• TABLE OF CONTENTS
HIDE
 Title Page
 Acknowledgement
 Table of Contents
 List of Tables
 List of Figures
 Abstract
 Introduction
 Secondary-fruit ontogeny
 Physiological studies on secondary-fruit...
 Growth regulator effects on fruit...
 Appendix
 Bibliography
 Biographical sketch






Title: Navel orange fruit drop
CITATION PDF VIEWER THUMBNAILS PAGE IMAGE ZOOMABLE
Full Citation
STANDARD VIEW MARC VIEW
Permanent Link: http://ufdc.ufl.edu/UF00097420/00001
 Material Information
Title: Navel orange fruit drop secondary-fruit ontogeny, physiological studies, and growth regulator effects
Physical Description: xiv, 141 leaves : ill. ; 28 cm.
Language: English
Creator: Lima, José Eduardo Oliveira de, 1953-
Publication Date: 1983
Copyright Date: 1983
 Subjects
Subject: Oranges   ( lcsh )
Abscission (Botany)   ( lcsh )
Horticultural Science thesis Ph. D
Dissertations, Academic -- Horticultural Science -- UF
Genre: bibliography   ( marcgt )
non-fiction   ( marcgt )
 Notes
Thesis: Thesis (Ph. D.)--University of Florida, 1983.
Bibliography: Bibliography: leaves 130-140.
Additional Physical Form: Also available on World Wide Web
General Note: Typescript.
General Note: Vita.
Statement of Responsibility: by José Eduardo Oliveira de Lima.
 Record Information
Bibliographic ID: UF00097420
Volume ID: VID00001
Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: alephbibnum - 000352639
oclc - 09799625
notis - ABZ0615

Downloads

This item has the following downloads:

PDF ( 5 MBs ) ( PDF )


Table of Contents
    Title Page
        Page i
        Page ii
    Acknowledgement
        Page iii
        Page iv
    Table of Contents
        Page v
        Page vi
    List of Tables
        Page vii
        Page viii
        Page ix
        Page x
    List of Figures
        Page xi
        Page xii
    Abstract
        Page xiii
        Page xiv
    Introduction
        Page 1
        Page 2
    Secondary-fruit ontogeny
        Page 3
        Page 4
        Page 5
        Page 6
        Page 7
        Page 8
        Page 9
        Page 10
        Page 11
        Page 12
        Page 13
        Page 14
        Page 15
        Page 16
        Page 17
        Page 18
        Page 19
        Page 20
        Page 21
        Page 22
        Page 23
        Page 24
        Page 25
        Page 26
        Page 27
        Page 28
        Page 29
        Page 30
        Page 31
        Page 32
        Page 33
        Page 34
        Page 35
        Page 36
        Page 37
        Page 38
        Page 39
        Page 40
        Page 41
        Page 42
        Page 43
        Page 44
        Page 45
        Page 46
    Physiological studies on secondary-fruit yellowing of navel orange
        Page 47
        Page 48
        Page 49
        Page 50
        Page 51
        Page 52
        Page 53
        Page 54
        Page 55
        Page 56
        Page 57
        Page 58
        Page 59
        Page 60
        Page 61
        Page 62
        Page 63
        Page 64
        Page 65
        Page 66
        Page 67
        Page 68
        Page 69
        Page 70
        Page 71
        Page 72
        Page 73
        Page 74
        Page 75
    Growth regulator effects on fruit drop, yield and quality of navel orange
        Page 76
        Page 77
        Page 78
        Page 79
        Page 80
        Page 81
        Page 82
        Page 83
        Page 84
        Page 85
        Page 86
        Page 87
        Page 88
        Page 89
        Page 90
        Page 91
        Page 92
        Page 93
        Page 94
        Page 95
        Page 96
        Page 97
        Page 98
        Page 99
        Page 100
        Page 101
        Page 102
        Page 103
        Page 104
        Page 105
        Page 106
        Page 107
        Page 108
        Page 109
        Page 110
        Page 111
        Page 112
        Page 113
        Page 114
        Page 115
        Page 116
        Page 117
        Page 118
        Page 119
    Appendix
        Page 120
        Page 121
        Page 122
        Page 123
        Page 124
        Page 125
        Page 126
        Page 127
        Page 128
        Page 129
    Bibliography
        Page 130
        Page 131
        Page 132
        Page 133
        Page 134
        Page 135
        Page 136
        Page 137
        Page 138
        Page 139
        Page 140
    Biographical sketch
        Page 141
        Page 142
        Page 143
        Page 144
        Page 145
Full Text














NAVEL ORANGE FRUIT DROP:
SECONDARY-FRUIT ONTOGENY, PHYSIOLOGICAL STUDIES,
AND GROWTH REGULATOR EFFECTS









By

JOSE EDUARDO OLIVEIRA DE LIMA


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



UNIVERSITY OF FLORIDA


1983































To my mother and father, whose encouragement and love

made it all possible.















ACKNOWLEDGEM4ENTS


The author expresses appreciation to his committee

chairman, Dr. Frederick S. Davies, Associate Professor,

Fruit Crops Department, for his most valuable encouragement

and assistance in conducting this research and preparing

this manuscript.

Appreciation is extended to Dr. James Soule, Dr. Terry

W. Lucansky and Dr. Howard Berg for allowing the use of

laboratory and microscope facilities, support and

suggestions during the course of morphological and

anatomical studies.

The assistance of Dr. R. Hilton Biggs, Dr. Dwain D.

Gull, and Dr. Jasper N. Joiner in the use of laboratory

facilities and suggestions during physiological studies is

greatly appreciated.

The help of Mr. Steven Hiss with photographic work is

appreciated.

The author is further indebted to Mr. Dixie Royal and

Mr. E. McCown for providing experimental trees and

assistance during field work.

This program was made possible through the sponsorship

by Empresa Brasileira de Pesquisa Agropecuaria, Embrapa,

whose support is greatly appreciated.


iii







Appreciation is also extended to the graduate students

and friends at the University of Florida for constant

support and encouragement during this program.

The author gladly acknowledges the support and love of

his parents, Sergio and Hilda de Lima, brothers, and sister.

Finally, the author wants to express eternal gratitude

for the constant support, understanding, company and love of

his wife Maria Ines, who patiently helped him through this

program and the homesickness for their dearly loved country,

and of their children,Jose Eduardo, Carolina, and others to

come, who unknowingly provided the foremost motivation for

this work.

















TABLE OF CONTENTS


ACKNOWLEDGEMENTS . . . .

LIST OF TABLES . . . . .

LIST OF FIGURES . . . .

ABSTRACT . . . . . .

INTRODUCTION . . . . .

CHAPTER

I SECONDARY-FRUIT ONTOGENY


Introduction . . . . . . .

Literature Review . . . . .

Materials and Methods . . . .

Results and Discussion . . . .

Summary . . . . . . .

II PHYSIOLOGICAL STUDIES ON SECONDARY-FRUIT
YELLOWING OF NAVEL ORANGE . . . .


Introduction . . . . .

Literature Review . . .

Materials and Methods . .

Results and Discussion . ..

Summary . . . . . .

III GROWTH REGULATOR EFFECTS ON FRUIT
YIELD AND QUALITY OF NAVEL ORANGE

Introduction . . . . .

Literature Review . . .

Materials and Methods . .


DROP,
* .


Page


. . . . . iii

. . . . . vii

. . . . . xi

. . . . . xiii

. . . . . 1




. . . . . 3
311


. . 47

. . 47

. . 49

. . 54

. . 60

. . 73



. . 76

S . 76

S . 77

. . 82








Page

Results and Discussion . . . . .. 88
Summary . . . . . . . . 115

APPENDIX . . . . . . . . ... . . 120

LITERATURE CITED . . . . . . . ... .130

BIOGRAPHICAL SKETCH . . . . . . . .. .141














LIST OF TABLES


Page

2.1. Secondary-fruit yellowing (SFY) of navel
orange as affected by date of induction by
fruit-stem ringing, 1982. . . . . .. 61

2.2. Eruit ethylene, secondary-fruit yellowing
(SFY), and fruit drop of navel and 'Hamlin'
orange following fruit-stem ringing and
ethephon (ETH) or 2,4-dichlorophenoxyacetic
acid (2,4-D) stylar-end treatments, 1982. . 62

2.3. Leaf abaxial diffusive resistance (ADR), xylem
water potential ('x), total nonstructural
carbohydrates (CHO), and secondary-fruit
yellowing (SFY) of navel orange as affected by
fruit-stem ringing, 1982. . . . . ... 67

2.4. Leaf abaxial diffusive resistance (ADR), xylem
water potential (Yx), total nonstructural
carbohydrates (CHO), and secondary-fruit
yellowing (SFY) of navel orange as affected by
branch sawing or scoring, 1982. . . . ... 68

2.5. Leaf totalnonstructural carbohydrates (CHO) of
navel orange as affected by trunk girdling,
1982. . . . . . . . . ... . 69

2.6. Secondary-fruit yellowing (SFY) of navel orange
following fruit-stem or branch ringing and leaf
removal treatments, 1980 to 1982. . . ... 70

2.7. Secondary-fruit yellowing (SFY) of navel orange
following fruit-stem ringing as affected by
2,4-dichlorophenoxyacetic acid (2,4-D)
applications, 1982. .. . . . . . .. 72

2.8. Secondary-fruit yellowing (SFY) of navel orange
following fruit-stem ringing as affected by
2,4-dichlorophenoxyacetic acid (2,4-D) whole-
tree application or trunk girdling, 1982. . 74


vii








Page


3.1. Fruit weight, diameter, secondary-fruit
diameter (SF), stylar-end aperture diameter
(SEA), equatorial peel thickness and presence
of secondary-fruit protrusions of healthy
fruit and fruit affected by secondary-fruit
yellowing (SFY) in navel orange, 1981. ... .92

3.2. Fruit weight, diameter, secondary-fruit
diameter (SF), stylar-end aperture diameter
(SEA), and equatorial peel thickness of
healthy fruit and fruit containing rind
protrusions (PRO) in navel orange, 1981. ... .93

3.3. Fruit weight, diameter, secondary-fruit
diameter (SF), stylar-end aperture diameter
(SEA), and equatorial peel thickness of
fruit sampled at the north bottom and south
top canopy positions in navel orange trees,
1981. . . . . . . . ... 94

3.4. Fruit weight, diameter, secondary-fruit
diameter (SF), stylar-end aperture diameter
(SEA), equatorial peel thickness, and presence
of rind protrusions of healthy fruit and fruit
affected by stylar-end decay (SED) in navel
orange, 1980 and 1981. ..... . . . 96

3.5. Fruit weight, diameter, secondary-fruit
diameter (SF), stylar-end aperture diameter
(SEA), and equatorial peel thickness of
healthy and split fruit in navel orange, 1980
and 1981. . . . . . . . . . 97

3.6. Bloom and June drop of reproductive structures
of navel orange as affected by gibberellic
acid (GA) and 2,4-dichlorophenoxyacetic acid
(2,4-D) applications, 1981. ... .. . ... 98

3.7. Summer fruit drop of navel orange as affected
by gibberellic acid (GA), 2,4-dichlorophenoxy-
acetic acid (2,4-D) and Promalin applications,
1980. . . . . ... . . . . .. 100

3.8. Summer fruit drop of navel orange as affected
by gibberellic acid (GA) and 2,4-dichloro-
phenoxyacetic acid (2,4-D) applications, 1981. 101

3.9. Summer fruit drop of navel orange as affected
by trunk girdling or 2,4-dichlorophenoxyacetic
acid (2,4-D) applications, 1982. . . . ... 102


viii








Page


3.10. Summer-fall fruit drop of navel orange as
affected by gibberellic acid (GA), 2,4-di-
chlorophenoxyacetic acid (2,4-D) and Promalin
applications, 1980. . . . . . . ... 104

3.11. Summer-fall fruit drop of navel orange as
affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . . ... 105

3.12. Summer-fall fruit drop of navel orange due to
stylar-end decay (SED), fruit splitting,
branch collapse, and other causes, as
affected by gibberellic acid (GA) and 2,4-di-
chlorophenoxyacetic acid (2,4-D) applications,
1901. . . . . . . . . .. ... .106

3.13. Navel orange yield as affected by
gibberellic acid (GA) and 2,4-dichloro-
phenoxyacetic acid (2,4-D) applications,
1981. . . . . . . . . ... ... .107

3.14. Fruit characteristics of navel orange as
affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . . ... 110

3.15. Fruit external color and rind puncture force
of navel orange as affected by gibberellic
acid (GA) and 2,4-dichlorophenoxyacetic acid
(2,4-D) applications, 1981. . . . . ... 114

3.16. Navel orange juice content, total soluble
solids (TSS), total titratablee) acid (TA),
and TSS:TA ratio as affected by gibberellic
acid (GA) and 2,4-dichlorophenoxyacetic acid
(2,4-D) applications, 1981. . . . . ... 116

3.17. Navel orange juice content, total soluble
solids (TSS), total titratablee) acid (TA),
and TSS:TA ratio as affected by 2,4-dichloro-
phenoxyacetic acid (2,4-D) applications,
1982. . . . . . . . . ... .117

A.I. Preharvest fruit drop of navel orange as
affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . . . 120








Page


A.2. Navel orange yield as affected by trunk
girdling or 2,4-dichlorophenoxyacetic acid
(2,4-D) applications, 1982. . . . .

