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1988 Genetics of Resistance to Reniform , Reniformis Linford and Oliveira, in Cotton, Gossypium Hirsutum L. Noor Muhammad Louisiana State University and Agricultural & Mechanical College

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Recommended Citation Muhammad, Noor, "Genetics of Resistance to Reniform Nematode, Linford and Oliveira, in Cotton, Gossypium Hirsutum L." (1988). LSU Historical Dissertations and Theses. 4586. https://digitalcommons.lsu.edu/gradschool_disstheses/4586

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Genetics of resistance to reniform nematode,RotyUnchulus rtniformis Linford and Oliveira, in cotton,Gossypium hirsutum L.

Muhammad, Noor, Ph.D.

The Louisiana State University and Agricultural and Mechanical Col., 1988

UMI 300 N. Zecb Rd. Ann Arbor. MI 48106 GENETICS OF RESISTANCE TO RENIFORM NEMATODE,

ROTYLENCHULUS RENIFORMIS LINFORD AND OLIVEIRA,

IN COTTON, GOSSYPIUM HIRSUTUM L.

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and Agr icultural and Mechanical College in partial fulfillment of the requirements for the degree of Doctor of Philosophy

in

The Department of Agronomy

by Noor Muhammad B. Sc. (Hons.), University of Agriculture, Faisalabad, Pakistan, 1972 M. Sc. (Hons.), University of Agriculture, Faisalabad, Pakistan, 1976 August 1988 ACKNOWLEDGEMENTS

The author wishes to express his admiration to his major professor,

Dr. Jack E. Jones, Professor of Agronomy, Louisiana State University,

for his encouragement, advice and dedication. The author expresses

sincere thanks to the members of his advisory committee, Dr. Jerry B.

Graves, Dr. M. S. Kang, Dr. B. G. Harville, Dr. S. A. Harrison and Dr.

E. C. McGawley for their interest and valuable comments.

Di. S G. Harville, not only provided the research equipment, but

also extended full cooperation during the course of this research.

The author gratefully acknowledges the advice, guidance, and

encouragement of Dr. E. C . McGawley regarding techniques in inoculat ion

and assessing populations of the reniform nematode used in this

research. The technical assistance of Ms. K. L. Winchell and Dr. J. P.

Beasley Jr. in similar activity is greatly appreciated.

Dr. M. S. Kang played a major role in genetic analyses of the data

and author gratefully acknowledges his efforts and guidance during the

course of this investigation. The author sincerely appreciates the

help, cooperation and guidance of Dr. A. M. Saxton in the statistical

analysis of the data and wishes to express his gratitude to him.

The author is greatly impressed by the enthusiasm, dedication and

professional guidance of Dr. S. A. Harrison from the very beginning of

this study. The assistance of research associates, Mr. J. I. Dickson,

Mr. W. C. Smith and Dr. S. J. Stringer is gratefully acknowledged.

The financial assistance by the U. S. Agency for International

Development and the opportunity provided by the Government of Pakistan

ii is greatly appreciated. The author is grateful to the program manager,

Mr. Kevin Schieffer, program specialists, Ms. Julie Ann Iselin and Ms.

Zaiga Robins, and staff of the Pakistan Participant Training Program,

Washington, D. C. for continuous moral support, and timely payment of

tuition fees and allowances.

The author expresses his love and appreciation to his mother and

late father, Saira and Muhammad Ali, for their patience, prayers,

support, and love during his education. He also wishes to thank his elder brothers, sisters, parents-in-law and other relatives for their

love, prayers, and support. The author sincerely appreciates the

patience and love of his wife, Surria and lovely children, Maimoona,

Naveed, Lubna, Aneela and Aisha.

Most of all, the author is thankful to the almighty God for his mercy and graciousness.

iii LIST OK TABI.ES

Tables Page

1 Inoculat ion dates, days for Rotylenchulus reniformis development and air temperature regimmes for three cotton crosses tested for reniform nematode resistance in the greenhouse...... 35

2 Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 1 (La. RN 910 x DP 41)...... 43

3 Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 2 (Auburn 612 RNR x DP41)...... 44

4 Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 3 (M 019-RNR x DP 41)...... 46

5 Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies oi crosses in Experiment 1 (La. RN 910 x DP 41)...... 48

6 Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 2 (Auburn 612 RNR x DP 41)...... 50

7 Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 3 (M 019-RNR x DP 41)...... 52

8 Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41)...... 54

9 Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 1 (La. RN 910 x DP 41)...... 55

10 Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41)...... 56

Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 1 (La. RN 910 x DP 41)...... 57

v 12 Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41)...... 58

13 Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 2 (Auburn 612 RNR x DP 41)...... 59

14 Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41)...... 60

15 Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 2 (Auburn 612 RNR DP 41)...... 61

16 Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 3 (M 019-RNR x DP 41)...... 62

17 Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41)...... 63

18 Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 3 (M 019-RNR x DP 41 )...... 64

19 Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41)...... 65

20 Estimates of number of gene pairs by which the two parents differed in resistance to R. reniformis in Experiments 1-3. 67

21 Broad-sense heritabilities (%) and standard errors (%) of resistance to R. reniformis in Experiments 1-3...... 67

22 Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 1 (La. RN 910 x DP 41)...... 69

23 Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 2 (Auburn 612 RNRx DP 41)...... 71

24 Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 3 (M 019-RNR x DP 41)...... 73

vi LIST OF APPENDIX TABLES

Tables Page

8 a Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41)...... 91

10a Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41)...... 92

11a Frequency distribution of number of eggs per gram of root for six populations at Planting 2 , Experiment 1 (La. RN 910 x DP 41)...... 93

14a Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41)...... 94

15a Frequency distribution of number of eggs per gram of root for six populations at Planting 2 , Experiment 2 (Auburn 612 RNR x DP 41)...... 95

18a Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 3 (M 019-RNR x DP 41)...... 96

19a Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41)...... 97

vii ABSTRACT

Four Gossyplum hirsutum L. lines (La. RN 910, Auburn 612 RNR, H

019-RNR, and 'Deltapine 41' (Dp 41)) were selected for a genetic study of their reaction to reniform nematode, Rotylenchulus renlformis Linford and Oliveira, based on observed differences. Dp 41 (susceptible) was crossed to each of the three moderately resistant lines. Fj generations were selfed and each was backcrossed to each of its parents. Estimates of mean gene effects and the three types of digenic epistatic effects affecting genetic variation of resistance to reniform were calculated using the generation means of Pjt P2 , Fj, F2 , PlFj, and P2 F 1 .

Six generations of the three crosses were evaluated for reniform resistance as seedlings grown in the greenhouse in plastic pots holding

500 grams of a sterilized mixture of Olivier silt loam soil and river sand. Each experiment (cross) consisted of two planting dates of two replications arranged in a randomized complete block design. Blocks of ten per replication for non-segregating generations (Pj, P2 * and

Fj)( blocks of 20 plants for PjFi and P2 P1 generations, and a block of

40 plants for the F2 generation were grown. At the first true leaf stage, each w inoculated with 2,000 reniform Juveniles. An

viii average growth period of 43 days was allowed in the winter and 32 days

in the summer. Ent ire root systems of plants were harvested to determine egg numbers.

Significant differences among generation means and generally higher

coefficients of variation for F2 and backcross populations than for parental and F^ populations indicated that differences between parents

for reniform resistance in 2 of 3 crosses were under genetic control.

Wide and non-discrete frequency distributions in segregating populations and high coefficients of variation indicated that resistance was quantitative in nature and greatly influenced by environment. No pattern was observed for the significance of additive and dominant gene effects.

Significant epistatic gene effects occurred in most cases and

transgressive segregation for susceptibility suggested that resistance

to reniform nematode in these cotton lines was controlled by at least two pairs of genes. Estimates of broad-sense heritability ranged from 0 to 86 % with a mean of 53% for eggs per root system and 0 to 85% with a mean of 57% for eggs per gram of root. Advancing generations to F^ or

F5 , while maintaining genetic variability prior to selection, may

improve selection efficiency by increasing homozygosity and thereby reducing nonadditive gene effects for resistance to reniform nematode.

ix INTRODUCTION

The reniform nematode, Rotylenchulus reniformis. Linford and

Oliveira (1940) was first identified on roots of upland cotton,

Gossypium hirsutum L., in Georgia (Smith, 1940). It was reported as a parasite of cotton in Louisiana in 1941 (Smith and Taylor). This nematode has become a serious pest in all coastal cotton-producing states (Birchfield and Jones, 1961; Bird et al,, 1973; Blasingame and

Patel, 1987; Fassuliotis and Rau, 1967; Lambe and Horne, 1963; Minton and Hopper, 1959; Neal, 1954).

The nematode female is a sedentary, endoparasite that feeds in the pericycle of the root and induces the formation of uninucleate giant cells (Veech, 1984). It reproduces abundantly on cotton, and causes a reduction in yield, a delay in maturity, a reduction in boll size, and in some years, a reduction in lint percent (Jones et al., 1959). It also causes dwarfing, premature decay with loss of secondary roots, and death of young cotton plants, which result in poor stands, grassy areas and yield reductions of 40-60% (Birchfield and Jones, 1961).

Nematicides, commonly used to suppress reniform before planting cotton (Birchfield, 1968; Birchfield and Jones, 1961;

Birchfield and Pinckard, 1964; Newsom and Jones, 1955; Thames et al.,

1969; Thames and Heald, 1974), do not prevent population build-up late in the season. This becomes a serious problem for the following crop and makes nematicide application imperative each year. Moreover, chemical control can lead to development of new nematode races that are resistant to nematicides, and create problems such as toxic soil

1 2 residues, environmental pollution, and biohazards. Rotation of cotton with non-host crops and fallow (weed free) land can subdue reniform nematodes (Braithwaite, 1974; Lambe and Horne, 1963; Rebois and Webb,

1979), but such practices may not be economically feasible. Utilization of resistant cultivars is an effective and economical means of reniform nematode control if resistant germplasm is available. Highly resistant cultivars could also be rotated with more desirable but susceptible cultivars to reduce or eliminate the necessity of pesticide application

(Rebois and Webb, 1979; Shepherd, 1982a; Williams et al., 1983).

While reniform nematode-resistant (Giveine max L. Merr) cultivars exist (Birchfield and Brister, 1969; McGawley et al., 1985), none are available for upland cotton growers. Although moderately reniform nematode-resistant lines have been identified in upland cotton

(Beasley, 1985; Jones et al., 1988; Yik and Birchfield, 1984), the genetic basis of this resistance is unknown.

Reniform nematode-resistant cultivars of , 'Pickett 71' and

'Dyer', are also resistant to the soybean cyst nematode, Heterodera glycines; but reniform and root-knot nematode, Meloidogyne incognita, resistance are not related (Rebois et al., 1970; Rebois et al., 1968).

Birchfield et al. (1971) have reported that not all soybean cyst nematode-res1stant breeding lines are also resistant to the reniform nematode and resistant genes for the two nematodes are separate but probably linked. Harville et al. (1985) reported that reniform nematode resistance in soybeans was quantitative in nature and controlled by two pairs of genes with unequal effects. Resistance to reniform nematode in (PI375937) is controlled by at least one dominant gene, which may 3

be closely linked to one or more genes for resistance to Heterodera

schachtii (Rebois et al., 1977),

The objective of the present study was to partition genotypic value

for resistance to reniform nematode, as measured by eggs per root system and eggs per gram of root of cotton plants, into additive, dominant, and

epistatic gene effects. Knowledge of estimates and relative importance

of these gene effects will hopefully be useful to plant breeders in

choosing breeding strategies and selection procedures for development of cotton cultivars with resistance to the reniform nematode. REVIEW OF LITERATURE

Distribution and Host Range of Reniform Nematode;

The reniform nematode, R. reniformis. was first reported as a parasite on , Vigna sinensis Endl., roots in Hawaii, by Linford and Oliveira (1940). The species was observed as early as 1935 by Yap on roots of , grown in a soil from a , Ananas comosus. field on the Island of Oahu, Hawaii (from Linford and Oliveira, 1940),

The common name 'reniform' was proposed because of the kidney shape appearance of the adult female body. Linford and Yap (1940) published a list of 65 host plants of reniform nematode, representing 30 families including pineapple, several weeds, and ornamentals, based on the presence of mature females with egg masses. Several plants were found to be unfavorable hosts, based on slow development of the females, maturation of few of them, and laying of very few eggs (Linford and

Yap, 1940). However, they did not mention cotton as a host plant. A species of Rotylenchulus was first identified by Steiner as a parasite of cotton near Cuthbert, Georgia (Smith, 1940).

Steiner (1947) reported cotton, tobacco, Nicotiana tabacum. coffee weed, Cassia tora L., and yew, Texus spp. as hosts of Rotylenchulus in the USA. He later found reniform nematodes attacking coffee weed and tomato, Lycoperslcon esculanturo L. In Tlorida and Jacquemontia tamnifolia L. Griaeb in Georgia (Steiner, 1949). Norton (1959) found R. reniformis in an ornamental nursery in Texas. It has been reported on cotton in Alabama (Minton and Hopper, 1959), Georgia (Bird et al.,

1973), Louisiana (Smith and Taylor, 1941), Texas (Lambe and Horne,

4 5

1963), Mississippi (Blasingame and Patel, 1987), and on ornamentals in

California (Allen and Magganti, 1959; Magganti and Allen, 1959; Sher

1959). It parasitizes sweet potatoes, 1pomea batatas, in Louisiana

(Martin, 1960), soybeans in South Carolina (Fassuliotis and Rau, 1967) and Alabama (Rebois and Cairns, 1968), roots of mango , ManRifera indica, in Florida (Van Weerdt et al., 1959b), An extension survey report by Blasingame and Patel (1987) showed that the two most damaging nematodes, reniform and root-knot [MeloidoRyne incognita (Kofoid and

White) Chitwood] are widely distributed in cotton-growing areas of the state of Mississippi.

R. reniformis has become a plant pathogen of considerable importance on a wide variety of crops in many areas of the world.

