloom stages. Working on inderterminate snap beans, Waddill et al.
(1984) noted that the removal of both primary leaves when only primary
leaves were present resulted in yield reduction of up to 65%. Vieira
(1981) reported that 66% defoliation of an indeterminate bean cultivar
at the flowering and pod formation stages was detrimental to yield.
Galvez et al. (1977) observed that total defoliation at the formation of
the first trifoliate leaves reduced yields of the bean cultivars ICA-
Guali and Porrillo-Sentetico by 34% and 49% respectively. These obser
vations indicate that the magnitude of yield loss due to defoliation
depends not only on the severity of defoliation but also on the growth
stage (time) the defoliation takes place and the cultivar of beans
grown. Thus, manual defoliation provides a useful estimate of the
host-pest relationship despite its imprecision in simulating pest damage
(Ruesink and Kogan, 1975).

55
This study was conducted to investigate the relationship between M.
incognita population levels, manual defoliation, and their interaction
to snap bean yield.
Materials and Methods
Two studies were conducted in a greenhouse at the Tropical Research
and Education Center in Homestead, Dade County, Florida, in the summer
and fall 1984. One experiment was an inoculation experiment with M.
incognita designed to determine the effect of this pest alone on yield,
and the other experiment examined the simultaneous effect of M.
incognita inoculation and manual defoliation. The first experiment (M.
incognita alone) was designed as a randomized complete-block replicated
four times, involving six different nematode population levels. The
other experiment was a 5x4x6 factorial replicated four times and
included the following treatments: 5 defoliation levels of 0, 25, 50,
75, and 100%; 4 nematode population levels of 0, 1,000, 10,000, and
100,000 eggs and juveniles per pot; and defoliation at the following 6
plant growth stages: primary leaf, first trifoliate leaf, third
trifoliate leaf, flower bud formation, full bloom, and pod set.
Preparations for both experiments were made by covering Rockdale
soil (3030 L) with a polyethylene sheet and fumigating it with
(R)
Dowfume MC2 (681g). Two -gallon, side-drain, plastic pots were
filled with 6.4 L soil and placed on corrugated greenhouse benches 0.91
m high. Fertilizer (8:16:16) was applied before planting at 3 g/pot
(448 kg/ha). Plants were top-dressed with the same fertilizer at 1.5
g/pot four weeks after germination. The M. incognita inoculation test

56
was planted on 25 June 1984 and harvested on 28 August 1984. The M.
incognita x defoliation study was planted on 24 October 1984 and
harvested on 26 December 1984 to 3 January 1985. In each experiment, a
plot comprised of 2 pots, each containing 3 plants. Irrigation was
provided by an automatic time-controlled, overhead, water-mist-producing
system twice a day.
The Meloidogyne incognita population used in each experiment was
originally obtained from Hausa potato (Coleus parviflorous Benth.) and
was maintained on greenhouse-grown tomato (Lycopersicon esculentum
Mill.) FloraDade' plants. Meloidogyne incognita eggs were extracted by
the sodium hypochlorite (NaOCl) method of Hussey and Barker (1973). A
(R)
0.525% NaOCl solution was made from Thrift King commercial bleach
(5.25% NaOCl) by dilution with cold tap water (25C). Tomato roots were
thoroughly washed of soil with running tap water. The clean roots were
cut into 2-3 cm long pieces and 120 g of the cut root material was
manually shaken in 200 ml of the NaOCl solution for 3.5 minutes. The
shaken material was serially filtered through 100-mesh, 230-mesh and
500-mesh sieves. The number of eggs and juveniles of M. incognita per
ml was determined by counting in a watch glass under a dissecting
microscope (20X). Appropriate dilutions of the nematode eggs and
juveniles were made according to the population levels used.
Plants were inoculated 10 days after planting by drenching their
bases with the nematode egg and juvenile suspension. The nematode
population levels of 0, 10, 100, 1,000, 10,000, and 100,000 per pot were
equivalent to 0, 0.16, 1.6, 15.6, 156 and 1562 eggs and juveniles per
100 ml soil, respectively.

57
In the experiment involving simultaneous nematode inoculation and
defoliation, plants were defoliated manually with a pair of scissors.
The correct number or proportion of leaves to be removed was determined
by leaf counts at each plant growth stage. To eliminate additional
(R)
uncontrolled defoliation, plants were sprayed with Ambushv (40 g
ai/ha) for bean leaf roller (Urbanus proteus L.) and cowpea curculio
00
(Chalcodermus aeneus Boh.) control; Trigardv (150 g ai/ha) for
(R)
leafminer (Liriomyza spp.) control, and Benlatev (550 g ai/ha) for
disease control. These pesticides applied at 14-day intervals. Yield
was taken from all six plants. Pods less than 7 cm in length and
diseased or damaged ones were discarded.
Yield data were subjected to analysis of variance and regression
analysis using the general linear models procedure of SAS (Ray, 1982).
Nematode inoculation data were also analyzed using Seinhorst's models
(Ekanayake and Di Vito, 1984; Ferris, 1984; Ferris et al. 1981; Sein-
horst, 1965).
Results
Meloidogyne incognita alone
Analysis of variance on the effect of Meloidogyne incognita popula
tion levels on yield showed a significant relationship with F = 26.2***
(Table 8). Regression analysis of the data produced models of the form
2
Y = a + bx or Y = a + bx + cx where Y = yield (g/plot) or log (yield),
x = log (M. incognita population + 1) (Table 9). Quadratic models
o
consistently gave somewhat higher coefficients of determination (R )
values. The predictive ability of the quadratic models was, however,

58
TABLE 8. Effect of M. incognita on snap bean yield. Data are means of
4 replicates.
No. of
M. incognita/Pot
Log (M. incognita
Population +1)
Yield (g/plot)a
Yield loss
(%)
0
0
128
0
10
1.0
103
19
100
2.0
79
38
1000
3.0
70
45
10000
4.0
68
47
100000
5.0
53
59
Data rounded off to the nearest whole number.
F = 26.2*** for M. incognita populations.

59
not greatly superior to that of linear models (Figure 11). Yield
reduction was initiated even by the lowest nematode population.
The data did not fit the Seinhorst (1965) model, which is of the
r p-Tl
form Y = m + (1-m) Z where Y = ratio between yield at nematode
population level p and in the absence of nematodes, m = minimum yield at
very high nematode population levels, T = tolerance limit (the nematode
population level below which yield reduction does not occur), and Z =
the proportion of the plant undamaged in the presence of parasitism or
infection by one nematode (Ferris et al., 1981; Ferris, 1984; Seinhorst,
2
1965 and 1972). The Seinhorst model produced a R -value of only 0.00661,
an indication that the data had a poor fit to the model. Data were,
moreover, linear in trend (Figure 11).
Table 10 shows the gross dollar values per hectare of snap beans.
These values were computed from the price ranges of $6.00 (low), $11.00
(medium), and $20.00 (high) per bushel (13.62 kg) of beans. These were
derived from gross yield and prices given above. They are what the
grower would get without subtracting the cost of production which
includes labor, pesticides, rent, farm machinery depreciation, and/or
interest in loans. The nematode-free plots produced at least twice as
much money as plots with 10,000 or 100,000 eggs and juveniles of M.
incognita per pot which were equivalent to 156 and 1562 eggs and juve
niles per 100 ml soil. Nematode-free plants produced $404, $740 and
$1345 per hectare at the low, medium, and high prices respectively
compared to $326, $598, and $1087 when pots were inoculated with 10 eggs
and juveniles (= 0.16/100 ml soil). This level of M. incognita popula
tion resulted in a 19% loss in gross dollar value per hectare ($77,
$142, and $258 loss at low, medium and high prices, respectively).

60
Meloidogyne incognita inoculation x defoliation
Analysis of variance on the relationship between yield and M.
incognita defoliation, and their interaction showed that there was no
significant interaction between defoliation and M. incognita at all
plant growth stages during which defoliation was done (Table 11). There
were, however, significant differences among M. incognita population
levels, and there were also significant differences in yield with defo
liation level at all plant growth stages (Table 11). Regression analy
sis produced models of the form Y = a + bx + cn, where Y = yield
(g/plot, x = defoliation level (as a fraction), and n = log (M.
incognita population + 1) (Table 12). Quadratic models of the form Y =
2 2
a + b^x + b^x + c^n + C£n gave somewhat higher coefficients of
2
determination (R ) (Table 12), but are more difficult to visualize
graphically. In general, yield was reduced much faster by nematodes
than when defoliation was held constant then when nematode populations
were held constant and defoliation levels were changed (Figure 13).
Table 13 shows data on the effects of defoliation, and M.
incognita, when they occurred simultaneously, on snap beans. The lowest
yields were obtained when 100% of the leaf area was removed at the first
trifoliate leaf stage or at flower bud formation and plants were inocu
lated with 100,000 eggs and juveniles/pot (Table 13). Total defoliation
at any time from the third trifoliate leaf stage through to pod set
reduced yield by at least 93% in the presence of an initial nematode
population density of 100,000 eggs and juveniles/pot. There was, gen
erally, no apparent synergistic effect when defoliation and M. incognita
occurred simultaneously (Tables 13 and 14), which was evidenced by the
lack of significance for the interaction term in the analysis of
variance.

61
TABLE 9. Regression equations for the relationship between M. incognita
population levels (x) and yield component (y).
Yield component
Model
Linear
Quadratic
Yield
Y = 118.6 -13.98x
Y = 126.7 26x + 2.4x2
R2 = 0.81*
R2 = 0.87*
Log (Yield)
Y = 2.1 O.lx
Y = 2.1-0.lx + 0.005x2
R2 = 0.84**
R2 = 0.85*
* R significant at 0.05
** R significant at 0.01

Yield (g)
62
Log (M. incognita Population + 1)
Figure 11
Effects of M. incognita on snap bean yield.

TABLE 10. The influence of M. incognita on the gross dollar value per
hectare of snap beans grown in a greenhouse.
Log (M. incognita
population +1)
Price range3
Gross dollar values
per hectare
0
low
404
medium
740
high
1345
1.04
low
326
medium
598
high
1087
2.0
low
248
medium
456
high
828
3.0
low
222
medium
407
high
740
4.0
low
215
medium
395
high
718
5.0
low
167
medium
307
high
558
a
Low = $6; medium = $11; high = $20/13.62 kg of snap beans.

TABLE 11. F-values and probability levels from analysis of variance for the effects of defoliation and M.
incognita and their interaction on snap bean yield.
Plant Growth Stage
Source
Primary First tri- Third tri- Flower bud Full
leaf foliate leaf foliate leaf formation bloom
Pod
set
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
Defoliation
17.67
0.0001
67.84
0.0001
66
0.0001
41.6
0.0001
100
0.0001
140
0.0001
Log (M. incognita
population +1)
23
0.0001
23.5
0.0001
35
0.0001
38
0.0001
28.95
0.0001
56.15
0.0001
Defoliation x
Log (M. incognita
population +1)
0.56
0.8871
0.54
0.8395
1.5
0.0962
0.72
0.7396
1
0.4601
0.38
0.9575

TABLE 12. Regressions equations for the relationship between M. incognita defoliation and yield.
Growth stage plants
were defoliated
Regression equation
a
Linear equations
Primary leaf stage Y =
First trifoliate leaf stage Y =
Third trifoliate leaf stage Y =
Flower bud formation stage Y =
Full bloom stage Y =
Pod set stage Y =
259-22.12N-25.lx (R2 = 0.62**)
256.51-21.04N-32.3x (R2 = 0.59**)
267.04-21.51N-33x (R2 = 0.72**)
258.15-24.17N-30.19x (R2 = 0.64**)
237.26-17.75N-31.34x (R2 = 0.58**)
25176-21.44N-29.77x (R2 = 0.58**)
Quadratic equations
Primary leaf stage Y
First trifoliate leaf stage Y
Third trifoliate leaf stage Y
Flower bud formation stage Y
Full bloom stage Y
Pod set stage Y
241.97 + 2.07N-5.13N2-7.9x-4.3x2 (R2 = 0.66**)
227.11 + 5.39N-5.61N2+8.09x-10.1x2 (R2 = 0.67**)
239.73 2.42N-4.03N2+7.42x-10.1x2 (R2 = 0.79**)
243.49 10.31N-2.94N2-10.53x-4.94x2 (R2 = 0.66**)
204.27-0.57-3.64N2+22.68x-13.5x2 (R2 = 0.79**)
218.66-0.24N-4.5N2+21.65x-12.86x2 (R2 = 6.68**)
aY = Yield (g/plot).
N = Log (M. incognita population + 1).
X = Defoliation level (fraction).
** = R significant at 0.01.

TABLE 13. Effects of M. incognita and defoliation on snap bean yield (g/plot). Data are means of 4
replicates.
Snap bean yield
(g/plot) by plant growth
stage
Defoliation
level
Log (M. incognita
population +1)
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower
bud
formation
Full
bloom
Pod
set
0
0
261
278
274
288
238
247
0
3.0
219
247
230
207
188
193
0
4.0
173
163
150
155
122
143
0
5.0
112
96
102
87
87
89
0.25
0
213
166
225
215
224
220
0.25
3.0
186
138
173
148
182
175
0.25
4.0
139
118
141
129
141
135
0.25
5.0
84
60
121
95
110
106
0.50
0
217
235
204
196
187
203
0.50
3.0
172
183
170
145
157
154
0.50
4.0
147
140
153
94
126
111
0.50
5.0
87
107
114
71
98
74
0.75
0
185
188
193
180
145
216
0.75
3.0
145
156
156
144
135
170
0.75
4.0
106
114
87
95
109
136
0.75
5.0
92
80
66
80
69
66
1.0
0
126
46
80
83
48
38
1.0
3.0
87
30
50
50
28
35
1.0
4.0
58
21
29
27
17
23
1.0
5.0
41
12
19
16
17
15
on
ON

T
H*
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C
T
D
Gross dollar value
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68
Figure 13: Effects of defoliation and M. incognita on snap
bean yield.

TABLE 14. The influence of M. incognita and defoliation on snap bean yield loss (%)
Yield loss
(%) by plant
growth stage
Defoliation
level
Log (M. incognita
population +1)
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower
bud
formation
Full Pod
bloom set
0
0
0
0
0
0
0
0
0
3.0
16
11
16
28
21
22
0
4.0
34
41
45
46
49
42
0
5.0
57
65
63
70
62
60
0.25
0
19
40
18
25
6
11
0.25
3.0
29
50
37
49
24
29
0.25
4.0
47
58
49
55
41
45
0.25
5.0
68
78
56
67
53
57
0.50
0
17
15
26
32
21
18
0.50
3.0
34
34
38
50
34
38
0.50
4.0
44
50
44
67
47
55
0.50
5.0
67
62
58
75
59
70
0.75
0
29
32
30
38
39
12
0.75
3.0
45
44
43
50
43
31
0.75
4.0
60
59
68
67
54
45
0.75
5.0
65
71
76
72
71
73
1.0
0
52
83
71
71
80
84
1.0
3.0
67
89
82
83
88
86
1.0
4.0
78
93
90
91
93
91
1.0
5.0
84
96
93
94
93
94
VO

TABLE 15. Effects of M. incognita and defoliation on the gross dollar values per hectare of snap
beans.
Gross dollar values by plant growth stage
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0
0
low
830
882
870
916
755
783
medium
1522
1617
1595
1679
1384
1435
high
2768
2940
2900
3053
2517
2609
0
3.0
low
697
785
731
659
596
611
medium
1278
1439
1340
1209
1094
1119
high
2325
2617
2436
2198
1988
2035
0
4.0
low
548
520
478
495
385
454
medium
1005
954
877
907
706
833
high
1827
1734
1595
1649
1283
1514
0
5.0
low
357
309
322
275
284
313
medium
655
566
590
504
521
574
high
1191
1029
1073
916
956
1044
0.25
0
low
673
529
714
687
710
697
medium
1233
970
1308
1259
1301
1278
high
2242
1764
2379
2289
2366
2323
0.25
3.0
low
589
441
548
467
574
556
medium
1081
808
1005
856
1052
1019
high
1965
1470
1827
1557
1913
1853
0.25
4.0
low
440
370
444
412
445
430
medium
807
679
813
756
817
789
high
1467
1235
1479
1374
1485
1435

TABLE 15. continued
Gross dollar values by plant growth stage
First Third
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
trifoliate
leaf
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0.25
5.0
low
266
194
383
302
355
337
medium
487
355
702
554
651
617
high
885
641
1276
1007
1184
1122
0.5
0
low
689
750
644
586
596
642
medium
1264
1375
1181
1075
1094
1177
high
2298
2499
2147
1954
1988
2140
0.5
3.0
low
548
582
540
458
586
485
medium
1005
1067
989
840
1075
890
high
1827
1940
1799
1527
1954
1618
0.5
4.0
low
465
441
482
302
398
352
medium
853
808
893
554
730
646
high
1551
1470
1624
1003
1327
1175
0.5
5.0
low
274
335
366
229
310
235
medium
502
615
670
420
568
430
high
912
1118
1219
764
1032
783
0.75
0
low
589
600
522
568
461
689
medium
1081
1099
957
1041
845
1263
high
1965
1999
1740
1893
1536
2297
0.75
3.0
low
457
494
496
458
430
540
medium
838
905
909
840
789
990
high
1523
1646
1653
1527
1435
1800

TABLE 15. continued
Gross dollar values by plant growth stage
First Third
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
trifoliate
leaf
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0.75
4.0
low
332
362
278
302
347
430
medium
609
663
510
554
636
789
high
1107
1206
928
1007
1157
1435
0.75
5.0
low
291
256
209
256
219
133
medium
533
469
383
470
401
244
high
969
853
696
855
730
443
1.0
0
low
399
150
252
266
151
125
medium
731
275
463
488
277
229
high
1329
499
841
885
504
417
1.0
3.0
low
274
97
153
155
91
110
medium
502
178
288
285
167
201
high
912
323
523
518
303
366
1.0
4.0
low
183
62
87
82
53
70
medium
335
113
160
151
97
129
high
609
206
291
275
176
235
1.0
5.0
low
133
35
61
55
53
47
medium
244
65
111
101
97
86
high
443
117
203
184
176
157

73
Gross dollar values are shown in Table 15 and Figure 12. These
values were computed from gross yield per hectare based on the following
price ranges $6.00 (low), $11.00 (medium), and $20.00 (high). Hence
these are gross values without deducting production costs. The gross
dollar values (Table 15) show that there was a wide range at each plant
growth stage. Generally highest dollar values were obtained when the
plants were nematode-free and when no defoliation occurred. The combi
nation of nematodes and defoliation had inconsistent effects on gross
dollar values. If defoliation were held constant at any level, gross
dollar values decreased as the nematode populations were increased
(Table 15). If nematode populations were held constant the decrease in
gross dollar values was not always consistent with the levels of defoli
ation. This is shown clearly at the primary leaf, first trifoliate
leaf, full bloom and pod set stages of plant growth. At these stages
some lower levels of defoliation have smaller dollar values than higher
defoliation levels. As expected, loss in dollar values was similar to
yield loss since the former was obtained from yield (g/plot), but varies
greatly depending on the current market price which can fluctuate
widely.
Discussion
There were significant differences in yield when M. incognita
inoculated to snap beans. In this test 10 eggs and juveniles/pot
reduced yield by at least 19%. In the defoliation M. incognita
interaction test, 1,000 eggs and juveniles/pot depressed yield by only
11-28% when plants were not defoliated (Table 14). Similar trends

74
occurred when 10,000 or 100,000 eggs and juveniles/pot were used. This
discrepancy in M. incognita effects on snap bean yield may be due to the
difference in the seasons in which the two trials were conducted. Tyler
(1933) reported that at temperatures ranging from 27.5C to 30C,
females of Meloidogyne spp. developed from infective juveniles to the
egg-laying stage in 17 days; at 24.5C in 21 to 30 days; at 20C, in 31
days; at 15.4C in 57 days; and at temperatures above 33.5%C or below
15.4C, females failed to reach maturity on tomato plants. Decker and
Casamayor-Garcia (1966) stated that one generation of M. incognita
developed on lettuce within 26 days at a mean temperature of 23.3C.
They further stated that from the time of larval invasion up to the
commencement of egg-laying required at least 19 days. Lamberti (1979)
observed that M. incognita rarely started to invade root tissues when
the soil temperature was below 18C. Since the two tests being discus
sed here were conducted at different times of the year, it is likely
that in the summer test soil temperatures were generally higher than in
the fall trial. Thus, the life cycle of the nematode may have been
completed in a shorter period of time in the summer than in the fall.
Consequently, more M. incognita generations (at least 3) may have been
completed during this season. There is also the possibility that the
quality of the inoculum was different in the two tests since the source
of the eggs and juveniles for the fail test was also exposed to
relatively lower temperatures than the summer inoculum. The verifi
cation of this phenomenon can only be obtained by conducting further
tests. The higher yields in the fall test may also be due to the fact
that the nematodes were not able to invade the root tissues of the
plants as fast as they could under optimum summer conditions. During

75
these tests soil temperatures were not taken which in fact precludes the
comparison of edaphic temperatures during the time the two tests were
conducted. Soil temperatures are generally warmer when air temperatures
are higher.
The results obtained in the test where M. incognita was used alone
indicate that the threshold population level was between 0 and 10 eggs
and juveniles/pot whereas in the fall experiment the threshold popula
tion level was between 0 and 1,000 eggs and juveniles/pot. During the
fall, 10 and 100 eggs and juveniles were not used hence the threshold
for this test could have possibly, been similar to the summer threshold
level. Generally, defoliation increased yield loss when combined with
nematodes (Table 14). The influence of defoliation level on yield was
not as drastic as expected. It is not apparent why defoliation had this
slight effect on yield. This is not, however, in agreement with the
general principle that nematodes predispose plants to diseases and other
pests. Statistical analysis showed that defoliation and M. incognita
acted independently in influencing yield.
The gross dollar values had a trend similar to that of yield since
they were computed from gross yield. These values are gross figures
from which one has to deduct production costs which include pesticides,
land rent, labor, interest on loans (if any), and farm machinery depre
ciation. Thus, net income would depend on the cost of production and
current market prices of snap beans. Snap bean production is costly
(Taylor and Wilkowske, 1984). Loss in gross income ranges compared to
nematode free plants were $470 to $1443; $441 to $2441; $521 to $2059;
$764 to $2167; $151 to $2201; and $286 to $2192 when plants were
defoliated at various levels at the primary leaf, first trifoliate leaf,

76
third trifoliate leaf, flower bud formation, full bloom, and pod set
stages respectively. If plants were not defoliated but were inoculated
with nematodes, loss in income ranged from $443 to $1577; $323 to $1911;
$464 to $1827; $855 to $2137; $529 to $1561; and $674 to $1565 when
plants were meant to be defoliated at the primary leaf, first trifoliate
leaf, third trifoliate leaf, flower bud formation, full bloom, and pod
set stages respectively. In each case the lower loss in income is for
the 1,000 eggs and juveniles/pot population level and the higher value
in loss was for the 100,000 eggs and juveniles/pot (Table 15). These
losses are based on yield loss disregarding production costs. The loss
in income has been computed using the high market price of snap beans.
The yields of snap beans in both studies were low, and thus, the grower
would have had a loss in income in both cases.

CHAPTER V
THE EFFECT OF BEAN RUST, UROMYCES PHASEOLI (PERS.) WINT.,
ON SNAP BEANS, PHASEOLUS VULGARIS L. 'Sprite'
Introduction
Bean rust, caused by the fungus Uromyces phaseoli (Pers.) Wint., is
a serious disease of beans, Phaseolus vulgaris L. (Agudelo, 1980; Allen,
1983, McMillan et al., 1982, Pohronezny et al., 1984). The disease
causes severe damage on winter and spring grown snap beans in south
Florida (McMillan, 1982; Pohronezny et al., 1984). Usually, IJ. phaseoli
first appears in January and becomes progressively more severe February
through May (Pohronezy et al., 1984). Initial inoculum is believed to
come from infected bean plant debris in abandoned fields. Losses of up
to 78% in pinto beans, 74.2% and 18.4% in 'Ex Rico 23' and 'Bat 308'
field beans, respectively, have been reported from severely infected
crops in the United States and Latin America (CIAT, 1983; Kelly, 1982).
Bean rust, £. phaseoli, is an autoecious polycyclic disease whose
rates of increase are affected by timing, amount of sporulation, light
intensity, relative humidity, and relative cultivar susceptibility
(Cohen and Rotem, 1970; Cook, 1978; Imhoff et al., 1982 a,b). Rotem et
al. (1973) reported that, in an automatic humidity chamber study,
humidity was inversely related to the sporulation of £. phaseoli.
Infection by IJ. phaseoli has, however, been reported to be favored by
prolonged periods of at least 95% relative humidity and moderate temper
atures (15-27C) (Augustin et al., 1972; Gonzalez, 1976; Schein, 1961).
77

78
Uromyces phaseoli progress on artificially inoculated beans, vulgaris
'Bountiful', depended more on length and frequency of wetting periods
than on temperature (Imhoff et al., 1982a).
Yield loss due to disease has been observed to be proportional to
the area under the disease progress curve or proportional to disease
severity at some critical stage of host growth (Madden et al., 1981;
Raymundo and Hooker, 1981; Romig and Calpouzos, 1970; Shaner and Finney,
1977; Teng et al. 1979). In many of these studies, area under the
disease progress curve satisfactorily explained the relationship between
diseases and yield losses. Disease severity at one or more points in
time and rate of increase of the disease were also satisfactory disease
parameters employed to explain the relationship between disease and
yield loss (James, 1974; James and Teng, 1979; Main, 1977).
Berger (1981) compared the logistic and Gompertz models for disease
progress curve fitting. It was observed that the Gompertz model consis
tently gave better fit to the data examined than the logistic model for
disease severity values outside the 0.05 < y < 0.6 range (Figure 14).
The Gompertz model was superior to the logistic model in linearizing 113
selected disease progress curves (Berger, 1981).
Growers often resort to routine fungicide sprays for disease
control. Currently, weekly sprays with mancozeb are applied for disease
control on beans (McMillan et al., 1982; Pohronezny et al. 1984). The
effectiveness of these sprays depends on spray coverage and disease
severity but in many cases disease control is less than satisfactory
(McMillan et al. 1982). Usually, sprays are initiated before disease
signs and/or symptoms are observed on the crop.
The present studies were conducted to determine the effect of bean
rust on 'Sprite' snap beans under field conditions.

V£>

80
Materials and Methods
Two trials were conducted at the Tropical Research and Education
Center in Homestead, Dade County, Florida, on Rockdale soil (pH ca.7.8).
The first trial was planted on 27 February 1985 and the second on 21
March 1985. Beans were harvested on 25 April 1985 and 13 May 1985
respectively.
In both trials plots were 3 rows wide (0.91 m row spacing) and 3 m
long. Beans were planted 7-10 cm apart within the row. Prior to
planting the herbicides Treflan^ (841 g ai/ha) and Dual^ (1.7 kg
ai/ha) were applied to the site. Fertilizer (8:16:16) was applied at
448 kg/ha before planting. Plants were top dressed at 224 kg/ha just
(R)
before flower bud formation. The crops were sprayed with Ambushv (40
g ai/ha) fortnightly for cowpea curculio (Chalcodermus aeneus Boh.)
(R)
control. Slugs and snails were controlled by Mesurolv (200 g ai/ha)
pellets. Plants were irrigated using an overhead sprinkler system.
In both trials five treatments were arranged in a randomized
complete block design with four replications. Fungicides were used as a
tool to manipulate disease levels. Treatments used were (a) no fungi
cide; (b) bitertanol (57 g ai/ha) at 7-day intervals; (c) mancozeb (0.7
kg ai/ha) tank-mixed with sulfur (4.5 kg ai/ha) at 4-5-day intervals;
(d) same as (c) but at 7-day intervals; and (e) same as (c) but at
(R)
14-day intervals. All sprays were applied with Helenav sticker or Nu
(R)
Film-17v as a spreader/sticker. Bitertanol plots were virtually
disease free.
Plants were inoculated at the primary leaf growth stage using
infected pole bean leaves collected from abandoned bean fields. The

81
infected leaves were clipped on to wire stakes 25 cm above ground level.
Two stakes were placed in each plot, one on each end, depending on
general wind direction. Disease progress was monitored by taking
trifoliate leaves at random from each plot once a week. At each
sampling occasion leaves were taken from the same relative level within
the canopy. Disease severity (proportion of leaf area infected by the
a_b
disease) was determined using the mathematical model y = where y =
disease severity, a = area of leaf before cutting out diseased tissue,
and b = area of leaf after cutting out diseased tissue. The mean of the
5 trifoliate leaves was the measure used in the final data. Leaf area
(R)
was determined by a LiCorv portable area meter (Model LI-300, Lombdar
Instruments Corp).
Disease progress curves are generally sigmoid in shape (Imhoff et
al., 1982a). The generalized progress curve is shown in Figure 14.
Progress curves of bean rust in these studies were obtained by determi
ning disease severity at weekly intervals as indicated above.
Area under the disease progress curve was computed using the general
model: y = £ [ [tl + 1 l1 ln "hich ? area under
the disease progress curve, x = disease severity at the i observation,
t^ = time (days) at the i*"*1 observation, and n = total number of obser
vations. The computations were facilitated by the use of a computer
program provided by Dr. R. D. Berger. The computer program employed the
following model: y = (((n (x) + n (x + l))/2) t (x)) where y = area
under the disease progress curve, x = disease severity, n = number of
disease severity values, and t = time (days) at which observation is
made. The rate of disease progress was determined by using the Gompertz
model which consistently gave better fit to the data (Berger, 1981).

32
Gross dollar values were obtained by multiplying yield (kg/ha) by
current market prices of 13.62 kg of snap beans which were $6 (low), $11
(medium), and $20 (high), respectively. Net returns from investment
were derived by subtracting the gross dollar values of unsprayed plants
and the cost of mancozeb from the values realized from sprayed plants.
No net returns for bitertanol spray were computed because this fungicide
was experimental and its price was not available.
Data were analyzed by the analysis of variance and regression
analysis using the general linear procedure of SAS (Ray, 1982).
Results
Figures 15 and 16 show disease progress in trials 1 and 2 respec
tively. Progress patterns were similar in both trials although disease
severity values were higher for trial 2.
Analysis of variance of yield data by treatments gave F values of
5.37 and 10.77 with probabilities of 0.01 and 0.005 for trials 1 and 2,
respectively. This showed that there was a significant relationship
between yield and disease severity. The disease free plants produced
higher yield in trial 2 than in trial 1 (Table 16). In trial 1 disease
severity ranged from 0.098 to 0.76 and yield was from 1102 g to 2723
g/plot whereas in trial 2 disease severity ranged from 0.4 to 0.86 and
yield was 276 g to 3214 g (Table 16). Generally, where the disease
occurred, yield was lower in trial 2 than in trial 1.
Figure 17 shows the relationship between yield loss and maximum
proportion of foliage infected (disease severity) for both trials. In
trial 1, 0.098, 0.46, 0.65, and 0.76 disease severity resulted in 21%,

83
17%, 35%, and 60% yield loss respectively whereas in trial 2 disease
severity of 0.4, 0.47, 0.71, and 0.86 lead to 55%, 56%, 80%, and 91%
yield loss respectively. Figure 18 is a representation of the relation
ship between area under the disease progress curve (AUDC) and yield for
trials 1 and 2. Both disease severity and AUDC were positively cor
related with yield loss which showed that these disease measures were
inversely related to yield. Figures 17 and 18 are similar in shape.
Regression equations between disease severity and snap bean yield shown
in Tables 17 and 18 are similar in nature. Regression analysis of the
data produced the model y = a + bx, y = yield (g/plot), and x = disease
parameters. Both disease severity and AUDC were significantly corre
lated with yield at full bloom and pod set in trial 1 (Table 17). When
pods were fully developed only disease severity was significantly
correlated with yield. In trial 2, disease severity and AUDC were
significantly correlated with yield at pod set through the stage when
pods were fully formed (Table 18). In trial 1 the coefficient of
2
determination (R ) at full bloom or later range from 0.46 to 0.93 (Table
2
17) while in trial 2 the coefficient of determination (R ) at pod set
or later ranged from 0.89 to 0.99 (Table 18). In trial 1 bean rust
severity was not significantly correlated with yield at the stage when
pods were half developed whereas the in trial 2 the disease was not
significantly related to yield at flower bud formation and full bloom
(Tables 17 and 18).
The gross dollar values per hectare of snap beans infected by bean
rust are shown in Table 19 and Figures 19 and 20. The virtually disease
free plants gave the highest dollar values per hectare in both trials.
These plants were sprayed with bitertanol an experiment fungicide, which
is currently not registered for rust control on beans. Therefore, no

84
Therefore, no price information on this product is given. In trial 1,
plants with a 0.46 disease severity gave a higher dollar value than
plants with a 0.098 disease severity. These dollar values corresponded
to spray intervals of mancozeb and sulfur of 7-days and 4-5 days (Table
19). Plants with a disease severity of 0.4 and 0.47 gave dollar values
of $2367 and $2327 respectively in trial 2. These disease severity
values corresponded with 4-5-day and 7-day spray schedules of mancozeb
and sulfur (Table 19). In trial 2, gross dollar values were consistent
ly inversely related to both disease parameters (Table 10). The plants
with the highest disease parameter produced the lowest gross dollar
value.
The relationship between disease severity and net returns from
investment per hectare of snap beans is shown in Table 20. No net
returns are shown for the virtually disease free plants because an
experimental fungicide with no price tag was used on them. In trial 1
there was no improvement on net returns by spraying beans at 4-5 day
intervals from the 7-day intervals. Actually, there was a loss in net
income by spraying plants more often (Table 20). In trial 1, there were
substantial increases in net returns when plants were sprayed at 7-day
intervals compared to the 14-day spray schedule. The increases in
returns were $1069, $578, and $306 at the high, medium and low prices at
the 7-day spray schedule from the 14-day schedule in trial 1. From the
14-day spray schedule to the 4-5-day spray interval there were increases
in net returns of $901, $475, and $239 at the high, medium, and low
prices respectively in trial 1. By increasing spray frequency from
7-day to 4-5-day intervals there were net losses of $168, $103, and $67
at the high medium, and low prices respectively. Thus, in trial 1 there

Proportion of Foliage Infected
85
Days After Inoculation
Figure 15. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at various frequencies (Trial 1).
A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals,
D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur
at 14-day intervals.

Proportion of Leaf Infected
86
Figure 16. Disease progress curves for Uromyces phaseoli on snap
beans sprayed with mancozeb at various frequencies
(Trial 1).
A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals,
D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur
at 14-day intervals.

87
TABLE 16. Effects of Uromyces phaseoli on snap bean yield.
Disease Parameters
Maximum Area under disease
Proportion of progress curve Yield (g/plot)
foliage
infested
(sq.
units)
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0.76
0.86
5.64
13.90
1102
276
0.65
0.71
5.93
12.85
1778
642
0.46
0.47
4.11
9.28
2248
1426
0.098
0.40
0.98
7.33
2158
1452
0.0
0.0
0.014
0.0
2723
3214

88
TABLE 17. Regression equations for the relationship between level bean
rust disease and snap bean yield (Trial 1).
Plant growth stage
Regression equation
Third trifoliate leaf
Flower bud formation
Full bloom
Pod set
Pods half developed
Pods fully developed
Disease incidence negligible
It II II
y =
2666.71
- 29738.55x
(R2
=
0.9**)
y =
2428.64
- 5481.99d
(R2
=
0.83*)
y =
2718.09
- 3407.12x
(R2
=
0.93**)
y =
2812.42
- 181.8d
(R2
=
0.84*)
y =
2622.1
- 605.23x
(R2
=
0.69NS)
y =
2441.97
- 1617.92d
(R2
=
0.46NS)
y =
2618.35
- 196.94x
(R2
=
0.71NS)
y =
2627.26
- 1689.48d
(R2
=
0.8*)
* R significant at P ^ 0.05
** R significant at P < 0.01
y = yield (g/plot)
x = area under the disease progress curve
d = proportion of foliage infected
NS = not significant at 0.05

89
TABLE 18. Regression equations for the relationship between bean rust
and snap bean yield (Trial 2).
Plant growth stage
Regression equation
First and Third trifoliate Disease incidence negligible
Flower bud formation
y
=
2492.5 -
7790.18x
(R2
=
0.39NS)
y
=
2443.73 -
23678.58d
(R2
=
0.36NS)
Full bloom
y
=
1940.83 -
1141.86x
(R2
=
0.16NS)
y
=
1734.26 -
1384.95x
(R2
=
0.21NS)
Pod set
y
=
3113.46 -
893.33x
(R2
=
0.89*)
y
=
2886.63 -
5340.87d
(R2
=
0.92**)
Pods half formed
y
=
3099.74 -
369.12x
(R2
=
0.98**)
y
=
3220.42 -
3224.52d
(R2
=
0.99**)
Pods fully formed
y
=
3165.01 -
203.23x
(R2
=
0.98**)
y
=
3288.9 -
2795.75d
(R2
=
0.98**)
* R Significant at 0.05
** R Significant at 0.01
y = yield (g/plot)
x = area under the disease progress curve
d = proportion of foliage infected
NS = not significant

Proportion of Loss in Yield
90
Maximum Proportion of Foliage Infected
Figure 17. The influence of the maximum proportion of foliage infected
by Uromyces phaseoli on snap bean yield.

Proportion of Loss in Yield
91
Figure 18. The influence of the area under disease progress curve
on snap bean yield.