A.3. Fruit weight, diameter, and stylar-end
aperture diameter (SEA) of navel orange as
affected by trunk girdling or 2,4-dichloro-
phenoxyacetic acid (2,4-D) applications,
1982 . . . . . . . . .

A.4. Fruit external color of navel orange as
affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . .

A.5. Fruit rind puncture force of navel orange
as affected by gibberellic acid (GA) and
2,4-dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . .

A.6. Fruit external color and rind puncture
force of navel orange as affected by 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1982. . . . . . .

A.7. Fruit juice content of navel orange as
affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . .


. 121





. 122




. 123




. 124




. 125




. 126


A.8. Total soluble solids in navel orange juice
as affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . . ... 127

A.9. Total titratablee) acid in navel orange juice
as affected by gibberellic acid (GA) and 2,4-
dichlorophenoxyacetic acid (2,4-D)
applications, 1981. . . . . . . ... 128

A.10. Ratio of total soluble solids (TSS) and
total titratablee) acid (TA) in navel
orange juice as affected by gibberellic acid
(GA) and 2,4-dichlorophenoxyacetic acid
(2,4-D) applications, 1981. . . . . ... 129














LIST OF FIGURES


Page

1.1. Primary-carpel primordia in flower buds
24 to 26 days before anthesis in navel
orange. . . . . . . . .... . 15

1.2. Secondary-carpel ontogeny in flower buds
21 to 23 days before anthesis in navel
orange. . . . . . . . .... . 18

1.3. Secondary-carpel ontogeny in flower buds
18 to 19 days before anthesis in navel
orange. . . . . . . . .... . 20

1.4. Secondary-carpel ontogeny in flower buds
12 to 15 days before anthesis in navel
orange. . . . . . . . .... . 22

1.5. Secondary-pistil ontogeny in flower buds
5 to 6 days before anthesis in navel
orange. . . . . . . . .... . 25

1.6. Secondary-pistil ontogeny in flower buds
4 days before anthesis in navel orange. ... . 27

1.7. Secondary-pistil development in flower buds
3 days before anthesis to 10 days after
petal fall in navel orange. . . . . .. 29

1.8. Primary- and secondary-ovary diameter after
1 mm flower bud stage in navel orange. . 32

1.9. Changes in volume of the primary and secondary
fruit, and in diameter of the stylar-end
aperture from anthesis until legal maturity
of the fruit in navel orange, 1980. . . ... 37

1.10. Tissue protrusions into primary-fruit
locules of navel orange. . . . . ... 40

1.11. Secondary-fruit abscission in navel orange . 43










2.1. Ethylene, cellulase activity, and fruit
removal force in the abscission zone at the
base of the primary and secondary fruit in
relation to stage of secondary-fruit
yellowing of navel orange, 1982. . . .


3.1. Distribution of causes of summer and summer-
fall fruit drop in 2 navel orange groves
near Eustis, Florida, 1979 to 1982 . . . .


xii


Page


S. 65







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



NAVEL ORANGE FRUIT DROP:
SECONDARY-FRUIT ONTOGENY, PHYSIOLOGICAL STUDIES,
AND GROWTH REGULATOR EFFECTS


By

JOSE EDUARDO OLIVEIRA DE LIMA

April 1983


Chairman: Dr. Frederick S. Davies
Major Department: Horticultural Science (Fruit Crops)


Navel orange (Citrus sinensis (L.) Osbeck) fruit drop,

which occurred after the fruit set period, was studied in 2

groves in the north central citrus region of Florida from

1979 to 1982. Fruit drop per year ranged from 63 to 200

fruit per tree. Major causes of drop were secondary-fruit

yellowing (SFY), stylar-end decay (SED), and fruit splitting.

These causes of fruit drop were related to the unusual

structure of the stylar-end of navel orange, which encloses

the secondary fruit.

The secondary fruit developed as an extra whorl of

carpels within the primary-carpel whorl. A complete

secondary pistil was formed entirely within the primary one.

Secondary ovary development followed a sigmoid growth curve

similar to that of the primary fruit.

An abscission zone was characterized at the base of the

secondary fruit, and abscission of the latter occurred prior


xiii








to SFY. Secondary-fruit abscission preceded and probably was

the cause of primary-fruit abscission in fruit affected by

SFY, as indicated by increases in cellulase activity and

ethylene levels in the abscission zone of the secondary

fruit prior to those in the abscission zone of the primary

fruit. Fruit-stem ringing and leaf removal experiments

suggested induction of SFY resulted from an interrupted

supply of bark-translocated leaf metabolites. Applications

of 2,4-dichlorophenoxyacetic acid (2,4-D) to the fruit or

fruit-stem reduced induction of SFY from fruit-stem ringing.

Fruit affected by SED and splitting had greater

diameter and thicker peel than healthy fruit, and their

aperture at the stylar-end had greater diameter. Rind

protrusions which usually occurred in segments of fruit

affected by SED consisted of secondary-carpel outgrowths

and were present early in fruit ontogeny.

A spray application of 20 ppm 2,4-D 5 to 9 weeks after

midbloom reduced SFY, but not other causes of fruit drop in

the 1980 to 1982 seasons, and increased yield in 1981 with

no adverse effect on fruit quality. Other 2,4-D, gibberellic

acid (GA), or 2,4-D + GA sprays at or within 19 weeks of

midbloom were less effective or ineffective.


xiv














I NTRODUCTION1


Navels constitute a group of early- to midseason-

maturing sweet orange cultivars (Citrus sinensis (L.)

Osbeck) characterized by the presence of a small, secondary

fruit, the navel, at the stylar end of the primary or main

fruit. Navel oranges are commercially grown throughout most

of the citrus areas of the world (51). Yields, however, are

usually erratic and lower than for other sweet oranges (23,

30,100,119). Navel yields in Florida are typically greater

than those in other citrus areas but frequently can be low

and fall below those of 'Hamlin' or 'Valencia' (67,68).

Although most authors attribute low navel yields to poor

fruit set (23,30,67,73,119), navel orange has been shown to

have significant fruit drop after the fruit-set period (79).

Causes of navel fruit drop after the fruit set period

in Florida include secondary-fruit yellowing (SFY), stylar-

end decay and fruit splitting (77,78,79). These fruit drop

problems are related to the unique structure of the stylar

end of the navel fruit, which encloses the secondary fruit

(77,79). Bouma (14) and Holtzhausen (52) studied fruit









development of navel orange, but little has been done

regarding secondary-fruit ontogeny (118).

Secondary-fruit yellowing accounted for more than half

of the fruit drop after fruit set in a navel grove in

Florida, in 1979 (77). The problem was related to physical

separation of the secondary from the primary fruit (77,79).

Pest or disease factors did not seem to be involved (77,

112). Fruit affected by SFY produced high amounts of

ethylene (112), but other physiological aspects of SFY and

related secondary-fruit separation have not been studied.

The objectives of this research were to study

secondary-fruit ontogeny, to investigate some physiological

aspects of SFY, and to control navel orange fruit drop in

Florida using growth regulator applications.















CHAPTER I

SECONDARY-FRUIT ONTOGENY






Introduction




Navel orange fruit drop after the fruit-set period has

been related to the unique structure of the fruit stylar-end

and of the secondary fruit (77,78,79). Secondary-fruit

yellowing, an important cause of fruit drop during late

spring and early summer, ma: result from the physiological

separation of the secondary fruit from the primary fruit.

This separation possibly involves an abscission zone at the

base of the secondary fruit (77,79). Fruit with larger

stylar-end aperture are more affected b:' stylar-end decay

and fruit splitting, which are important causes of navel

fruit drop during late summer and early fall (79). In

addition, rind-like tissue protrusions that originate from

the secondary fruit occur in the decayed primary-fruit

locules of most fruit affected by stylar-end decay (77,79).

Bouma (14) in Australia, and Holtzhausen (52) in South

Africa, studied fruit development of navel orange but made









no reference to the secondary fruit. Some aspects of

secondary-fruit structure and development have been briefly

discussed (12,105,118), but the available information on the

ontogeny of the secondary fruit is limited.

Developmental morphology of the secondary fruit in

navel orange was studied with emphasis on early ontogeny,

characterization of a secondary-fruit abscission zone and

the occurrence of secondary-fruit tissue protrusions within

the primary-fruit locules.





Literature Review




Fruit Ontogeny


Flower induction in sweet oranges typically occurs

during late fall and early winter (8). Differentiation of

the vegetative meristem into a reproductive meristem,

however, occurs only at the onset of the spring growth flush

(4). The floral apical meristem is characterized by a

flattening of the apical dome and the production of closely-

spaced successive whorls of flower organ primordia. Each

whorl of primordia is located slightly above and inside the

preceding whorl and gives rise to the various floral organs

(105).

The innermost whorl of primordia of floral organs

around the floral meristem consists of ca. 10 carpel









primordia (105). These primordia are first observed around

the floral meristem when the flower bud is 1-1.5 mm in

diameter. The carpel primordia are initially free, finger-

like and inwardly curved. Marginal meristems on the sides of

each primordium produce lamina-like wings which grow toward

the expanding apical meristem or axis (105). Developing

carpels appear horseshoe-shaped in transaction with the open

ends toward the center of the meristem. The carpel surface

at the periphery of the meristem is analogous to the abaxial

leaf surface (34). Carpels become fused at the abaxial

surface of their lamina-like wings, and are fused to the

expanding floral axis (i.e., the central axis) by their

margins (37). Carpels develop as chambers that are open at

the top, and are fused into one single pistil, which

composes the gynoecium of the flower.

A fully developed citrus pistil has a well-

differentiated ovary, style and stigma. Each carpellary

chamber is a locule in the ovary, and the fused wings of the

carpels become the septa (105). The carpellary chambers are

continuous throughout the style and stigma and form the

stylar canals. Stylar canals, one per carpel, open onto the

surface of the stigma and on the upper portion of the

locules between the two rows of ovules (37).

Ovules develop early in the ontogeny of the carpels

from tissue that is produced by the fusion of the 2

carpellary margins to the central axis in the ovary (105).








Two rows of ovules are present per locule. Placentation is

axile.

Juice sacs develop from the adaxial carpel surface as

club-shaped emergences prior to or at anthesis. They occupy

the entire locule when the ovaries are about 10 mm in

diameter (37).

A secondary set of carpels develops in some instances

at the apex of the floral axis within the primary pistil

(118). These secondary carpels become the characteristic

navel or secondary fruit of some citrus cultivars, such as

in navel orange (51).




Fruit Morphology and Anatomy



A citrus fruit is a hesperidium,which differs from other

berries by the presence of a hard or leathery rind (105).

The pericarp is composed of the exocarp, mesocarp and

endocarp. The exocarp, or flavedo, is the pigmented outer

portion of the fruit derived from the abaxial surface of the

carpels and is composed of an epidermis and several layers

of cells adjacent to it. The epidermis is covered by a thick

waxy cuticle and contains stomata interspersed among

epidermal cells (106). Some authors (12,106) recognize a

hypodermis composed of 1 to 3 rows of collenchyma cells.

Several layers of chlorenchyma cells, oil glands and the

endings of vascular bundles occur beneath the hypodermis.









Chlorophyll and carotenoid pigments in the chlorenchyma

cells give the fruit its characteristic green to orange

color (32).

The mesocarp, or albedo, is the usually white, spongy

tissue found internal to the exocarp. This tissue is

analogous to the spongy mesophyll of a leaf. The mesocarp is

composed of aerenchyma tissue which consists of lobed

parenchyma cells and schizogenous intercellular spaces (105).

An extensive network of vascular bundles permeatesthe albedo

(12). The exocarp plus the mesocarp are commonly referred to

as the peel or rind of a citrus fruit.

The endocarp originates from the adaxial surface of the

carpel primordia and surrounds the locular cavities. This

tissue is composed of an inner epidermis and several layers

of parenchyma cells (105). A portion of the endocarp

differentiates into the membrane that lines the locules.