Martin (1955) reported its occurrence on baobad , Adansonia digitate

L., bamboo, Bambos vulgaris Schrad., and corn, Zea mays L., in Rhodesia and Nyasaland. Steiner (1960) found reniform nematodes attacking pigeonpea, Cajanus indicus Spreng., and reported it as being one of the most common nematode species on the island of Puerto Rico. Other hosts were added in an anonymous publication (Anonymous, 1960). Roman (1961) observed it in sugarcane, , fields in Puerto Rico, and Ayala (1961) found it to be widely distributed and in high density in pineapple fields in Puerto Rico. Ayala and Ramirez (1964) published a list of 201 different host plants from 15 countries, including 89 from

Puerto Rico, 15 of which were new hosts to Rotylenchulus spp. and 74 to

R. reniformis. Reniform nematode has also been reported from West

Africa (Luc and Guiran, 1960), Japan (Tanka, 1954), Taiwan (Hung, 1961),

Peru (Sasser et al., 1962), Egypt (Khadr et al., 1972; Oteifa and Salem 6

1972), Jamaica (Van Weerdt et al., 1959a), Guam (Reinking and Radewald,

1961), and Pakistan (Timm, 1956). Reniform nematode was included among several other spec ies known and suspected to be noxious to some grasses and white clover, Trifolium repens L., in the Netherlands meadows

(Oostenbrink, 1961). It was reported on tobacco, clover, and tea,

Camellia sinensis, in Java; on clover in Sumatra and the Philippines

(Thorne, 1961); on palms in Togo and Ghana (Luc and Hoestra,

1960), on tomatoes and papaws, Asimina triloba, in Queensland, Australia

(Colbran, 1960); and on citrus in Montserrat (Pennock, 1959). Eighteen host plants of R. reniformis. including sorghum, Sorghum vulgare, and corn, were reported from Accra, the capital of Gold Coast, West Africa

(Peacock, 1956a). Peacock (1956b) observed the reniform nematode on a wide range of food crops and found that it can maintain itself under fallow conditions on at least two common weeds (Amaranths spinous L,, and Synedrella nodiflora Gaertn) in the Gold Coast. In Trinidad, West

Indies, it is a pest of sweet (Brathwaite, 1972) and tobacco

(Singh, 1974). In India, the reniform nematode attacks castor, Ricinus communis L. and tomato (Nath et al., 1969), coffee roots (Siddiqi and

Basir, 1959), and tobacco (Gopalachari, 1978; Patel, 1986). In

Venezuela, it occurs in great numbers on tobacco (Heald and Meredith,

1987).

Birchfield and Brister (1962) tested 43 agronomic plants and listed

11 plants as non-host (barnyard grass, Echinocola crus-galli. dallis grass, Paspalum dilatatum. mustard, 'Florida', . ,

'Fulghum', Avena sativa. onion, 'Evergreen*, Allium cepa, pepper of two species, sweet, bell, 'California Wonder', annuum, and red, 7

hot, var. fasciculatum, rice, ’Blue Bonnet’ Oriza

sativa, sweet sorghum, sugarcane, 'C. P. 44-101', turnip, 'Purple Top',

'White Globe’, Brassica rapa and wild barley, ) and six

host plants, all legumes (crimson clover, Trifolium incarnatum, red

clover, Trifolium spp. , white clover, hairy vetch, Vicia vi1losa.

Singletary pea, Lathyyrus hirsutus. and Spanish peanuts, Arachis

hypogaea). Nath et al. (1969) studied 15 plant varieties, representing

11 species within five families, and found that castor, Ricinus communis

L,, and tomato were the two most susceptible hosts of the reniform nematode.

Disease Interact ions:

The first research report of a disease of cotton (Atkinson, 1892) dealt with the vascular wilt, oxysporum f. vasinfectum, and the pronounced effect of root-knot nematode on the expression of this disease. Smith (1954) reported that in addition to direct yield loss, nematodes provide an opening to the vascular system of cotton plants for the wilt pathogen, F. oxysporum f . vasinfectum, and thus increase their susceptibility. Martin et al. (1956) reported that root-knot nematode, M. incognita and M. incognita acrita, significantly increased the incidence of Fusarium wilt in the cotton cultivars 'Deltapine 15'

(wilt-susceptible) and 'Coker 100 Wilt' (wilt-resistant). Minton and

Minton (1963) observed abundant F. oxysporum f. vasinfectum growth in M. incognita acrita-induced giant cells as well as in the xylem. The apparently entered the xylem through decaying tissue (Minton and

Minton, 1963). Shepherd (1970) reported that root-knot nematode 8 resistance was a major factor in resistance of cotton plants to Fusarium wilt.

Smith and Taylor (1941) reported that a high infestation of the reniform nematode occurred in the regional wilt plots on the roots of both cotton and cowpeas collected from Baton Rouge. This was the first published record of the species having been found on cotton in association with Fusarium wilt. Further reniform nematode-wiIt association was reported by Neal (1953). He later indicated that increased incidence of Fusarium wilt in a susceptible variety of cotton in the Baton Rouge area was dependent on the presence in the soil of high populations of the reniform nematode (Neal, 1954), Jones et al.

(1959) found that reniform nematode increased wilt development on wilt- susceptible cotton varieties, but not on wilt-resistant ones. The presence of reniform nematodes resulted in a marked increase in the percentage of wilt infection, and the increase was more than doubled in wilt-susceptible varieties (Khadr et al., 1972).

Brodie and Cooper (1964) reported that cotton seedlings grown in soil infested with any one of three Meloidogyne spp. or R. reniformis or

Hoplolaimus tylenchiformis were susceptible to the Rhizoctonia solani longer than were seedlings grown in nematode-free soil, and that prolonged susceptibility to R . solani was associated with reduction in seedling growth. Reynolds and Hanson (1957) reported that an increase in post-emergence damping-off of cotton, caused by Rhizoctonia solani

Kuhn, was associated with an increase in the incidence of M . incognita acrita Chitwood and a corresponding decrease in plant size and survival. 9

Histopathology:

The responses of root tissues of various crop plants to

by Rotylenchulus reniformis are well documented (Birchfield, 1962 and

1972; Carter, 1974 and 1981; Cohn, 1973 and 1976; Heald, 1975; Linford

and Oliveira, 1940; Nath et al., 1969; Rebois et al., 1968 and 1970;

Robinson and Orr, 1980; Yik and Birchfield 1979 and 1981). Linford and

Oliveira (1940) originally reported feeding of the R. reniformis on

cowpea roots to be in the cortex. A later study (Birchfield, 1962)

indicated that young R. reniformis initiated infection in young roots of

upland cotton by extending the anterior portion of their bodies through

epidermis and cortical parenchyma to feed in phloem tissue. Sivakumar

and Seshadrin (1972) made similar observations on tomato and papaya.

Carica papaya. Birchfield1s observations were further supported by Nath

et al. (1969), who considered R. reniformis a phloem feeder on tomato

and castor, and reported that the cells around the nematode head

disintegrated to form a cavity. Browning and necrosis of root tissue

away from the initial feeding site was also reported (Birchfield, 1962;

Nath et al., 1969). Studies with other plants (, bean, pea, corn, and sugarcane) led Birchfield (1972) to conclude that the parasite preferred the pericycle of all plants studied except corn and sugarcane.

Cohn (1973 and 1976) observed a consistent pattern of parasitism by

R. reniformis in several species of plants including cotton (G. hirsutum). He observed that R. reniformis cosies to rest in the endodermis and induces a syncytium composed solely or mainly of pericycle cells extending around the roots to either side of the initial 10

feeding cell. Affected cells were described as hypertrophied,

uninucleate, and with intact cell walls. Oteifa (1970) found that R.

reniformis caused hypertrophy of pericycle cells in cotton roots and

noted that this symptom was the major histological response for the

debilitated root system. Rebois et al. (1970) and Oteifa & Salem

(1972), using soybeans and Egyptian cotton, G. barbadense respectively,

as hosts, designated the pericycle as the feeding site of the nematode

and described formation of giant cells in this region of the root. In a

subsequent electron-microscopic study on soybean, Rebois et al. (1975)

described R. reniformis as generally stopping with the lips pressed

against an endodermal cell which they termed the prosyncyte. This event was followed by hypertrophy of the prosyncyte and the adioining pericycle, a distance of approximately 3-10 cells away. Affected cells were essentially uninucleate but comprised a syncytium formed by the coalescing of cytoplasm after the partial dissolution of radial pericycle walls. Heald (1975), working concurrently with cantaloupe,

Cucumis melo cantalupensis, also described R. reniformis as generally feeding on an endodermal cells with pericycle involvement. The feeding cell and pericycle cells were uninucleate with prominent nucleoli.

Similar observations on sunflower, Helianthus spp., were reported by

Robinson and Orr (1980), where affected cells were essentially uninucleate and the feeding cell or prosyncyte was endodermal in 23 of

28 cases.

Yik and Birchfield (1979) reported that in sweet potato, the reniform nematode juveniles penetrated intercellularly and fed in the single-layered endodermis. A single endodermal cell at the nematode 11 head hypertrophied into a giant cell. The uniseriate pericycle adjacent to the giant cell reacted to the infection and hypertrophied into a curved sheet of syncytia encompassing 7-10 cells on either side of the

infection site. The infected pericycle cells showed dense cytoplasmic contents.

In susceptible cotton plants, Yik and Birchfield (1981) observed typical pericycle hypertrophy, enlarged nuclei and nucleoli, cell wall dissolutions, and granular cytoplasm, whereas in resistant plants, endodermal and pericycle cells were collapsed or killed at feeding sites

(hypersensitivity) or syncytial development was restricted. Yik (1981) observed enlargement and hypertrophy of the entire pericycle in susceptible cotton plants.

Resistance Mechanisms:

The most apparent sign of resistance is a rapid death of host cells

immediately adjacent to the nematode head (Dropkin, 1980). Dropkin

(1980) found that phenols occur in higher concentrations in resistant plants. After the feeding has begun and the reniform nematode females become sedentary, they are subject to any changes in the host (Powell,

1971). Toxic metabolites of the resistant host or starvation may cause death of sedentary nematodes (Carter, 1974; Yik, 1981).

Resistance in plants is due to defense mechanisms which either restrict penetration or inhibit reproduction of the parasite (Montasser,

1986). Rohde (1960) indicated that resistance in plants was dependent upon physiological differences, which rendered a plant unable to respond to nematode stimulation and, therefore, unable to meet the metabolic 12 requirements of the parasite, or the inactivation of the stimulatory substances released by the parasite. In some cases, toxic substances were released by resistant plants which slow down plant parasite interactions (Rohde, 1960). The general metabolic response of plants to infection by nematodes or other disease agents is the accumulation of phenols and the buildup of oxides. Phenols are oxidized to quinones, which in turn polymerize, forming complex products such as tannins, lignins, and melanins, giving necrotic tissue its characteristic brown color (Rohde, 1972).

Riddle and Bird (1985) stated that responses to chemical attractants may affect the distribution of the reniform nematode in its natural environment. They observed that second-stage (J2 ) reniform nematode juveniles were attracted to several inorganic salts, cyclic

AMP, AMP, and germinated tomato seeds. The order of attractiveness was

Cl” > Na+ > C2 H302” > Mg2+, NH/(+, S0A2"; and 3, 5 cyclic AMP was a stronger attractant than 5 AMP. The reniform nematodes developed rapidly in the presence of high amounts of potassium (Dropkin, 1980).

Large populations of reniform nematode cause nutritional imbalances in host plants by reducing nitrogen, potassium, and manganese (Dropkin,

1980).

Life Cycle:

The life cycle of the reniform nematode varies with host plant and environmental conditions. It differs from the root-knot nematode in that the J2, J3 , J4 juveniles and males do not feed, and sex ratio is not regulated by post infection factors (Veech, 1984). The infective 13

stage is the immature female that starts egg production after 8-9 days

of infection (Birchfield, 1962). Eggs (50-80) are deposited in a

gelat inous matrix that cover the ent ire protruding port ion of the female

body (Heald et al., 1981). The mean number of eggs produced on cotton

in 22 days is 66 per egg mass (Peacock, 1956b). Sexual reproduction

occurs and non-parasitic males are often seen coiled around parasitic

females (Veech, 1984). Bird (1983) reported that the freshly hatched

second stage juvenile takes two weeks to complete moulting and to become

a mature male or immature female. The reniform nematode takes 29 days

to complete its life cycle on castor seedling at a temperature range of

30-32°C (Nath et al., 1969). Its life cycle is completed in 17-23 days

on cotton at room temperature (Birchfield, 1962), and in 15 to 22 days

on soybeans (Peacock, 1956b). Peacock (1956a) observed that the life

cycle of the reniform nematode on soybeans, in Gold Coast, West Africa, was shorter than that described by Linford and Oliveira (1940) on

cowpeas in Hawaii.

Factors Affecting Reniform Nematode Populations:

Ayala and Ramirez (1964) suggested that humidity, elevation, temperature, and soil pH were not limiting factors in the occurrence and distribution of the reniform nematode. It occurs more in loamy soils but clayey and sandy soils with little organic matter support large numbers if host plants are suitable. Baker and Olthof (1976) Indicated that the tolerance of a crop to a given species of nematode varies with several environmental factors, such as soil type, moisture, temperature, host nutrition, and the presence or absence of microflora and 14 microfauna, as well as age and size of plants, and population or race of the nematode present.

Hollis's (1963) theory stressed the importance of population levels, the effects on young plants, and the biological control of plant-parasitic nematodes, and suggested designs for basic tests of their parasitic and pathogenic action. According to Hollis, small grasses are resistant to root-knot and reniform nematodes because their roots are of insufficient diameter to support development and reproduction of the females.

Smith (1954) reported that nematodes are a major cotton disease problem on the lighter soils of the rain belt and the lighter soils of the irrigated regions. However, Birchfield et al. (1966) reported that reniform nematodes occurred frequently on fine-textured Harlingen,

Laredo, and Cameroon alluvial clay loams near the Rio Grande River, and less frequently and in fewer numbers on the coarser-textured sands farther from the river. However, these nematodes occurred to some extent in most soil types in the area.

Rebois et al. (1968) examined soil samples for nematodes from soybean fields in 28 major soybean producing counties in Alabama, North

Florida, and Georgia. They observed that the number of genera and the total number of plant parasitic nematodes recovered per pint of soil were higher in the light textured soils than in the clay soils.

The effect of soil water content on R. reniformis infectivity was investigated by Rebois (1973b) on 'Lee' soybean roots in an autoclaved sandy clay loam soil. Nematodes were introduced into soil masses maintained at constant soil wetness levels from 3.4 to 193! by weight. 15

Nematode infectivity was greatest when the soil water content was

maintained just below field capacity in the 7.2 (-1/3 bar) to 13.0% (-

1/7 bar) ranges. Nematode invasion of roots was reduced in the wetter

15.5 (-1/10 bar) to 19.0J[ (-1/20 bar) soil moisture ranges and in the

dryer 3.4 (-15 bar) to 5.8% (-3/4 bar) soil moisture ranges.