TABLE 19. The influence of Uromyces phaseoli on the gross dollar value per hectare of snap beans
(Phaseolus vulgaris 'Sprite').
Spray
frequency
(days)
Disease Parameters at harvest
Maximum Proportion of Area under disease
foliage infected progress curve
Price range
Gross dollar values
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0
0.76
0.86
5.64
13.9
high
1797
449
medium
987
247
low
539
135
14
0.65
0.71
5.93
12.85
high
2574
1047
medium
1416
576
low
772
314
7
0.46
0.47
4.11
9.28
high
3665
2327
medium
2016
1280
low
1100
698
4-5
0.098
0.4
0.98
7.33
high
3520
2367
medium
1936
1302
low
1056
710
7 3
0.0
0.0
0.014
0.0
high
4446
5244
medium
2446
2884
low
1334
1573
a
Sprayed with bitertanol

TABLE 20. The relationship between disease severity and net return per hectare of snap beans
sprayed with mancozeb and sulfur.
No. of
sprays
Fungicide
spray frequency
Maximum
Proportion of
foliage infected
Price range
Net return ($)
in investment
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0
0
0
0.76
0.86
high
0
0
medium
0
0
low
0
0
3
14-day
14
0.65
0.71
high
743
564
medium
395
295
low
199
145
5 (6) a
7-day
7
0.46
0.47
high
1812
1810
medium
973
965
low
505
495
7(8)
4-5-day
4-5
0.098
0.4
high
1644
1828
medium
870
965
low
438
485
a
Number of sprays in trial 2 are in parentheses.

94
ai
3
CO
>
)-i
CO
O
a
en
en
o
u
O
5000
4000
3000
2000
1000
0
0 0.098 0.46 0.65 0.76
Disease Severity
Figure 19. Influence of disease severity on gross dollar value per
hectare of 'Sprite' snap beans in trial 1.

95
5500
5000
4000
3000
2000
1000
0
0 0.40 0.47 0.71 0.86
Disease Severity
Figure 20. Influence of disease severity on gross dollar value per
hectare of 'Sprite' snap beans in trial 2.

96
was no benefit from spraying the plants more often than 7-day intervals.
By spraying at 4-5-day intervals there was a net loss in returns
compared to the 7-day intervals. The total cost per spray occasion per
hectare was $11.29.
In trial 2, there were increases in net returns, by spraying plants
at 7-day intervals compared to the 14-day spray schedule, of $1246,
$670, and $350 at the high, medium, and low prices. From the 14-day to
the 4-5 day spray schedules, there were increases in net returns of
$1264, $670, and $340 at the high, medium, and low prices. When plants
were sprayed at 7-day and 4-5-day intervals there was an increase in net
returns of $22 and $10 at the high and low prices but there was no net
increase in returns at the medium price. Thus, in trial 2 there was no
apparent benefit in spraying plants at 4-5-day intervals. The
additional sprays hardly paid for the extra cost of spraying more often.
Discussion
Significant relationships were found between yield and both disease
severity and area under the disease progress curves. Yields were
generally higher in trial 1 than in trial 2. The disease was clearly
established earlier in trial 2 than in trial 1.
The relationship between yield or loss in yield and bean rust was
well described using both parameters: maximum severity and area under
the disease progress curve (AUDC). Rate of disease progress was as well
2
correlated with yield as disease severity or AUDC in both trials with R
of 0.71 and 0.79 for trial 1 and trial 2 respectively. In trial 2,
o
disease severity and AUDC had coefficients of determination (R ) of 0.98

97
at the same growth stage. This discrepancy in correlation of rate of
disease progress and yield in these trials may be due to the early onset
of the disease in trial 2 where the rate of disease progress may have
started to decrease since most of the foliage had already been infected.
There is also a possibility that rates of disease progress were sporadic
in trial 2. Although AUDC was an equally good predictor of yield at
full bloom and pod set in trial 1 and at pod set through fully formed
pods in trial 2, it was a difficult parameter to use since it required
the use of computers. Computers are not readily accessible to extension
specialists who work closely with local farmers. Disease severity, on
the other hand, can be estimated by the use of graph papers or tracing
paper for the determination of leaf area before and after cutting out
diseased leaf tissue. Thus, disease severity would be a more convenient
parameter for the description of the relationship between bean rust and
snap bean yield. Moreover, AUDC is derived from disease severity. Area
under the disease progress curve is, however, a better representation of
the magnitude of the disease for the growing season. Disease severity
is, however, easier to measure and does not require complicated calcula
tions. Therefore, disease severity may be useful for the prediction of
yield at specific bean plant growth stages.
The Horsfall-Barratt disease rating system was not used in these
trials because of its subjectivity. This system has a lot of variabi
lity according to the person assessing the disease and would require the
comparison of several people's data to be relied on. The Horsfall-Barratt
method could, however, be adapted based on standardized diagrams put out
by extension.
At disease severity levels (maximum proportion of foliage infected)
below 0.5 (50%), it was not worth the expense of the shorter 4-5-day

98
mancozeb + sulfur spray interval in both trials. The grower would end
up losing income by spraying at this frequency. There was no apparent
explanation for the slight decrease in yield at the 0.098 disease
severity level in trial 1 (Table 16). The decrease in yield may have
been caused by adverse effects of the fungicide mixture on plants. The
fungicides used to manipulate the disease had zinc and sulfur which are
required by plants for proper growth. It was, however, not clear
whether the adverse effect was due to over supply of these elements or
due to the interaction of these elements and factors. This phenomenon
may have been a chance effect which requires further investigation under
similar growing conditions.
In trial 2, yield was consistently inversely related to disease
severity and AUDC. In this trial lower disease severities were correla
ted better with higher yields than the higher disease severities. The
lower disease severity corresponded with more frequent sprays of mancozeb
+ sulfur. These low disease parameters resulted from more frequent
sprays which cost more money. Thus, returns were reduced substantially.
Generally, the shorter 4-5-day mancozeb + sulfur intervals were more
costly than the 7-day intervals and the extra cost did not produce
enough yield to pay for itself. The optimum spray schedule for beans
was, therefore, every 7 days.
From these trials, the grower would be ill-advised to spray snap
beans at 4-5-day intervals if the disease severity is below 0.1 and the
onset of the disease is late. The economic threshold of bean rust
severity was below 0.1 since 0.098 disease severity resulted in 21%
yield loss.

99
There are many methods of determining disease severity in the
field. These methods include the use of leaf area meters, the Horsfall-
Barratt disease rating system, and pictorial keys. In these trials it
has been shown that the use of leaf area is a more convenient method for
disease severity assessment. Hence, in order to determine disease
severity the grower or extension specialist would have to take random
leaf samples at specific intervals and estimate the proportion of
foliage infected at each stage. Leaf area determination would be
facilitated if a leaf area meter were available but where this was not
the case then graph or trace paper could be used. When computers are
accessible to the grower/extension specialist then AUDC can be deter
mined based on disease severity. The choice of the variable would
depend on availability of expertise and equipment.
Yields were inversely related to disease parameters at specific
bean plant growth stages. Rate of disease progress, AUDC and disease
severity correlated well with yield. Therefore, the extension worker or
grower can choose the parameter of disease to measure. These disease
parameters are, however, time consuming which may increase labor and
other costs.

CHAPTER VI
THE EFFECT OF DEFOLIATION, METAM-SODIUM, AND BEAN RUST ON SNAP BEANS
Introduction
Beans, Phaseolus vulgaris L., are subject to defoliation by a wide
range of factors including insects, diseases, adverse environmental
conditions, mammals, and farm machinery (Agudelo, 1980; Allen, 1983;
Cook, 1978; Costa and Rossetto, 1972; Ruppel and Idrobo, 1962; Vargas,
1980). The extent to which these factors influence yield depends on the
plant growth stage at which they are attacked. Among the most important
insect pests feeding on bean leaves are leafminers (Liriomyza spp.),
cabbage loopers ((Hub.) Trichoplusia ni), leafrollers (Urbanus proteus
L.), Mexican bean beetles (Epilachna varivestis Muls.), and Chrysomelid
beetles (Schoonhoven and Cardona, 1980). Bean rust, Uromyces phaseoli
(Pers.) Wint., is one of the most damaging diseases of bean in bean
producing regions of the world (Acland, 1971; Allen, 1983; Cook, 1978;
Crispin and Dongo, 1962; Iraneta and Rodrigez, 1983; Martinez, 1983;
Schwartz et al., 1979; Stoetzer and Omunyin, 1983; Vargas, 1980). Other
diseases which affect beans include anthracnose (Colletrotrichum
lindemuthianum (Sacc. and Mgr.) Bri. and Cav.), angular leaf spot
(Isariopsis griseola Sacc.), halo blight, (Pseudomonas syringae pu.
phaseolicola (Burkh.) Young, Dye and Wilkie, common blight (Xanthomonas
campestris pu. phaseoli (Smith) Dye, and bean common mosaic virus
(Acland, 1971; Allen, 1983; Martinez, 1983; Stoetzer and Omunyin, 1983).
Root rots caused by Rhizoctonia solani Kuhn, Macrophomina phaseolina
100

101
(Tassi) Gold, Sclerotium rolfsii Sacc., Pythium spp., and Fusarlum spp.
have also been reported to initiate wilting and eventually defoliation
(Martinez, 1983). Many nematode species are found in association with
bean roots in various parts of the world (Agudelo, 1980, Allen, 1983).
The root-knot nematode complex (Meloidogyne spp.) is among the most
damaging on beans (Agudelo, 1980; Allen, 1983; Ngundo, 1977; Ngundo and
Taylor, 1974, 1975a,b). Thus, an understanding of the relationship
between these factors and yield is a prerequisite for the development of
a sound pest management strategy.
Information on the relationship between leaf damaging pests and
yield has been obtained through pest damage simulation by manually
defoliating plants at various growth stages and at several defoliation
levels (Edje and Mughogho, 1976a,b; Edje et al., 1972, 1973, 1976;
Galvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho,
1983; Vieira, 1981; Waddill et al., 1984). Ruesink and Kogan (1975)
observed that manual defoliation is not precise in simulating pest
damage. The imprecision in pest damage simulation may be due to the
exact timing of manual defoliation and careful determination of the
proportion of the foliage to be removed, which pests cannot do. More
over, manual defoliation does not introduce saliva and possible phyto
toxins which may be important factors in the plant damage caused by
specific pests.
Rarely are crop plants attacked by one pest species only. More
often several species attack a crop at the same time. Thus, McSorley
and Waddill (1982) studied the effect of insect and nematode pests on
squash (Cucrbita pepo L.). They partitioned yield loss into insect and
nematode components by using multiple regression procedures. It was

102
observed that the prediction of yield loss was more accurate when both
insect and nematode pests were present. Bookbinder and Bloom (1980)
reported that Meloidogyne spp. interacted with bean rust, Uromyces
phaseoli, on beans. The root-knot nematodes and the disease had an
additive effect on the suppression of shoot and root weights of bean
plants. Meloidogyne incognita infections reduced uredial diameter of _U.
phaseoli. Similar effects were observed if IJ. phaseoli was inoculated
first. Bookbinder and Bloom (1980) observed that rusted plants had 62%
less M. incognita than uninfected plants. They suggested this was due
to suppressed translocation of photosynthates to the roots. _U. phaseoli
infection did not affect M. incognita egg hatch.
This study was conducted to determine the relationship between
defoliation, nematodes and bean rust and yield of snap beans.
Materials and Methods
General
Three trials were conducted at the Tropical Research and Education
Center in Homestead, Dade County, Florida. Experimental sites were on
Rockdale soil (pH ca. 7.8) planted in fall 1984 and early spring 1985.
The fields had been previously cropped in tomato (Lycopersicon
esculentum Mill.). In all three trials snap beans (Phaseolus vulgaris
L. 'Sprite') were used. Prior to planting, the herbicides Treflan^^
(R)
(841 ai/ha) and Dual" (1.7 kg ai/ha) were applied to the site.
Fertilizer (8:16:16) was applied preplant at 448 kg/ha according to the
University of Florida Extension recommendation (Stall and Sherman,
1983). The plants were topdressed with 224 kg/ha fertilizer at flower

103
bud formation. Metam-sodium was used to manipulate nematode
populations. Subsequent weeding was done by cultivation. Irrigation
was provided by an overhead sprinkler system.
The effect of metam-sodium on snap beans
The crop was planted on 26 November 1984. Individual plots consis
ted of 3 rows 3m long with 0.91m between rows. Seeds were planted at
8-10 cm spacing. Treatments were replicated 4 times in a randomized
complete block. Metam-sodium was applied preplant at 0, 47, 94, 187,
281, and 374 L/ha. Preplant soil samples consisted of a composite
mixture of 10 soil scoops (to a 6-8 cm depth) from each plot 12 days
after fumigation. The 12-day period was based on the observations made
by McSorley and Parrado (1984). Aliquots of 100 ml soil were processed
by sieving and centrifugal flotation (Jenkins, 1964). Subsequent soil
samples at mid-season and harvest were taken from the root zone and
similarly extracted. Only live nematodes were counted in the preplant
samples but in the later samples, nematodes were first killed by gentle
heating in a water bath (55-60C) and counted. Beans were harvested on
31 January 1985.
Yield data were subjected to analysis of variance and regression
analysis using the general linear models procedure of SAS (Ray, 1982).
The effect of metam-sodium and defoliation on snap beans
The crop was planted on 26 November 1984. Individual plots consis
ted of 4 rows 3 m long with 0.91 m between rows. Seeds were planted at
8-10 cm spacing within the row. A split-plot design was used in this
trial to investigate the effect of metam-sodium (main plots) and defoli
ation (subplots) on snap beans. Treatments were replicated 4 times.
Each subplot consisted of a 3 m long row. Defoliation levels

104
investigated were 0%, 25%, 50%, and 75% and metam-sodium was applied at
0, 47, 94, 187, and 374 L/ha. Metam-sodium was applied preplant.
Preplant soil samples were taken 12 days after fumigation by compositing
10 soil scoops (to a depth of 6-8 cm) from each main plot. Aliquots of
100 ml soil were processed by sieving and centrifugal flotation.
Subsequent soil samples were taken at midseason and harvest from the
root zone and similarly treated. Only live nematodes were counted from
the preplant soil samples and in the later samples nematodes were killed
before counting.
Defoliation was accomplished by removing the lamina from the distal
end of the petiole using a pair of scissors. Plants were defoliated
once at flower bud formation. Beans were harvested on 31 January to 1
February 1985.
Data were analyzed by analysis variance and covariance followed by
regression analysis using the general linear models procedure of SAS
(Ray, 1982).
The effect of defoliation, metam-sodium and bean rust on snap beans
A 4x3x2 factorial experiment was conducted at the Tropical Research
and Education Center in Homestead, Dade County, Florida, on Rockdale
soil (pH ca. 7.8). The crop was planted on 26 March 1985. Plots
consisted of 3 rows, 3 m long with 0.91 m between rows. Seeds were
planted at 8-10 cm spacing within the row. Fertilizer (8:16:16) was
applied preplant at 448 kg/ha and plants were topdressed at flower bud
formation at 224 kg/ha.
Metam-sodium was applied at 935 L/ha and a fumigated control was
included. Defoliation levels investigated were 0, 25%, and 50%. Bean
rust was manipulated by sprays of bitertanol (57 g ai/ha) at 7-day

105
Intervals, mancozeb (1.7 kg ai/ha) plus sulfur (4.5 kg ai/ha) at 7-day
and 14-day Intervals. A no-spray plot was included to ensure a high
(R)
disease level at specific times of assessment. Helenav sticker or Nu
(R)
Filmv -17 was used as a surfactant in all fungicide sprays.
Preplant soil samples were taken by compositing 10 soil scoops from
each plot 12 days after fumigation. Aliquots of 100 ml soil were
processed by sieving, and nematodes were extracted by centrifugal
flotation (Jenkins, 1964). Subsequent soil samples taken at mid season
and at harvest were similarly handled. Only live nematodes were counted
in the preplant soil samples. In the later samples the nematodes were
first killed by heating in a water bath and then counted.
Plants were subjected to rust inoculum at the primary leaf stage by
clipping infected pole bean leaves on to wire stakes just east of the
test plots and 25 cm off the ground. Disease progress was monitored by
taking 5 trifoliate leaves from each plot once a week. The leaf area
before and after cutting diseased leaf tissue was determined. The leaf
samples were taken from the same general position in the canopy at each
sampling.
For the defoliation treatments, plants were manually defoliated
with pairs of scissors. The foliage was removed from the distal end of
the petiole. Defoliation was done only once, at full bloom because
results from other workers indicated that this was a critical growth
stage for beans (Hohmann and DeCarvalho, 1983).
Data were subjected to analysis of variance and regression analysis
using the general linear models procedure of SAS (Ray, 1982).

106
Results
Effect of metam-sodlum on snap beans
Tables 21, 22 and 23 show the numbers of nematodes in 100 ml
aliquots of soil at preplant, mid-season and harvest respectively. Two
nematode genera, Criconemella and Rotylenchulus, were found in the soil
at preplant (Table 21). At mid-season, four nematode genera were
detected in the soil: Helicotylenchus, Meloidogyne, Rotylenchulus, and
Tylenchorhynchus (Table 22). Meloidogyne had the highest numbers in
unfumigated plots (Table 22), but at this time, the effect of metam-
sodium on nematode population was not proportional to its rate of
application. Criconemella, Helicotylenchus, Meloidogyne, and
Rotylenchulus were found in the soil at harvest (Table 23).
Metam-sodium had no significant effects on nematode numbers at any
sampling (Tables 21, 22, and 23).
Analysis of variance on snap bean yield showed that there were
significant differences among metam-sodium rates (F = 3.4*). Yield
responses were, however, not consistently proportional to metam-sodium
rates (Table 21). Regression analysis on the relationship between
2
metam-sodium and yield produced models of the form Y = -0.004x + 345.75
(R2 = 0.17) and Y = -0.00004x3 0.03x2 + 4.9x + 287.5 (R2 = 0.22) where
Y = yield (g/plot), and x = metam-sodium rate (L/ha). The cubic model
gave a higher coefficient of determination than the quadratic model
(Figure 21). The coefficients of determination were very low. The
linear model of the form Y = a + bX gave an R of only 0.12 indicating
that yield was not linearly related to metam-sodium rate. Multiple
regression on nematode general effects on yield gave the model Y = a +
bl X1 + b2 X2 + '* + bn xn where Y = yield, x = log (nematode population

107
2
+ 1). The linear models gave low R at all soil sampling times (Table
24).
Gross dollar values per hectare are shown in Table 25. The highest
dollar value was obtained with the 374 L/ha soil fumigation, as
expected, since this was the rate at which the highest yield was
achieved. The gross dollar values were based on yield per hectare and
the following prices $6.00 (low), $11.00 (medium), and $20.00 (high) per
bushel of snap beans (13.62 kg). Net income is shown in Table 26.
Yields obtained in this study were generally low. These yields were low
probably because of the less ideal temperatures at the time the study
was conducted. Under these conditions the grower would have made a
profit at 47 L/ha, and 94 L/ha at the low, medium, and high snap bean
prices respectively (Table 26). The dollar values were based on the
metam-sodium cost of $1.59/L (McSorley and Pohronezny, 1984); no other
expenses have been used in economic analyses.
The effect of metam-sodium and defoliation on snap beans
Analysis of variance on the influence of metam-sodium, defoliation,
and their interaction on snap beans showed that there was no significant
interaction between metam-sodium and manual defoliation (Table 27).
There were also no significant differences among metam-sodium rates (F =
1.5 NS) on yield. There were, however, significant differences among
defoliation levels (F = 20.22**) (Table 27). Regression analysis of
2 2
yield data on metam-sodium rates produced the model y = a + bx cx (R
= 0.34) where y = yield (g/plot), x = metam-sodium rate (gal/acre)
(Figure 21). The cubic model of the form y = a + bx + cx + dx (R =
0.46) was obtained when yield data were analyzed by regression against
manual defoliation (Figure 28).

108
TABLE 21. Effect of metam-sodium on snap bean yield and populations of
nematode genera in preplant soil samples. Data are means of
3 replicates.
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Yield Yield loss
(g/plot)
0
1
21
360
49
47
1
27
596
16
94
4
72
650
8
187
1
17
680
4
281
0
42
581
18
374
0
25
707
0

Yield (g/plot)
109
0.00004x3
Figure 21.
Effect of metam-sodium on snap bean yield.

TABLE 22. Effect of metam-sodium on snap bean yield and populations of nematode genera in midseason
soil samples. Data are means of 3 replicates.
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus Meloidogyne Rotylenchulus Tylenchorhynchus Yield
(No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml) (g/plot)
0 0
47 0
94 2
187 0
281 0
374 0
3 12 0
8 8 0
4 0 1
1 7 0
3 110
1.3
3
1
4
8
0
1
5
2
360
596
650
680
581
707
110

TABLE 23. Effect of metam-sodium on snap bean yield and populations of nematode genera in final soil
samples (at harvest).
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Meloidogyne
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Yield
(g/plot)
0
1
21
4
25
360
47
1
27
6
1
596
94
4
72
1
4
650
187
1
17
0
0
680
281
0
42
0
18
581
374
0
25
0
1
707
Ill

112
TABLE 24. Regression equations for the relationship between various
nematode populations, during the growing season, and snap
bean yield. Nematode populations were transformed to log
(population +1).
Time
Regression Equation
a
Coefficient of
determination (R )
Preplant
y
=
497.3-21x,+7.lx.
1 4
0.025
NS
y
=
538.4-20.3Xj
0.018
NS
Midseason
y
=
894.6-191.lx,-11.lx-25.4x-66.6x.-8.2x_
1 2 3 4 5
0.17
NS
y
=
902.7-200.6x -12.3x -29.6x -67.2x.
12 3 4
0.17
NS
y
=
841.7-220.7xj-24.8x2~20.3x3
0.14
NS
y
=
767.7-187.2Xj-30.4x2
0.13
NS
y
=
770.4-251.61Xj
0.11
NS
Harvest
y
=
797.3-157.6x.-35.5x0-28.6x+9.4x,
12 3 4
0.17
NS
y
=
783.5-142.8Xj-27.lx2-25.8x3
0.17
NS
y
=
731.7-132xj-27.3x2
0.15
NS
y
=
682.8-137Xj
0.13
NS
a
= Criconemella
= Helicotylenchus
= Meloidogyne
= Rotylenchulus
= Tylenchorhynchus
NS = Not significant at 0.05

113
TABLE 25. The effect of metam-sodium on gross dollar values per hectare
of beans.
Price Range
Vapam
(liters/ha)
low
medium
high
0
176
323
587
47
292
535
972
94
318
583
1061
187
333
610
1094
281
285
522
948
374
346
634
1153

114
TABLE 26. Net returns on investment per hectare of snap beans.
Metam-sodium
(Liters/ha)
Price Range
low
medium
high
0
47
94
187
281
0
0
0
41
137
311
- 7
112
325
-141
- 10
210
-338
-247
- 85
-425
-283
- 29
374

115
TABLE 27. F-values from analysis of variance for the effects of metam-
sodium, defoliation and their interaction on snap bean yield.
Source
F
Probability of F
Metam-sodium
1.5
0.22
Defoliation
20.22
0.0001
Defoliation x
metam-sodium
0.62
0.82

TABLE 28. Nematode genera found in soil samples collected (preplant). Data are means of 4
replicates
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Tylenchorhynchus
(No./lOO ml)
0
0
0
4
4
47
0
1
6
2
94
0
3
0
2
187
0
0
2
.0
374
1
0
2
0
116

117
TABLE 29. Nematode genera found in soil samples (midseason). Data are
means of 4 replicates.
Metam-sodium Helicotylenchus Meloidogyne Rotylenchulus
(Liters/ha) (No./lOO ml) (No./lOO ml) (No./lOO ml)
0
3
15
5
47
3
19
3
94
4
15
0
187
8
19
4
374
0
19
3

TABLE 30. Nematode genera found in soil samples (harvest). Data are means of 4 replicates.
Metam-sodium Criconemella Helicotylenchus Meloidogyne Rotylenchulus Tylenchorhynchus
(Liters/ha) (No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml)
0
47
94
187
374
0
0
2
0
0
8
5
2
2
0
4
8
6
5
3
9
9
9
9
4
0
1
1
1
0
00

119
TABLE 31. Mean snap bean yield of plants defoliated at various levels
on plots fumigated with metam-sodium.
Defoliation level
(proportion of foliage)
Metam-sodium
(Liters/ha)
Yield (g/plot)
Yield loss (%)a
0
0
545
36
0
47
551
35
0
94
738
13
0
187
851
0
0
374
568
33
0.25
0
469
45
0.25
47
417
51
0.25
94
596
30
0.25
187
403
53
0.25
374
423
50
0.50
0
264
69
0.50
47
253
70
0.50
94
371
56
0.50
187
330
61
0.50
374
335
61
0.75
0
263
69
0.75
47
186
78
0.75
94
276
68
0.75
187
194
77
0.75
374
284
67
£
Yield loss compared to
maximum yield of
851 g.

120
TABLE 32. The effect of defoliation and metam-sodium
value per hectare of snap beans.
on the
gross
dollar
Gross
dollar
values
Defoliation
level
Metam-sodium
by
price
range
(proportion of
foliage)
(Liters/ha)
Low
Medium
High
0
0
267
489
889
0
47
269
494
898
0
94
361
662
1204
0
187
416
763
1388
0
374
278
510
927
0.25
0
230
421
765
0.25
47
204
374
681
0.25
94
292
535
972
0.25
187
197
362
658
0.25
374
207
279
690
0.5
0
129
237
431
0.5
47
124
227
412
0.5
94
182
333
605
0.5
187
162
296
539
0.5
374
164
301
547
0.75
0
129
236
429
0.75
47
91
167
303
0.75
94
135
247
450
0.75
187
95
174
315
0.75
374
164
255
463

121
TABLE 33. Net returns on investment ($) per hectare of snap beans.
Plants were defoliated at various levels and soil treated
with metam-sodium.
Defoliation level Metam-sodium Net returns by price range
(proportion of foliage) (Liters/ha) Low Medium High
0
47
- 27
- 25
- 21
0
94
34
112
255
0
187
29
154
379
0
374
-229
-220
-203
0.25
0
- 37
- 68
-124
0.25
47
- 93
-145
-238
0.25
94
- 35
- 14
- 23
0.25
187
-190
-247
-351
0.25
374
-300
-350
-440
0.50
0
-137
-252
-458
0.50
47
-173
-292
-507
0.50
94
-145
-217
-345
0.50
187
-225
-313
-470
0.50
374
-343
-429
-583
0.75
0
-138
-253
-460
0.75
47
-206
-352
-616
0.75
94
-192
-301
-499
0.75
187
-292
-436
-694
0.75
374
-343
-475
-666

122
TABLE 34-1. F values from the analysis of variance for yield.
Source
F value
Probability of F
Defoliation
3.06
0.07
Metam-sodium
0.12
0.73
Defoliation x
Metam-sodium
2.20
0.12
Fungicide
7.62
0.0002
Defoliation x
Fungicide
1.15
0.34
Metam-sodium x
Fungicide
1.41
0.25
Defoliation x
Fungicide x
Metam-sodium
0.62
0.72

123
TABLE 34-2. Mean snap bean yield per plot sprayed with fungicides.
Fungicide
Spray
Frequency
Q
Mean yield (g/plot)
No fungicide
0
284a
Mancozeb
14-day
396a
Mancozeb
7-day
342a
Bitertanol
7-day
616b
Means followed by the same letter are not significantly different at
P 0.05 (Duncan's multiple range test).

TABLE 35. Regression equations for the effect of defoliation, nematodes, and bean rust on snap beans.
Time of assessment of disease
Regression equation
Flower bud formation
(preplant nematode counts)
Full bloom
(preplant nematode counts)
Pod formation
(preplant nematode counts)
Pods half-developed
(preplant nematode counts)
Pods fully formed
(preplant nematode counts)
y = 637.7-262.7x^-6608.9x2+26.3x^-62x^+2.8x^
y = 638.8-332.3x^6497.6x2
y = 547.6-6164.IX2
y = 539.6-234.8x,-3269.5x+55.lx+25.5x.-1.32xc
1 2 3 4 5
y = 653.56-314.82x^299.2
y = 572.46-295.5x2
y = 695.7-290x1-947.7x+7.6x_-102.6x.+9.8x[;
1 2 3 4 5
y = 646.1-348.4x^-925x2
y = 549.6-870.6x2
y = 740.4-275.2x,-569.lx0-2.6x_-78.lx.+2.lxc
1 2 3 4 5
y = 665.9-301.2x^-564x2
y = 590.4-563.3x2
y = 702.8-283.4x^398.3x2+2.2x3-ll.lx4-2.3x5
(R
(R2
(R2
(R2
(R2
(R2
(R2
(R2
(R2
(R
(R2
(R2
(R
0.45**)
0.42**)
0.26**)
0.67**)
0.61**)
0.47**)
0.56**)
0.52**)
0.36**)
0.62**)
0.61**)
0.48**)
0.66**)
124

TABLE 35. Continued.
Time of assessment of disease
Regression equation
1
Pods fully formed
(final nematode counts)
Pods fully formed
(final nematode counts)
y = 687.1-288.9x^397.7x2
y = 616.7-401x2
y = 702.8-283.4x^398.3x2+2.2x3-11.1x4-2.3x5
y = 687.1-288.9x^397.7x2
y = 616.7-401x2
(R2 = 0.66**)
(R2 = 0.54**)
(R2 = 0.66**)
(R2 = 0.66**)
(R2 = 0.54**)
y = 869.8-230.4x -415.9x0+6x0-18.3xc+6x,-42.4x_, /n2
1 j 3 0 / V.K
y = 734.6-282.5x^410.7x2-33x? (R2
y = 616.7-401.4x2 (R2
0.69**)
0.66**)
0.54**)
y = yield (g/plot; x^ = defoliation level; x2 = disease severity; x^ = Heliotylenchus log ( x+1);
x, = Meloidogyne log ( x+1); x,. = Rotylenchulus log ( x+1); x, = Tylenchorhynchus log ( x+1); x7 =
Criconemella log ( x+1).
** R significant at P < 0.01.
125

TABLE 36. Nematode genera and disease severity values found in plots fumigated with metam-sodium.
Figures are means of 3 replicates (Preplant soil samples).
Defoliation
(proportion of
level
foliage)
Disease severity
(maximum
Metam-sodium proportion of
(Liters/ha) foliage infected)
Helicotylenchus
(No./lOO ml)
Meloidogyne
(no./100 ml)
Rotylenchulus
(No./lOO ml)
0
0
0.02
0
0
19
0
935
0.05
0
0
16
0.25
0
0.04
5
0
5
0.25
935
0.04
0
0
2
0.50
0
0.03
0
0
3
0.50
935
0.03
5
0
2
0
0
0.01
5
0
36
0
935
0.01
0
0
15
0.25
0
0
5
0
37
0.25
935
0.03
0
0
15
0.50
0
0.0
5
0
37
0.50
935
0.0
0
0
15
0
0
0.02
0
0
13
0
935
0.04
5
0
27
0.25
0
0.02
0
0
9
0.25
935
0.02
0
0
9
0.50
0
0.02
0
5
2
0.50
935
0.5
0
0
12
0
0
0.04
10
0
12
0
935
0.02
10
0
45
0.25
0
0.03
0
0
6
0.25
935
0.02
0
0
3
0.50
0
0.03
10
0
30
0.50
935
0.01
0
0
40
126

TABLE 37. Nematode genera + disease severity values found in plots fumigated with metam-sodium. Figures are
means of 3 replicates (Soil samples at harvest).
Disease severity
Defoliation level (maximum
(proportion
of foliage)
Metam-sodium
(Liters/ha)
proportion of
foliage infected)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Tylenchorhynchui
(No./lOO ml)
0
0
0.87
0
13
85
10
0
935
0.83
0
2
77
41
0.25
0
0.64
0
13
93
13
0.25
935
0.71
0
8
32
10
0.5
0
0.77
0
5
134
8
0.5
935
0.74
2
2
47
3
0
0
0.02
2
28
66
23
0
935
0
0
15
77
2
0.25
0
0
2
7
125
15
0.25
935
0.01
2
23
108
25
0.5
0
0
0
2
78
7
0.5
935
0
2
2
83
18
0
0
0.56
2
18
87
7
0
935
0.59
0
13
77
7
0.25
0
0.7
0
5
98
3
0.25
935
0.66
0
17
67
18
0.5
0
0.68
2
5
103
15
0.5
935
0.71
0
8
118
3
0
0
0.71
0
23
135
20
0
935
0.74
0
7
45
5
0.25
0
0.75
2
13
120
3
0.25
935
0.83
2
7
45
12
0.5
0
0.72
0
8
100
17
0.5
935
0.81
0
20
129
28
127

Proportion of Foliage Infected
128
Days After Inoculation
Figure 22. Disease progress curve for Uromyces phaseoli on snap
beans not sprayed with mancozeb, defoliated at various
levels and soil fumigated with metam-sodium.
1 = no defoliation, no metam-sodium and no fungicide, 2 = no defoliation,
935 L/ha metam-sodium, and no fungicide, 3 = 25% defoliation, no metam-
sodium and no fungicide, 4 = 25% defoliation, 935 L/ha metam-sodium and
no fungicide, 5 = 50% defoliation, no metam-sodium and no fungicide, 6 =
50% defoliation, no metam-sodium and no fungicide.

129
Figure 23. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at 7 day intervals, defoliated at
various levels and soil fumigated with metam-sodium.
13 = no defoliation, no metam-sodium and mancozeb + sulfur at 7-day inter
vals, 14 = no defoliation, 935 L/ha metam-sodium and mancozeb + sulfur at
7-day intervals, 15 = 25% defoliation, no metam-sodium and mancozeb 4-
sulfur at 7-day intervals, 16 = 25% defoliation, 935 L/ha metam-sodium
and mancozeb + sulfur at 7-day intervals, 17 = 50% defoliation, no metam-
sodium and mancozeb + sulfur at 7-day intervals, and 18 = 50% defoliation,
935 L/ha metam sodium and mancozeb + sulfur at 7-day intervals.

130
Days After Inoculation
Figure 24. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at 14-day intervals, defoliated at
various levels and soil fumigated with metam-sodium.
19 = no defoliation, no metam-sodium and mancozeb + sulfur at 14-day
intervals, 20 = no defoliation, 935 L/ha metam-sodium and mancozeb +
sulfur at 14-day intervals, 21 = 25% defoliation, no metam-sodium and
mancozeb + sulfur at 14-day intervals, 22 = 35% defoliation, 935 L/ha
metam-sodium and mancozeb + sulfur at 14-day intervals, 23 = 50% de
foliation, no metam-sodium and mancozeb + sulfur at 14-day intervals,
and 24 = 50% defoliation, 935 L/ha metam-sodium and mancozeb + sulfur
at 14-day intervals.

TABLE 38. Effect of defoliation, metam-sodium and bean rust on yield of gross dollar value per hectare of
snap beans. Data are means of 3 replicates.
Disease severity Gross dollar values
Defoliation level
(proportion of foliage)
Metam-sodium
(Liters/ha)
(proportion of
foliage infested)
Yield
(g/plot)
high
Price Range
medium
low
0
0
0.87
184
300
165
90
0
935
0.83
365
596
328
179
0.25
0
0.64
260
424
233
127
0.25
935
0.71
345
563
310
169
0.5
0
0.77
220
359
197
108
0.5
935
0.74
338
551
303
165
0
0
0.02
807
1316
724
395
0
935
0
827
1349
742
405
0.25
0
0
570
930
512
279
0.25
935
0.01
499
814
448
244
0.5
0
0
446
727
400
219
0.5
935
0
548
895
492
268
0
0
0.56
349
569
313
171
0
935
0.59
573
935
514
281
0.25
0
0.70
520
848
467
255
0.25
935
0.66
276
450
247
135
0.5
0
0.68
258
420
231
126
0.5
935
0.71
143
233
128
70
0
0
0.71
363
593
326
178
0
935
0.74
406
662
364
199
0.25
0
0.75
190
310
171
93
0.25
935
0.83
359
585
322
176
0.5
0
0.72
327
533
293
160
0.5
935
0.81
397
648
357
196
131

132
TABLE 39. Net
returns ($) on
investment per
hectare
of snap beans.
Defoliation
Net Income
level
Metam-sodium
Disease
Price Range
(proportion
of foliage)
(Liters/ha)
severity
high
medium low
0
0
0.87
300
165
90
0
935
0.83
-1191
-1324
-1398
0.25
0
0.64
124
68
38
0.25
935
0.71
-1224
-1342
-1408
0.5
0
0.77
59
32
9
0.5
935
0.74
-1236
-1349
-1562
0
0
0.02
1016
589
305
0
935
0
- 438
910
-1172
0.25
0
0
670
346
189
0.25
935
0.01
- 973
-1204
-1332
0.5
0
0
427
235
128
0.5
935
0
- 892
-1160
-1308
0
0
0.56
213
92
81
0
935
0.59
- 908
-1194
-1352
0.25
0
0.7
420
245
108
0.25
935
0.66
-1394
-1461
-1498
0.5
0
0.68
64
10
- 20
0.5
935
0.71
-1610
-1580
-1563
0
0
0.71
263
161
54
0
935
0.74
-1158
-1321
-1412
0.25
0
0.75
- 24
- 28
- 31
0.25
935
0.83
-1235
-1364
-1435
0.5
0
0.72
199
94
36
0.5
935
0.81
-1194
-1451
-1436

133
Tables 28, 29, and 30 show nematode genera detected in the soil at
preplant, midseason, and at harvest. Metam-sodium rate had no signifi
cant effects on nematode numbers at all sampling times.
Total yield (g/plot) and yield loss are shown in Table 31. The
lowest yield was obtained from plots fumigated at 47 L/ha metam-sodium
and plants defoliated at the 0.75 level. Gross dollar values are given
in Table 32 and net income is shown in Table 33. These gross dollar
values were computed from total yield (per hectare) multiplied by the
following price ranges: $6.00 (low), $11.00 (medium), and $20.00 (high)
per bushel (13.62 kg). Net dollar values were obtained by subtracting
the cost of metam-sodium per hectare from the gross dollar values.
Gross dollar values generally decreased with the increase in defoliation
level. There were net returns on investment when plants were not
defoliated (Table 33). There were no net returns on investment when
plants were defoliated at any level (Table 33).
The effect of defoliation, metam-sodium, bean rust, and their interaction
on snap bean yield
Analysis of variance showed that there were no significant
differences among defoliation levels and metam-sodium rates at the 0.05
level (Table 34-1). There was no significant interaction among
defoliation levels, metam-sodium rates, and fungicides at the 0.05 level
(Table 34-1). There were, however, significant differences among
fungicide sprays (F = 7.62**). When analysis of variance was used for
yield data analysis with disease severity as one of the independent
factors, there were no significant differences in yield based on disease
severity (maximum proportion of foliage infected) (F < 1). There was no
significant interaction among defoliation levels, metam-sodium rates,

Proportion of Loss in Yield
134
Figure 25.
The relationship between area under disease progress curve
and yield loss of snap beans.