Juice sacs develop in the endocarp (primarily on the dorsal

wall of the locules) from the epidermis and adjoining cell

layers. The locules of a fruit are separated by sept formed

by fusion of adjacent carpellary walls. A locule that

contains juice sacs, seeds, and the surrounding membrane is

called a segment (105). Segments are clustered around the

central axis of the fruit and form the edible pulp of the

latter. The central axis originates from expansion of the

floral meristem. Both the central axis and the portion of

the septa between locular membranes are composed of

mesocarp-like aerenchyma tissue (12).








Major vascular bundles in a citrus fruit are restricted

to the mesocarp and the central axis. Five bundles occur in

each carpel (37). Carpellary bundles in the mesocarp consist

of a prominent dorsal bundle opposite the locule and two

septal or lateral bundles opposite the septa. Two marginal

bundles opposite the septa or carpel margins occur in the

distal half of the central axis. Lateral bundles of 2

adjacent carpels, as well as two adjacent marginal bundles,

are fused. All major carpellary bundles are collateral

bundles (37). Dorsal and lateral bundles diverge from axial

bundles at the base of the ovary (37). Axial bundles,

however, continue into the proximal half of the central axis

of the fruit and then terminate after giving rise to ovular

traces and marginal bundles (105). The marginal bundles are

inverted collateral bundles which branch and fuse with

septal bundles and then give rise to concentric stylar

bundles (37).




Fruit Growth and Development



The growth of a citrus fruit, as determined by changes

in volume, equatorial diameter or dry or fresh weight,

typically follows a sigmoid pattern (10,14,52). Bouma (14)

and Holtzhausen (52) defined three stages of development for

navel orange. Stage I, the cell division stage, lasts until

a few weeks after anthesis and is characterized by small









growth rates. Fruit growth during stage I is due mainly to

growth of the peel that results from cell division and

enlargement in the mesocarp and exocarp tissues. The

majority of cells found in a fully developed fruit is

produced during stage I, although cell divisions continue in

the outer peel throughout fruit development. Stage II is

characterized primarily by rapid enlargement of cells in the

fruit segments and lasts until color break. At the end of

this stage of development the fruit has almost attained

final diameter and dry weight. The peel reaches maximum

thickness early in stage II and then decreases to almost

final thickness. Stage III is characterized by slow fruit

growth, and the latter is due primarily to development of

the fruit segments. The rind becomes orange during stage III,

accompanied by a decrease in titratable acidity of the juice

and other changes indicative of maturity. The fruit

typically reach maximum fresh weight during stage III, and

then fresh weight gradually decreases.




Fruit Abscission



Detachment of the young citrus fruit from the plant

usually is caused by the separation of cells in one of the 2

abscission zones which are located at the base of the

pedicel and at the base of the ovary (32). Sclerenchyma

tissue which forms in the abscission zone at the base of









the pedicel limits abscission to the abscission zone at the

base of the developing fruit (19).

The abscission or separation zone is the zone of

tissues proximal to the structure to be shed. This zone may

or may not be anatomically distinguishable from adjacent

tissues prior to abscission (11,123). The abscission zone is

composed typically of an epidermis, cortex, vascular bundles

and pith (11,34,72). The epidermis and pith are usually

similar to corresponding tissues on either side of the

abscission zone. Cortical parenchyma cells in the abscission

zone, however, usually are close-packed, thin-walled,

densely protoplasmic, and smaller than adjacent cells.

Xylary and phloic fibers of vascular bundles in the

abscission zone are unusually small or absent, and the

number of vessels reduced (17,80). The separation or

abscission layer typically refers to 1 or 2 layers of cells

in which separation actually occurs. The protective layer is

the layer of suberized cells which develop over exposed

surfaces of the plant following fruit abscission. The

protective layer typically consists of a periderm which is

usually continuous with the periderm of the stem (11,34).

The abscission zone of a citrus fruit is not easily

distinguished from adjacent tissues by anatomical

characteristics (124). Parenchyma cells in the separation

zone often are slightly smaller than in surrounding tissue,

and the tracheary elements in the abscission zone are

compressed. The occurrence of starch grains in parenchyma









cells of the separation layer is the first obvious

indication of an abscission zone in citrus (19,124). Starch

grains can be observed in cortical parenchyma cells on both

sides of the separation layer and in the intercellular

spaces between separated cells after abscission has occurred.

No cell division has been detected in association with

citrus fruit abscission (19,124).

No protective layer forms in the proximal (stem) side

after fruit abscission in citrus, and the pedicel usually

dessicates and dies (124). A protective layer does develop,

however, on the distal (fruit) side of the abscission zone

(19,124).






Materials and Methods



Navel orange trees (Citrus sinensis (L.) Osbeck) 12- to

25-years-old, on sour orange rootstock (Citrus aurantium L.)

were used in this study. The trees were growing in a grove

near Eustis (Lake County), in the north central citrus

region of Florida.

Samples for secondary-fruit ontogenetic studies were

collected from late February until fruit maturity in late

October, from 1980 to 1982. Two to 10 flower buds or fruit

per tree from 5 to 30 randomly selected trees were collected

daily until anthesis, weekly until stylar abscission and

then monthly.








Anatomical studies of secondary-fruit separation from

the primary-fruit were done on samples of tissues from the

central axis of the primary fruit, taken just below the

secondary ovary. Approximately 50 fruit which were sampled

at different stages of secondary-fruit yellowing during June

and July were examined in 1981 and 1982.

Primary- and secondary-fruit development were followed

by measuring their equatorial diameter on 30 randomly

sampled fruit weekly. Stylar-end aperture diameter was also

measured on these fruit.




Light Microscopy



Plant material for anatomical and ontogenetic studies

was fixed in formalin-acetic acid-50% ethanol (5:5:90 v:v:v),

for a few days to several months. The material was rinsed in

50% ethanol, dehydrated through an ethanol series,

infiltrated with paraffin, sectioned on a rotary microtome

at 8 to 25 pm, mounted on glass slides and stained with

either safranin-fast green (58,102), ruthenium red (57), or

toluidine blue (91). In addition, iodine-potassium iodide

(IKI) staining (57) was utilized in abscission zone studies.

Hesperidin crystals, which form during tissue dehydration

and interfere with the study of thin sections (37), were

removed by adding 0.25% NaOH to a 50% ethanol step in the

staining sequence (104).









Scanning Electron Microscopy



Flower buds and young pistils for studies at the

scanning electron microscope were killed and fixed in 2%

glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2) at 20 to

25 C for 24 hrs, rinsed with buffer for 30 min, post-fixed

in 1% osmium tetroxide for 2 to 3 days, dehydrated through

an ethanol series, critical-point dried, and gold coated.

Specimens were examined in a Hitachi S-450 scanning electron

microscope operated at 20 KV.






Results and Discussion




Secondary-Fruit Ontogeny



Primary-carpel primordia of navel orange were first

observed as the innermost whorl of primordia around the

floral apical meristem in flower buds ca. 0.8 mm in length

(Fig. 1.1 A). Carpel primordia in longisections of 1 mm-

length flower buds were finger-like protuberances around the

floral meristem and inwardly curved (Fig. 1.1 B). Marginal

meristems on the primordia produced lamina-like wings which

grew toward the center of the apical meristem (Fig. 1.1 C).

Carpels were initially free, but the wings of adjacent





























Fig. 1.1.


Primary-carpel primordia in flower buds 24 to 26
days before anthesis in navel orange. A, 0.8 mm-
length flower bud; B-D, 1 mm-length flower bud:
B, longisection, C, top view, and D, top view;
E-F, 1.2 mm-length flower bud: E, transaction,
and F, top view. FM, floral apical meristem; PE,
petal; PC, primary-carpel primordium; SE, sepal;
ST, stamen primordium.




15






,.






SEF



STT
C

AF


50pm 250Tm

















100pmn 25pm
PC PC















250pm 250pm








carpels became fused early in their ontogeny (Fig. 1.1 D).

Carpel primordia were horseshoe-shaped in transaction with

the open ends toward the center of the floral meristem (Fig.

1.1 E,F).

Secondary-carpel primordia were first observed within

the primary-carpel whorl in flower buds approximately 1.5 mm

in length (Fig. 1.2 A,B,C). No sepal, petal or stamen

primordia were associated with secondary-carpel development.

Secondary-carpel primordia in longisections of flower buds

1.8 to 2 mm in length (Fig. 1.2 D,E,F) were finger-like,

inwardly curved protuberances around the floral apical

meristem. Marginal meristems on the primordia produced

lamina-like wings which grew toward the center of the floral

meristem (Fig. 1.2 F) in a manner similar to primary-carpel

primordia (37,105). Secondary-carpel primordia in

transactions of developing pistils from 2.7 to 3 mm-length

flower buds (Fig. 1.3 A,B,C,D) appeared horseshoe-shaped

with the open ends toward the center of the meristem (Fig.

1.3 B,E). Secondary carpels were initially free (Fig. 1.3 B,

E,F), but their lamina-like wings fused with one another

early in their ontogeny.

Secondary carpels were entirely within primary pistils

in flower buds 6 to 8 mm in length (Fig. 1.4 A,B,C,D,E,F).

Contrary to the uniform and symmetrical development of

primary carpels (37,105), secondary carpels grew

asymmetrically and frequently overlapped each other (Fig.

1.4 D,E,F). The space for secondary-carpel development was

















0 C



--4-


1 4-4 E



f- --- I -,-
0 I II



4-Q n 0 "( a-
TC F- 04 Q
O--- -I ,- | J 0
-UI 0 E iE

0'- C E C -. O '

Cro = ( O ~ D



-1 0 .- 0 0
0, r-F- l u 4




: -i U












n D (V 0 0 4
O r-1 :)-
-O r -0 E I
-4 1 i 4-i -
-'U I ra >1i *U U) ro


0 r-1 O i 0 0
OC **J O *
-I '-l Ti ^-D0

UE 0 (D

a.E U) 0 U


c: .-1 .-) 0, U
a) U) D) Q r-i ra

4J '4 f i ra fa
C 0U 4-i J






U C'U 4-4 0








rI ,.
.-> 4-J0 *
r. -o IwD Lc



CU C MU-l 0

c --U -A 00
0t -C r 4J F= E


1l r5 -Q I a.


C.
C D ( I

O0 r- '




























o
I e


1 L
































Fig. 1.3. Secondary-carpel ontogeny in flower buds 18 to 19
days before anthesis in navel orange. A-B, 2.7 mm-
length flower buds: A, primary pistil, and B,
secondary-carpel primordia; C-F, 3mm-length
flower buds: C, primary pistil, D, longisection of
primary pistil, E, secondary-carpel primordia, and
F, secondary-carpel primordia. FMi, floral apical
meristem; PC, primary carpel; PP, primary pistil;
SC, secondary-carpel primordium.



















lot
c`


50jum


c:'^f-a


Kl
































Fig. 1.4.


Secondary-carpel ontogeny in flower buds 12 to 15
days before anthesis in navel orange. A-D, 6 mm-
length flower bud: A, longisection, B, transaction,
C, primary pistil, and D, secondary carpels; E-F,
8 mm-length flower buds: E, longisection of
primary pistil, and F, secondary carpels. ND,
nectar disc; PE, petal; PO, primary ovary; PP,
primary pistil; SC, secondary carpel; SE, sepal;
ST, stamen.













































































SI









limited and, in some instances, a carpel bent over and grew

toward the floral meristein (Fig. 1.5 A). Most secondary

carpels remained partially or totally free above the base

(Fig. 1.5 B,C,D), although fusion at their base (secondary-

ovary region) was generally complete (Fig. 1.5 B,D).

Secondary carpels generally developed into a complete

secondary pistil entirely within the primary pistil before

anthesis (Fig. 1.6 A). Incomplete fusion of secondary

carpels frequently resulted in multiple secondary styles and

stigmas (Fig. 1.6 A,B,C,D) and a single, well-fused

secondary ovary (Fig. 1.6 E,F). Conversely, primary carpels

are almost always fused and develop into pistils with a

single ovary, style, and stigma (37,105).

Stylar canals were present in the stigmas and styles of

secondary pistils (Fig. 1.6 B). As in primary carpels (37,

105), the stylar canals of secondary styles opened into the

locules, and formed continuous chambers within secondary

carpels. Stigmatic surfaces composed of papillose hairs were

present on the distal half of the secondary styles (Fig. 1.6

C,D). Secondary styles and stigmas were less distinct than

the primary ones (Fig. 1.7 A,B,C). The constriction at the

distal end of the ovary that marks the beginning of the

style was less noticeable (Fig. 1.7 A,C,D), and generally

there was no expansion of the distal extremity of the styles

to delineate the stigma (Fig. 1.7 B,C).

