Different aspects of the biology of phlebotomine sand flies (Diptera: psychodidae) and their age structure in a focus of...


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Different aspects of the biology of phlebotomine sand flies (Diptera: psychodidae) and their age structure in a focus of cutaneous Leishmaniasis in northeastern Colombia
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xix, 290 leaves : ill., photos. ; 28 cm.
Mahmood, Farida, 1953-
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Thesis (Ph. D.)--University of Florida, 1990.
Includes bibliographical references (leaves 262-287).
Statement of Responsibility:
by Farida Mahmood.
General Note:
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University of Florida
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Copyright 1990


Farida Mahmood

I dedicate my dissertation to my parents who always

encouraged my search for knowledge.


I thank all the people in Gainesville, Medical

Entomology Laboratory and in Florida Medical Entomology

Laboratory, Vero Beach Florida, who assisted me during the

course of my studies.

I greatly appreciate the close supervision of my

committee chairman and cochairman for taking a close

interest in my work during this study. I especially thank

Dr. Jerry F. Butler for allowing me to use his laboratory

facilities. I am indebted to Dr. Butler for providing me

with ample supply of photography materials during my stay at

Gainesville and I especially thank him. I also thank Dr. S.

Zam for providing me with his wealth of knowledge and kind


I enjoyed the pleasant company of all the graduate

students at Building 62 especially Nedeweso Kiwia, Al

Gettman, Bruce Alexander and Ben Beard and I thank them for

their help. I especially thank Diana Simon, Margo Duncan,

Debbie Boyd and Edna Mitchell for their help and assistance

during my studies in Gainesville.

I thank Dr. R. H. Baker and the administration and

staff of Florida Medical Entomology Laboratory, for making

my stay a pleasant lasting experience. I especially thank

Bonnie K. Pattok for helping me in times of need and

providing me with her advice for the art work. I especially

thank Mrs Alice Helleso at Vero Beach for her help and good

company. Finally I thank my parents and family for

providing me with love and encouragement.



LIST OF TABLES. ... . . .ix


ABSTRACT . . .xviii


Medical Importance of Sand flies . .1
American Leishmaniases . 2
Effects of Leishmania Infection on Sand Flies 4
Life Cycle of Phlebotomine Sand Flies . 9
Objectives . . 11


Reproductive Biology of Sand Flies. .. ... 13
Morphology of the Male Reproductive System 16
Digestion of Food by Sand Flies. . ... 19
Digestive System of Phlebotomine Sand flies .19
Different Enzymes Secreted during Digestion of
Food. . . 32
Female Age Grading . .. 46
Reproductive System of Female Sand Flies .46
Different Methods of Age Grading Sand Flies 53
Hormonal Control of Insect Reproduction and Role of
Juvenile Hormone III in Adult Insects .. 61


Introduction . . .72
Materials and Methods .... .. .. 73
Investigation of Epigamic Behavior .75
The Effect of Age on Mating Competence. .. 75
Effects of Age and Mating on the Morphology of
Male Reproductive System .. ... 78


Results .... .. . ..... .79
Mating Behavior of Sand Flies .. 79
Effect of Age on the Mating Competence of
Lu. anthophora .... .. 83
Morphology of the Male Reproductive System .85
Effects of Age and Mating on the Morphology of
the Reproductive System . .. .90
Discussion . . 105


Introduction . . .. 113
Materials and Methods . .. 115
Duration of Larval Instars . .. 116
Quantitative Determination of Trypsin- and
Chymotrypsin-like Enzymes .. 116
Polyacrylamide Gel Electrophoresis (PAGE) and
Fluorography . . 119
Statistical Analysis . .. 121
Results . ... . 124
Duration of Different Larval Instars .. 124
Trypsin- and Chymotrypsin-like Enzymes of Lu.
anthophora . . 129
Polyacrylamide Gel Electrophoresis and
Fluorography of Trypsin- and Chymotrypsin-
like Isozymes . .. 143
Discussion . . .. 151


Introduction . . 156
Materials and Methods . .. 159
Results . . 167
Discussion . . .168


Introduction . . 172
Materials and Methods .. .. 173
Rate of Egg Development in Lu. anthophora .173
Sampling Methods . .. 178
Tree Trunk Collections . .. 178
Human Bait Collections . .. 179
Light Trap Collections . .. 179
Leaf Litter Collections . .. 179


Indoor Resting Collections . .. .180
Resting Collections from the Rocks 180
Dissections Methods ..... . 180
Calculations. . . 182
Results . . 182
Morphology of Female Reproductive System and
Rate of Egg Development in Lu. anthophora .182
Changes in the Reproductive Status and Age
Structure of Wild Caught Sand Flies .194
Discussion . . ... .237


REFERENCES. .. . . ... .262




Table Page

3-1. Effects of age on the mating ability of
virgin males of Lu. anthophora ... .84

3-2. Effects of age on the morphology of the
reproductive system of virgin male Lu.
anthophora. ..... .... 89

3-3. Effects of matings on the morphology of the
male reproductive system of Lu. anthophora 94

4-1. Age related changes in the width of head
capsule and the length of the body of
different larval instars of Lutzomyia
anthophora . .. .125

4-2. Time course of the production of serine
proteases in immature and adult Lutzomyia
anthophora . ... .. 130

4-3. Time course of the production of serine
proteases in blood fed Lutzomyia anthophora 131

6-1. Rate of egg development after blood feeding
in Lutzomyia anthophora . .184

6-2. Phlebotomine sand flies collected at Arboledas
and Durania, Colombia. . ... .189

6-3. Changes in age structure of Lutzomyia
cayennensis collected by various methods in
Durania, Colombia during January 1987 to
January 1988 .... .. 192

6-4. Changes in the age structure of Lutzomyia
ovallesi, collected resting on tree trunks in
Arboledas, Colombia, during August 1985 to
September 1986 . . 199

6-5. Changes in age structure of Lutzomyia
ovallesi, collected by different methods in
Durania, Colombia, During January 1987 to

Table Page

January 1988. .... . .. .200

6-6. Age structure of Lutzomyia serrana,
collected resting on tree trunks in
Arboledas, Colombia during August 1985 to
September 1986 .. . .. 205

6-7. Changes in the age structure of Lutzomyia
shannoni, collected resting on tree trunks
and during human bait biting collections in
Arboledas, Colombia during August 1985 to
September 1986 ... .. 212

6-8. Changes in age structure of Lutzomyia
shannoni collected by different methods in
Durania, Colombia during January 1987 to
January 1988 .... .214

6-9. Changes in longevity of Lutzomyia
spinicrassa collected resting on tree trunks
and during human bait collections in Arboledas,
Colombia during August 1985 to September 1986. .222

6-10. Changes in age structure of Lutzomyia
spinicrassa, collected by different methods
in Durania, Colombia, during January 1987
to January 1988. .. .224

6-11. Age structure of Lutzomyia trinidadensis,
collected by different methods in Durania,
Colombia, during January 1987 to January
1988. .. ........ . 230

6-12. Changes in the structure of Lutzomyia
venezulensis, collected by different methods
in Durania, Colombia, during January 1987 to
January 1988. ... .232

6-13. Changes in the age structure of Lutzomyia
gomezi collected during human bait
collections from Arboledas, Colombia, during
August 1985 to September 1986 ... .235

6-14. Changes in the age structure of Lutzomyia
erwindonlodoi collected resting on tree
trunks and in human bait collections in
Durania, Colombia, during March 1987 to
October 1987 ..... .. ..236


6-15. Number of gonotrophic cycles completed by
different Lutzomyia species during 1985-1988
in Colombia . ... .238


Figure Page

2-1. Male reproductive system of Lutzomyia sauroida .15

2-2. Internal anatomy of Lutzomyia sand flies
showing various organs. .... 18

2-3. Digestive system of Lu. anthophora 20

2-4. Diagramatic representation of the activation
of chymotrypsin resulting in the formation of
a chymotrypsin . .. 38

2-5. Charge relay system of chymotrypsin molecule. 39

2-6. Acylation reaction during the hydrolysis of a
peptide bond by chymotrypsin .. ... 41

2-7. Deacylation reaction during the hydrolysis of a
peptide bond by chymotrypsin molecule 42

2-8. Irreversible deactivation of trypsin-like
enzyme by DFP . .. 43

2-9. Diagramatic representation of the reproductive
system of a female sand fly ... 45

2-10. Different stages of the follicular development
in Lutzomyia anthophora. . .48

2-11. Digramatic representation of the different
stages that are found during the formation of
dilatations. .. . 57

3-1. The reproductive system of male Lutzomyia
anthophora. . . 77

3-2. Spermathecae of a mated Lu. anthophora .81

3-3. Different parts of the reproductive system of
Lutzomyia anthophora . 86

3-4. Effects of age on the testes of virgin



Lutzomyia anthophora. . ... 87

3-5. Effects of matings on the testes of 7-day-old
Lutzomyia anthophora showing progressive
decrease in the size of sperm reservoir and
amount of spermatozoa (Sp) after successive
matings. . . .. 100

3-6. Effects of succesive matings on the anterior
portion of the accessory glands and seminal
vesicles of 7-day-old male Lu. anthophora,
showing progressive decrease in size and
depletion of Light yellow MAGS (Lt) and
spermatozoa (Sp). . .. .. 102