The effect of soil temperature on parasitism and development of R.

reniformis on resistant ('Peking' and 'Custer') and susceptible ('Hood'

and 'Lee') soybean cultivars was also studied by Rebois (1973a). He

maintained soil temperatures of 15, 21.5, 25, 29.5 and 36°C i 1 in

temperature tanks in greenhouse. R. reniformis developed best at 25 and

29.5°C. The female life cycle can be completed within 19 days after

inoculation under favorable conditions at 29.5°C that is higher than the

temperature (21.5°C) at which plant root growth was best. During a 25- day period, no egg masses were present on nematodes feeding on roots grown at 15°C and 36°C.

Population Dynamics:

Bird et al. (1973) observed that reniform nematode populations

increased rapidly from June to August, with populations of 100/g of soil as late as the end of October. They reported that populations started declining slowly with the decrease in feeder root production. They further stated that the rate of decline increased in spring until 45 days after planting a new crop. Minton et al. (1960) studied population build-up and pathogenicity of reniform, root-knot, lance, and spiral nematodes on cotton, soybean, and tomato in field bins. They reported that the average reniform nematode count was 132,000/pint of soil one 16

year after infesting the duplicate field bins, following eradication of

existing nematodes. Minton (1964) determined population dynamics of

seven nematode species including R. reniformis, in concrete-bordered

bins containing Independence loamy fine sand, under 10 cotton

selections. He observed that R. reniformis attained its maximum

population in two years and its numbers were the highest of seven

nematode species studied. He further stated that maximum populations of

all nematode species studied occurred during late fall or early winter,

and minimum populations occurred in late spring.

Economic Threshold:

McGawley et al. (1985) indicated that the reniform nematode 3 threshold is between 500 and 750 individuals/500 cm of soil at the time of planting soybeans, and suggested that threshold values may be lower

if plants are under stress induced by other pathogens or by unfavorable environmental condit ions. Thakar and Yadav (1986) reported that damaging levels for the susceptible and resistant varieties of pigeonpea, Cajanus cajan. were 1,000 and 10,000 juveniles of reniform nematode per 700 g of soil, respectively.

Survival:

Blrchfield and Martin (1970) reported that reniform nematodes were still present and capable of infecting sweet potatoes 668 days after the soil was air-dried (3.33S moisture) and stored in a covered can. The supply of air-dried soil was exhausted after 6 6 8 days. The survival time was 1,276 days in moist stored soil (10.2Z moisture). Reniform nematode 17 eggs were resistant to drying and have hatched after two years storage in soil at 5% moisture (Heald et al., 1981).

Crop Losses:

Reniform nematode reproduced abundantly and caused severe injury to all cotton cultivars tested by Jones et al. (1959). It caused reduction in lint yield, delayed maturity, reduced boll size, and, in some years, reduced lint percent. Heald et al. (1981) stated that extremely high numbers of juveniles, males and females of the reniform nematode occur in the rhizosphere of infected cotton roots. Minton et al. (1960) observed that the reniform nematode was pathogenic on all entries of cotton and caused stunting, delayed maturity, and reduced yield. Minton et al. (1964) reported that reniform nematodes reduced seedling emergence, plant height and yield of seven G. hirsutum L., one G. barbadense L. and two G. arboreum L. (1394, 1402) genotypes tested. The

G. hirsutum entries, Auburn 56 and H 257, were tolerant with respect to yield. The nematode caused early crop maturity and increased Fusarium wilt.

A survey by Birchfield and Jones (1961) revealed that the reniform nematode was associated with cotton failures distributed over 2000-2500 acres (810-1012.5 ha) in two Louisiana parishes. The nematodes caused dwarfing, premature decay with losses of secondary roots and death of young cotton, which resulted in poor stands, grassy areas and yield reductions of 40-60% of the crop.

Cotton plants growing in soil heavily infested with reniform nematodes were dwarfed and chlorotic, and showed loss of secondary 18

roots. Young plants were occasionally killed (Lambe and Horne, 1963).

R. reniformis caused severe damage to sweet potatoes in a greenhouse

test (Martin, 1960). Williams and Birchfield (1974) found that reniform

nematodes caused root decay, unthrifty growth, and up to 10% yield

reduction in susceptible soybean cultivars.

McGawley et al. (1985) identified 26 nematode species, representing

10 genera, as potentially important parasites of soybean varieties

produced in Louisiana. They reported that soybean cyst, root-knot, and

reniform nematodes were probably the most damaging species, and that

reni form nematodes, although not as widely distributed as cyst and root -

knot nematode, can cause severe damage to soybeans. They observed two

races of reniform nematode on soybeans in Louisiana.

Chemical Control:

Nematicides and rotation with non-host crops are commonly used to

control reniform nematodes on cotton. Newsom and Jones (1955) obtained

about one-half bale per acre (269 kg/ha) yield increase of cotton lint

through control of the reniform nematode-wilt complex by fumigating with

Dichloropropene (D-D mixture) at 13 gallons (121.49 liters/ha), ethylene

dibromide W-85 at 4 gallons (37.38 liters/ha), Nemagon (1,2 dibromo-3-

chloropropane) at 0.5 and at 1.0 gallon per acre (4.67 and 9,35

liters/ha), respectively, three weeks before planting. Roman (1961)

suggested that emulsifiable soil fumigants such as Nemagon and Fumazone are less complicated and easier to use for control of nematode

parasites, including the reniform nematode, on sugarcane in Puerto Rico.

Birchfield and Pinckard (1964) observed that a combination of 19

pentachloronitrobenzene (PCNB) with 1,2 dibromo-3-chloropropane (DBCP)

reduced reniform nematode infection of cotton seedlings more than DBCP

alone, PCNB (15%), dieldrin (7.5%), and DBCP (25%), combined as a seed

treatment (NDT), 1 lb/10 lb (.373 kg/37.30 kg) of cotton seed [2 lb/acre

(1.84 kg/ha)], were more effective than might be expected with such a

small amount of DBCP. They further found that both DBCP and NDT

increased yields of seed cotton. DBCP-T (DBCP and PCNB) most

effectively reduced nematode infection but failed to increase seed

cotton yield.

Birchfield (1968) observed that the best nematode control and highest cotton yields were obtained with the following nematicides: 1,2 dibromo-3-chloropropane; 1,3-dichloropropene mixture; 1,3- dichloropropenes and related C^ hydrocarbons; 0,0-diethyl S-{2-

(ethylthio)ethyl] phosphorodithioate-PCNB mixture; 2-methyl-2-

(methylthio) propionaldehyde 0 -(methyl-carbamoyl) oxime; 0 ,0 -diethyl 0 -

((£-methylsulfinylJphenyl) phosphorothioate; 0 and ethyl S ,S-dipiropyl phosphorodi-thioate.

Thames et al. (1969) reported that cotton yields were significantly increased by fumigation, and post-treatment nematode counts were negatively correlated with yield in one year. In the following year, yields were significantly different only in one trial, and post- treatment counts were negatively correlated with yield in three trials.

Thames and Heald (1974) tested deep (20-inch) and shallow (10-inch) placement of soil fumigants, in paired trials, for control of R . reniformis on cotton following cotton, and on cotton following sorghum.

They obtained significantly higher seed cotton yields from chemical 20 treatments than checks, when cotton followed four years of continuous cotton. In 1969, pretreatment nematodes were 10 times greater when cotton followed cotton than when cotton followed grain sorghum.

However, in 1970, there were no significant differences in yields between soil fumigants and checks when cotton followed one year in cotton. Nematode counts were negatively correlated with yields unti1 near harvest, when correlation changed to positive.

Rotat ions:

Cotton following sorghum, planted on reniform-infested land made normal growth in south Texas (Lambe and Horne, 1963). Population dynamics of the reniform nematode varies with the crop and soil fumigation (Brathwaite, 1974). Corn is as effective as fallow in reducing the nematode population of soil to levels which may not be economically damaging to sweet potato (Brathwaite, 1974). Rebois and

Webb (1979) suggested the use of reniform nematode-resistant potato cultivars as a primary or rotated crop in reniform nematode problem areas. Williams et al. (1983) demonstrated that growing a resistant soybean cultivar ('Pickett 71') for two consecutive years would eliminate the expense of fumigation for reniform nematode control when growing cotton or a susceptible soybean cultivar in the following year.

Soil Solarization;

Heald and Robinson (1987) evaluated soil solarization for control of R. reniformis in the lower Rio Grande Valley of Texas. They observed reduced soil nematode population densities 0-15 cm deep and increased 21 yields of lettuce and cowpea (Vigna unguiculata) with soi1 solarization.

The time required for 9 Q Z mortality in the laboratory varied from 25 to less than 1 hour between 41°C and 47°C. In water, juveniles and eggs required up to 10 days to recover from sublethal thermal stress, and lethal time-temperatures in laboratory were in general agreement with field results (Heald and Robinson, 1987),

Host Plant Resistance:

Baker and Olthof (1976) pointed out that the concepts of host efficiency and host sensitivity often are characterized by the terms resistant, susceptible, tolerant, and intolerant. They further stated that since reference to two variable phenomena with a single term is often confusing, they suggested that usage of either term should include a qualification, indicating whether or not the reference is to plant damage or to pest reproduction. Lim and Castillo (1979) used root necrosis, nematode recovery from roots and soil, and egg production as criteria for resistance or susceptibility to the reniform nematode in soybeans, and suggested that any parameter could be used to identify resistance. Egg production is more reliable as a measure of host resistance than invasion or number of females per root (Gaur, 1986).

Varying degrees of resistance to reniform nematode on different crops has been reported in recent years. Two of 53 cultivars (cvs) of mungbean, Vigna radiata (Patel and Thakar, 1986); five of 17 cvs of green gram, Phaseolus aureus (Vigna radiata); and three of 17 cvs of black gram, Vigna mungo (Routaray et al., 1986) showed resistance to R. reniformis in India. Montasser (1986) evaluated 20 tomato cvs for their 22

reaction to R. reniformis and rated cvs 'Peto-108', 'Peto-95r and 'Nema-

1400' as highly resistant, and 'Ace-551 and 'Petopride' as resistant

hosts. Gaur (1986) tested seven cvs of cowpea and one of green gram (cv

'PS-16') and confirmed the latter to be susceptible in terms of

invasion, development, and egg production. However, cowpea cvs were

resistant at one or more of these stages; cv 'RC-48' was highly

resistant based on egg production (Gaur, 1986). Sahoo et al. (1986)

reported that 7 of 49 bengal gram (Cicer aretinum) cvs were resistant,

while another 20 were moderately resistant. Out of 60 cowpea 1ines,

only 11 were moderately resistant based on pot inoculation and root

staining with acid fuchsin/lactophenol 12 days later (Patel et al.,

1986), Rebois and Webb (1979) reported that cvs of potato, La Rouge and

Red La Soda, showed high levels of resistance to the reniform nematode

19 9 and the resistance was not linked to H , H , or H genes for resistance

to races of potato cyst nematodes, Globodera rostochiensis and G.

pallida. Three tobacco cvs evaluated by Heald and Meredith (1987) were

suitable hosts of the reniform nematode, and M . incognita-resistant

'NC95' tobacco was susceptible to R. reniformis.

Resistance to reniform nematode in tomato (PI375937) is controlled

by at least one dominant gene, which may be closely linked to one or more genes for resistance to Heterodera schachtii (Rebois et al., 1977).

Martin et al. (1966) reported that of 24 sweet potato selections, the cultivar 'Goldrush', susceptible to M . incognita, was the least suitable host of reniform nematode.

Six of 43 soybean cvs currently recommended by the Louisiana

Cooperative Extension Service are resistant to reniform nematodes, 13 to 23 cyst nematodes, and 12 have root-knot nematode resistance (McGawley et al., 1985). Whereas, several cvs of soybeans contain resistance to two of these three nematodes, the cvs 1 Forrest1 (Group V) and 'Centennial*

(Group VI) contain resistance to common races (race 1 of reniform nematode and race 2 of root-knot nematode) of all three (McGawley et al., 1985). Soybean cyst nematode-resistant cvs, Pickett 71 and ’Dyer', were also resistant to the reniform nematode; however, resistance to R. reniformis and M. incoRnita were not related (Rebois et al., 1970;

Rebois et al., 1968). Birchfield et al. (1971) reported that all soybean cyst nematode resistant breeding lines were not resistant to the reniform nematode. He further indicated that genes controlling resistance to the two nematodes in soybeans were separate but probably linked. Birchfield and Brister (1969) confirmed a high degree of resistance in Pickett 71 and Dyer of soybean cvs. Harville et al.

(1985) reported that reniform resistance in soybeans is controlled by two pairs of genes with unequal effects, and cvs 'Dare* and Pickett 71 are moderately-resistant and resistant, respectively.

Birchfield and Brister (1963) studied 24 cotton cvs, lines, and crosses for infection and reproduction of R. reniformis. They found that all plants were susceptible to infection and nematodes reproduced on all. However, some were more susceptible than others based on numbers of infect ing nematodes. Some of the least suscept ible ones included Auburn 56, 'DPL 15', 'Stoneville 7A* , 'Acala', and 'Dixie

King1 (Birchfield and Brister, 1969). Immune G. longlcalyx did not support the development of a penetrating reniform nematode female (Yik and Birchfield, 1984). Based on female development and egg production 24

Yik and Birchfield (1981 and 1984) reported that G. stocksi i. G.

somalanse and G. raimondii It9 were highly resistant or resistant; the G.

barbadense race stock Texas 110 and the G. hirsutum race stocks Texas

893 and Texas 903 were highly resistant or resistant; and the G. arboreum genotypes P.I. 417895, P.I, 417891, and CB 3839, and the G. herbaceum genotype P.I. 408775 were highly resistant or resistant.

Carter (1974 and 1981) reported that G. arboreum 'Nanking' (C.B.

1402) was highly resistant to the reniform nematode. Minton (1964) also observed lower numbers of reniform nematode juveniles on G. arboreum

(1394 and 1402) as compared to G. hirsutum (Clevewilt 6-3-5, Auburn 56,

H257, Coker 100A, Empire 6 -8 , Deltapine 15, and Mexican Wild-Jack Jones) and G. barbadense var. darwinii.