Expansion of the secondary style caused at times

splitting of the primary style. Abscission of the primary






























Fig. 1.5. Secondary-pistil ontogeny in flower buds 5 to 6
days before anthesis in navel orange. A, 10 mm-
length flower bud, longisection of primary pistil
and secondary carpels; B-D, 12 mm-length flower
bud: B, longisection of primary and secondary
pistils, C, transaction of primary and secondary
ovaries, and D, longisection of secondary pistil.
AX, axial vascular bundle; FM, floral apical
meristem; ND, nectar disc; PG, primary stigma; PO,
primary ovary; PS, primary style; SC, secondary
carpel; ST, stamen; TC, tertiary carpel primordium.





















t


Imm
..m.


i..
2501pr
oiwtWAla


1250jm
.A t t- c<


f! 1 j
r*l






























Fig. 1.6.


Secondary-pistil ontogeny in flower buds 4 days
before anthesis in navel orange. A-F, 15 mm-length
flower buds: A, longisection of primary pistil, B,
transaction of primary and secondary styles, C,
longisection of primary style and secondary
stigmas, D, transaction of primary style and
secondary stigmas, E, transaction of flower bud,
and F, longisection of secondary ovary. AX, axial
vascular bundle; CA, stylar canal; DB, dorsal
vascular bundle; PE, petal; PO, primary ovary; PP,
primary pistil; PS, primary style; SE, sepal; SG,
secondary stigma; SL, secondary-ovary locule; SO,
secondary ovary; SP, secondary pistil; SS,
secondary style; ST, stamen.




27






5CA.











AA
pp -







1mm A




















25 OAm 250Pm





SE








s ST





1mm E 259p





























Fig. 1.7.


Secondary-pistil development in flower buds 3 days
before anthesis to 10 days after petal fall in
navel orange. A-B, 16 mm-length flower buds, 3
days before anthesis: A, longisection of primary
and secondary pistil, and B, secondary style and
stigma; C-D, at anthesis: C, secondary pistil, and
D, longisection of primary and secondary ovaries;
E-F, at 10 days after petal fall: E, longisection
of primary and secondary ovaries, and F,
longisection at the stylar-end of primary and
secondary ovaries. FL, flavedo; PO, primary ovary;
PS, primary style; PV, primary ovule; SA, stylar
abscission zone; SG, secondary stigma; SO,
secondary ovary; SS, secondary style; SV,
secondary ovule; TC, tertiary carpel; TG, tertiary
stigma.













PS


500Jur
bl^ncs'kt


15 -T








style occurred about 8 days after petal fall and usually

preceded secondary-style abscission (Fig. 1.7 E,F).

Secondary styles, however, were simply broken off by the

detachment of the primary style in some instances.

Secondary ovaries were generally well differentiated at

the base of secondary pistils (Fig. 1.7 C,D). These ovaries

were ca. 1.5 mm in diameter at anthesis (Fig. 8).

Pericarp tissues of the mature secondary ovary were

similar anatomically to primary-ovary tissues. Most

secondary fruit had their mesocarp partially fused to

primary-fruit mesocarp at the stylar end of the primary

fruit (Fig. 1.7 D,E). Flavedo or exocarp tissue developed on

the abaxial surface of the secondary ovary in areas where no

fusion to the primary ovary occurred (Fig. 1.7 E). The

distal half of a secondary ovary typically had a well-

developed exocarp (Fig. 1.7 E,F). Secondary-fruit endocarp

consisted of segments clustered around a central axis. As in

the primary fruit (105), segments were composed of a

membrane-lined locule, which contained juice sacs and seeds,

and were separated by septa. Juice sacs were observed

shortly after anthesis as randomly arranged, dome-shaped

protuberances from the locular wall which is adjacent to the

mesocarp. The juice sacs in fully developed secondary fruit,

however, were arranged in clusters along the major vascular

bundles which abut the wall of the segment.













$::rord




~I (r) 'V

woow,

> 4

d>1
Li 44J- -4

> .> 3
oP LI H ~

L14 C.I

Uu



LI 'V 4-) H
'-0 Li O-l



Y- awa -



'WL J4- I
-4
4-) CT) L' (0r


I LQr
'V>1 ;T

ETC) OC)H -

H C0 -1 H E

(L) 4k
>1 LU) -r :

;rrj U-
0i m (i

I -I-q .7


>- E o 4N -4 T
C) >i 4J 0
r C U)


UtC I4 4-I
DCT) S= L U)
cU) rT) rq Q

>O H'V ~
m a4 aC


LM c LIU)
M -'04 Li)



ra -q !-- a a)
Hrq> rOa
Li rO C S: C)
CJ4 C "1 "0 k


C






IL4
























cn
U
EC~v


0


M E
C -.0
' *-----

U ^>


- *3


0 U


E


(LULU) I313WVIa )IVAO


0
o -

C-e
LU


0


--
U-



.o



I-



U-
UJ

-


>>-
O C0

< z
0
- U

LU
* ) I


r I 1









The secondary-fruit central axis was an extension of

the primary-fruit one. In fact, the latter axis functions as

the pedicel for the secondary fruit (23).

Axial bundles of the primary fruit continued

acropetally into the distal half of the central axis of the

primary fruit and entered the secondary ovary (Fig. 1.6 F).

These bundles, however, typically terminate after giving

rise to ovular traces and marginal bundles in primary fruit

without a secondary fruit (37,105). Each secondary carpel

had 5 major vascular bundles. Bundles in the mesocarp

consisted of a dorsal bundle opposite the locule and 2

lateral bundles opposite the septa. Two marginal bundles

opposite the septa or the carpellary margins occurred in the

distal half of the central axis of the secondary fruit. All

major carpellary bundles were collateral bundles, and

marginal bundles were inverted. Dorsal and lateral bundles

diverged from axial bundles at the base of the secondary

ovary. Marginal bundles, however, diverged from axial

bundles after the latter had penetrated the proximal half of

the central axis of the secondary fruit. This type of

carpellary vascularization is similar to that of the primary.

carpels as described by Schneider (105). As in the primary

fruit, lateral bundles of 2 adjacent carpels, as well as 2

adjacent marginal bundles, were fused. In addition to the

major vascular bundles, minor ones extended from the primary

fruit into the secondary fruit through their fused mesocarps.








Considerable variability was observed in the size and

development of the secondary fruit. Secondary fruit ranged

from barely noticeable rind tissue only to well-developed

fruit similar to the primary fruit, but much smaller.

Development of the secondary ovary into the secondary

fruit results in a unique fruit structure in navel orange.

The secondary fruit typically is contained within the stylar

end of the primary fruit. As a result, this stylar end

usually is open. The borders of the stylar-end aperture and

the tissues lining the stylar-end cavity are extensions of

the rind of the primary and secondary fruit. The aperture at

the stylar-end of the primary fruit varied from very small

or absent, to about 50 mm in diameter. The peel at the

stylar-end of the primary fruit extended over the secondary

fruit in some instances. As a result, small apertures

sometimes enclosed large secondary fruit. Conversely, large

stylar-end apertures sometimes exposed small secondary fruit.

No correlation has been found between the size of the

stylar-end aperture and that of the secondary fruit (77).

The stylar-end of secondary fruit varied from ridged

to hemispherical and rarely exhibited a stylar-end aperture.

A ridged stylar end usually reflected incomplete fusion of

secondary carpels. Each ridge represented the distal portion

of a secondary carpel. Similarly, carpels of the primary and

secondary fruit of the 'Buddha's Hand' citron are partially

free, and their distal portions vary from ridged to finger-

like (51).









A tertiary fruit frequently was observed within well-

developed secondary fruit. Tertiary carpel primordia were

observed as an additional whorl of primordia within the

secondary-carpel whorl around the floral meristem (Fig. 1.5

D). Tertiary carpels developed into complete pistils

entirely within secondary pistils shortly after anthesis

(Fig. 1.7 E,F). The ontogeny of the tertiary fruit was

similar to that of the secondary fruit.




Secondary'-Fruit Development



Primary and secondary fruit in navel orange followed a

sigmoid pattern of growth (Fig. 1.9) as observed for

primary-fruit growth in navel (14,52) and 'Valencia' (10)

sweet oranges. The stage I of development of the secondary

fruit lasted until late May, approximately 8 weeks after

anthesis. Growth of the secondary fruit during stage I was

due primarily to cell divisions in the peel. The stage II

lasted from late May until early October and was

characterized by rapid cell enlargement in the segments of

the secondary fruit. The stage III consisted of slow growth

of the secondary fruit accompanied by a gradual change of

the color of the peel from green to orange. The beginning of

the cell-enlargement stage (stage II) of the secondary fruit

occurred approximately 2 weeks after that of the primary

fruit. The overall developmental pattern of the secondary

fruit, however, was similar to that of the primary fruit.































Fig. 1.9. Changes in volume of the primary and secondary
fruit, and in diameter of the stylar-end aperture
from anthesis until legal maturity of the fruit
in navel orange, 1980. Each value represents the
mean of a 30-fruit sample.

















0-
.* **
PRIMARY FRUIT
0


eem





A M J 'A S


SECONDARY FRUIT


* 0


*


SM J**


A' M J J A' S 0


STYLAR-END
APERTURE **


0.0 I


0*


A M J J A S O


250-
;

E 150-


E
S50-

0-
0-


5-


c;o
E

u 2-
E
=

0.


9-


E 6
E


S3-
0-

0-








The aperture at the stylar-end of the primary fruit was

not noticeable until mid-May, but then developed rapidly

and reached final diameter in early August (Fig. 1.9).


Secondary-Fruit Protrusions


Three types of tissue protrusions extended from the

secondary fruit into primary-fruit locules, namely, abnormal

placentae, free secondary carpels and secondary-carpel

outgrowths. An abnormally large placenta developed and

almost filled the locular cavity in a few instances (Fig.

1.10 A,B). These abnormally large placentae established

tissue connections with secondary-fruit carpels (Fig. 1.10

A) but could not be distinguished from other types of

protrusions at later stages of primary-fruit development.

Another type of protrusion resulted from free

secondary carpels extending into adjacent primary-fruit

locules (Fig. 1.10 C). These protrusions had juice sacs and

carpellary vascularization and were found in all stages of

primary-fruit development. Development of free secondary

carpels caused separation of the primary-carpel margins in

affected locules.

The most common type of protrusion resulted from

secondary-carpel rind outgrowths extending into the primary-

fruit locules (Fig. 1.10 D). These protrusions also caused

separation of the primary-carpel margins as they extended

into the locules and frequently established direct contact





























Fig. 1.10. Tissue protrusions into primary-fruit locules of
navel orange. A-B, at 20 days after anthesis: A,
abnormally large placenta in a transaction of the
primary and secondary ovaries, and B, normal and
abnormal placentae; C, free secondary carpel in
median longisection of 60 mm-diameter fruit; D,
secondary-carpel rind outgrowths in transaction
at the base of secondary fruit in the primary-
fruit stylar-end. AE, abnormal placenta; CO,
secondary-carpel outgrowth; FC, free secondary
carpel; JS, juice sacs; NE, normal placenta; OV,
ovule; PF, primary fruit; PL, primary-fruit
locule; PO, primary ovary; SF, secondary fruit;
SL, secondary-fruit locule; SO, secondary ovary;
VB, vascular bundle.













































U;








between a locular cavity and the outside of the primary

fruit. Secondary-carpel rind outgrowths into the locules of

primary fruit were found at all stages of primary-fruit

development. These protrusions have been linked to stylar-

end decay (77).




Secondary-Fruit Abscission



An abscission layer was detected in the central axis at

the base of the secondary ovary. The abscission zone was not

easily distinguishable anatomically from adjoining tissues

prior to abscission, as is true for the primary-fruit

abscission zone (19,124). The abscission zone at the base of

the secondary fruit was composed of mesocarp-like tissue

with parenchyma cells surrounding xylem and phloem of the

axial vascular bundles, and parenchyma cells of the pith.

Parenchyma cells in the abscission zone of the secondary

fruit were thin-walled, isodiametric, slightly smaller, and

had fewer intercellular spaces than adjacent parenchyma

cells. Xylary and phloic fibers were less numerous in

vascular bundles in the abscission zone than on either side

of it.

Secondary-fruit abscission was observed only in fruit

with symptoms of secondary-fruit yellowing (Fig. 1.11 A,B),

which results in fruit drop of navel orange from early June

until early August (77,78,79). Abscission or separation of
















C O-1
- 0 f-
C O *H -H
0 t 0 -1 U) X
4 r, .- 4- U). (0
4-i ,-I )4 U)
O O ,- 4 l O L
*H 0 *H 0. XI -,4

H- n -I U, Q U4
I 4- - (l n ( 0.