3-7. Effects of matings on the posterior portion
of the accessory glands of 7-day-old male
Lutzomyia anthophora showing progressive
decrease in size and depletion of dark
yellow MAGS (Dk) after successive matings. 103

4-1. Relationship between the body length and age
(days) in the larvae of Lutzomyia anthophora 122

4-2. Relationship between head capsule width and
age (days) of the larvae of Lutzomyia
anthophora . . 123

4-3. Relationship between body length and head
capsule width of Lutzomyia anthophora larvae. 126

4-4. Calibration curve of trypsin, showing
relationship between trypsin concentration
(ng/100l) and [1,3-3H]DIP-trypsin
derivatives . . 127

4-5. Calibration curve for the determination of
chymotrypsin, showing relatinoship between
chymotrypsin concentration (ng/100 pl) and
[1,3- H]DIP-chymotrypsin derivatives 128

4-6. Time course of the changes in trypsin- and
chymotrypsin-like enzyme concentrations
(ng/midgut) of larvae and pupae of Lutzomyia
anthophora. . .. 133

4-7. Time course of the changes in trypsin- and
chymotrypsin-like midgut equivalents
(ng/midgut) of 3-day-old sugar fed and



3-day-old blood fed females of Lutzomyia
anthophora at different hours after blood
feeding. . . .. 135

4-8. Time course of the changes in trypsin ratio
compared with other serine proteases (% per
midgut) of larvae and pupae of Lutzomyia
anthophora. . .. 137

4-9. Time course of the changes in trypsin ratio
compared to total serine proteases midgutgut)
of 3-day-old blood fed Lutzomyia anthophora
females . . 138

4-10. PAGE Fluorography of [1,3-3H]DIP-trypsin- and
-chymotrypsin-like isozymes of first and
second instar larvae of Lutzomyia
anthophora . . 140

4-11. PAGE Fluorography of [1,3-3HIDIP-trypsin- and
-chymotrypsin-like isozymes of second-fourth
instar larvae of Lutzomyia anthophora. 141

4-12. PAGE Fluorography of [1,3-3H]DIP-trypsin- and
-chymotrypsin-like isozymes of the fourth
instar larvae of Lutzomyia anthophora. 142

4-13. PAGE Fluorography of [1,3- H]DIP-trypsin- and
-chymotrypsin-like isozymes of the pupae of
Lutzomyia anthophora. . ..145

4-14. PAGE Fluorography of [1,3-3H]DIP-trypsin- and
-chymotrypsin-like isozymes of larvae, pupae
and females of Lutzomyia anthophora. 146

4-15. PAGE Fluorography of [1,3- 3H]DIP-trypsin- and
-chymotrypsin-like isozymes of blood fed
females of Lutzomyia anthophora. 147

4-16. PAGE Fluorography of [1,3-3H]DIP-trypsin and
-chymotrypsin-like derivatives from blood fed
female Lutzomyia anthophora. ... 148

5-1. Diagrammatic illustration of dorsal view of
stomatogastric nervous system and associated
endocrine gland in phlebotomine sand flies. .158

5-2. In vitro incubation of head-thorax complex
of Lutzomyia anthophora for the biosynthesis
of JH III. . .160



5-3. C1, Reversed phase HPLC of JH III, [12-3H]JH
III, [12-3H]methyl farnesoate and JH I. .163

5-4. A typical example of C18 Reversed phase HPLC
of [12-3H]JH III, JH III, JH I and
[12-3H]methyl farnesoate. .. ... 164

5-5. Time course of in vitro biosynthesis of
[12-3H]JH III in the life of female Lutzomvia
anthophora. . . ..166

6-1 Map of Colombia showing locations of Arboledas
and Durania .... . .174

6-2. Rainfall in Arboledas (Colombia) during 1985. .177

6-3. Morphology of the female reproductive system
Lutzomyia anthophora .. 181

6-4. Different parts of female reproductive system
of Lutzomyia anthophora. . 183

6-5. Different stages of ovarian follicles of
Lutzomyia anthophora. . ... .186

6-6. Different Stages of ovarian dilatation
formation in Lu. anthophora. ... 187

6-7. Temporal changes in the reproductive status of
Lu. cayennensis collected resting on tree
trunks in Durania, Colombia during January
1987 to January 1988 . 191

6-8. Temporal changes in the ovarian development
stages of Lu. cayennensis collected resting on
tree trunks in Durania, Colombia during
January 1987 to January 1988. ... 191

6-9. Temporal changes in the reproductive status
of Lu. ovallesi collected resting on tree
trunks in Arboledas, Colombia during May 1985
to December 1986. . 196

6-10. Temporal changes in the ovarian development
stages of Lu. ovallesi collected resting on
tree trunks in Arboledas, Colombia during May
1985 to December 1986. . 196

6-11. Temporal changes in the reproductive status of
Lu. ovallesi collected resting on tree trunks




in Durania, Colombia during January 1987 to
January 1988. .. ...... .. .198

6-12. Temporal changes in the ovarian development
stages of Lu. ovallesi collected resting on
tree trunks in Durania, Colombia during
January 1987 to January 1988 .. .198

6-13. Temporal changes in the reproductive status
of Lu. serrana collected resting on tree
trunks in Arboledas, Colombia during November
1985 to September 1986. ... .. 204

6-14. Temporal changes in the ovarian development
stages of Lu. serrana collected resting on
tree trunks in Arboledas, Colombia during
November 1985 to September 1986 ..204

6-15. Temporal changes in the reproductive status
of Lu. shannoni collected resting on tree
trunks in Arboledas, Colombia during August
1985 to September 1986. .... ..209

6-16. Temporal changes in the ovarian development
stages of Lu. shannoi collected resting on
tree trunks in Arboledas, Colombia during
August 1985 to September 1986. ... 209

6-17. Temporal changes in the reproductive status
of Lu. shannoni collected resting on tree
trunks in Durania, Colombia during January
1987 to January 1988 . .... 211

6-18. Temporal changes in the ovarian development
stages of Lu. shannoni collected resting on
tree trunks in Durania, Colombia during
January 1987 to January 1988 .. 211

6-19. Temporal changes in the reproductive status
of Lu. spinicrassa collected resting on tree
trunks in Arboledas, Colombia during October
1985 to September 1986 .. .. .. 218

6-20. Temporal changes in the ovarian development
stages of Lu. spinicrassa collected resting on
tree trunks in Arboledas, Colombia during
October 1985 to September 1986. .. .218

6-21. Temporal changes in the reproductive status
of Lu. spinicrassa collected resting on tree



trunks in Durania, Colombia during January
1987 to December 1987. . .220

6-22. Temporal changes in the ovarian development
stages of Lu. spinicrassa collected resting
on tree trunks in Durania, Colombia during
January 1987 to December 1987. ... 220

6-23. Temporal changes in the reproductive status
of Lu. trinidadensis collected resting on
tree trunks in Durania, Colombia during
January 1987 to January 1988 . .229

6-24. Temporal changes in the ovarian development
stages of Lu. trinidadensis collected
resting on tree trunks in Durania, Colombia
during January 1987 to January 1988 .. .229


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




August 1990

Chairman: Dr. D. G. Young
Cochairman Dr. D. Borovsky
Major Department: Entomology and Nematology

Different biological aspects of the reproduction of Lu.

anthophora were investigated. Longevity of different

Lutzomyia species from a focus of leishmaniasis in the

Department of Norte de Santander, Colombia, was determined.

Maximum number of 1-day-old females was inseminated by

4-day-old males. Males became sexually mature 13 to 24 hrs

after emergence and two-day-old males inseminated the

maximum number of females. Male reproductive organs became

smaller due to aging and multiple matings. The males

accessory gland contained 2 types of secretions.

Trypsin- and chymotrypsin-like enzymes were

quantitatively determined in immature and adult sand flies.

More chymotrypsin-like enzymes were present in the larvae,


newly emerged, sugar fed and females 72 hrs after blood

feeding. Blood fed females had more trypsin-like enzymes.

Twelve, 8, 4, 4, 5 and 11 different isozymes were secreted

by larvae, early pupae, late pupae, newly emerged females,

sugar fed females and blood fed females respectively. Eight,

1, 1 and 4 isozymes were trypsin-like in larvae, early

pupae, late pupae and blood fed females respectively and the

rest were chymotrypsin-like isozymes.

Biosynthesis, of JH III by the corpora allata in vitro

was analyzed by high pressure liquid chromatography and gas

chromatography. Peak JH III biosynthesis occurred 24 hrs

after emergence. The rate of JH III biosynthesis declined

in 3-day-old sugar fed females and immediately following

blood feeding. The biosynthesis however, peaked at 4 hrs.

A rapid decline occurred after 4 hrs and the minimum rate of

biosynthesis was found 30 hrs after blood feeding. The rate

of JH III biosynthesis increased at 38 hrs after blood

feeding and the maximum biosynthesis was after 96 hrs.

The females were capable of leishmania transmission

throughout the year and completed 1-4 gonotrophic cycles.