Two Fjq lines of G. hirsutum, designated A 623 and A 61, having the highest known resistance to root-knot nematodes were developed by

Shepherd (1974). A 623 originated from an family of Clevewilt-6 X

Mexico Wild-Jack Jones (a root-knot tolerant primitive G. hirsutum from

Mexico). Both parents were identified by Jones et al. (1958) as sources of resistance to root-knot. A 61 was developed from an F^ family of

Hybrid 257 X Mexico Wild-Jack Jones. Both A 623 and A 61 were transgressive segregates for resistance. Root-knot resistance in the Fj generation from resistant A 623 X susceptible 'Stoneville 213,' 'Coker

201,' and 'Dixie King II' was incompletely dominant. Fusarium wilt

[Fusarium oxysporum Schlecht. f. vasinfectum (Atk.) Snyd. & Hans.] resistance in the above material was highly associated with root-knot resistance (Shepherd, 1974).

Shepherd (1982a) reported that cotton line Auburn 623 RNR without 25

fumigation was more effective for controlling root-knot nematodes than

Auburn 56 and Deltapine 16 with fumigation and was equally effective as

those cottons with fumigation for controlling Fusarium . He

suggested that if the high level of resistance exhibited by Auburn 623

RNR can be bred into cotton cvs, it should reduce much of the present

root-knot nematode and Fusarium wilt damage to cotton and to susceptible

crops following resistant cottons in rotations.

Three breeding lines of cotton (G. hirsutum L.), Auburn 566 RNR,

Auburn 612 RNR, and Auburn 634 RNR, each with exceptionally high

resistance to root-knot nematodes and Fusarium wilt were developed using

A 623 as the resistant parent and released cooperatively by ARS-USDA and

the Alabama Agric. Expt. Stan. (Shepherd, 1982b). The narrow genet i c

base of A634 led Shepherd (1983) to evaluate 471 primitive race stocks,

18 (3.82) of which were resistant. However, none was as resistant as

Auburn 634 RNR, although several approached it.

The following 12 (4 BC3F4 and 8 BC2F4 ) root-knot nematode

resistant, noncommercial, flowering germplasm 1 ines of upland cotton

involving G. hirsutum L. race accessions were developed and released by

Shepherd and co-workers (Anonymous, 1987) from the Agricultural

Research Service of the United States Department of Agriculture and the

Mississippi Agricultural and Forestry Experiment Station: M 019-RNR, M

022-RNR, M 025-RNR, M 026-RNR, M 027-RNR, M 028-RNR, M 070-RNR, M 075-

RNR, M 078-RNR, M 188-RNR, M 487-RNR, and M 495-RNR. M 019 (BC2FZ(), a day-neutral (DN)-converted flowering line, was developed by recurrently backcrossing photoperiodic primitive Texas race stock 19 (race 26

richmondii), collected from Chiapas, Mexico, to ’Deltapine 16' with the

former serving as the male parent (Anonymous, 1987).

In a greenhouse test based on egg production per gram of root,

Beasley (1985) observed that G. hirsutum L. line La 434-1031-4-4 (La.

RN 4-4) was resistant to R. reniformis. According to him, the following race stocks and day-neutral converted race stocks (seed of which were obtained from Shepherd) also expressed resistance: TR 19, DN-converted

TR 19, TR 26, DN-converted TR 26, DN-converted TR 75, DN-converted TR

78, TR 176, DN-converted TR 176 and Texas 110. G. longicalyx was immune, or nearly so, and data from G. longicalyx X G. hirsutum triploids and hexaploids suggested that immunity behaved as a dominant character. He further reported that in a field test, Auburn 80-180,

Auburn 634, La 434-1031-810909 (La. RN 909), and La 434-1031-810910 (La.

RN 910) were judged to be resistant on the basis of egg production, juvenile population, and effects on yield components.

Four cotton, G. hirsutum L., germplasm lines with resistance to reniform (R. reniformis Linford and Oliveira) and root-knot nematode have been released by the Louisiana Agricultural Experimental Station

(Jones et al., 1988): La. RN 4-4, La. RN 909, La. RN 910, and La. RN

1032. These lines also exhibited resistance to Fusarium wilt, good yielding ability, and enhanced fiber quality equal to or greater than that of 'Stoneville 825* and 'Deltapine 41'. These agronomically enhanced breeding lines represent the first germplasm releases of cotton with known resistance to reniform nematode. The four germplasm lines were developed from selections within La. 434-RKR (root-knot resistant).

LA. 434-RKR originated from a cross of Bayou 7769 X Deltapine 16. Bayou 27

7769 is a RKN-resistant, high fiber quality selection from a cross of

'Deltapine 15' X 'Clevewilt-6 1 (Jones et al., 1988).

Smith (1954) found that root-knot nematode resistance in cotton is

inherited recessively and may be polygenic. Resistance to the root-knot nematode in the wild G. barbadense var. darwinii type is controlled by

two recessive genes (Malo, 1964). Jones et al, (1958) reported that

resistance to root-knot nematode in cotton was inherited in a quantitative manner, greatly influenced by environment, and controlled

by atleast two pairs of genes.

Information regarding the inheritance of reniform nematode

resistance in cotton is very limited. The present study represents the

first genetic study of reniform nematode resistance in cotton (Muhammad and Jones 1988).

Techniques for Evaluation of Host-Plant Resistance:

Several techniques were reviewed for greenhouse evaluation of test plants (Beasley, 1985; Beasley and Jones, 1985; Bugbee and Sappenfield,

1972; Carter, 1981; Gaur, 1986; Harville at al., 1985; Khadr et al.,

1972; Rebois, 1973a and 1973b; Rebois et al., 1970; Shepherd, 1983;

Turcotte et al., 1963; Williams et al., 1979; Yik and Birchfield, 1984).

To evaluate resistance of different crop plants to nematodes, researchers have used different inoculum levels, times of inoculation, length of growth period of plants, and inoculation methods.

Yik and Birchfield (1984) inoculated cotton seedlings three days after transplanting with 2,000 R. reniformis young females, males and 28 juveniles, obtained from a reniform nematode-infested field. Plants were grown for an average of 35 days at 20-32°C. Entire root systems of the test plants were harvested to measure egg production. The soil was removed by soaking the roots in water without injuring them. Roots were blotted dry with paper towels and weighed. Roots with egg masses were cut into 1 cm lengths and placed in 0.5% sodium hypochlorite solution for 10 minutes to free the eggs from the egg matrix. Roots were blended for 5 seconds to dispense the eggs. Eggs were separated from the root debris with a 45-pm mesh sieve, collected on an 18-pm mesh sieve, and washed with tapwater to remove the hypochlorite. Eggs were suspended in

100 ml of water from which two 10 ml aliquots were counted, and the mean counts corrected for 100 ml. Eggs per gram of root was determined for each plant.

Rebois (1973b) inoculated soybean plants 15 days after planting with 400 and 800 reniform nematodes (juveniles, males and infective females) in Tests 1 and 2, respectively. At 15 and 10 days after inoculation in Tests 1 and 2, respectively, plants were removed from the cups, roots and tops were weighed and the roots were fixed and stained with lactophenol-acid fuchsin to facilitate counting of nematodes and eggs. Nematodes were extracted from the entire soil mass by the elutriation method. He used total number of nematodes per root system and per gram of root as criterion for measuring the invasion rate, since the post inoculation periods were too short for significant second- generation hatch and invasion of roots (Rebois, 1973a). He used the numbers of mature females per root, mature females per gram of root, and eggs per egg matrix as indicators of nematode development rates. 29

Rugbee and Sappenfield (1972) inoculated plants 2 or 6 weeks old with F. oxispormn f. vasinfectum, Verticillimn albo-atrum, or M. incognita acrita (2 or 4 juveniles/ml in 50 ml samples), individually and in all combinations, to determine an inoculum procedure that would permit an evaluation for multiple disease resistance. Four cotton varieties of known disease reaction were used for comparison.

Differences in varietal responses to F. oxlsporum were best expressed in the younger plants, and to V. albo-atrum in older plants. Most stunting occurred when plants growing in root-knot nematode-infested soil were inoculated with either or both pathogens. However, only the most susceptible and most resistant varieties could be distinguished when inoculated with all three pathogens.

Turcotte et al. (1963) evaluated cotton plants for root-knot nematode resistance after AO days of planting. Each plant was examined and given a relative root-knot index rating from 0 to 4: 0 represented no infection; 1 , a trace to light galling; 2 , light to moderate galling;

3, moderate to heavy galling; and A, heavy to severe galling. Plants with a rating of 1 and 2 were considered resistant, and those with higher rating were considered susceptible.

Carter (1981) grew 15 seedlings, 5 per 15 cm plastic pot, of each of 12 cultivars of G. arboreum (CB 1402, 27, 32, 41, 20, 36, AA, 30, 42,

28, 47, 16) and G. hirsutum (Deltapine 16). He infested each pot with

5,000 reniform nematode juveniles per seedling with a syringe 4 cm into the soil next to each 10 day old seedling, and harvested all plants 30 days after inoculation; he weighed individual root systems; counted numbers of females, egg bearing egg masses, and numbers of eggs per egg 30 mass: and converted the counts to number per gram of root. He suggested that the use of resistance rating, based on egg production, is potentially useful in categorizing resistance to R. reniformis.

Shepherd (1983) used methyl bromide-fumigated soil in 7.6 X 7.0 cm pots, wetted and placed on greenhouse benches. About 8,000 root-knot nematode eggs were deposited into a hole 2 cm deep in the center of each pot . Eggs were covered with dry soil which was wetted immediately.

Seven to 10 days later, a newly emerged cotton seedling was transplanted into each pot. About A0 days after transplanting, soil was washed from roots, and root-knot nematode egg masses on roots were counted.

Gaur (1986) suggested that uniform pre-sowing inoculation more closely simulates field conditions than post-germination or post­ planting inoculation. Khadr et al. (1972) used natural and steam- sterilized loamy soil to test the effect of R. reniformis alone or in combination with the cotton wilt pathogen on Egyptian cotton varieties.

In the first experiment they transferred natural field soil infested with about 3,000 reniform nematodes and 250 other plant parasitic nematodes per pint of soil to a wooden box measuring 105 x 65 x 45 cm

(internal dimensions). Another box was filled with steam-sterilized soil. They inoculated the soil in both boxes, maintained in the greenhouse at a temperature of 25 to 30°C, with cotton wilt pathogen at the rate of 10 g/kg of soil; planted 10 seeds of each of the 16 cotton varieties; and recorded the percentage of wilt infection at eight weeks after seeding cotton. In the second experiment, Khadr et al. (1972) inoculated steam-sterilized loamy soil, potted in 25 cm diameter clay pots arranged at random in the greenhouse, with a reniform nematode 31 juvenile suspension at the rate of 6,000 juveniles/kg of soil and a conidial suspension of the fungus at the rate of 4,000 spores/g of dry soil. Egyptian cotton varieties highly (six), moderately (two) and lightly (three) susceptible to wilt pathogen were grown. Treatments of the experiment, with four replications were as follows: non-inoculated control; R, reniformis only; Fusarium only; reniform nematode and

Fusarium added at seeding time; reniform nematode and Fusarium, with the fungus added at one week after seeding time. Wilt infection was observed by vascular discoloration in the roots and the base of the stem, and was confirmed by the recovery of the fungus from infected tissue on potato-dextrose agar medium.

Harville et al. (1985) planted soybean cultivars Bragg, Dare,

Davis, Pickett 71, and Fj, and F2 generations of a particular cross, 2.5 cm deep in 7.5 cm diameter plastic pots filled with reniform nematode infested soil with minimal population of 14,000 juveniles per liter of soil. Pots were placed on a layer of sand on greenhouse benches in a completely randomized block design. Each pot with one seed represented an experimental unit. Thirty days after planting, they removed individual plants, gently washed their roots, and rated them for reniform nematode infection on a scale of 0 -6 , where 0 ■ 0% of roots bearing egg masses, 1 * 1-10%, 2 - 11-20Z, 3 * 21-30Z, 4 ■ 41-50Z, and 6

« 51-100Z.

Williams et al. (1979) effectively evaluated soybean plants

(resistant Pickett 71 and susceptible 'Sohoma') for reniform nematode reaction, between 21 and 31 days after planting at juvenile populations of 1800 to 2800 per 500 cm3 of soil. Plants were sampled at six dates 32 beginning 1A days after planting, and twice weekly thereafter. An egg- mass index of 1 to 6 was used, where 1 = 0 to 10% of the roots with egg masses and 6 - 51 to 100% of the roots infected.

Rebois et al. (1970) conducted a soybean resistance screening test on a raised transite greenhouse bench filled to a depth of 12 cm with a heat-treated fine sandy loam soil. They leveled the surface of the soil bed and punched rows of holes each 1 cm deep. Holes within rows were 5 cm apart and rows were 10 cm apart. They added a 1 ml of aliquot containing approximately 1,500 surface sterilized juveniles to each hole before planting one soybean seed per hole. Each of ten soybean cultivars was divided into four replications of five seeds. Plants were grown for 7 weeks at soil temperatures ranging from 18 to 33°C. Roots were then gently washed, photographed, weighed, fixed in hot lactophenol acid fuchsin for 30 sec, and cleared in lactophenol. Juveniles and mature females with egg masses on the stained roots were counted separately. A female matrix which contained one or more eggs was considered an egg mass for resistance evaluation purposes (Rebois et al.

1970).

Beasley and Jones (1985) and Beasley (1985) collected reniform nematode juveniles from an Olivier silt loam at the Perkins Road Farm near Baton Rouge, and added 1,500 to 2,000 juveniles per pot (500 cm3) of sterile soil before sowing cotton seed. Plants were grown for a period of A0 to A9 days before counting eggs per root system and converting to eggs per gram of root. Beasley (1985) also tried evaluating resistance based on an index of reniform egg masses/root system but concluded that resistance based on relative number of eggs 33

per gram of root was a better method. He also concluded that juveniles were better than eggs for inoculation. MATERIALS AND METHODS

Four cotton lines, La. RN 910, Auburn 612 RNR, M 019-RNR, and

Deltapine 41 (Dp 41), were selected as parents for a genetic study, based on their reaction to the reniform nematode under field and greenhouse conditions. La. RN 910 is resistant to reniform and root- knot nematode (Jones et al., 1988); Auburn 612 RNR and M 019-RNR, both provided by Dr. Raymond Shepherd and known to be resistant to root-knot nematode (Shepherd, 1982b, 1983), are also resistant to reniform nematode (Beasley, 1985); Dp 41 is susceptible to reniform nematode

(Beasley, 1985). These lines differ in their yielding ability, fiber quality, and agronomic desirability.