>iC! 40
4 0 ) ., .C -I



4. O 4
f .. $-4 (d "A ,- 0 )

1 44 -- 0 4 >0 ,


Q ro0 0 ) 0 .,-H
C C co C Q) I 4
O0 o 0 En O
0-,- ) ( () 4- U -
01 g >i U C >1 a
U) 0 f 0 . ..
a) ,-C 4-) O 0)
> 4-) a04 H c
- E C U -, H *


i >,-I 0 r 04 X M
0I OI 00 -C -.
) r 0 C -i'-4 C 4-

-I C -i ( 0
: --i U C ( U
*HI 1 r 0 -0( C=
U) ( U) O rod: m4 -)
01 C r-I (1) JU U 00
-- 1 () (U) >I C
01 O 4-) r- --I U -H

(3 4 M C (0 E1
() 3: 0 0 1 4- H


4-4- 0 U ( 4-
*-I (U 4 O C! 0 <-1 *



^ O (0 *H -A
44 -1n4 O4-) U 3M 44
* H t3 0 Xn >i
4 S O- C En

0 *-Al 0 C (0 (a


rO U r ( -( <- ro
c% W (U 4 4 4 3 C
0 W 4J 4J W 0 0 0
U >1 > 01 U)
()O -44 (0 (T 3 ()
-HO c U O-r > 0






1-1



h(



















IA
































Fii"
...........
..........








.........






.......
.ilF~i;...... ......ii









cells commenced in parenchyma cells that surround the axial

vascular bundles in the abscission layer, and extended into

parenchyma cells of the pith. Parenchyma cells of the

abscission zone underwent cytolysis, which resulted in

development of a gelatinous layer initially around the axial

bundles only (Fig. 1.11 C), but later extending into the

pith (Fig. 1.11 D). Abscission at the base of the primary

fruit, however, starts in the pith and progresses outwardly

to the cortex and epidermis (19,124).

The abscission layer located at the base of the

secondary fruit is typically 2 to 3 cells thick and

approximately 5 to 10 mm in diameter (Fig. 1.11 E).

Parenchyma cells in the abscission layer of the secondary

fruit possessed numerous starch grains during abscission, as

reported for other abscission layers in citrus (19,124).

Starch grains were present also in parenchyma cells on both

sides of the separation layer and in the lysogenous

intercellular spaces created by the disintegration of cells

after abscission of the secondary fruit (Fig. 1.11 F).

Secondary-fruit abscission resulted in the decay of the

secondary fruit, but the latter did not fall since its

mesocarp is typically fused to the mesocarp of the primary

fruit. The decay of the secondary fruit frequently extended

into the primary fruit and caused the drop of the latter.

The primary fruit remained unaffected in some instances,

however, and a protective layer of suberized cells (i.e., a

periderm) was formed on both sides of the secondary-fruit








abscission zone and sealed off the exposed tissues of the

primary fruit. The periderm in the protective layer

consisted of a phellogen which produced a phelloderm (2 to 5

layers of thin-walled parenchyma cells) and a phellem. The

latter was composed of small cork cells and thick-walled,

isodiametric parenchyma cells. As a result of the formation

of the protective layer, the secondary fruit remained in the

stylar-end cavity of the primary fruit as a mummified

structure and the primary fruit did not fall. These primary

fruit, however, frequently were affected by stylar-end decay

later on.






Summary




The secondary fruit (navel) of navel orange develops

as a whorl of secondary-carpel primordia within the

primary-carpel whorl around the floral meristem when the

flower bud is about 1.5 to 2 mm in length. A complete

secondary pistil with fused ovary but separate styles and

stigmas develops entirely within the primary pistil before

anthesis. Stigmas and styles are not as distinct in

secondary carpels as in primary ones. Secondary styles

usually abscise following abscission of the primary style,

but may simply break off.








Secondary and primary fruit have similar sigmoid growth

curves, except the onset of the cell-enlargement stage in

the former lags approximately 2 weeks.

Three types of tissue protrusions from the secondary

fruit into primary-fruit locules were detected, namely,

abnormal placentae, free secondary carpels and secondary-

carpel outgrowths. Abnormal placentae almost fill the

primary-fruit locules and are continuous with secondary-

fruit rind tissue. Free secondary carpels and secondary-

carpel outgrowths extend into the primary-fruit locules and

cause separation of carpel margins. Secondary-carpel

outgrowths were the most common type of protrusion and

related to stylar-end decay.

An abscission layer is present in the central axis of

the primary fruit at the base of the secondary ovary. The

abscission layer is anatomically indistinguishable from

adjacent tissues before abscission of the secondary fruit.

Parenchyma cells in this layer have large numbers of starch

grains during abscission. Secondary-fruit abscission was

detected only during late spring and early summer in fruit

affected by secondary-fruit yellowing.















CHAPTER II

PHYSIOLOGICAL STUDIES ON SECONDARY-FRUIT
YELLOWING OF NAVEL ORANGE






Introduction





Lower yields of navel orange trees have been attributed

to excessive drop of reproductive structures (23,30,67,73,

119). Previous research on fruit drop of navel orange in

Florida included only the fruit-set period from bloom until

early June (26,68,125). However, summer drop between early

June and early August has been significant in 3 out of 4

seasons from 1978 to 1981 with up to 101 fruit, or 15% of

the crop, falling per tree in a single season, virtually

all with symptoms of secondary-fruit yellowing (78).

Secondary-fruit yellowing consists of an early

discoloration of the secondary fruit, or navel, while the

primary fruit remains sound. Secondary decay-organisms and

insects are observed only at late stages of secondary-fruit

yellowing. The problem appears to result from secondary-

fruit abscission (77).









Fruit abscission is typically promoted by ethylene (82,

116), but inhibited by auxins (29,39,53,113). Secondary-

fruit yellowing can be induced by ethylene treatment (112),

but effects of auxins on the development of the disorder

have not been studied.

Water stress may cause abscission of young citrus fruit,

particularly those of navel orange (23,26,119). Carbohydrate

levels in the plant also may influence young fruit drop in

citrus (59,60,74). Neither water nor carbohydrate status

have been studied, however, in relation to summer drop of

navel orange.

Secondary-fruit yellowing was induced by fruit-stem

ringing and effects of time of ringing, distance and number

of leaves between the ring and a fruit and application of

2,4-dichlorophenoxyacetic acid were studied. Levels of

cellulase and ethylene in the abscission zone of the primary

and secondary fruit, and fruit removal force were determined

at different stages of secondary-fruit yellowing. In

addition, the role of leaf abaxial diffusive resistance,

xylem water potential and nonstructural carbohydrates in

secondary-fruit yellowing was investigated.









Literature Review




Secondary-Fruit Yellowing



Secondary-fruit yellowing (SFY) of navel orange occurs

during late spring and early summer when fruit are

approximately 35 to 65 mm in diameter (77,79). The

secondary-fruit rind exposed through the aperture at the

stylar end of the primary fruit becomes yellow, but the

secondary fruit decays only at advanced stages of SFY. The

primary fruit remains green and sound during development of

SFY, but finally becomes yellow at the stylar-end and

abscises.

Insects and microorganisms do not damage secondary

fruit at early stages of SFY (77,79). Later, however,

secondary fruit decays and attracts sap beetles (Coleoptera,

Uitidulidae). Southwick et al. (112) isolated fungi from

decaying fruit affected by SFY and reinoculated healthy

fruit in the presence or absence of ethylene. They concluded

fungi and ethylene are related to SFY but do not appear to

be causal factors. Moreover, SF' was not controlled by

benomyl or malathion sprays (77,78).

Secondary fruit of fruit with early symptoms of SFY

were found to have separated from the primary fruit through

a zone at the base of the secondary ovary resembling an

abscission zone. Separation of the secondary fruit in fruit









affected by SFY in the absence of primary insect or fungi

damage suggests the possibility of a physiological cause for

summer drop of navel orange in Florida. Fruit affected by

SFY produce large amounts of ethylene (112), but other

physiological aspects of SFY and secondary-fruit abscission

have not been studied.




Fruit Abscission



The abscission process. Abscission is brought about by

the loss of cementing capability of cell wall material in

the separation layer, followed by mechanical breakage of

nonliving vascular elements and the epidermis (11,92,116).

Leopold (72) divided the abscission process into 5 stages.

Differentiation of an abscission zone usually occurs prior

to the time of most active fruit enlargement. A holding

stage follows in which no weakening occurs in the abscission

zone. This stage usually lasts for the functional life of

the subtending organ. The holding stage is followed by

stages of structural weakening of the abscission zone,

separation, and healing. Not all these stages, however, are

essential, e.g., the abscission zone at the base of citrus

fruit is not clearly differentiated, and no healing occurs

of the exposed surface on the plant after fruit abscission

(92,124).

Abscission of a citrus fruit begins with the swelling

of cell walls in the separation layer followed by cell wall









dissolution and exudation of cellular contents into the

break created by separation. The separation layer becomes

gelatinous with no distinguishable intact cell structures,

except for tracheary elements and the epidermis (124).

Cytolysis during abscission has been linked to

increased enzymatic activity. Wilson and Hendershott (124)

detected significant demethylation of pectins in the fruit

abscission zone, which produced a distinct band of cells

low in methylated pectins, the separation layer. Pectins act

as cementing substances between cells and are linked by

calcium and magnesium ions (101). Loss of these ions from

cells in the separation layer during abscission indicates

the involvement of pectinases, e.g., polygalacturonase (13,

87,97,114,124).

Biggs (13) related the swelling of cell walls in

separation layers of citrus to a decrease in cross-linkage

among polymers and endobreaking of polymers. He concluded

that cellulases are involved in the abscission of citrus

fruit, as reported previously (1,93,97). Cellulase activity

has been related to reduced fruit removal force in citrus

(42,63). Other enzymes-peroxidase, dehydrogenase, and acid

phosphatase-also have been associated with the abscission

process (11,72).

Accumulation of starch has been reported in cells of

the separation layer prior to abscission in several plants

(11) including citrus (19,124). No explanation for this

phenomenon has been proposed. Large numbers of accumulated









starch grains which remain intact throughout abscission

eventually appear in the intercellular spaces formed after

cell separation (19,124).



Growth regulators. Growth regulators are intimately

involved in fruit abscission (82,116). Abscission can be

inhibited or promoted by auxins but is inhibited in most

cases (13,45,72,116). Abscission promotion by auxins appears

to result from the induction of high levels of ethylene,

especially at high concentrations of auxin (116). The delay

in abscission by auxins occurs principally through

inhibition of passage out of the holding stage (72).

Progression from the holding into the structural weakening

stage apparently is due to a decline in endogenous auxin

levels (72,116). Inhibition of abscission by auxins has been

related in some instances to decreased activity of cell wall

macerating enzymes such as cellulases and polygalacturonase

in the abscission zone (13,41,97,99). Auxin sprays prevent

fruit abscission in several crops (11), including citrus (5,

29,39,83,84,89). The effect, however, is dependent upon

concentration and species or cultivar.

Ethylene generally has a stimulating effect on

abscission (3,72,82,116), and may be the major regulator of

the process, with other factors acting through inhibition or

promotion of ethylene (116). Ethylene is a strong promoter

(24,25,92) as well as a natural product of citrus abscission

(92). Ethylene affects abscission primarily at the stage of









structural weakening when senescence has commenced in the

tissues distal to the separation layer (72). Abscission in

young citrus fruit is not promoted or hastened by ethylene

or ethylene-generating compounds in some instances (53,64,

85). This nas been attributed to the lack of tissue

sensitivity (53,75) and high levels of auxin (85). A

decline in auxin levels with aging increases tissue

sensitivity to ethylene and allows abscission. Ethylene

appears to promote abscission by increasing the activity of

cell wall hydrolases, particularly cellulases (3,13,43,97)

and polygalacturonase (43,99), and by inducing senescence

(1,2,3).

Gibberellins, abscisic acid and cytokinins also

influence abscission (27,72,96,116). Gibberellins seem to

promote abscission when applied to abscission zones (116)

or entire citrus trees (68), but prevent abscission when

individual fruit are treated (21,68). Abscisic acid

generally enhances fruit abscission (3,11,116), probably by

promoting senescence of distal tissues (72) and stimulating

ethylene and cellulase biosynthesis (3,72). Cytokinins

weakly inhibit abscission and are thought to act like auxins

by delaying the completion of the holding stage or slowing

down senescence (72,82,116).

Hormone interaction and balance have been suggested to

control abscission (3,18,116). Most hypotheses involve an

auxin-ethylene balance, because the effects of these

hormones on abscission are better known (18).








Materials and Methods




Plant Material



Navel orange (Citrus sinensis (L.) Osbeck) trees on

sour orange (Citrus aurantium L.) rootstock, planted in 1957

were used for experiments in 1980. Similar trees, planted in

1969, were used in 1981 and 1982. Trees were located near

Eustis, in the north central citrus region of Florida.

Experimental trees received general care, including

irrigation, typical of groves with fruit destined for the

the fresh-fruit market.