Proportion of Loss in Yield
135
Figure 26. The relationship between yield loss and maximum proportion
of foliage infected.

Yield (g/plot)
136
igure 27.
The effect of metam-sodium on nsap bean yield when plants
were not defoliated.

930
880-
780-
680
580
480
380
280
180
-gure
137
y = 650.4-772.4x-45x2+462.4x3
R2=0.46
0.25 0.5
Defoliation Level
0.75
The general relationship between defoliation and
snap bean yield at all metam-sodium rates.

138
and disease severity. Since there were significant differences among
fungicides means were separated using Duncan's multiple range test
showed that there were no significant differences between mancozeb
sprays and the unsprayed treatment. There was, however, significant
difference between bitertanol and mancozeb sprays (Table 34-2).
Regression analysis on the effect of defoliation, nematodes, and
disease severity on snap bean yield produced models of the form: y = a
+ b. X, + b0 X. + ... + bnxn, where y = yield (g/plot), x, ...x =
l l z z In
independent variable (defoliation level, disease severity, and log
(nematode population + 1) (Table 35). Stepwise regression showed that
disease severity contributed most to the coefficients of determination
2
(R ) (Table 35). Since there were no significant differences among
defoliation levels and metam-sodium rates, further explanations in this
section will be restricted to disease severity or area under the disease
progress curve (AUDC). Thus, figures 25 and 26 show the relationship
between yield loss and AUDC and disease severity, respectively. The
relationship between these disease parameters and yield should be the
inverse of their relationship with yield loss. Regression analysis of
yield data produced models of the form y = a-bx, where y = yield
(g/plot), x = either disease severity or AUDC (Table 35). The
2
coefficients of determination (R ) for disease severity and AUDC were
0.54 and 0.52 respectively when the disease was assessed at harvest.
Coefficients of determination for disease severity for the other times
of disease assessment are given in Table 35. Since disease assessment
2
severity consistently had better R than AUDC, it was generally used in
yield data analysis. Moreover, a set of disease severity points results
in AUDC. Disease progress curves are shown in figures 22, 23, and 24.

139
Disease progress curves in unsprayed controls and those sprayed with
mancozeb + sulfur at 14-day intervals were similar (Figures 22 and 24).
The 7-day spray schedule of mancozeb produced progress curves with
maximum disease severity below those of the other two spray schedules
(Figure 23).
Vapam had no significant effect on nematode populations at preplant
and harvest (Tables 36 and 37). There were virtually no nematodes
detected at midseason.
Yield and gross dollar values per hectare are shown in Table 38.
The highest yield and gross dollar values were obtained from fumigated
plots with plants kept virtually disease free and not defoliated. The
lowest yield was obtained from fumigated plots with plants which had
0.50 defoliation level and 0.71 disease severity (Table 38). Generally,
yield responses were not proportional to defoliation levels and
metam-sodium rates. Fumigation resulted in net loss of income regard
less of defoliation level and disease severity (Table 39). No defolia
tion, 0.83 disease severity, and fumigation with vapam at 374 L/ha
resulted in loss of income of $1191, $1320 and $1398 at the high, medium
and low prices respectively whereas plots not fumigated, plants not
defoliated, and having 0.87 disease severity had net gains in income of
$300, $165 and $90 at the high, medium and low prices respectively
(Table 39). Net gains in income varied in magnitude depending on
disease severity and defoliation.
Discussion
The lack of consistent reduction in nematode populations on
treatment with metam-sodium may be related to application technique.

140
There may have been less than adequate retention of the nematicide by
the soil. This less adequate retention of the nematicide may have
contributed to its apparent poor efficacy. Nematode populations were
also generally low at all sampling times in the three tests.
In the study where metam-sodium alone was used, there was signifi
cant relationship between metam-sodium rates and yield. This signifi
cant difference may be due to metam-sodium controlling some soilborne
plant diseases on beans since the nematicide had no significant effect
on nematode populations and populations were very low.
Metam-sodium had no single or interactive effects on yield when it
was used in combination with defoliation and/or bean rust in these
tests. Defoliation was, however, the more important factor affecting
yield when simultaneously used with metam-sodium. The non-significant
interaction between defoliation and metam-sodium may be due to the
generally low nematode populations on the site the study was conducted.
In the study where defoliation, metam-sodium, and bean rust were
used simultaneously there were no significant interactions among the
three factors. There were no significant differences in yield among
defoliation levels, disease severity levels and metam-sodium rates.
There were, however, significant differences in yield among fungicide
sprays. Lack of interaction among the three factors may be due to low
nematode populations and overriding effects of the disease. The disease
was continuously associated with the crop from the time of inoculation
to harvest. The overriding effects were shown by regression analysis
which indicated that disease severity contributed most to the coeffi
cient of determination. Thus, the continuous association of the disease
with the crop may have influenced the physiology of the plants which

141
which was reflected in yield. The overriding effect of bean rust on
yield was complicated by the fact that it was manipulated by fungicides
which had micronutrients required for plant growth. Duncan's multiple
range test showed that the micronutrient effect was not significant. To
elucidate this micronutrient effect of fungicides on crop yield, further
studies need to be conducted. Bitertanol may have controlled other
pathogens not yet known, hence the higher yield.
Under the conditions these studies were conducted, the grower would
have been better off not fumigating the soil due to the high cost of
metam-sodium and low nematode populations. Fungicides, however, improved
yield and hence gross income. The net return from fungicide sprays
dpended on the frequency of spray. The optimum spray frequency was a
7-day schedule which indicated that spraying beans at intervals shorter
than seven days was not beneficial. The loss a grower would incur
depended on the price of snap beans and the cost of pesticides. Regres
sion analysis appeared to be the best method of predicting yield, hence
income, given various pest combinations. Regression analysis was able
to show which factor contributed more to the coefficient of determination.

CHAPTER VII
THE EFFECT OF INOCULATION METHOD AND INITIAL POPULATION DENSITY OF
MELOIDOGYNE INCOGNITA (KOIFOID AND WHITE) CHITWOOD ON SNAP BEANS
(PHASEOLUS VULGARIS L.) 'SPRITE'
Introduction
The root-knot nematode Meloidogyne incognita (Kofoid and White)
Chitwood poses a serious threat to bean (Phaseolus vulgaris L.) produc
tion in many bean growing areas of the world (Agudelo, 1980; Allen,
1983; Ngundo, 1977; Singh et al. 1981a; Sharma and Guazelli, 1982).
Meloidogyne incognita infections have been reported to decrease the
apparent photosynthetic rate of vulgaris 'Topnotch Golden Wax' as
well as other physiological growth factors (Melakberhan et al., 1983).
The limitation on bean production by root-knot nematodes may be due to
root galling which interferes with nitrogen fixation by Rhizobium spp.
and also interference with nutrient uptake. Yield losses of 50-90% have
been reported from fields infested with root-knot nematodes (Agudelo,
1980; Freire and Ferraz, 1977; Ngundo, 1977; Varn and Galvez, 1974).
Damage functions ascribable to nematodes are influenced by many
factors (McKenry, 1983). Nematode management decisions should, there
fore, be based on environmental factors and the crops grown (Ferris,
1980). Environmental factors such as soil temperature, texture, and
structure, and water infiltration rates influence moisture regimes of
soil profiles which in turn affect nematode damage functions (McKenry,
1983). Noe and Barker (1983) related 24 edaphic variables to the field
distribution of Meloidogyne spp. Work done on the influence of these
142

143
environmental factors on crop damage functions of root-knot nematodes
has shown that these factors significantly affect the establishment of
the nematodes and the growth of the crop and yield (Ferris, 1980;
McKenry, 1983; Noe and Barker, 1983; Roberts, 1983).
The derivation of mathematical models relating nematode densities
to crop damage has been discussed by Ferris (1980, 1984) and Seinhorst
(1965, 1972). In these models, the relationship between the initial
density of root-infesting nematodes and yield or other growth parameters
of infected plants are expressed the assumptions that (i) up to a
certain density the yield is not affected and (ii) a certain minimum
yield remains unaffected by the nematodes even at the highest densities
(Seinhorst, 1965).
The relationship between M. incognita and other Meloidogyne spp.
initial population densities and plant growth and/or yield has been
reported on tobacco (Barker et al., 1981, Ekanayake and Di Vito, 1984),
beans (Melakberhan et al, 1983), tomato (Barker et al., 1976) and pepper
(Di Vito et al., 1982). In these studies, inoculum consisted of eggs of
M. incognita or other Meloidogyne spp. eggs extracted by the sodium
hypochlorite method (Hussey and Barker, 1973) a factor which may be
critical to the development of damage functions in these studies. Vrain
(1977) evaluated the infectivity of three types of inocula which consis
ted of intact egg masses, eggs extracted with 0.53% NaOCl, and larvae
hatched from NaOCl-treated eggs. The data obtained showed the limita
tion of egg masses, low infectivity from NaOCl-extracted eggs and
sensitivity of larvae to relatively high temperatures.
Sodium hypochlorite dilutions have been used for the sterilization
of the surface of nematodes and their eggs in laboratory studies

144
(Briggs, 1946; Feder and Feldmesser, 1955). Sodium hypochlorite has
also been used for sterilizing processing substrates and equipment in
diagnostic nematology laboratory work (Esser, 1972), so its adverse
effects on nematodes at higher concentrations is well known.
The main objectives of this study were twofold (1) to determine the
influence of the initial M. incognita population density on yield of
beans and effects of inoculation method on the establishment of M.
incognita on beans, and (2) to determine the effect of sodium hypochlo
rite (NaOCl) concentration on the number of M. incognita eggs and
juveniles extracted from infected bean and tomato plants, and the
influence of NaOCl concentration on egg hatch.
Materials and Methods
Meloidogyne incognita (Kofoid and White) Chitwood, obtained from
Hausa potato (Coleus parviflorous Benth.), was maintained on greenhouse-
grown tomato (Lycopersicon esculentum Mill Floradade). Infected bean
roots were obtained from an earlier experiment conducted in a greenhouse
at the Tropical Research and Education Center in Homestead, Dade County,
Florida. Sodium hypochlorite (NaOCl) solutions (0.13, 0.26, 0.525, 1.3
and 2.6%) were made from Thrift King^ commercial bleach (5.25% NaOCl)
diluted serially with cold tap water (25C). Roots were cleaned of
soil, cut into 2-3 cm pieces, mixed, and 120 g of the mixture was used
for egg and juvenile extraction by the sodium hypochlorite method
(Hussey and Barker, 1973), except that a 230-mesh sieve was used instead
of the 200-mesh. Comparisons of egg and juvenile numbers extracted by
the various concentrations of NaOCl were made.

145
For the egg-hatch test, 1 ml of M. Incognita eggs and juveniles
suspended in tap water was put in a watch glass and 2 ml of tap water
was added to the suspension. The initial number of eggs and juveniles
was determined by counting them under a dissecting microscope. The
watch glasses were kept at room temperature (24-30C). Subsequent egg
hatch in each treatment were assessed every 2 days until hatching
levelled off (Vrain (1977).
The inoculation method studies were established on 31 August 1984,
in 1-quart side-drain black plastic pots filled with 1 L of soil (1 part
(R)
sand to 3 parts Palmetto Rich Earth' The soil was inoculated with
eggs and juveniles extracted with the 0.525% NaOCl solution. One series
of pots was inoculated by thoroughly mixing the inoculum with the soil.
The other series was inoculated by drenching the seeds with the
inoculum. A third series was inoculated by placing the appropriate
number of galls in the pot. Each gall contained an average of 246 eggs
and juveniles.
The egg and juvenile populations investigated were 0, 10, 100,
1,000, 10,000, and 100,000 per pot and the numbers of galls were 0, 1,
10, 100, and 500 galls per pot. Since each gall contained an average of
246 eggs and juveniles the gall inoculum was, therefore, equivalent to
0, 246, 2,460, 24,600 and 123,000 eggs and juveniles per pot. Three
seeds were planted in each pot and plants were thinned to one plant/pot
after germination. Treatments were replicated four times in a randomized
complete block. Pots were placed on corrugated benches 0.91 m high in
an open greenhouse and watered twice daily using an automatic time-
controlled water mist-forming system. Beans were harvested on 25

146
October 1984 and root gall indices determined following the method
outlined by Taylor and Sasser (1978).
Yield data were subjected to regression analysis using the general
linear models of SAS. Seinhorst model curve fitting was also attempted.
Results
The relationship between NaOCl concentration and log^ (number of
eggs and juveniles + 1) extracted is shown in figure 29. Data fit the
equations Y = a + b LnX or Y = a where Y = log (number of eggs and
juveniles + 1), X = concentration (%) of NaOCl, and a and b are con-
2
stants. The coefficient of determination (R ) values were 0.69* to
0.75* for the bean and tomato curves, respectively. Figure 29 shows
that the number of eggs and juveniles extracted increased more rapidly
at low NaOCl concentrations (0 to 0.26%) and levels off at high
concentrations (0.525-2.6%).
Table 40 shows the number of Meloidogyne incognita eggs and juve
niles extracted form 120 g of infected plant roots at various NaOCl
concentrations. These eggs and juveniles were extracted from bean and
tomato roots 63 and 50 days after inoculation respectively.
Figures 30 and 31 show the total number of juveniles which emerged
from eggs extracted at various NaOCl concentrations. On the day of
extraction, sodium hypochlorite-extracted eggs hatched more than the
water-extracted eggs. The highest number of juveniles had emerged from
eggs extracted with the 0.525% NaOCl solution from bean roots over the
next 6 days after extraction (Figure 30). Eggs extracted from tomato
roots had a different hatch trend from that of beans (Figure 31). The

147
0.525% solution gave the best overall hatch of nematode eggs extracted
from both bean and tomato roots (Figure 30 and 31).
The percentage hatch of M. incognita eggs is shown in figure 32.
The lowest percentage hatch was obtained from the 2.6% NaOCl-extracted
eggs. Water-extracted eggs had a percentage hatch comparable to that of
the 0.13, 0.26 and 0.525% NaOCl-extracted eggs. The 0.525% NaOCl-
extracted eggs had the highest proportion of eggs hatched among all
treatments by day 8, leading to the choice of the 0.525% NaOCl-extracted
eggs in the inoculation of bean plants and maximum yield of eggs over
time as above.
The relationship between initial M. incognita densities and snap
bean yield is shown in figure 33. There was a significant (P = 0.01)
negative correlation between nematode densities and yield for all three
inoculation systems (correlation coefficients were -0.96, -0.78 and
-0.96 for seed drench, soil mix and gall inoculation). As the nematode
densities increased, the snap bean yield decreased. The lowest yield
was obtained from plants inoculated with 500 galls/pot (= 123,000
eggs/pot). Negative impacts on yield were greater in seed-drench
inoculated pots than soil-mix inoculated ones (Figure 33). There were
significant differences in yield among nematode densities in the
seed-drench and gall inoculation methods with F values of 14.34** and
26.18**, respectively. There were, however, no significant differences
in yield among nematode densities in the soil-mix inoculation method
with an F value of 3.09.
Gall indices were comparable in all three inoculation methods as
shown in Table 41. Among inoculated plants, the lowest gall index was
observed on plants inoculated with 10 eggs and juveniles per pot.

148
TABLE 40. Total number of eggs and juveniles extracted from 120 g of bean
and tomato roots. Data are means of 3 replicates.
NaOCl Concentration
(%)
Inoculum Source
Bean
Tomato
0
135,000
320,000
0.13
290,000
704,000
0.26
517,000
1,566,000
0.525
642,000
2,120,000
1.3
864,000
2,886,000
1.6
1,062,000
3,228,000

Log10(No. eggs and juveniles extracted)
149
y=5.8O0.74 In x
y=5.80x
0.12
y=5.38+0.65 In x
y=5.38x0'12
Concentration (%) of
Sodium hypochlorite
Log^CNo. eggs and juveniles extracted) at various
NaOCl concentrations.
Figure 29.

Number of juveniles
150
Figure 30
No. juveniles hatched from eggs obtained from bean plants.

Number of juveniles
151
Figure 31. No. juveniles hatched from eggs obtained from tomato plants.

152
Figure 32. Egg hatch (%) at various NaOCl concentrations.

153
TABLE 41. Yield and root gall indices of bean plants treated with low
to high initial densities of Meloidogyne incognita eggs and
juveniles. Data are means of 4 replicates.
Mean initial number of
eggs and juveniles/lL soil
Yield
(g/plot)
Gall indices
(means)
Untreated
check
19
0
10
(Seed drench)
18
2
100
fl II
14
4
1,000
II II
13
5
10,000
II II
11
5
100,000
II II
4
5
10
(Soil mix)
12
3
100
II II
15
4
1,000
II II
15
4
10,000
II II
8
5
100,000
II II
9
5
246
(1 gall)
13
5
2,460
(10 galls)
4
5
24,600
(100 galls)
1
5
123,000
(500 galls)
1
5
Gall index data based on the scale
of Taylor and
Sasser
(1978) as
follows:
no galls or egg masses =
0; 1-2 galls i
or egg masses = 1;
3-10 = 2
; 11-30 = 3; 31-100 = 4; more than 100 =
5.

154
Log10(lnitial nematode density+1)
Relation between log^g (initial nematode density +1)
and yield.
Figure 33.

155
Plants inoculated with one gall/pot had a similar gall index to that of
the plants inoculated with 100 or 1,000 eggs and juveniles/pot as
anticipated. Ten galls per pot gave gall indices similar to these of
10,000 or 100,000 eggs and juveniles/pot. Plants inoculated with more
than 10 galls gave the maximum gall index. The controls had no root
galls.
Data obtained in this study did not fit the Seinhorst model
(Seinhorst, 1965, 1972) which is of the form Y = M + (l-M)Z^ ^ where Y
= ratio between the yield at nematode population P and at P < T, M =
relative minimum yield, P = initial nematode population density, T =
tolerance limit for the nematode density, and Z = a constant. Coeffi
cients of determination were 0.031, 0.003, and 0.28 for the soil mix,
seed drench, and gall inoculation systems. Linear regression analysis
of yield data is shown in Figure 33. The linear model produced signifi
cant coefficients of determination at the 0.01 probability level.
Discussion
The largest number of M. incognita eggs and juveniles was extracted
with the 2.6% NaOCl solution, due to the better dissolution of the
gelatinous matrix enclosing the nematode eggs at the highest NaOCl
concentration tested. The rate of increase of eggs and juveniles
extracted, however, decreased dramatically above the 0.525% concentra
tion probably because the number of egg masses or galls per sample was
the same. Further the increase in NaOCl did not improve egg and juve
nile extraction that much.

156
There were more eggs and juveniles extracted from tomato than from
bean roots (Table 40). This may be an indication that tomato is a more
comparitible root-knot nematode host than beans. Consequently the
nematodes may have reproduced at a faster rate on tomato than on beans.
It is possible that this difference in extracted eggs and juveniles may
be a reflection of age differences. Eggs and juveniles were extracted
63 days after inoculation from bean roots whereas eggs and juveniles
were extracted from tomato plants 50 days after inoculation. Thus,
there might have been more juveniles which had already emerged from bean
root galls than on tomato root galls. Consequently, more juveniles may
have been washed off along with soil from bean roots which may have lead
to the low numbers of eggs and juveniles extracted. Moreover, weighing
did not imply the same number of galls or eggs on the root material
used. Weighing may thus have biased the number of egg masses used
towards the tomato source. There is little information in the
literature pertaining to variation in the number of eggs per gall in
various hosts. This suggests that variation in egg numbers per gall on
various hosts be investigated.
More juveniles of M. incognita may have emerged on the day of
extraction from NaOCl-treated eggs than from water-treated eggs, probab
ly because the NaOCl reacted with the egg shell, thereby inducing early
hatch in eggs with fully developed juveniles. This effect of NaOCl on
egg hatch had, however, some negative effects when the concentration was
above 0.525%. The negative effect of NaOCl on eggs was particularly
evident with the 2.6% NaOCl treated eggs. This high concentration of
NaOCl gave rise to numbers of juveniles comparable to those obtained
from water-treated eggs, despite the 2.6% solution having extracted the

157
highest number of eggs from either crop. The 0.525% solution had the
highest number of juveniles that emerged from eggs obtained from either
beans or tomato. Thus, a 0.525% NaOCl solution may be the optimum
concentration for M. incognita egg extraction for inoculation studies.
The low emergence of juveniles from water-treated eggs may imply that
under natural conditions root-knot nematode eggs hatch over a long
period. This may be a survival mechanism for this nematode.
There were no significant differences in the proportion of eggs
that had hatched between water and NaOCl solutions of 0.13, 0.26 and
0.525%. This was because there were far fewer eggs extracted with water
than with the NaOCl treatments. The lowest percentage hatch was
obtained from eggs extracted with 2.6% NaOCl. This low hatch was
probably due to the adverse effects of high NaOCl concentration on eggs
which may have arrested the development of embryos of juveniles.
Gall inoculations had a more marked effect on yield and gall
indices (as evidenced by greater slope) than extracted eggs probably
because eggs in intact egg masses hatched in their natural environment
and there was no loss from mortality due to NaOCl. The effect of egg
extraction may have been exacerbated by the inoculation of soil before
seeds germinated. Thus, juveniles may have been rendered less infective
before seeds germinated. Root-knot nematodes are more infective at the
second juvenile than any other stage (Dropkin, 1980). Seeds germinated
5 days after planting and nematode inoculation. A good proportion of
eggs may have hatched well before the plants produced roots. Nematode
juveniles, especially root-knot nematodes, are sensitive to relatively
high edaphic temperatures (McKenry, 1983). Root gall indices were

158
comparable in all three inoculation methods. The extent of galling was,
however, not reflected in yield in the soil-mix inoculated plants.
There were significant differences in snap bean yield among
nematode egg and juvenile densities in the seed-drench method (F =
14.34, P 0.01). This may be due to some juveniles ecloding from the
eggs in the vicinity of the germinating seeds since the nematode eggs
hatched over a period of up to 14 days. Hence many of them may have
been able to penetrate bean radicles soon after hatching. There were no
significant differences in snap bean yield (F = 3.09, P ^ 0.1) among
nematode egg and juvenile levels in the soil-mix inoculation method
probably because some of the eggs and juveniles leached out with irriga
tion water. Moreover, juveniles may not have been able to swim up to
the vicinity of germinating seeds against the downward flow of water.
The seed drench method apparently concentrated nematodes, more, within
the vicinity of the plant root system so the local population density
was higher than if much of the inoculum was scattered through the entire
volume of soil. Gall-inoculated plants produced the lowest yield and
differences in yield among gall treatments were significant at P = 0.01
(F = 26.18). This may be due to the natural environment in the galls
within which juveniles ecloded over a long period.
There was lack of association between yields obtained from
2
seed-drench and soil-mix inoculated plants (X = 7.74, 0.10 < P < 0.20).
This lack of association does not necessarily prove that the yields were
different from each other but there was a chance that they were
different.
The data did not fit the Seinhorst model probably because the
nematode population levels were in a logarithmic progression. In most

159
of the work in which this model was able to explain the relationship
between yield and nematode population densities, the intervals between
population levels were smaller particularly at low densities. Thus,
where nematode population levels are far apart, the Seinhorst model may
not be the best for explaining the relationship between plant growth
parameters and nematode population levels.
Results presented here indicate that M. incognita may cause yield
loss in snap beans regardless of the method of inoculation. If the soil
is mixed with 0.525 NaOCl-extracted eggs the influence on yield may be
less profound than if the plants were inoculated with galls or via seed
drench. It has, also, been shown that NaOCl concentrations have a
direct positive relationship to the numbers of nematode eggs and juve
niles extracted. Indicated in the results is the fact the NaOCl has an
egg-hatch inducing effect, compared to eggs allowed to stand in tap
water at room temperature, which is less pronounced as the concentration
increases above 0.525%.

CHAPTER VIII
SUMMARY AND CONCLUSIONS
Pest damage and disease infection on beans rarely result in total
leaf abscision under field conditions. In the experiments conducted in
this study, various defoliation levels up to total defoliation were
included. Bean rust was manipulated by fungicide sprays at various
frequencies and nematode infestation was studied under field as well as
greenhouse conditions. These factors were studied individually and in
combination.
Bean plants were most sensitive to defoliation at full-bloom and
pod-set growth stages. Twenty five percent leaf removal at the primary
leaf and first trifoliate leaf stages resulted in yield losses of up to
36% and 29% in the greenhouse and field respectively. In the greenhouse
50%, 75%, and 100% defoliation resulted in up to 34%, 39%, and 74% yield
loss, respectively. In the field 50%, 75%, and 100% defoliation caused
yield losses of up to 68%, 70%, and 95% respectively depending on the
plant growth stage. Yield loss fluctuated in such a way that in some
cases lower defoliation levels resulted in higher yield losses than
higher defoliation levels. This fluctuation may have been due to
various factors, one of which could be better exposure of photosynthe-
tically active foliage to light.
Root-knot nematodes, Meloidogyne incognita, caused yield loss in
snap beans when plants were inoculated with eggs and juveniles. Ten
eggs and juveniles per pot resulted in 19% yield loss and 100, 1,000,
10,000, and 100,000 eggs and juveniles resulted in 38%, 45%, 47%, and
160

161
59% yield loss. Thus, the threshold level of M. incognita was between 0
and 10 eggs and juveniles per pot. Combining defoliation and root-knot
nematodes showed that there was no significant interaction between them.
When defoliation was held constant, yield reduction rate was greater
than when root-knot nematode populations were held constant. Equations
were devised to express this relationship between the two factors.
Bean rust caused yield loss in snap beans, which was substantial at
the highest disease severity (maximum proportion of foliage infected).
In the first trial, disease severity levels of 0.098, 0.46, 0.65, and
0.76 caused yield losses of 21%, 17%, 35%, and 60% respectively.
Disease severity levels of 0.4, 0.47, 0.71, and 0.86 resulted in yield
losses of 55%, 56%, 80%, and 91% in trial 2. Thus, the disease severity
level which could be tolerated was below 0.1. When bean rust was used
simultaneously with defoliation and nematodes, it was demonstrated by
regression analysis of the data that the disease had an overriding
effect on yield loss. The disease may have had this overriding effect
due to its continual adverse effects on the plants from its onset to
bean harvest. In these experiments, nematodes and defoliation had
little effect on yield. There was no significant interaction among
nematodes, defoliation and bean rust. As expected the best yield was
obtained from plants free of all stress.
From these results, several general conclusions can be drawn. Some
defoliation may result in exposure of photosynthetically active foliage
to light and also may stimulate rapid growth of new leaves which are
highly photosynthetically active. Yield reduction due to defoliation
depends on the growth stage at which leaf removal occurs. This may be
due to the differences in partitioning of photosynthates at these
various growth stages.

162
The nematode, M. incognita, caused substantial yield loss when
plants were inoculated in pots, but hardly had any effect on yield under
field conditions. The lack of apparent effect on yield by nematodes may
be due to interaction of several biotic as well as abiotic factors in
field soils. Moreover, the nematode populations in the field were very
low, likely were below threshold levels. In the greenhouse, plants were
grown in pots in which nematodes were probably concentrated on the upper
soil layer thus making the nematode density higher. This discrepancy
between nematode effects on yield in the greenhouse and field may be an
indication that threshold levels determined in the greenhouse may not
necessarily apply under field conditions.
Bean rust caused apparent yield loss even at the lowest maximum
severity level measured. This may be due to the continual adverse
effects of the disease on the physiology and other bean plant growth
factors. A low disease severity, however, required the use of mancozeb
+ sulfur more often hence increasing production costs. Increasing
production costs in turn reduced net returns from the crop. The improve
ment in yield by mancozeb + sulfur may also have been due to increased
supply of micronutrients, such as zinc, which stimulated plant growth.
Thus, yield increases in beans may not necessarily be due to bean rust
control/prevention only.
There may not necessarily be a close relationship between experi
mental results obtained in the greenhouse and the field. Thus, pest
control recommendations should not be based only on greenhouse/laboratory
experimental results. Recommendations for control of a pest complex
should be based on the population dynamics of the key pests supplemented
by the presence of other factors. Pest population levels can be obtained
by close and routine population monitoring throughout the growing season.

163
Yield was generally low in these experiments, probably because of
insect pest pressure, especially cowpea curculio and other leaf-feeding
insects. In the defoliation, metam-sodium, and bean rust disease
experiment the low yields may have been due to the late planting thus
going into a season where rainfall was erratic and insect pests were
more abundant. Generally, the weather conditions were not optimum for
snap bean production during the period some of the field experiments
were conducted.
A multipest assessment on beans in the field was not easy because
it was time consuming and tedious. Thus, a grower/extension worker
would have to assess pests separately and act on them accordingly. This
has been shown to be a valid approach in these experiments by the
general lack of interaction among defoliation, nematodes or nematicide,
and bean rust disease.

APPENDIX
EFFECTS OF DEFOLIATION AND FUNGICIDES ON SNAP BEANS
Introduction
Bean rust, caused by Uromyces phaseoli (Pers.) Wint., is one of the
most prevalent diseases on beans (Phaseolus vulgaris L.) grown in
Florida and other bean producing areas of the world (Agudelo, 1980;
Allen, 1983; Cook, 1978; Kidney, 1980). Bean rust is common in warm,
(15-27C) moist weather (Augustin et al., 1972; Gonzalez, 1976;
Shurtleff, 1966). Uromyces phaseoli is an autoecious/monoecious fungus
which infects mainly plants in the legume family (Cook, 1978; Shurtleff,
1966). Symptoms of the disease are confined largely to leaves, but
young stems and branches are occasionally infected. On severe infection
symptoms or signs may appear on pods (Cook, 1978; Vargas, 1980).
Lesions first appear as small slightly raised spots that are light in
color and more often on the lower leaf surfaces. These pustules gra
dually enlarge to 1-2 mm in size and may rupture the leaf epidermis 8
days after infection (Allen, 1983). A ring of secondary sori usually
develops around the original infection locus on susceptible varieties.
Bean rust is commonly controlled by weekly sprays of mancozeb
tank-mixed with sulfur or other of the ethylene bis-dithiocarbamate
fungicides (Pohronezny et al., 1984). A few experimental fungicides are
available for bean rust control. One of these more potent fungicides
for bean rust control is bitertanol. Thus, this study was conducted to
164

165
investigate the effect of manual defoliation and bitertanol and mancozeb
sprays at various frequencies on bean rust and snap bean yield, the
original objective of which had been to determine the effects of bean
rust disease and defoliation on yield. However, due to lack of disease
development, the test was limited to evaluation of the direct effects of
the fungicides and defoliation on snap bean yield.
Materials and Methods
00
Soil was fumigated with Dowfume MC2V at a rate of 314 kg/ha.
Two-gallon side drain plastic pots were filled with 6.4 L of soil and
placed in the field. A plot consisted of 2 pots each with 3 plants.
Plots were 1.5 m apart. Treatments were replicated 4 times in a ran
domized complete block. Treatments were arranged in a 5x5x2 factorial
design. Fertilizer (8:16:16) was applied at 448 kg/ha (to pots and
incorporated) and plants were top dressed at 224 kg/ha just before
(R)
flowering. Trigardv (150g/ha) was applied for leafminer control.
Mesurol was applied for slug and snail control as needed. Other
leaf-eating insects were controlled with sprays of permethrin or
endosulfan.
Beans (Phaseolus vulgaris L., 'Sprite') were planted on 25 January
1985 and harvested on 25 April 1985. Irrigation was provided by an
overhead sprinkler system twice a week. Plants were defoliated at two
growth stages, the primary leaf stage and flower bud formation. Defoli
ation levels investigated were 25%, 50%, 75%, and 100% plus an undefo
liated control. The foliage was removed from the distal end of the
petiole using a pair of scissors.

166
Two fungicides were tested for the manipulation of bean rust.
These were; bitertanol (57 g ai/ha) applied at 7-day intervals, and
mancozeb (1.7 kg/ha) tank-mixed with sulfur (4.2 kg ai/ha) applied at
4-5, 7, and 14-day intervals. An unsprayed control was also included.
Inoculum exposure was done at the primary leaf stage by clipping
infected pole bean leaves on a string tied across experimental plots 1.5
m above the ground. Fungicide sprays were initiated soon after inocula
tion. Disease progress was assessed by the Horsfall-Barratt rating
system.
Data were subject to analysis of variance.
Results
Analysis of variance on the effect of fungicides and defoliation on
snap bean yield showed that there were significant differences among
fungicides and manual defoliation levels (F = 18.32** and 29.34** (P <_
0.01) respectively) (Table 42-1). There were, however, no significant
interactions between fungicides and manual defoliation levels (Table
42-1). There were no significant differences between defoliation times
(F = 1.39 N.S.) and interaction among manual defoliation levels and time
(F = 1.09 N.S.). Duncan's multiple range test showed that there were no
significant differences in yield between the unsprayed plants and those
sprayed with mancozeb and sulfur at 7 and 14-day intervals (Table 42-2).
There were, however, significant differences in yield between mancozeb
at all schedules and bitertanol (Table 42-2).
The highest snap bean yield was obtained from plots sprayed weekly
with bitertanol at all defoliation levels and both times of defoliation

167
TABLE 42-1. F-values and probabilities from analysis of variance on the
effects of defoliation, fungicides and their interaction on
snap bean yield.
Source
F
Probability F
Defoliation
29.34
0.0001
Time
1.39
0.24
Defoliation x
Time
1.09
0.36
Fungicide
18.32
0.0001
Defoliation x
Fungicide
0.74
0.75

168
TABLE 42-2. Mean snap yield per hectare sprayed with various fungicides.
Fungicide
Spray frequency
(days)
Mean yield (g/plot)
No fungicide
0
114 cd
Mancozeb + sulfur
14
98 d
Mancozeb + sulfur
7
119 c
Mancozeb + sulfur
4-5
137 b
Bitertanol
7
166 a
Means with the same letter are not significantly different at P<^ 0.05
(Duncan's multiple range test).

169
TABLE 43. Mean snap bean yield from plants defoliated at various levels
and various fungicide spray frequencies.
Defoliation
levels (%)
Plant
Fungicide Primary leaf
Growth
Flower
Stage
bud formation
0
No fungicide
143
152
0
Bitertanol (7-day)a
212
232
0
Mancozeb (4-5-day)
185
157
0
Mancozeb (7-day)
144
155
0
Mancozeb (14-day)
126
131
25
No fungicide
112
126
25
Bitertanol (7-day)
181
172
25
Mancozeb (4-5-day)
162
152
25
Mancozeb (7-day)
147
149
25
Mancozeb (14-day)
116
73
50
No fungicide
99
95
50
Bitertanol (7-day)
180
168
50
Mancozeb (4-5-day)
154
153
50
Mancozeb (14-day)
106
128
75
No fungicide
104
98
75
Bitertanol (7-day)
135
134
75
Mancozeb (4-5-day)
121
100
75
Mancozeb (7-day)
108
106
75
Mancozeb (14-day)
109
59
100
No fungicide
100
74
100
Bitertanol (7-day)
93
106
100
Mancozeb (4-5-day)
81
113
100
Mancozeb (7-day)
60
77
100
Mancozeb (14-day)
76
56
a
Figures in parentheses are spray intervals.