Seed of the four cotton lines were planted in a field nursery on an

Olivier silt loam (fine-silty, mixed, thermic Aquic Fragiudalf) soil at the Louisiana Agri cultural Experimental Station (LAES) Perkins Road

Agronomy Farm near Baton Rouge, Louisiana in 1985. At anthesis, each line was self-pollinated, and single crosses between the three resistant parents and the susceptible parent (Dp 41) were made. Selfed parental and single cross seed were sent to a winter nursery in Mexico during the fall of 1985 for the production of selfed and crossed generations

[parental lines, V j * backcross of Fj to parent one (Pi^iK ancl backcross of Fj to parent two (P2 F1 )] of the three crosses, La. RN 910 X

DP 41, Auburn 612 RNR X DP 41, and M 019-RNR X DP 41 (Table 1). Seed of these generations were available for greenhouse evaluation in 1986.

The seed supply permitted four replications for each cross. Two replications of each cross were planted at each of two dates to

34 35

Table 1. Inoculation dates, days for Rotylenchulus reniformis development and air temperature regimes for three cotton crosses tested for reniform nematode resistance in the greenhouse.______

Cross Parentage Planting Inoculation Days for Air temperature number date date nematode min. max. devlopment °C----- 1 La. RN 910 X 1 12-19-1986 50$ 2 1 $$ 31$$ Deltapine Al 2 5-20 1987 32 1l 2A 39

2 Auburn 612 RNR 1 2-27-1987 Al^l 21 3A X Deltapine Al 2 5-20-1987 32* 2A 39

3 M 019-RNR X 1 11-06-1986 AO* 20 27 Deltapine Al 2 7 - 1 A-1987 32l! 25 A2

§ Inoculated with eggs.

11 Inoculated with juveniles.

§§ Average during nematode development.

facilitate individual plant observations and to test for genotype X environment interaction. Cotton seed of each generation were acid- delinted and treated with a solution mixture of captan 300 [Captan N-

(trichloromethyl)thio-A-cyclohexene-1,2-dicarboximide], Vitavax-30C

[Carboxin (5,6-dihydro-2-methyl-l, A-oxathiin-3-carboxanilide)], malathion [0,0-Dimethyl S-(1,2-dicarbethoxyethyl) phosphorodithioate], methoxychlor 300 [Methaxychlor 2,2-bis (p-methoxyphenyl)-1,1,1- trichloroethane), and Apron [Metalaxyl:N-(2,6 -dimethylphenyl)-N-

(methoxyacetyl)alanine methyl ester].

The first planting was on October 21, 1986, and successive plantings continued through the end of 1987. A block of 10 plants for each of the three non-segregating generations (Pj, P2 » and Fj), a block of 20 plants for each backcross generation (P^Fj and P2Fj)» and a block 36 of 40 plants for the generation were included in each replication.

More plants were grown for segregating generations to ensure satisfactory representation of segregates. Blocks were arranged in rows of 10 experimental units on a layer of sand on a greenhouse bench in a randomized complete block design. Each experimental unit consisted of one plant grown in an individual 1,000 cm3, plastic pot holding about

500 g of soil. Pots were half buried in sand that was kept moist to avoid hi gh soil temperatures.

A 50:50 mixture of Commerce silt loam soil and river sand was steam sterilized and used for the 12-19-1986 Planting. Subsequent plantings were in a 50:50 mixture of Olivier silt loam soil and river sand that was fumigated with methylbromide. Two seed per pot were planted, and after emergence plants were thinned to one per pot. At the first true leaf stage, each plant was inoculated with a 1 ml suspension containing a mixture of 2,000 reniform nematodes (juveniles, males, infective females) along with a few other species of mostly non-plant-parasitic nematodes. One exception to this procedure was that plants in the first planting of Cross 1 were inoculated with 3,000 to 3,500 reniform nematode eggs. Inoculum was dispensed into 5-cm deep depressions with a pipette at two locations in the root zone. Dry soil was placed into the holes before and after nematode inoculum to avoid leaching of the inoculum.

Soil from a localized area of a cotton field (breeding nursery) at

Perkins Road Agronomy Farm near Baton Rouge, Louisiana, heavily infested with reniform nematode and not contaminated with root-knot nematodes, was screened by the modified LSU method (Beasley, 1985) to collect 37 inocula. A soil-water suspension (1:4) was screened with an 80-mesh sieve to remove the debris and the suspension collected in 4 1 plastic jars. This slurry was then poured over a 325-mesh sieve to collect nematodes. The decant was collected in a 500 cm3 beaker and poured over another 325-mesh, 9 cm diameter sieve containing a piece of filter- paper. The filter-paper lined sieve was then placed in a 10 cm diameter petri dish lid, half filled with water. After 48 hours, active juveniles in various stages of development were collected and diluted to a concentration of 2,000 reniform nematodes per ml of water. A 120 liter capacity plastic can full of contaminated soil was enough to provide inoculum for 220 plants.

Plants in the greenhouse were fertilized biweekly with 20:20:20 (N-

P2O 5 -K2O) water-soluble fertilizer (0.03 g/plant), and pesticides were used to protect plants from insect and disease organisms. A period of

40 and 32 days for the first and second planting dates, respectively, were allowed for the infective females to infect and reproduce (Table

1). Extra time was allowed for the planting inoculated with eggs over those inoculated with juveniles and more days were allowed for development for winter plantings than summer plantings (Table 1). Air temperature ranges for the first planting dates of the three experiments were 21°C to 31°C, 21°C to 34°C, and 20°C to 27°C, respectively (Table

1). Air temperature ranges for the second planting dates of the three experiments were 24°C to 39"C, 24°C to 39 °C, and 25°C to 42°C, respectively (Table 1). In general, these temperature regimes are suitable for nematode development. Rebois (1973a) has reported that R 38

reniformis developed best at 25 and 29.5°C soil temperatures on soybeans

in temperature tanks in greenhouse.

Entire root systems of the plants were harvested to estimate

nematode egg product ion. Soil was removed by soaking the root bal1 in water. Each root system was held over a 20 X 25 cm sieve with 3 x 3 mm mesh while washing to minimize breakage and loss of secondary roots.

The washed root system of each plant was packed in a labeled

polyethylene bag and stored in a refrigerator. Bags of each generation were grouped to facilitate egg counting. Individual root systems were

blotted with a paper towel and weighed. Each root system was then

placed in a 125-ml flask containing 100 ml of a 5% Clorox (5.257. sodium hypochlorite) solution for 10 minutes to free the eggs from egg masses.

Roots in each of 20 flasks were agitated as a group for four minutes on an electricshaker to disperse eggs. Eggs were separated from the root debris by pouring the suspension through nested 140 and 500-mesh sieves.

Eggs on the 500-mesh sieve were then washed with tap water to remove residual sodium hypochlorite and backwashed into a beaker. The volume of the suspension in each beaker was standardized to 50 ml. A 10-ml aliquot from each beaker was counted in a graduated plastic petri dish at 15X with a stereo microscope. Then counts were corrected for 50 ml.

Data for each plant were expressed as number of eggs per root system and number per gram of root.

Statistical and Genetic Analysis of Data:

Analyses of variance were done separately for each experiment, using individual plant data of eggs per gram of root and eggs per root 39

system. Generation means were compared using least square differences.

Phenotypic variances among plants within each generation were calculated

on a single plant basis and functioned as estimates of generation

variance in the equations for broad sense heritability (H3), its

standard error, and estimates of gene numbers. Frequency distribution

tables showing sample sizes, generation means, standard deviations of

means, and coefficients of variation were prepared.

Minimum Gene Number:

The following Castle-Wright formula was used to estimate the

difference in minimum gene number for resistance to reniform in the two

parents in each experiment (from Frey, 1949).

N = ______Df______8 (S2 F2 - S 2 F})

Where:

N is the estimate of gene numbers;

D is the difference between means of the two parents;

S 2 Fj is the Fj mean square; and

S2 F2 is the F2 mean square.

Heritability:

Broad-sense heritability was estimated using Allard's approach

(from Ginkel and Scharen, 1987).

H 2 - S2F?-(S2PT + S2Po + S 2F1)/3 s 2f2 AO

McNew's equation (from Ginkel and Scharen, 1987) was used to estimate the standard error of the H2 equation:

S.E. H 2 = U *___2_ * {(S 2P 1+S2P-t+S2F1 )2 + ( S ^ ) + (S2F?) + (S2Fj)>]*■ 9 (S3F2 )2 dfF2 dfPj dfP2 dfFj

Where:

dfPj, dfP2, dfFj, and dfF2 * Degrees of freedom of the P^ , P2 , Fj,

and F2 populations, respectively. S2P^, SZP2, S2F p and S2F2 are

variances of Pj , P2 , Fj, and F2 , respectively.

Generation Mean Analysis:

For each experiment, genotypic values of number of eggs per gram of root and number of eggs per root system were partitioned into additive, dominant, and epistatic gene effects by generation mean analysis, following Gamble's (1962) method. Trigenic and higher interactions were assumed to be negligible. In the analysis of the data, population means for each experiment were calculated from the individual plant data obtained from two replications in each planting date as well as combined data over dates. The variances of the population means were used to estimate the variance of the estimated parameters. Estimates of these parameters provide an indication of the relative importance of the various types of gene effects affecting the total genetic variation of a plant characteristic. The model for a generation mean (Y) is:

Y * m + a a + B d + a2 aa + 2 op ad + P2 dd

Where: 41

m ~ overall mean effect using the F2 population mean as a

reference,

a = pooled additive gene effects,

d * pooled dominance gene effects,

aa = pooled additive X additive epistatic gene effects,

ad = pooled additive X dominance epistatic gene effects, and

dd = pooled dominance X dominance epistatic gene effects.

a and fl = the appropriate coefficients for the additive and

dominance effects for a particular generation.

Means of six populations, that is , P2 , Fj, 4 F2 , 2PjF^, and 2P2Fi were used to estimate the six parameters as follows:

m = F2

— “ a pl*l P2Fi

d s - + + 2 i Pi - 4 P2 + Fl - 4F2 2PlPl P ^ i

= + 2 1 + aa - 4F2 Pj"f 2 P2Pl

ad = - + - 4 Pi + 4 P2 PjFi *2*1

s + 2 - - dd Pi ? 2 + ?! + 4F2 4 ? ^ RESULTS AND DISCUSSION

Parental Differences in Reproduction of Reniform Nematode:

To estimate the genetic effects of reniform nematode resistance in cotton, it was essential that parental lines differed in their responses to nematode infection as measured by mean numbers of reniform nematode eggs produced.

Exper iment 1:

Results of Experiment 1 indicate that the mean number of reniform nematode eggs per root system produced on La. RN 910 at Plantings 1, 2, and combined were 72, 44, and 587., respectively, of those produced on

the susceptible parent, Dp 41 (Table 2). Mean numbers of eggs per gram of root produced on La. RN 910 in Experiment 1, Plantings 1, 2, and combined, expressed as a percent of those produced on Dp 41, were 43,

37, and 40%, respectively. Except at Planting 1, differences between

La. RN 910 and Dp 41 parent in eggs per root system and eggs per gram of root were significant at the \% level of probability. These results confirm previous findings of resistance by Beasley, 1985.

Experiment 2 :

Results of Experiment 2 indicate that the mean numbers of reniform nematode eggs per root system produced on Auburn 612 RNR at Plantings

1 , 2, and combined were 6 8 , 58, and 63%, respectively, of those produced on the susceptible parent, Dp 41 (Table 3). Mean numbers of eggs per gram of root produced on Auburn 612 RNR in Experiment 2, Plantings 1, 2,

42 43

Table 2 : Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 1 (La. RN 910 x DP 41),$

Planting dates

Generations Combined Planting 1^ Planting 2

Reniform nematode eggs per root system ( 1 0 0 0 's)

Pi (La. RN 910) 6.10c§§ 0.18c 10.83c P2 (DP 41) 1 4 .85a 0.25c 24.34a Fj 9.19b 0.62ab 16.91b F2 10.23b 0. 76a 18.98b P ^ j 7 .99bc 0.33bc 15.65b P2 Fj 9.74b 0.73a 16.50b

Px as a % of P2 58%** 72% 44%**

Reniform nematode eggs per gram of root (1 0 0 0 's)

Pj (La. RN 910) 0.73d 0 . 06d 1.27d P2 (DP 41) 2.12a 0.14cd 3.41a Fi 1.18c 0 .22bc 2.04c F2 1.59b 0.30ab 2 .7Bab P ^ j 1.24c 0.18bcd 2.30bc P2Fj 1.41bc 0.35a 2.19bc

AA Pj as a % P2 407.** 43% 37%

§ Means based on two replications in each planting and two plantings in combined analysis.

11 Inoculated with eggs.

§§ Means followed by a letter in common do not differsignificantly at 0.05 probability level according to "LSD" test.

** Significant at the 0.01 probability level according to "LSD" test. 44

Table 3: Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 2 (Auburn 612 RNR x DP 41).§

Planting dates

Generations Combined Planting 1 Planting 2

Reniform nematode eggs per root system (1 0 0 0 ' s)

Pj (Auburn 612 RNR) 4.47c11 2.96b 5.97b P? (DP 41) 7.39ab 4.33a 10.29a 2 8 6 F 1 7.85a . b 1 2 .10a f2 6.63ab 2.74b 10.42a PjFi 6 .2 1 b 2.99b 9.60a 2 8 6 P2 F 1 7.73ab . b 1 2 .0 0 a

■fr A ft ft Pj as a 7. of P2 63% 6 8 % 58%**

Reniform nematode eggs per gram of ’•oot ( 1 0 0 0 's)

Pj (Auburn 612 RNR) 0.62c 0.50b 0.75b P? (DP 41) 1.14a 0. 76a 1 . 50a

F 1 0.99ab 0.43b 1,46a f2 0.91b 0.40b 1,41a P1F 1 1 .0 0 ab 0. 52b 1.50a P2Fj 1 .16a 0.54b 1.71a ft ft ft ft ft ft P^ as a % P2 58% 6 6 % 50%

§ Means based on two replications in each planting and two plantings in combined analysis.

11 Means followed by a letter in common do not differ significantly at 0.05 probability level according to "LSD" test.

** Significant at the 0.01 probability level according to "LSD" test. 45

and combined, expressed as a percent of those produced on Dp 41, were

6 6 , 50, and 58%, respectively. In all cases, the differences between

the two parents in eggs per root system and eggs per gram of root were

significant at the 1% level of probability. These results confirm

previous findings of resistance by Beasley, 1985.