Induction of Secondary-Fruit Yellowing



Secondary-fruit yellowing (SFY) was induced by fruit-

stem ringing from late May until late July. This consisted

of the removal of a 1 cm ring of bark from the fruit stem,

5 to 10 cm from the fruit, and excision of all leaves

between the ringed area and the fruit. Secondary-fruit

yellowing was also achieved without leaf removal so long as

the ringed area was within 10 cm of the fruit.



Application of 2,4-D and Ethephon


Applications of 2,4-dichlorophenoxyacetic acid (2,4-D)

were made to whole trees, whole fruit, or to parts of fruit









or fruit stem. Whole-tree applications consisted of 20 ppm

2,4-D sprays in 55 liters of solution on April 19 or May 6,

1982. Whole-fruit applications consisted of 4 to 8 biweekly

sprays of 10 ppm 2,4-D before or after fruit-stem ringing.

Applications of 2,4-D to the ringed area or the stylar-end

of stem-ringed fruit were made at the time of ringing as a

1000 or 2000 ppm slurry in lanolin paste. Control fruit

received lanolin without 2,4-D. Control fruit and trees were

not sprayed. Applications of ethephon (2-chloroethyl-

phosphonic acid) to the stylar-end of unringed fruit were

made as a 10 sec dip in a 300 ppm solution.




Ethylene, Cellulase and Fruit Removal Force



Ethylene concentration and cellulase activity in the

abscission zone at the base of the primary and secondary

fruit, and fruit removal force (FRF) were determined at

different stages of SFY. Secondary-fruit yellowing stages

were 1, green (healthy secondary fruit), 2, early (slight

discoloration of the secondary fruit), 3, moderate

(yellowing but no secondary decay), 4, advanced (yellowing

and secondary decay), and 5, very advanced (advanced decay

and yellowing symptoms spreading to primary-fruit areas

around the secondary fruit). Ten to 20 fruit were collected

at each stage and determinations made on the same day.

Twenty to 40 fruit were analyzed daily. Determinations were









made in 7- to 10-day periods during the peak of SFY

incidence in late June to early July in 1981 and 1982.

Ethylene concentration was determined by gas

chromatography on samples taken with disposable syringes

from central axis tissues which included either the primary-

or the secondary-fruit abscission zone. Fruit were cut in

half transversally under water, a hypodermic needle was

placed either into the central axis of the stem half toward

the primary-fruit abscission zone or of the stylar half

toward the secondary-fruit abscission zone, and a 1 ml gas

sample withdrawn. Samples were injected into a Hewlett-

Packard 5710A gas chromatograph equipped with a flame

ionization detector and a stainless steel 1 m x 6.35 mm ID

column packed with activated alumina. Flow rate was 75 ml

per min using nitrogen as a carrier gas. Oven temperature

was 100 C.

Cellulase activity was determined by viscometry (76).

Samples of 30 to 50 mg of tissue containing the abscission

zone plus a small amount of adjacent tissue were excised

with a scalpel. Cellulases were extracted by grinding the

tissue sample with mortar and pestle in 2 ml of 20 mM sodium

phosphate buffer (pH 6.5) and centrifuging at 10,000 g for

10 min (63). Extraction was done at temperatures under 4 C.

Cellulase activity was assayed in the supernatant by

measuring the reduction in viscosity of a 1.2% solution of

carboxymethylcellulose (CMC) type 7H3SF (Hercules Powder

Co., Wilmington, Del.) in 20 mM sodium phosphate buffer









(pH 6.5) (98). The assay mixture consisted of 0.4 ml of CNC

solution and 0.2 ml of the enzyme extract. Viscosity was

determined by measuring drainage time of the assay mixture

through a calibrated portion of a 0.1 ml pipette between the

0.00 and 0.03 ml marks (76). The change in viscosity from

time zero (time of mixing enzyme and substrate) and 1 to 2

hours of incubation at 24 C was used to measure enzyme

activity. Data were converted to units of cellulase activity

per abscission zone and time (hour) (7,76).

Fruit removal force (FRF) was measured by clamping the

fruit-explant stem to a Chatillon strain gauge (John

Chatillon & Sons, New York, NY) and applying tension

manually by pulling the fruit until separation from the stem

occurred. The FRF was the maximum force required for

separation.




Water and Carbohydrate Status



Secondary-fruit yellowing was induced by fruit-stem

ringing with no removal of leaves between the ringed area

and the fruit. Control fruit were not ringed. Four fruit

were ringed per replication with 5 replications per

treatment.

Branches were sawed halfway at 3 locations

approximately 5 cm apart on alternate, opposite halves of

the axis in an attempt to restrict xylem translocation









although phloem translocation was also restricted. Branch

scoring consisted of a single knife-cut made through the

bark around the main axis in an attempt to restrict phloem

translocation only. Branches sawed or scored, and control

untreated branches were ca. 2.5 cm in diameter and held an

average of 13 fruit. Ten branches were used per treatment.

Trunk girdling consisted of a single knife-cut through

the bark around the trunk at 10 to 20 cm above the bud

union. Control trees were not girdled. Five single-tree

replications were used per treatment.

Leaf abaxial diffusive resistance, xylem water

potential, and total nonstructural carbohydrates were

determined on fully expanded spring-flush leaves after

induction of SFY by fruit-stem ringing, and after branch

sawing or scoring. Total nonstructural carbohydrates of

leaves were also determined after trunk girdling. Leaf

measurements were made on 1 subtending leaf (closest to the

fruit) per ringed fruit, 2 to 3 leaves per sawed or scored

branch, or 10 leaves per girdled tree. Leaves from treated

branches or trees were collected from fruitless shoots

located randomly on the outside of the canopy at 1 to 2 m-

height.

Leaf abaxial diffusive resistance was measured with a

LI-COR LI-1600 steady state porometer. Leaf xylem water

potential was obtained using the Scholander pressure chamber

technique (103). Leaf abaxial diffusive resistance and xylem

water potential were measured between 12 and 2 PM. Total








nonstructural leaf carbohydrate level was determined by

treating leaf extract suspensions with invertase,

amyloglucosidase and takadiastase (81). Enzyme-digested

suspensions were then analyzed for reducing sugars according

to a modified nielson copper reduction test (90,108).



Evaluation of Secondary-Fruit Yellowing



Secondary-fruit yello':ing incidence was evaluated after

fruit-stem ringing and growth regulator treatments through

weekly inspection of treated fruit for 25 to 45 days after

ringing. Total number of SFY-affected fruit was obtained for

the entire period and expressed as % of treated fruit.

Secondary-fruit yellowing after branch sawing or scoring was

estimated by counting the number of affected fruit from June

10 until August 6 and expressed as % of the initial number

of fruit per branch.




Experimental Designs



Completely randomized or randomized complete block

designs were used. Mean separation was done by Duncan's

multiple range test, utilizing Kramer's adaptation for

experiments with a variable number of replications (65).








Results and Discussion




Maximum SFY was obtained from fruit-stem ringing on

May 26 and 28 in 1982 (Table 2.1). The % secondary-fruit

yellowing was lower on dates prior to or after these, with

no SFY induction achieved on May 6 or August 6. Natural SFY

occurred from early June until early August with a peak

during late June or early July. It is possible that the time

of peak natural induction of SFY was also during late May in

1982. Similar results were obtained in 1981, with maximum

SFY induction obtained from fruit-stem ringing performed on

June 15 (data not shown), or about 2 weeks later than in

1982. Full bloom and petal fall in 1981 also occurred 2 to 3

weeks later than in 1982. As a result, fruit size at the

time of peak SFY was comparable in both seasons and ranged

from 45 to 55 mm in diameter.

Fruit-stem ringing of navel orange induced higher

ethylene levels in the stylar-end of the fruit 7 days after

ringing, accompanied by SFY, which was followed by fruit

drop (Table 2.2). Although ethylene is produced in plant

tissues after wounding (24,82,116), the responses to fruit-

stem ringing probablywere not due to high ethylene levels in

treated fruit. No increase in ethylene concentration was

detected after fruit-stem ringing when SFY was inhibited by

2,4-D application (Table 2.2). In addition, ringing did not

increase ethylene levels or produce SFY symptoms or fruit

drop in 'Hamlin' orange (Table 2.2), which typically does




61



















Table 2.1. Secondary-fruit yellowing (SFY) of navel orange
as affected by date of induction by fruit-stem
ringing, 1982.


Date of
fruit-stem ringing


SFY
(% of treated fruit)z


May 6 Oc'
May 21 60b
May 26 90a
May 28 90a
June 3 65b
August 6 Oc


Ten to 30 fruit treated per date. Evaluation of SFM' was
done until 25-40 days after ringing.
'Unlike letters indicate significant differences by Duncan's
multiple range test, It level.
















Table 2.2. Fruit ethylene, secondary-fruit yellowing (SFY),
and fruit drop of navel and 'Hamlin' orange
following fruit-stem ringing and ethephon (ETH)
or 2,4-dichlorophenoxyacetic acid (2,4-D) stylar-
end treatments, 1982.


Fruit ethylene (ppb)y
SFYX Fruit dropx
Treatments Stem-end Stylar-end (%) (%)

Navel
No ringing 70aw 74b Oc Oc
Ringing 96a 347a 90a 90a
No ringing + ETH -v 65b 65b
Ringing + 2,4-D 88a 126b Oc Oc
'Hamlin'
No ringing 71a 97b -
Ringing 97a 82b -


ZTwo to 6 fruit per replication, 5 replications per
treatment, applied on May 28. Stylar-end treatments
consisted of a 300 ppm ethephon dip or an application of
lanolin paste containing 1000 or 2000 ppm 2,4-D.
YSampled from the central axis at the primary-fruit
abscission zone in the stem-end, and at the secondary-fruit
abscission zone in the stylar-end, 7 days after treatment,
7 fruit per treatment.
XEvaluated until July 16. Fruit drop followed SFY in all
instances and both are expressed as % of treated fruit.
Unlike letters within columns indicate significant
differences by Duncan's multiple range test, 1% level.
Not determined or not applicable.








not have a secondary fruit (51). Fruit growth, however,

ceased after fruit-stem ringing in both navel and 'Hamlin'

(data not shown). It appears that higher fruit ethylene

levels after fruit-stem ringing in navel orange occur after

rather than before secondary-fruit abscission and SFY.

Fruit abscission is typically promoted by ethylene

(116), but inhibited by au:xins (29,39,53). Navel oranges

treated with 2,4-D at the stylar-end did not respond to

fruit-stem ringing. Dipping the fruit stylar end in a 300

ppm ethephon solution for 10 sec, however, did result in

SFY, followed by fruit drop (Table 2.2), as previously

reported (109,112).

Ethylene levels and cellulase activity began to

increase in the abscission zone of the secondary fruit

during stage 2 of SFY, when visible discoloration was first

detected (Fig. 2.1). It appears that secondary-fruit

abscission preceded development of SFY symptoms and

subsequent invasions by sap beetles and decay organisms.

Conversely, high levels of ethylene and cellulase activity,

as well as reduced fruit removal force, were observed in the

primary-fruit abscission zone only at advanced stages of SFY

(Fig. 2.1). Fruit with SFY symptoms produce high amounts of

ethylene (112). Ethylene induces abscission probably by

stimulating cellulase activity (3,13) which in turn reduces

fruit removal force (12,63). It appears that secondary-fruit

abscission precedes and is the cause of primary-fruit

abscission during summer drop.






























Fig. 2.1. Ethylene, cellulase activity, and fruit removal
force in the abscission zone at the base of the
primary and secondary fruit in relation to stage
of secondary-fruit yellowing of navel orange, 1982.
Each value represents the mean of a 10- to 20-
fruit sample SE. Stages of secondary-fruit
yellowing were 1, green (healthy secondary fruit,
SF), 2, early (slight discoloration of SF), 3,
moderate (yellowing but no decay), 4, advanced
(yellowing with decay), and 5, very advanced
(yellowing with advanced decay).














Abscission Zone
*Primary Fruit
oSecondary Fruit


. 44


1 2 3 4 5



T i


15
2=_
-

3
S10-

5-

0-


S A


I i
2 3
of Secondary-fruit


4 5
Yellowing


S12-
U
U-
- 8-
CD
0
E
C 4-

L-
LL_


i
Stage








Water stress has been implicated in the abscission of

young citrus fruit, particularly those of navel orange (23,

26,50,122). Carbohydrate levels in the plant also may

influence fruit drop during the fruit set period (59,60,74).

Fruit-stem ringing resulted in no change in leaf abaxial

diffusive resistance, xylem water potential or total

nonstructural carbohydrates but did induce secondary-fruit

yellowing (Table 2.3). Conversely, branch sawing and scoring

treatments resulted in changes in leaf water and

nonstructural-carbohydrate status during the SFY-induction

period in late May but did not influence summer fruit drop

in 1981 (data not shown) and in 1982 (Table 2.4). In

addition, trunk girdling in early May increased the levels

of nonstructural carbohydrate in the leaves but had no

effect on summer fruit drop or yield (Table 2.5 and Chapter

III, Table 3.9). Apparently, the water andnonstructural-

carbohydrate status of fully expanded spring-flush leaves

are not associated with SFY.