170
TABLE 44. Effect of fungicides
and defoliation
on gross
dollar values
per
hectare of snap
beans.
Plant
Growth Stage
of
defoliation
Defoliation
Price
Primary
Flower bud
level (%)
Fungicide
range
leaf
formation
low
1122
1193
0
No fungicide
medium
2054
2186
high
3740
3975
low
1663
1820
0
Bitertanol
medium
3049
3337
(7-day)
high
5544
6067
low
451
1232
0
Mancozeb
medium
2661
2258
(4-5-day)
high
4838
4106
low
1130
1216
0
Mancozeb
medium
2071
2230
(7-day)
high
3766
4054
low
989
1028
0
Mancozeb
medium
1812
1884
(14 days)
high
3295
3426
low
879
973
25
No fungicide
medium
1611
1784
high
2929
3243
low
1420
1349
25
Bitertanol
medium
2604
2474
(7-day)
high
4734
4498
low
1271
1193
25
Mancozeb
medium
2330
2186
(4-5-day)
high
4237
3975
low
1153
1169
25
Mancozeb
medium
2114
2143
(7-day)
high
3844
3897
low
1028
973
25
Mancozeb
medium
1884
1784
high
3426
3243
low
777
745
50
No fungicide
medium
1424
1366
high
2589
2485

171
TABLE 44. Continued
Defoliation
level (%)
Fungicide
Price
range
Plant
of
Primary
leaf
Growth Stage
defoliation
Flower bud
formation
low
1412
1318
50
Bitertanol
medium
2589
2417
(7-day)
high
4707
4393
low
1208
1200
50
Mancozeb
medium
2215
2201
(4-5-day)
high
4028
4001
low
842
1004
50
Mancozeb
medium
1544
1841
(7-day)
high
2807
3348
low
910
573
50
Mancozeb
medium
1669
1050
(14-day)
high
3034
1909
low
816
769
75
No Fungicide
medium
1496
1410
high
2720
2563
low
1059
1051
75
Bitertanol
medium
1942
1927
(7-day)
high
3531
3504
low
949
785
75
Mancozeb
medium
1740
1438
(4-5-day)
high
3164
2615
low
847
832
75
Mancozeb
medium
1568
849
(7-day)
high
2824
2772
low
855
463
75
Mancozeb
medium
1568
849
(14-day)
high
2851
1543
low
785
581
100
No Fungicide
medium
1438
1064
high
2615
1935
low
730
832
100
Bitertanol
medium
1338
1525
(7-day)
high
2432
2772

172
TABLE 44. Continued
Defoliation
level (%)
Fungicide
Price
range
Plant
of
Primary
leaf
Growth Stage
defoliation
Flower bud
formation
low
636
887
100
Mancozeb
medium
1165
1625
(4-5-day)
high
2118
2955
low
471
604
100
Mancozeb
medium
863
1108
(7-day)
high
1569
2014
low
596
439
100
Mancozeb
medium
1093
806
(14-day)
high
1988
1465

173
TABLE 45. Net income (dollars) per hectare of snap beans defoliated at
the primary leaf and flower bud formation stages and sprayed
with various fungicides
Plant growth
Plants were
Defoliation
level(%)
stage
defoliated
Fungicide
Primary leaf
Flower bud formation
0
No fungicide
0
0
0
Bitertanol (7-day)
1805
2092
0
Mancozeb (4-5-day)
929
- 39
0
Mancozeb (7-day)
- 87
- 34
0
Mancozeb (14-day)
- 501
- 662
25
No fungicide
- 811
- 732
25
Bitertanol (7-day)
694
523
25
Mancozeb (4-5-day)
328
- 169
25
Mancozeb (7-day)
8
191
25
Mancozeb (14-day)
- 370
- 789
50
No fungicide
-1151
-1491
50
Bitertanol (7-day)
968
418
50
Mancozeb (4-5-day)
118
- 143
50
Mancozeb (7-day)
-1046
- 741
50
Mancozeb (14-day)
- 763
-2123
75
No fungicide
-1020
-1412
75
Bitertanol (7-day)
- 209
- 471
75
Mancozeb (4-5-day)
- 745
-1529
75
Mancozeb (7-day)
-1028
-1316
75
Mancozeb (14-day)
- 946
-2489
100
No fungicide
-1125
-2040
100
Bitertanol (7-day)
-1308
-1203
100
Mancozeb (4-5-day)
-1791
-1189
100
Mancozeb (7-day)
-2284
-2074
100
Mancozeb (14-day)
-1809
-2567

174
except for total defoliation (Table 43). The unsprayed plots gave
yields comparable to those obtained from fortnightly mancozeb-sprayed
plants. There was no apparent difference between the 4-5-day and 7-day
mancozeb spray schedules (Table 43).
Gross dollar values per hectare of snap beans are shown in Table
44. Bitertanol-sprayed plots consistently gave the highest dollar
values at all defoliation levels and both growth stages. The gross
dollar values were computed from the following price range, $6.00 (low),
$11.00 (medium), and $20.00 (high) per bushel (13.62 kg/) of snap beans
multiplied by the yield per hectare. Net income (Table 45) was derived
from the no spray value as the base line. This was deducted from the
values of the other treatments and the cost of fungicide sprays deducted
from this difference. Bitertanol is an experimental fungicide which has
not been registered for use on beans, hence the price is not known.
Thus, the net value of beans for this fungicide excludes its cost.
Consequently, the net values for bitertanol may not be a true
reflection. Under the conditions this study was conducted, the grower
would have made a profit if he sprayed his crop with mancozeb at 4-5 day
intervals even if 35% or 50% of the leaf area were removed at the
primary leaf stage (Table 45). There was no positive effect on net
income from fungicide sprays if plants were defoliated at the flower bud
formation stage.
Discussion
There were significant differences among fungicide spray schedules
based on yield. Bitertanol gave the highest yield and by derivation the

175
largest dollar values. Four to five-day sprays of mancozeb were
generally better than 7-day spray schedules on yield. Bitertanol gave
the highest yield and highest dollar values. Four to five-day sprays of
mancozeb and sulfur were generally better than 7-day sprays. This may
have been due to the presence of zinc, an element required for plant
growth. May be the more frequent sprays provided more of this element
than the 7-day spray schedule of mancozeb and sulfur rather than the
efficacy of the fungicide since there was virtually no apparent disease
build up on the plants. It is possible that the more frequent sprays
controlled other diseases resulting in higher yield. This same fact may
be true for the fortnightly spray schedule which generally gave the
lowest yield among the fungicide sprays.
The net income value picture is not clear with regard to bitertanol
since its market price was not known. Moreover, the net income depends
on snap bean prices which fluctuate widely. Furthermore, these values
do not include other production costs such as farm machinery, labor and
interest on loans. There was no apparent benefit from the shorter (4-5
day) spray schedule. The extra sprays did not produce enough increase
in yield to justify added cost. From these results the ideal mancozeb
spray schedule was the 7-day interval. Biteranol increased snap bean
yield substantially probably because this chemical controlled some other
subtle disease and it may have plant growth regulating properties. This
has been suggested by teh non-establishment of bean rust disease in this
study. Bitertanol has no known micronutrients which ruled out the
micronutrient effect.

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de Fitopalogia. ICA, Palmira, Colombia. 10 pp.
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BIOGRAPHICAL SCKETCH
Afete Divelias Gadabu was born on 19 March 1949, in Lilongwe,
Malawi. He attended primary school at Kamzimbi Primary School and went
to Bwaila Secondary School where he got his Junior Certificate of
education in 1968. He went to Blantyre Secondary School in 1969 and
obtained a Cambridge School Certificate in 1970. In 1971, he received a
scholarship to the University of Malawi from which he graduated with a
Bachelor of Science in biology and chemistry in 1975.
In the same year he joined the Civil Service as a research officer
in the Department of Agricultural Research of the Ministry of Agricul
ture. In 1976 he received a scholarship to the University of Newcastle
upon Tyne, U.K. He graduated with a Master of Science in applied
entomology in 1978 and returned to Malawi. He worked for the Department
of Agricultural Research as a research entomologist. In 1982, he
received a scholarship to the University of Florida where he has been a
graduate student in the Department of Entomology and Nematology from
1983 to 1986. Upon fulfilling the requirements for the degree of Doctor
of Philosophy, he is returning to Malawi to continue working as a
research entomologist.
194

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
V. H. Waddill, Chairman
Professor of Entomology
and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
J. R. Strayer, Co-Chairman
Professor of Entomology
and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
R. T. McSorley
Associate Professor of
Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
K. L. Pohronezny
Associate Professor of Plant
Pathology

This dissertation was submitted to the Graduate Faculty of the College
of Agriculture and to the Graduate School and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
May 1986
t. £
Dean( /College of Agriculture
&
Dean, Graduate School



97
at the same growth stage. This discrepancy in correlation of rate of
disease progress and yield in these trials may be due to the early onset
of the disease in trial 2 where the rate of disease progress may have
started to decrease since most of the foliage had already been infected.
There is also a possibility that rates of disease progress were sporadic
in trial 2. Although AUDC was an equally good predictor of yield at
full bloom and pod set in trial 1 and at pod set through fully formed
pods in trial 2, it was a difficult parameter to use since it required
the use of computers. Computers are not readily accessible to extension
specialists who work closely with local farmers. Disease severity, on
the other hand, can be estimated by the use of graph papers or tracing
paper for the determination of leaf area before and after cutting out
diseased leaf tissue. Thus, disease severity would be a more convenient
parameter for the description of the relationship between bean rust and
snap bean yield. Moreover, AUDC is derived from disease severity. Area
under the disease progress curve is, however, a better representation of
the magnitude of the disease for the growing season. Disease severity
is, however, easier to measure and does not require complicated calcula
tions. Therefore, disease severity may be useful for the prediction of
yield at specific bean plant growth stages.
The Horsfall-Barratt disease rating system was not used in these
trials because of its subjectivity. This system has a lot of variabi
lity according to the person assessing the disease and would require the
comparison of several people's data to be relied on. The Horsfall-Barratt
method could, however, be adapted based on standardized diagrams put out
by extension.
At disease severity levels (maximum proportion of foliage infected)
below 0.5 (50%), it was not worth the expense of the shorter 4-5-day


153
TABLE 41. Yield and root gall indices of bean plants treated with low
to high initial densities of Meloidogyne incognita eggs and
juveniles. Data are means of 4 replicates.
Mean initial number of
eggs and juveniles/lL soil
Yield
(g/plot)
Gall indices
(means)
Untreated
check
19
0
10
(Seed drench)
18
2
100
fl II
14
4
1,000
II II
13
5
10,000
II II
11
5
100,000
II II
4
5
10
(Soil mix)
12
3
100
II II
15
4
1,000
II II
15
4
10,000
II II
8
5
100,000
II II
9
5
246
(1 gall)
13
5
2,460
(10 galls)
4
5
24,600
(100 galls)
1
5
123,000
(500 galls)
1
5
Gall index data based on the scale
of Taylor and
Sasser
(1978) as
follows:
no galls or egg masses =
0; 1-2 galls i
or egg masses = 1;
3-10 = 2
; 11-30 = 3; 31-100 = 4; more than 100 =
5.


Proportion of Loss in Yield
134
Figure 25.
The relationship between area under disease progress curve
and yield loss of snap beans.


19
Identification and Etiology of the Pathogen
Bean rust is caused by the fungus Uromyces phaseoli [Pers.] Wint.
(= U. phaseoli typica (Reben) Wint. = U. appendiculatus [Pers.] Unger).
The fungus was first described in Germany in 1795 (Cook, 1978). The
pathogen is an autoecious macrocyclic rust fungus (Kolmer et al., 1984;
Laundon and Waterston, 1965). This pathogen is parasitic on the legumi
nous genera Dolichos, Phaseolus, and Vigna (Laundon and Waterston,
1965). The fungus is transmitted generally through wind-borne uredo-
spores. Uredospores are rusty orange in color and ellipsoidal to
obvoidal in shape, 20-30 x 20-26 yum in measurement (Laundon and Waters
ton, 1965).
The aecial and pycnial stages are rare in U. phaseoli (Harter et
al., 1935). Harter et al. (1935) did not observe any aecia or pycnia of
this fungus in the field. In Queensland, Ogle and Johnson (1974) did
not report seeing mature aecia or pycnia of IJ. phaseoli. The absence of
aecia on U. phaseoli under field conditions has also been reported in
Maryland (Marcus, 1952). The aecial stage of this fungus has, however,
been observed and reported in New York and Virginia (Fromme and Wingard,
1921; Jones, 1960). Both aecia and pycnia were reported to occur on
field grown beans in North Dakota by Venette et al. (1978).
Fromme and Wingard (1918) and Harter et al. (1935) reported that
telia form under unfavorable conditions for the development of the
pathogen such as low temperatures, decreased host vigor, and increased
host resistance. The propensity of the pathogen to form telia was
suspected to be an innate character of the fungal isolate (Harter et
al., 1935). It has been reported that teliospores do not occur in
Florida (Townsend, 1939). Consequently, Townsend (1939) suggested


TABLE 37. Nematode genera + disease severity values found in plots fumigated with metam-sodium. Figures are
means of 3 replicates (Soil samples at harvest).
Disease severity
Defoliation level (maximum
(proportion
of foliage)
Metam-sodium
(Liters/ha)
proportion of
foliage infected)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Tylenchorhynchui
(No./lOO ml)
0
0
0.87
0
13
85
10
0
935
0.83
0
2
77
41
0.25
0
0.64
0
13
93
13
0.25
935
0.71
0
8
32
10
0.5
0
0.77
0
5
134
8
0.5
935
0.74
2
2
47
3
0
0
0.02
2
28
66
23
0
935
0
0
15
77
2
0.25
0
0
2
7
125
15
0.25
935
0.01
2
23
108
25
0.5
0
0
0
2
78
7
0.5
935
0
2
2
83
18
0
0
0.56
2
18
87
7
0
935
0.59
0
13
77
7
0.25
0
0.7
0
5
98
3
0.25
935
0.66
0
17
67
18
0.5
0
0.68
2
5
103
15
0.5
935
0.71
0
8
118
3
0
0
0.71
0
23
135
20
0
935
0.74
0
7
45
5
0.25
0
0.75
2
13
120
3
0.25
935
0.83
2
7
45
12
0.5
0
0.72
0
8
100
17
0.5
935
0.81
0
20
129
28
127


55
This study was conducted to investigate the relationship between M.
incognita population levels, manual defoliation, and their interaction
to snap bean yield.
Materials and Methods
Two studies were conducted in a greenhouse at the Tropical Research
and Education Center in Homestead, Dade County, Florida, in the summer
and fall 1984. One experiment was an inoculation experiment with M.
incognita designed to determine the effect of this pest alone on yield,
and the other experiment examined the simultaneous effect of M.
incognita inoculation and manual defoliation. The first experiment (M.
incognita alone) was designed as a randomized complete-block replicated
four times, involving six different nematode population levels. The
other experiment was a 5x4x6 factorial replicated four times and
included the following treatments: 5 defoliation levels of 0, 25, 50,
75, and 100%; 4 nematode population levels of 0, 1,000, 10,000, and
100,000 eggs and juveniles per pot; and defoliation at the following 6
plant growth stages: primary leaf, first trifoliate leaf, third
trifoliate leaf, flower bud formation, full bloom, and pod set.
Preparations for both experiments were made by covering Rockdale
soil (3030 L) with a polyethylene sheet and fumigating it with
(R)
Dowfume MC2 (681g). Two -gallon, side-drain, plastic pots were
filled with 6.4 L soil and placed on corrugated greenhouse benches 0.91
m high. Fertilizer (8:16:16) was applied before planting at 3 g/pot
(448 kg/ha). Plants were top-dressed with the same fertilizer at 1.5
g/pot four weeks after germination. The M. incognita inoculation test


Proportion of Foliage Infected
128
Days After Inoculation
Figure 22. Disease progress curve for Uromyces phaseoli on snap
beans not sprayed with mancozeb, defoliated at various
levels and soil fumigated with metam-sodium.
1 = no defoliation, no metam-sodium and no fungicide, 2 = no defoliation,
935 L/ha metam-sodium, and no fungicide, 3 = 25% defoliation, no metam-
sodium and no fungicide, 4 = 25% defoliation, 935 L/ha metam-sodium and
no fungicide, 5 = 50% defoliation, no metam-sodium and no fungicide, 6 =
50% defoliation, no metam-sodium and no fungicide.


26
in cowpea (Thomason et al., 1959). Interaction of root-knot nematodes
is not limited to nematode-fungus or nematode bacterium complexes.
Meloidogyne incognita has also been reported to interact with other
nematodes (Thomas and Clark, 1983). Thus, Thomas and Clark (1983)
reported that early season counts of M. incognita and Rotylenchulus
reniformis Linford and Oliveira were positively correlated with later
counts of the same nematode, but negative correlations were found
between early M. incognita and subsequent R_. reniformis, and between
early R. reniformis and subsequent M. incognita counts. The authors
suggested that a competitive interaction existed with each species
capable of inhibiting the other and becoming the dominant population.
Bookbinder and Bloom (1980) reported the interaction of Meloidogyne
spp. with bean rust, Uromyces phaseoli (Pers.) Wint. They observed that
IJ. phaseoli and M. incognita were synergistic in suppressing shoot and
root weights of beans. Meloidogyne incognita infections reduced uredial
diameter of U. phaseoli significantly. It was observed that
simultaneous inoculations of _U. phaseoli and M. incognita resulted in
reduced mean numbers of galls per gram of root tissue. Similar effects
were observed when U. phaseoli was inoculated first. Meloidogyne
incognita numbers were consequently reduced by 62% in rusted plants.
This reduction in nematode numbers was probably due to suppressed
translocation of photosythates to the roots (Bookbinder and Bloom,
1980). Egg hatch was, nevertheless, not affected by the fungus.


TABLE 14. The influence of M. incognita and defoliation on snap bean yield loss (%)
Yield loss
(%) by plant
growth stage
Defoliation
level
Log (M. incognita
population +1)
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower
bud
formation
Full Pod
bloom set
0
0
0
0
0
0
0
0
0
3.0
16
11
16
28
21
22
0
4.0
34
41
45
46
49
42
0
5.0
57
65
63
70
62
60
0.25
0
19
40
18
25
6
11
0.25
3.0
29
50
37
49
24
29
0.25
4.0
47
58
49
55
41
45
0.25
5.0
68
78
56
67
53
57
0.50
0
17
15
26
32
21
18
0.50
3.0
34
34
38
50
34
38
0.50
4.0
44
50
44
67
47
55
0.50
5.0
67
62
58
75
59
70
0.75
0
29
32
30
38
39
12
0.75
3.0
45
44
43
50
43
31
0.75
4.0
60
59
68
67
54
45
0.75
5.0
65
71
76
72
71
73
1.0
0
52
83
71
71
80
84
1.0
3.0
67
89
82
83
88
86
1.0
4.0
78
93
90
91
93
91
1.0
5.0
84
96
93
94
93
94
VO


183
Iraneta, M. and R. Rodrigez. 1983. Agrotecnia del frijol, pp. 71-75
In: Curso Intensivo de Postgrado en la produccin de frijol, 40,
Mantazas, Cuba, 1983. Conferencias. Cuba, Ministerio de Agricul
tura.
James, W. C. 1974. Assessment of plant diseases and losses. Annu.
Rev. Phytopathol. 12: 27-48.
James, W. C. and P. S. Teng. 1979. The quantification of production
constraints associated with plant diseases. Appl. Biol. 4:
201-267.
Jenkins, W. R. 1964. A rapid centrifugal flotation technique for
separating nematodes from soils. Plant Dis. Rep. 44: 809
Jimenez, R. M. 1976. Eficacia de phenamiphos, ditrapex y DBCP en el
control de nematodo cecidogeno Meloidogyne spp. en el cultivo del
pijol en Valle de Azapa. Idezia 4: 115-119.
Jones, E. D. 1960. Aecia stage of bean rust found in New York State.
Plant Dis. Rep. 44: 809.
Kalton, R. P., C. R. Weber and J. C. Eldridge. 1945. The effect of
injury simulating hail damage to soybeans. Iowa Agrie. Expt. Sta.
Res. Bull. 357: 733-796.
Kelly, J. D. 1982. Varietal and class diversity and accentuation of
disease problems in a major production area. pp. 12-14 In: Rept.
Bean Improv. Coop, and Nat. Bean Res. Conf. 1982. 5-7 January.
Univ. of Fla., Gainesville.
Keularts, J. L. W. 1980. Effect of the vegetable leafminer, Liriomyza
satiuae Blanchard, and the associated plant pathogens on yield and
quality of the tomato, Lycopersican esculentum Mill. Ph.D.
Dissertation. Univ. of Fla., Gainesville. 154 pp.
Kidney, B. A. 1980. Quantifying expression of resistance to Uromyces
phaseoli. M.S. Thesis. Univ. of Fla., Gainesville. 92 pp.
Kobriger, K. M. and D. J. Hagedorn. 1983. Determination of bean root
rot potential in vegetable production fields of Wisconsin's central
sands. Plant Dis. Rep. 67: 177-178.
Kolmer, J. A., B. J. Christ, and J. V. Groth. 1984. Comparative
virulence of mononaryotic and dikaryotic stages of five isolates of
Uromyces appendiculatus. Phytopathology 74: 111-113.
Kucharek, T. and G. Simone. 1980. Florida plant disease control guide.
IFAS FI. Coop. Ext. Service. Univ. of Fla., Gainesville.
Lamberti, F. 1971. Nematode-induced abnormalities of carrot in
southern Italy. Plant Dis. Rep. 55: 111-113.


Gross dollar value
46
0 25 50 75 100
Defoliation Level (%)
Figure 8. Influence of defoliation on gross values per' hectare of
'Sprite' snap beans defoliated at the primary leaf stage
in the field.


140
There may have been less than adequate retention of the nematicide by
the soil. This less adequate retention of the nematicide may have
contributed to its apparent poor efficacy. Nematode populations were
also generally low at all sampling times in the three tests.
In the study where metam-sodium alone was used, there was signifi
cant relationship between metam-sodium rates and yield. This signifi
cant difference may be due to metam-sodium controlling some soilborne
plant diseases on beans since the nematicide had no significant effect
on nematode populations and populations were very low.
Metam-sodium had no single or interactive effects on yield when it
was used in combination with defoliation and/or bean rust in these
tests. Defoliation was, however, the more important factor affecting
yield when simultaneously used with metam-sodium. The non-significant
interaction between defoliation and metam-sodium may be due to the
generally low nematode populations on the site the study was conducted.
In the study where defoliation, metam-sodium, and bean rust were
used simultaneously there were no significant interactions among the
three factors. There were no significant differences in yield among
defoliation levels, disease severity levels and metam-sodium rates.
There were, however, significant differences in yield among fungicide
sprays. Lack of interaction among the three factors may be due to low
nematode populations and overriding effects of the disease. The disease
was continuously associated with the crop from the time of inoculation
to harvest. The overriding effects were shown by regression analysis
which indicated that disease severity contributed most to the coeffi
cient of determination. Thus, the continuous association of the disease
with the crop may have influenced the physiology of the plants which


Proportion of Loss in Yield
91
Figure 18. The influence of the area under disease progress curve
on snap bean yield.


CHAPTER I
INTRODUCTION
Beans, Phaseolus vugaris L., are the major protein source for many
people in the world, especially in developing countries (Yamaguchi,
1978). Consequently beans are considered an important crop in the
tropics, subtropics, and warm temperate areas of the world (Zaumeyer and
Meiners, 1975). Zaumeyer and Meiners (1975) stated that the leading
world bean producers were Brazil, Mexico, and the United States of
America (U.S.A.) in descending order. Beans are grown for fresh market,
processing, and dry seed. In some countries the leaves are also used as
a vegetable.
Snap beans are known by various names in different areas. These
names include French beans, green beans, pole beans, string beans, and
wax beans (Yamaguchi, 1978). Snap beans are grown in many states in the
U.S. where 132,720 metric tons (mt.) were harvested for fresh-market
consumption from 35,847 hectares (ha.) in 1981 (Anon., 1981). In the
same crop year, 671,640 mt. of snap beans were harvested for processing
from 93,324 ha. (Anon., 1981). The gross value for snap beans in 1981
was $199,282,000 in the U.S. (Anon., 1981).
Florida is the largest producer of fresh-market snap beans in the
U.S., producing nearly 40% of the crop (Anon., 1972, 1982; Rose, 1975;
Ware and McCollum, 1980). In the 1981-82 production year, Florida
produced 60,600 mt. of bush and pole beans from 37,206 ha. (Anon.,
1982). Southeast Florida is the major producing area for snap beans in
1


5
many crops (Allen, 1983; Stoetzer and Omunyin, 1983). Metcalf (1975)
reported that repeated use of pesticides has sometimes led to a decline
in acreage or yield of the crop. As a consequence of this subtle
decline in crop yield due to repeated pesticide use, an integrated pest
management (IPM) approach has been advocated (Huffaker and Smith, 1980;
Waddill et al., 1981). IPM aims at better understanding of the signifi
cance of biological, ecological, and economic processes in the production
of the crop, and the population dynamics of the pest complex, their
predators and parasites, and other factors affecting the system in the
field (Huffaker and Smith, 1980).
Research was undertaken during the period 1984-1985 to investigate
the effects of manual defoliation, root-knot nematode population levels,
and bean rust on snap bean yield. These factors were used separately,
in combinations of two or more of them together, and were studied as
stated in order to determine their threshold levels singly and when they
occurred simultaneously.
Specific experiments were conducted to determine
1. the effect of defoliation on yield of snap beans under field and
open greenhouse conditions,
2. the effect of Meloidogyne incognita population levels on snap bean
yield,
3. the effect of bean rust, Uromyces phaseoli on snap bean yield,
4. the effect of defoliation and M. incognita population levels on
snap bean yield,
5. the effect of defoliation and several nematode genera on snap bean
yield,


108
TABLE 21. Effect of metam-sodium on snap bean yield and populations of
nematode genera in preplant soil samples. Data are means of
3 replicates.
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Yield Yield loss
(g/plot)
0
1
21
360
49
47
1
27
596
16
94
4
72
650
8
187
1
17
680
4
281
0
42
581
18
374
0
25
707
0


98
mancozeb + sulfur spray interval in both trials. The grower would end
up losing income by spraying at this frequency. There was no apparent
explanation for the slight decrease in yield at the 0.098 disease
severity level in trial 1 (Table 16). The decrease in yield may have
been caused by adverse effects of the fungicide mixture on plants. The
fungicides used to manipulate the disease had zinc and sulfur which are
required by plants for proper growth. It was, however, not clear
whether the adverse effect was due to over supply of these elements or
due to the interaction of these elements and factors. This phenomenon
may have been a chance effect which requires further investigation under
similar growing conditions.
In trial 2, yield was consistently inversely related to disease
severity and AUDC. In this trial lower disease severities were correla
ted better with higher yields than the higher disease severities. The
lower disease severity corresponded with more frequent sprays of mancozeb
+ sulfur. These low disease parameters resulted from more frequent
sprays which cost more money. Thus, returns were reduced substantially.
Generally, the shorter 4-5-day mancozeb + sulfur intervals were more
costly than the 7-day intervals and the extra cost did not produce
enough yield to pay for itself. The optimum spray schedule for beans
was, therefore, every 7 days.
From these trials, the grower would be ill-advised to spray snap
beans at 4-5-day intervals if the disease severity is below 0.1 and the
onset of the disease is late. The economic threshold of bean rust
severity was below 0.1 since 0.098 disease severity resulted in 21%
yield loss.


161
59% yield loss. Thus, the threshold level of M. incognita was between 0
and 10 eggs and juveniles per pot. Combining defoliation and root-knot
nematodes showed that there was no significant interaction between them.
When defoliation was held constant, yield reduction rate was greater
than when root-knot nematode populations were held constant. Equations
were devised to express this relationship between the two factors.
Bean rust caused yield loss in snap beans, which was substantial at
the highest disease severity (maximum proportion of foliage infected).
In the first trial, disease severity levels of 0.098, 0.46, 0.65, and
0.76 caused yield losses of 21%, 17%, 35%, and 60% respectively.
Disease severity levels of 0.4, 0.47, 0.71, and 0.86 resulted in yield
losses of 55%, 56%, 80%, and 91% in trial 2. Thus, the disease severity
level which could be tolerated was below 0.1. When bean rust was used
simultaneously with defoliation and nematodes, it was demonstrated by
regression analysis of the data that the disease had an overriding
effect on yield loss. The disease may have had this overriding effect
due to its continual adverse effects on the plants from its onset to
bean harvest. In these experiments, nematodes and defoliation had
little effect on yield. There was no significant interaction among
nematodes, defoliation and bean rust. As expected the best yield was
obtained from plants free of all stress.
From these results, several general conclusions can be drawn. Some
defoliation may result in exposure of photosynthetically active foliage
to light and also may stimulate rapid growth of new leaves which are
highly photosynthetically active. Yield reduction due to defoliation
depends on the growth stage at which leaf removal occurs. This may be
due to the differences in partitioning of photosynthates at these
various growth stages.


TABLE 28. Nematode genera found in soil samples collected (preplant). Data are means of 4
replicates
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Tylenchorhynchus
(No./lOO ml)
0
0
0
4
4
47
0
1
6
2
94
0
3
0
2
187
0
0
2
.0
374
1
0
2
0
116


162
The nematode, M. incognita, caused substantial yield loss when
plants were inoculated in pots, but hardly had any effect on yield under
field conditions. The lack of apparent effect on yield by nematodes may
be due to interaction of several biotic as well as abiotic factors in
field soils. Moreover, the nematode populations in the field were very
low, likely were below threshold levels. In the greenhouse, plants were
grown in pots in which nematodes were probably concentrated on the upper
soil layer thus making the nematode density higher. This discrepancy
between nematode effects on yield in the greenhouse and field may be an
indication that threshold levels determined in the greenhouse may not
necessarily apply under field conditions.
Bean rust caused apparent yield loss even at the lowest maximum
severity level measured. This may be due to the continual adverse
effects of the disease on the physiology and other bean plant growth
factors. A low disease severity, however, required the use of mancozeb
+ sulfur more often hence increasing production costs. Increasing
production costs in turn reduced net returns from the crop. The improve
ment in yield by mancozeb + sulfur may also have been due to increased
supply of micronutrients, such as zinc, which stimulated plant growth.
Thus, yield increases in beans may not necessarily be due to bean rust
control/prevention only.
There may not necessarily be a close relationship between experi
mental results obtained in the greenhouse and the field. Thus, pest
control recommendations should not be based only on greenhouse/laboratory
experimental results. Recommendations for control of a pest complex
should be based on the population dynamics of the key pests supplemented
by the presence of other factors. Pest population levels can be obtained
by close and routine population monitoring throughout the growing season.


104
investigated were 0%, 25%, 50%, and 75% and metam-sodium was applied at
0, 47, 94, 187, and 374 L/ha. Metam-sodium was applied preplant.
Preplant soil samples were taken 12 days after fumigation by compositing
10 soil scoops (to a depth of 6-8 cm) from each main plot. Aliquots of
100 ml soil were processed by sieving and centrifugal flotation.
Subsequent soil samples were taken at midseason and harvest from the
root zone and similarly treated. Only live nematodes were counted from
the preplant soil samples and in the later samples nematodes were killed
before counting.
Defoliation was accomplished by removing the lamina from the distal
end of the petiole using a pair of scissors. Plants were defoliated
once at flower bud formation. Beans were harvested on 31 January to 1
February 1985.
Data were analyzed by analysis variance and covariance followed by
regression analysis using the general linear models procedure of SAS
(Ray, 1982).
The effect of defoliation, metam-sodium and bean rust on snap beans
A 4x3x2 factorial experiment was conducted at the Tropical Research
and Education Center in Homestead, Dade County, Florida, on Rockdale
soil (pH ca. 7.8). The crop was planted on 26 March 1985. Plots
consisted of 3 rows, 3 m long with 0.91 m between rows. Seeds were
planted at 8-10 cm spacing within the row. Fertilizer (8:16:16) was
applied preplant at 448 kg/ha and plants were topdressed at flower bud
formation at 224 kg/ha.
Metam-sodium was applied at 935 L/ha and a fumigated control was
included. Defoliation levels investigated were 0, 25%, and 50%. Bean
rust was manipulated by sprays of bitertanol (57 g ai/ha) at 7-day


ACKNOWLEDGEMENTS
I wish to express my sincere gratitude to many people for their
support and guidance:
Dr. V. H. Waddill, my advisor and committee chairperson, for his
guidance, encouragement, suggestions, and assistance throughout my
Doctor of Philosophy program at the University of Florida.
Dr. J. R. Strayer, co-chairperson, for his constructive suggestions,
encouragement, and providing materials for computer work whenever I
needed them.
Dr. R. McSorley, for serving on my committee, encouragement,
suggestions, and invaluable assistance with statistical data analysis.
Dr. K. Pohronezny, for serving on my committee, encouragement,
suggestions, and assistance with data analysis.
Dr. S. H. Kerr, for his guidance and constructive suggestions
during my Ph.D. program.
I am indebted to Dr. R. D. Berger for his advice, helpful sugges
tions on disease progress analysis, and allowing me to use his computer
programs.
The invaluable technical assistance and friendship of Nancy Shivers,
Diane Putnal, W. (Hank) Dankers, Jorge Parrado, James Reynolds, John
Sarvich, and Ingeborg Stough during the long and difficult field and
laboratory hours are appreciated. I am also indebted to William Meyers,
Joyce Francis, Jeanette Viola, and all staff at Homestead TREC for their
ii


TABLE 15. continued
Gross dollar values by plant growth stage
First Third
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
trifoliate
leaf
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0.25
5.0
low
266
194
383
302
355
337
medium
487
355
702
554
651
617
high
885
641
1276
1007
1184
1122
0.5
0
low
689
750
644
586
596
642
medium
1264
1375
1181
1075
1094
1177
high
2298
2499
2147
1954
1988
2140
0.5
3.0
low
548
582
540
458
586
485
medium
1005
1067
989
840
1075
890
high
1827
1940
1799
1527
1954
1618
0.5
4.0
low
465
441
482
302
398
352
medium
853
808
893
554
730
646
high
1551
1470
1624
1003
1327
1175
0.5
5.0
low
274
335
366
229
310
235
medium
502
615
670
420
568
430
high
912
1118
1219
764
1032
783
0.75
0
low
589
600
522
568
461
689
medium
1081
1099
957
1041
845
1263
high
1965
1999
1740
1893
1536
2297
0.75
3.0
low
457
494
496
458
430
540
medium
838
905
909
840
789
990
high
1523
1646
1653
1527
1435
1800


CHAPTER VI
THE EFFECT OF DEFOLIATION, METAM-SODIUM, AND BEAN RUST ON SNAP BEANS
Introduction
Beans, Phaseolus vulgaris L., are subject to defoliation by a wide
range of factors including insects, diseases, adverse environmental
conditions, mammals, and farm machinery (Agudelo, 1980; Allen, 1983;
Cook, 1978; Costa and Rossetto, 1972; Ruppel and Idrobo, 1962; Vargas,
1980). The extent to which these factors influence yield depends on the
plant growth stage at which they are attacked. Among the most important
insect pests feeding on bean leaves are leafminers (Liriomyza spp.),
cabbage loopers ((Hub.) Trichoplusia ni), leafrollers (Urbanus proteus
L.), Mexican bean beetles (Epilachna varivestis Muls.), and Chrysomelid
beetles (Schoonhoven and Cardona, 1980). Bean rust, Uromyces phaseoli
(Pers.) Wint., is one of the most damaging diseases of bean in bean
producing regions of the world (Acland, 1971; Allen, 1983; Cook, 1978;
Crispin and Dongo, 1962; Iraneta and Rodrigez, 1983; Martinez, 1983;
Schwartz et al., 1979; Stoetzer and Omunyin, 1983; Vargas, 1980). Other
diseases which affect beans include anthracnose (Colletrotrichum
lindemuthianum (Sacc. and Mgr.) Bri. and Cav.), angular leaf spot
(Isariopsis griseola Sacc.), halo blight, (Pseudomonas syringae pu.
phaseolicola (Burkh.) Young, Dye and Wilkie, common blight (Xanthomonas
campestris pu. phaseoli (Smith) Dye, and bean common mosaic virus
(Acland, 1971; Allen, 1983; Martinez, 1983; Stoetzer and Omunyin, 1983).
Root rots caused by Rhizoctonia solani Kuhn, Macrophomina phaseolina
100


6
6. the effect of defoliation, nematodes and bean rust on snap bean
yield, and
7. the effect of inoculation system on the establishment of M.
incognita in beans (Phaseolus vulganis L.).


159
of the work in which this model was able to explain the relationship
between yield and nematode population densities, the intervals between
population levels were smaller particularly at low densities. Thus,
where nematode population levels are far apart, the Seinhorst model may
not be the best for explaining the relationship between plant growth
parameters and nematode population levels.
Results presented here indicate that M. incognita may cause yield
loss in snap beans regardless of the method of inoculation. If the soil
is mixed with 0.525 NaOCl-extracted eggs the influence on yield may be
less profound than if the plants were inoculated with galls or via seed
drench. It has, also, been shown that NaOCl concentrations have a
direct positive relationship to the numbers of nematode eggs and juve
niles extracted. Indicated in the results is the fact the NaOCl has an
egg-hatch inducing effect, compared to eggs allowed to stand in tap
water at room temperature, which is less pronounced as the concentration
increases above 0.525%.


119
TABLE 31. Mean snap bean yield of plants defoliated at various levels
on plots fumigated with metam-sodium.
Defoliation level
(proportion of foliage)
Metam-sodium
(Liters/ha)
Yield (g/plot)
Yield loss (%)a
0
0
545
36
0
47
551
35
0
94
738
13
0
187
851
0
0
374
568
33
0.25
0
469
45
0.25
47
417
51
0.25
94
596
30
0.25
187
403
53
0.25
374
423
50
0.50
0
264
69
0.50
47
253
70
0.50
94
371
56
0.50
187
330
61
0.50
374
335
61
0.75
0
263
69
0.75
47
186
78
0.75
94
276
68
0.75
187
194
77
0.75
374
284
67
£
Yield loss compared to
maximum yield of
851 g.


TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS
ABSTRACT vi
CHAPTER I INTRODUCTION 1
CHAPTER II LITERATURE REVIEW ON DEFOLIATION, AND THE IDENTI
FICATION AND CONTROL OF ROOT-KNOT NEMATODES (Meloidogyne
spp.) AND BEAN RUST (UROMYCES PHASEOLI [PERS.] WINT.) ... 7
Introduction 7
Simulated Leaf Damage on Crop Plants 8
Nematodes Associated with Beans 10
Occurrence and Importance of Root-knot Nematodes ... 10
Epidemiology and Life Cycle of Meloidogyne spp. ... 13
Control of Root-knot Nematodes 15
Identification of Root-knot Nematodes 16
The Importance of Bean Rust 18
Identification and Etiology of the Pathogen 19
Symptoms 20
Epidemiology of the Disease 22
Control of the Disease 2A
Interaction of Root-knot Nematodes and other Pathogens 25
CHAPTER III THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN
YIELD 27
Introduction 27
Materials and Methods 29
Results 31
Discussion 49
CHAPTER IV THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION
ON SNAP BEANS 52
Introduction 52
Materials and Methods 55
Results 57
Discussion 73
CHAPTER V THE EFFECT OF BEAN RUST ON SNAP BEANS 77
Introduction 77
Materials and Methods 80
Results 82
Discussion 96
CHAPTER VI THE EFFECT OF DEFOLIATION, VAPAM, AND BEAN RUST ON
SNAP BEANS 100
Introduction 100
Materials and Methods 102
Results 106
Discussion 139
iv


TABLE 6. Effects of defoliation and defoliation time on gross dollar values per hectare of 'Sprite' snap beans
grown in the greenhouse.
Defoliation
level (%)
Price range3
Gross
Time of
dollar values
defoliation
per hectare
(growth stage)
Primary leaf
First trifoliate
leaf
Third
trifoliate
leaf
Flower bud
formation
Full bloom
Pod set
0
low
292
258
265
270
256
258
medium
535
472
485
495
468
472
high
974
859
885
900
852
859
25
low
187
178
207
189
199
204
medium
343
326
379
347
365
373
high
623
593
689
630
664
679
50
low
193
152
209
189
167
216
medium
353
279
384
347
307
397
high
643
507
698
630
558
772
75
low
178
175
172
165
197
178
medium
327
321
316
302
361
326
high
594
584
574
549
656
593
100
low
96
137
115
59
110
67
medium
177
250
211
109
201
123
high
332
455
384
198
366
223
a
low $7/13.62 kg; medium = $11.20/13.62 kg; and high = $20/13.62 kr of snap beans


CHAPTER III
THE EFFECT OF MANUAL DEFOLIATION ON SNAP BEAN
(PHASEOLUS VULGARIS L.) YIELD
Introduction
Snap beans, Phaseolus vulgaris L., are defoliated by leaf eating
insects, diseases, mechanical injury, and adverse weather conditions
(Agudelo, 1980; Costa and Rossetto, 1972; Ruppel and Idrobo, 1962;
Schoonhoven and Cardona, 1980; Vargas, 1980). Thus, an understanding of
the yield-loss relationship between pest infestations and a crop is
essential for the development of an integrated pest management program.
Much information on the relationship between a crop and pest
infestations has been obtained by simulating pest attack through manual
defoliation of plants (Edje et al., 1972, 1973, 1976; Edje and Mughogho,
1976a, b; Galvez et al., 1977; Greene and Minnick, 1967; Vieira, 1981;
Waddill et al.; 1984). Manual defoliation is not a precise simulator of
pest defoliation (Ruesink and Kogan, 1975); however, it provides a good
estimate of the host-pest relationship. To minimize or avoid
defoliation by pests, producers often resort to preventive pesticide
applications on their crop (Greene and Minnick, 1967). These pesticide
applications are a form of insurance on the crop when little knowledge
on the pest damage-yield loss relationship is available.
Beans are defoliated by a wide range of leaf-eating insects includ
ing leafminers (Liriomyza spp.), cabbage looper (Trichoplusia ni
27


10
stages have been established at 30% and 50% respectively (Keularts,
1980). Extensive research on artificial defoliation effects on tomato
has been conducted by various writers (Keularts, 1980; Wolk et al.,
1983). The effect of leaf removal has also been studied on cotton
(Ludwig, 1926), grain sorghum (Stickler and Pauli, 1961) and wheat and
oats (White, 1962; Wotmack and Thurman, 1962).
Nematodes Associated with Beans
Many nematodes have been found in and around the roots of beans
(Agudelo, 1980; Allen, 1983). Among the nematodes associated with
beans, root-knot nematodes, Meloidogyne spp., are the most important in
tropical and subtropical regions (Agudelo, 1980; Allen, 1983). Table 1
shows the nematode species associated with beans in various bean produc
ing areas (Agudelo, 1980; Ayala and Ramirez, 1964; Bridge, 1973; Bridge
et al., 1977; Castillo and Litsinger, 1978; Caveness et al., 1975,
Feakin, 1973; Hague, 1980; Sinclair and Shurtleff, 1975; Singh and
Farrell, 1972).
Of the four main species of Meloidogyne, M. hapla Chitwood has a
more northerly distribution than M. arenaria (Neal) Chitwood, M.
incognita (Kofoid and White) Chitwood, and M. javanica (Treub) Chitwood
which are cosmopolitan in warmer regions (Allen, 1983; Roberts and
Boothroyd, 1984). The distribution of the other nematode genera is
shown on Table 1.
Occurrence and Importance of Root-knot Nematodes
The most common species of root-knot nematodes are Meloidogyne
arenaria, M. incognita, M. hapla, and M. javanica. Meloidogyne arenaria,
M. incognita, and M. javanica occur worldwide warmer regions whereas M.
hapla has a more northerly distribution (Agudelo, 1980; Allen, 1983;


117
TABLE 29. Nematode genera found in soil samples (midseason). Data are
means of 4 replicates.
Metam-sodium Helicotylenchus Meloidogyne Rotylenchulus
(Liters/ha) (No./lOO ml) (No./lOO ml) (No./lOO ml)
0
3
15
5
47
3
19
3
94
4
15
0
187
8
19
4
374
0
19
3


Log10(No. eggs and juveniles extracted)
149
y=5.8O0.74 In x
y=5.80x
0.12
y=5.38+0.65 In x
y=5.38x0'12
Concentration (%) of
Sodium hypochlorite
Log^CNo. eggs and juveniles extracted) at various
NaOCl concentrations.
Figure 29.


3
(Acland, 1971; Cook, 1978; Crispin and Dongo, 1962; Iraneta and
Rodrigez, 1983; Martinez, 1983; Schwartz et al., 1979; Stoezer and
Omunyin, 1983; Vargas, 1980, Zaumeyer and Thomas, 1957). Other major
bean disease include anthracnose (Colletotrichum lindemuthianum Sacc.
and Magn.), angular leaf spot (Isariopsis griseola Sacc.), halo blight
(Pseudomonas phaseolicola (Burk.) Dows.), common blight (Xanthomonas
phaseoli (E. F. Smith)), and bean common mosaic virus (Acland, 1971;
Allen, 1983; Martinez, 1983; Stoetzer and Omunyin, 1983). Martinez
(1983) stated that root rots caused by Macrophomina phaseolina (Tassi)
Gold.,, Sclerotium rolfsii Sacc., Rhizoctonia solani Kuhn, Pythium spp.
and Fusarium spp. were among the most important diseases of beans.
Fifty-seven races of the bean rust fungus, Uromyces phaseoli, have
been reported in the U.S. (Laundon and Waterston, 1965). The number of
races of _U. phaseoli is, however, not fixed, due to controversy on what
constitutes a physiologic race of a pathogen (Crispin and Dongo, 1972;
Davidson and Vaughan, 1963; Groth and Shrum, 1971; McMillan, 1972).
Uromyces phaseoli has been reported to reduce the translocation of
photosynthetic products from the foliage to the roots and developing
seeds (Daly, 1976; Livne, 1962; Montalbini, 1973; Zaki and Durbin,
1965). The reduction of photosynthetic products translocation is
exacerbated by increased water loss through the damaged leaf cuticle
despite a decrease in transpiration (Vargas, 1980). Water loss in
creases as infection becomes more severe. Infection by IJ. phaseoli
predisposes bean plants to other pathogens such as Pythium spp.,
Rhizoctonia spp. F_. phaseolicola, £. lindemuthianun and many others
(Vargas, 1980).


Proportion of Foliage Infected
85
Days After Inoculation
Figure 15. Disease progress curves for Uromyces phaseoli on snap beans
sprayed with mancozeb at various frequencies (Trial 1).
A = no fungicide spray, C = mancozeb + sulfur at 4-5 day intervals,
D = mancozeb + sulfur at 7-day intervals, and E = mancozeb + sulfur
at 14-day intervals.


20
uredospores blown in from the north serve as primary inocula in Florida.
Later, Kidney (1980) observed telia in Alachua and Dade counties,
contrary to Townsend's findings.
Uredospores overwinter in infected crop debris and trellis poles
(Davison and Vaughan, 1963). These overwintered uredospores are known
to initiate the disease cycle in the next growing season in Oregon and
Maryland (Davison and Vaughan, 1963; Marcus, 1952). In Florida, colder
temperatures than those normal for that state are apparently required
for the uredospores to be viable for relatively long periods (Davison
and Vaughan, 1963).
Disease development is frequently initiated by uredospores under
natural conditions. The uredospore produces a germ tube upon germina
tion. An appressorium which molds itself into the stomatal ledge is
formed when the germ tube gets in touch with the stoma (Mendgen, 1973).
An infection peg develops, from the appressorium and pushes the guard
cells apart until the fungal cytoplasm is transferred into the substoma-
tal vesicle (Vargas, 1980). Enzymes, lipid bodies, and glycogen parti
cles are contained in the vesicle (Mendgen, 1973). The fungus develops
infection hyphae and haustoria as it proceeds inter-cellularly in the
host tissue (Mendgen and Heitefuss, 1975; Vargas, 1980).
The bean rust fungus may complete its life cycle within 10-15 days
after inoculation (Yarwood, 1961). Uredospores are released passively
from pustules and disseminated by farm implements, insects, animals, and
wind currents (Yarwood, 1961; Zaumeyer and Thomas, 1957).
Symptoms
Apparently, bean rust is primarily a foliar disease which occasion
ally occurs on pods, stems and branches (Fromme and Wingard, 1918;
Laundon and Waterston, 1965; Vargas, 1980).


28
(Hub.)) leafroller (Urbanus proteus L.), and beetles (Systates spp.).
Pohronezny et al (1978) reported that Liriomyza spp. were considered by
many local farmers in Dade County, Florida, as the most serious pest on
their crops. Farmers expect yield loss as long as same leaf damage is
observed, but Harris (1974) showed that leaf consumption by pests does
not necessarily result in yield reduction.
Greene and Minnick (1967) reported that snap bean yield was not
significantly reduced until more than 33% of the leaf surface was
removed during blooming. It was later observed that snap bean plants
tolerated up to 66% defoliation if damage occurred before flowering
(Greene, 1971). Vieira (1981) found that 66% leaf loss on an indeter
minate bean cultivar reduced yield when defoliation occurred during
flowering and pod formation. At the first trifoliate leaf stage, Galvez
et al. (1977) found that total (100%) defoliation decreased yield of two
bean cultivars by 34 and 49% respectively. Total defoliation of plants
when only primary leaves were present resulted in yield reduction of 65%
on pole beans in Dade County (Waddill et al. 1984).
Kalton et al. (1945) and Weber (1955) reported that up to 75% leaf
removal in soybeans had little effect on yield if plants were defoliated
prebloom. Defoliation in the bloom and pod formation stages, however,
resulted in significant yield losses (Begum and Eden, 1963, 1964; Camery
and Weber, 1953; Kalton et al., 1945; McAlister and Krober, 1958). Todd
and Morgan (1972) reported significant yield reductions on soybeans with
33%, 67% and 100% defoliation at 2 wk, 4 wk, and 8 wk after first bloom.
Research on the effects of defoliation on crop yield has also been
conducted on tomato, cotton, corn, wheat, oats, and other grain crops
(Dungan, 1930; Keularts, 1980; Ludwig, 1926; Stickler and Pauli, 1961;
White, 1946; Wolk et al. 1983; Womack and Thurman, 1962).


141
which was reflected in yield. The overriding effect of bean rust on
yield was complicated by the fact that it was manipulated by fungicides
which had micronutrients required for plant growth. Duncan's multiple
range test showed that the micronutrient effect was not significant. To
elucidate this micronutrient effect of fungicides on crop yield, further
studies need to be conducted. Bitertanol may have controlled other
pathogens not yet known, hence the higher yield.
Under the conditions these studies were conducted, the grower would
have been better off not fumigating the soil due to the high cost of
metam-sodium and low nematode populations. Fungicides, however, improved
yield and hence gross income. The net return from fungicide sprays
dpended on the frequency of spray. The optimum spray frequency was a
7-day schedule which indicated that spraying beans at intervals shorter
than seven days was not beneficial. The loss a grower would incur
depended on the price of snap beans and the cost of pesticides. Regres
sion analysis appeared to be the best method of predicting yield, hence
income, given various pest combinations. Regression analysis was able
to show which factor contributed more to the coefficient of determination.


167
TABLE 42-1. F-values and probabilities from analysis of variance on the
effects of defoliation, fungicides and their interaction on
snap bean yield.
Source
F
Probability F
Defoliation
29.34
0.0001
Time
1.39
0.24
Defoliation x
Time
1.09
0.36
Fungicide
18.32
0.0001
Defoliation x
Fungicide
0.74
0.75


TABLE 19. The influence of Uromyces phaseoli on the gross dollar value per hectare of snap beans
(Phaseolus vulgaris 'Sprite').
Spray
frequency
(days)
Disease Parameters at harvest
Maximum Proportion of Area under disease
foliage infected progress curve
Price range
Gross dollar values
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0
0.76
0.86
5.64
13.9
high
1797
449
medium
987
247
low
539
135
14
0.65
0.71
5.93
12.85
high
2574
1047
medium
1416
576
low
772
314
7
0.46
0.47
4.11
9.28
high
3665
2327
medium
2016
1280
low
1100
698
4-5
0.098
0.4
0.98
7.33
high
3520
2367
medium
1936
1302
low
1056
710
7 3
0.0
0.0
0.014
0.0
high
4446
5244
medium
2446
2884
low
1334
1573
a
Sprayed with bitertanol


75
these tests soil temperatures were not taken which in fact precludes the
comparison of edaphic temperatures during the time the two tests were
conducted. Soil temperatures are generally warmer when air temperatures
are higher.
The results obtained in the test where M. incognita was used alone
indicate that the threshold population level was between 0 and 10 eggs
and juveniles/pot whereas in the fall experiment the threshold popula
tion level was between 0 and 1,000 eggs and juveniles/pot. During the
fall, 10 and 100 eggs and juveniles were not used hence the threshold
for this test could have possibly, been similar to the summer threshold
level. Generally, defoliation increased yield loss when combined with
nematodes (Table 14). The influence of defoliation level on yield was
not as drastic as expected. It is not apparent why defoliation had this
slight effect on yield. This is not, however, in agreement with the
general principle that nematodes predispose plants to diseases and other
pests. Statistical analysis showed that defoliation and M. incognita
acted independently in influencing yield.
The gross dollar values had a trend similar to that of yield since
they were computed from gross yield. These values are gross figures
from which one has to deduct production costs which include pesticides,
land rent, labor, interest on loans (if any), and farm machinery depre
ciation. Thus, net income would depend on the cost of production and
current market prices of snap beans. Snap bean production is costly
(Taylor and Wilkowske, 1984). Loss in gross income ranges compared to
nematode free plants were $470 to $1443; $441 to $2441; $521 to $2059;
$764 to $2167; $151 to $2201; and $286 to $2192 when plants were
defoliated at various levels at the primary leaf, first trifoliate leaf,


This dissertation was submitted to the Graduate Faculty of the College
of Agriculture and to the Graduate School and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
May 1986
t. £
Dean( /College of Agriculture
&
Dean, Graduate School


Yield (g)
62
Log (M. incognita Population + 1)
Figure 11
Effects of M. incognita on snap bean yield.


MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS
BY
AFETE DIVELIAS GADABU
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1986


16
Crop rotation has been used to reduce nematode numbers in bean
fields (Agudelo, 1980). Beans are planted once every 2 or 4 years in
rotation with a crop such as corn, which is not particularly susceptible
to many nematodes parasitic on beans. Cover crops such as marigold
(Tagetes minuta), rattle box (Crotalaria spectablilis), or hairy indigo
(Indigofera hirsuta) have been used for this purpose (Eguiguren et al.,
1975; Navarro and Barriga, 1970; Rhoades, 1976; Zaumeyer and Thomas,
1957). Other cultural practices used to reduce nematode numbers include
long fallow periods, deep plowing, and flooding (Crispin et al., 1976;
Vieira, 1967).
There are many bean cultivars resistant to M. incognita (Blazey et
al., 1964; Christie, 1959; Fassuliotis et al., 1970; Hartman, 1968;
Ngundo and Taylor, 1974; Rhoades, 1976; Varn and Galvaz, 1974; Wester
et al., 1958). In some cases resistance to M. incognita is broken by
simultaneous infection of M. incognita and M. javanica (Ngundo, 1977).
Ngundo (1977) reported that seven bean lines were resistant to M.
incognita and M. j avanica when they occurred simultaneously.
Identification of Root-knot Nematodes
Maggenti (1981) and Taylor and Sasser (1978) state that root-knot
nematodes were first described by Berneley in England in 1855. M.
incognita was studied, independently, in the U.S. by Neal and Atkinson
in 1889 (Maggenti, 1981). Maggenti (1981) reported that Neal indicated
that root-knot nematodes occurred in Florida before 1805. During these
early studies, Meloidogyne spp. were described under the species names
Heterodera marioni or H. radicicola (Maggenti, 1981). Chitwood and
Chitwood (1950), as a result of their work on the taxonomy of root-knot
nematodes, placed them under the genus Meloidogyne. Chitwood and
Chitwood recognized five species of Meloidogyne and one subspecies.


180
Edje, 0. T., L. K. Mughogho and U. W. U. Ayonoadu. 1972. Agronomy
experiments on beans, Phaseolus vulgaris L. (Sav.). Bunda College
of Agriculture Res. Bull. 3: 20-36.
Edje, 0. T., L. K. Mughogho, and U. W. U. Ayonoadu. 1973. Agronomy
experiments on Phaseolus beans. Bunda College of Agriculture Res.
Bull. 4: 38-67.
Edje, 0. T., L. K. Mughogho and Y. P. Rao. 1976. Effects of
defoliation on bean yield. Bean Improv. Coop. Annu. Rept. 19:
26-29.
Eguiguren, R. G., G. Robalino, and G. Jijn. 1975. Influence of
different crops on nematode populations in Guayllabamba Valley.
Proc. Amer. Phytopath. Soc. 2: 22.
Ekanayake, H. M. R. K. and M. Di Vito. 1984. Effect of population
densities of Meloidogyne incognita on growth of susceptible and
resistant tomato plants. Nemat. Medit. 12: 1-6.
Esser, R. P. 1972. Effect of sodium hypochlorite concentrations on
selected genera of nematodes. Proc. Helminthol. Soc. Washington
39: 108-114.
Esser, R. P., V. G. Perry, and A. L. Taylor. 1976. A diagnostic
compendium of the genus Meloidogyne (Nematoda: Heteroderidae).
Helminthol. Soc. Washington 43: 138-150.
Fassuliotis, G., J. R. Deakin, and J. C. Hoffman. 1970. Root-knot
nematode resistance in snap beans: Breeding and nature of
resistance. J. Amer. Soc. Hort. 95: 640-645.
Feakin, S. D. (ed.). 1973. Pest Control in Groundnuts. PANS Manual
No. 2. Centre of Overseas Pest Control, London. 197 pp.
Feder, W. A. and J. Feldmesser. 1955. Progress report on studies on
the reproduction of the burrowing nematode, Radopholus similis
(Cob) Thorne, on citrus seedlings growing in petri dishes. Plant
Dis. Rep. 39: 395-396.
Ferris, H. 1980. Nematology-Status and Prospects: Practical
implementation of quantitative approaches to nematology. J.
Nematol. 12: 164-170.
Ferris, H. 1984. Nematode damage functions: The problems of
experimental and sampling error. J. Nematol. 16: 1-9.
Ferris, H., W. D. Turner, and L. W. Duncan. 1981. An algorithm for
fitting Seinhorst curves to the relationship between plant growth
and preplant nematode densities. J. Nematol. 13: 300-304.
Fisher, H. H. 1952. New physiologic races of bean rust (Uromyces
phaseoli typica). Plant Dis. Rep. 36: 103-105.


171
TABLE 44. Continued
Defoliation
level (%)
Fungicide
Price
range
Plant
of
Primary
leaf
Growth Stage
defoliation
Flower bud
formation
low
1412
1318
50
Bitertanol
medium
2589
2417
(7-day)
high
4707
4393
low
1208
1200
50
Mancozeb
medium
2215
2201
(4-5-day)
high
4028
4001
low
842
1004
50
Mancozeb
medium
1544
1841
(7-day)
high
2807
3348
low
910
573
50
Mancozeb
medium
1669
1050
(14-day)
high
3034
1909
low
816
769
75
No Fungicide
medium
1496
1410
high
2720
2563
low
1059
1051
75
Bitertanol
medium
1942
1927
(7-day)
high
3531
3504
low
949
785
75
Mancozeb
medium
1740
1438
(4-5-day)
high
3164
2615
low
847
832
75
Mancozeb
medium
1568
849
(7-day)
high
2824
2772
low
855
463
75
Mancozeb
medium
1568
849
(14-day)
high
2851
1543
low
785
581
100
No Fungicide
medium
1438
1064
high
2615
1935
low
730
832
100
Bitertanol
medium
1338
1525
(7-day)
high
2432
2772


31
sprinkler system. The foliage was removed from the distal end of the
petiole and the correct proportion of the foliage removed was determined
from leaf counts.
Ambush^ (40 g ai/ha) or Pydrin^ (250 g ai/ha) was sprayed at 14
(R)
day intervals for leafroller and cowpea curculio control. Benlatev
(550 g ai/ha) and Trigard (150 g ai/ha) were applied fortnightly for
disease and leafminer control respectively. Mesurolv (200 g ai/ha)
was applied as needed for snail and slug control. Treatments were
replicated four times in a randomized complete block. Fresh pod weights
were determined and yield loss computed from the undefoliated plot data
at each growth stage. Yield data were analyzed by analysis of variance
and regression utilizing the general linear models procedure of SAS
(Ray, 1982).
Results
Defoliation had a significant effect on yield in both the green
house and the field, with F-values of 50.16 and 39.95 (p < 0.001)
respectively (Table 2). There was no significant interaction between
time of defoliation and defoliation levels under greenhouse conditions
(F = 0.79). There was significant interaction between defoliation
levels and time of defoliation in the field (F = 3.83). Analysis of
variance on the effects of time of defoliation showed that there were
significant differences between defoliation times (F = 6.23) in the
field but not in the greenhouse.
Regression analysis of the relationship between yield in g/plot and
defoliation level produced models of the form: Y = a+ bx where Y = log
(yield), x = defoliation level and a = intercept. The quadratic model


Manual defoliation caused the highest snap bean yield losses at
full-bloom and pod-set both in the field and the greenhouse experiment.
Snap bean yield loss was observed at the 25% defoliation level in both
experiments. Total defoliation resulted in the highest yield losses.
Yield was negatively correlated to Meloidogyne incognita (Kofoid
and White) Chitwood population levels when plants were grown in pots and
the nematodes were used alone. Yield was also inversely related to
nematode population levels when manual defoliation occurred on nematode
inoculated plants. Yield loss was observed on plants grown in soil
inoculated with 10 eggs and juveniles per pot.
The bean rust disease, Uromyces phaseoli (Pers.) Wint., was
manipulated by fungicide sprays. It was observed that plants with the
highest disease severity gave the lowest yield whereas plants which were
virtually disease free had the highest yield. Generally, fungicide
sprays increased yields. In some cases yield increases were not high
enough to pay for the extra cost of fungicide sprays.
Soil funigation with metam-sodium increased yields slightly. The
optimum metam-sodium application rate was 187 L/ha.
Yield was affected most by the bean rust disease when defoliation,
metam-sodium and the disease were used simultaneously.
vii


165
investigate the effect of manual defoliation and bitertanol and mancozeb
sprays at various frequencies on bean rust and snap bean yield, the
original objective of which had been to determine the effects of bean
rust disease and defoliation on yield. However, due to lack of disease
development, the test was limited to evaluation of the direct effects of
the fungicides and defoliation on snap bean yield.
Materials and Methods
00
Soil was fumigated with Dowfume MC2V at a rate of 314 kg/ha.
Two-gallon side drain plastic pots were filled with 6.4 L of soil and
placed in the field. A plot consisted of 2 pots each with 3 plants.
Plots were 1.5 m apart. Treatments were replicated 4 times in a ran
domized complete block. Treatments were arranged in a 5x5x2 factorial
design. Fertilizer (8:16:16) was applied at 448 kg/ha (to pots and
incorporated) and plants were top dressed at 224 kg/ha just before
(R)
flowering. Trigardv (150g/ha) was applied for leafminer control.
Mesurol was applied for slug and snail control as needed. Other
leaf-eating insects were controlled with sprays of permethrin or
endosulfan.
Beans (Phaseolus vulgaris L., 'Sprite') were planted on 25 January
1985 and harvested on 25 April 1985. Irrigation was provided by an
overhead sprinkler system twice a week. Plants were defoliated at two
growth stages, the primary leaf stage and flower bud formation. Defoli
ation levels investigated were 25%, 50%, 75%, and 100% plus an undefo
liated control. The foliage was removed from the distal end of the
petiole using a pair of scissors.


45
O 25 50 75 100
Defoliation Level
Figure 7. Influence of defoliation on gross dollar value per hectare
of 'Sprite' snap beans defoliated at pod set in the
greenhouse.


17
Esser et al. (1976), however, recognized 35 species in this genus.
Dickson (unpublished) reported that more than 50 species of Meloidogyne
were identified. The number of species in this genus fluctuates due to
various identification procedures used and discovery of new species each
year.
Root-knot nematode speciation is based on perineal patterns, the
distance between stylet knobs and the dorsal esophageal gland opening,
the second-stage juvenile morphology, chromosome number, electrophoresis,
and host range (Maggenti, 1981; Taylor and Sasser, 1978). Host differen
tials are also used to separate races of the same species (Taylor and
Sasser, 1978).
Meloidogyne incognita is the most widely distributed species of
root-knot nematode, comprising 52% of a world collection (40N to 33S)
in areas where annual temperatures are normally within the 18-30C range
(Taylor and Sasser, 1978). This species has four host races as follows:
race 1 does not infect 'Deltapine' 16 cotton, 'NC95' tobacco, and
'Florunner' peanuts; race 2 does not infect 'Deltapine' cotton, and
'Florunner' peanuts; race 3 does not infect 'NC95' tobacco and
'Florunner' peanuts; and race 4 does not infect 'Florunner' peanuts
only. All four races infect 'California Wonder' pepper, 'Charleston
Grey' watermelon and 'Rutgers' tomato (Taylor and Sasser, 1978).
Meloidogyne incognita has a very extensive host range and frequently
coexists with M. javanica (Dickson, unpublished; Santacruz, 1983).
Meloidogyne javanica is the second most widely distributed species,
forming 31% of a world collection (Taylor and Sasser, 1978). Meloidogyne
javanica has no known host races but exhibits variation in chromosome
numbers. M. hapla and M. arenaria comprised 8 and 7% of a world


76
third trifoliate leaf, flower bud formation, full bloom, and pod set
stages respectively. If plants were not defoliated but were inoculated
with nematodes, loss in income ranged from $443 to $1577; $323 to $1911;
$464 to $1827; $855 to $2137; $529 to $1561; and $674 to $1565 when
plants were meant to be defoliated at the primary leaf, first trifoliate
leaf, third trifoliate leaf, flower bud formation, full bloom, and pod
set stages respectively. In each case the lower loss in income is for
the 1,000 eggs and juveniles/pot population level and the higher value
in loss was for the 100,000 eggs and juveniles/pot (Table 15). These
losses are based on yield loss disregarding production costs. The loss
in income has been computed using the high market price of snap beans.
The yields of snap beans in both studies were low, and thus, the grower
would have had a loss in income in both cases.


23
Fifty-seven races of U. phaseoli have been identified in the U.S.
(Stavely, 1984). Laundon and Waterston (1965) reported 35 races of U.
phaseoli. Races 1 and 2 were identified from specimens obtained from
Washington, D.C. and California (Harter et al., 1935). Twenty races of
_U. phaseoli were differentiated according to their reaction on seven
bean cultivars (Harter and Zaumeyer, 1941). Fisher (1952) isolated 10
races from the Rocky Mountain states and Maryland. Race 31 was identi
fied from New Mexico and race 32 from Maryland (Sappenfield, 1954;
Zaumeyer, 1960). Hikida (1961, 1962) isolated and identified races 33
and 34 in Oregon. Race 35 was isolated by McMillan (1972) from the bean
cultivar Dade, which was bred for resistance to previously known races
of £. phaseoli in Florida. McMillan (1972) reported that races 1, 2, 5,
9, 10, 11, and 35 occur in Florida.
There is tremendous variability in the reaction pattern of U.
phaseoli races (Groth and Shrum, 1977). In many areas where several
races occur, there is usually one race which is greatly predominating
(Fisher, 1952).
Uromyces phaseoli races have also been identified outside the U.S.
Thirty-one races have been identified in Mexico (Crispin and Dongo,
1962), 10 races in Colombia (Zuniga and Victoria, 1975), 46 races in
Brazil (Pereira and Chaves, 1977), 12 races in Puerto Rico (Lopez,
1976), 4 races in Nicaragua, 5 races in Honduras (Vargas, 1969, 1970), 7
races in Guatemala (Vargas, 1970), 5 races in El Salvador (Vargas,
1971), 4 races in Peru (Guerra and Dongo, 1973), 11 races in Costa Rica,
11 races in Australia, and 8 races in East Africa (Ballantyne, 1974,
1975; Fisher, 1952; Ogle and Johnson, 1974).


189
Santacruz de la Rosa, E. 1983. Estudios sobre nematodos fitoparasitos
en cultivos associados de dafe y frijol. Tesis Mag. Se. Univ.
Nacional, Bogota, Inst. Comombiano Agro Pecuario. 74 pp.
Sappenfield, W. P. 1954. A new physiologic race of bean rust (Uromyces
phaseoli typica) from New Mexico. Plant Dis. Rep. 38: 282.
Sasser, J. N., C. B. Lucas, and H. R. Powers, Jr. 1955. The relation
ship of root-knot nematodes to black-shank resistence in tobacco.
Phytopathology 45: 459-461.
Sasser, J. N., J. C. Wells, and L. A. Nelson. 1968. The effect of nine
parasitic nematode species on the growth, yield and quality of
peanuts as determined by soil fumigation and correlation of nema
tode populations with host response. Nematologica 14: 15 (Abstr.)
Schein, R. D. 1961. Some aspects of temperature during the colonization
period of bean rust. Phytopathology 51: 674-680.
Schmitt, D. P., F. T. Corbin, and L. A. Nelson. 1983. Population
dynamics of Heterodera glycines and soybean responses in soil
treated with selected nematicides and herbicides. J. Nematol. 15:
432-437.
Schipper, A. L., Jr. and C. J. Mirocha. 1969. The histochemistry of
starch depletion and accumulation in bean leaves at rust infection
sites. Phytopathology 59: 1416-1422.
Schoonhoven, A. Van and C. Cardona. 1980. Insects and other bean pests
in Latin America, pp. 363-412 In: Bean Production Problems. H.
F. Schwartz and G. E. Galvez (eds.), CIAT, Cali, Colombia.
Schuster, M. L. 1959. Relation of root-knot nematodes and irrigation
water to the incidence and dissemination of bacterial wilt of bean.
Plant Dis. Rep. 43: 27-32.
Schwartz, H. F., J. M. Castao and C. R. Rivero. 1974. Enfermedades
del frijol. Centro Internacional de Agricultura Tropical, Cali,
Colombia. 28 pp.
Seinhorst, J. W. 1965. The relation between nematode density and
damage to plants. Nematoligica 11: 137-154.
Seinhorst, J. W. 1972. The relationship between yield and square root
of nematode density. Nematologica 18: 585-590.
Shaner, G and R. E. Finney. 1977. The effect of nitrogen fertilization
on the expression of slow-mildew resistence in Knox wheat. Phyto
pathology 67: 1051-1056.
Sharma, R. D. and R. J. Guazelli. 1982. Evaluation of bean breeding
lines for resistance to root-knot nematode, M. javanica. Reuniao
Porasileira de Nematologia 5: 99-107.


89
TABLE 18. Regression equations for the relationship between bean rust
and snap bean yield (Trial 2).
Plant growth stage
Regression equation
First and Third trifoliate Disease incidence negligible
Flower bud formation
y
=
2492.5 -
7790.18x
(R2
=
0.39NS)
y
=
2443.73 -
23678.58d
(R2
=
0.36NS)
Full bloom
y
=
1940.83 -
1141.86x
(R2
=
0.16NS)
y
=
1734.26 -
1384.95x
(R2
=
0.21NS)
Pod set
y
=
3113.46 -
893.33x
(R2
=
0.89*)
y
=
2886.63 -
5340.87d
(R2
=
0.92**)
Pods half formed
y
=
3099.74 -
369.12x
(R2
=
0.98**)
y
=
3220.42 -
3224.52d
(R2
=
0.99**)
Pods fully formed
y
=
3165.01 -
203.23x
(R2
=
0.98**)
y
=
3288.9 -
2795.75d
(R2
=
0.98**)
* R Significant at 0.05
** R Significant at 0.01
y = yield (g/plot)
x = area under the disease progress curve
d = proportion of foliage infected
NS = not significant


Proportion of Loss in Yield
135
Figure 26. The relationship between yield loss and maximum proportion
of foliage infected.


32
Gross dollar values were obtained by multiplying yield (kg/ha) by
current market prices of 13.62 kg of snap beans which were $6 (low), $11
(medium), and $20 (high), respectively. Net returns from investment
were derived by subtracting the gross dollar values of unsprayed plants
and the cost of mancozeb from the values realized from sprayed plants.
No net returns for bitertanol spray were computed because this fungicide
was experimental and its price was not available.
Data were analyzed by the analysis of variance and regression
analysis using the general linear procedure of SAS (Ray, 1982).
Results
Figures 15 and 16 show disease progress in trials 1 and 2 respec
tively. Progress patterns were similar in both trials although disease
severity values were higher for trial 2.
Analysis of variance of yield data by treatments gave F values of
5.37 and 10.77 with probabilities of 0.01 and 0.005 for trials 1 and 2,
respectively. This showed that there was a significant relationship
between yield and disease severity. The disease free plants produced
higher yield in trial 2 than in trial 1 (Table 16). In trial 1 disease
severity ranged from 0.098 to 0.76 and yield was from 1102 g to 2723
g/plot whereas in trial 2 disease severity ranged from 0.4 to 0.86 and
yield was 276 g to 3214 g (Table 16). Generally, where the disease
occurred, yield was lower in trial 2 than in trial 1.
Figure 17 shows the relationship between yield loss and maximum
proportion of foliage infected (disease severity) for both trials. In
trial 1, 0.098, 0.46, 0.65, and 0.76 disease severity resulted in 21%,


33
TABLE 2. F-values from analysis of variance for the effects of defoli
ation, time (plant growth stage) and their interaction on snap
bean yield.
Field
Greenhouse
Source
F
Prob. F
F
Prob. F
Defoliation
50.16
0.0001
39.95
0.0001
Growth Stage
6.23
0.0001
0.35
0.92
Defoliation X
Growth Stage
3.83
0.001
1.04
0.79


CHAPTER IV
THE EFFECT OF ROOT-KNOT NEMATODES AND DEFOLIATION ON SNAP BEANS
Introduction
Root-knot nematodes, especially Meloidogyne incognita (Kofoid and
White) Chitwood, are a serious threat to bean (Phaseolus vulgaris L.)
production in many bean-producing areas of the world (Agudelo, 1980;
Allen, 1983; Ngundo, 1977; Sharma and Guazelli, 1982; Singh et al.
1981a). M. incognita has been reported to cause extensive root-galling
on bean plants, which interferes with nitrogen fixation by Rhizobium
spp. as well as nutrient uptake by the root system. Moreover, M.
incognita has been reported to increase the severity of other pathogens
through predisposition of host plant root tissues (Carter, 1975a,b;
Golden and Van Gundy, 1975; Powell, 1971; Powell and Nusbaum, 1960;
Porter and Powell, 1967; Sasser et al., 1955). Thus, yield loss caused
by M. incognita and related species is not always a unitary effect, but
often a result of interaction of these nematodes with other plant-
parasitic organisms.
Yield losses of 50 to 90% in field beans have been reported due to
severe root-knot nematode infections (Agudelo, 1980; Freire and Ferraz,
1977; Ngundo, 1977; Varn and Galvez, 1974). Yield decreases caused by
M. incognita are also well known in other crops (Allen, 1983; Lambert!,
1979). Crop yield is usually expected to be inversely related to
nematode counts (Sasser et al., 1968). In view of this theory, Barker
52


139
Disease progress curves in unsprayed controls and those sprayed with
mancozeb + sulfur at 14-day intervals were similar (Figures 22 and 24).
The 7-day spray schedule of mancozeb produced progress curves with
maximum disease severity below those of the other two spray schedules
(Figure 23).
Vapam had no significant effect on nematode populations at preplant
and harvest (Tables 36 and 37). There were virtually no nematodes
detected at midseason.
Yield and gross dollar values per hectare are shown in Table 38.
The highest yield and gross dollar values were obtained from fumigated
plots with plants kept virtually disease free and not defoliated. The
lowest yield was obtained from fumigated plots with plants which had
0.50 defoliation level and 0.71 disease severity (Table 38). Generally,
yield responses were not proportional to defoliation levels and
metam-sodium rates. Fumigation resulted in net loss of income regard
less of defoliation level and disease severity (Table 39). No defolia
tion, 0.83 disease severity, and fumigation with vapam at 374 L/ha
resulted in loss of income of $1191, $1320 and $1398 at the high, medium
and low prices respectively whereas plots not fumigated, plants not
defoliated, and having 0.87 disease severity had net gains in income of
$300, $165 and $90 at the high, medium and low prices respectively
(Table 39). Net gains in income varied in magnitude depending on
disease severity and defoliation.
Discussion
The lack of consistent reduction in nematode populations on
treatment with metam-sodium may be related to application technique.