Experiment 3:

Results of Experiment 3 indicate that the mean numbers of reniform

nematode eggs per root system produced on M 019-RNR at Plantings 1, 2,

and combined were 8 6 , 99, and 93%, respectively, of those produced on

the susceptible parent, Dp 41 (Table 4), Mean numbers of eggs per gram

of root produced on M 019-RNR in Experiment 3, Plantings 1, 2, and

combined, expressed as a percent of those produced on Dp 41, were 70,

91, and 81%, respectively. Except at Planting 1, where differences

between the M 019-RNR and Dp 41 parent in eggs per gram of root were

significant at the 5% level of probability, differences between the two

parents for the two traits were nonsignificant. These results only

partially confirm previous report of resistance by Beasley (1985), who

tested M 019 against Dp 41 as day-neutral converted Texas race stock 19.

In this study, the resistance level of M 019-RNR was lower than that

reported by Beasley.

Analysis of Variance for Generations:

It was essential to have significant differences among generations

in order to proceed with generation mean analysis. 46

Table 4 : Mean reniform nematode eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 3 (M 019-RNR x DP 41).§

Planting dates

Generat ions Combined Planting 1 Planting 2

Reniform nematode eggs per root system (1 0 0 0 's)

P: (M 019-RNR) 7.27b11 ll.OObc 3. 37c F? (DP 41) 7.55b 12.72abc 3.42c

F 1 8.70ab 12.OOabc 5.57abc F 2 1 0 .11a 13.02ab 7.34a 4.95bc P 1F 1 7.76b 10.56c P2 f 1 1 0 .21a 14.18a 6.14ab

Pj as a % of P2 93% 867. 99%

Reniform nematode eggs per gram of root ( 1 0 0 0 's)

P : (M 019-RNR) 1 .50b 2. 24b 0.72c Pt (DP 41) 1 .8 6 ab 3.21a 0 . 79be

F 1 1.63ab 2.36b 0.94abc F 2 2.06a 2.98ab 1.19a P 1F 1 1.59b 2.28b 0.89abc 1.1 P2 F 1 2 .0 2 a 2.91ab lab

Pi as a 7. P2 81% 70%* 91%

§ Means based on two replications in each planting and two plantings in combined analysis.

11 Means followed by a letter in common do not differ significantly at 0,05 probability level according to "LSD" test.

** Significant at the 0,01 probability level according to "LSD" test. 47

Experiment 1:

Analyses of variance of Experiment 1 indicate that at least one of

the generations was significantly different from others at the 17. level of probability for both traits at both planting dates as well as in combined analysis (Table 5). Differences between planting dates were also significant for both traits at the 17. level of probability.

Planting 2 had 31 times more eggs per root system and 10 times more eggs per gram of root than Planting 1. These differences may have been partly caused by seasonal effects because Planting 1 was grown in early winter and Planting 2 in late spring. But, the differences were more

likely caused by inoculation methods. Plants at Planting 1 were

inoculated with reniform nematode eggs while plants at Planting 2 were inoculated with reniform juveniles. The coefficients of variation

(C.V.) were 104, 46, and 62% at Planting1, Planting 2, and combined over plantings, respectively, for numbers of eggs per root system. The

C.V.'s were 89, 50, and 63% for numbers of eggs per gram of root at

Planting 1, Planting 2, and combined over plantings, respectively. The

lower C.V, for both traits at Planting 2 was, perhaps, due to a higher infestation level, and, therefore, probably to more infection sites and fewer escapes. It is probable that inoculation with juveniles was better than with eggs even though more eggs (3,000 per ml) were used than juveniles (2,0 0 0 per ml) and 18 more days (10 more days because of eggs and 8 more days because of winter) were allowed for nematode development at Planting 1 than at Planting 2. Results of Experiment 1 also suggest that numbers of eggs per root system and numbers of eggs per gram of root were equally efficient in differentiating between 48

Table 5 : Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 1 (La. RN 910 x DP 41).

Planting dates

Combined Planting 1 Planting 2

Sources of df MS df MS df MS variat ion

Reniform nematode egRs per root system (1 0 0 0 's) A Jr Replication 1 243.23 1 2.07* 1 397.02* Planting date (PD) 1 20963.51** Generation (G) 5 194.66** 5 1.82** 5 436.02** PD x G 5 180.98** Error 394 35.25 182 0. 35 211 64. 74

C.V. 62% 104% 46%

R 2 69% 15% 16%

Reniform nematode eggs per gram of root ( 1000 *s)

Replication 1 2.91 1 0.49** 1 2.80 Planting Date (PD) 1 337.27** Generat ion (G) 5 6.18**it i 5 0.28** 5 12.30** PD x G 5 4.84 Error 394 0.81 182 0.05 211 1.47

C.V. 63% 89% 50%

R 2 63% 19% 17%

Significant at the 0.05 and 0.01 probability levels, respectively, according to F test of significance. 49 levels of nematode resistance. However, results of this experiment are unique and based on only two environments with two replications each; therefore, they may not be representative of the general situation.

Interactions between planting dates (PD) and generations (G) were significant for both variables in Experiment 1 at the 1% level of probability. Because of significant PD x G interactions, estimates of gene effects were calculated by individual planting dates as well as over combined dates.

Experiment 2 :

Analyses of variance of Experiment 2 (Table 6 ) indicate that at least one of the generations was significantly different from the others at either the 5% or 1% level of probability for both traits at both planting dates as well as in the combined analysis. Differences between planting dates were also significant for both traits at the 1% level of probability. Planting 2 averaged about three times more eggs per root system or per gram of root than Planting 1. Again these differences were probably due to seasonal environments, because Planting 1 was planted in late winter and Planting 2 in late spring. The C.V.'s for numbers of eggs per root system were 59, 53, and 63% and for numbers of eggs per gram of root, 67, 58, and 6 6% at Planting 1, Planting 2, and combined over plantings, respectively. Results of Experiment 2 also suggest that numbers of eggs per root system and numbers of eggs per gram of root were equally efficient in differentiating between levels of nematode resistance. Again, results of this experiment are unique and based on only two environments and two replications, and, therefore, may 50

Table 6 : Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 2 (Auburn 612 RNR x DP 41).

Planting dates

Combined Planting 1 Planting 2

Sources of df MS df MS df MS variat ion

Reniform nematode eggs per root system (1 0 0 0 's) •tt

C.V. 63% 59% 53%

R 2 51% 16% 27%

Reniform nematode eggs per gram of root (1 0 0 0 's) & lit Jfc A Replication 1 19.90** 1 2.70 1 21 .44 Planting Date (PD) 1 60.25** A it Generation (G) 5 1.75** 5 0.46 5 2.52 PD x G 5 1.24* Error 412 0.41 201 0.11 210 0.68

C.V. 6 6 % 67% 58%

R2 43% 18% 19%

Significant at the 0.05 and 0.01 probability levels, respectively, according to F test of significance. 51 not be representative of the general situation.

Interactions between PD and G in Experiment 2 for eggs per root system and eggs per gram of root were significant at the 1% and 5%

levels of significance, respectively. Because of significant PD x G

interactions, estimates of gene effects by individual planting dates as well as combined dates were calculated.

Experiment 3:

Analyses of variance of Experiment 3 (Table 7) indicate that significant differences occurred among generations for both traits at both planting dates and in the combined analysis. Differences between planting dates were also significant for both traits at the 1% level of probability. Plant ing 1 averaged twice the number of eggs per root system and three times the number of eggs per gram of root at Planting

2. Again, these differences were probably due to seasonal environments, because Planting 1 was planted in late fall and Planting 2 in mid summer, and the greenhouse environment was differentially affected by these dates (Table 1). The C.V.'s were 36, 75, and 49X at Plantings 1, and 2 , and combined over plantings, respectively, for numbers of eggs per root system. The C.V.'s were 53, 71, and 49% for numbers of eggs per gram of root at Planting 1, Planting 2, and combined over plantings, respectively. Thus, as in Experiments 1 and 2, tests grown under a more temperate climate (spring, fall) were less variable and more discriminating than those grown under less temperate climate

(summer, winter). Results of Experiment 3 suggest that numbers of eggs per root system were more discriminating than numbers of eggs per gram 52

Table 7: Analyses of variance of resistance to reniform nematode as measured by number of eggs per root system and eggs per gram of root of the parents and progenies of crosses in Experiment 3 (M 019-RNR x DP 41).

Planting dates

Combined Planting 1 Planting 2

Sources of df MS df MS df MS variat ion

Reniform nematode eggs per root system (1 0 0 0 's)

4 Replicat ion 1 776.98** 1 614.51 1 215.12** Planting Date (PD) 1 4095.00** Generation (G) 5 122.51** 5 66.34 5 91.01** PD x G 5 32.03 Error 416 19.54 204 19.87 211 19.06

C.V. 49 % 36% 75%

R 2 44% 19% 14% —* r— o Reni form nematode eggs per gram of root o O

Replicat ion 1 1.78 1 0.92 1 7 .92** Planting Date (PD) 1 240.31** A A Generation (G) 5 4.66 5 5.03* 5 1.25* PD x G 5 1.59 Error 416 1.31 204 2.09 211 0.52

C.V. 49% 53% 71%

R 3 44% 6% 11%

Significant at the 0.05 and 0.01 probability levels, respectively, according to F test of significance. 53 of root in evaluating nematode resistance. Again, results of this experiment are unique and based on only two environments and two replications per environment, and, therefore, may not be representative of general situation. Interaction between PD and G in Experiment 3 for both variables were not significant.

Because there was a lack of consistency in the estimates of gene effects by planting dates (environments) and because each environment was represented by only two replications, major emphasis in interpretation of data of each experiment was given to the planting date with the lower C.V. value, and to the combined analysis, but it is recognized that the confidence level of these estimates was lowered by a highly significant PD x G interaction in Experiments 1 and 2.

Frequency Distribution:

Frequency distribution, number of plants observed, standard deviation, coefficients of variation, and mean numbers of eggs per root system and eggs per gram of root in each planting date of the three experiments are given in Tables 8 to 19. Observations occurring five or more intervals away on each extreme were excluded in the frequency distribution tables as well as in each analysis. However, respective complete frequency tables are given in the appendix. Large spreads in observations, high coefficients of variation, and high standard deviations in most populations indicated tremendous environmental variation. Spread of segregating populations beyond the ranges of parents in most cases suggests the presence of transgressive segregation for susceptibility. Spread of parents close to zero scale indicate that Table 8. Frequency distribution of number of eggs per root system for six populations at Planting I, Experiment 1 (La. RN 910 x DP 41).

Frequency n x S.E C.V

Pi 11 3 2 16 .18 .14 75

?2 5 5 3 13 .25 .16 65

Pi 1 7 4 2 1 3 18 .62 .47 75

P2 12 18 16 6 3 3 5 2 1 1 2 1 1 1 1 72 .76 .81 108

PlFj 25 6 2 3 2 1 1 40 .33 .48 142

P2F! 3 7 3 6 4 1 3 2 1 30 .73 .44 60

0 .4 .8 1.2 1.6 2 2.4 2.8 3.2 3.6 4

Number of eggs per root system (1000's) Table 9. Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 1 (La. RN 910 x DP 41).

Frequency n x S.E C.V

n 3 4 1 4 4 2 1 1 20 10.83 3.89 36

P2 1 2 1 1 2 2 1 4 1 1 1 1 1 1 20 24.34 9.12 37

Fl 1 2 2 1 3 2 5 1 1 I 1 20 16.91 6.83 40 f2 1 1 2 2 4 7 4 11 9 7 7 2 7 2 5 1 1 1 2 78 18.98 8.81 46

PlFi 3 1 2 5 3 3 4 3 3 4 3 3 3 40 15.65 7.10 45

P2Fj 2 7 5 2 1 6 6 2 2 3 1 1 1 1 40 16.50 9.28 56 // 0 4 8 12 16 20 24 28 32 36 40 44

Number of eggs per root system (1000's)

on On Table 10. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41).

Frequency n x S.E C.V

Pi 8 6 2 16 .06 .03 61

?2 3 2 2 2 2 1 1 13 .14 . 10 69

Pi 4 3 3 I 2 1 1 2 1 18 .22 .14 61 f2 6 9 10 9 9 4 4 1 3 2 2 1 4 2 2 2 1 1 72 .30 .26 86

PlFi 14 9 2 5 2 1 1 2 1 1 2 40 .18 .24 130

P2 Fj 2 1 3 3 2 6 1 1 2 1 2 2 2 2 30 .35 .24 67

0 .1 .2 .3 .4 .5 .6 .7 .8 .9 1.0

Number of eggs per gram of root (1000's)

i/i o Table 11. Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 1 (La. RN 910 x DP 41).

Frequency n x S.E C.V

Pi 6 4 4 2 4 20 1.27 0.46 36

P2 1 1 1 1 2 5 1 1 I 2 2 1 1 20 3.41 1.41 41

Pi 1 2 2 2 3 2 1 1 2 1 I 2 20 2.04 1.00 49

F2 1 1 2 4 4 11 7 8 6 7 3 3 8 2 5 1 1 2 1 1 78 2.78 1.36 49

PlFi 2 2 2 5 3 4 5 2 3 4 2 4 1 1 40 2.30 1.07 46

P2 ?l 4 6 7 4 1 2 2 5 2 3 1 1 1 I 40 2.19 1.28 58 i t it 0 .6 1.2 1.8 2.4 3 3.6 4.2 4.8 6 6.3 7.2

Number of eggs per gram of root (1000's) Table 12. Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41).

Frequency n x S.E C.V

Pi 1 1 1 1 2 3 2 4 3 1 20 2.96 1.49 50

1 1 1 1 2 5 1 1 2 1 19 4.332.07 48 n 2 2 2 2 1 1 1 1 2 1 1 1 17 2.86 1.80 63 f2 8 6 7 10 5 8 5 2 3 1 7 3 1 5 3 1 2 77 2.74 2. 12 77

PlFi 2 4 5 3 6 4 2 3 1 2 2 3 1 2 40 2.99 1.8461

P2Fi 2 3 7 3 7 4 1 1 2 3 1 I 35 2.86 1.45 51

0 .8 1.6 2.4 3.2 4.0 4.8 5.6 6.4 7.2 8

Number of eggs per root system (1000's)

Ln 00 Table 13. Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 2 (Auburn 612 RNR x DP 41).

