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Doctoral Dissertations 1896 - February 2014
1-1-1983
Location and structure of the pheromone gland, and physiological control of sex pheromone release in the female gypsy moth, Lymantria dispar (L.).
Andrew Lee Hollander University of Massachusetts Amherst
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Recommended Citation Hollander, Andrew Lee, "Location and structure of the pheromone gland, and physiological control of sex pheromone release in the female gypsy moth, Lymantria dispar (L.)." (1983). Doctoral Dissertations 1896 - February 2014. 5624. https://scholarworks.umass.edu/dissertations_1/5624
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LOCATION AND STRUCTURE OF THE PHEROMONE GLAND, AND
PHYSIOLOGICAL CONTROL OF SEX PHEROMONE RELEASE IN THE
FEMALE GYPSY MOTH, Lymantria dispar (L.).
A Dissertation Presented
By
ANDREW LEE HOLLANDER
Submitted to the Graduate School of the University of Massachusetts in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
September 1983
Entomology Andrew Lee Hollander
All Rights Reserved
/ LOCATION AND STRUCTURE OF THE PHEROMONE GLAND, AND
PHYSIOLOGICAL CONTROL OF SEX PHEROMONE RELEASE IN THE
FEMALE GYPSY MOTH, Lymantria dispar (L.).
A Dissertation Presented
By
ANDREW LEE HOLLANDER
Approved as to style and content by:
ill DEDICATION
To my brothers; Ted, Paul and Jim. Without their example, intellectual stimulation and moral support, I would not be where I am today.
iv ACKNOWLEDGEMENTS
I wish to express my appreciation to Dr. Chih-Ming Yin for
his work, guidance and constructive criticisms as major advisor
for this work. I also wish to thank Drs. John G. Stoffolano and
Gordon Wyse for their time and effort as graduate committee
members and instructors.
Dr. Charles Schwalbe of the Otis Gypsy Moth Methods
Development Center provided the experimental animals for the
experiments carried out in this work. Financial support was
provided by the Massachusetts Agricultural Experiment Station.
I am grateful to Dr. Michael Ma of University of Maryland,
and Dr. Ring Card£ and Ralph Charlton of University of
Massachusetts for their help and use of equipment in the
experiments using the aerial pheromone collection techniques.
For sharing a laboratory and their assistance, I wish to
thank Gaylen Jones, Zong-Shun Wang, Wei-Di Chaw, and Wu-Kang
Peng .
A debt of gratitude goes to the rest of the faculty, staff and graduate students of the Department of Entomology, and other
friends at UMASS and in the Amherst community for making this a most worthwhile learning experience.
v ABSTRACT
LOCATION AND STRUCTURE OF THE PHEROMONE GLAND, AND
PHYSIOLOGICAL CONTROL OF SEX PHEROMONE RELEASE IN THE
FEMALE GYPSY MOTH, Lymantria dispar (L.).
September 1983
Andrew L. Hollander, B.S., Iowa State University,
B.A., Iowa State University, M.S., Oklahoma State University,
Ph.D., University of Massachusetts
Directed by: Professor Chih-Ming Yin
The physiological control of calling and pheromone release in the female gypsy moth was investigated. The structure of the pheromone gland and associated regions were observed using histological preparations, scanning electron microscopy, and
Nomarski interference microscopy. The pheromone glands appear to
be located in two discrete patches: one on the dorsal and one on
the ventral surface of the inter segmental membrane between the
eighth and ninth (terminal) abdominal segments.
Micro-surgical techniques were used to determine whether the
endocrine system was involved in pheromone release and calling
behavior. No humoral controls of pheromone release or calling
behavior were demonstrated from the brain, corpora allata,
corpora cardiaca or ovaries.
Nervous control was demonstrated for both calling behavior
vi and pheromone release. Severing the ventral nerve cord at any
level anterior to the pheromone glands up to, and including, the circumesophageal connectives, stopped sex pheromone release.
These data strongly indicate that control of pheromone release resides in the brain. The immediate controlling mechanism over calling behavior, however, appears to be a motor pattern generator associated with the terminal abdominal ganglion.
Quantitative verification of differences in pheromone release due to surgical manipulation was attempted. The ventral nerve cords of the gypsy moths were severed and aerial pheromone collections were made. These collections were compared with collections from control and sham operated animals. Two techniques to collect and assay released pheromone were evaluated.
Attempts were made to elicit pheromone release by stimulating the nerves that run from the terminal abdominal ganglion to the region of the pheromone glands. No evidence for an increase of pheromone release due to nerve stimulation was shown.
vii TABLE OF CONTENTS
TITLE PAGE. .7 COPYRIGHT PAGE.. 11 APPROVAL PAGE.111 DEDICATION. 1V ACKNOWLEDGEMENTS . ABSTRACT. .V1 TABLE OF CONTENTS.viil LIST OF TABLES . LIST OF ILLUSTRATIONS.xii
Chapter I. INTRODUCTION . 1
II. SEX PHEROMONE GLAND LOCATION: BIOASSAY, MORPHOLOGY, AND HISTOLOGY . 7
Introduction . 7 Methods and Materials . 11 Insects. 11 Bioassay for female sex pheromone . 12 Location of the sex pheromone- producing gland. 13 Scanning electron microscopy . 14 Histology of the inter segmental membrane. 14 Interference Microscopy . 14 Results. 15 Localization of the female sex pheromone gland . 15 Microscopic examination of the sex pheromone gland . 16 Discussion. 28
III. HUMORAL CONTROL. 33
Introduction . 33 Methods and Materials . 35 Experimental animals . 35 Operations. 36 Allatectomies . 37 Corpora cardiaca - corpora allata removal. 37 Ovariectomies . 38 Bioassays. 38 Results. 39
viii Discussion . 44 Allatectomies . Corpora cardiaca - corpora allata removal. 45 Ovariectomies . 46
IV. NERVOUS CONTROL. 50
Introduction . 50 Methods and Materials . 54 Experimental animals . 54 Operations. 55 Operations on ventral nerve cord . 55 Operations on the brain. 55 Bioassay. 55 Aerial collection of pheromone . 56 Ma-Schnee technique . 56 Char1ton-Card& technique . 57 Stimulation of nerves from the terminal abdominal ganglion . 58 Results. 59 Anatomical relationship of the nervous system to the pheromone gland. 59 Experimental effects . 60 Discussion. 76
V. SUMMARY AND CONCLUSION. 82
LITERATURE CITED . 86
APPENDIX. 96
ix LIST OF TABLES
1. Gypsy moth wing fanning bioassay for female sex pheromone. All treatments with the exceptions of blank controls and calling females were tissue extracts from virgin females from 24 to 48 hours post adult eclosion. 17
2. Effect of removal of the corpora allata (CA) from the last larval instar on calling behavior and pheromone release by adult Lymantria dispar females. 41
3. Effect of removal of the corpora cardiaca and corpora allata (CC-CA) from the last larval instar on calling behavior and pheromone relesase by adult Lymantria dispar females ... 42
4. Effect of ovariectomy in the last larval instar on calling behavior and pheromone release by adult female Lymantria dispar . 43
5. Effect of operations conducted on the pupal nervous system with respect to calling behavior and pheromone release by adult female Lymantria dispar . 63
6. Effect of severing the circumesophageal connectives (CEC) in the pupal stage on calling behavior and pheromone release by adult female Lymantria dispar . 64
7. Effect of stimulation of the fifth and sixth major nerves of the terminal abdominal ganglion on calling behavior and pheromone release by adult female Lymantria dispar . 75
8. Standard curve data for disparlure. Generated with an Envirochem Model 780B gas chromatograph . 97
9. Effect of stimulation of the fifth and sixth major nerves of the terminal abdominal ganglion on calling behavior of adult female Lymantria dispar. 98
10. Number of times males were exposed (n) to female Lymantria dispar preparations, and the males
x exhibi ting the wing fanning response to the prepar ations before and duri ng electrical stimul ation of the female’s fifth and sixth major terminal abdominal gan glion nerves . 99
xi LIST OF ILLUSTRATIONS
1. Extended position of the female gypsy moth’s eighth and ninth abdominal segments bordering the intersegmental membrane in typical calling behavior. 10
2. Retracted position of the female gypsy moth’s terminal abdominal segments in typical calling behavior. 10
3. Pheromone bioassay apparatus . 19
4. Scanning electron micrograph with a dorsal view of a female gypsy moth's terminal abdominal segments. 19
5. Scanning electron micrograph of the lateral aspect of a female gypsy moth’s terminal abdominal segments . 22
6. Scanning electron micrograph of the ventral aspect of a female gypsy moth’s terminal abdominal segments . 22
7. Longitudinal section through dorsal convolutions of a virgin female showing columnar cells (CO), cuboidal cells (CU), cuticular domes (CD), circular-oblique muscles (CM), and dorsal longitudinal muscles (DLM) . 25
8. Longitudinal section through the ventral convolutions of a virgin female showing columnar cells (CO), cuboidal cells (CU), cuticular domes (CD), circular-oblique muscles (CM), and ventral longitudinal muscles (VLM) .... 25
9. Cross section through dorsal convolutions of a virgin female showing the circular-oblique muscles (CM) and dark granules (DG). 27
10. Cross section through the ventral convolutions of a virgin female showing direct insertion of circular-oblique muscles to the cuticle .... 27
11. Diagramatic dorsal view of the gypsy moth
xii nervous system 53
12. Diagramatic ventral view of the gypsy moth terminal abdominal ganglion, ventral nerve cord and six major nerves . 62
13. Standard dilution curve for concentrations of disparlure ( cis-7,8- epoxy -2- methyl- octadecane), assayed using an Envirochem Model 780B gas chromatograph, with a flame ionization detector and a capillary column . 68
14. Representative chromatogram from the Envirochem Model 780B gas chromatograph. The sample was a 24 hour aerial collection from an adult control female . 70
15. Representative chromatogram from the Varian 3700 gas chromatograph. The sample was from a 12 hour aerial collection from a control with no female. The internal standard was cis-9,10-epoxy-eicosane . 72
16. Representative chromatogram from the Varian 3700 gas chromatograph. The sample was from a 12 hour aerial collection from a control with no female. 40 ng of synthetic disparlure and 40 ng of an internal standard, cis-9, 10-epoxy-eicosane . were added to the sample . 72
17. Representitive chromatogram from the Varian 3700 gas chromatograph. The sample was from a 12 hour aerial collection from a stock colony female. 40 ng of cis-9, 10-epoxy- eicosane were in the sample as an internal standard . „ 71
• • • Xlll CHAPTER I
INTRODUCTION
During the past two decades we have seen a revolution in the way people view insect pest control. Social and political demands for environmental quality, compounded by increasing resistance of insects to classical chemical pesticides have necessitated investigations into other areas that may yield possible alternatives to outdated control strategies. Many areas of a pest insect's biology must be studied to aid development of new techniques and compounds for pest control. Further, new techniques and compounds must not only maximize restraints on the pest populations in question, but must also minimize damage to man and his environment. Hopefully, these insights will lead to effective integrated pest management systems.