Fruit-stem ringing at 5 to 10 cm from the fruit, with

or without leaf removal, resulted in induction of SFY in 80

to 100% of treated fruit (Table 2.6). Significantly less SFY

induction was achieved as the distance of the ringed area

from the fruit, and the number of leaves above the ringed

area increased. Furthermore, removal of all leaves up to 30

to 45 cm from the fruit in the absence of fruit-stem

ringing did not induce SFY (Table 2.6). Secondary-fruit

yellowing, therefore, can be induced when bark-translocation






















Table 2.3. Leaf abaxial diffusive resistance(ADR), xylem
water potential (Vx), totalnonstructural
carbohydrates (CHO), and secondary-fruit
yellowing (SFY) of navel orange as affected by
fruit-stem ringing, 1982.

Leaf measurements"
SFY
z -1 -1
Treatments ADR(s cm ) Yx(MPa) CHO(mg g d..) (A)

Control 0.19 -1.55 51.9 0
Ringing 0.32ns -1.39ns 41.5ns 65**


Four fruit per replication, 5 replications per treatment,
applied on June 3.
'One fully expanded spring-flush leaf subtending each fruit.
ADR and 'x were evaluated on June 10 and CHO on June 29.
xEvaluated until July 16 as % of treated fruit.

ns,**Honsignificant (ns) or significant at 1% level (**) by
a t-test.





















Table 2.4. Leaf abaxial diffusive resistance (ADR), xylem
water potential ('x), total nonstructural
carbohydrates (CHO), and secondary-fruit
yellowing (SFY) of navel orange as affected by
branch sawing or scoring, 1982.

Leaf measurements
SFYX
Treatments ADR(s.cm ) Yx(MPa) CHO(mg g d.w.) (%)

Control 4.31bw -1.30b 98.2a 7.8a
Sawing 8.58a -1.59a 72.5b 7.6a
Scoring 4.04b -1.34b 115.6a 9.3a


ZTen branches (2.5 cm-diameter) per treatment, 13 fruit per
branch, applied on May 8. The main axis was sawed halfway
transversally at 3 locations 5 cm apart. The cuts were made
on alternate opposite halves of the axis in an attempt to
restrict xylem translocation although phloem translocation
was also restricted. Scoring consisted of a single knife-
cut through the bark around the main axis in an attempt to
restrict phloem translocation.
YDetermined on May 21 using 2 to 3 fully expanded spring-
flush leaves per branch.
xFrom June 10 to August 6. Expressed as % of initial number
of fruit per branch.
wUnlike letters within columns indicate significant
differences by Duncan's multiple range test, 5% level.



























Table 2.5. Leaf totalnonstructural carbohydrates (CHO) of
navel orange as affected by trunk girdling, 1982.

-1 .
CHO(mg g d.w.)

Treatments May 15 flay 21 June 29

Control 106 98 65
Trunk girdling (May 6) 106ns 99ns 72*


ZTrunk girdling consisted of a single knife-cut through the
bark around the trunk 10 to 20 cm above the bud union.
YTen fully expanded spring-flush leaves per tree and 5 trees
per treatment were used at each date.
ns,*Nonsignificant (ns) or significant at 5% level (*) by a
t-test.





















rU L r







o o o
r-0 00


I
*4

N4 00
H o
oo






Sr-f
4-4

Oco








,-4
r--4

O





r0










U)r-



-4J
O









4-1
(




o r


0

O -

> (0







E-4






-Q
IC






ki
*H











.0
4
*~-,If
3 Uo

^1Cu
a, n
I


3 o~


a,
4-1

rd



O
olo
r-4

0
o,





00
I- --k





>


U)
a,
O C
f0 O





E3-
.0 Q) 4-
0



















N


a,
E
'4-
0a,
0(


mC
0 I
(c


U

O

>


0
4-)
a4





an 44
(a1



> 4-4
rd 0
a,







-H
U
rnm





0
0
S4E

o s-
2 4-1


x
.-
0











>1
a




























44



0
-,-I

.H
Cd

*r-

*H >
O 0
E
0 z(
Z 0


E

0

o>
O
I (




tn >
~cn


-4l a,


o o
0
r-4


0
' %D


I 1







I I


on

o
0
(N














tUo


I f
0 >
n

4-1 (-)

3

0H a

"-4 0


a0

4-



Q
S,
O-
-H 0


LH H
0

,4 u)-
03 O



rn





*H
Q) 0



a,


03






aO
a,-I





O O
ko



0




0a>
Sa,


Q)
















OO
N-)
m r




0 4


0
r-I
0, >1
O)
N a
41


0 rfl


3 0

N


U)



C
0







aQ

4)










Ini
rd
.a

o












4
0









-a,
n



U)










- )








4-,-
.H (1)
-4-4






0)







-P
(U

4-,


3
0












Q)


E
U4

C0 r-

I >
n oo

4-1 ,)
E ,
c-a,



-H a,

*r 0
-HO c


n >
r-A 0
E


44(0
r-d
Cd
*H a,

r 0








of leaf metabolites to the fruit is limited. Little SFY,

however, results from fruit-stem ringing if enough leaves

remain between the ringed area and the fruit, and sufficient

leaf metabolites are supplied to the fruit. Brown (16) and

Powelland Krezdorn (95) found that movement of 14C-

metabolites from the leaf to the fruit influenced citrus

fruit development and young-fruit abscission.

Induction of SFY by fruit-stem ringing was prevented

by applying 2,4-D to the ringed area, the stylar-end, or as

whole-fruit sprays prior to or after ringing (Table 2.7).

The only exception was with 2000 ppm 2,4-D applied to the

ringed area. High auxin concentrations may promote

abscission by stimulating ethylene biosynthesis (116).

Secondary-fruit yellowing appears to result from secondary-

fruit abscission; hence, reduction of SFY by 2,4-D may

occur through inhibition of abscission of the secondary

fruit. Auxins act as strong abscission antagonists in citrus

(13,45,53) as well as in other plants (72,116). Leopold (72)

suggested that the progression from the holding into the

structural weakening stage of abscission is due to a

decline in endogenous auxin levels, which increase tissue

sensitivity to ethylene and allow abscission (85). It is

possible that auxins or auxin-precursors are some of the

bark-translocated leaf metabolites that would act similarly

to exogenous 2,4-D. Ringing and leaf removal would limit

auxin supply to the abscission zone of the secondary fruit,

thereby allowing abscission and the development of SFY














Wa,

II -40) 0l

S- o o ua
-1 0 04 0 0 Q

*d ro C) CD CD0 1 C l
3 0 -4 o 4-i U) ( r
r-i >1 r- (10) 4

S* ( O co rO j
r-j 4--i 4- a00 0 0 (1
00 0- Q4 Q4 ., C
SU j 04 4 0 '( Wro )
( O 1 -0 rd 01 a

x 4 O CC o o I T fo (U) 0
f 0 44 1 (1) m r- Q4 .-I un 4 4H
4CO 0 1n >4 O 0d c4
0 a rn r-1 cn -
Oa Z 0 .) O C 0
-1 0 (0 44 H Q) r-. O 0 -1J
> 0 -i 0 *0 4o
CO O -, -> 1 .
0o -O Ir-I' i Cl)
> O L i a Q > 4
S 0 4- *o0 Wo C-
U 4) rl Q a 4-00
4- O 4 3 0 1 4 --


0 i U ,--I ( a--I

,--( ) 44 o o
Or C -a o l a) c
I dP 4 -.O .-. () 0- -




O' ,-
00 4 0 I O -O 4 r-
>1 r- O- OA (a 4- j 3 >C
4-) I 0 0 0- U
U) 'ar t0 c(, 0


So o o 4U 0\0 Id -I4


Sr- 0 .0 .4 1 O Cd'

Z l N 0) *H 4- r tP Z


N 4-) En 4- () r. *i 4 f
Crr a Q -r. f
r0l E- d C 0 4t ) f-
5 40i 4- 00+N 0 r-I 50 C O4
04 r- C C -I -0 .' -I JO Cd c (>









a, 4 o : C4 0 -a, C:wz
Nn Cd 0 4-) C 41 4- (0




4 U r rl u O O -l S- ra
-4 d C -I 00 C >1 0






C 4 Q C 4 -i O O- 0 Q c
c *d 0 I I I I 1 0 U) 1 0 i 4H
: (c1 0 ::1 -- O4 C: ( ) -4
Or-i n N 1 N N I N 5H N0 -r-4 l (d 1 0W


0 01 0 0 ao 0-*d C P -
S4 rU) 0 -4 4O -1 a ,




E-U) 4 N >, C *( C
a, *4 .4 L i W0'. -0 .I 4)




(I a 0 *H -* o o +C d -4-4

,N > X








symptoms. Accordingly, SFY which occurs during summer fruit

drop may result from limited supplies of auxins or auxin-

precursors to tne developing fruit. Whole-tree 2,4-D sprays

as early as full bloom were found to prevent SFY and

significantly reduce summer fruit drop (Chapter III, Tables

3.7,3.8,3.9). In addition, attempts to induce SFY by fruit-

stem ringing of fruit on 2,4-D sprayed trees were

unsuccessful (Table 2.8). Sprays of 2,4-D were applied 3 to

5 weeks prior to fruit-stem ringing which made the fruit

unresponsive to SFY induction.

Trunk girdling increases gibberellin levels (06,120)

and fruit set in citrus (66,69,86). Trunk girdling of navel

orange during early May resulted in increased leaf

nonstructural-carbohydrate levels within 7 weeks of

treatment (Table 2.5), but did not prevent SFY and summer

fruit drop (Chapter III, Table 3.9), or induction of SFY

by fruit-stem ringing (Table 2.8).






Summary




Induction of secondary-fruit yellowing (SFY) of navel

orange by fruit-stem ringing or stylar-end ethephon

treatment and its prevention by 2,4-D application suggest

a physiological nature for the disorder. This contention is

























Table 2.8. Secondary-fruit yellowing (SFY) of navel orange
following fruit-stem ringing as affected by 2,4-
dichlorophenoxyacetic acid (2,4-D) whole-tree
application or trunk girdling, 1982.

Fruit-stem SFYX
Treatments ringingy (%)

No treatment No Obw
No treatment Yes 89a
2,4-D on April 19 Yes 25b
2,4-D on May 6 Yes 15b
Girdling on May 6 Yes 80a


ZFive to 10 trees per treatment. Sprays of 2,4-D were made
at 20 ppm in 55 liters of solution per tree. Trunk girdling
consisted of a single knife-cut through the bark around the
trunk 10 to 20 cm above the bud union.
YRinged on May 28.
XEvaluated until July 16 as % of treated fruit.
WUnlike letters indicate significant differences by Duncan's
multiple range test, 1% level.








further supported by a cause-and-effect relationship between

secondary-fruit abscission and SFY.

Secondary-fruit abscission precedes primary-fruit

abscission and the levels of both ethylene and cellulase

activity increase in the abscission zone of the secondary

fruit prior to that of the primary fruit affected by SFY.

Reduced fruit removal force also occurs only at advanced

stages of SFY. Abscission of the primary fruit, which

follows SFY, may result from increased ethylene levels after

secondary-fruit abscission.

Induction of SFY by fruit-stem ringing was not

associated with changes in ethylene levels of the fruit or

the water and nonstructural-carbohydrate status of

subtending leaves. Moreover, changes in water and

nonstructural-carbohydrate status of leaves induced by

branch sawing or scoring, or by trunk girdling had no

significant effect on SFY incidence.

Fruit-stem ringing and leaf removal experiments

indicated induction of SFY resulted from an interrupted

supply of bark-translocated leaf metabolites to the fruit.

Applications of 2,4-D to the whole tree, fruit, or to the

ringed area on the fruit stem, before or after ringing,

however, reduced induction of SFY from fruit-stem ringing.















CHAPTER III

GROWTH REGULATOR EFFECTS ON FRUIT DROP,
YIELD AND QUALITY OF NAVEL ORANGE






Introduction




Fruit drop after the fruit-set period is significant

(77,78,79), and may be responsible for lower yields of navel

orange trees in Florida. Three major causes of fruit drop

have been characterized, secondary-fruit yellowing, stylar-

end decay, and fruit splitting, all of which have been

associated with the morphology and anatomy of the stylar-end

of the navel fruit, which encloses the secondary fruit (77,

79). Secondary-fruit yellowing which causes most of the

summer fruit drop appears to result from abscission of the

secondary fruit. Stylar-end decay and fruit splitting, the

major causes of the summer-fall drop, more frequently affect

fruit with larger stylar-end aperture.

Growth regulators influence fruit abscission (116).