Lu. spinicrassa, a probable vector of Le. brazilensis

completed 2 gonotrophic cycles from May-December.



Medical Importance of Sand flies

Sand flies are the sole vectors of Leishmania, a group

of parasites that causes a number of enzootic and zoonotic

diseases collectively referred to as leishmaniasis.

According to a report of WHO (1984) this protozoan pathogen

annually torments more than 400,000 people worldwide. Human

leishmaniasis is caused by at least 14 different species of

the genus Leishmania. Sand flies are widely distributed and

their ranges extend from approximately 500 N to 400 S. In

the Old World, leishmaniasis is transmitted by sand flies

belonging to the genus Phlebotomus Rodani and Bert. It's

distribution is mostly temperate and encompasses areas such

as Central Asia, India, North Africa, and Mediterranean

Europe. In the New World, leishmaniasis is transmitted by

sand flies of the genus Lutzomyia Franca and its

distribution is mostly tropical except for few autochthonous

cases that were recorded in Texas (Lawyer 1984).

Sand flies are also vectors of other diseases. In the

New World, a bacterium, Bartonella bacilliformis is commonly

referred to as Carrion's disease that has two clinical


forms, Oroya fever and Verruga peruana. The disease has a

limited distribution, having been recorded from Peru,

Ecuador, and Colombia. Lutzomyia verrucarum (Townsend) is

the probable vector (Hertig 1939).

Arboviruses belonging to three families (Rhabdoviridae,

Reoviridae, and Bunyaviridae) are transmitted by sand flies

(Tesh et al. 1971b; Tesh et al. 1987; Tesh 1988; Tesh et al.

1989). Transovarian transmission of Phleboviruses has been

shown for three Lutzomyia species (Endris et al. 1983;

Jennings & Boorman 1980; Tesh & Modi 1983). Viruses of

seven New World serotypes were isolated from male sand flies

(Tesh 1988), and Phlebovirus was also isolated from both

sexes of sand flies in Panama (Tesh et al. 1974). In the

Old World, viruses belonging to the above three families

have been isolated from Phlebotomus sand flies. The best

known disease in Old World is papatacci fever, or sand fly

fever, or 3-day fever reported from India, China, Central

Asia, and the Mediterranean region.

American Leishmaniases

A large number of Leishmania species coexist in the

Americas, and together with these a number of

morphologically similar nonleishmanial trypanosomatids

parasites occur in many localities (Young & Lawyer 1987).

This coexistence complicates the studies of vector

implication of different sand flies in an area. The


situation is further complicated due to the presence of

certain host specific mammalian leishmaniasis species (Le.

hertigi of porcupines) that have never been reported to

infect humans (Young & Lawyer 1987). The occurrence of

sympatric morphospecies of Lutzomyia in certain foci of

leishmaniasis further complicates the situation (Young &

Lawyer 1987).

In the New World two clinical forms of leishmaniasis,

visceral and cutaneous are often recognized, but the

symptoms alone do not indicate the identity of the

parasite. In certain types of visceral leishmaniasis,

dermal lesions have been observed. The genus Leishmania is

divided into two sub-genera, Leishmania and Viannia

according to their mode of growth and other parameters. The

subgenera correspond to two sections, Peripylaria and

Suprapylaria (Young & Lawyer 1987). The section Peripylaria

contains Le. braziliensis braziliensis, Le. b. quyanensis,

Le. b. panamensis, Le. b. peruviana and species of

Leishmania from armadillos. Section Suprapylaria contains

Le. donovani chagasi, Le. hertigi hertigi, and Le. h. deanei

(from tree porcupines in the neotropics). Also included are

Le. mexicana mexicana, Le. m. aristedesi (from rodents), Le.

m. enreittii (From domestic guinea pigs), Le. m. qarnhami,

Le. m. venezulensis, Le. m. subspecies from Matto Grosso

State, Brazil, Le. m. subspecies from Minas Gerais State,

Brazil, and different Leishmania subspecies from the


Dominican Republic. The distribution and vectors of all

these species were reviewed by Young and Lawyer (1987).

Effects of Leishmania Infection on Sand Flies

Recent improvements in the rearing techniques of sand

flies (Young et al. 1981; Endris et al. 1982) have enabled

many scientists to experimentally infect sand flies with

different species of Leishmania and study their course of

development. Sand flies are pool feeders and their mouth

parts and digestive system are described in chapter 2. Sand

flies that feed on the periphery of the leishmaniasis lesion

ingest the parasite in a form that is called an amastigote.

Since sand flies are pool feeders, they can ingest only a

few amastigotes with their blood meal. These amastigotes

are then released due to the rupture of haemocytes during

feeding perhaps by the cibarial teeth (Lewis 1975). The

number of amastigotes ingested by sand flies partly depends

on the length of their mouth parts (Young & Lawyer 1987).

Sand flies inflict injury to the host by lacerating the

tissues. This damage may attract macrophages in the

vicinity of the feeding site, thus increasing the chances

for the parasite to reach its site of extrinsic development

in the sand fly midgut.

Reptilian Leishmania and Trypanosoma are capable of

thriving in their sand fly vectors even when they are

ingested with nucleated reptile blood (Adler 1964).


Although human and mammalian blood causes mortality in

Leishmania cultures, this effect is immediately removed

after the ingestion of the parasite by the sand flies (Adler

& Theodor 1930). Human serum can inhibit the rate of

trypsin production in the midgut of sand flies. Low rate of

blood digestion is favorable for the growth of Leishmania

(Borovsky & Schlein 1987).

After reaching the sand fly midgut, the amastigotes are

released from the macrophages and are now present inside the

peritrophic membrane where they undergo either one or two

divisions and transform into flagellate forms,

promastigotes. The rate of formation of the peritrophic

membrane may be directly involved in the susceptibility or

refractoriness of sand flies to various parasites because

leishmanial promastigotes cannot penetrate it (Killick-

Kendrick 1979).

The Old World species of Leishmania can be transmitted

only by their particular host sand flies. In Lu. abonnenci

(Floch & Chassignet) females, fed on mice infected with L.

mexicana mexicana, the parasites were embedded in the

peritrophic membrane as early as 18 hrs after a blood meal

(Walters et al. 1987). The first division of the parasite

occurs within the first 24 hrs after the ingestion of an

infective blood meal (Strangeway Dixon & Lainson 1966).

These actively dividing parasites lie within the bag of the

peritrophic membrane (Leany 1977). Thus, it may be possible


that rate of formation of the peritrophic membrane varies in

different species of sand flies. In some species it acts as

a barrier to massive transfer and establishment of the

promastigotes in the microvilli of the midgut epithelial


The nature of the peritrophic membranes of sand flies

also affects their susceptibility to Leishmania. Feng

(1951) studied the peritrophic membrane in Ph. chinensis

(Newst), Ph. mongolensis (Sinton), and Sergentomyia

squamirostris (Newstead). He postulated that in poor hosts

of Leishmania such as Ph. mongolensis, the peritrophic

membrane remained intact and the parasites passed out in

feces within it. On the other hand, in susceptible species

such as Ph. chinensis the peritrophic membrane was broken

three days after a blood meal. Thus, leishmanial

promastigotes were able to escape and establish an infection

in the anterior midgut. The transformation of amastigotes

to promastigotes takes 2-3 days (Molyneux & Killick-Kendrick

1987). The promastigotes develop flagellar expansion or

hemidesmosomes that help to attach them to either microvilli

of midgut or cuticle of the pylorus or ileum and transform

into paramastigotes. Paramastigotes undergo an extensive

multiplication before migrating to the esophagus and pharynx

and transform into promastigotes. Later the promastigotes

migrate to the proboscis.


Free floating flagellates of Leishmania mexicana were

observed in the lumen of the Malpighian tubules and the

pylorus of field collected Lutzomyia olmeca (Fairchild &

Theodor)(Williams 1970) and also in many laboratory infected

sand flies (Walters et al. 1987; Christensen & Herrer 1980).

The Malpighian tubules may serve as sites providing

nutrition to the parasites after the complete digestion of

the blood meal (Christensen & Herrer 1980; Lawyer 1984;

Walters et al. 1987).

At the completion of the extrinsic development of the

Leishmania parasites, small actively moving forms are found

within the mouth parts and the different parts of the

midgut. These small forms may attach to the chitin or may

occur free in the pharynx or in the mouth parts of the sand

flies (Killick-Kendrick 1979). The attachment of the

parasites in the mouth parts results in impaired function of

the cibarial receptors (Killick-Kendrick et al. 1977b).

Infected Lu. lonqipalpis females take a very small

subsequent blood meal or sometimes are unable to feed at all

(Adler & Bur 1941). Such flies transmitted maximum number

of L. mexicana amazonensis infections (Killick-Kendrick et

al. 1977b).

The salivary gland lysate of Lu. lonqipalpis (Lutz &

Neiva) possesses a property of enhancing the extent of

cutaneous infection by Leishmania major in BALB/C mice,

which are genetically susceptible to Leishmania infection


and CBA mice which are genetically resistant to Leishmania

infection. Injection of Leishmania major together with

salivary lysate results in cutaneous lesions which are five

to ten times as large and contain as much as 5,000 times

more parasites than the controls. In inocula that contains

smaller number of parasites, the parasites are detected in

the lesion only when they are injected together with the

salivary homogenates. The lysate is capable of producing an

enhancing effect in quantities as small as 10% of the total

gland homogenate (Titus & Riberio 1988).