One area that has stirred considerable interest and research is that of insect pheromones. These compounds are produced in small quantities by the insects. Pheromones serve to influence or change the behavior of other individuals of the same species.
These chemicals are usually of low toxicity (Beroza et al. 1975), need only be used in minute quantities to elicit the expected response and are highly specific for the target organism.
There are a number of different kinds of pheromones, including trail pheromones, alarm pheromones, aggregation
pheromones, epideictic pheromones, and maturation pheromones. One
class of pheromones that has been studied extensively is the sex 2
attractants. These are chemicals emitted by one sex of the
species to elicit behaviors from the other sex to enhance the
chances of mating. Since von Siebold (1837) first reported that
male moths were attracted to female moths by an airborne substance
these materials have been thought to have potential in insect
control. The rationale for using sex pheromones for pest control
is that by successfully disrupting mate-seeking behavior with
synthetic pheromone, mating can be prevented. Reducing
successful matings in a population enough will lead to a
decreased population size in the next generation. The goal of
such an undertaking would be to reduce and keep the pest
population below an economic threshold.
The existence of sex pheromones has been well documented in
Lepidoptera. Because many species in this order are of economic
importance, considerable research has been devoted to the characterization of these pheromones (Brand et al . 1979), their behavioral effects (Bartell 1977) and possible iraplementions in control schemes (Roelofs 1979).
Past control efforts utilizing pheromones have highlighted the great need for more basic research into the behavior and physiology of the producers of these materials. Some areas of investigation include: the biochemistry of production (Law and
Regnier 1971, Howard and Blomquist 1982), quantification of pheromone release (Shorey and Gaston 1965, Sower et al . 1971b,
Caro et al. 1978, Charlton and Card£ 1982, Ma and Schnee 1983), 3
and environmental influences on calling behavior and pheromone release (Sower et al. 1971a, Shorey 1974, Sower and Fish 1975,
Webster and Card* 1982). Electrophysiology, in the form of electroantennograms (Priesner 1975, Miller et al. 1977, Schneider et al. 1977) and neuroanatomy of the male antennal lobes
(Schweitzer et al. 1976) has also become an area of interest.
Only three publications to date have addressed the endocrinological aspects of pheromone release by female
Lepidoptera, (Roller et al. 1963, Steinbrecht 1964, Riddiford
and Williams 1971). No previous work has dealt directly with the
relationship of the moth’s nervous system to pheromone release.
Knowledge of the relationships among endocrinology, neurobiology
and pheromone physiology is greatly lacking. Basic research in
this area is important for identifying possible weak links that
may be exploited in an integrated pest management system.
My main objectives for this work were in two areas of
physiology dealing with sex pheromone release. The first
objective was to ascertain the role of humoral and hormonal
factors on pheromone release and associated behaviors. Hormone
producing organs examined were the brain, corpora cardiaca,
corpora allata, and the ovaries. The second objective was to
establish the role of nervous control and location of the
controlling centers involved with calling behavior and pheromone
release.
The gypsy moth, Lymantria dispar (L.) (Lepidoptera:
Lymantriidae), was chosen as the model with which to study the 4
control systems of pheromone release because it is 1) an important pest species, 2) readily available, 3) easy to manipulate in the laboratory, and 4) the adult’s behavior is relatively uncomplicated.
The North American gypsy moth originated from Europe where it is a relatively innocuous insect. The species was first introduced to the United States by the French naturalist, Leopold
Trouvelot to develop a high yield silk producing moth (Forbush and Fernald 1896). The gypsy moth’s accidental release in
Medford, Massachusetts, around 1869 has lead to the increase and spread of one of the northeast’s most conspicuous forest, shade tree and ornamental pests (Doane and McMannus 1981). Due to the very conspicuous nature of the damage the gypsy moth inflicts, a great deal of research and monetary support has been targeted at various aspects of the biology of this insect, and its control. A current and comprehensive compendium on the results of previous research in this area is provided by Doane and McMannus (1981).
There is a precedent for using the gypsy moth in physiological studies. Koped (1922), in his classic head ligation experiment, used the gypsy moth to show that the head was important in the production of a humoral factor for retarding premature pupal development. This was the initial study that lead to the discovery and characterization of insect juvenile hormone. Unfortunately, no further physiological work has been done with the gypsy moth.
The search for the active compounds in the gypsy moth’s sex 5
pheromone started with the report of a chemical named "gyptol" ,
(Acree 1953). Gyptol was tentatively identified as the sex
pheromone, and was chemically characterized as (+) -10- acetoxy
-1- hydroxy - cis -7- hexadecene by Jacobson et al. (I 960).
The lack of significant behavioral activity of gyptol and
gyplure, a synthetic homologue, lead to a reinvestigation. As a
result, gyptol and gyplure are considered to be behaviorally
inactive with the gypsy moth (Jacobson et al. 1970). Ci_s
_7f8_ epoxy -2- methyloctadecane was proposed as the pheromone
structure by Bierl et al. (1970, 1972). The synthetic compound
with this structure was dubbed "disparlure" . Iwaki et al.
(1974) found that the ( + ) enantiomer of disparlure was by far
the most potent in eliciting male responses as compared with a
racemic mixture or the (-) enantiomer alone. Most moth sex
pheromones that have been studied have several components, as
many as four in a number of cases, but to date, cis -7»8-
epoxy -2- methyloctadecane is the only compound identified from
the naturally produced pheromone of the gypsy moth.
The sex pheromone of the gypsy moth is one of the most
studied for its use in population control. Collins and Potts
(1932) suggested that the male’s attraction to the female from
great distances might be used to trap out the males. For an
overview of the efforts and theories over the years on trapping
gypsy moth males, refer to Cameron (1981).
Studies of the exact nature and location of the gland(s) 6
that produce and release the sex pheromone are lacking. To fill
this void in our knowledge, research was conducted to elucidate the structure and location of the pheromone glands.
The three areas studied were: the location, morphology and histology of the sex pheromone gland, the role of humoral
controls in pheromone release and the role of the nervous system
in controlling pheromone release. The following chapters detail
the experiments, results, conclusions and questions resulting
from these investigations. CHAPTER II
SEX PHEROMONE GLAND LOCATION: BIOASSAY, MORPHOLOGY AND HISTOLOGY
Introduction
A considerable amount of literature exists on the structure of scent glands of Lepidoptera. Urbahn (1913) first described the
scent organs of a number of moths and butterflies. More recently,
Jacobson (1972) and Percy and Weatherston (1974) have published
comprehensive reviews. Other recent studies cover species
including Trichoplusia ni (Htlbner) (Percy 1 970 ), Plutella
xylostella L. (Su and Lee 1977) and Argysataenia velutinana
(Walker) (Feng and Roelofs 1977).
Research on gypsy moth sex attractants has been focused
primarily on pheromone identification, behavior, population
monitoring, periodicity of pheromone release, and improvement of
trap efficacy. Disparlure, ci s -7,8- epoxy-2-methyloctadecane ,
was isolated from female gypsy moths and synthesized by Bierl et
al. (1970). In field tests, traps baited with 5 ng of disparlure
effectively attracted males (Beroza et al. 1971). These findings
have stimulated much work in gypsy moth pheromone-related studies
over the past decade.
Despite the intensive study of the gypsy moth pheromone, the
exact location and structure of the pheromone-producing gland(s)
remain unknown. It has been shown that the female gypsy moth 8
produces a potent sex attractant from the apical portion of the abdomen (Forbush and Fernald 1896, Collins and Potts 1932). Gypsy moth male attraction is associated with an overt female act, referred to as calling behavior. Calling behavior is characterized by the rhythmic extention and retraction of the eighth and ninth (terminal) abdominal segments (Figs. 1 and 2).
In a thorough study of pheromone release, it is necessary to understand the nature of the gypsy moth pheromone gland . Such information could provide some insight into the control mechanisms and functions of the gland(s).
A closely related species of the gypsy moth, the white-marked tussock moth (Orgyia leucostigma J. E. Smith) has been the subject of morphological and ultrastructural studies involving the pheromone gland (Percy et al. 1971, Percy 1975).
The tussock moth pheromone gland is a crescent-shaped area on the dorsal inter segmental membrane between the eighth and ninth abdominal segment. The glandular epidermal cells are described as being hypertrophied and included cuboidal, columnar and goblet-shaped cells.
The locations of the gypsy moth’s pheromone glands were determined with a bioassay technique performed with portions of the terminal abdominal segments. The bioassay was designed to register male preflight wingbeat excitement caused by extracts from different terminal abdominal segment preparations. The number of activated male gypsy moths was compared to the number excited by calling female, solvent and blank controls. Wing
9
th' s Fig. 1. Extended position of the female gypsy mo the eighth (E) and ninth (N) abdominal segments bordering r . inter segmental membrane (I) in typical calling behavio Abdominal hair has been removed for clarity. (15X)
gypsy moth's Fig . 2. Retracted position of the female ninth (terminal) abdominal segments in typical calling eighth and clarity. (15X) behavior. Abdominal hair has been removed for 10 11
vibration is an overt event in the activation of resting males by the sex pheromone. Bartell (1977) indicated that wing fanning may serve to raise the temperature of flight muscles in preparation for flight. Wing fannig was often preceded by elevation of the antennae and walking. However, exhibition of these lesser degrees of excitement alone was considered negative in scoring the assays.
The morphology and histology of the pheromone glands and associated structures were examined with both optical microscopy and scanning electron microscopy. This work forms the foundation for following studies on the regulatory systems important in the control of pheromone release and calling behavior.