179
Coyne, D. P. and M. L. Schuster. 1975. Genetic and breeding strategy
for resistance to rust (Uromyces phaseoli (Reben) Wint.) in beans
(Phaseolus vulgaris L.) Euphytica 24: 795-803.
Crispin, A. and S. Dongo. 1962. New physiologic races of bean rust,
Uromyces phaseoli typica from Mexico. Plant Dis. Rep. 46: 411-413.
Crispin, A., J. A. Sifuentes, and J. Campos. 1976. Enfermedades y
plagas del pijol en Mexico. Inst. Nac. Invest. Agr. Mexico Foil.
Tec. No. 39: 22-24.
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Physiological Plant Pathology. R. Heitefuss and P. H. Williams
(Eds.) Springer-Verlag, Berlin.
Daly, J. M., A. A. Bell, and L. R. Krupka. 1961. Respiration changes
during development of rust diseases. Phytophathology 51: 461-471.
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race 33 of Uromyces phaseoli var. phaseoli in storage.
Phytopathology 53: 736-737.
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la presencia de nematodes formadonas de agallas en las raices
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Technologia, Universidad Central de las Villas 1(2): 19-29.
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resistant and susceptible tomato. Nemat. Medit. 11: 151-155.
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Coop. Annu. Rept. 19: 34-35.


9
yield by 11, 20, 20, and 70% respectively. At the same levels of leaf
area reduction, defoliation at initiation of flowering decreased yield
by 18, 12, 19, and 55% respectively. At the formation of the third
trifoliate leaves only total defoliation affected the yield.
Kalton et al. (1945) and Weber (1955) reported that 50% and 75%
leaf removal in soybeans had little effect on yield when defoliation
occurred at the prebloom stage. Significant yield losses were, however,
observed when plants were heavily defoliated at the bloom or pod set
stages (Begum and Eden, 1963, 1964; Camery and Weber, 1953; Kalton et
al., 1945; McAlister and Krober, 1958). Todd and Morgan (1972) observed
significant yield reduction on soybeans with 33, 67, and 100% leaf
removal at 2 wk, 4 wk, and 8 wk after first bloom. Wilkerson et al.
(1984) reported that all defoliations on Florunner' peanuts resulted in
lower stem weight to length ratios and lower pod numbers and weights.
It was observed that defoliation altered the normal partitioning of
photosynthates between plant parts in peanuts. Wit (1983) reported that
during the most sensitive period (July) in the Netherlands, 60% defolia
tion induced a maximum yield reduction of 35% in Brussel sprouts. He
also noted that when partial defoliation was carried out 15 wk after
transplanting or later, no effect on yield was observed. Douglas et al.
(1981) observed a grain yield reduction of 77% in corn when complete
defoliation was carried out at silking. Grain yield losses decreased
with delay in defoliation toward maturity. Less severe defoliations,
however, resulted in smaller reductions in yield. Generally, grain
yield was tolerant of post-silking defoliation and yield losses
exceeding 20% were recorded only after 67% of the leaves were removed.
Defoliation action thresholds for tomato for the prebloom and postbloom


CHAPTER II
LITERATURE REVIEW ON DEFOLIATION AND THE IDENTIFICATION AND CONTROL
OF ROOT-KNOT NEMATODES (MELOIDOGYNE SPP.) AND BEAN RUST
(UROMYCES PHASEOLI [PERS.] WINT.)
Introduction
Bean insect pests such as leafminers (Liriomyza spp.), bollworms
(Heliothis armigera Hbn.) and leaf rollers (Urbanus proteus L.) as well
as diseases including rust (Uromyces phaseoli [Pers.] Wint.)> web blight
(Thanatephones cucumeris [Frank] Donk) and angular leaf spot (Isariopsis
griseola Sacc.) not only destroy plant foliage but also cause physio
logical damage in some cases (Acland, 1971; Galvez et al., 1977).
Root-knot nematodes, Meloidogyne spp., cause prolific galls on the root
system of plants which may lead to the following above-ground symptoms:
incipient wilting, stunted growth, and chlorotic leaves, often with
burnt out edges (Agudelo, 1980). A combination of these organisms on
beans usually leads to great losses in yield. Control of these pest
problems has been mainly by the use of chemical pesticides (Acland,
1971; Agudelo, 1980).
This review is a summary of the problems encountered in the identi
fication of root-knot nematodes and bean rust and the use of simulated
leaf damage on beans.
Simulated Leaf Damage on Crop Plants
Bean plants are susceptible to defoliation by insects, diseases,
hail, moisture stress, and mechanical injury resulting from farm
7


166
Two fungicides were tested for the manipulation of bean rust.
These were; bitertanol (57 g ai/ha) applied at 7-day intervals, and
mancozeb (1.7 kg/ha) tank-mixed with sulfur (4.2 kg ai/ha) applied at
4-5, 7, and 14-day intervals. An unsprayed control was also included.
Inoculum exposure was done at the primary leaf stage by clipping
infected pole bean leaves on a string tied across experimental plots 1.5
m above the ground. Fungicide sprays were initiated soon after inocula
tion. Disease progress was assessed by the Horsfall-Barratt rating
system.
Data were subject to analysis of variance.
Results
Analysis of variance on the effect of fungicides and defoliation on
snap bean yield showed that there were significant differences among
fungicides and manual defoliation levels (F = 18.32** and 29.34** (P <_
0.01) respectively) (Table 42-1). There were, however, no significant
interactions between fungicides and manual defoliation levels (Table
42-1). There were no significant differences between defoliation times
(F = 1.39 N.S.) and interaction among manual defoliation levels and time
(F = 1.09 N.S.). Duncan's multiple range test showed that there were no
significant differences in yield between the unsprayed plants and those
sprayed with mancozeb and sulfur at 7 and 14-day intervals (Table 42-2).
There were, however, significant differences in yield between mancozeb
at all schedules and bitertanol (Table 42-2).
The highest snap bean yield was obtained from plots sprayed weekly
with bitertanol at all defoliation levels and both times of defoliation


193
Wit, A. K. H. 1983. The relation between artificial defoliation and
yield in brussel sprouts as a method to assess the quantitative
damage induced by leaf-eating insects. Z. Angew. Entomol. 94:
425-431.
Wolk, T. 0., D. W. Kretchman, and D. G. Ortega. 1983. Responses of
tomato to defoliation. J. Amer. Hort. Sci. 108: 536-540.
Womack, D. and R. L. Thurman. 1962. Effect of leaf removal on the
grain yield of wheat and oats. Crop Sci. 2: 423-426.
Yamaguchi, M. 1978. World's Vegetables: Principles, Production and
Nutritive Values. Univ. of Ca., Davis. 223 pp.
Yarwood, C. E. 1961. Uredospore production by Uromyces phaseoli.
Phytopathology 51: 22-27.
Yen, D. E. and R. M. Brien. 1960. French bean rust (U. appendiculatus).
Studies on resistance and determination of rust races in New
Zealand. N. Z. J. Agrie. Res. 3: 358-363.
Yoshii, K. 1977. The therapeutic effect of fungicides in the control
of bean rust. Fitopatologa 12: 99-100.
Yoshii, K. and G. E. Galvez. 1975. The effect of rust on yield
conponents of dry beans (£. vulgaris). Proc. Phytopath. Mtg.,
Caribbean Div., (Abstr.).
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translocation of photosynthetic products from diseased leaves.
Phytopathology 55: 528-529.
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Rep. 44: 459-462.
Zaumeyer, W. J. and J. P. Meiners. 1975. Disease resistance in beans.
Annu. Rev. Phytopathol. 13: 313-334.
Zaumeyer, W. J. and H. R. Thomas. 1957. A monographic study of bean
diseases and methods of their control. USDA Agrie. Bull. No. 868.
255 pp.
Zuniga de Rodriguez, J. E. and J. I. Victoria. 1975. Determinacin de
las razas fisiolgicas de la roya del frijol (Uromyces phaseoli
var. typica) Arth. en el Valle del Cauca. Acta Agron. 25: 75-85.


115
TABLE 27. F-values from analysis of variance for the effects of metam-
sodium, defoliation and their interaction on snap bean yield.
Source
F
Probability of F
Metam-sodium
1.5
0.22
Defoliation
20.22
0.0001
Defoliation x
metam-sodium
0.62
0.82


192
Varn, F. H. and G. E. Galvez. 1974. Informe anual de Labores Programa
de Fitopalogia. ICA, Palmira, Colombia. 10 pp.
Venette, J. R., B. M. Olson, and J. B. Nayes. 1978. Bean rust pycnia
and aecia in North Dakota. Annu. Rept. Bean Improv. Coop. 21: 49.
Vieira, C. 1967. Feijoeiro-comum-cultura, Doencas e Melhoramento. pp.
84-124. In: Imprensa Universitaria, Vicosa, Brazil.
Vieira, C. 1981. Effect of artificial defoliation on the yield of two
indeterminate bean CP. vulgaris) cultivars. Turrialba 31: 383-385.
Villamonte, R. 1965. Ensayo comparativo de nematicide en el cultivo
del tomatero. An Ciento 3: 206-214.
Vrain, T. C. 1977. A technique for the collection of larvae of
Meloidogyne spp. and comparison of eggs and larvae as inocula. J.
Nematol. 9: 249-251.
Waddill, V. H., R. McSorley, and K. Pohronezny. 1981. Field monitoring:
basis for integrated management of pests on snap beans. Trop.
Agrie. 58: 157-159.
Waddill, V. H., K. Pohronezny, R. McSorley, and H. H. Bryan. 1984.
Effect of manual defoliation on pole bean yield. J. Econ. Entomol.
77: 1019-1023.
Walker, J. L. 1965. Patologia vegetal. Trad. Antomio Aguirre. (Omega
edit.) Barcelona, Spain. 818 pp.
Wang, D. 1961. The nature of starch accumulations at the rust
infection site in leaves of pinto bean plants. Can. J. Bot. 39:
1595-1604.
Ware, G. W. and J. P. McCollum. 1980. Producing vegetable crops.
Interstate printers, Danville. 607 pp.
Weber, C. R. 1955. Effects of defoliation and toppings simulating hail
injury to soybeans. Agron. J. 47: 262-266.
Wester, R. E., H. B. Cordner and P. H. Massey, Jr. 1958. Nemagreen, a
new Lima. Amer. Veg. Grower 6: 31-32.
White, L. V. 1962. Root-knot and seedling disease complex of cotton.
Plant Dis. Rep. 46: 501-504.
Wilkerson, G. G., J. W. Jones, and S. L. Poe. 1984. Effect of
defoliation on peanut (Arachis hypogaea cv. Florunner) plant
growth. Crop Sci. 24: 526-531.
Wimalajeewa, D. L. S and P. Thavam. 1973. Fungicidal control of bean
rust disease. Trop. Agrie. 129: 61-66.


132
TABLE 39. Net
returns ($) on
investment per
hectare
of snap beans.
Defoliation
Net Income
level
Metam-sodium
Disease
Price Range
(proportion
of foliage)
(Liters/ha)
severity
high
medium low
0
0
0.87
300
165
90
0
935
0.83
-1191
-1324
-1398
0.25
0
0.64
124
68
38
0.25
935
0.71
-1224
-1342
-1408
0.5
0
0.77
59
32
9
0.5
935
0.74
-1236
-1349
-1562
0
0
0.02
1016
589
305
0
935
0
- 438
910
-1172
0.25
0
0
670
346
189
0.25
935
0.01
- 973
-1204
-1332
0.5
0
0
427
235
128
0.5
935
0
- 892
-1160
-1308
0
0
0.56
213
92
81
0
935
0.59
- 908
-1194
-1352
0.25
0
0.7
420
245
108
0.25
935
0.66
-1394
-1461
-1498
0.5
0
0.68
64
10
- 20
0.5
935
0.71
-1610
-1580
-1563
0
0
0.71
263
161
54
0
935
0.74
-1158
-1321
-1412
0.25
0
0.75
- 24
- 28
- 31
0.25
935
0.83
-1235
-1364
-1435
0.5
0
0.72
199
94
36
0.5
935
0.81
-1194
-1451
-1436


TABLE 15. Effects of M. incognita and defoliation on the gross dollar values per hectare of snap
beans.
Gross dollar values by plant growth stage
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
First
trifoliate
leaf
Third
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0
0
low
830
882
870
916
755
783
medium
1522
1617
1595
1679
1384
1435
high
2768
2940
2900
3053
2517
2609
0
3.0
low
697
785
731
659
596
611
medium
1278
1439
1340
1209
1094
1119
high
2325
2617
2436
2198
1988
2035
0
4.0
low
548
520
478
495
385
454
medium
1005
954
877
907
706
833
high
1827
1734
1595
1649
1283
1514
0
5.0
low
357
309
322
275
284
313
medium
655
566
590
504
521
574
high
1191
1029
1073
916
956
1044
0.25
0
low
673
529
714
687
710
697
medium
1233
970
1308
1259
1301
1278
high
2242
1764
2379
2289
2366
2323
0.25
3.0
low
589
441
548
467
574
556
medium
1081
808
1005
856
1052
1019
high
1965
1470
1827
1557
1913
1853
0.25
4.0
low
440
370
444
412
445
430
medium
807
679
813
756
817
789
high
1467
1235
1479
1374
1485
1435


Gross dollar value
1200
1000
800
600
400
200
0
0 25 50 75 100
Defoliation Level (%)
Figure 9. Influence of defoliation on gross dollar values per
hectare of 'Sprite' snap beans defoliated at the third
trifoliate leaf stage in the field.


49
a higher dollar value than the undefoliated control. Removal of 25% of
the foliage at the primary leaf stage and full bloom also resulted in
slightly higher dollar values than the undefoliated control. Generally,
increased defoliation resulted in lower dollar values per hectare.
Discussion
Total defoliation when only primary leaves were present resulted in
69% and 55% yield loss in the greenhouse and field respectively. This
level of defoliation resulted in 57%, 95%, 74% and 92% yield reduction
in the greenhouse and field when plants were defoliated at full bloom
and pod set, respectively. The lower yield reduction in the greenhouse
may be due to the better controlled environmental conditions. The only
growth stage at which total defoliation resulted in less yield loss in
the field was at the primary leaf stage. This may have been due to
better recovery of plants from the total defoliation in the field.
Removing 25% of the foliage resulted in yield loss of at least 20%
in the greenhouse at all growth stages. This is a substantial loss in
terms of dollar values. Thus, it appears that the economic threshold
under greenhouse conditions was between 0 and 25% defoliation. In the
field the economic threshold level varied with the plant growth stage
(Table 4). The increase in yield in the field may have been due to
compensatory reactions of the plant. The compensation may have resulted
from increased photosynthesis due to increased exposure of the remaining
photosynthetically active foliage to light. The same may apply to the
increase in yield of plants with 75% defoliation at the first trifoliate
leaf stage. Reducing foliage may have increased air circulation among


188
Rhoades, H. L. 1976. Effect of Indigofera hirsuta on Belonolaimus
longicaudatus, Meloidogyne Incognita, and M. javanica and subse
quent crop yields. Plant Dis. Rep. 60: 384-386.
Rivera, G. 1977. Incorporacin de resistencia a la raza 29 de la roya
del frijol comum (Uromyces appendiculatus Pers.) Fr. en el cultivar
Pacuaral Vaina Morada. Tesis Ing. Agr. Univ. de Costa Rica, 55 pp.
Robbins, R. T., 0. J. Dickerson, and J. H. Kyle. 1972. Pinto bean
yield increased by chemical control of Pratylenchus spp. J.
Nematol. 4: 28-32.
Roberts, D. A. and C. W. Boothroyd. 1984. Fundamentals of Plant
Pathology. Freeman, New York, 432 pp.
Roberts, P. A. 1983. Influence of carrot planting date on Meloidogyne
inconita infection and damage. J. Nematol. 15: 488 (Abstr.).
Rodriguez, V. A. 1976. Evaluacin de variadades crillas e introducidas
de frijol comum resistentes a roya (Uromyces phaseoli var. typica)
en El Salvador, pp. 30-39 In: Reunion Anual del Programa Coopera
tivo Centro Americano para el Mejoramiento de cultivos Alimentorios
22a, San Jose, Costa Rica.
Romig, R. L. and L. Calpouzos. 1970. The relationship between stem
rust and loss in yield of spring wheat. Phytopathology 60:
1801-1805.
Rose, C. N. 1975. Snap bean production in Florida: a historical data
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Reporting Service.
Rotem, J., Y. Ben-Joseph and R. Reuveni. 1973. Design and use of an
automatic humidity chamber in phytopathological research. Phyto-
parasitica 1: 39-45.
Rudolf, K. and N. Baykal. 1978. Diseases of bean (Phaseolus vulgaris)
in south and western Turkey. Annu. Rept. Bean Improv. Coop. 21:
45-47.
Ruesink, W. G. and M. Kogan. 1975. The quantitative basis of pest
management: Sampling and measuring pp. 309-351 In: Introduction
to Pest Management, R. L. Metcalf and W. H. Luckman (eds.). Wiley,
New York. 587 p.
Ruppel, R. F. and E. Idrobo. 1962. Lista preliminar de insectos y
otros animales que daman frijoles en America. Agr. Trop. 18:
651-679.
Saka, V. W. 1982. International Meloidogyne Project Report in Malawi,
pp. 31-36 In: Research Planning Conference on root-knot nematodes,
Meloidogyne spp., 3rd, regions IV and V, IITA, Ibadan, Nigeria,
1981.


83
17%, 35%, and 60% yield loss respectively whereas in trial 2 disease
severity of 0.4, 0.47, 0.71, and 0.86 lead to 55%, 56%, 80%, and 91%
yield loss respectively. Figure 18 is a representation of the relation
ship between area under the disease progress curve (AUDC) and yield for
trials 1 and 2. Both disease severity and AUDC were positively cor
related with yield loss which showed that these disease measures were
inversely related to yield. Figures 17 and 18 are similar in shape.
Regression equations between disease severity and snap bean yield shown
in Tables 17 and 18 are similar in nature. Regression analysis of the
data produced the model y = a + bx, y = yield (g/plot), and x = disease
parameters. Both disease severity and AUDC were significantly corre
lated with yield at full bloom and pod set in trial 1 (Table 17). When
pods were fully developed only disease severity was significantly
correlated with yield. In trial 2, disease severity and AUDC were
significantly correlated with yield at pod set through the stage when
pods were fully formed (Table 18). In trial 1 the coefficient of
2
determination (R ) at full bloom or later range from 0.46 to 0.93 (Table
2
17) while in trial 2 the coefficient of determination (R ) at pod set
or later ranged from 0.89 to 0.99 (Table 18). In trial 1 bean rust
severity was not significantly correlated with yield at the stage when
pods were half developed whereas the in trial 2 the disease was not
significantly related to yield at flower bud formation and full bloom
(Tables 17 and 18).
The gross dollar values per hectare of snap beans infected by bean
rust are shown in Table 19 and Figures 19 and 20. The virtually disease
free plants gave the highest dollar values per hectare in both trials.
These plants were sprayed with bitertanol an experiment fungicide, which
is currently not registered for rust control on beans. Therefore, no


95
5500
5000
4000
3000
2000
1000
0
0 0.40 0.47 0.71 0.86
Disease Severity
Figure 20. Influence of disease severity on gross dollar value per
hectare of 'Sprite' snap beans in trial 2.


43
O 25 50 75 100
Defoliation Level
Figure 5. Influence of defoliation on gross dollar value per hactare
of 'Sprite' snap beans defoliated at the primary leaf
stage in the greenhouse.


170
TABLE 44. Effect of fungicides
and defoliation
on gross
dollar values
per
hectare of snap
beans.
Plant
Growth Stage
of
defoliation
Defoliation
Price
Primary
Flower bud
level (%)
Fungicide
range
leaf
formation
low
1122
1193
0
No fungicide
medium
2054
2186
high
3740
3975
low
1663
1820
0
Bitertanol
medium
3049
3337
(7-day)
high
5544
6067
low
451
1232
0
Mancozeb
medium
2661
2258
(4-5-day)
high
4838
4106
low
1130
1216
0
Mancozeb
medium
2071
2230
(7-day)
high
3766
4054
low
989
1028
0
Mancozeb
medium
1812
1884
(14 days)
high
3295
3426
low
879
973
25
No fungicide
medium
1611
1784
high
2929
3243
low
1420
1349
25
Bitertanol
medium
2604
2474
(7-day)
high
4734
4498
low
1271
1193
25
Mancozeb
medium
2330
2186
(4-5-day)
high
4237
3975
low
1153
1169
25
Mancozeb
medium
2114
2143
(7-day)
high
3844
3897
low
1028
973
25
Mancozeb
medium
1884
1784
high
3426
3243
low
777
745
50
No fungicide
medium
1424
1366
high
2589
2485


30
(7.5 L) pots were filled with 6.4 L soil and placed on a corrugated
bench 0.91 m high. Six seeds were planted in each pot and thinned to
three after emergence. A plot consisted of three pots with three
plants/pot. The crop was irrigated twice a day using an automatic
time-controlled, water-mist-producing overhead system. The foliage was
removed from the distal end of the petiole and the correct number of
leaves removed at a particular growth stage was determined by leaf
counts.
The treatments were replicated four times and randomized in a
complete block. Fresh weights of pods were determined. Yield loss
(percentage) was computed from the untreated control yield at each
growth stage. The dollar economic value was computed by extrapolating
plot data to a per hectare basis. Plot yield data were subjected to
analysis of variance and regression using the general linear models
procedure of SAS (Ray, 1982).
Field experiment
The herbicides Treflan^ (841 g ai/ha) and Dual^ (1.7 kg ai/ha)
were applied to the site prior to planting. Plots were kept as weed-
free as possible by mechanical cultivation. Plots were three rows wide
(0.91 m row spacing) and 6 m long. Seeds were mechanically planted at
8-10 cm spacing within the row. The crop was irrigated using an overhead


TABLE 15. continued
Gross dollar values by plant growth stage
First Third
Defoliation
level
Log (M. incognita
population +1)
Price
range
Primary
leaf
trifoliate
leaf
trifoliate
leaf
Flower bud
formation
Full
bloom
Pod set
0.75
4.0
low
332
362
278
302
347
430
medium
609
663
510
554
636
789
high
1107
1206
928
1007
1157
1435
0.75
5.0
low
291
256
209
256
219
133
medium
533
469
383
470
401
244
high
969
853
696
855
730
443
1.0
0
low
399
150
252
266
151
125
medium
731
275
463
488
277
229
high
1329
499
841
885
504
417
1.0
3.0
low
274
97
153
155
91
110
medium
502
178
288
285
167
201
high
912
323
523
518
303
366
1.0
4.0
low
183
62
87
82
53
70
medium
335
113
160
151
97
129
high
609
206
291
275
176
235
1.0
5.0
low
133
35
61
55
53
47
medium
244
65
111
101
97
86
high
443
117
203
184
176
157


168
TABLE 42-2. Mean snap yield per hectare sprayed with various fungicides.
Fungicide
Spray frequency
(days)
Mean yield (g/plot)
No fungicide
0
114 cd
Mancozeb + sulfur
14
98 d
Mancozeb + sulfur
7
119 c
Mancozeb + sulfur
4-5
137 b
Bitertanol
7
166 a
Means with the same letter are not significantly different at P<^ 0.05
(Duncan's multiple range test).


105
Intervals, mancozeb (1.7 kg ai/ha) plus sulfur (4.5 kg ai/ha) at 7-day
and 14-day Intervals. A no-spray plot was included to ensure a high
(R)
disease level at specific times of assessment. Helenav sticker or Nu
(R)
Filmv -17 was used as a surfactant in all fungicide sprays.
Preplant soil samples were taken by compositing 10 soil scoops from
each plot 12 days after fumigation. Aliquots of 100 ml soil were
processed by sieving, and nematodes were extracted by centrifugal
flotation (Jenkins, 1964). Subsequent soil samples taken at mid season
and at harvest were similarly handled. Only live nematodes were counted
in the preplant soil samples. In the later samples the nematodes were
first killed by heating in a water bath and then counted.
Plants were subjected to rust inoculum at the primary leaf stage by
clipping infected pole bean leaves on to wire stakes just east of the
test plots and 25 cm off the ground. Disease progress was monitored by
taking 5 trifoliate leaves from each plot once a week. The leaf area
before and after cutting diseased leaf tissue was determined. The leaf
samples were taken from the same general position in the canopy at each
sampling.
For the defoliation treatments, plants were manually defoliated
with pairs of scissors. The foliage was removed from the distal end of
the petiole. Defoliation was done only once, at full bloom because
results from other workers indicated that this was a critical growth
stage for beans (Hohmann and DeCarvalho, 1983).
Data were subjected to analysis of variance and regression analysis
using the general linear models procedure of SAS (Ray, 1982).


Proportion of Loss in Yield
90
Maximum Proportion of Foliage Infected
Figure 17. The influence of the maximum proportion of foliage infected
by Uromyces phaseoli on snap bean yield.


185
McGregor, W. C., D. R. Hansen, and A. I. Magee. 1953. Artificial
defoliation of field beans. Can. J. Agrie. Sci. 33: 125-131.
McKenry, M. V. 1983. Soil environment and nematode damage to plants.
J. Nematol. 15: 484 (Abstr.).
McMillan, R. T., Jr. 1972. A new race of bean rust on pole beans in
Florida. Plant Dis. Rep. 56: 759-760.
McMillan, R. T., Jr., G. Ellal and H. H. Bryan. 1982. Fungicides for
the control of squash powdery mildew and bean rust. Proc. Fla.
State Hort. Soc. 95: 304-307.
McSorley, R. and J. L. Parrado. 1984. Nematode persistence after
fumigation: A methodological problem. J. Nematol. 16: 209-211.
McSorley, R. and K. Pohronezny. 1984. Cost effectiveness of nematode
control by fumigation with SMDC on Rockdale soils. Proc. Soil and
Crop Sci. Soc. Fla. 43: 188-192.
McSorley, R., K. Pohronezny and W. M. Stall. 1981. Aspects of nematode
control on snap bean with emphasis on the relationship between
nematode density and plant damage. Proc. Fla. State Hort. Soc.
94: 134-136.
McSorley, R. and V. H. Waddill. 1982. Partitioning yield loss on
yellow squash into nematode and insect components. J. Nematol. 14:
110-118.
Meiners, J. P. 1974. International cooperation on bean rust research.
Annu. Rept. Bean Improv. Coop. 17: 55-57.
Meiners, J. P. 1977. Sources of resistance to U.S. bean rust popula
tions. Annu. Rept. Bean Improv. Coop. 20: 82-83.
Melakberhan, H., J. M. Webster, and R. C. Brooke. 1983. Time related
effects of Meloidogyne incognita on Phaseolus vulgaris. J.
Nematol. 25: 484 (Abstr.).
Mendgen, K. 1973. Feinbau der infektionsstrukturen von Uromyces
phaseoli (Electronmicroscopy of the bean rust infection
structures). Phytopath. Z. 78: 109-120.
Mendgen, K. and R. Heitefuss. 1975. Micro-autoradiographic studies on
host-parasite interactions. I. The infection of Phaseolus vulgaris
with tritium-labelled uredospores of Uromyces phaseoli. Arch.
Micribiol. 105: 193-199.
Metcalf, R. L. 1975. Insecticides in pest management, pp. 235-273
In: Introduction to Insect Pest Management, R. L. Metcalf and W.
H. Luckmann (eds.), Wiley, New York.


191
Teng, P. S., R. C. C. Close, and M. J. Blackie. 1979. Comparison of
models for estimating yield loss caused by leaf rust (Puccinia
hordei) on Zephyr Barley in New Zealand. Phytopathology 69: 1239
1244.
Thomas, R. J. and C. A. Clark. 1983. Population dynamics of M.
incognita and R. reniformis alone and in combination, and in their
effects on sweet potato. J. Nematol. 15: 204-211.
Thomason, I. J., D. C. Irwin, and M. J. Garber. 1959. The relationship
of the root-knot nematode, M. javanica to Fusarium wilt of cowpea.
Phytopathology 49: 602-606.
Thomason, I. S. and M. McKenry. 1975. Chemical Control of Nematode
Vectors of Plant Viruses, pp. 423-439 In: Nematode Vectors of
Plant Viruses. F. Lamberti, C. E. Taylor and J. W. Seinhorst
(Eds.).
Thorne, G. 1961. Principles of Nematology. McGraw Hill, New York.
552 p.
Todd, J. W. and L. W. Morgan. 1972. Effects of hand defoliation on
yield and seed weight of soybeans. J. Econ. Entomol. 65: 567-570.
Townsend, G. R. 1939. Diseases of beans in South Florida. Agrie.
Expt. Sta. Bull. 336. 60 pp.
Townsend, G. R. 1947. Diseases of beans in South Florida. Agrie.
Expt. Sta. Bull. 439. 56 pp.
Tyler, J. 1933. Development of root-knot nematodes as affected by
temperature. Hilgardia 7: 391-415.
Van Gundy, S. D., J. D. Kirkpatrick and J. Golden. 1977. The nature
and role of metabolic leakage from root-knot nematode galls and
infection b Rhizoctonia solani. J. Nematol. 9: 113-121.
Vargas, E. 1969. Determinacin de razas fisiolgicas de la roya del
frijol en Nicaragua y Honduras, en la primera siembra de 1968. In:
reunion Anual del Programa Cooperativo Centroamericano para el
Mejoramiento de cultivos Alimenticios 15a, San Salvador, El
Salvador.
Vargas, E. 1970. Determination de la razas fisiolgicas de la roya del
frijol en Nicaragua y Honduras en la segunda siembra de 1968. In:
Reunion Anual del Programa Cooperativo Centroamericano apra Mejora
miento de Cultivos Alimenticios 16a, Antigua, Guatemala.
Vargas, E. 1971. Determinacin de la razas fisiolgicas de la roya del
frijol en El Salvador. In: Reunion Anual del programa Cooperativo
Centroamericano para el Mejoramiento de cultivos Alimenticios 17a,
Panama.
Vargas, E. 1980. Rust, pp 17-36 In: Bean Production problems, H. F.
Schwartz, and G. E. Galvez (eds.). CIAT, Cali, Colombia.


Log (Yield)
39
Figure 3. Effects of defoliation and time of defoliation on snap
bean yield under field conditions (linear models).
Letters represent the plant growth stage defoliation occurred.
A = primary leaf, B = first trifoliate leaf, C = thrid trifoliate
leaf, D = flower bud formation, E = full bloom, and F = pod set.


99
There are many methods of determining disease severity in the
field. These methods include the use of leaf area meters, the Horsfall-
Barratt disease rating system, and pictorial keys. In these trials it
has been shown that the use of leaf area is a more convenient method for
disease severity assessment. Hence, in order to determine disease
severity the grower or extension specialist would have to take random
leaf samples at specific intervals and estimate the proportion of
foliage infected at each stage. Leaf area determination would be
facilitated if a leaf area meter were available but where this was not
the case then graph or trace paper could be used. When computers are
accessible to the grower/extension specialist then AUDC can be deter
mined based on disease severity. The choice of the variable would
depend on availability of expertise and equipment.
Yields were inversely related to disease parameters at specific
bean plant growth stages. Rate of disease progress, AUDC and disease
severity correlated well with yield. Therefore, the extension worker or
grower can choose the parameter of disease to measure. These disease
parameters are, however, time consuming which may increase labor and
other costs.


57
In the experiment involving simultaneous nematode inoculation and
defoliation, plants were defoliated manually with a pair of scissors.
The correct number or proportion of leaves to be removed was determined
by leaf counts at each plant growth stage. To eliminate additional
(R)
uncontrolled defoliation, plants were sprayed with Ambushv (40 g
ai/ha) for bean leaf roller (Urbanus proteus L.) and cowpea curculio
00
(Chalcodermus aeneus Boh.) control; Trigardv (150 g ai/ha) for
(R)
leafminer (Liriomyza spp.) control, and Benlatev (550 g ai/ha) for
disease control. These pesticides applied at 14-day intervals. Yield
was taken from all six plants. Pods less than 7 cm in length and
diseased or damaged ones were discarded.
Yield data were subjected to analysis of variance and regression
analysis using the general linear models procedure of SAS (Ray, 1982).
Nematode inoculation data were also analyzed using Seinhorst's models
(Ekanayake and Di Vito, 1984; Ferris, 1984; Ferris et al. 1981; Sein-
horst, 1965).
Results
Meloidogyne incognita alone
Analysis of variance on the effect of Meloidogyne incognita popula
tion levels on yield showed a significant relationship with F = 26.2***
(Table 8). Regression analysis of the data produced models of the form
2
Y = a + bx or Y = a + bx + cx where Y = yield (g/plot) or log (yield),
x = log (M. incognita population + 1) (Table 9). Quadratic models
o
consistently gave somewhat higher coefficients of determination (R )
values. The predictive ability of the quadratic models was, however,


Gross dollar value
44
Figure 6
1000
800
600
400
200
high
Price range medium
low
25 50 75
Defoliation Level
Influence of defoliation on gross dollar value per
hectare of 'Sprite' snap beans defoliated at the
third trifoliate leaf stage in the greenhouse.