Many methods of Leishmania transmission are suggested.

The first mode of transmission is by the bite of an infected

sand fly. The second method known as "blocked fly" theory

suggests that in infected sand flies the leishmanial

parasites infecting the stomodeal valve can be back washed

into the feeding site by the blood meal. This happens due

to the blockage of the normal blood flow in the posterior

midgut after the congestion of the stomodeal valve by the

parasites (Jefferies et al. 1986). The concept of blocked

flies is further supported by the presence of gel-like

matrix in the cardia and stomodeal valve regions of the

infected Lu. abonnenci (Floch & Chassignet) and Lu.

diabolica (Hall)(Lawyer 1984; Walters et al. 1987). Few

enzymes are found in the saliva of hematophagous insects.

Presence of salvia is necessary for the intake of a blood

meal in pool feeders such as tsetse flies compared to


capillary feeders like mosquitoes (Benjamini & Feingold


Sugar is an essential component of the diet of sand

flies (Kirk & Lewis 1951; Lewis & Domoney 1966; Chaniotis

1974; Young et al. 1980). Its presence in the diet of sand

flies enhances their chance of transmitting leishmaniasis to

susceptible hosts (Smith et al. 1940, 1941; Swaminath et al.

1942; Short 1945; Killick-Kendrick et al. 1977b; Young et

al. 1980). Presence of sugars in the crop of infected flies

also enhance the rate of development of Leishmania (Killick-

Kendrick 1978).

These sugars also act as food for the leishmanial

parasites present in the foregut, which is lined with

relatively impermeable cuticle (Davis 1967). Motile

flagellates of Leishmania mexicana mexicana were

occasionally seen in the crop of Lutzomyia abonnenci

(Walters et al. 1987). Similarly, Hertig and McConnell

(1963) also reported Leishmania promastigotes in the crops

of Lutzomyia sanquinaria (Fairchild & Hertig).

Life Cycle of Phlebotomine Sand Flies

A newly emerged female usually seeks a blood meal 12-24

hrs after emergence. Males usually emerge earlier than

females and are ready to mate after one day. Mating of sand

flies may take place at the breeding site or on or near a

host (Chaniotis 1967). Males usually outnumber females at


the resting sites (Alexander 1988). The male to female

ratio observed in the colony cages is mostly 1:1; perhaps in

nature females move to blood feeding sites shortly after

mating. Females can complete 1 to 4 gonotrophic cycles in

the laboratory (Ward 1977; Chaniotis 1986) and also in

nature (see chapter 6). The number of eggs that develop

during one gonotrophic cycle depends upon the protein

content of the blood meal (Ready 1979). In laboratory

colonies of Lu. trapidoi (Fairchild & Hertig), survival of

females increased in the presence of high male to female

ratios during oviposition and females completed upto 3

gonotrophic cycles (Chaniotis 1986). Females that laid eggs

within 4 days after blood feeding laid large number of eggs

and survived longer than those that laid 9 days after blood

feeding (Chaniotis 1986).

Females that take a sugar meal before blood feeding

develop more eggs than those that are starved before a blood

meal (Ward 1977). In nature some species are autogenous and

lay their first batch of eggs without blood feeding (Johnson


Phlebotomine sand flies deposit their eggs in the

detritus and the eggs hatch after about 8 days in non-

diapausing species. Larvae feed on organic detritus, dead

leaves, and fungi (Johnson & Hertig 1961). The first instar

larvae are whitish in color, lack legs, and possess a pair

of caudal setae. The first instar larvae molt after 5 or


more days to second instar. The second instar molts to

third instar, fourth instar and finally to pupae. The whole

growth takes from 15-30 days.

Sand flies diapause in the larval stage and it is often

due to changes in day length and temperature. In New World

species diapause in larvae is induced due to insufficient

humidity at the start of the dry season or may occur in the

egg stage (Hanson 1968). Pupae eclose to adult sand flies

after 8 or more days depending on temperature and sand fly

species. No information is available about the occurrence

of circadian rhythms during adult emergence.


The present investigation was carried out to obtain

information about some important aspects of sand fly

biology. These includes the following studies:

1) The mating behavior of males and females and the

effects of age and matings on the morphology of male

reproductive system.

2) Production of trypsin- and chymotrypsin-like

isozymes was investigated in both immature and adult Lu.

anthophora. Poor information is available about Trypsin-

and chymotrypsin-like isozymes of adult sand flies (Schlein

et al. 1983; Schlein & Romano 1986; Borovsky & Schlein 1987)

and no information is available about the enzymes in

immature sand flies.


3) Juvenile hormone III (JH III) production was

investigated in female Lu. anthophora. Previous studies on

mosquito physiology has shown that some of the most

important aspects of an insect's life, such as host seeking

behavior, mating, and vitellogenesis are controlled by

juvenile hormones (Meola & Petralia 1980; Borovsky 1984;

Meola & Readio 1987). The recent advances in tissue culture

techniques of corpora allata (Borovsky unpublished) greatly

facilitated the in vitro production of JH by the corpora

allata of unfed, sugar fed and blood fed female Lu.


4) All the above topics are important features of egg

development. Thus, egg development was studied in Lutzomvia

anthophora (Addis). Finally, the most important aspect that

results from the above processes is oviposition and survival

of sand flies in nature. Eight different Lutzomyia species,

collected during 1985-1988 from two leishmaniasis foci,

Arboledas and Durania from Norte de Santander in Colombia,

were age graded according to the methods of Detinova (1962).

Their longevity in relation to Leishmania transmission is



Reproductive Biology of Sand Flies

Reproduction, the most important aspect of an

insects's life, is an intricate process. It encompasses

many other aspects such as larval nutrition, environmental

factors, mating, host finding, feeding, egg development and

oviposition. Each one of the above processes is an entity

in itself and is comprised of a network of behavioral and

physiological events.

Mating consist of behaviors like courtship or epigamic

behaviors that bring the two sexes together and results in

the insemination of the female. After mating, host finding

behavior is invoked. In some mosquito species, release of

juvenile hormone (JH) is the trigger for host seeking

behavior (Meola & Petralia 1980; Meola & Readio 1988),

whereas in others, JH does not seem to play a part in host

seeking behavior (Lea 1963; Bowen & Davis 1989).

Certain sand fly species develop their first clutch of

eggs without a blood meal autogenouss), whereas others

require a blood meal (anautogenous) for the completion of

first and later gonotrophic cycles. Rate of blood digestion

depends on factors such as environmental temperatures (Weitz

& Buxton 1953; Yang & Davis 1968; Honda et al. 1985),

presence or absence of parasites in the blood meal (Borovsky

& Schlein 1987), and mating (Edman 1970; Downe 1975; Adlakha

& Pillai 1976a; Houseman & Downe 1986).

In female mosquitoes and other haematophagous

insects, different consequences of mating, alteration of

blood meal size (Adlakha & Pillai 1976a), stimulus for

oviposition (Leahy & Craig 1965), and initiation of a

refractory period to further mating (Young & Downe 1982,

1983) have been observed. Blood proteins are utilized for

the development and maturation of eggs. Presence of JH

makes the ovaries receptive to egg development

neurosecretory hormone (EDNH) and induces responsiveness in

the fat bodies to 20-hydroxyecdysone (Flanagan & Hagedron

1977; Shapiro & Hagedron 1982).

The reproductive biology of haematophagous insects

represents a network of processes that begins from larval

nutrition and exposure to different environmental conditions

and ends after oviposition. In autogenous species of

mosquitoes the number of eggs laid by a female is directly

dependent on the larval nutrition, whereas in anautogenous

species it also depends on the size of the blood meal and

the species of the host. Many autogenous species of

mosquitoes and sand flies require a blood meal for egg

development during the second gonotrophic cycle.







Figure 2-1. Male reproductive system of Lutzomyia sauroida


Morphology of the Male Reproductive System

The male reproductive system consists of a pair of

rounded, oblong, or oval testes, a large pear-shaped,

conical or almost spherical vesicula seminalis, the sperm

pump, and the sperm tubes (Perfil'ev 1968; Sherlock &

Carneiro 1964). The vasa efferens vary in size in different

species of sand flies and open into the vesicula seminalis

by pressing part of its wall inward (Fig. 2-1). The

vesicula seminalis opens into a narrow ejaculatory duct

which leads to the genital pump (sperm pump). The genital

pump is a syringe-shaped organ having two long sperm tubes

or filaments arising from it (Sherlock & Carnerio 1964;

Downes 1968). The filaments lie in the grooves of the

aedeagus where they can move freely. The filament emerges

from the apex of the aedeagus, or slightly proximal to it

depending on the shape of the aedeagus.

In Phlebotomus papatasi (Scopoli) the filaments are

short in length and are only inserted into the tip of the

spermathecal orifices. In other species the filaments are

longer and are inserted well inside the spermathecal ducts,

sometimes reaching into the spermathecae (Sinton 1925). In

species having common spermathecal opening, the filaments

remain paired; e.g., in Phlebotomus melloni (Sinton) the

filaments or the aedeagal ducts remain paired and are

inserted together in the spermathecal duct and at the apex


the filaments are inserted into the separate spermathecae

(Sinton 1932).