Materials and Methods
Insects. A weekly supply of third instar gypsy moth
caterpillars was obtained through the mail from the Otis Gypsy
Moth Methods Development Center. Larvae were housed from the
first instar to pupation in XE-6 (Sweetheart Plastics) cups in
groups of 12 per cup. Larval cups initially contained 75-90 ml
of a standard wheat germ diet (Bell et al. 1981). Upon arrival,
the larvae were kept in a Percival growth chamber at 26 + 2°
C at approximately 40% relative humidity and 16 hour light : 8
hour dark photoperiod and reared to pupation. Pupae were
collected and sexed daily and transfered to 16 oz unwaxed paper
cups (Sweetheart Plastics) with clear plastic lids until adult 12
emergence. Under these conditions, larval and pupal stages lasted 14-21 days after arrival and 9-12 days thereafter, respectively.
Bioassay for female sex pheromone. Adult male wing fanning behavior upon exposure to female sex pheromone (Card& and
Hagamen 1979) was the basis for the present assay. The system contained a vacuum source, a rotometer and a glass assay tube (5.8 cm diameter by 54 cm long) (Fig. 3). Male gypsy moths were held in the middle section of the assay tube by two brass screens placed approximately 10 cm apart. Female gypsy moths or samples for bioassay were placed in the open end of the tube on clean
Whatman No. 1 filter paper. The other end of the tube was closed by a No. 12 rubber stopper with an air collecting funnel inserted in the center hole. The funnel stem was connected to the rotometer and vacuum pump with Tygon tubing. The vacuum source was adjusted to generate a constant air flow of 8 cm/sec in the assay tube. For each set of bioassays, two unmated males were placed in the assay tube and allowed to acclimate for at least 10 min. Before each test, a three minute control test was run with blank filter paper at the specified air flow. Only males not responding to the blank were used for testing biological materials. Test materials were considered biologically active if they aroused male preflight wingbeat behavior within three minutes at an 8 cm/sec air flow rate. Materials that failed to 13
elicit wingbeat activity were considered inactive. The sample chamber was cleaned thoroughly with soap and water followed by acetone after each set of bioassays to avoid contamination from one test to another. To reduce the potential of habituation to the pheromone, males were subjected to no more than 10 bioassay trials before being discarded. All bioassays were conducted between 6.5 to 9 hour after the onset of photophase to reduce po ssible discrepancies stemming from the diel pheromone release rhythm (Card& et al. 1974).
Location of the sex pheromone-producing gland. To locate the sex pheromone gland, pieces of tissue or hexane extract from various parts of the terminal abdominal segments were bioassayed for pheromone activity. Assayed portions included: entire terminal abdominal segments (eighth abdominal segment, ninth abdominal segment and their inter segmental membrane) , dorsal and ventral halves of these segments, isolated eighth abdominal segments, ninth abdominal segments, and isolated inter segmental membranes from between the eighth and ninth abdominal segments.
Each bioassay sample contained extracts of tissue portions from three different individuals. These samples were pooled in 1/4 ml of chromatography-grade hexane and soaked for 10 - 20 min. At the completion of the soak, a strip of Whatman No. 1 filter paper (1 x 3 cm) was placed in the hexane. After the filter paper had absorbed the hexane solution, the hexane was allowed to evaporate completely before the filter paper was placed in the bioassay 14
tube. Only virgin females between 24 and 48 hour post-emergence were used for this study. Bioassays were accompanied by blank, solvent, and calling female controls.
Scanning electron microscopy. Ten terminal abdominal segments
(segments eight and nine) from virgin female moths were fixed in their extended position (Fig. 1) and prepared as described by
Sabatini et al. (1963) for scanning electron microscopic studies.
After critical-point dehydration in liquid CC>2 (Anderson
1951), the terminal segments were mounted on metal stubs with conductive cement, coated with gold—palladium at 15 mA for 3 min
(approximately 495 8) in a vacuum evaporator, and examined
with an ISI Super III-A scanning electron micoscope using a tilt
angle of 40° and an accelerating voltage of 30 kV. Micrographs
were recorded on Type 55 Polaroid positive/negative film.
Histology of the inter segmental membrane. Extended apical
portions of the female abdomens were fixed and prepared for
histological studies as in Yin and Chippendale (1973). Serial
sections (8 urn) were stained with Delafield’s hematoxylin or
Mallory-Heidenhain’s one-step triple stain (Humason 1967).
Photomicrographs were taken using a Wild M-20 microscope equipped
with a Pentax MX camera system.
Interference microscopy. To supplement the scanning electron 15
microscopic examination of surface morphology, whole-mount preparations of the female terminal abdominal segments were observed using a Zeiss compound microscope equipped with interference optics (Nomarski prisms). Specimens were cleared in
10% KOH, flattened and mounted on a glass slide for viewing.
The interference microscopy revealed the surface contour of the specimen.
Localization of female sex pheromone gland. Initially, tests were made to reconfirm the pheromone gland’s location on the abdominal tip (Collins and Potts 1932). This was done by running male wing fanning bioassays on different preparations including the isolated terminal abdominal segments from calling females, females with the terminal segments removed, and unoperated calling females serving as controls. Only preparations that included the female gypsy moth’s eighth and ninth (terminal) abdominal segments elicited positive responces from the males in the bioassay.
Two series of experiments were run to more precisely indicate the location of the pheromone gland. In the first series, the terminal segments were carefully dissected into three parts: the eighth segment, the ninth segment, and the inter segmental membrane between the eighth and ninth segments. 16
Bioassays were performed on these preparations along with blank tubes and calling females as controls. The results show that the intersegmental membrane between the eighth and ninth segment was the portion that elicited male wingbeat response (Table 1).
The second series of experiments were designed to indicate whether the sex pheromone gland was on the dorsal or ventral side of the intersegmental membrane. Bioassays were performed on hexane extracts of dorsal and ventral halves of terminal abdominal segments from calling females. Pure hexane and hexane extract of the intact terminal abdominal segments served as controls. The bioassay results indicate that the dorsal and ventral halves are about equally potent in their ability to stimulate male wingbeat response (Table 1).
Microscopic examination of the sex pheromone gland. With the above information, I turned to morphology and histology for supporting evidence to help establish the inter segmental membrane as the location of the sex pheromone gland. External morphology of the terminal abdominal segments was observed using scanning electron microscopy. The dorsal view of the inter segmental membrane shows a well-defined highly convoluted integumentary area
(Fig. 4). Since the lengthy process of preparing specimens for electron microscopy may occasionally introduce structural artifacts, the existence of the dorsal convoluted area was confirmed with another method, i.e. interference microscopy.
The scanning electron micrograph shows that the lateral 17
TABLE 1. Gypsy moth wing fanning bioassay for female sex pheromone. All treatments, with the exceptions of blank controls and calling females, were tissue extracts from virgin females from 24 to 48 hours post adult eclosion.
Wing fanning response
Treatment n % positive
Blank control 124 0.0
Solvent control 24 0.0
Calling female 35 88.6
8th abdominal segment 20 0.0
8th-9th intersegmental membrane 20 60.0
9th abdominal segment 20 10.0
Entire terminal abdominal segments 10 90.0
Dorsal halves of terminal segments 21 61.9
Ventral halves of terminal segments 20 75.0 18
Fig. 3. Pheromone bioassay apparatus showing the incurrent end of the bioassay tube (T), the rotometer (R), and the vacuum pump (P) .
Fig. 4. Scanning electron micrograph with a dorsal view of a female gypsy moth’s eighth (E) and ninth (N) abdominal segments. This micrograph shows the highly convoluted nature of the inter segmental membrane (I) where glandular epithelium is located. The right part of the figure is an enlargement of the rectangle in the left half, showing the cuticular convolutions. (Left 90X, Right 270X, Scale bar = 0.067 mm). 19 20
aspects of the intersegmental membrane between the eighth and
ninth abdominal segments (Fig. 5) are relatively smooth and
featureless. The ventral surface (Fig. 6), however, is
convoluted similarly to that of the dorsal surface. Interference
microscopy also confirmed the existence of the ventral
convolutions.
Serial paraffin sections of the terminal abdominal segments
revealed some of the histological characteristics of the
intersegmental membrane. The cells in the longitudinal sections
of the dorsal convolutions are enlarged (Fig. 7). The dorsal
cells range from 0.01 to 0.015 mm wide by 0.025 to 0.06 mm thick.
Microscopic examination indicates these cells are columnar in
shape, although some cuboidal and goblet shaped cells are also
present. The columnar cells contain a large nucleus with one distinct nucleolus although several nuclei seem to have more than one nucleoli. The nuclei are usually located in the apical 2/5 to the center of these columnar cells. Large numbers of dark granules are often found at the apical and/or basal part of the cells (Fig. 9).
Observed epidermal cells of the inter segmental region underlying the smooth cuticle were generally cuboidal or squamous. These cells ranged in size from 0.008 to 0.015 mm in length by 0.00« to 0.006 mm in thickness.
The cuticle of the lateral intersegmental membrane and undifferentiated dorsal and ventral surfaces was usually thinner 21
Fig. 5. Scanning electron micrograph of the lateral aspect of a female gypsy moth’s eighth (E) and ninth (N) abdominal segments. Note the lack of convolutions on the intersegmental membrane (I). The right half of the figure is an enlargement of the rectangle in the left half of the picture. (Left 75X, Right 225X, Scale bar = 0.067 mm).
Fig. 6. Scanning electron micrograph of the ventral aspect of a female gypsy moth’s terminal abdominal segments. The convoluted ventral portion of the in ter segmental membrane is also underlaid by glandular epidermis. The right half of the figure is an enlargement of the rectangle in the left half of the picture showing the ventral inter segmental convolutions (VC) and part of the ninth abdominal segment (N). (Left 70X, Right 210X, Scale bar = 0.07 mm) . 22 23
(approximately 0.008 mm) than the cuticle of the convoluted regions (approximately 0.012 mm). The lateral epidermal cells were not enlarged and were predominantly squamous in shape with some cuboidal cells present.
The histological features of the ventral convolutions resemble, in many respects, those of their dorsal counterparts
(Fig. 8). However, in many sections the enlarged ventral epidermal cells are more cuboidal than columnar in shape and are somewhat smaller than the dorsal cells. The ventral epidermal cells in the convolutions measure 0.01 to 0.015 mm wide by 0.015 to 0.05 mm long. The position of the nucleus and presence of dark granules in the cytoplasm of the ventral epithelium were
similar to those of the dorsal epidermal cells.
Longitudinal sections show that there are two layers of muscles in the region of the pheromone glands. A layer of circular or oblique (circular-oblique) muscles is located between the epidermal cells and the massive layer of longitudinal muscles
(Figs. 7 and 8). The intrasegraental circular-oblique muscles make direct insertions to the endocuticle overlying the pheromone
gland cells (Figs. 9 and 10). By contrast, the longitudinal muscles attach themselves from the anterior margin of the eighth
to the anterior margin of the ninth abdominal segment and are
inter segmental in nature.