Frequency n x S.E C.V

Pi 5 2 3 6 1 1 1 1 20 5.97 3.25 54

P2 1 4 5 3 2 1 1 1 2 20 10.29 4.16 40

Pi 2 2 2 1 1 2 1 1 1 1 2 2 1 1 20 12.10 7.88 65

P2 7 10 9 8 7 9 4 6 2 1 3 6 3 1 1 2 79 10.42 6.74 65

P1F l 2 7 3 4 6 5 5 3 2 1 38 9.60 3.63 38

P2 F! 6 5 2 5 3 2 3 2 1 1 1 2 1 4 1 1 40 12.00 7.43 62 // 0 3 6 9 12 15 18 21 24 30

Number of eggs per root system (lOOO's) Table 14. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41).

Frequency n x S.E C.V

Pi 1 1 1 1 2 1 1 3 1 1 1 2 L i I 20 .50 .37 73

P2 1 1 1 3 1 1 2 2 2 2 19 .76 .33 44

Pi 1 3 1 3 1 1 1 1 1 2 1 1 17 .43 .35 82

F2 7 4 12 5 8 4 5 3 3 2 3 1 3 1 1 i 2 1 2 2 3 2 1 1 77 .40 .34 85

PlFi 6 1 2 5 2 2 1 3 2 1 3 1 1 2 2 1 1 1 I 1 1 40 .52 .40 78

P2Fi 1 3 3 2 2 3 2 3 1 2 2 3 2 1 3 1 1 35 .54 .30 55 // 0 .1 .2 .3 .4 .5 .6 .7 .8 .9 1 1.1 1.2 1.3 1.4 1.5

Number of eggs per gram of root (1000's) Table 15. Frequency distribution of number of eggs per gram of root root for six populations at Planting 2, Experiment 2 (Auburn 612 RNR x DP 41).

Frequency n x S.E C.V

Pi 2 3 2 5 4 2 1 1 20 .75 .46 61

5 2 3 1 1 4 1 20 1.50 .58 *2 3 39

Fl 2 2 1 1 2 3 1 2 2 1 1 1 1 20 1.46 .84 57

F2 1 5 9 U 6 7 9 4 5 3 5 I 2 5 1 I 2 1 1 79 1.41 .92 65

PlFi 3 3 4 4 6 3 3 3 2 1 2 3 1 38 1.50 .69 46

P2 F! 5 5 4 4 5 2 1 3 3 1 2 1 2 1 1 40 1.71 1.21 71 // i i it 0 .4 .8 1.2 1.6 2 2.4 2.8 3.2 3.8 4.4 5

Number of eggs per gram of root (1000’s) Table 16. Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 3 (M 019-RNR x DP 41).

Frequency n x S.E C.V

Pi 2 2 2 2 4 3 4 1 20 11-00 4.16 38

P2 1 2 1 3 2 4 1 1 1 16 12.71 3.55 28

Fj 1 1 1 3 4 3 1 1 2 1 19 12.00 6.44 54 f2 3 4 2 9 5 8 5 14 9 5 3 4 1 76 13.02 4.80 37

P 1 Fj 1 9 1 3 6 6 3 7 1 1 2 40 10.56 4.29 41

P2 Fj 1 1 4 1 9 3 5 5 3 3 1 1 40 14.18 4.96 35

0 3 6 9 12 15 18 21 24 27

Number of eggs per root system ( 1000's)

MO' Table 17. Frequency distribution of number of eggs per root system for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41).

Frequency n x S.E C.V

Pi 2 4 4 1 3 2 3 19 3.37 2.04 61

P2 3 3 5 3 1 2 ] 20 3.42 2.93 86

Fl 1 1 1 2 5 2 2 3 3 20 3.57 2.71 49

P2 8 8 7 8 3 3 3 8 3 5 3 4 3 2 4 1 3 1 1 1 1 80 7.34 5.57 76

PlFi 2 8 5 1 3 8 2 3 1 2 2 1 40 4.95 3.22 65

P2Fi 3 8 4 2 1 5 4 1 3 1 1 1 4 1 39 6.14 5.13 84 // 0 2 4 6 8 10 12 14 16 18 20 22

Number of eggs per root system (1000's)

O' w Table 18. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, " Experiment 3 (M 019-RNR x DP 41).

Frequency n x S.E C.V

Pi 2 6 2 1 4 2 2 1 20 2.24 1.09 48

Pz 1 1 3 6 1 2 1 1 16 3.21 1.28 40

FL 1 3 5 4 1 1 2 1 I 19 2.36 1.36 58

P2 6 9 13 8 12 5 5 3 6 2 2 1 1 1 1 1 76 2.98 1.82 61

PlFi 4 6 9 6 6 3 2 3 1 40 2.28 1.07 47

p2f. 1 2 8 4 5 9 6 1 2 2 40 2.91 1 .18 41

0 1 2 3 4 5 6 7 8 9

Number of eggs per gram of root (1000's) Table 19. Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41).

Frequency n X S.E (:.v

Pi 3 3 2 5 2 2 1 1 19 .72 .48 66

P2 2 3 2 7 3 1 1 1 20 .79 .61 78

Fl 1 1 2 5 4 1 2 2 1 1 20 .93 .51 55

P2 10 7 11 6 3 3 6 9 6 7 2 4 2 1 3 1 80 1.19 .81 69

PlFt 4 8 2 7 5 3 1 4 1 3 1 1 40 .89 .66 73

P2Fi 4 10 1 4 1 4 4 1 2 1 2 1 2 2 39 1.11 .93 83

0 .4 .8 1.2 1.6 2 2.4 2.8 3.2

Number of eggs per gram of root (1000's)

O' on 66

there was not enough opportunity for the expression of transgressive

segregation for resistance.

Number of Genes:

There is no accurate method for determining the number of segregating genes for quantitatively inherited traits. In this study, an approximate number of minimum genes that could be segregating was estimated by utilizing the Castle-Wright formula (N = D 2/8 (S2F2 - S 2F1).

Values obtained in each planting date within an experiment ranged from

-1.22 to 0.74, which suggest that there is a one gene difference between the susceptible parent Dp 41 and the resistant parents (Table 20). This is a minimum estimate, however, since this formula assumes that a maximum range exists between parents, that dominance is absent, that the gene action is additive and that each gene is equal in its effect.

Failure to meet these assumptions results in an under-estimation.

Indications of epistasis and transgressive segregation through generation mean analysis and frequency distributions suggest that most of these assumptions were not met in these experiments. Therefore, it seems reasonable to conclude that this gene number is under-estimated.

Broad-sense Heritabilities;

Broad-sense heritability (H2) estimates and their standard errors are presented in Table 21. The H2 establishes the upper limit of the narrow-sense heritability (h2), because, the numerator in the equation of the H2 includes total genetic variance as opposed to only additive variance in the case of h2. 67

Table 20: Estimates of number of gene pairs by which the two parents differed in resistance to R. reniformis in Experiments 1-3.

Experiment 1 Experiment 2 Experiment 3

Errs per root system

P lanting 1: 0.001 0. 19 -0.02 P lanting 2: 0.74 -0.14 0.00

Errs per gram of root

Plant ing 1: 0.03 -1.22 0.08 Planting2: 0.67 0.50 0.02

Table 21: Broad-sense heritabilities (Z) and standard errors (%) of resistance to R. reniformis in Experiments 1-3

'

Experiment 1 Experiment 2 Experiment 3

Errs per root system

Plant ing1: 86 ± 4 28 ± 2 -3 ± 0.6 P lanting 2: 38 + 0.3 34 ± 0.2 78+0.1

Errs per Rram of root

Planting 1: 85 ± 32 -6 ± 115 53 + 2 P lanting2: 42 ± 4 51 ± 7 56 + 8 68

Estimates of broad-sense heritability ranged from 0 to 86 % with a mean of 53% for eggs per root system and 0 to 85% with a mean of 57% for eggs per gram of root.

Generation Mean Analysis:

Experiment 1:

In Experiment 1, involving the parents La. RN 910 and Dp 41t estimates of genetic components of genotypic values indicated that additive gene effects significantly reduced eggs per root system and eggs per gram of root at 5 and 1% levels of probability, respectively, at Planting 1 (Table 22). Since coefficients of variation were very high and parents did not differ significantly at Planting 1, the level of confidence of these results is low. Significant additive effects were not detected in Planting 2 nor in the combined analysis. The low

R 2 values reflect high environmental effects and/or poor fit of the model. Dominant effects were significant for both eggs per root system and eggs per gram of root at Planting 2 and for eggs per gram of root in combined analysis, but, their direction was consistently negative for both traits. The negative sign associated with dominance component indicated that in these hybrid combinations, number of eggs were decreased relative to the mid-parent (Ginkel and Scharen, 1987).

Comparisons of Fj and backcross generation means with the mid-parent value show partial dominance for resistance, the desired direction for a

Fj hybrid cotton breeding program.

Among epistatic effects, estimates of pooled aa gene effects were 69

Table 22: Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 1 (La. RN 910 x DP 41).

Planting dates

Gene effects Combined Planting 1$ Planting 2

Number of eggs per root system (1 0 0 0 1s)

11.02 + 1 .00** 0.87 ± 0.08** 20.33 + 1.06** a -1.68 ± 1.70 -0.37 + 0.14* -0.85 ± 1.80 d -6.92 ± 5.26 -0.55 ± 0.44 -12.29 ± 5.58* aa -5.67 ± 4.82 -0.96 + 0.40* -11.61 + 5.12* ad 2.66 ± 2.12 -0.35 ± 0. 18+ 5.91 ± 2.20** dd 9.77 ± 8.71 0.59 ± 0.73 16.29 ± 9.19+

C.V. 108% 105% 46%

R 2 4% 15% 16%

Number of eggs per gram of root (1 0 0 0 's) m 1 .68 ± 0.14** 0.36 ± 0.03* 2.89 ± 0.16** a -0.16 ± 0.23 -0.16 ± 0.05** 0.10 + 0.27 d -1. 34 ± 0.72+ -0.05 + 0.16 -2.44 ± 0.84** aa -1 . 10 + 0 .66+ -0.18 ± 0.15 -2.14 ± 0.77** ad 0.53 + 0.29+ -0.12 ± 0.07+ 1.17 ± 0.33** dd 1.05 i 1 .20 -0.22 + 0,27 1.91 ± 1.38

C.V. 101% 89% 50%

R 2 5% 19% 17%

§ Inoculated with eggs.

11 m»mean of the F2 generation; a*pooled additive effects; d“pooled dominance effects; aa, ad, and dd“pooled additive X additive, additive X dominance, and dominance X dominance effects, respectively.

+, *, ** Significantly different from zero at the 0.10, 0.05, and 0.01 probability levels, respectively, according to "t" test of significance. 70

negative and generally significant for both numbers of eggs per root

system and eggs per gram of root. These estimates are in the desirable

direction, and aa gene effects contribute to the heritability of

resistance to reniform nematode. Estimates of pooled ad gene effects

for both traits were negative and significant (10% level) at Planting 1

but were positive and highly significant at Planting 2. Estimates of

pooled dd gene effects for both traits were generally positive and

nonsignificant. The ad and dd gene effects are in the undesirable

direction and difficult to manipulate in a cotton breeding program.

Experiment 2:

In Experiment 2, involving Auburn 612 RNR and Dp 41 parents,

estimates of genetic components of genotypic values indicated that

pooled additive gene effects generally were in the resistant (negative)

direction and pooled dominant effects in the susceptible (positive)

direction, but both were either nonsignificant or significant at a low

level (10%) of probability (Table 23). The dominant gene effects are in

the undesirable direction to be useful in an Fj hybrid cotton breeding

program. Among estimates of epistatic effects, pooled aa effects for

both traits were in the positive direction and, in the case of eggs per

gram of root, significant at Planting 1 and combined analysis at the 5

and 10% levels of probability, respectively. Estimates of pooled ad

gene effects were mostly positive and consistently nonsignificant.

Estimates of pooled dd gene effects were generally negative and

significant for eggs per gram of root. Directions of the d, aa, and dd 71

Table 23: Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 2 (Auburn 612 RNR x DP 41).

Planting dates

Gene effects Combined Planting 1 Planting 2

Number of eggs per root system (1 0 0 0 1s) m§ 4.87 ± 0.53** 2.11 + 0.24** 7.68 ± 0.71** a -1.64 + 0.91 + -0.04 ± 0.41 -2.40 ± 1.23+ d 3.37 ± 2.79 0.06 ± 1.26 5.32 ± 3.78 aa 1.51 + 2.56 0.90 + 1.15 1.35 + 3.47 ad -0.16 ± 1.12 0.74 + 0.50 -0.24 + 1 .50 dd -2.13 ± 4.63 0.18 ± 2.10 -4.08 ± 6.26

C.V. 847. 59% 53%

R 3 11% 16% 27%

Number of eggs per gram of root (1 0 0 0 's) m 0.69 + 0.07** 0.29 ± 0.04** 1,10 ± 0 .11** a -0.18 + 0.13 -0.04 ± 0.08 -0.20 ± 0.19 d 0.76 ± 0.40+ 0.32 ± 0.23 1.09 ± 0.57+ aa 0.78 t 0.36+ 0.54 ± 0.21* 0.75 + 0.53 ad 0.08 + 0,16 0.09 ± 0.09 0.17 ± 0.23 dd -1 .29 ± 0.66* -0.58 ± 0.39 -1.99 + 0.95*

C.V. 82% 67% 58%

R 3 10% 18% 19%

§ m*mean of the ?2 generation; a*pooled additive effects; d“pooled dominance effects; aa, ad, and dd«pooled additive X additive, additive X dominance, and dominance X dominance effects, respectively.

+, *, ** Significantly different from zero at the 0.10, 0.05, and 0.01 probability levels, respectively, according to "t" test of significance. 72

gene effects are generally opposite to those observed for these effects

in Experiments 1 and 3.

Experiment 3:

In Experiment 3, involving M 019-RNR and Dp 41 parents, although PD x G interaction was not significant, parents differed significantly at the 5% level of probability for mean numbers of eggs per gram of root at

Plant ing 1 only (Table 4). Estimates of genetic components of genotypic values are, therefore, relevant for only eggs per gram of root at

Planting 1 (Table 24). Additive and dominant gene effects significantly reduced eggs per gram of root over mid-parent value at the 10% level of probability. Among epistatic effects, the estimate of pooled aa gene effects was negative and significant at the 10% level of probability, other estimates of epistatic effects were nonsignificant.

Discussion of Generation Mean Analysis:

Instead of estimating genetic variation within generations, generation mean analysis, a type of genetic analysis of gene action, determines the relative importance of genetic effects estimated from the means of different generations. Genetic variances, on the other hand, are determined from the summation of squared effects for each locus.