Sherlock and Carnerio (1964) described the male

reproductive system of different sand flies and emphasized

that the differences observed in fresh specimens could be

used to identify the different species. They described the

male reproductive system as having two testes, two efferent

tubules, one seminal vesicle, one deferent tubule

(= ejaculatory duct of Perfil'ev 1968), one ejaculatory pump

(= genitial pump of other authors). They noted that

spermatozoa were cane shaped. During mating spermatozoa

passed through the seminal vesicles and were deposited

together with the seminal liquid into the female

spermathecae. In males which had not mated the spermatozoa

were in the form of balls in each testis.

Dallai et al. (1984) studied the evolution of sperm in

the Phlebotominae and their relation to other Nematoceran

Diptera. The spermatozoa of Ph. papatasi (Scopoli) are

thread like, 13 um long and 0.5 jm wide, head is 2.5 uim long

with a compact electron-dense fusiform nucleus and chromatin

is absent in its anterior end. The mature spermatozoon has

a longitudinally arranged distal centriole that is continued

into a 10 pm long axoneme. The axoneme has a "9 + 9 + 0"

arrangement of the microtubules. The axonemes are motile

inside the spermathecae. The structure of the tail part is

similar to other insects.



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Digestion of Food by Sand Flies

Digestive System of Phlebotomine Sand flies

Mouth parts. Sand flies have a piercing sucking

fascicle or syntrophium (Jobling 1976), and an archaic type

proboscis (Lewis 1974). The proboscis extends into the

buccal cavity or cibarium (Fig. 2-2). Variable number of

horizontal, vertical, and lateral teeth are present in the

cibarium of Lutzomyia species. From the ventral aspect, the

horizontal or hind teeth have either pointed or blunt tips

(Young 1979). The vertical teeth are distal to the

horizontal teeth and sometimes appear as dark dots. The

lateral teeth occur next to horizontal teeth on both sides

of the cibarium. Which connects posteriorly to the pharynx

and lies within the clypeus (Young 1979).

Lutzomyia sand flies that feed on mammals have long

labra with hooked teeth. These teeth are also present on

the paired maxillae. In comparison, most species that feed

on reptiles have ridged tipped maxillae (Lewis 1975). Two

types of sensillia are present on the mouth parts of sand

flies. The first type occurs on the proboscis; the second

type is on the cibarium (Killick-Kendrick & Molyneux 1981;

Lewis 1984).

An aggregation pheromone occurs in the maxillary palps

of Ph. papatasi. The response of sand flies to this







Figure 2-3. Digestive system of Lu. anthophora.





pheromone is olfactory and temperature dependent, suggesting

it is volatile in nature (Schlein et al. 1984).

Sand flies have a pair of salivary glands situated at

the junction of the head to the thorax (Jobling 1987; Lewis

1974) (Fig. 2-2).

Forequt. The alimentary tract of sand flies has three

major regions, foregut, midgut, and hindgut (Fig. 2-2). The

foregut consists of a proboscis, cibarium, pharynx, crop,

oesophagus, and stomodeal valve. The pharynx is divided

into an anterior and a posterior pharynx. The latter has an

armature that is composed of three chitinous plates. Most

Lutzomyia species have transverse unarmed ridges on this

region, and in few species it is armed with spines (Young

1979). The pharynx opens into the oesophagus, which is

connected via a stomodeal valve to a small wide slightly

bulbous portion, the cardia or cardiac stomach (Gemetchu

1974; Walters et al. 1987) (Fig. 2-3). The stomodeal valve

in Phlebotomus longipes (Parr) is formed by the protrusion

of the oesophagus as a thick ring of elongated cells into

the lumen of the cardia. On the hemocoelic aspect, this

ring is covered with fine muscle strands.

Crop. The crop is ballon shaped and opens at the

junction of the oesophagus to the cardia (Figs. 2-2 & 2-3).

Sugar solutions imbibed by the sand flies go into the crop

where they play an important role in maintaining energy for

the growth of Leishmania parasites.


Midqut. The midgut is divided into an anterior and a

posterior midgut. The cardia is the first part of the

anterior midgut and is continuous with the thoracic portion

of the anterior midgut. Both the cardia and anterior midgut

are found between thoracic muscles (Jobling 1987; Lewis

1974; Walters et al. 1987) (Figs. 2-2 & 2-3). Some authors

consider the whole anterior midgut lying in the thoracic

muscles as the cardia (Adler & Theodor 1926; Hertig &

McConnell 1963), whereas, Davis (1967) considers the

slightly bulbous area following oesophageal-crop junction as

the cardia (Fig. 2-3). The cells of the anterior midgut are

cuboid, with striated borders or microvilli (Gemetchu 1974).

The anterior midgut is continuous with the posterior midgut.

Ultrastructure of the posterior midgut was described

for two species of sand flies (Gemetchu 1974; Rudin & Hecker

1982). Ultrastructure of the posterior midgut of Lu.

longipalpis (Lutz & Nieva) shows distinct differences

between sugar fed and blood fed females (Rudin & Hecker

1982). The midgut of sugar fed female shows a single

layered epithelium of highly polarized elongated cells of

relatively small size compared to cells of blood fed

females. An oval nucleus, 10% of the cell volume, occurs in

the center of the cell. Triglyceride-like structures are

found around the nuclear region in association with the

myelinated bodies and lysosome like structures. About 15%

of the cell cytoplasmic volume contains randomly distributed


mitochondria. Free ribosomes are present throughout the

cells. Smooth endoplasmic reticulum occurs in association

with the golgi bodies that are regularly distributed

throughout the cells. Various amount of vesicles,

cisternae, and whorls of rough endoplasmic reticulum are

present in the cells and accounts for 5.5 m2/ m3 of the


The epithelial cells show densely packed microvilli,

which are about 1 mm long and make up to 8% of the

cytoplasmic volume in Lu. longipalpis. The cells also show

a cytoskeleton composed of many microtubules and

microfilaments. Cells of other haematophagous insects also

have a large amount of endoplasmic reticulum and enzyme

synthesis may occur there. The endoplasmic reticulum

involved in enzyme synthesis is secluded from the rest of

the cell cytoplasm to prevent it from auto-digestion

(Richards 1975).

A thin basal lamina separates the stomach epithellium

of Lu. lonqipalpis from the hemocoel. Gaps and septate

junction are present between the lateral cell walls. Only

few regenerative cells are present. The midgut is

surrounded with longitudinal and circular muscles (Rudin &

Hecker 1982). The inner surface of the midgut is devoid of

cuticle but this does not mean that inner midgut epithelium

is directly exposed to the blood meal. An acellulary


secreted peritrophic membrane is present between the midgut

epithelium and the blood meal.

A brush border formed of microvilli is present in both

the anterior and posterior midgut portions of Ph. longipes

and Ph. sergenti (Parr)(Perfil'ev 1928; Gemetchu 1974),

while it is present only in the anterior midguts of Ph.

orientalis (Pass) and Ph. papatasi (reference not seen, but

referred in Gemetchu 1974).

In unfed insects, the midgut cells are inactive and the

ribosomes are localized on balls, whorls or sheets of rough

endoplasmic reticulum (Gemetchu 1974; Richards 1975). The

midgut epithelium of Ph. longipes and Lu. lonqipalpis

becomes flattened and the whorls of the endoplasmic

reticulum open after the ingestion of blood meal. Mean

volume of microvilli, rough endoplasmic reticulum, and

smooth endoplasmic reticulum of the midgut epithelial cells

of Lu. lonqipalpis increases within 24 hrs after the

ingestion of a blood meal. Some of the parameters described

above decrease 2 days after ingestion of a blood meal (Rudin

& Hecker 1982). Under different physiological conditions,

similar changes occur in the midgut structure of Ph.

longipes (Gemetchu 1974).

Electron microscopical studies of the midgut of

mosquito Aedes aegypti (L) show that it has three types of

cells, a. digestive cells (Hecker et al. 1971), b. endocrine


cells (Brown et al. 1985) and c. regenerative cells (Hecker

et al. 1971).

The midguts of mosquitoes show changes similar to those

observed in sand flies before and after the ingestion of a

blood meal. In unfed female Ae. aeQvyti the digestive cells

of the midgut epithelium contain no secretary granules

(Hecker et al. 1971). In blood fed mosquitoes, the surface

of the rough endoplasmic reticulum greatly increases by the

opening of its tight whorls, a condition existing in sugar

fed and/or starved mosquitoes (Hecker et al. 1971; Hecker &

Brun 1975; Bauer et al. 1977). The secretary granules

appear in increasing number 8 and 24 hrs after a blood meal

(Rudin & Hecker 1979).

The above observations are also supported by a recent

report in which free and membrane-bound ribosomes of

intestinal cells of female Ae. aegypti were quantified

during the digestive cycle (Gander et al. 1980). Free

ribosomes increase up to 24 hrs after blood feeding, reach a

plateau and decline to the unfed level after 96 hrs.