Preliminary transmission electron microscopic studies have
revealed that the cuticle of the dorsal convolutions consists of
a thin epicuticle and a thick lamellate endocuticle. The 24
Fig. 7. Longitudinal section through dorsal convolutions of a virgin female showing columnar cells (CO), cuboidal cells (CU), cuticular domes (CD), circular-oblique muscles (CM), and dorsal longitudinal muscles (DLM). In the extended state, the columnar cells are closely packed, but not compressed longitudinally. (300X, Scale bar = 0.1 mm) .
Fig . 8. Longi tudinal sec tion through the ventral con volut ions of a v irgin femal e showing columnar cells (CO), cub oidal cel Is (CU) , cuticular domes (CD), circular-oblique mus cles (CM) , and v entral long itudinal muscles (VLM). (300X,
Sea le bar = 0.1 mm) • 25 26
Fig. 9. Cross s ection through dorsal convolut ions o f a virgin female showing the circular-oblique muscles (CM) and dark granules (DG). When the convolutions are exposed, their epidermal cells are compressed by the contracting circular-oblique muse les. (250X, Scale bar = 0.1 mm) .
Fi g. 10. Cross section through the v entral convolutions of a virgin female showing direct insertion of circular-oblique muscles to the cuticle (arrows). (330X, Scale bar = 0.1 mm). 27 dome-shaped protrusions observed over the entire inter segmental
membrane surface contain a corresponding thickening of
endocuticle, but the thickness of the overlying epicuticle
remains uniform. No cuticular ducts or openings were observed.
The glandular columnar cells appear ductless and possess well
developed apical microvilli (Yin unpublished).
Discussion
The first set of pheromone bioassays on the abdominal tip
confirmed that the gypsy moth sex pheromone did come from the
terminal abdominal segments as noted by Collins and Potts (1932)
Results from pheromone bioassays of the eighth abdominal
segment, the ninth abdominal segment and the inter segmental membrane between the eighth and ninth segment suggest that the
female sex pheromone glands are located on the inter segmental membrane. Pheromone glands of female moths are commonly situated on the inter segmental membrane between the eighth and ninth
segments (Percy and Weatherston 1971). The types of glands, however, differ from species to species. They may take the form of a ring gland, dorsal sac, ventral sac, lateral sac or dorsal
fold (Percy and Weatherston 1971).
Further bioassays conducted on the dorsal and ventral halve of the terminal abdominal segments indicate that the attractant emanates from both the dorsal and ventral surfaces. From these 29
data it may be concluded that there are both dorsal and ventral components of the pheromone gland. This could mean that the gland is a ring gland as in Vi tula edmandsae (Packard)
(Weatherston and Percy 1968) or that it has discrete dorsal and ventral components on the inter segmental membrane between the eighth and ninth abdominal segments.
Scanning electron microscopy clearly shows two areas of highly convoluted inter segmental membrane cuticle. These regions are on the dorsal and ventral aspects and are separated by smooth lateral inter segmental membrane. Histological sections and interference microscopy of inter segmental membrane preparations confirm the locations of the convolutions. The cells underlying both the dorsal and ventral convolutions were enlarged columnar epidermal cells and appeared similar to pheromone gland cells described for a number of other species
(Jefferson et al. 1966, Jefferson et al. 1968, MacFarlane and
Earle 1970, Percy et al. 1971, Percy and Weatherston 1971, Percy
1975, Lew and Ball 1978). Therefore, the morphology is consistent with bioassay results which suggest comparable
potency of the dorsal and ventral halves of the terminal
abdominal segments.
The gypsy moth seems to have a slightly different pheromone
gland configuration than those listed above. In the case of the
gypsy moth both dorsal and ventral regions of the inter segmental
membrane appear to be modified for the production of pheromone.
Qrgyia leucostigma, a closely related species, like the gypsy 30
moth, does not evert the pheromone gland, rather it is exposed by
rhythmic telescoping movements of the terminal abdominal
segments (Percy and George 1979). However, the pheromone gland
of 0^ leucostigma consists of only the dorsal fold; no
ventral component has been described (Percy et al. 1971).
The putative glandular cells underlying the cuticular
convolutions appear to be ductless, but have well developed
apical microvilli. Under the system of classification given by
Noirot and Quennedy (197*0* the gypsy moth pheromone glands
should be called class-1 epidermal glands.
Due to the lipoidal nature of disparlure, no good techniques
could be found to directly identify the pheromone in histological
sections. Any of the standard histochemical techniques that
would be applicable to disparlure would have been nonspecific for
a great number of similar lipoidal compounds. This makes the
specific identification of disparlure by histochemical techniques unlikely. Because of the lack of direct evidence, the
information provided here is not conclusive as to the real nature of the pheromone gland. However, strong curcumstantial evidence is now available for use as a basis for further work in the identification of the pheromone glands.
Although the present study was not focused on the musculature governing the exposure of the pheromone gland, a brief discussion is needed for completeness. Movement of the terminal abdominal segments of female moths are associated with 31
muscules attached to paired anterior and posterior apophyses
(Matsuda 1976). Percy and George (1979) carried out a
comparative study to identify muscles associated with these
apophyses. They reported up to 9 pairs of dorsal and lateral
apophyseal muscles in Choristoneura fumiferana (Clem),
Tr ichoplus i a n^ (HUbner) and 1 e u c o s t i g m a . However
they did not mention any non-apophyseal muscles. The
observations presented in this study suggest that, in addition
to apophyseal muscles, coordinated contraction and relaxation of
longitudinal and circular-oblique muscles may also contribute to
the rhythmic protrusion and retraction of the terminal abdominal
segments. The arrangement of circular-oblique muscles
underlying the convolutions of the putative phermone glands of
the gypsy moth (Figs. 8 and 10) suggest that the contractions of
these muscles during calling may actually aid in pheromone
release, similar to squeezing a lemon peel.
Hemolymph pressure has been cited as being a possible
mechanism involved in calling movements (Jefferson and Rubin
1970). However, in studies to be discussed in Chapter IV, I have
noted that even when the abdomen of the gypsy moth female was
opened down to the terminal abdominal segments, the moth was
still able to extend and retract the terminal abdominal segments.
In addition, Percy and George (1979) showed the musculature and
mechanics in the terminal abdominal segments of the species
mentioned above are sufficient to function strictly with the
neuromuscular system. This does not, however, exclude hemolymph 32
pressure from havng a minor role in eversion of the terminal abdominal segments, and/or pheromone gland. CHAPTER III
HUMORAL CONTROL
Introduction
To identify the control mechanisms of calling behavior and
pheromone release in the female gypsy moth, previous work had dealt with the neurological aspects (Hollander and Yin 1982).
That work provided strong evidence for the importance of the nervous system in the control of female gypsy moth calling behavior and sex pheromone release. This study involves the
humoral control mechanisms that might be necessary for the
insect’s regulation of calling behavior and/or pheromone
release. Three endocrine related organs were examined for their
possible role(s) in the regulation of calling behavior and
pheromone release in the gypsy moth. These were the corpora
allata corpora cardiaca and ovaries.
The corpora allata were suspected of being a potential
producer or releaser of humoral agents that might control calling
behavior and/or pheromone release. Earlier findings have shown
that juvenile hormone from the corpora allata was involved in
other aspects of reproductive physiology, such as obgenesis,
and behavior in other insects (Raabe 1982). Lessman et al.
( 1 982 ), working with Danau s p1e xippus plexippus L., found a
juvenile hormone dose-dependent increase in ovary weight and
number of mature o&cytes. A number of examples of 34
reproductive behavior being influenced by the corpora allata are
discussed by Barth and Lester (1973).
Previous experiments with Galleria melonella (L.)
(ROller et al. 1963)* Bombyx mori L. (Steinbrecht 1964),
Antheraea polyphemus Cramer and Hyalophora cecropia (L.)
(Riddiford and Williams 1971) indicate that removal of corpora
allata from pupae does not affect pheromone release. The
studies above, however, did not take into account the
possibility that the pupal corpora allata had already released
humoral factors that would effect the pheromone release in the
adult stage. In the present experiments the gypsy moth corpora
allata were extirpated in the mid last larval instar, thus
allowing the larvae time to recover from the surgery before
pupation. Larval allatectomized individuals were freed from the
effect of any corpus allatum factor that may be released during
the late last larval, pupal and adult stages.
Riddiford and Williams (1971) also removed the corpora cardiaca and corpora allata together from pupal Hyalophora cercropia. They reported that the resulting adult moths showed reduced calling and pheromone release, although no method for quantitatively measuring pheromone release was employed.
Since they found no effect by removing the corpora allata, the corpora cardiaca would be a likely source organ for humoral factors controlling calling behavior and pheromone release in the gypsy moth. 35
Mating and reproduction are the major events in the life of an adult female gypsy moth. The ovaries, being central in this process, were logical candidates for exerting control over reproductive behaviors and physiological processes. These processes include the mechanisms involved in attraction of the male, ie. calling behavior and pheromone release. Ovarian influences on reproductive physiology and behavior have been demonstrated for other insects such as Periplaneta americana
L. (Bell 1969), Diploptera punctata (Eschaltz) (Stay and Tobe
1981) and Aedes aegypti (L.) (Rossignol et al . 1981).
Gypsy moth ovaries were removed during the last larval instar to show whether they had any effect on pheromone release or calling behavior. Because of the difficulty of closing the wound after the removal of the ovaries from soft-bodied larval lepidoptera, two alternative methods for closing the surgical wound were developed, i.e. a clamp technique and a cyanoacrylate ester glue technique. The effectiveness of these two wound closure techniques were evaluated.
Methods and Materials
Experimental animals. Second and third instar larval gypsy moths, Lymantria dispar (L.), were obtained from the Gypsy
Moth Methods Development Center, Otis Air Base, Massachusetts,
02542. Shipments were received weekly with the larvae packaged
in 8 oz. Sweetheart containers with a standard wheat germ diet 36
(Bell et al. 1981). The diet provided in the containers could
sustain the larvae until pupation. After being received, the
gypsy moth containers were kept in an environmental chamber with
a 16:8 Light/Dark cycle, 26 _+ 2° C. and a relative humidity
of approximately 40%.