TABLE 3. Regression equations for the relationship between yield and defoliation
Plant Growth Stage Greenhouse Field
Linear
Primary leaf
y
=
127.5
- 76x
R2 =
0.48
y
=
1525 -
805x
R2 = 0.41
(y
=
2.11 -
0.39x)a
(R2 =
0.49)
(y
=
3.18 -
0.35x)
(R2 = 0.34)
y
=
108.1
- 4.64x
R2 =
0.22
y
=
1197 -
686x
RZ = 0.16
First trifoliate
leaf
(y
=
2.01 -
0.24x)
(R2 =
0.22)
(y
=
3.08 -
0.62x)
(R2 = 0.25)
y
=
119.5
- 57.8x
R2 =
0.35
y
=
1420.4
- 922x
RZ = 0.59
Third trifoliate
leaf
(y
=
2.08 -
0.31x)
(R9 =
0.38)
(y
=
3.2 0.55x)
(R2 = 0.58)
y
=
117 -
64.5x
R2 -
0.42
y
=
1286.5
- 863x
RZ = 0.61
Flower bud formation
(y
=
2.07 -
0.36x)
(R, =
0.42)
(y
=
3.14 -
0.54x)
(R2 = 0.53)
y
=
115.9
- 55.9x
E2 *
0.30
y
=
1266.2
- 999x
RZ = 0.66
Full bloom
(y
=
2.05 -
0.31x)
(R2 "
0.26)
(y
=
3.36 -
1.47x)
(R2 = 0.46)
y
=
125.2
- 76.2x
R2 =
0.51
y
=
1441 -
1390.3x
R2 = 0.73*
Pod set
(y
=
2.13 -
0.48x)
(R2 =
0.51)
(y
=
3.25 -
1.06x)
(R = 0.81)
Quadratic
Primary leaf
y
=
129.2
- 89.3x
<- 13.3x2
2
R2
=
0.48
y
=
1446 -
173.6x -
631.4x2
R2
=
0.43
(y
=
2.07 -
0.14x -
0.25xZ)
cr;
=
0.22)
(y
=
3.15 -
0.08x 0
.26x2)
(r;
=
0.36)
y
=
117 -
117.3x +
70.9x2
K
=
0.27
y
=
1005.6
+ 845.4x
- 1531.4x
R2
=
0.24
First trifoliate
leaf
(y
=
2.05 -
0.54x +
0.29x2)
(Ro
=
0.25)
(y
=
2.85 +
1.16x 1
. 78x2)
(r;
=
0.43)
y
=
117.3
- 39.9x
- 17.8xZ
R2
=
0.34
y
=
1269.8
+ 283.7x
- 1205.7x
R2
=
0.68*
Third trifoliate
leaf
(y
=
2.05 -
O.Olx -
0.23xZ)
(Ro
=
0.41)
(y
=
3.07 +
0.52x 1
.07x2)
CR,
=
0.77)
y
=
115.7
- 53.5x
- llxZ
R2
=
0.43
y
=
1284.7
- 848.7x
- 14.3xZ
r7
=
0.61
Flower bud formation
(y
=
2.03 -
0.06x -
0.3xZ)
(R,
=
0.45)
(y
=
3.1 (
D.17x 0.
37x2)
(R
=
0.55)
y
=
116.3
- 58.9x
- 3x2
R2
=
0.29
y
=
1129.8
+ 91.lx -
1090.9x
R2
=
0.73*
Full bloom
(y
=
2.03 -
0.16x -
0.15x1
(R2
=
0.27)
(y
=
2.94 +
1.86x 3
32x2)
(Ro
=
0.67)
y
=
114.A
+ 10.7x
- 86.9x2
R2
=
0.55
V
=
1552
2282.6x +
892.3x
R2
=
0.87*
Pod set
(y
=
2.02 -
0.38x -
0.86x )
(R2
=
0.65)
(y
=
3.17 -
0.42x 0
.64x )
(R2
=
0.84)
Figures in parentheses are y = log (yield); x = defoliation as proportion.
* R significant at 0.05


96
was no benefit from spraying the plants more often than 7-day intervals.
By spraying at 4-5-day intervals there was a net loss in returns
compared to the 7-day intervals. The total cost per spray occasion per
hectare was $11.29.
In trial 2, there were increases in net returns, by spraying plants
at 7-day intervals compared to the 14-day spray schedule, of $1246,
$670, and $350 at the high, medium, and low prices. From the 14-day to
the 4-5 day spray schedules, there were increases in net returns of
$1264, $670, and $340 at the high, medium, and low prices. When plants
were sprayed at 7-day and 4-5-day intervals there was an increase in net
returns of $22 and $10 at the high and low prices but there was no net
increase in returns at the medium price. Thus, in trial 2 there was no
apparent benefit in spraying plants at 4-5-day intervals. The
additional sprays hardly paid for the extra cost of spraying more often.
Discussion
Significant relationships were found between yield and both disease
severity and area under the disease progress curves. Yields were
generally higher in trial 1 than in trial 2. The disease was clearly
established earlier in trial 2 than in trial 1.
The relationship between yield or loss in yield and bean rust was
well described using both parameters: maximum severity and area under
the disease progress curve (AUDC). Rate of disease progress was as well
2
correlated with yield as disease severity or AUDC in both trials with R
of 0.71 and 0.79 for trial 1 and trial 2 respectively. In trial 2,
o
disease severity and AUDC had coefficients of determination (R ) of 0.98


TABLE 22. Effect of metam-sodium on snap bean yield and populations of nematode genera in midseason
soil samples. Data are means of 3 replicates.
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus Meloidogyne Rotylenchulus Tylenchorhynchus Yield
(No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml) (g/plot)
0 0
47 0
94 2
187 0
281 0
374 0
3 12 0
8 8 0
4 0 1
1 7 0
3 110
1.3
3
1
4
8
0
1
5
2
360
596
650
680
581
707
110


73
Gross dollar values are shown in Table 15 and Figure 12. These
values were computed from gross yield per hectare based on the following
price ranges $6.00 (low), $11.00 (medium), and $20.00 (high). Hence
these are gross values without deducting production costs. The gross
dollar values (Table 15) show that there was a wide range at each plant
growth stage. Generally highest dollar values were obtained when the
plants were nematode-free and when no defoliation occurred. The combi
nation of nematodes and defoliation had inconsistent effects on gross
dollar values. If defoliation were held constant at any level, gross
dollar values decreased as the nematode populations were increased
(Table 15). If nematode populations were held constant the decrease in
gross dollar values was not always consistent with the levels of defoli
ation. This is shown clearly at the primary leaf, first trifoliate
leaf, full bloom and pod set stages of plant growth. At these stages
some lower levels of defoliation have smaller dollar values than higher
defoliation levels. As expected, loss in dollar values was similar to
yield loss since the former was obtained from yield (g/plot), but varies
greatly depending on the current market price which can fluctuate
widely.
Discussion
There were significant differences in yield when M. incognita
inoculated to snap beans. In this test 10 eggs and juveniles/pot
reduced yield by at least 19%. In the defoliation M. incognita
interaction test, 1,000 eggs and juveniles/pot depressed yield by only
11-28% when plants were not defoliated (Table 14). Similar trends


TABLE 11. F-values and probability levels from analysis of variance for the effects of defoliation and M.
incognita and their interaction on snap bean yield.
Plant Growth Stage
Source
Primary First tri- Third tri- Flower bud Full
leaf foliate leaf foliate leaf formation bloom
Pod
set
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
F
Prob.
F
Defoliation
17.67
0.0001
67.84
0.0001
66
0.0001
41.6
0.0001
100
0.0001
140
0.0001
Log (M. incognita
population +1)
23
0.0001
23.5
0.0001
35
0.0001
38
0.0001
28.95
0.0001
56.15
0.0001
Defoliation x
Log (M. incognita
population +1)
0.56
0.8871
0.54
0.8395
1.5
0.0962
0.72
0.7396
1
0.4601
0.38
0.9575


LITERATURE CITED
Acland, J. D. 1971. East African Crops and Introduction to the Produc
tion of Field and Plantation Crops in Kenya, Tanzania and Uganda
Longman, London. 252 pp.
Agudelo, F. V. De. 1980. Nematodes, pp 315-326 In: Bean Production
Problems. H. F. Schwartz and G. E. Garvez (Eds). CIAT, Cali,
Colombia.
Allen, D. J. 1983. The Pathology of Tropical Food Legumes; Disease
Resistance in Crop Improvement. Wiley and Sons, New York. 413 pp.
Almeida, A. M. R., G. M. Chaves, and L. Zambolim. 1977. Influcia da
poca de ataque de Uromyces phaseoli typica Arth. sobre o
rendimento de duas variadades de feijoeiro (Phaseolus vulgaris L.)
en casa-de-vegetacao. Fitopatologia Brasileira 2: 17-21.
Andersen, A. L. 1975. Bean rust. Extension Bull. E-893, Mich. State
Univ. East Lansing 2 pp.
Anonymous. 1972. Vegetables: Annual Summary; Acreage, Yield and
Value. C.R.B. Statistical Reporting Service, USDA, Washington DC.
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Value. C.R.B. Statistical Reporting Service, USDA, Washington DC.
Anonymous. 1982. Vegetables, Summary: Florida Agrie. Statistics.
Fla. Crop and Livestock Reporting Service. IFAS-USDA, Orlando.
Augustin, E., D. P. Coyne, and M. L. Schuster. 1972. Inheritance of
resistance in Phaseolus vulgaris to Uromyces phaseoli typica
Brazilian rust race Bll and of plant habit. J. Am. Soc. Hort. Sci.
97: 526-529.
Ayala, A. and C. T. Ramirez. 1964. Host range, distribution, and
bibliography of the reniform nematode, Rotylenchulus reniformis,
with species reference to Puerto Rico. J. Agrie. Univ. Puerto Rico
48: 140-161.
Ballantyne, B. J. 1974. Resistance to rust in beans. Annu. Rept. Bean
Improv. Coop. 17: 19-20.
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differential varieties and a standard nomenclature of races. Proc.
Bean Rust Workshop, Oct. 1974, CIAT, Cali, Colombia.
176


103
bud formation. Metam-sodium was used to manipulate nematode
populations. Subsequent weeding was done by cultivation. Irrigation
was provided by an overhead sprinkler system.
The effect of metam-sodium on snap beans
The crop was planted on 26 November 1984. Individual plots consis
ted of 3 rows 3m long with 0.91m between rows. Seeds were planted at
8-10 cm spacing. Treatments were replicated 4 times in a randomized
complete block. Metam-sodium was applied preplant at 0, 47, 94, 187,
281, and 374 L/ha. Preplant soil samples consisted of a composite
mixture of 10 soil scoops (to a 6-8 cm depth) from each plot 12 days
after fumigation. The 12-day period was based on the observations made
by McSorley and Parrado (1984). Aliquots of 100 ml soil were processed
by sieving and centrifugal flotation (Jenkins, 1964). Subsequent soil
samples at mid-season and harvest were taken from the root zone and
similarly extracted. Only live nematodes were counted in the preplant
samples but in the later samples, nematodes were first killed by gentle
heating in a water bath (55-60C) and counted. Beans were harvested on
31 January 1985.
Yield data were subjected to analysis of variance and regression
analysis using the general linear models procedure of SAS (Ray, 1982).
The effect of metam-sodium and defoliation on snap beans
The crop was planted on 26 November 1984. Individual plots consis
ted of 4 rows 3 m long with 0.91 m between rows. Seeds were planted at
8-10 cm spacing within the row. A split-plot design was used in this
trial to investigate the effect of metam-sodium (main plots) and defoli
ation (subplots) on snap beans. Treatments were replicated 4 times.
Each subplot consisted of a 3 m long row. Defoliation levels


Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fullfillment
of the Requirements for the Degree of
Doctor of Philosophy
MULTIPEST ECONOMIC THRESHOLDS ON SNAP BEANS
By
Afete Divelias Gadabu
May, 1986
Chairman: Dr. Van H. Waddill
Major Department: Entomology and Nematology
During 1984 and 1985, a number of greenhouse and field experiments
were carried out at the Tropical Research and Education Center at
Homestead in Dade County, Florida, to determine the effect of manual
defoliation, Meloidogyne incognita, bean rust, and various other nema
todes on Sprite' snap beans (Phaseolus vulgaris L.). Treatments
consisted of total defoliation (100%), 0%, 25%, 50%, and 75% defoliation
at various plant growth stages; 0, 10, 100, 1,000, 10,000 and 100,000 M.
incognita eggs and juveniles per pot, fungicide sprays which included
bitertanol at 7-day intervals, mancozeb tank-mixed with sulfur at
4-5-day, 7-day, and 14-day intervals respectively, and soil fumigation
with metam-sodium at 0, 47, 94, 187, 281, and 374 L/ha and a separate
one at 935 L/ha. Experiments were conducted with each series of
treatments as well as with combinations of two or more types of
treatments simultaneously.
vi


I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
V. H. Waddill, Chairman
Professor of Entomology
and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
J. R. Strayer, Co-Chairman
Professor of Entomology
and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
R. T. McSorley
Associate Professor of
Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
K. L. Pohronezny
Associate Professor of Plant
Pathology


121
TABLE 33. Net returns on investment ($) per hectare of snap beans.
Plants were defoliated at various levels and soil treated
with metam-sodium.
Defoliation level Metam-sodium Net returns by price range
(proportion of foliage) (Liters/ha) Low Medium High
0
47
- 27
- 25
- 21
0
94
34
112
255
0
187
29
154
379
0
374
-229
-220
-203
0.25
0
- 37
- 68
-124
0.25
47
- 93
-145
-238
0.25
94
- 35
- 14
- 23
0.25
187
-190
-247
-351
0.25
374
-300
-350
-440
0.50
0
-137
-252
-458
0.50
47
-173
-292
-507
0.50
94
-145
-217
-345
0.50
187
-225
-313
-470
0.50
374
-343
-429
-583
0.75
0
-138
-253
-460
0.75
47
-206
-352
-616
0.75
94
-192
-301
-499
0.75
187
-292
-436
-694
0.75
374
-343
-475
-666


174
except for total defoliation (Table 43). The unsprayed plots gave
yields comparable to those obtained from fortnightly mancozeb-sprayed
plants. There was no apparent difference between the 4-5-day and 7-day
mancozeb spray schedules (Table 43).
Gross dollar values per hectare of snap beans are shown in Table
44. Bitertanol-sprayed plots consistently gave the highest dollar
values at all defoliation levels and both growth stages. The gross
dollar values were computed from the following price range, $6.00 (low),
$11.00 (medium), and $20.00 (high) per bushel (13.62 kg/) of snap beans
multiplied by the yield per hectare. Net income (Table 45) was derived
from the no spray value as the base line. This was deducted from the
values of the other treatments and the cost of fungicide sprays deducted
from this difference. Bitertanol is an experimental fungicide which has
not been registered for use on beans, hence the price is not known.
Thus, the net value of beans for this fungicide excludes its cost.
Consequently, the net values for bitertanol may not be a true
reflection. Under the conditions this study was conducted, the grower
would have made a profit if he sprayed his crop with mancozeb at 4-5 day
intervals even if 35% or 50% of the leaf area were removed at the
primary leaf stage (Table 45). There was no positive effect on net
income from fungicide sprays if plants were defoliated at the flower bud
formation stage.
Discussion
There were significant differences among fungicide spray schedules
based on yield. Bitertanol gave the highest yield and by derivation the


81
infected leaves were clipped on to wire stakes 25 cm above ground level.
Two stakes were placed in each plot, one on each end, depending on
general wind direction. Disease progress was monitored by taking
trifoliate leaves at random from each plot once a week. At each
sampling occasion leaves were taken from the same relative level within
the canopy. Disease severity (proportion of leaf area infected by the
a_b
disease) was determined using the mathematical model y = where y =
disease severity, a = area of leaf before cutting out diseased tissue,
and b = area of leaf after cutting out diseased tissue. The mean of the
5 trifoliate leaves was the measure used in the final data. Leaf area
(R)
was determined by a LiCorv portable area meter (Model LI-300, Lombdar
Instruments Corp).
Disease progress curves are generally sigmoid in shape (Imhoff et
al., 1982a). The generalized progress curve is shown in Figure 14.
Progress curves of bean rust in these studies were obtained by determi
ning disease severity at weekly intervals as indicated above.
Area under the disease progress curve was computed using the general
model: y = £ [ [tl + 1 l1 ln "hich ? area under
the disease progress curve, x = disease severity at the i observation,
t^ = time (days) at the i*"*1 observation, and n = total number of obser
vations. The computations were facilitated by the use of a computer
program provided by Dr. R. D. Berger. The computer program employed the
following model: y = (((n (x) + n (x + l))/2) t (x)) where y = area
under the disease progress curve, x = disease severity, n = number of
disease severity values, and t = time (days) at which observation is
made. The rate of disease progress was determined by using the Gompertz
model which consistently gave better fit to the data (Berger, 1981).


15
nutrients from the plant. Plant cells in the vicinity of the nematode
juvenile increase both in number and size (hyperplasia and hypertrophy),
causing the characteristic giant cells (synctia) (Dropkin, 1980; Taylor
and Sasser, 1978). The giant cells usually form near the juvenile's
head by the fusion and enlargement of plant cells in response to
nematode feeding. These giant cells eventually become apparent in the
form of galls on the root system. Injury to plant root systems usually
becomes apparent 10 days after infection. Five to six weeks after
infection, epidermal cells of the roots collapse after females have
deposited eggs near the outer root surface (Ngundo and Taylor, 1975a).
Control of Root-knot Nematodes, Meloidogyne spp.
The economic importance of plant-parasitic nematodes is commonly
assessed by the use of soil fumigants (Mountain, 1965). Usually, an
inverse relationship between yield and nematode numbers is expected
(Sasser et al., 1968). The relationship between yield and nematode
counts is not always inverse (Robbins et al., 1972). In many bean-
producing regions, nematicides are extensively used on a preventative
basis (Agudelo, 1980). The world farming community has many nematicides
available depending on supply and legal registration. These nematicides
include dicholoropropene-dichloropropane (DD), ethylene dibromide (EDB),
phenamiphos, methyl bromide, aldicarb, metam-sodium, and DBCP (Jimenez,
1976; Parisi et al., 1972; Rhoades, 1976; Sosa Moss and Wrihs, 1973).
In these operations, no attempt is made to eradicate nematodes (Thomason
and McKenry, 1975). These nematicide applications are aimed at reducing
the nematode populations by 80-90% in the upper 40-60 cm of the soil and
are considered adequate to provide economic control (Thomason and
McKenry, 1975).


TABLE 20. The relationship between disease severity and net return per hectare of snap beans
sprayed with mancozeb and sulfur.
No. of
sprays
Fungicide
spray frequency
Maximum
Proportion of
foliage infected
Price range
Net return ($)
in investment
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0
0
0
0.76
0.86
high
0
0
medium
0
0
low
0
0
3
14-day
14
0.65
0.71
high
743
564
medium
395
295
low
199
145
5 (6) a
7-day
7
0.46
0.47
high
1812
1810
medium
973
965
low
505
495
7(8)
4-5-day
4-5
0.098
0.4
high
1644
1828
medium
870
965
low
438
485
a
Number of sprays in trial 2 are in parentheses.


154
Log10(lnitial nematode density+1)
Relation between log^g (initial nematode density +1)
and yield.
Figure 33.


147
0.525% solution gave the best overall hatch of nematode eggs extracted
from both bean and tomato roots (Figure 30 and 31).
The percentage hatch of M. incognita eggs is shown in figure 32.
The lowest percentage hatch was obtained from the 2.6% NaOCl-extracted
eggs. Water-extracted eggs had a percentage hatch comparable to that of
the 0.13, 0.26 and 0.525% NaOCl-extracted eggs. The 0.525% NaOCl-
extracted eggs had the highest proportion of eggs hatched among all
treatments by day 8, leading to the choice of the 0.525% NaOCl-extracted
eggs in the inoculation of bean plants and maximum yield of eggs over
time as above.
The relationship between initial M. incognita densities and snap
bean yield is shown in figure 33. There was a significant (P = 0.01)
negative correlation between nematode densities and yield for all three
inoculation systems (correlation coefficients were -0.96, -0.78 and
-0.96 for seed drench, soil mix and gall inoculation). As the nematode
densities increased, the snap bean yield decreased. The lowest yield
was obtained from plants inoculated with 500 galls/pot (= 123,000
eggs/pot). Negative impacts on yield were greater in seed-drench
inoculated pots than soil-mix inoculated ones (Figure 33). There were
significant differences in yield among nematode densities in the
seed-drench and gall inoculation methods with F values of 14.34** and
26.18**, respectively. There were, however, no significant differences
in yield among nematode densities in the soil-mix inoculation method
with an F value of 3.09.
Gall indices were comparable in all three inoculation methods as
shown in Table 41. Among inoculated plants, the lowest gall index was
observed on plants inoculated with 10 eggs and juveniles per pot.


CHAPTER VII
THE EFFECT OF INOCULATION METHOD AND INITIAL POPULATION DENSITY OF
MELOIDOGYNE INCOGNITA (KOIFOID AND WHITE) CHITWOOD ON SNAP BEANS
(PHASEOLUS VULGARIS L.) 'SPRITE'
Introduction
The root-knot nematode Meloidogyne incognita (Kofoid and White)
Chitwood poses a serious threat to bean (Phaseolus vulgaris L.) produc
tion in many bean growing areas of the world (Agudelo, 1980; Allen,
1983; Ngundo, 1977; Singh et al. 1981a; Sharma and Guazelli, 1982).
Meloidogyne incognita infections have been reported to decrease the
apparent photosynthetic rate of vulgaris 'Topnotch Golden Wax' as
well as other physiological growth factors (Melakberhan et al., 1983).
The limitation on bean production by root-knot nematodes may be due to
root galling which interferes with nitrogen fixation by Rhizobium spp.
and also interference with nutrient uptake. Yield losses of 50-90% have
been reported from fields infested with root-knot nematodes (Agudelo,
1980; Freire and Ferraz, 1977; Ngundo, 1977; Varn and Galvez, 1974).
Damage functions ascribable to nematodes are influenced by many
factors (McKenry, 1983). Nematode management decisions should, there
fore, be based on environmental factors and the crops grown (Ferris,
1980). Environmental factors such as soil temperature, texture, and
structure, and water infiltration rates influence moisture regimes of
soil profiles which in turn affect nematode damage functions (McKenry,
1983). Noe and Barker (1983) related 24 edaphic variables to the field
distribution of Meloidogyne spp. Work done on the influence of these
142


146
October 1984 and root gall indices determined following the method
outlined by Taylor and Sasser (1978).
Yield data were subjected to regression analysis using the general
linear models of SAS. Seinhorst model curve fitting was also attempted.
Results
The relationship between NaOCl concentration and log^ (number of
eggs and juveniles + 1) extracted is shown in figure 29. Data fit the
equations Y = a + b LnX or Y = a where Y = log (number of eggs and
juveniles + 1), X = concentration (%) of NaOCl, and a and b are con-
2
stants. The coefficient of determination (R ) values were 0.69* to
0.75* for the bean and tomato curves, respectively. Figure 29 shows
that the number of eggs and juveniles extracted increased more rapidly
at low NaOCl concentrations (0 to 0.26%) and levels off at high
concentrations (0.525-2.6%).
Table 40 shows the number of Meloidogyne incognita eggs and juve
niles extracted form 120 g of infected plant roots at various NaOCl
concentrations. These eggs and juveniles were extracted from bean and
tomato roots 63 and 50 days after inoculation respectively.
Figures 30 and 31 show the total number of juveniles which emerged
from eggs extracted at various NaOCl concentrations. On the day of
extraction, sodium hypochlorite-extracted eggs hatched more than the
water-extracted eggs. The highest number of juveniles had emerged from
eggs extracted with the 0.525% NaOCl solution from bean roots over the
next 6 days after extraction (Figure 30). Eggs extracted from tomato
roots had a different hatch trend from that of beans (Figure 31). The


178
Carter, W. W. 1975a. Effects of soil temperature and inoculum levels
of Meloidogyne incognita and Rhizoctonia solani on seedling disease
of cotton. J. Nematol. 7: 229-233.
Carter, W. W. 1975b. Effects of soil texture on the interaction
between Rhizoctonia solani and Meloidogyne incognita on cotton
seedling. J. Nematol. 7: 234-236.
Carvalho, J. M., S. Ferraz, and A. A. Cardoso. 1981. Bean seed
dressing with oxamyl dissolved in acetone or ethanol for the
control of nematodes. Revista Ceres 28: 580-587.
Castillo, M. B. and J. A. Litsinger. 1978. Plant parasitic nematodes
of mung beans in Philippines, pp. 195-200 In: Proc. 1st
Internat. Mungbean Symp. Asian Vegetable Research and Development
Center, Taiwan.
Cauquil, J. and R. L. Shepherd. 1970. Effect of root-knot nematode-
fungi combinations on cotton seedling diseases. Phytophathology
60: 448-451.
Caveness, F. E. 1967. Nematology studies. Nigeria Ministry of
Agriculture and Natural Resources, Western Region, Lagos. 135 pp.
Caveness, F. E., R. M. Gilmer, and R. J. Williams. 1975. Transmission
of cowpea mosaic virus by Xiphinema basiti in Western Nigeria, pp.
289-290 In: Nematode Vectors of Plant Viruses. F. Lamberti, C.
E. Taylor and J. W. Seinhorst (eds.). Plenum Press, London and New
York.
Chitwood, B. G and M. B. Chitwood. 1950. An introduction to
nematology. University Park Press, Baltimore, Maryland. 334 pp.
Christie, J. R. 1959. Plant Nematodes; their Bionomics and Control.
H. and W. B. Drew, Jacksonville. 256 pp.
Centro Internacional de Agricultura Tropical. 1983. Rust. pp. 39-40
In: Bean Program Annu. Report for 1983. Cali, Colombia.
Cohen, Y. and J. Rotem. 1970. The relationship of sporulation to
photosynthensis in some obligatory and facultative parasites.
Phytopathology 60: 1600-1604.
Cook, A. A. 1978. Diseases of Tropical and Subtropical Vegetables and
Other Plants. Hafner Press, New York. 381 pp.
Costa, A. S. 1972. Anais do I Simposio Brasileiro de Feijao. pp.
311-316. Universidade Federalde Vicosa, Minas Gerais, Brazil.
Costa, C. L. and C. J. Rossetto. 1972. Investigacoes sobre pragas de
feijo eiro no Brasil. Anais do I Simposio Porasileiro de Feijao.
Campinas, 22-29 August, 1971, 29. Vol. Impr. Univ. Vicosa, Minas
Gerais, Brazil.


Log (Yield + 1)
38
2.5
2.0
1.5
1.0
0.5
0
Figure 2.
A Primary leaf stage
B First trifoliate leaf
C Third trifoliate leaf
Effects of defoliation and time of defoliation on snap
bean yield in the greenhouse (quadratic models).
Letters represent the plant growth stage defoliation occurred.


187
Ogle, H. J. and J. C. Johnson. 1974. Physiologic specialization and
control of bean rust (Uromyces appendiculatus) in Queensland. J.
Agrie. Sci. 31: 71-82.
Parisi, C. R. A., C. J. Torres and C. Sosa Moss. 1972. Incorporacin
de una nematicida sistemico a la planta de frijol por inmersin de
semillas. Nematropica 2: 22.
Pereira, A. A. and G. M. Chaves. 1977. Differential varieties and a
ternary system of nomenclature to designate races of Uromyces
phaseoli typica Arth. Annu. Rept. Bean Improv. Coop. 20: 85.
Pohronezny, K., J. Francis, and J. S. Reynolds. 1984. Efficacy of
selected fungicides against bean rust: a preliminary report on
alternative control programs compatible with Canadian fungicide
tolerances. Homestead TREC Res. Rept. SB84-1, 7 pp.
Pohronezny, K., V. H. Waddill, W. M. Stall, and W. Dankers. 1978.
Integrated control of the vegetable leafminer (Liriomyza sativae
Blanchard) during the 1977-78 tomato season in Dade County,
Florida. Proc. Fla. State Hort. Sci. 91: 264-267.
Porter, D. M. and N. T. Powell. 1967. Influence of certain Meloidogyne
spp. in Fusarium-wilt development in flue-cured tobacco.
Phytopathology 57: 282-285.
Powell, N. T. 1971. Interaction of plant parasitic nematodes with
other disease-causing agents. Vol. II. pp. 119-136. In: Plant
Parasitic Nematodes, B. M. Zuckerman, W. F. Mai and R. A. Rohde.
(Eds.) Academic Press, New York.
Powell, N. T. and C. J. Nusbaum. 1960. The black shank-root knot
complex in flue-cured tobacco. Phytopathology 50: 899-906.
Raggi, V. 1978. Photorespiration, respiration, photosynthesis and
their correlation with the C0 compensation point in French bean
leaves mildly infected by rust (Uromyces phaseoli). Phytopathol.
Medit. 17: 105-109.
Ray, A. A. (ed.). 1982. SAS User's Guide: Basics. 1982 Edition. SAS
Institute Inc., Cary, North Carolina. 921 pp.
Raymundo, A. and A. L. Hooker. 1981. Measuring the relationship
between northern corn leaf blight and yield losses. Plant Dis.
Rep. 65: 325-327.
Rey, G. J. V. and T. C. J. Lozano. 1961. Estidies fisiolgicas de la
roya del frijol (Phaseolus vulgaris L.) causadado por el Uromyces
phaseoli var. typica Arth. Acta Agron., Palmira 11: 147-186.
Reynolds, H. W. and R. G. Hanson. 1957. Rhizoctonia disease of cotton
in the presence or absence of the cotton root-knot nematode in
Arizona. Phytopathology 47: 256-261.


78
Uromyces phaseoli progress on artificially inoculated beans, vulgaris
'Bountiful', depended more on length and frequency of wetting periods
than on temperature (Imhoff et al., 1982a).
Yield loss due to disease has been observed to be proportional to
the area under the disease progress curve or proportional to disease
severity at some critical stage of host growth (Madden et al., 1981;
Raymundo and Hooker, 1981; Romig and Calpouzos, 1970; Shaner and Finney,
1977; Teng et al. 1979). In many of these studies, area under the
disease progress curve satisfactorily explained the relationship between
diseases and yield losses. Disease severity at one or more points in
time and rate of increase of the disease were also satisfactory disease
parameters employed to explain the relationship between disease and
yield loss (James, 1974; James and Teng, 1979; Main, 1977).
Berger (1981) compared the logistic and Gompertz models for disease
progress curve fitting. It was observed that the Gompertz model consis
tently gave better fit to the data examined than the logistic model for
disease severity values outside the 0.05 < y < 0.6 range (Figure 14).
The Gompertz model was superior to the logistic model in linearizing 113
selected disease progress curves (Berger, 1981).
Growers often resort to routine fungicide sprays for disease
control. Currently, weekly sprays with mancozeb are applied for disease
control on beans (McMillan et al., 1982; Pohronezny et al. 1984). The
effectiveness of these sprays depends on spray coverage and disease
severity but in many cases disease control is less than satisfactory
(McMillan et al. 1982). Usually, sprays are initiated before disease
signs and/or symptoms are observed on the crop.
The present studies were conducted to determine the effect of bean
rust on 'Sprite' snap beans under field conditions.


29
This study on snap beans (Phaseolus vulgaris L., 'Sprite') was
conducted to determine the plant growth stage most sensitive to defolia
tion, and the effects of defoliation on yield.
Materials and Methods
Two defoliation experiments were conducted in the summer and fall,
1984 in the greenhouse and field, respectively. Bush snap beans
(Phaseolus vulgaris L. 'Sprite') were grown at the Tropical Research and
Education Center in Homestead, Dade County, Florida. The crops were
grown on Rockdale soil (pH ca. 7.8). The greenhouse and field experi
ments were planted on 25 June 1984, and 24 October 1984, respectively.
Fertilizer (8:16:16) was applied at the rate of 448 kg/ha according to
the University of Florida Extension recommendations (Stall and Sherman,
1983). Benlate^ (550g ai/ha) was applied fortnightly for control of
(R)
certain diseases and sprays of Trigardv (150 g/ha) were applied at the
(R)
same frequency for leafminer (Liriomyza spp.) control. Ambushv (40 g
(R)
ai/ha) or Pydrinv (250 g ai/ha) was applied at 14-day intervals for
cowpea curculio (Chalcodermus aeneus Boh.) control.
Defoliation levels investigated were total (100%), 25%, 50%, and
75%. An undefoliated control was included. Plants were defoliated at
the primary leaf stage, first trifoliate leaf, third trifoliate leaf,
flower bud formation, full bloom, and pod set. Beans were harvested on
8-20 August 1984 and 18-26 December 1984. The harvest was not graded
since cowpea curculio feeding damage to pods was extensive.
Greenhouse experiment
Rockdale soil (3030 L) was fumigated with Dowfume^ MC2 (681 g) on
a cement slab under a tightly sealed polyethylene sheet. Number two


56
was planted on 25 June 1984 and harvested on 28 August 1984. The M.
incognita x defoliation study was planted on 24 October 1984 and
harvested on 26 December 1984 to 3 January 1985. In each experiment, a
plot comprised of 2 pots, each containing 3 plants. Irrigation was
provided by an automatic time-controlled, overhead, water-mist-producing
system twice a day.
The Meloidogyne incognita population used in each experiment was
originally obtained from Hausa potato (Coleus parviflorous Benth.) and
was maintained on greenhouse-grown tomato (Lycopersicon esculentum
Mill.) FloraDade' plants. Meloidogyne incognita eggs were extracted by
the sodium hypochlorite (NaOCl) method of Hussey and Barker (1973). A
(R)
0.525% NaOCl solution was made from Thrift King commercial bleach
(5.25% NaOCl) by dilution with cold tap water (25C). Tomato roots were
thoroughly washed of soil with running tap water. The clean roots were
cut into 2-3 cm long pieces and 120 g of the cut root material was
manually shaken in 200 ml of the NaOCl solution for 3.5 minutes. The
shaken material was serially filtered through 100-mesh, 230-mesh and
500-mesh sieves. The number of eggs and juveniles of M. incognita per
ml was determined by counting in a watch glass under a dissecting
microscope (20X). Appropriate dilutions of the nematode eggs and
juveniles were made according to the population levels used.
Plants were inoculated 10 days after planting by drenching their
bases with the nematode egg and juvenile suspension. The nematode
population levels of 0, 10, 100, 1,000, 10,000, and 100,000 per pot were
equivalent to 0, 0.16, 1.6, 15.6, 156 and 1562 eggs and juveniles per
100 ml soil, respectively.


101
(Tassi) Gold, Sclerotium rolfsii Sacc., Pythium spp., and Fusarlum spp.
have also been reported to initiate wilting and eventually defoliation
(Martinez, 1983). Many nematode species are found in association with
bean roots in various parts of the world (Agudelo, 1980, Allen, 1983).
The root-knot nematode complex (Meloidogyne spp.) is among the most
damaging on beans (Agudelo, 1980; Allen, 1983; Ngundo, 1977; Ngundo and
Taylor, 1974, 1975a,b). Thus, an understanding of the relationship
between these factors and yield is a prerequisite for the development of
a sound pest management strategy.
Information on the relationship between leaf damaging pests and
yield has been obtained through pest damage simulation by manually
defoliating plants at various growth stages and at several defoliation
levels (Edje and Mughogho, 1976a,b; Edje et al., 1972, 1973, 1976;
Galvez et al., 1977; Greene and Minnick, 1967; Hohmann and De Carvalho,
1983; Vieira, 1981; Waddill et al., 1984). Ruesink and Kogan (1975)
observed that manual defoliation is not precise in simulating pest
damage. The imprecision in pest damage simulation may be due to the
exact timing of manual defoliation and careful determination of the
proportion of the foliage to be removed, which pests cannot do. More
over, manual defoliation does not introduce saliva and possible phyto
toxins which may be important factors in the plant damage caused by
specific pests.
Rarely are crop plants attacked by one pest species only. More
often several species attack a crop at the same time. Thus, McSorley
and Waddill (1982) studied the effect of insect and nematode pests on
squash (Cucrbita pepo L.). They partitioned yield loss into insect and
nematode components by using multiple regression procedures. It was


TABLE 5. Relationship between defoliation, time and yield (loss (%)) of snap beans in the greenhouse
and field.
Yield (Z loss by plant growth stage
Defoliation
level (Z)
Primary
leaf
First
triboliate
leaf
Third
trifoliate
leaf
Flower
bud formation
Full bloom
Pod set
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
0
0
0
0
0
0
0
0
0
0
0
0
0
25
36
-3
31
29
22
16
30
14
22
0
21
29
50
34
19
41
20
21
12
30
48
35
39
16
68
75
39
23
32
-4
35
37
39
52
23
29
31
70
100
69
55
47
74
57
74
64
73
57
95
74
92


58
TABLE 8. Effect of M. incognita on snap bean yield. Data are means of
4 replicates.
No. of
M. incognita/Pot
Log (M. incognita
Population +1)
Yield (g/plot)a
Yield loss
(%)
0
0
128
0
10
1.0
103
19
100
2.0
79
38
1000
3.0
70
45
10000
4.0
68
47
100000
5.0
53
59
Data rounded off to the nearest whole number.
F = 26.2*** for M. incognita populations.


68
Figure 13: Effects of defoliation and M. incognita on snap
bean yield.


80
Materials and Methods
Two trials were conducted at the Tropical Research and Education
Center in Homestead, Dade County, Florida, on Rockdale soil (pH ca.7.8).
The first trial was planted on 27 February 1985 and the second on 21
March 1985. Beans were harvested on 25 April 1985 and 13 May 1985
respectively.
In both trials plots were 3 rows wide (0.91 m row spacing) and 3 m
long. Beans were planted 7-10 cm apart within the row. Prior to
planting the herbicides Treflan^ (841 g ai/ha) and Dual^ (1.7 kg
ai/ha) were applied to the site. Fertilizer (8:16:16) was applied at
448 kg/ha before planting. Plants were top dressed at 224 kg/ha just
(R)
before flower bud formation. The crops were sprayed with Ambushv (40
g ai/ha) fortnightly for cowpea curculio (Chalcodermus aeneus Boh.)
(R)
control. Slugs and snails were controlled by Mesurolv (200 g ai/ha)
pellets. Plants were irrigated using an overhead sprinkler system.
In both trials five treatments were arranged in a randomized
complete block design with four replications. Fungicides were used as a
tool to manipulate disease levels. Treatments used were (a) no fungi
cide; (b) bitertanol (57 g ai/ha) at 7-day intervals; (c) mancozeb (0.7
kg ai/ha) tank-mixed with sulfur (4.5 kg ai/ha) at 4-5-day intervals;
(d) same as (c) but at 7-day intervals; and (e) same as (c) but at
(R)
14-day intervals. All sprays were applied with Helenav sticker or Nu
(R)
Film-17v as a spreader/sticker. Bitertanol plots were virtually
disease free.
Plants were inoculated at the primary leaf growth stage using
infected pole bean leaves collected from abandoned bean fields. The


106
Results
Effect of metam-sodlum on snap beans
Tables 21, 22 and 23 show the numbers of nematodes in 100 ml
aliquots of soil at preplant, mid-season and harvest respectively. Two
nematode genera, Criconemella and Rotylenchulus, were found in the soil
at preplant (Table 21). At mid-season, four nematode genera were
detected in the soil: Helicotylenchus, Meloidogyne, Rotylenchulus, and
Tylenchorhynchus (Table 22). Meloidogyne had the highest numbers in
unfumigated plots (Table 22), but at this time, the effect of metam-
sodium on nematode population was not proportional to its rate of
application. Criconemella, Helicotylenchus, Meloidogyne, and
Rotylenchulus were found in the soil at harvest (Table 23).
Metam-sodium had no significant effects on nematode numbers at any
sampling (Tables 21, 22, and 23).
Analysis of variance on snap bean yield showed that there were
significant differences among metam-sodium rates (F = 3.4*). Yield
responses were, however, not consistently proportional to metam-sodium
rates (Table 21). Regression analysis on the relationship between
2
metam-sodium and yield produced models of the form Y = -0.004x + 345.75
(R2 = 0.17) and Y = -0.00004x3 0.03x2 + 4.9x + 287.5 (R2 = 0.22) where
Y = yield (g/plot), and x = metam-sodium rate (L/ha). The cubic model
gave a higher coefficient of determination than the quadratic model
(Figure 21). The coefficients of determination were very low. The
linear model of the form Y = a + bX gave an R of only 0.12 indicating
that yield was not linearly related to metam-sodium rate. Multiple
regression on nematode general effects on yield gave the model Y = a +
bl X1 + b2 X2 + '* + bn xn where Y = yield, x = log (nematode population


18
collection respectively (Taylor and Sasser, 1978). These two species
are known to have 2 host races each (Dickson, unpublished).
The Importance of Bean Rust
Bean rust is known to occur whenever beans are grown (Vargas,
1980). Bean rust is the most important bean disease in Central and
South America (Augustin et al., 1972; Crispin and Dongo, 1962; Makram et
al., 1973; Zaumeyer and Meiners, 1975). Bean rust has been reported to
reduce the yield of snap beans in New Zealand, Egypt, and Australia
(Ballantyne, 1974; Makram et al., 1973; Yen and Brien, 1960). Yields of
dry beans have been lowered by infections of bean rust in Kenya and
Turkey (Mukumya, 1974; Rudolph and Baykal, 1978).
Although the occurrence of bean rust was characterized as sporadic
in the U.S. (Harter et al., 1935), Vargas (1980) reported yield losses
as high as 40-80% in the U.S. are caused by this disease. Brazil is
reported to incur losses of 35-50% due to bean rust infections (Vargas,
1980).
Bean rust was reported to be responsible for the bulk of the yield
losses in Navy beans in Michigan (Andersen, 1975). The disease was
reported to be troublesome in snap bean fields of North Dakota and
Minnesota (Meiners, 1977). Zaumeyer and Meiners (1975) reported that
prior to 1945, bean rust was a major disease in irrigated fields in
Colorado, western Nebraska, Wyoming and Montana. In their review,
Zaumeyer and Meiners (1975) reported bean rust was no longer a problem
in those areas, although it was still occasionally important in fall
snap bean crops along the Atlantic seaboard and in winter crops grown in
Florida.