Similarly, membrane-bound ribosomes increase up to 36 hrs

and decline to the unfed level after 96 hrs (Gander et al.

1980). Increase in the amount of rough endoplasmic

reticulum after a blood meal is thought to be due to the

production of trypsin-like enzymes (Rudin & Hecker 1979).

Contrary to previous reports, in mosquito Anopheles


stephensi (Liston) secretary granules were found before the

intake of a blood meal (Berner et al. 1983).

In Ph. lonqipes some increase in the vacuolation of the

cytoplasm and in the labyrinth spaces of the basal membrane

occurs after the ingestion of a blood meal. This increase

is probably due to the transport of digestion products

across the epithelial cells and through the basal membrane

to the haemocoel (Gemetchu 1974).

Peritrophic membrane. Hematophagous insects, except

fleas, synthesize a peritrophic membrane after ingesting a

blood meal (Richards & Richards 1971).

Peritrophic membrane is absent in unfed and sugar fed

female sand flies (Gemetchu 1974). After blood feeding

development of the peritrophic membrane is initiated within

30 minutes after ingestion of a blood. A secretion, present

between the microvilli within 12 hrs post feed, extends to

the microvilli tips and reaches into the lumen of the

midgut. A thin sac of this secretion is present around the

blood bolus within 24 hrs post feed. This sac gradually

thickens and becomes stronger. After 48 hrs the peritrophic

membrane can be dissected away from the midgut epithelium.

It is present mainly in the midgut but a small extension

continues in the posterior end of the foregut and this

portion is much folded (Gemetchu 1974).

Two and three days after ingestion of a blood meal, the

digested blood, now dark brown, is present in the form of a


rod in the anterior part of the peritrophic membrane.

Similar rod shaped particles are also present inside the

blood meal and in the ectoperitrophic space.

The ectoperitrophic space occurs outside the

peritrophic membrane and inside the midgut epithelium. The

peritrophic membrane starts to break down at its anterior

end at about the third day. It breaks around the blood meal

by the fourth and fifth day. Sand flies defecate the

undigested blood and remaining peritrophic membrane on the

sixth or seventh day after blood feeding (Gemetchu 1974).

If a partially blood fed Ph. longipes refeeds, the

second blood meal envelopes the first peritrophic membrane

containing the first blood meal. This results in a slower

rate of disintegration of the first peritrophic membrane.

The rate of digestion of the first blood meal is also slower

and may be due to its separation from the midgut epithelium

by the second blood meal (Gemetchu 1974).

The electron microscopy of the peritrophic membrane

just after engorgement reveals an irregular lattice like

structure and a network of fibrils. The peritrophic

membrane, rich in proteins and some of the hexoses, is

insoluble in water, alcohol and KOH (Gemetchu 1974).

The process of peritrophic membrane secretion may

involve the interaction of proteolytic secretions at their

interface with the blood mass. This process may become weak


as blood digestion proceeds to its completion (Gemetchu


Thickness of the peritrophic membrane varies in

different species of insects and also in different instars

of the same species. It may be a granular sheath or it may

have layers of fibrous material under the granular layer.

The microfibrous layer may be arranged randomly or in order

of hexagonal or orthogonal arrays (Peters 1976).

Insects ingest their food in macromolecular form, such

as polysaccharides, proteins, and lipids (triglycerides,

phospholipids and glycolipids). These large molecules can

not pass the peritrophic membrane and midgut epithelium;

therefore, they are broken down in to small peptides, amino

acids by trypsin-like enzymes. These enzymes are reported

from many larvae and adults of insects, such as stable fly

Stomoxys calcitrans (L), house fly Musca domestica(L),

common blow fly Calliphora vomitoria (L), mosquito Ae.

aegypti, and Culex niqripalpus (Theobald) (Champlain & Fisk

1956; Yang & Davis 1971; Borovsky 1985; Patterson & Fisk

1958; Greenberg & Parestky 1955; Patel & Richards 1960;

Fraser et al. 1961; Gooding 1966a, 1966b; Borovsky 1986).

There is direct evidence that midgut epithelium cells of

mosquitoes produce these proteases (Briegal & Graf 1989). A

high level of proteolytic activity, with specificity to the

trypsin- and chymotrypsin-like substrates, was also detected

in the midgut of detritus-feeding larvae of the crane fly


Tipula abdominalis (Say) (Sharma et al. 1984). Who

suggested that these enzymes were secreted from the midgut

epithelium cells and not from the microbes originating from

the detrital food.

The peritrophic membrane is selectively permeable to

small peptides, amino acids, water, enzyme amylase, and poly

peptide of up to 23,000 dalton molecular weight (Borovsky

1986). Borovsky (1986) proposed a system for blood

digestion in mosquitoes, and wrote the following:

A signal is sent to the midgut epithelial cells to
initiate trypsin synthesis, and trypsin is secreted
into the ectoperitrophic space. The trypsin traverses
the pores in the peritrophic membrane and begins to
digest the blood clot around its periphery, minimizing
exposure to trypsin-inhibitory subfractions. Peptides,
amino acids, and polypeptides, of molecular weight
smaller than 23,000 daltons, diffuse out through the
peritrophic membrane into the ectoperitrophic space.
Poly peptides and peptides that has accumulated in the
ectoperitrophic space are further digested into free
amino acids, which are transported through the midgut
epithelial cells into the hemolymph. The free amino
acids are then used by the fat bodies to synthesize
vitellogenin (egg yolk proteins). pp. 158-159.

The above theory is also supported by studies on Ae.

aegypti that shows 60% of total proteolytic activity occurs

around the peritrophic area, and the rest of the activity is

inside the midgut epithelium and inner to the peritrophic

membrane (Graf & Briegal 1982). This theory is further

supported by a study in which polycolonal antibodies were

developed against trypsin from the midgut of blood fed Ae.

aeqvpti. These antibodies were employed to detect the

trypsin activity in the midgut at different hours after a


blood meal. Trypsin activity was detected in the lumen of

the midgut 8 hrs after blood feeding. The trypsin activity

was localized in the posterior distensible part of the

midgut and was on the periphery of the peritrophic membrane

and the blood bolus. Maximum trypsin activity was detected

24 hrs after a blood meal (Graf et al. 1986).

Secretory granules, labelled with anti-trypsin antibody

and protein A-colloidal gold, were first detected 12 hrs

after the blood meal, by electron microscopy (Graf et al.

1986). At 18 hrs post blood meal secretary pathway was

traced from the cisternae of the golgi body where granules

were formed to their release by exocytosis in the lumen of

the midgut (Graf et al. 1986). Graf et al. (1986) also

suggested that the changes in rough endoplasmic reticulum,

after blood feeding, were not due to production of proteases

as was suggested by many others (Frevogel & Staubli 1965;

Hecker 1977; Berner et al. 1983), but were due to the

formation of peritrophic membrane which is secreted in

greater amount than the proteases.

Hindqut. The posterior midgut opens into the hindgut

via the pyloric valve, which is similar to the stomodeal

valve of Ph. lonqipes (Walters et al. 1987). Four

Malpighian tubules arise at this junction (Figs. 2-2 & 2-3).

Two Malpighian tubules arise together from a common origin

on each side (Jobling 1987). The function of the Malpighian

tubules is excretion of urine.


The hindgut is divided into the anterior intestine and

the posterior intestine. The anterior intestine has three

portions, i.e., pylorus, ileum, and colon (Figs. 2-2 & 2-3).

A pyloric armature has been observed in the New World sand

flies. The armature is spiculate and occupies more than

posterior 1/3 of the pylorus. It is similar in structure to

the pharyngeal armature of several Neotropical species and

has posteriorly directed spines. It may function in the

disruption of the un-digested hematin blood meal residue and

its enclosing peritrophic membrane during the ultimate

passage through the hindgut (Christensen et al. 1971).

The pylorus is connected to ileum, which is a narrow

long tube (Figs. 2-2 & 2-3). The ileum of Ph. papatasi is

surrounded by circular muscles, which in turn are surrounded

by longitudinal muscles. The whole ileum is enveloped on

the outside with a peritoneal sheath (Jobling 1987). The

ileum leads to the rectum, which has a pair of leaf like

rectal papillae and opens to the outside through the anus.

The posterior intestine is subdivided into the rectal

sac containing the rectal papillae and rectum. The inner

surface of the hindgut, like the foregut, is lined with

cuticle that is continuous with the cuticle on the outside

of the animal. The cuticle of the foregut is relatively

impermeable compared to that of the hindgut. Most of the

absorption of nutrients occurs in the midgut. The rectum

opens to the outside by an opening called the anus.

Different Enzymes Secreted during Digestion of Food

Enzymes are organic catalysts that enhance the rate of

chemical reactions without changing themselves. The first

array of digestive enzymes is found in the salivary glands

of the blood-sucking insects. The next enzyme system occurs

in the midgut of insects.