Operations. Female gypsy moths were removed from the stock
colony three to five days after ecdysis to the last larval
instar. In all the following experiments, the larvae were placed
into three groups. The first was anesthetized with CC>2 only,
to serve as the unoperated controls. The second set was
anesthetized and operated on, but the organ(s) in question were not removed. These were designated as sham operated individuals.
The last group had the appropriate organ(s) removed in the operation.
Larvae were held with metal restrainers in a dissecting dish with Sylgard silicon rubber covering the bottom of the dish.
Larvae were anesthetized by passing CC>2 gas over them throughout the operation. After the operations, larvae of the same experimental group were housed with up to six individuals to a cup. These larvae continued to feed and develop. Pupated individuals were removed and placed in individual containers where they could continue to develop and eclose to adults.
Unilateral extirpation of paired glands and organs was ruled out as a possible control regime. Tobe (1977) showed that there 37
was asymmetrical release of humorally mediated compounds. Such
asymmetrical release would reduce the usefulness of this control
below acceptable levels.
Allatectomies. An incision was made dorsally in the post
cranial inter segmental membrane, exposing the tracheae,
esophagus, and muscles associated with the neck and
postoccipital foramen of the head. By pushing aside the
trachea, the CA could be exposed, one on each side of the
esophagus. Watchmakers forceps were used to grasp the corpus
allatum, usually by the nerve leading from the corpus cardiacum
and pulling the corpus allatum free. This procedure was
repeated bilaterally. After removing the corpora allata, the
head restraint was removed allowing the head to be retracted
back against the first thoracic segment. Dental wax was melted
over the neck region to prevent the loss of hemolymph after the
insect revived from the anesthesia. Sham operated individuals
were treated as above. After locating the corpora allata, a
small amount of fat body, approximately equal to the corpora
allata’s volume, was removed in place of the corpora allata.
Corpora cardiaca - corpora allata removal. Larval gypsy
moths were anesthetized and operated on in the same manner as the
allatectomies. The corpora cardiaca were located on either side
of the esophagus between the brain and the corpora allata.
Watchmakers forceps were used to grasp the connections of the
corpus cardiacum with the brain and remove the corpus cardiacum
and attached corpus allatum from the head. This procedure was 38
repeated bilaterally.
Ovariectomies. A dorsal longitudinal incision was made on the fourth abdominal segment of the larvae. Watchmaker’s forceps were used to grasp and pull the ovaries, one at a time, to the incision where the connections with the surrounding tissue could be cut free and the ovaries removed. Both the sham operated controls and ovariectomized larvae were composed of two
sub-groups. Clamps and dental wax were used to close the wounds in the first group and cyanoacrylate ester glue was
applied to the wounds in the other. The clamp was made from thin malleable wire, hammered flat, bent and cut to the appropriate / o — _ tm v size and shape. The cyanoacrylate ester glue (Super Glue )
was manufactured by Woodhill Permatex.
Bioassays. All surviving experimental individuals were
observed for calling behavior and bioassayed for sex pheromone
release 24 - 48 hr after adult eclosion. To minimize the
confounding factors of diel periodicity of male activity
(Cardfc et al. 1974) and sex pheromone release (Charlton and
Card& 1982), bioassays were performed between 6.5 to 9 hours
after the onset of photophase. The bioassay used was described
by Hollander et al. (1982). Briefly, the female moth was placed
at the open end of a glass tube, 5.8 cm. in diameter and 54 cm
long. Two newly emerged male gypsy moths were held downstream
between two brass screens approximately 10 cm apart in the tube. 39
The other end of the tube was closed by a rubber stopper with
a small funnel's stem protruding from it. Tygon tubing
connected the funnel stem to a rotometer and then to a vacuum
pump. Air was drawn through the bioassay tube at 8 cm/sec.
Males were allowed to acclimate for at least 10 min in the
bioassay tube before a pheromone source was introduced to
determine if the males were receptive to the sex pheromone. Males
showing no response were replaced. Blank controls were run prior
to each individual bioassay. No more than 10 bioassays were run
with any one pair of male moths to avoid habituation of the males
to the pheromone.
Excitation of the male gypsy moths to continual wing fanning
(Hollander et al. 1982) was scored in the bioassay as a positive
response. Males not wing fanning were scored as negative.
Calling behavior was scored by visually observing for the
presence or absence of the rhythmic extention and retraction of
the terminal abdominal segments (ovipositor) while the female was
being bioassayed. Following the bioassays, female gypsy moths
were dissected to ensure that the intended organ(s) had been
completely removed by the operation.
There was less than a 4% difference of males responding
among the control, sham and corpora allata removal groups.
Allatectomy had no effect on calling behavior or actual 40
pheromone release (Table 2).
Females that had both the corpora cardiaca and corpora allata removed showed little difference from the sham group
(Table 3). Two-thirds of the males responded to females that had both the CA and CC removed. About 96% of those females still exhibited calling behavior.
There was little difference between sham and ovariectomized groups with respect to percentages of females calling and males responding for both the cyanoacrylate ester glue and the clamp and dental wax techniques (Table 4). Individuals of both groups showed only minor scarring after molting into the pupal stage.
Neither the clamp nor the glue hindered the process of ecdysis.
The adult females that resulted from operated larvae could only be differentiated from control females by dissection. Control female gypsy moths had large abdomens, filled with eggs, ready to be fertalized and laid. The abdomens of ovariectomized moths had no eggs. Nevertheless, the abdomens of ovariectomized females appeared equally large. Dissection of these individuals revealed an abdominal cavity filled with traces of fat body, the midgut and the hind gut which may have been somewhat larger than the controls. Trachea which would have supplied the ovarioles, and hemolymph filled the remaining volume.
The overall mortality sustained by the groups closed with
clamp and dental wax glue was 6.0% as compared to 13.2% for the
glue technique. The operations with cyanoacrylate ester glue TABLE 2. Effect of removal of the corpora allata (CA) from the last larval instar on calling behavior and pheromone release by adult Lymantria dispar females. l-i rH i—I c CO 13 03 CO -H s a> E a 03 O 0) TO * * E—' 4-3 -P CO i- 13 C bO 13 c a; E p aj v—/ ^—V T— o CO cn LTi o o ON m o o -p i—1 u c O o o • • C\J /—\ .=r vO T— OO C\J on on o < cn 00 J3 c 03 E o • • /—. '—/ «J0 co OJ c— r— o^ on r*H [ o < O i. 13 e O > 03 • • •H O •H o TO > 2= O *H O •rH CO TO 3 •H £ TO JO >-! -H JO C * 1-. i—H 40 . > CO TO O 4-5 03 (U *H1 3 a; t- jo c 03 a. co i- 13 C 3 03 TO 03 J- E £> 03 p >> C TO CO CO l- C 4-> 3 13 13 l-i O 03 > -H CO 13 O. E 03 •rH > •rH 70 •rH 4-> •rH •H JC• JO JO 13 -P o SI 13 * * VI JO40 l-i 03 JO 03 JO i—1 *rH •rH CO •rH *H rH J-03 4-3 4-> O TO •H 1h s_ •rH C C S- C -rH CO l. CO >» CO bO 13 CX r-H CO —'13 13 O *H (0 o CO 'rH 13 3 X CO 13 C3 S- 4-3 O O JC C 13 13 13 rH E -P 13 E O JO13 E D 03 OE o co 13 C 3 <~ c a bO JO C E c o 03 O C 13 bO Cm JO 4-3 jQ CO 13 13 JO > 1_( O 13 >. 41 TABLE 3. Effect of removal of the corpora cardiaca and corpora allata (CC-CA) in the last larval instar on calling behavior and pheromone release by adult Lymantria dispar females. ft*. 0-. CO ** 05 rH C (U rH s 2 CO " CO *H e o. CO O 05 TO * * H 4-5 4-5 c to u O c w L. <15 s <15 CO =T in on t— o o o OO O O 4-5 rH LTi c i- o o o • • CM VO -- CO CVI o o o
were performed in mid summer when the naturally occurring nuclear polyhedrosis virus that attacks larval gypsy moths was quite prevalent. Despite great care, the colony and experimental animals were not completely free of this pathogen.
Discussion
Allatectomies. Juvenile hormone, normally released from the corpora allata, has been linked to sexual receptivity in a number of species (Engelman 1968, Roth and Barth 1964). Juvenile hormone also plays a significant role in ovarian maturation and development (Koeppe et al. 1980, Mwangi and Goldsworthy 1980).
Removal of the corpora allata from Galleria mellonella
(Roller et al. 1963), Bombyx mori (Steinbrecht 1964),
Anther aea Polyphemus and Hyalophor a £££_r £ £££ (Riddiford and Williams 1971)• however, had no effect on calling behavior or pheromone release. In the last species mentioned, by severing the nerves leading to the corpora cardiaca from the brain there was a decreased ability to call (Riddiford and Williams 1971).
In this work it has become evident that the corpora allata in the gypsy moth plays no role in calling behavior or pheromone release. Unlike the previously mentioned research, the operations were conducted on mid last instar larvae, in stead of the pupal stage. A method to indicate actual pheromone release
(i.e. the male wingbeat bioassay) was employed. The only
indicator of moth pheromone release used in previous studies 45
(Riddiford and Williams 1971) was the overt calling behavior.
The bioassay was deemed necessary since calling behavior and actual pheromone release had been shown to be separate physiological phenomena, (Hollander and Yin 1982).
Corpora cardiaca - corpora allata removal* Extracts from cockroach corpora cardiaca have a pronounced influence on efferent activity of the cockroach terminal abdominal ganglion
(Milburn and Roeder 1962). It is possible that the gypsy moth corpora allata or corpora cardiaca also releases compounds to control efferent activity of the terminal abdominal ganglion.
However, the removal of the corpora cardiaca from the gypsy moth has nearly no effect on calling behavior or pheromone release.
If similar compounds exist in the gypsy moth corpora cardiaca,
they might directly effect the terminal abdominal ganglion, the
muscles involved in calling behavior or even the pheromone
producing cells themselves. The studies presented here indicate,
however, these agents would not be a necessary prerequisite for
calling behavior or pheromone release.
It is possible that pheromone release or calling behavior
may be influenced or controlled by extra-ganglionic
neurosecretory cells such as those described by Griffith and
Finlayson (1982) in Agrotis segetum Schiff. These multipolar
peripheral neurosecretory cell soma were found within the
transverse nerves of the abdomen, but nothing is known as to
their roles in the regulation of the insect’s physiology or 46
their roles in the regulation of the insect’s physiology or behavior.