The intralocus gene action at a given locus may involve dominance, additiveness, partial dominance or overdominance type of gene effects.

Dominance refers to the modification of the expression of one member of a pair of alleles by the other that results in a deviation from the 73

Table 24: Estimates of genetic components of generation means and their standard errors for number of reniform nematode eggs per root system and eggs per gram of root in Experiment 3 (M 019-RNR x DP 41).$

Planting dates

Gene effects Combined Planting 1 Planting 2

Number of eggs per root system (1000's) ml' 8.74 ± 0.52** 11.31 + 0.60** 6.34 ± 0.57** a -2.47 + 0.88** -3.61 ± 1.00** -1.21 ± 0.98 d -3.07 ± 2.73 -2.25 ± 3.12 -4.91 + 3.02 aa -4.45 ± 2.50+ -2.59 ± 2.86 -7.11 ± 2.77* ad -2.36 + 1.09* -2.86 ± 1.25* -1.16 ± 1.21 dd 0.67 ± 4.52 0.77 ± 5.15 2.75 ± 5.00

C.V. 617. 36% 75%

R 2 10% 19% 14%

Number of eggs per gram of root (1000's)

1.99 ± 0.13** 3.04 ± 0.19** 0.99 ± 0.09** a -0.43 ± 0.23+ -0.62 ± 0.32+ -0.22 ± 0.16 d -1.07 ± 0.70 -1.91 ± 1.01+ -0.54 ± 0.50 aa -1.02 ± 0.65 -1.54 ± 0.93+ -0.73 + 0.46 ad -0.25 + 0.28 -0.13 ± 0.40 -1.18 ± 0.20 dd 0.43 ±1.16 1.34 ± 1.67 0.08 ± 0.83

C.V. 77% 53% 71%

R 2 3% 6% 11%

§ No significant interaction between generations and planting dates was detected for number of eggs per root system and eggs per gram of root.

II nmnean of the F2 generation; a~pooled additive effects; d»pooled dominance effects; aa, ad, and dd~pooled additive X additive, additive X dominance, and dominance X dominance effects, respectively.

+, *, ** Significantly different from zero at the 0.10, 0.05, and 0.01 probability levels, respectively, according to F test of significance. 74

simple additive scheme. In case of no dominance or additive type of

intalocus interaction , the phenotype of the heterozygote falls midway

between that of the two homozygous parents. The heterozygote may

achieve a value closer to one parent than the other. This is termed partial dominance. If the heterozygote has a value outside the range

between the two parents, it is called overdominance. Just as dominance

at any particular locus results in a deviation from the simple additive

scheme, so also interlocus interactions (among genes at different loci) result in epistatic deviations.

Generation mean analysis, as used by Gamble (1962), involves the following assumptions to simplify statistical procedures: 1) multiple alleles absent; 2) linkage absent; 3) lethal genes absent; 4) constant viability for all genotypes; 5) environmental effects additive with the genotypic value. Failure to meet these assumptions may result in serious bias to the estimates.

There would be no serious bias expected in the estimates of the parameters from assumptions 1, 3, and 5. Since the only segregating populations used in this study are the F2 and first backcross generations of a cross between two presumably homozygous lines, multiple alleles would be present only if the parental lines were not homozygous or if mutations occurred. Lethal genes are not likely to be present in the crosses, since the parental lines used in this study have been maintained by selfing and/or by growing in an area with minimum or no outcrossing. Viability perhaps was not constant for all genotypes but was satisfactory in the greenhouse tests and negligible bias would be expected. 75

According to Hayman (1960), estimates of a and d effects are biased

by epistatic effects and linkage disequilibrium (if present).

Estimation of digenic epistatic effects is unbiased if linkage of

interacting loci and higher order epistatic effects are absent. Only

early generations of a cross are considered in this study and an

equilibrium of linkage relations is improbable in a quantitative trait

like yield (Comstock and Robinson 1952; Mather 1949), therefore, if

there is epistasis, bias due to linkage relations may be present in the

estimates of gene effects (Kempthorne 1957). The most serious bias

would be expected to occur in the estimates of additive x additive and

the dominance x dominance effects. However, apparent linkage bias might

be due to trigenic or higher epistasis. Because of the bias in the

estimates of a and d effects, the relative importance of a and d effects

vs. epistatic effects cannot be directly assessed. Some indication of

these relative effects may be gained by comparing residual sums of

squares after fitting the three-parameter (m, a, and d) and the six-

parameter (m, a, d, aa, ad, and dd) models. In this study, low R 2

values for estimates of genetic components (3 to 27Jt), using six-

parameter model, indicate that this genetic model did not account for much of the total variation. This suggests presence of trigenic or

higher epistatic effects and/or tremendous environmental influence.

Therefore, three-parameter model was not fitted.

The assumption that environmental effects are additive with the genotypic value is not expected to be realized in the material under

study. The bias caused by genotype-environment interactions in the estimates of various parameters is of unknown magnitude and direction; 76 furthermore, the bias may not be the same for each parameter. In this study, the bias due to genotype-environment interact ions may have been reduced somewhat in the estimates of gene effects since each experiment was divided into two replications and grown at two planting dates.

However, it is realized that number of environments was not sufficient.

Generation mean analysis has some advantages in comparison with mating designs used for estimation of genetic components of variance.

The errors are inherently smaller because means (first order statistics) rather than variances (second order statistics) are used in generation mean analysis. Smaller experiments are required for generation mean analysis to obtain the same degree of precision.

Estimation of heritability and prediction of genetic advance is not possible with generation mean analysis because estimates of genetic variances are not available. In this study, heritability was estimated by using phenotypic variances of different generations that were estimated from individual plant data. Cancellation of effects may be a significant disadvantage because, for example, dominance effects may be present but opposing at various loci in the two parents and cancel each other. Generation mean analysis does not reveal opposing effects.

Primary function of generation mean analyses is to obtain specific genetic information about a specific pair of lines. For quantitative traits, the estimates of genetic effects could be quite different for different pairs of lines, depending on the relative frequency of opposing and reinforcing effects for the specific pair of lines studied.

Generation mean analysis is useful in determining the occurance of epistatic effects and if not present in estimating the relative 77 importance of nonadditive genetic effects for the justification of a hybrid breeding program in self-pollinated species, because limited hand pollinations are required to produce the different generations.

Significant epistatic effects for both traits in the three experiments of this study suggest that both traits are controlled by two or more pairs of genes. Since the susceptible parent , Dp 41, is common in all three experiments, differences in the direction of gene effects would be due to the genetics of resistant parents. In general the direction of d and aa effects in La. RN 910 and M 019-RNR was negative and that of dd effects was positive, whereas, direction of these effects was opposite in Auburn 612 RNR. Differences in direction of these gene effects suggest that La. RN 910 and M 019-RNR lines have different genetic mechanism for resistance to reniform nematode than Auburn 612,

RNR and their combination through breeding may lead to increased resistance to reniform.

Since these results are unique, more experiments with greater precision and higher inoculation levels are required to draw general conclusions. It is suggested that advancing generations to F/, or F5, while maintaining genetic variability prior to selection, may improve selection efficiency by increasing homozygosity and thereby reducing nonadditive gene effects for resistance to reniform nematode.

Significant epistatic gene effects in most cases and transgressive segregation for susceptibility suggest that resistance to reniform nematode in these cotton lines is controlled by at least two pairs of genes. Wide frequency distributions and relatively high coefficients of variation indicate that environmental variation greatly influenced 78 reniform nematode egg production and phenotypic differences among cotton genotypes in relative resistance. Further studies on the genetics of resistance to this nematode may be enhanced by the use of higher and more consistent inoculation levels and by the use of effective growth chambers or, if unavailable, by limiting studies in the greenhouse to the spring and fall times of the year. LITERATURE OXTED

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138 Yik, C. P. and W, Birchfield. 1984. Resistant germplasm in Gossypium species and related plants to Rotylenchulus reniformis. J. Nematology 16(2):146-153. APPENDIX

90 Table 8a. Frequency distribution of number of eggs per root system for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41).

Frequency

Pi 11 3 2 I5

P2 5 5 3

Pi 1 7 4 2 1 3 I

P2 12 18 16 6 3 3 5 2 1 1 2 1 1 1 1

PfPl 25 6 2 3 2 1 1

P2Fi 3 7 3 6 4 1 3 2 1

0 .4 .8 1.2 1.6 2 2.4 2.8 3.2 3.6 4

Number of eggs per root system (1 0 0 0 's)

S ■ number underlined was omitted for analysis purposes. Table 10a. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 1 (La. RN 910 x DP 41).

Frequency

Pi 8 6 2 I5

?2 3 2 2 2 2 1 1 I

Pi 4 3 3 1 2 1 1 2 1 _1

*2 6 9 10 9 9 4 4 1 3 2 2 1 4 2 2 2 1 1

PlFi 14 9 2 5 2 1 1 2 I 1 2

P2 Fi 2 1 3 3 2 6 I 1 2 1 2 2 2 2 I 1 // // // 0 .1 .2 .3 .4 .5 .6 .7 .8 1.0 1.2 1.5 1.6

Number of eggs per gram of root (1000's)

§ ■ number underlined was omitted for analysis purposes.

* NJ Table 11a. Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment I (La. RN 910 x DP 41).

Frequency

Pi 6 4 4 2 4

P2 1 1 1 1 2 5 1 1 1 2 2 1 I

Pi 1 2 2 2 3 2 1 1 2 1 1 2

F2 1 1 2 4 4 11 7 8 6 7 3 3 8 2 5 1 1 2 1 1 J.

PlFi 2 2 2 5 3 4 5 2 3 4 2 4 1 1

P2Fi 4 6 7 4 1 2 2 3 2 3 1 1 1 1 Si // // // 0 .6 1.2 1.8 2.4 3 3.6 4.2 4.8 6 6.3 7,2 9.0 14.0

Number of eggs per gram of root (1000’s)

S » number underlined was omitted for analysis purposes. Table 14a. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 2 (Auburn 612 RNR x DP 41).

Frequency

Pi 2 2 3 4 2 3 2 1 1

*2 1 1 1 2 4 3 4 2 1 I§

Pi 1 4 4 1 2 1 2 1 I _1

P2 11 17 12 8 5 4 4 2 3 4 5 1 1 3

PlFi 6 3 7 3 5 1 3 2 2 2 2 1 1 1 1

P2Fi 4 5 5 2 4 4 5 1 4 1 I I 1 1 1

0 .2 .4 .6 .8 1.0 1.2 1.4 1.6 1.8 2.0 2.2 2.4 2.6 2.8 3.0

Number of eggs per gram of root (1000's)

S * number underlined was omitted for analysis purposes. Table 15a. Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 2 (Auburn 612 RNR x DP Al).

Frequency

Pl 2 3 2 5 A 2 1 1

P2 5 2 3 3 1 1 A 1 Fl 2 2 1 1 2 3 1 2 2 L 1 1 1

F2 1 5 9 11 6 7 9 A 5 3 5 1 2 5 1 1 2 1 1 _1*

PlFi 3 3 AA 6 3 3 3 2 1 2 3 1 1

P2fl 5 5 A A 5 2 1 3 3 1 2 1 2 1 1 // // // // 0 .4 .8 1.2 1.6 2 2.A 2.8 3.2 3.8 4.A 5 6.A

Number of eggs per gram of root (1000's)

$ * number underlined was omitted for analysis purposes.

<£> Ln Table 18a. Frequency distribution of number of eggs per gram of root for six populations at Planting 1, Experiment 3 (M 019-RNR x DP 41).

Frequency

Pi 2 6 2 1 4 2 2 1

P 2 1 1 3 6 1 2 1 1 I5

Pi 1 3 5 4 1 1 2 1 1 f 2 6 9 13 8 12 5 5 3 6 2 2 1 1 1 1 1 I

PlFi 4 6 9 6 6 3 2 3 i

P2Fi 1 2 8 4 5 9 6 1 2 2

0 1 2 3 4 5 6 7 8 9 10 11 12

Number of eggs per gram of root (1000’s)

§ * number underlined was omitted for analysis purposes.

o O' Table 19a. Frequency distribution of number of eggs per gram of root for six populations at Planting 2, Experiment 3 (M 019-RNR x DP 41).

Frequency

Pi 3 3 2 5 2 2 1 1 I 5

P2 2 3 2 7 3 1 1 1

F1 1 1 2 5 4 1 2 2 i 1

f2 10 7 11 6 3 3 6 9 6 7 2 4 2 1 3 1 1

PlFi 4 8 2 7 5 3 1 4 1 3 1 1

P2Fi 4 10 1 4 1 4 4 2 1 2 1 2 2 IJ 0 .4 .8 1.2 1.6 2 2.4 2.8 3.2 3-4 3.6 5.0

Number of eggs per gram of root (1000's)

} * number underlined was omitted for analysis purposes. VTTA

Noor Muhammad was born in Multan (Pakistan) on November 1, 1949.

He received his elementary education from Mauza, Bootey Wala (Multan) and graduated from Islamia High School, Am Khas Bagh, Multan in 1967. A

Bachelor of Science (Hons.) Agriculture and Masters of Science (Hons.)

Agriculture with specialization in Plant Breeding and Genetics was received in 1972 and 1976, respectively, from the University of

Agr iculture Faisalabad (Pakistan). He joined the Cotton Research

Station, Multan, Punjab (Pakistan) in May, 1975.

On the recommendations of the selection committee, Governor of the

Punjab nominated him in 1984 for a three year U . S . training program leading to a Ph. D. in plant breeding, under financial aid of the U. S.

Agency for International Development. He enrolled in the Graduate

School of Louisiana State University during the Spring of 1985. He is now a candidate for a Ph. D. degree in Agronomy with a minor in

Entomology.

He is an associate member of the scientific research society Sigma

Xi and a member of the Gamma Sigma Delta Honor Society, American Society of Agronomy, and Crop Science Society of America.

98 DOCTORAL EXAMINATION AND DISSERTATION REPORT

Candidate: Noor Muhammad

Major Field: Agronomy

Title ol Dissertation: Genetics of Resistance to Reniform Nematode, Rotylenchulus Reniformis Linford and Oliveira, in Cotton, Cossypium Hirsutum L.

Approved:

Major Professoi Cha irm an

Dean of the Graduate Scl

EXAMINING COMMITTEE:

ci

Date of Examination:

_Ju_Ty 15a 1988