Salivary enzymes and their functions

Sand flies are pool feeders and have very small mouth

parts, which can penetrate only the superficial layers of

skin (Adler & Theodor 1926; Lewis 1975; Jobling 1987). They

feed by lacerating the capillary loops in the superficial

skin layers and ingest the blood pooling into the resulting

hematomas (Riberio et al. 1986). The Old World sand fly Ph.

papatasi possesses salivary anticoagulants which have an

apyrase activity, but lack an erthyma-inducing substance

(EIS) (Alder & Theodor 1926; Riberio 1987). On the other

hand the New World sand fly Lu. longipalpis feeds by

capillary action and possesses antiplatelet activity, an

apyrase, and an erthyma-inducing substance (EIS). The

presence of apyrase and EIS factors, together with the short

mouth parts, helps Lu. longipalpis to feed on the blood

released from the superficial blood capillaries. Treatment

with trypsin or heating for one minute in boiling water


destroys the activity of EIS and suggests that EIS is a

peptide (Riberio et al. 1986).

Crop contents and their function

Gas bubbles of aspirated oxygen are present inside the

crop of mosquitoes and sand flies. The complete absence of

carbon dioxide in them indicates that no fermentation occurs

in the crop (Fisk 1950). Although these bubbles are present

in the crop and also in the midgut of unfed and sugar fed

sand flies (Lewis 1974); no information is available about

their chemical nature. The pH of the crop contents of Culex

pipiens (Say), Ae. aeqypti, and An. quadrimaculatus (Say) is

acidic compared to the basic pH of its midgut (MacGregor


In Lu. lonqipalpis the final destination of the fluid

meal is a function of the method of feeding rather than the

chemical composition of the fluid. Thus, sugar solution

goes directly into the crop and the blood meal, which is

imbibed through a membrane, goes to the midgut (Ready 1978).

In black flies the sugar solution is not routed to the crop

in all flies, in some it goes to the midgut (Disney 1970).

Different enzymes secreted during blood digestion

The pH of the midgut is not easily determined in blood

fed females because mammalian red blood corpuscles have a

buffering capacity which over masks the gut pH, whereas the

midgut epithelium cells remain slightly acidic in nature


(Fisk 1950). No such studies about the pH of midgut and

crop contents of sand flies are available.

Different types of serine proteases are secreted before

and after a blood meal by the midgut epithelial cells.

These proteases are classified by activity and pH optima

(Gooding 1975). Serine proteases are present in mosquitoes

like Ae. aeqypti and Cx. niqripalpus (Graf & Briegal 1985;

Borovsky 1986). To understand their mode of action one has

to answer the following questions: (1) What is the state of

enzyme synthesis prior to the intake of a blood meal? (2)

What initiates the rise in enzyme synthesis? (3) What is the

relationship between the size of the blood meal and the

amount of trypsin- and chymotrypsin-like enzymes secreted?

(4) What is the relationship in the amount of total protein

in the blood meal and the quantity of the secreted enzymes?

(5) What is the effect of blood meal on the changes in the

pH of the midgut? (6) What is the effect of proteolytic

enzyme inhibitors on the rate of blood digestion? and (7)

What effect proteolytic enzymes have on the growth of

endoparasites and how the parasites withstand the changes in

the midgut pH values?

In mosquitoes, ingestion of a blood meal follows within

first 30-35 minutes by rapid diuresis (Boorman 1960;

Williams et al. 1983; Mahmood & Nayar 1989). Several

changes occur in the structure of the secretary cells of the

midgut as described previously. These changes are


accompanied by synthesis of serine proteases by midgut

epithelium. The enzymes found in the midgut are active;

however, synthesis of inactive zymogen that is later

secreted into the midgut and activated there cannot be

discarded. Although there is no example in which insects

secrete an inactive zymogen, the digestive secretary cells

of Ae. aeqvDti contain very few zymogen grannules per cell

compared to vertebrate midgut exocrinee cells) and

trypsinogen might occur in the digestive cells of Ae.

aeqypti (Graf & Briegal 1985).

The midgut of mosquitoes has only one type of secretary

cells and absorptive cells are absent. The function of

absorption is also performed by the secretary cells. They

have well developed microvilli, an elaborate basal

labyrinth, and many mitochondria in the basal and apical

region of the cell. The secretary cells have few secretary

granules per cell (Graf et al. 1986). In Ae. aeqypti both

m-RNA and trypsin are synthesized after the ingestion of a

blood meal (Gooding 1973). Alkaline proteases are detected

4 hrs after blood feeding and reach their peak activity at

24-35 hrs depending on the mosquito species (Gooding 1966a,

1966b, 1975; Briegal & Lea 1975; Borovsky 1986; Graf et al.


The pH optima for inset proteolytic enzymes such as

chymotrypsin, aminopeptidases, carboxypeptidase-A and

carboxypeptidase-B are alkaline. Different substrates and


inhibitors are used to characterize these enzymes (Gooding

1969). Trypsin and chymotrypsin are the major enzymes

present in the midgut of mosquitoes Ae. aegypti, Culex

fatigans (Weid), goat lice Melophaqus ovinus (L), and human

lice Pediculus humanus (L). Both enzymes are absent in the

midgut of blood fed bed bug Cimex lectularius (L), and

kissing bug Rhodinus prolixus (Stal). These insects contain

a high molecular weight protease that is active at pH 5

(Gooding 1969). Gooding (1969) also suggested that insect

trypsin and chymotrypsin probably have serine and histidine

at their active centers. The amount of trypsin secreted

after a blood meal is directly correlated to size of blood

meal (Gooding 1973; Graf & Briegal 1985). A similar

relationship was shown between the size of blood meal and

the amount of total proteases (Shambaugh 1954). The midgut

and gut lumen of tsetse fly Glossina morsitans (Westwood)

shows at least 6 different types of proteolytic enzymes

(Gooding & Rolseth 1976).

The rate of trypsin-like enzyme secretion in certain

hematophagous insects such as horn fly Haematobia irritans

(L) and stable fly S. calcitrans depends on the ambient

temperature (Honda et al. 1985). In blood fed H. irritans

trypsin-like activity is detected earlier at 300C than at

250C. The rate of protein digestion is 2 fold faster at

250C than at 300C (Honda et al. 1985).


The rate of blood meal digestion in sand flies depends

on the source of the blood meal. Schlein et al. (1982a,

1982b, 1983) reported vector incompatibility between field

collected Ph. papatasi and Le. major due to the presence of

turkey blood meal. They also showed that if sand flies were

fed leishmania parasite in saline 24 hrs prior to a blood

meal, the rate of infection was low (Schlein et al. 1983).

This observation was contrary to the earlier studies in

which sand flies showed a high rate of infection even when

they were fed on saline containing promastigotes of a local

strain of Le. tropica (Adler 1938).

Serine proteases and their mode of action

Structure of serine proteases. Serine proteases are a

large group of enzymes with a serine residue at their active

site. These enzymes are irreversibly inhibited by

diisopropylflurophosphate (DFP). DFP is a flouridated

phosphate ester and is cleaved by trypsin or chymotrypsin.

The enzymes are classified as endopeptidases because the

cleavage of the terminal peptide is inhibited by the charge

on the amino or carboxyl group of terminal residue.

Chymotrypsin and trypsin are classified as serine proteases.

Mammalian enzymes are secreted in the form of zymogen or

proenzyme, an inactive form of an enzyme. Mammalian trypsin

and chymotrypsin are synthesized as trypsinogen and

chymotrypsinogen at the pancreas endoplasmic reticulum

(Stryer 1975). The enzymes are then transferred to the

1 245


I1 15 1


., ^m +
,/ Ser- Arg
/ 14-15

1 A1 J13

I I 146
] 146



Figure 2-4. Diagramatic representation of the activation of
chymotrypsin resulting in the formation of cm
chymotrypsin (adapted from Stryer 1975).


Asp 102-C-O-H-N/N--- H-O- Ser 195

His 57

II y
Asp 102 -CO-0-H ...N N -H- -- Ser 195

B His 57

Figure 2-5. Charge relay system of chymotrypsin molecule.
A = normal charge distribution in a
chymotrypsin molecule; B = Conversion of
chymotrypsin into a powerful nucleophile. The
removal of one proton by aspartate 102 from
serine 105 via histidine 57 results the
conversion of serine 195 into a powerful
nucleophile (adapted from Stryer 1975).


golgi apparatus where a lipid membrane is secreted around

them. Membrane bound trypsinogen and chymotrypsinogen are

transported into the gut lumen. The mammalian

chymotrypsinogen molecule has 245 amino acid residues. It

is activated by the action of trypsin that brings about

cleavage of the molecule at arginine 15 and isoleucine 16.

The resulting n-chymotrypsin is completely active but is

unstable. It is further cleaved by the action of other n-

chymtrypsin molecules into a-chymotrypsin, a stable molecule

(Fig. 2-4).

The mammalian a-chymotrypsin molecule has three peptide

chains which are interconnected by two disulfide bonds (Fig.

2-4). It has a compact ellipsoidal shape and all the

charges are on the surface of the molecule except the three

most important charges that play a crucial role in its

activity. These residues are serine 195 (Ser), histidine 57

(His) and aspartic acid 102 (Asp). Serine 195 is a strong

nucleophile and plays an active role in the reactivity of

both trypsin and chymotrypsin.

The high nucleophile activity of the serine 195 is

contributed to the presence of charge relay system between

serine 195, histidine 57 and aspartic 102 (Fig. 2-5).