Ovariectomies, In many other insects, the ovaries play an important role in humoral mediation of sexual behavior. The mosquito ovaries, for example, seem to act as the central humoral receiver and broadcaster. Under normal conditions, the taking of a blood meal elicits release of egg development neurosecretory hormone (EDNH) from the corpora cardiaca which may stimulate the ovaries to release 20-hydroxyecdysone. By removing the ovaries,
Borovsky (1982) found that there was no vitellogenin produced, and no EDNH was produced. His associated experiments indicate that in addition to the previously noted role of the ovaries in mosquitoes, it appears that the ovaries are also stimulated into releasing a humoral agent by another humoral agent associated with the taking of a blood meal. The agent released by the ovaries is apparently the material responsible for the corpora cardiaca's release of EDNH.
The ovaries in cockroaches, a not so well characterized
system, are found to be important for the exhibition of calling
behavior and pheromone release (Barth and Lester 1973)*
The results and observations show that the morphology,
pheromone production, and development of the pheromone gland,
(and its associated structures), as well as calling behavior,
were not affected by ovariectomy in the last larval instar.
Larval ovariectomized females still released sex pheromone and 47
attracted males in the bioassay. This is strong evidence against any humoral involvement by the ovaries in these processes after the last larval instar. The lack of ovarian control, along with the lack of control from the corpora allata and corpora cardiaca, is consistent with Barth's hypothesis
(Barth 1965) that relatively long lived adult insects with multiple ovarian cycles will be more likely to use endocrine controls to coordinate their reproductive activities than short-lived adults with only a single ovarian cycle, such as the gypsy moth.
The removal of the ovaries seemed to have no effect on the external morphology of the female gypsy moth. Dissection
revealed the abdominal cavities to be filled with trachea, gut,
accessory glands and hemolymph. Since the fat bodies were nearly
nonexistent in the abdominal cavity of these ovariectomized
individuals, it is likely the fat body material was metabolized
into soluble forms and discharged into the hemolymph as in the
cockroach (Engelman 1978). Under normal circumstances these
materials would be actively removed from the hemolymph by the
developing obcytes. Since obcytes are not present, the
proteins and other material may remain in the hemolymph. It
would be interesting to expand upon the work of Brown and Mazzone
(1977) with comparisons of blood proteins from normal and
ovariectomized females and typical egg proteins. This would
provide some insight into where the materials from the fat bodies 48
go.
An effective method of closing the wound was crucial to the success of the ovariectomy operations. In most forms of insect microsurgery, the incision is made in or near substantially sclerotized areas of the cuticle. The sclerotized area gives a sturdy inflexible surface for wax to adhere. In the larval ovariectomy, the incision is made on very flexible membranous cuticle. If wax alone is applied to seal the wound the membranous cuticle will be flexed by the revived larva, loosening the wax closure from the cuticle. Hemolymph loss results.
Often, the midgut and hindgut herniate through the incision. In either case the death of the insect follows.
Two methods for closing the incisions were devised. The
first involved clamping the two edges of the incision together with a very small wire clip. When the clamp was in place, low melting point dental wax was applied to seal the rest of the wound. When the insect became active the clamp acted as a stable base for the wax and the cuticle to ensure a tight seal.
The second method involved applying cyanoacrylate ester
glue to the area of the incision. The properties of
cyanoacrylate ester glue allow it to adhere to the cuticle and
harden into an inflexible covering without the need for the
clamp type anchor discribed for the dental wax technique. The
glue could be applied even if the cuticle was slightly damp with
hemolymph, and yet still adhere well. The glue must dry
completely, however, before the insect is removed from the 49
anesthesia. Curing of the glue takes from 30 seconds to a minute. Unhardened glue may be dislodged from the cuticle much the same as dental wax without a clamp.
The results of the experiments illustrate that both of the techniques described work equally well. The higher mortality with the cyanoacrylate glue technique may be attributable to the presence of the nuclear polyhedrosis virus. Both techniques have the desired property of closing the incision and both take approximately an equal amount of time, effort and dexterity to use. The major criterion for the use of either of these techniques is the availability of the materials. CHAPTER IV
NERVOUS CONTROL
Introduction
Influences on the release of pheromones by female
Lepidoptera have been studied using environmental variables
(Sower et al. 1971a, Cardd and Roelofs 1973. Card<§ et al.
1975, Baker and Card6 1979, Castrovillo and Card<§ 1 979,
Bjostad et al. 1980, Alford and Hammond 1982) and classical endocrinological methods (Roller et al. 1963, Steinbrecht 1964,
Riddiford and Williams 1971). To date, however, no work has been devoted to establishing direct neural control over production and/or release of the sex pheromone in moths.
The calling behavior of the gypsy moth female is characterized by the rhythmic protrusion and retraction of the eighth and ninth (terminal) abdominal segments. The rhythmic calling behavior and attraction of the males in the wild occur during the day, mostly in the afternoon (Card£ et al. 1974).
When not calling, the two small terminal abdominal segments are retracted into a depression in the seventh abdominal segment.
There they are concealed by a thick covering of abdominal hairs.
During calling the pheromone glands are exposed by the extension of the terminal abdominal segments beyond the hair covering.
Previous research indicates the pheromone glands are located on 51
the dorsal and ventral aspects of the inter segmental membrane,
between the eighth and ninth abdominal segments. They appear as
two highly convoluted integumentary areas with enlarged glandular
epidermal cells (Hollander et al. 1982).
Since calling is associated with pheromone release, a neural
control mechanism was suspected in the coordination of the
abdominal movements (calling) and pheromone release. This study was designed to explore the basic nervous controls involved.
The adult gypsy moth nervous system is composed of a brain and subesophageal ganglion in the head, two thoracic ganglia and
four abdominal ganglia (Fig.11). The main connections to be examined were those between the terminal abdominal ganglion and the terminal abdominal segments, and between the brain and the terminal abdominal segments. These experiments provided information on the influences that the brain and terminal abdominal ganglion have on pheromone release and calling behavior.
The male wingbeat bioassay used gave a good indication of whether pheromone was being released, however, the bioassay is still a qualitative assay. Currently, there are no quantitative methods to directly measure production of pheromone. Pheromone gland washs quantified with gas chromatography, such as those decribed by Coffelt et al. (1978), only provide information on how much pheromone is present on the gland at a particular time.
Aerial collection of emitted pheromone promised both measurable amounts of pheromone and an indication of production over a 52
Fig. 11. Diagramatic dorsal view of the gypsy moth nervous system including the corpora cardiaca (CC), corpora allata (CA), terminal abdominal ganglion (TAG), and terminal abdominal segments (TAS). 53
_c CC k- m 54
relatively long span of time. Two methods for aerial collection
of gypsy moth sex pheromone and gas chromatographic analysis
have been described (Charlton and Card* 1982, Ma and Schnee
1983). These two techniques were used in an attempt to quantify
the difference in overall pheromone production between controls and female moths with the ventral nerve cord severed.
Since the brain appears to be responsible for neurological control of pheromone release, stimulation of the nerves leading to the pheromone gland might elicit pheromone release. To test this hypothesis, female gypsy moths were prepared by severing the ventral nerve cord. Severing the ventral nerve cord stops pheromone release, but calling behavior continues (Hollander and
Yin 1982). Nerves leading from the terminal abdominal ganglion were stimulated electrically and pheromone release was monitored with the bioassay technique.
Methods and Materials
Experimental animals. Gypsy moth larvae, Lymantria dispar, were acquired from the Otis Gypsy Moth Methods Development
Center, Otis Air Base, Mass., and reared on an artificial gypsy moth diet provided with the larvae. The colony was kept in an environmental chamber at 26 + 2° C, on a 16 hour light : 8 nour dark cycle at approximately 40% relative humidity. 55
Operations. Females were operated on within 24 hours after
pupal ecdysis. Pupae were restrained in a plasticine cradle,
under CO^ gas anesthesia. Incisions on the operated
individuals were sealed with melted dental wax. For each set of operated individuals, a comparable number of sham operated and
CO^ anesthetized unoperated pupae served as controls.
Operations on ventral nerve cord. An incision was made on
the ventral side of the 6th abdominal segment. The ventral nerve cord was exposed and cut anterior to the terminal abdominal ganglion, or the terminal abdominal ganglion was removed, or the nerves leading posteriorly from the terminal abdominal ganglion were severed.
Operations on the brain. A window was cut through the pupal cuticle directly over the pupal brain. The brain was then removed or the circumesophageal connectives were sectioned. This technique was patterned after Zarrow et al. (1964). The square of cuticle that had been removed was replaced to its original position. Dental wax was melted over the wound to seal it.
Experimental moths were allowed to develop to the adult stage in individual containers. After adult eclosion, moths were observed for calling behavior and bioassayed for pheromone release.
Bioassay. Experimental and control female moths were bioassayed for pheromone release 24 - 48 hr after adult eclosion.
All bioassays were run between 6.5 - 9 hr after the onset of the photophase. The bioassay used was essentially the same as that 56
described by Hollander et al. (1982). Briefly, the female was
placed at the head of a glass tube 5.8 cm in diameter and 54 cm
long. Two newly emerged male gypsy moths were held downstream in
the tube by a disposable cheese-cloth barrier. Air was forced
through this no-choice olfactometer at a flow rate of 2 to 3
cm/sec from a compressed air cylinder. Males were allowed to
acclimate for at least 10 min before females were introduced.
A normal, calling female was first introduced into the
bioassay tube for up to 3 min to establish that the males were
receptive to the pheromone. Blank controls were run prior to
every individual bioassay. No more than ten bioassays were done
with any one pair of male moths. Excitation of the males to
continual wing fanning after exposure to a female (Card* and
Hagaman 1979) was scored as positive for pheromone release.
Calling was determined by visual observation for the presence or
absence of the rhythmic extensions and retractions of the
terminal abdominal segments while the female was being
bioassayed. Operated females were dissected after being
bioassayed, to ensure the operation had been successful.
Aerial collection of pheromone.
Ma - Schnee technique. For full details, see Ma and
Schnee (1983). Adult female gypsy moths were held for a 24 hr
period in a vertical glass tube 3 cm in diameter and 10 cm long.