Auxin applications delayed or prevented preharvest fruit

drop of sweet oranges in California (20,113) and of

'Pineapple' and seedling sweet oranges in Florida (39). In









addition, applications of 2,4-dichlorophenoxyacetic acid

(2,4-D) in combination with gibberellic acid (GA) to

individual flowers caused the peel of the primary fruit to

extend over the secondary fruit resulting in smaller or

absent stylar-end aperture in navel orange (68). Growth

regulators, however, have not been examined in terms of

their effects on summer and summer-fall fruit drop of navel

oranges in Florida.

The objectives of this research were to study the

effects of growth regulator sprays on summer and summer-fall

fruit drop and on the external and internal characteristics

of the fruit of navel orange in Florida.






Literature Review




Citrus Fruit Drop



Mature fruit usually develop from less than 1% of the

reproductive structures produced by a sweet orange tree (33,

79). Shedding of reproductive structures takes place during

all stages of development from flower buds to ripe fruit,

typically occurring in cycles or waves (19,22,23,73,79). The

first wave, bloom drop, starts at the flower-bud stage, has

a peak during bloom and involves abscission of flower buds,

flowers and young fruit (19,33). Abscission generally occurs








at the base of the pedicel. Most of the reproductive

structures of sweet oranges fall during bloom drop.

Erickson and Brannaman (33) reported 97.5% of the fruit of

navel orange and 96% of those of 'Valencia' were shed before

they reached 10 mm in diameter. Similarly, in Florida, 88.7%

of navel orange ovaries were shed during the bloom drop

period (79). The cause of bloom drop has not been determined,

but malfunction of reproductive structures (19,54) and

competition for metabolites (32) have been suggested.

Senescing citrus flowers produce high levelsof ethylene, but

fruitlet abscission appears independent of ethylene

production (111). Bloom drop may be intensified by diseases

(35,36), insects (44) and temperature or water stress (23,

50).

A second peak of fruit drop, "June drop", occurs from

mid-May to early July in California (22) and from late April

to early June in Florida (19,46,78,125). Sweet orange losses

during June drop range from insignificant (33) to almost

total (122). Abscission typically occurs at the base of the

ovary.

June drop is probably caused by competition between

fruitlets for metabolites (32). High temperature, water

stress, or both appear to accentuate the problem (23,56,122).

Several fungi have been isolated from fruit during June

drop, the most prevalent being Alternaria spp. (23,112,119).

These fungi, however, also have been detected in healthy

fruit, and attempts to control June drop with fungicides









have been unsuccessful (23,79,119). Fruit remaining on the

tree after the June drop period typically reach commercial

maturity with little further drop in most sweet oranges. The

period from bloom until the end of June drop is considered

the fruit-set period (67).

A third wave of drop in sweet oranges is preharvest

drop. This drop affects fully developed fruit and becomes

increasingly important during on-tree storage of fruit (5,

31,39,55). Some causes of preharvest drop are fruit

splitting, brown rot (Phytophthora spp.), black rot

(Alternaria citri Ellis & Pierce), stem-end rot (Diplodia

natalensis Pole-Evans), twig dieback, and freeze injury (31,

55,62). Many apparently sound fruit, however, also fall for

unknown reasons during prolonged storage on the tree (31).

Fruit drop continues after the fruit set period and

before preharvest drop for navel orange in Florida (78,79).

Summer drop, which occurs from early June until early

August, and summer-fall drop,which occurs from late August

through October, have been characterized (78,79). SufLmmer and

summer-fall fruit drop reduced yield by as much as 17% and

15%, respectively, in a navel grove in Florida in 1979 (79).

The major causes of navel fruit drop after fruit set

are related to the morphology and anatomy of the fruit

stylar-end and the secondary fruit. Summer drop is caused by

a yellowing of the secondary fruit resulting from its

abscission from the primary-fruit central axis (Chapter I,

p. 41, and Chapter II, p. 63). Fruit splitting, stylar-end








decay, branch collapse and brown rot (Phytophthora spp.)

are responsible for most of the summer-fall drop (79). Fruit

splitting and stylar-end decay more frequently affect

fruit with larger stylar-end aperture. In addition, most

fruit affected by stylar-end decay have rind tissue

protrusions from the secondary fruit into the affected

locules (79). These protrusions are present in fruit locules

before any decay has taken place and consist of secondary-

carpel rind outgrowths (Chapter I, p. 38).





Control of Citrus Fruit Drop



Several attempts have been made to increase fruit set

in citrus. Cross-pollination is recommended as a means of

increasing fruit set and yield in Florida for 'Orlando'

tangelo and other tangerine-type hybrids (67). Cross-

pollination of navel orange with other citrus cultivars

has been successful in South Africa (28) and Egypt (30), but

not in Florida (67). Cross-pollination by bees is difficult

because navel flowers produce little pollen and attract

nectar-gathering but not pollen-gathering bees (28).

Girdling or scoring of the trunk or major branches

during bloom generally reduces fruit drop during the fruit-

set period and increases yield (66,67,71,86). Navel oranges,

however, do not respond consistently to girdling. Yield

increases were achieved in the first year of treatment in








some instances (66), but girdling over several successive

years did not result in higher cumulative yield compared to

untreated trees (6,107). Azzouni and Mahmoudi (9) reported

girdling to be ineffective in increasing yield of nav.el

orange in Egypt.

Some researchers have suggested excessive abscission of

navel fruitlets to result from high temperature, wind, or

water stress (23,50,67,73,122). Accordingly, overhead

misting or application of an antitranspirant spray shortly

after bloom resulted in higher leaf water potential and

increased fruit set of navel orange in Florida (26). In

addition, overhead sprinkling of navels in California

during periods of high temperature significantly reduced

air and leaf temperature and increased fruit set (15).

Hormones influence fruit set and development (40,61,

125). Applications of auxins (38,94,113) or cytokinins (38,

88,110), however, usually have little or inconsistent

effects on citrus fruit set. Gibberellin (GA) applications

to individual flowers, flower clusters or small branches

consistently result in increased fruit set in navel orange

as well as other citrus (38,48,68,109). Results for whole-

tree GA applications, however, generally have been

inconsistent. Hield et al. (48) obtained increased navel

fruit set initially in California, but further attempts

resulted in severe leaf drop. Similar attempts in Florida

have failed to increase navel yield (68). Fruit set and

yield of 'Orlando' tangelo and other tangerine hybrids,









however, usually are increased by whole-tree sprays of GA

during bloom (21,69). More recently, spray applications of

GA alone or in combination with Ca(H2PO4)2 and/or 6-benzyl-

adenine resulted in increased fruit set of navel orange in

Florida (110). Yield data, however, were not obtained for

these experiments. No material is presently recommended to

improve fruit set of navel orange in Florida.

Auxin sprays can be used to control preharvest fruit

drop of citrus in Florida (5,39) and California (31,62,113).

Auxin action may be indirect through the control of fruit-

stem dieback and water spot of navel orange (31,62), or

direct by inhibition or delay of mature fruit abscission (5,

39,84).

Little information is available on the effects of

growth regulator applications on summer and summer-fall

fruit drop of navel orange in Florida. Sprays of 2,4-D at

bloom or a few weeks later appeared to prevent summer drop

(78), but information on summer-fall drop and yield was

incomplete.






Materials and Methods




Plant Material


Fruit drop was studied from 1979 to 1982 in 2 navel

orange (Citrus sinensis (L.) Osbeck)groves located near








Eustis in the north central citrus region of Florida. The

rootstock in both groves was sour orange (Citrus aurantium

L.). Grove A, planted in 1957 was used in 1979 and 1980 but

later abandoned because of severe freeze injury. Grove B,

planted in 1969, and unhurt by the 1981 and 1982 freezes,

was chosen for studies in 1981 and 1982.




Experiments 1 and 2, 1980



Two experiments to control fruit drop were carried out

in 1980. Each experiment consisted of 5 to 6 spray

treatments with 5 single-tree replications arranged in a

randomized complete block design. Treatments in experiment 1

were 20 ppm gibberellic acid (GA), 20 ppm 2,4-dichloro-

phenoxyacetic acid (2,4-D), 20 ppm 2,4-D + 20 ppm GA,

1000 ppm Promalin (Ciba-Geigy, 1.8% GA4+7 plus 1.81 N-phenyl-

methyl-l-H-purin-6-amine), and an unsprayed control. All

treatments were applied on May 27, at 8 weeks after midbloom.

Those of experiment 2 were 20 ppm 2,4-D at midbloom + 11

weeks (June 17), 20 ppm GA or 20 ppm 2,4-D at midbloom + 15

weeks (July 17), 20 ppm GA or 20 ppm 2,4-D at midbloom + 19

weeks (August 14), and an unsprayed control. Treatments were

applied to run-off as sprays of 75 liters of solution per

tree with a hand-gun sprayer. Midbloom occurred from March

25 to 31.









Experiment 3, 1981



Ten spray treatments for fruit drop control were

evaluated in 1981 at grove B, using 4 single-tree

replications in a randomized complete block design.

Treatments consisted of 20 ppm GA, 10 or 20 ppm 2,4-D, and

10 or 20 ppm 2,4-D + 20 ppm GA applied at 3 dates, March 20

(midbloom), May 1 (midbloom + 5 weeks), and June 26

(midbloom + 13 weeks), and an unsprayed control. Treatments

were applied to run-off as sprays of 75 liters of solution

per tree with a hand-gun sprayer. Midbloom occurred from

March 20 to 26.


Experiment 4, 1982



The 1982 experiment at grove B consisted of 5

treatments using 5 to 10 single-tree replications in a

completely randomized design, trunk girdling at early bloom

(February 19) or midbloom + 9 weeks (May 6), 20 ppm 2,4-D at

midbloom + 6 weeks (April 19) or midbloom + 9 weeks (May 6),

and an unsprayed, ungirdled control.Spray treatments were

applied to run-off at 55 liters of solution per tree with a

hand-gun sprayer. Trunk girdling consisted of a single

knife-cut through the bark around the trunk at 10 to 20 cm

above the bud union. Midbloom occurred from March 1 to 5.








Fruit Drop and Yield Evaluation, 1980 to 1982



Fruit drop was evaluated through catch-frame counts

from bloom until the end of the fruit-set period in mid-

June in 1981 (experiment 3), and,from mid-June until

harvesting, by whole-tree counts in 1980 to 1982

(experiments 1 to 4). The catch-frame method consisted of

placing a Saran cloth frame (2 x 1 m) tangentially to the

canopy drip line in a similar location underneath each tree.

Flower buds, flowers and fruit collected in the frames were

counted and removed at 7- to 14-day intervals. The canopy

projection area was estimated from diameter measurements.

Total drop of reproductive structures per tree was

calculated from the drop counts in the 2 m catch-frames.

Whole-tree fruit drop counts were made by counting fruit

underneath each tree at 7- to 14-day intervals. An

examination to determine cause of fruit drop was carried-out

at this time.

Yield was determined in 1981 and 1982 by harvesting

all remaining fruit on the trees shortly after commercial

maturity in late November to early December. Number of

boxes (40.8 kg each) per tree was obtained. Number of fruit

per tree was calculated in 1981 from the number of fruit in

each box, which in turn was estimated from fruit-diameter

measurements made on 40 fruit per treatment (117).









Fruit External and Internal Characteristics, 1981 and 1982



The effects of spray treatments on fruit external and

internal characteristics were evaluated in 1981 and 1982.

Random samples of 10 fruit per tree were used for

determination of external color, rind puncture force, juice

content, total soluble solids (TSS), total titratablee) acid

(TA), weight, equatorial diameter, stylar-end aperture

diameter, peel thickness and secondary-fruit diameter.

External color was determined in a Hunter color

difference meter using 2 readings per fruit taken at the

fruit equator through a 5 cm-diameter viewing window.

Lightness (L), redness (A) and yellowness (B) readings were

recorded. The instrument was standardized using a white

enamel plaque with values L = 94.0, A = -1.2, and B = 2.2.

Fruit color was expressed as a ratio between the values in

the green-to-red (A) and blue-to-yellow (B) scales.

Fruit rind puncture force was determined using a

Chatillon DPP-30 strain gauge equipped with a cylindrical

plunger beveled to a 1 x 6 mm rectangular surface at the

tip. Two measurements per fruit were made at the fruit

equator.

Juice extraction and analyses were performed as

recommended by Wardowski et al. (121). Juice content was

expressed as % by weight. Total soluble solids (TSS) were

determined using a Brix hydrometer calibrated to read

directly in % sucrose. Total titratablee) acid (TA) was




University of Florida Home Page
© 2004 - 2010 University of Florida George A. Smathers Libraries.
All rights reserved.

Acceptable Use, Copyright, and Disclaimer Statement
Last updated October 10, 2010 - - mvs