CHAPTER VII THE EFFECT OF INOCULATION SYSTEM ON Meloidogyne
incognita RACE ESTABLISHMENT ON BEANS 142
Introduction 142
Materials and Methods 144
Results 144
Discussion 155
CHAPTER VIII SUMMARY AND CONCLUSIONS 160
APPENDIX THE EFFECT OF FUNGICIDES ON SNAP BEANS 164
Introduction 164
Materials and Methods 165
Results 166
Discussion 174
LITERATURE CITED 176
BIOGRAPHICAL SKETCH 194
v


177
Barker, K. R., P. B. Shoemaker, and L. A. Nelson. 1976. Relationship
of initial population densities of Meloidogyne incognita and M.
hapla to yield of tomato. J. Nematol. 8: 232-239.
Barker, K. R., F. A. Todd, W. W. Shane, and L. A. Nelson. 1981. Inter
relationship of Meloidogyne spp. with flue cured tobacco. J.
Nematol. 13: 67-79.
Begum, A. and W. G. Eden. 1963. When to treat soybeans for worm
control. Highlights of Agrie. Res. 10(2): 1-4.
Begum, A. and W. G. Eden. 1964. Influence of defoliation on yield and
quality of soybeans. J. Econ. Entomol. 58: 591-592.
Berger, R. D. 1981. Comparison of the Compertz and logistic equations
to describe plant disease progress. Phytopathology 71: 716-719.
Blazey, D. A., P. G. Smith, A. Gentile, and S. T. Miyagama. 1964.
Nematode resistance in common bean. J. Hered. 55: 20-23.
Bonnefil, L. 1965. Las plagas del frijol en centro America y su
combate, pp. 61-88 In: XI Reunion del PCCMCA, Panama, March
17-19, 1965.
Bookbinder, M. G. and J. R. Bloom. 1980. Interaction of Uromyces
phaseoli and Meloidogyne incognita on beans. J. Nematol. 12:
177-182.
Bridge, J. 1973. Hoplolaimus seinhorsti, an endoparasitic nematode of
cowpea in Nigeria. Plant Dis. Rep. 57: 748-799.
Bridge, J., W. S. Bos, L. J. Page, and D. McDonald. 1977. The biology
and possible importance of Aphelenchoides arachidis, a seed-borne
endoparasitic nematode of groundnuts from northern Nigeria.
Nematologica 23: 253-259.
Briggs, M. P. 1946. Culture methods for a free living nematode. M. A.
Thesis. Stanford University. 50 pp.
Brodie, B. B. and W. E. Cooper. 1964. Relation of parasitic nematodes
to postemergence damping-off of cotton. Phytopathology 54:
1023-1027.
Brodie, B. B. and P. 0. Dukes. 1972. The relationship between tobacco
yield and time of infection with M. javanica. J. Nematol. 4:
80-83.
Brown, J. S. and R. J. Holmes. 1983. Guidelines for use of foliar
sprays to control stripe rust of wheat in Australia. Plant Dis.
67: 485-487.
Camery, M. D. and C. R. Weber. 1953. Effects of certain simulated hail
injury on soybeans and corn. Iowa Agrie. Expt. Sta. Res. Bull.
404. 39 pp.


138
and disease severity. Since there were significant differences among
fungicides means were separated using Duncan's multiple range test
showed that there were no significant differences between mancozeb
sprays and the unsprayed treatment. There was, however, significant
difference between bitertanol and mancozeb sprays (Table 34-2).
Regression analysis on the effect of defoliation, nematodes, and
disease severity on snap bean yield produced models of the form: y = a
+ b. X, + b0 X. + ... + bnxn, where y = yield (g/plot), x, ...x =
l l z z In
independent variable (defoliation level, disease severity, and log
(nematode population + 1) (Table 35). Stepwise regression showed that
disease severity contributed most to the coefficients of determination
2
(R ) (Table 35). Since there were no significant differences among
defoliation levels and metam-sodium rates, further explanations in this
section will be restricted to disease severity or area under the disease
progress curve (AUDC). Thus, figures 25 and 26 show the relationship
between yield loss and AUDC and disease severity, respectively. The
relationship between these disease parameters and yield should be the
inverse of their relationship with yield loss. Regression analysis of
yield data produced models of the form y = a-bx, where y = yield
(g/plot), x = either disease severity or AUDC (Table 35). The
2
coefficients of determination (R ) for disease severity and AUDC were
0.54 and 0.52 respectively when the disease was assessed at harvest.
Coefficients of determination for disease severity for the other times
of disease assessment are given in Table 35. Since disease assessment
2
severity consistently had better R than AUDC, it was generally used in
yield data analysis. Moreover, a set of disease severity points results
in AUDC. Disease progress curves are shown in figures 22, 23, and 24.


163
Yield was generally low in these experiments, probably because of
insect pest pressure, especially cowpea curculio and other leaf-feeding
insects. In the defoliation, metam-sodium, and bean rust disease
experiment the low yields may have been due to the late planting thus
going into a season where rainfall was erratic and insect pests were
more abundant. Generally, the weather conditions were not optimum for
snap bean production during the period some of the field experiments
were conducted.
A multipest assessment on beans in the field was not easy because
it was time consuming and tedious. Thus, a grower/extension worker
would have to assess pests separately and act on them accordingly. This
has been shown to be a valid approach in these experiments by the
general lack of interaction among defoliation, nematodes or nematicide,
and bean rust disease.


148
TABLE 40. Total number of eggs and juveniles extracted from 120 g of bean
and tomato roots. Data are means of 3 replicates.
NaOCl Concentration
(%)
Inoculum Source
Bean
Tomato
0
135,000
320,000
0.13
290,000
704,000
0.26
517,000
1,566,000
0.525
642,000
2,120,000
1.3
864,000
2,886,000
1.6
1,062,000
3,228,000


184
Lamberti, F. 1975. Fumiganti e nematocidi sistemici nella lotta contro
i fitoelminti ipogei. Report presented at the Round Table of
S.I.F., Cagliari, Italy.
Lamberti, F. 1979. Economic importance of Meloidogyne spp. in subtrop
ical and Mediterranean climates, pp. 340-356 In: Root-knot
nematodes (Meloidogyne spp.): Systematics, Biology and Control.
F. Lamberti and C. E. Taylor (eds.). Academic Press, New York.
Laundon, G. F. and J. M. Waterston. 1965. Uromyces appendiculatus.
CMI descriptions of pathogenic fungi and bacteria No. 57.
Livne, A. 1962. Photosynthesis in healthy and rust affected tissues.
Phytopathology 52: 739 (Abstr.).
Lopez, G. M. A. 1976. Identification de razas de la roya (Uromyces
appendiculatus (Pers.) Unger) del pijol (P^. vulgaris) en Puerto
Rico. Ph.D. Dissertation. Univ. de Puerto Rico, Mayaquez, 50 pp.
Ludwig, C. A. 1926. Some effects of late defoliation on cotton. South
Carolina Agrie. Expt. Sta. Bull. 238, 23 pp.
Madden, L. V., S. P. Pennypacker, C. E. Antle, and C. H. Kingslover.
1981. A loss model for crops. Phytopathology 71: 685-689.
Madriz, R. and E. Vargas. 1975. Evaluacin de la resistencia de
cultivares de prijol a la roya (Uromyces phaseoli var. typica)
mediante tres mtodos diferentes. In: Reunion Annual del Programa
Cooperativo Centroamericano para el Mejoramiento de Cultivas
Alimenticios 21a, San Salvador, El Salvador.
Maggenti, A. 1981. General Nematology. Springer-Verlag, New York.
372 pp.
Main, C. E. 1977. Crop destruction The raison d etre of Plant
Pathology, pp. 55-78 In: Plant Disease, an Advanced Treatise.
Vol. I. How Disease is Managed, J. G. Horsfall and E. B. Cowling
(eds.) Academic Press, New York.
Makram, M. W., S. T. Sidky, S. H. Nassar, and F. S. Faris. 1973.
Producing a variety of beans resistant to the rust disease,
Uromyces phaseoli. Agrie. Res. Rev. 51: 145-152.
Marcus, C. P. 1952. A new physiologic race of rust (Uromyces phaseoli
typica Arth.) causing losses to beans in Maryland. Phytopathology
42: 342.
Martinez, M. S. 1983. Principales enfermedades fungosas del frijol,
pp. 12-32 In: Curso Intensivo de postgrado en la produccin de
frijol, 40, Mantazas, Cuba, 1983. Conferencias, Cuba.
McAlister, D. F. and C. A. Krober. 1958. Response of soybeans to leaf
and pod removal. Agron. J. 50: 674-677.


157
highest number of eggs from either crop. The 0.525% solution had the
highest number of juveniles that emerged from eggs obtained from either
beans or tomato. Thus, a 0.525% NaOCl solution may be the optimum
concentration for M. incognita egg extraction for inoculation studies.
The low emergence of juveniles from water-treated eggs may imply that
under natural conditions root-knot nematode eggs hatch over a long
period. This may be a survival mechanism for this nematode.
There were no significant differences in the proportion of eggs
that had hatched between water and NaOCl solutions of 0.13, 0.26 and
0.525%. This was because there were far fewer eggs extracted with water
than with the NaOCl treatments. The lowest percentage hatch was
obtained from eggs extracted with 2.6% NaOCl. This low hatch was
probably due to the adverse effects of high NaOCl concentration on eggs
which may have arrested the development of embryos of juveniles.
Gall inoculations had a more marked effect on yield and gall
indices (as evidenced by greater slope) than extracted eggs probably
because eggs in intact egg masses hatched in their natural environment
and there was no loss from mortality due to NaOCl. The effect of egg
extraction may have been exacerbated by the inoculation of soil before
seeds germinated. Thus, juveniles may have been rendered less infective
before seeds germinated. Root-knot nematodes are more infective at the
second juvenile than any other stage (Dropkin, 1980). Seeds germinated
5 days after planting and nematode inoculation. A good proportion of
eggs may have hatched well before the plants produced roots. Nematode
juveniles, especially root-knot nematodes, are sensitive to relatively
high edaphic temperatures (McKenry, 1983). Root gall indices were


TABLE 35. Regression equations for the effect of defoliation, nematodes, and bean rust on snap beans.
Time of assessment of disease
Regression equation
Flower bud formation
(preplant nematode counts)
Full bloom
(preplant nematode counts)
Pod formation
(preplant nematode counts)
Pods half-developed
(preplant nematode counts)
Pods fully formed
(preplant nematode counts)
y = 637.7-262.7x^-6608.9x2+26.3x^-62x^+2.8x^
y = 638.8-332.3x^6497.6x2
y = 547.6-6164.IX2
y = 539.6-234.8x,-3269.5x+55.lx+25.5x.-1.32xc
1 2 3 4 5
y = 653.56-314.82x^299.2
y = 572.46-295.5x2
y = 695.7-290x1-947.7x+7.6x_-102.6x.+9.8x[;
1 2 3 4 5
y = 646.1-348.4x^-925x2
y = 549.6-870.6x2
y = 740.4-275.2x,-569.lx0-2.6x_-78.lx.+2.lxc
1 2 3 4 5
y = 665.9-301.2x^-564x2
y = 590.4-563.3x2
y = 702.8-283.4x^398.3x2+2.2x3-ll.lx4-2.3x5
(R
(R2
(R2
(R2
(R2
(R2
(R2
(R2
(R2
(R
(R2
(R2
(R
0.45**)
0.42**)
0.26**)
0.67**)
0.61**)
0.47**)
0.56**)
0.52**)
0.36**)
0.62**)
0.61**)
0.48**)
0.66**)
124


21
The uredia (uredinia) are the major diagnostic sign of the pathogen
(Fromme and Wingard, 1918). In a susceptible reaction, symptoms of bean
rust first appear on the lower leaf surface as minute, whitish, slightly
raised spots about 5-6 days after infection. These spots enlarge to
form mature reddish-brown pustules which rupture the epidermis and
obtain a diameter of 1-2 mm, 10-12 days after infection (Vargas, 1980).
The uredia reach a diameter of 5 mm by the 14th day after infection (Rey
and Lozano, 1961). The size of the uredia varies depending on environ
mental conditions as well as the host. The uredia may appear powdery
due to uredospores protruding from them (Fromme and Wingard, 1918).
Uredia often appear on both leaf surfaces. The uredia are frequently
surrounded by chlorotic halos and eventually by rings of secondary and
tertiary sori (Zaumeyer and Thomas, 1957). As infection progresses, the
leaf becomes debilitated and the chlorotic areas surrounding pustules
coalesce, while tissue ramified by the fungus remains green, apparently,
as a result of starch accumulation (Wang, 1961). Severe rust infections
may cause premature abscission. Bean rust rarely causes small, circular
necrotic lesions on pods (Kucharek and Simone, 1980).
Rust infection has been reported to cause increased respiratory
rates in susceptible hosts (Daly et al., 1961). Twenty-four hours after
infection starch accumulation decreases sharply in the vicinity of the
fleck. Accumulation of starch at the perimeter of the lesion, however,
resumes 96-120 hr after infection. The quantity of starch in this area
decreases at the time the fungus sporulates (Schipper and Mirocha,
1969). Rust infections cause leakage of ions, amino acids, and sugars
in susceptible plant leaves (Hoppe and Heitefuss, 1974a). Hoppe and
Heitefuss (1974b) presented evidence that rust infection caused damage


155
Plants inoculated with one gall/pot had a similar gall index to that of
the plants inoculated with 100 or 1,000 eggs and juveniles/pot as
anticipated. Ten galls per pot gave gall indices similar to these of
10,000 or 100,000 eggs and juveniles/pot. Plants inoculated with more
than 10 galls gave the maximum gall index. The controls had no root
galls.
Data obtained in this study did not fit the Seinhorst model
(Seinhorst, 1965, 1972) which is of the form Y = M + (l-M)Z^ ^ where Y
= ratio between the yield at nematode population P and at P < T, M =
relative minimum yield, P = initial nematode population density, T =
tolerance limit for the nematode density, and Z = a constant. Coeffi
cients of determination were 0.031, 0.003, and 0.28 for the soil mix,
seed drench, and gall inoculation systems. Linear regression analysis
of yield data is shown in Figure 33. The linear model produced signifi
cant coefficients of determination at the 0.01 probability level.
Discussion
The largest number of M. incognita eggs and juveniles was extracted
with the 2.6% NaOCl solution, due to the better dissolution of the
gelatinous matrix enclosing the nematode eggs at the highest NaOCl
concentration tested. The rate of increase of eggs and juveniles
extracted, however, decreased dramatically above the 0.525% concentra
tion probably because the number of egg masses or galls per sample was
the same. Further the increase in NaOCl did not improve egg and juve
nile extraction that much.


TABLE 1. Nematodes commonly found In association with roots of beans.
Species
Distribution
Reference
Meloidogyne arenaria (Neal) Chitwood
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980; Castillo
and Litsinger, 1980
M. hapla Chitwood
N. Europe, Japan, U.S.A., Agudelo, 1980; Sinclair
Canada, & warmer regions of and Shurtleff, 1975
Africa and Middle East
M. incognita (Kofoid & White) Chitwood
M. javanica (Treub) Chitwood)
Pratylenchus brachyurus (Godfrey) Filipjev
Aphelenchoides spp.
Rotylenchulus reniformis Linford & Oliveira
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980
Cosmopolitan, tropical to warm
temperate regions
Agudelo, 1980;
& Shurtleff,
Sinclair
1975
Cosmopolitan
Agudelo, 1980;
1973
Bridge,
Nigeria
Agudelo, 1980;
et al., 1977
Bridge
W. Africa, U.S.A., Indonesia,
Agudelo, 1980;
Ayala &
Philippines Ramirez, 1964; Singh
& Farrell, 1972
Helicotylenchus spp.
Cosmopolitan
Agudelo, 1980; Bridge,
1973; Hague, 1980
Criconemella spp.
Widespread
Agudelo, 1980; Feakin,
1973


84
Therefore, no price information on this product is given. In trial 1,
plants with a 0.46 disease severity gave a higher dollar value than
plants with a 0.098 disease severity. These dollar values corresponded
to spray intervals of mancozeb and sulfur of 7-days and 4-5 days (Table
19). Plants with a disease severity of 0.4 and 0.47 gave dollar values
of $2367 and $2327 respectively in trial 2. These disease severity
values corresponded with 4-5-day and 7-day spray schedules of mancozeb
and sulfur (Table 19). In trial 2, gross dollar values were consistent
ly inversely related to both disease parameters (Table 10). The plants
with the highest disease parameter produced the lowest gross dollar
value.
The relationship between disease severity and net returns from
investment per hectare of snap beans is shown in Table 20. No net
returns are shown for the virtually disease free plants because an
experimental fungicide with no price tag was used on them. In trial 1
there was no improvement on net returns by spraying beans at 4-5 day
intervals from the 7-day intervals. Actually, there was a loss in net
income by spraying plants more often (Table 20). In trial 1, there were
substantial increases in net returns when plants were sprayed at 7-day
intervals compared to the 14-day spray schedule. The increases in
returns were $1069, $578, and $306 at the high, medium and low prices at
the 7-day spray schedule from the 14-day schedule in trial 1. From the
14-day spray schedule to the 4-5-day spray interval there were increases
in net returns of $901, $475, and $239 at the high, medium, and low
prices respectively in trial 1. By increasing spray frequency from
7-day to 4-5-day intervals there were net losses of $168, $103, and $67
at the high medium, and low prices respectively. Thus, in trial 1 there


122
TABLE 34-1. F values from the analysis of variance for yield.
Source
F value
Probability of F
Defoliation
3.06
0.07
Metam-sodium
0.12
0.73
Defoliation x
Metam-sodium
2.20
0.12
Fungicide
7.62
0.0002
Defoliation x
Fungicide
1.15
0.34
Metam-sodium x
Fungicide
1.41
0.25
Defoliation x
Fungicide x
Metam-sodium
0.62
0.72


61
TABLE 9. Regression equations for the relationship between M. incognita
population levels (x) and yield component (y).
Yield component
Model
Linear
Quadratic
Yield
Y = 118.6 -13.98x
Y = 126.7 26x + 2.4x2
R2 = 0.81*
R2 = 0.87*
Log (Yield)
Y = 2.1 O.lx
Y = 2.1-0.lx + 0.005x2
R2 = 0.84**
R2 = 0.85*
* R significant at 0.05
** R significant at 0.01


186
Minton, N. A., M. B. Parker, and R. A. Flowers. 1975. Response of
soybean cultivars to Meloidogyne incognita and to combined effects
of M. arenaria and Sclerotium rolfsii. Plant Dis. Rep. 59:
920-923.
Montalbini, P. 1973. Effect of infection by Uromyces phaseolus (Pers.)
Wint. on electron carrier quiones in bean leaves. Physiol. Plant
Pathol. 3: 437-441.
Morrell, J. J. and J. R. Bloom. 1981. Influence of M. incognita on
Fusarium wilt of tomato at or below minimum temperature for wilt
development. J. Nematol. 13: 57-60.
Mountain, W. B. 1965. Pathogenesis by soil nematodes, pp. 285-301
In: Ecology of Soil Borne Plant Pathogens, Prelude to Biological
Control. K. F. Baker and W. C. Snyder (eds.). John Murray,
London.
Mukumya, D. M. 1974. Bean diseases in Kenya. Annu. Rept. Bean Improv.
Coop. 17: 57-59.
Nasser, L. C. B. 1976. Efeito da ferrugem em differentes estadios de
desenvolvimiento do feijoeiro e dispersao de esporos de Uromyces
phaseoli var typica Arth. Tesis M.S., Univ. Federal de Vicosa,
Minas Gerais, Brazil, 79 pp.
Navarro, A. R. and 0. R. Barriga. 1970. Control de nematoda fitopara-
sitos pormeido de rotacin con cultivos resistantes a estos
organismos. Revista (ICA) 5: 173-184.
Nemec, S. and L. S. Morrison. 1972. Histopathology of Thuja orientalis
and Juniperus horizontales plumosa infected with M. incognita. J.
Nematol. 4: 72-74.
Ngundo, B. W. 1977. Screening of bean cultivars for resistance to
Meloidogyne spp. in Kenya. Plant Dis. Rep. 61: 991-993.
Ngundo, B. W. and D. P. Taylor. 1974. Effects of Meloidogyne spp. on
bean yield in Kenya. Plant Dis. Rep. 58: 1020-1023.
Ngundo, B. W. and D. P. Taylor. 1975a. Comparative histopathology of
six bean cultivars infected with M. incognita and M. javanica. E.
African Agrie. For. J. 41: 76-80.
Ngundo, B. W. and D. P. Taylor. 1975b. Some factors affecting
penetration of bean roots by larvae of M. incognita and M.
javenica. Phytopathology 65: 175-178.
Noe, J. P. and K. R. Barker. 1983. Edaphic variables related to the
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Norton, D. C. 1960. Effects of combinations of pathogenic organisms at
different temperatures on the cotton seedling disease. Texas
Agrie. Expt. Sta. Mise. Publ. 412, 20 pp.


4
Root-knot nematodes, Meloidogyne spp., also infect bean plants.
Meloidogyne spp. are most prevalent in light sandy soils with good
drainage and moderately warm temperatures (25-30C) (Crispin et al.,
1976). Roberts and Boothroyd (1984), however, stated that M. incognita
(Kofoid and White) Chitwood is more common in southern states of the
U.S. and M. hapla Chitwood is commonly found in northern states.
Meloidogyne spp. reportedly limit the production of beans by interfering
with nitrogen fixation by Rhizobium spp. and causing root galls (Ngundo,
1977; Sharma and Guazelli, 1982; Singh et al. 1981 a). Meloidogyne
incognita has been observed to predispose beans to Fusarium wilt (Singh
et al., 1981 b). Severe root-knot nematode infections may lead to
50-90% yield loss (Freire and Ferraz, 1977; Ngundo, 1977; Varn and
Galvez, 1974).
Control of these pests and diseases has been based on chemical
pesticides and cultural methods (Acland, 1971; Allen, 1983; Carvalho et
al., 1981; Martinez, 1983; Rhoades, 1976; Robbins et al., 1972; Shorey
and Hall, 1963; Stoetzer and Omunyin, 1983; Villamonte, 1965; Yoshii,
1977; Zaumeyer and Meiners, 1975). The use of resistant varieties and
flooding has been part of the management strategies for bean rust and
root-knot nematodes (Crispin et al., 1976; Martinez, 1983; Ngundo, 1977;
Singh et al., 1981a; Vieira, 1967).
One of the problems research scientists are confronted with, in
crop pest management, is the development of multi-pest threshold levels
to be used in determining the type of management strategies and the
extent to which these pest complexes have to be controlled so that
maximum yield is obtained with minimum disruption of the environment.
Pesticides are, however, the main means of controlling pest complexes on


CHAPTER V
THE EFFECT OF BEAN RUST, UROMYCES PHASEOLI (PERS.) WINT.,
ON SNAP BEANS, PHASEOLUS VULGARIS L. 'Sprite'
Introduction
Bean rust, caused by the fungus Uromyces phaseoli (Pers.) Wint., is
a serious disease of beans, Phaseolus vulgaris L. (Agudelo, 1980; Allen,
1983, McMillan et al., 1982, Pohronezny et al., 1984). The disease
causes severe damage on winter and spring grown snap beans in south
Florida (McMillan, 1982; Pohronezny et al., 1984). Usually, IJ. phaseoli
first appears in January and becomes progressively more severe February
through May (Pohronezy et al., 1984). Initial inoculum is believed to
come from infected bean plant debris in abandoned fields. Losses of up
to 78% in pinto beans, 74.2% and 18.4% in 'Ex Rico 23' and 'Bat 308'
field beans, respectively, have been reported from severely infected
crops in the United States and Latin America (CIAT, 1983; Kelly, 1982).
Bean rust, £. phaseoli, is an autoecious polycyclic disease whose
rates of increase are affected by timing, amount of sporulation, light
intensity, relative humidity, and relative cultivar susceptibility
(Cohen and Rotem, 1970; Cook, 1978; Imhoff et al., 1982 a,b). Rotem et
al. (1973) reported that, in an automatic humidity chamber study,
humidity was inversely related to the sporulation of £. phaseoli.
Infection by IJ. phaseoli has, however, been reported to be favored by
prolonged periods of at least 95% relative humidity and moderate temper
atures (15-27C) (Augustin et al., 1972; Gonzalez, 1976; Schein, 1961).
77


CHAPTER VIII
SUMMARY AND CONCLUSIONS
Pest damage and disease infection on beans rarely result in total
leaf abscision under field conditions. In the experiments conducted in
this study, various defoliation levels up to total defoliation were
included. Bean rust was manipulated by fungicide sprays at various
frequencies and nematode infestation was studied under field as well as
greenhouse conditions. These factors were studied individually and in
combination.
Bean plants were most sensitive to defoliation at full-bloom and
pod-set growth stages. Twenty five percent leaf removal at the primary
leaf and first trifoliate leaf stages resulted in yield losses of up to
36% and 29% in the greenhouse and field respectively. In the greenhouse
50%, 75%, and 100% defoliation resulted in up to 34%, 39%, and 74% yield
loss, respectively. In the field 50%, 75%, and 100% defoliation caused
yield losses of up to 68%, 70%, and 95% respectively depending on the
plant growth stage. Yield loss fluctuated in such a way that in some
cases lower defoliation levels resulted in higher yield losses than
higher defoliation levels. This fluctuation may have been due to
various factors, one of which could be better exposure of photosynthe-
tically active foliage to light.
Root-knot nematodes, Meloidogyne incognita, caused yield loss in
snap beans when plants were inoculated with eggs and juveniles. Ten
eggs and juveniles per pot resulted in 19% yield loss and 100, 1,000,
10,000, and 100,000 eggs and juveniles resulted in 38%, 45%, 47%, and
160


181
Franklin, M. T. 1978. Meloidogyne. pp. 98-124 In: Plant Nematology,
J. F. Southey (ed.). MAFF, HMSO, London.
Freire, F. C. 0. and S. Ferraz. 1977. Nematoides asociados ao feijoeiro
na zona da Mata, Minas Gerais, e efeitos do parasitismo de M.
incognita e M. javanica sobre o cultivar Rico 23. Revista Ceres
24: 141-149.
Frenhani, A. A., E. A. Bulisani, E. Issa, and S. G. P. de Silveira.
1971. Controle da ferrugem (Uromyces phaseoli var. typica Arth.)
do feijoeiro (Phaseolus vulgaris L.), com fungicida sistemico 0
Biolgico 37: 25-30.
Fromme, F. D. and S. A. Wingard. 1918. Bean rust: Its control through
the use of resistant varieties. Va. Agr. Expt. Stat. Bull. 220 18
pp.
Fromme, F. D. and S. A. Wingard. 1921. Varietal susceptibility of
beans to rust. J. Agrie. Res. 21: 385-404.
Galvez, G. E., J. J. Galindo, and G. Alvarez. 1977. Artificial
defoliation for estimating losses from foliage damage in beans.
Turrialba 27: 143-146.
Gilvonio, H. V. and A. M. Ravines. 1971. Estudio del effecto del
inuculo de Meloidogyne acrita en frijol. Nematropica 1: 43.
Golden, J. K and S. D. Van Gundy. 1975. A disease complex of okra and
tomato involving the nematode M. incognita and the soil inhabiting
fungus Rhizoctonia solani. Phytopathology 65: 265-273.
Gonzalez, A. M. 1976. Investigaciones sobre el compartamiento de
variedades de frijol prente al patgeno causante de la roya
(Uromyces phaseoli var. typica Arth.). pp. 26-32 In: Acad, de
Ciencias de Cuba.
Greene, C. L. 1971. Economic damage levels of bean leaf roller
populations on snap beans. J. Econ. Entomol. 64:673-674.
Greene, C. L. and D. R. Minnick. 1967. Snap bean yields following
simulated insect defoliation. Proc. Fla. State Hort. Soc. 80:
132-134.
Groth, J. V. and B. D. Shrum. 1977. Virulence in Minnesota and
Wisconsin bean rust collections. Plant Dis. Rep. 61: 756-760.
Guerra, E. and S. Dongo. 1973. Determinacin de razas firiologicas del
hengo Uromyces phaseoli var. typica Arth. en el Peru. Invest.
Agropec. 3: 92-94.
Hague, N. C. M. 1980. Nematodes of legume crops, pp. 199-205 In:
Advances in Legume Science, R. J. Summerfield and A. H. Bunting
(eds.) HMSO, London.


51
inconsistency was due to the imprecise nature of manual defoliation in
simulating insect damage. It is possible that removing whole fractions
of leaf surfaces had a different effect on plants from damage done by
leaf feeding pests which usually occurs at random.
Although the influence of defoliation on yield was not consistent
at all plant growth stages, plants showed more sensitivity at full bloom
and pod set. This was an indication that leaf damaging pests should be
managed before these plant growth stages. If left unchecked and if
plants become heavily defoliated, substantial loss in yield would be
expected. Insecticides are generally applied as soon as insect pest
infestations are detected. Insecticide application normally starts
before the pest populations exceed threshold levels. Disease control
chemicals are primarily preventatives applied well before the diseases
are observed. Since pesticide sprays against diseases and insect pests
were the same at all defoliation levels and plant growth stages, the
grower would incur loss in gross dollar values proportional to loss in
yield. In this study the threshold level for defoliation was below 25%.
At all plant growth stages.


TABLE 23. Effect of metam-sodium on snap bean yield and populations of nematode genera in final soil
samples (at harvest).
Metam-sodium
(Liters/ha)
Criconemella
(No./lOO ml)
Helicotylenchus
(No./lOO ml)
Meloidogyne
(No./lOO ml)
Rotylenchulus
(No./lOO ml)
Yield
(g/plot)
0
1
21
4
25
360
47
1
27
6
1
596
94
4
72
1
4
650
187
1
17
0
0
680
281
0
42
0
18
581
374
0
25
0
1
707
Ill


Yield (g/plot)
109
0.00004x3
Figure 21.
Effect of metam-sodium on snap bean yield.


TABLE 1. Continued.
Species
Distribution
Reference
Belonolaimus spp.
Southeastern U.S.A.
Agudelo, 1980; Feakin,
1973; Sinclair &
Shurtleff, 1975
Trichodorus spp.
Widespread
Agudelo, 1980; Feakin,
1973
Xiphinema spp.
Widespread
Agudelo, 1980; Caveness
et al., 1975; Feakin,
1973


24
Control of the Disease
Cultural control measures of this disease include crop rotation and
removal of old plant debris (Vieira, 1967). Reduced plant density and
planting date adjustment for specific production areas may reduce rust
incidence (Vargas, 1980). Resistant varieties of beans have been used
for the control of rust (Augustine et al., 1972; Ballantyne, 1974; Coyne
and Schuster, 1975; Crispin et al., 1976; Madriz and Vargas, 1975;
Meiners, 1974; Rivera, 1977; Rodriguez, 1976).
Fungicidal sprays are usually recommended to help manage bean rust.
Since bean rust reduces yields more severely when infection occurs
before flowering than when infection is initiated after flowering,
fungicidal sprays are, therefore, more effective if applied during early
plant development (Yoshii and Galvez, 1975). Of the older fungicides,
sulfur dusts have given relatively good control (Ballantyne, 1975;
Harter et al., 1935; Zaumeyer and Thomas, 1957). Sulfur is usually
applied at the rate of 25-30 kg/ha every 7-10 days. Generally, protec
tant fungicides fail in areas where rainfall is frequent because
deposits are washed off too soon. Other preventative chemicals applied
at schedule similar to that of sulfur are chlorothalonil (225 g/ha),
maneb (4-5 kg/ha), and mancozeb (3-4 kg/ha) (Costa, 1972; Crispin et
al., 1976; Hilty and Mullins, 1975; Vieira, 1967; Wimalajeewa and
Thavam, 1973).
Plantvax (Oxycarboxin) is somewhat therapeutic when sprayed 20 to
40 days after planting at the rate of 1.8-2.5 kg/ha (Costa, 1972;
Frenhani et al., 1971; Hilty and Mullins, 1975). McMillan et al. (1982)
reported effective control of bean rust when bean plants were sprayed
weekly with bitertanol or triadimefon. These fungicides are not regis
tered for use on beans at this time. While certain fungicides are


87
TABLE 16. Effects of Uromyces phaseoli on snap bean yield.
Disease Parameters
Maximum Area under disease
Proportion of progress curve Yield (g/plot)
foliage
infested
(sq.
units)
Trial 1
Trial 2
Trial 1
Trial 2
Trial 1
Trial 2
0.76
0.86
5.64
13.90
1102
276
0.65
0.71
5.93
12.85
1778
642
0.46
0.47
4.11
9.28
2248
1426
0.098
0.40
0.98
7.33
2158
1452
0.0
0.0
0.014
0.0
2723
3214


TABLE 30. Nematode genera found in soil samples (harvest). Data are means of 4 replicates.
Metam-sodium Criconemella Helicotylenchus Meloidogyne Rotylenchulus Tylenchorhynchus
(Liters/ha) (No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml) (No./lOO ml)
0
47
94
187
374
0
0
2
0
0
8
5
2
2
0
4
8
6
5
3
9
9
9
9
4
0
1
1
1
0
00


TABLE 4. Effect of defoliation and defoliation time on snap bean yield (g/plot) in the greenhouse and
field. Data are means of 4 replicates.
Snap
bean yield
(g/plot)
by plant growth
stage
Primary
leaf
First
trifoliate
leaf
Third
trifoliate leaf
Flower
bud formation
Full bloom
Pod set
Defoliation
level (I)
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
Greenhouse
Field
0
138
1361
122
1168
125
1293
128
1295
121
1138
122
1530
25
89
1403
84
835
97
1210
89
1113
94
1140
96
1085
50
92
1105
72
933
99
1140
89
673
79
690
102
495
75
84
1050
82
1215
81
820
78
620
93
813
84
457
100
46
613
65
120
54
335
46
350
52
52
32
125


152
Figure 32. Egg hatch (%) at various NaOCl concentrations.


8
machinery. Sometimes defoliation is initiated by chemical defoliants to
facilitate harvesting (McGregor et al., 1953). In most cases defolia
tion occurs due to pest problems and adverse environmental conditions.
To reduce defoliation by pests, preventative spray programs are usually
followed (Greene and Minnick, 1967). These sprays are usually applied
regardless of the anticipated crop loss. Hence, it would be desirable
to determine the relationship between defoliation levels and yield
losses to maximize the efficiency and rationale for spray treatments.
A wide range of yield losses due to artificial defoliation has been
observed on various bean cultivars (Edje and Mughogho, 1976a, 1976b;
Edje et al., 1973, 1976; Garvez et al., 1977; Greene and Minnick, 1967;
Hohmann and De Carvalho, 1983; Vieira, 1981; Waddill et al., 1984).
Edje et al. (1973, 1976), Edje and Mughogho (1976a, 1976b), Vieira
(1981) and Waddill et al. (1984) manually defoliated indeterminate bean
cultivars. Waddill et al. (1984) reported that complete defoliation
when only primary leaves were present reduced yield by about 65% and
repeated weekly defoliation of 50% resulted in 34% yield loss. Vieira
(1981) reported that 66% leaf area removal during the flowering and pod
formation stages was detrimental to yield. Galvez et al. (1977) ob
served that 100% defoliation at formation of the first trifoliate leaves
decreased yields of the bean cultivars ICA-Guali and Porrillo-Sentetico
by 34% and 49% respectively. Greene and Minnick (1967) indicated that
yield reduction in snap beans begins somewhere between 33% and 50%
defoliation when defoliation occurs in the prebloom and bloom stages,
respectively. Hohmann and DeCarvalho (1983) reported that removal of
25, 50, 75, and 100% of the leaf area at the pod formation stage reduced


74
occurred when 10,000 or 100,000 eggs and juveniles/pot were used. This
discrepancy in M. incognita effects on snap bean yield may be due to the
difference in the seasons in which the two trials were conducted. Tyler
(1933) reported that at temperatures ranging from 27.5C to 30C,
females of Meloidogyne spp. developed from infective juveniles to the
egg-laying stage in 17 days; at 24.5C in 21 to 30 days; at 20C, in 31
days; at 15.4C in 57 days; and at temperatures above 33.5%C or below
15.4C, females failed to reach maturity on tomato plants. Decker and
Casamayor-Garcia (1966) stated that one generation of M. incognita
developed on lettuce within 26 days at a mean temperature of 23.3C.
They further stated that from the time of larval invasion up to the
commencement of egg-laying required at least 19 days. Lamberti (1979)
observed that M. incognita rarely started to invade root tissues when
the soil temperature was below 18C. Since the two tests being discus
sed here were conducted at different times of the year, it is likely
that in the summer test soil temperatures were generally higher than in
the fall trial. Thus, the life cycle of the nematode may have been
completed in a shorter period of time in the summer than in the fall.
Consequently, more M. incognita generations (at least 3) may have been
completed during this season. There is also the possibility that the
quality of the inoculum was different in the two tests since the source
of the eggs and juveniles for the fail test was also exposed to
relatively lower temperatures than the summer inoculum. The verifi
cation of this phenomenon can only be obtained by conducting further
tests. The higher yields in the fall test may also be due to the fact
that the nematodes were not able to invade the root tissues of the
plants as fast as they could under optimum summer conditions. During


TABLE 35. Continued.
Time of assessment of disease
Regression equation
1
Pods fully formed
(final nematode counts)
Pods fully formed
(final nematode counts)
y = 687.1-288.9x^397.7x2
y = 616.7-401x2
y = 702.8-283.4x^398.3x2+2.2x3-11.1x4-2.3x5
y = 687.1-288.9x^397.7x2
y = 616.7-401x2
(R2 = 0.66**)
(R2 = 0.54**)
(R2 = 0.66**)
(R2 = 0.66**)
(R2 = 0.54**)
y = 869.8-230.4x -415.9x0+6x0-18.3xc+6x,-42.4x_, /n2
1 j 3 0 / V.K
y = 734.6-282.5x^410.7x2-33x? (R2
y = 616.7-401.4x2 (R2
0.69**)
0.66**)
0.54**)
y = yield (g/plot; x^ = defoliation level; x2 = disease severity; x^ = Heliotylenchus log ( x+1);
x, = Meloidogyne log ( x+1); x,. = Rotylenchulus log ( x+1); x, = Tylenchorhynchus log ( x+1); x7 =
Criconemella log ( x+1).
** R significant at P < 0.01.
125