The moths were allowed to hang on to a tubular aluminum screen 57
inside the glass tube. Both ends of the tube were stoppered, the bottom stopper having a hole 6 cm in diameter running through it to allow air passage into the chamber. The top stopper had two holes large enough to accommodate a 2.5 mm inside diameter, 3»5 cm long glass tube in each. Each glass tube was packed with
Porapack Q or Tenax. The packing material was washed with hexane, then clean filtered nitrogen gas was passed through it at a temperature of 160° C for six hours or more. The Porapack Q or Tenax was held in place in the collection tubes by fiberglass plugs. Both tubes (aerial collection filters) were connected with Tygon tubing to a vacuum pump running with a negative pressure of 10 PSI to draw air through the system.
The aerial collection filters were removed after 24 hr of collection. The tubes were then placed in vials and stored at
-4° C until they were analyzed. Filters were eluted with a total of 0.5 ml of methanol. Five microliters of the elutent was injected into an Envirochem Series 4 sample concentrating gas chromatograph with a fused silica column containing OV 101, and flame ionization detector. Analysis of the chromatograms was accomplished with a Spectrophysics 4100 micro-computer.
Charlton - Card& technique. For full details, see
Charlton and Cardfc (1982). This method consisted of a glass holding chamber with ground glass connectors. The female gypsy moth hung in the chamber from a suspended metal screen. Above the moth was an incurrent filter of glass beads. Below the moth was a larger tube that curved 90° and ended with a ground glass 58
fitting. The pheromone collection filter connected to this fitting. The filter had a glass frit in the middle of the tube, was packed with 1 mm diameter glass beads behind the frit, and was stoppered with glass wool. Air was pulled through the chamber and filter at approximately 2 ml per second. Aerial collectors were eluted with two hexane washes of 2 ml each. The hexane from the washes was stored at -2° C until the samples could be analysed.
An internal standard, cis -9,10- epoxy-eicosane, was added
at 1000 ng/sample. Samples were evaporated with nitrogen then
reconstituted with 25 microliters of carbon disulfide (CS^).
One microliter of the concentrate was injected onto a Varian 3700
gas chromatograph. The chromatograph was equipped with a 60 m,
SP 2100 capillary column and a flame ionization detector and used
nitrogen as the carrier gas. The injection port and detector
were kept at a constant 250° C. The column oven was programmed
to hold at 90° C for two minutes after sample injection, then
increase linearly at six degrees per minute until a temperature
of 210° C was attained. This temperature was maintained for 25
minutes before returning to the starting temperature. The
resulting chromatograms were analyzed with a Hewlett-Packard 3390
recording integrator, with an attenuation setting of 64.
Stimulation of nerves from the terminal abdominal ganglion.
Operations on the ventral nerve cord were performed as described 59
earlier. Control and sham operated groups were also included as before. Female gypsy moths, between 24 - 48 hr after adult eclosion, were pinned upside down through the thorax in a dish.
The ventral aspect of the abdomen was opened to expose the terminal abdominal ganglion. Silver hook electrodes, attached to a Model SD9 Grass Stimulator, were placed under the last two pairs of major nerves (the fifth and sixth major nerves as counted from anterior to posterior) of the terminal abdominal ganglion (Fig. 12). The nerves were stimulated at 16 Hz, 2 msec duration and 0.4 V. To determine if the females were releasing pheromone as a result of the stimulation, a slightly modified bioassay system was used. A funnel was placed at the incurrent opening of the bioassay tube, the stem pointing outward so it could be brought very close to the terminal abdominal segments of the pinned female moth. The position of the funnel stem would draw any pheromone being released into the biassay apparatus. The observations on female calling and male response were scored as in the bioassay section above.
Results
Anatomical relationship of the nervous system to the pheromone gland. Several studies to determine if and/or how the pheromone gland might be associated with the nervous system were attempted.
The histological preparation of the terminal abdominal segments
(Hollander et al. 1982) did not differentiate nervous tissue, so 60
another approach was necessary. Bronskill’s wholeraount technique
(Bronskill 1970) allowed for an overall view of the regions involved, but also did not differentiate nervous tissue. Cobalt chloride injections (Peters 1976, Bacon and Altman 1977) of the nerves leading posteriorly from the terminal abdominal ganglion
were attempted, but due to the extremely low success rate, were not practical. The best results were obtained using a methylene blue technique (Humason 1967). Observations made from
dissections in conjunction with the methylene blue technique
indicated that there are six major nerve pairs leading from the
terminal abdominal ganglion, not including the two large ventral
nerve cords (Fig. 12). Two of these paired nerve bundles, the
fifth and sixth, have branches that lead to the terminal
abdominal segments in the region of the pheromone glands. It is
possible that there may be direct innervation of the cells that
produce and release the sex pheromone. However, it is likely
that the vast majority of the nerve axons in these bundles
innervate the circular-oblique and longitudinal muscles
associated with this region.
Experimental effects. The results of the surgeries are
summarized in Tables 5 and 6. The operations performed had
little effect on either metamorphosis or adult eclosion of the
females. Severing the ventral nerve cord, terminal abdominal
ganglion removal, cutting the nerves running posteriorly from the 61
Fig. 12. Diagraraatic ventral view of the gypsy moth terminal abdominal ganglion (TAG), ventral nerve cord and six major nerves (numbered 1 through 6 ). 62
6 5 TABLE 5. Effect of operations conducted on the pupal nervous system with respect to calling behavior and pheromone release by adult female Lymantria •H | ■Ol M to! cx\ co! ** Vh i—I T3 i-H CO C
#*»* TAG = Terminal abdominal ganglion 63 TABLE 6. Effect of severing the circumesophageal connectives (CEC) in the pupal stage on calling behavior and pheromone release by adult female Lymantria dispar . tr*
terminal abdominal ganglion, brain removal and severing the eircumesophageal connectives all disrupted the ability to release pheromone (Table 5). The only operations that severely curtailed calling were removal of the terminal abdominal ganglion or severing the nerves leading posteriorly from the TAG. Isolation of the terminal ganglion from higher centers by section of the ventral nerve cord, cutting the eircumesophageal connectives or removal of the brain had no observable effect on calling. Calling by operated females generally appeared normal at the time of
their bioassay.
Brainless females were able to eclose and call, but were
unable to expand their wings after eclosion. These females also
had great difficulty in the coordination of posture and
locomotion. These problems are similar to those noted by Truman
(1979) in silkmoths after brain removal.
The brain is not only a neurological organ, but also an
important humoral organ (Raabe 1982). By removing the brain in
the pupal stage, the humoral agents to be released by the brain
will also be removed. To show the neurological component of the
brain’s control, the eircumesophageal connectives were severed,
but the brain left intact. This experiment had the same results
as those of brain removal. Those individuals with the
eircumesophageal connectives cut, were still able to exhibit
calling behavior, but were unable to release pheromone (Table
6). Dissection of experimental adult females subsequent to being
bioassayed showed no regenereation of nerves or ganglia. 66
Aerial pheromone collections were made from individual female gypsy moths in three groups: C02 controls, sham operated, and ventral nerve cord cut anterior to the terminal
abdominal ganglion. The Ma - Schnee technique and the Charlton -
Card* technique were used in the attempt to quantify the
differences in the amounts of pheromone released by these
experimental groups. Neither the Ma - Schnee method nor the
Charlton - Card* method provided satisfactory quantitative
results. The standard dilution curve of synthetic disparlure for
the Ma - Schnee method may be found in Fig. 13. Data for the
standard curve are located in Table 8. Chromatograms from the Ma
- Schnee technique were difficult to read and no pheromone peak
could be discerned from the background (Fig. 14). The Charlton -
Card* technique suffered from a similar problem. The pheromone
peak from the aerial collections, if present, were not disernable
from the background. Representitive chromatograms of aerial
collectons show the lack of a pheromone peak in a 12 hour
collection with no female (Fig. 15) and the location of the
synthetic disparlure peak added to another blank control (Fig.
16). No corresponding pheromone peak was found in 12 hr
collections from calling control female gypsy moths (Fig. 17).
Direct stimulation of the fifth and sixth major terminal
abdominal ganglion nerves of female gypsy moth resulted in no
significant increase in male response (Table 7). There was a 10%
increase in males responding during stimulation in the VNC cut 67
Fig. 13. Standard dilution curve for concentrations of disparlure (cis-7,8- epoxy -2- methyloctadecane), assayed using an Envirochera-Model 780B gas chromatograph, with a flame ionization detector and a capillary column. Correlation coefficient, r = 0.977. The linear equasion for the regression line is Y = 64.05 - 757.0. See Table 8 in the Appendix for data . DISPARLURE STANDARD CURVE eejv >|Becj 68 69
Fig. 14. Representative chromatogram from the Envirochem Model 780B gas chromatograph. The sample was a 24 hr aerial collection from an adult control female. Average retention time for disparlure was 29.43 min and 20.84 min for the hexadecane internal standard. 70
■nj 71
Fig. 15. Representative chrom atogram from the Varian 3700 gas c hromatograph. The sample was a 12 hr aerial collection from a control with no female. One microl iter of sample was in j ec ted (1/25 of the total reconst ituted sample) with 40 ng of cis - 9,10- epoxy - eicosane (inter nal st andard , retention time 16.84 min) .
Fig. 16. Representative chromatogram from the Varian 3700 gas chromatograph. The sample was a 12 hr aerial collection from a control with no female. One microliter of the sample was injected (1/25 of the total reconstituted sample) with 40 ng of disparlure (retention time 14.96 min) and 40 ng of cis -9,10- epoxy - eicosane (internal standard, retention time 16.86 min) . 72 73
Fig. 17. Representative chromatogram from the Varian 3700 gas chromatograph. The sample was a 12 hr aerial collection from a stock colony female. One microliter of the sample was injected (1/25 of the total reconstituted sample) with 40 ng of cis —9,10— epoxy — eicosane (internal standard, retention time 16.86 min). 74 TABLE 7. Effect of stimulation of the fifth and sixth major nerves of the terminal abdominal ganglion on calling behavior and pheromone release by adult female Lymantria dispar. X UJ E-* CO < X O cc UJ Q UJ H Ou > z cc <0 -J UJ cc UJ o O O CO > cc Q 33 H O O z H OS o -J **
group. This may show some increase in release of pheromone, but was not significantly different from the prestimulation regimes
(95% confidence level). Data was transformed by the arcsin square root transformation of the percentages. Statistical inferences were made from the result of 2 way ANOVAs calculated with the transformed data. There were significant differences among the control, sham operated and VNC cut groups. The VNC cut percentages were significantly lower than either the control or sham operated groups. There were no statistically significant differences among rows or columns for percentages of calling females. The data for the number of calling females and responding males may be found in Tables 9 and 10.
Even though the preparation for nerve stimulation required a large ventral abdominal incision the gypsy moths typically continued to call throughout the duration of the procedure. If the female was not calling at the start of the operation, a slight stimulation of the terminal abdominal segments with a pin or pair of forceps would prompt the continuous rhythmic extention and retraction of the terminal abdominal segments.
Discussion
Hitherto, humoral influences have been the main target of control systems research on pheromone production and release
(Barth and Lester 1973, Riddiford 1974). The present study clearly shows that the central nervous system has an important 77
role in pheromone release and associated behaviors of the gypsy
moth. To understand how the nervous system exerts control over
the release of pheromone, the relationship between the nervous
system and pheromone gland tissue must be determined. Because of
the effect that cutting the nerves anterior to the terminal
abdominal ganglion has, and the presence of nerves leading into
the vicinity of the pheromone gland, direct innervation of the
pheromone gland is a distinct possibility. However, direct
innervation remains to be shown experimentally.
The brain appears to be the control center for pheromone
release in female gypsy moths. The brain of L. dispar has
been found to be an important humoral organ for diapause and
diapause development in maturing embryos of the gypsy moth (Loeb
and Hayes 1980). By removing the brain, cessation of pheromone
release might have been caused by loss of humoral agents.
However, the same results may be obtained with the brain intact by severing the circumesophageal connectives or cutting the ventral nerve cord anterior to the terminal abdominal ganglion.
These data do not rule out a humorally mediated control
system, however. Possibly both neural and hormonal mechanisms
play roles in the control of the pheromone gland. Humoral
agents, however, are not sufficient by themselves. This work indicates that humoral controls would have to be dependent on
intact nervous connections for those agents to be effective.
There are a number of cases where glandular tissues have been 78
shown to be neurally controlled. Such examples include the
nervous control of secretion of adipokinetic hormone from the
corpora cardiaca of Locusta (Rademaker 1977) and cholinergic
neurons regulating salivary fluid secretions in ixodid ticks
(Teel et al. 1978). Although neural control is not unique among
glandular systems, nervous control of pheromone release is a new
finding.
Calling behavior, in contrast, does not require the central
nervous system anterior to the terminal abdominal ganglion.
Calling appeared to continue unaffected, even when the brain had
been removed or the ventral nerve cord had been severed anterior
to the terminal abdominal ganglion. The moth’s ability to control
the rhythmic extensions and retractions of the terminal abdominal
segments was lost only after removing the terminal abdominal
ganglion or severing the nerves leading posteriorly from the
terminal abdominal ganglion.
These results indicate that the terminal abdominal ganglion
is important in central or peripheral pattern generation for the rhythmic calling behavior, and is sufficient to initiate calling behavior without nervous communication with the rest of the central nervous system. Comparable pattern generators have been reported by Roeder et al. (I960), Truman (1979) and Delcomyn
(1980). Even so, there may still be humoral influences such as those found in the cockroach (Milburn and Roeder 1962) and the giant silkworm moth (Truman 1979).
In the Orthoptera, severed motor neurons have been found to 79
regenerate to the thoracic leg muscles (Denburg et al. 1977,
Pearson and Bradley 1972, Young 1972, Wood and Usherwood 1979,
Donaldson and Josephson 1981, Whitington 1979). The typical time
course for nerve regeneration is around 60 to 90 days in adults.
This is considerably longer than the 11 to 13 days between the
operations to sever nerves and the bioassays on the gypsy moth
females. However, if damage occurs in a nymphal stage of
Periplaneta americana, the regeneration is considerably
faster. Regeneration rates as fast as 12 days (Guthrie 1967), 13
to 14 days (Denburg et al. 1977) and 30 days (Jacklet and Cohen
1967) have been reported. These periods are much closer to the
time frame in which the experiments described here were carried out. Dissections of experimental adult gypsy moths showed no sign of regeneration of nerves at any of the locations where nerves were cut.
In the silkworm moths, the site of the biological clock responsible for timing of eclosion and flight rhythms is in the brain (Truman 1972, 1974). In both cases the brain’s message is humorally mediated. Calling behavior of brainless female gypsy moths observed during the bioassay appeared to be normal.
Observations were only made between 6.5 and 9 hr after the onset of photophase, so the roles of neural and humoral control systems in the gypsy moth’s diel rhythm of calling behavior can not be addressed at this time.
The demonstration of neural control over calling behavior 80
leads to speculation on the nervous connections involved. There
are many ways that neural networks could be organized to perform
the observed functions, but from the data presented, many of the
simpler networks may be ruled out. Inhibition is the classic
control function of the brain for such acts as reproductive
behavior in the male praying mantis (Roeder 1935). For calling
behavior, neural inhibition by the brain could very well be
important. Calling is present, even when the brain is removed
from nervous connection with the terminal abdominal ganglion. The experiments conducted in this study were designed to test for a removal of the behavior when released from active controls, rather than an inhibition of behavior by the brain. Therefore, these experiments are not conclusive on this point. The behavior could just as easily be inhibited humorally at the appropriate times during its diel rhythm.
The results of the experiments dealing with pheromone release were not consistent with a hypothesis of classical inhibition by the brain. If the brain is acting as an inhibitor, when the ventral nerve cord is severed, there should be a loss of inhibition leading to pheromone release. The ability to release sex pheromone was lost when the nervous connections between the brain and the pheromone gland were severed. This inability to release pheromone indicates the brain must actually be stimulating rather than inhibiting release.
If the brain is responsible for the stimulation of pheromone release, then artificial stimulation might prompt pheromone 81
release. In the experiment to test for pheromone release with electrical stimulation of the nerves leading to the area of the pheromone gland, only a minor increase in male response was observed. This increase is statistically insignificant, so does not support the hypothesis that the glands can be stimulated into releasing pheromone by exogenous electrical stimulation.
It is possible that the correct voltage and/or duration of stimulation was not tested, leaving this question open to further investigation. 82
CHAPTER V
SUMMARY AND CONCLUSION
In the research discussed to this point, the goal has been
to shed new light on controlling factors involved in the female
gypsy moth’s calling behavior and sex pheromone release. To do
this, I had to first determine where the pheromone gland was
located. Bioassays of sections of the two terminal abdominal
segments, histology and scanning electron microscopy were used.
These techniques indicated that the sources of the gypsy moth sex
pheromone are two discrete regions: one on the dorsal and one on the ventral aspects of the inter segmental membrane between the last two segments of the abdomen. These areas have a highly convoluted cuticle underlaid by columnar and goblet shaped epithelial cells as opposed to the squamous and cuboidal cells that line the non-convoluted regions of the inter segmental membrane.
Microsurgical extirpation of corpora allata, corpora cardiaca, and ovaries in last larval instar females had little or no effect on the insect’s ability to call or release pheromone.
This indicates that the humoral agents associated with these organs are not directly involved in pheromone release and/or calling behavior. In addition, this indicates the aforementioned organs are not necessary for developmental aspects of pheromone release or calling behavior since they were 83
removed before the onset of pheromone gland tissue
differentiation. These results do not rule out the possibility
that there are secondary sources of humoral agents normally
associated with the corpora allata, ovaries and brain. If
secondary sources do exist, their products might affect calling
behavior and pheromone release even in the absence of the
classical source organs.
The results indicate that the brain lacks direct humoral
controls over calling and pheromone release. The moths not only
lost the ability to release pheromone after the brain was
removed, but also when the circuraesophageal connectives or
abdominal ventral nerve cord were severed. If there are any humoral factors from the brain involved in pheromone release, they are apparently overshaddowed by the disruption of nerve connections between the brain and the pheromone gland region.
Neither the severing of the circumesophageal connectives nor brain removal seem to hinder the ability of the adult female to elicit calling behavior. This is strong evidence against the involvement of humoral or nervous factors from the brain in the promotion of calling behavior. If anything, the brain may act as a source of behavioral inhibition, much the same as the classical examples of inhibition by the brain in praying mantids, and cockroaches as described by Roeder (1937) and
Roeder et al. ( 1 960).
By severing the female gypsy moth' s ventral nerve cord 84
anywhere between the brain and the pheromone glands, pheromone release does not occur. Calling behavior, however, will continue unless the terminal abdominal ganglion is removed or the nerves leading from the terminal abdominal ganglion to the pheromone gland region are severed. By removing the terminal abdominal ganglion or severing the nerves leading to the pheromone gland, both pheromone release and calling behavior are terminated. The results strongly suggest that the controlling center for calling behavior is located in the terminal abdominal ganglion. The motor pattern generator for calling behavior that is associated with the terminal abdominal ganglion can undoubtedly be affected by nervous and/or humoral influences. However, in the absence of such influences, the motor pattern generator is able to function autonomously to drive calling behavior.
The methylene blue staining of the abdominal nerves showed that there are six major pairs of nerve bundles originating from the terminal abdominal ganglion. Of these six, the fifth and sixth nerves, (numbered in anterior to posterior sequence) have branches that end in the regions of the dorsal and ventral convolutions of the inter segmental membrane between the eighth and ninth abdominal segments. Although the nerves were not traced down to their ending in the current study, many of the endings are assumed to be associated with the muscles in the terminal abdominal segments. Some of these nerves could be associated with the pheromone gland cells directly. Although direct innervation of the gland cells has not been shown, the 85
results of these experiments would be consistent with the
hypothesis that stimulatory nerves synapse onto the pheromone
gland cells in the gypsy moth.
Calling behavior and pheromone release have been used
synonymously in a number of previous studies (Riddiford and
Williams 1971, Leppla 1972, Sanders and Lucuik 1972, Percy 1974,
Shorey 1974, Richardson et al. 1976, Percy and George 1979). The
data presented here, however, show that calling behavior and the
actual release of pheromone are physiologically separate
entities. Therefore, the practice of using these terms
synonymously should be discouraged in the future.
It appears that the physiological relationships and control
systems in gypsy moth reproduction are not as complex as those of
some other insects. Elaborate control systems for female
reproductive behavior may be unnecessary because of the gypsy moths’ short single purposed adult life. This fits well with
Barth’s hypothesis (Barth 1965) that insects capable of multiple
reproductive cycles are more likely to have elaborate humoral
control systems for the synchronization of reproductive
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Correlation coefficient r = 0.98 97 Table 9. Effect of stimulation of the fifth and sixth major nerves of the terminal abdominal ganglion on calling behavior of adult female Lymantria dispar. (n = the number of females bioassayed) O H co o 2 DC O _J o UJ