Mode of action of serine proteases. The serine

proteases hydrolyze peptide bonds by attacking the carbonyl

group of the peptide bond through the oxygen atom of the

hydroxyl group of serine 195. This reaction is brought


CH2-- R-N -H

His 57


0\ /R
-CH2-O/ \





Figure 2-6. Acylatation reaction during the hydrolysis of a
peptide bond by chymotrypsin (adapted from
Stryer 1975).

-CH2 -O/

His 57


-CH2--/ \


-CH2--O O-H



Figure 2-7. Deacylation reaction during the hydrolysis of a
peptide bond by chymotrypsin molecule (adapted
from Stryer 1975).

0 Ser 195
A s p 1 0 2 ) O H -
O-H ... -H...O 1

t -
/ II \
CH3 0 CH3

Asp 102 k-

His 57

'. .

Ser 195


F CH3 0 CH3
I.C-- P-O-CH

(His 57

Asp 102)O
0 ser 195

*- O^*^ P-O-CH
H' CH3y CH3

His 57
Asp 102>), -
0-H --^ -H

Ser 195

CH3 r CH3


Figure 2-8. Irreversible deactivation of trypsin-like
enzyme by DFP.



about by a charge relay system that helps to draw a proton

from the hydroxyl group of serine 195 (Fig. 2-5). The

nitrogen atom of the susceptible peptide bond then receives

a proton from Histidine 57 and the peptide bond is cleaved.

This brings about the completion of the acylation stage of

the hydrolytic reaction and a complex results in which the

amine component is hydrogen bonded to histidine 57, and the

acid component of the substrate is esterified to serine 195

(Fig. 2-6).

In the next step a water molecule is added resulting in

several steps of proton removal and ionic attack of the

carbonyl carbon of the acyl group (Fig. 2-7). As a result

the active site is regenerated.

Inhibitors of digestive enzymes. Certain compounds act

as proteinase inhibitors, such as certain factors in the

serum from mammalian blood can cause in vivo inhibition of

Ae. aegypti proteinase (Fisk & Shambaugh 1952; Gooding

1966a, 1966b, 1972, 1974; Huang 1971). Certain other

compounds, DFP (Irreversible inhibitor of both trypsin and

chymotrypsin) binds irrevesibly to serine 195 at the active

site (Fig. 2-8), TPCK (tosylamide-2-phenyl chloromethyl

ketone; chymotrypsin inhibitor) and TLCK (tosyl-L-lysine

chloromethyl ketone HC1; trypsin inhibitor), have been used

for qualitative and quantitative determination of

proteolytic enzymes (Graf & Briegel 1985; Brovosky & Schlein

1987; Brovosky & Schlein 1988).



.."'.., COMMON

Figure 2-9. Diagramatic representation of the reproductive
system of a female sand fly.


Female Age Grading

Reproductive System of Female Sand Flies

The reproductive system of female sand flies consists

of a pair of fusiform ovaries, each of which has a separate

lateral oviduct that opens into a common oviduct. The

latter is a short, wide tube lined with cuticle and which

has circular and longitudinal muscles on the outside. The

common oviduct opens into a wider portion, or vagina, and is

surrounded by arms of a chitinous furca (Perfil'ev 1968).

Two elongated accessory glands are present on the sides of

the ovaries and also open into the vagina. In newly emerged

females, these glands are pale, transparent, and lack

granular substance, whereas the granular substance is

present in fed and/or parous females (Lewis 1965; Lewis et

al. 1970) (Fig. 2-9). These granular secretions are

necessary for protecting eggs against fungal and bacterial

infections (Adler & Mayrink 1961).

Two spermathecae open on the sides of the bladder-

shaped vagina by their respective spermathecal ducts in some

species (Sinton 1925; Sinton & Braud 1928) or by a common

spermathecal duct in others (Sinton 1925). In the majority

of the Diptera, three spermathecae and three spermathecal

openings are the primitive condition. This condition

corresponds to the aedeagal openings in the male. The

female spermathecal openings are coadapted to the aedeagal


openings of the male (Downes 1968). The size of the

spermathecal duct varies in different species of sand flies.

The spermathecae are of variable shapes and are useful in

the identification of different species of sand flies (Young


Downes (1968) considered the presence of two

spermathecal ducts, which open separately a primitive

character compared to a common spermathecal duct and a

single spermathecal opening. Gerber (1970) studied the

evolution of spermatophores in Pterygotan insects. He

considered the presence of spermatophore as the most

primitive condition and the deposition of accessory gland

secretions, together with sperms directly inside the

spermathecae and/or bursa copulatrix, or vagina, as the most

advanced character. The presence of a mating plug formed by

the accessory gland substance and the sperms (a condition

found in some mosquitoes) was considered as an intermediate

character between the above two conditions.

Each spermatheca has a round or conical head, present

at the apex of the spermathecal neck. The head is

surrounded by numerous gland cells which open into the neck

of spermatheca by different number of chitinous canals. The

gland cells form a large rounded gland that can be easily

seen in fresh preparations. It is destroyed by fixation

processes using alcohol or KOH (Perfil'ev 1968). The gland

produces a fluid into the spermathecal head that serves to



Figure 2-10. Different stages of the follicular development
in Lutzomyia anthophora. A = stage N; B =
stage I; C = stage I-II; D = stage II; E-F =
stage III; G-H = stage IV; and I = stage V.


preserve the spermatozoa (Perfil've 1968). The vagina

narrows down to a tube that opens to the outside as the

vulva. The vulva is situated near the anus and is separated

from it by a muscular thickening.

The ovaries are enclosed in an ovarian sheath that

continues with the oviduct on lower end and on the upper end

forms a soft filament like structure (Fig. 2-9). The

filament or ligament of each ovary connects it to the body

cavity (Fig 2-2). The ovaries are of polytrophic meriostic

type. Each ovary has a variable number of ovarioles,

radially attached to the internal oviducts on each side. The

part of the internal oviduct where the pedicle of the

ovariole attaches to the oviduct is called the calyx. Each

ovariole is attached to the ovarian sheath by a filament and

has its own ovariole sheath. The part of the ovariole where

active cell division occurs is called the germarium which

has a variable number of oogonia. Each developing follicle

(e.g., in Ph. papatasi) contains about seven nurse cells and

an oocyte (Jobling 1987). Each follicle, present in its own

follicular tube, has its own follicular epithelium. At a

given time, at least three follicles are present in

different stages of development. The size of the follicles

depends on their stage of development (Fig. 2-10).

In gonoactive females, the last follicle in each

follicular tube begins to develop after the ingestion of a

blood meal. The follicles of newly emerged females contain


eight monomorphic nuclei, seven of which are nurse cells and

the remainder is the oocyte or ovum. The function of the

nurse cells is to nourish the ovum. The development of the

follicle was first described in anopheline mosquitoes by

Christophers (1911), who divided the whole developmental

cycle into stages I-V. Mer (1936) recognized two more

stages in the initial growth period of the follicle and

called them as N and I-II (Fig. 2-10)

Detinova (1962) described the following stages of ovum

maturation and which are essentially as applied to sand

flies due to similarities in their growth pattern.

Stage N. A spherical follicle that consists of 8

monomorphic cells (Fig. 2-10-A).

Stage I. The follicle is spherical or slightly oval.

Seven nurse cells lie above a distally located oocyte (Fig.


Stage I-II. The follicle is distinctly oval in shape

and one or two rows of yolk granules encircle the oocyte

nucleus (Fig. 2-10-C).

Stage II. The egg grows larger due to accumulation of

large amount of yolk granules around the oocyte nucleus

(Fig. 2-10-D). The ovum is considerably larger than the

nurse cells and occupies half of the follicle.

Stage III. The follicle is oblong in shape and the

nucleus is hidden by the yolk granules that cover 1/2 to 3/4

of the follicle (Fig. 2-10-E & 2-10-F).


Stage IV. The follicle is longer and larger and the

nurse cells occupy only the upper most part of it. More

than 9/10 of the follicle is occupied by oogonium, that is

full of yolk (Fig. 2-10-G & 2-10-H).

Stage V. The nurse cells are pushed towards the tip of

the follicle due to increased yolk deposition. The follicle

is covered with a well developed chorion. A micropyle is

formed at this stage (Fig 2-10-I). In certain species of

mosquitoes the nurse cells are pushed out of the micropyle.

The above classification of different developmental

stages of the follicle has been successfully applied to

different species of sand flies (Magnarelli et al. 1984).

Certain species of sand flies can develop their eggs

autogenously. This characteristic introduces an error in

determining the epidemiological importance of such species

in the transmission of nontransovarially transmitted

pathogens such as Leishmania. Autogenous females can

maintain transovarially transmitted viruses without blood

feeding; e.g., several sand-fly born viruses of vertebrates

are transovarially transmitted in their insect hosts (Tesh &

Chaniotis 1975; Endris et al. 1983; Tesh & Modi 1983).

Comparision of egg development between autogenous Ph.

papatasi held on fructose solution only, with those provided

with a blood meal, showed no difference in the total number

of eggs produced (Magnarelli 1984). Twenty-two percent of

the females were autogenous and produced eggs without a


blood meal. The rate of egg development was similar in both

autogenous and anautogenous females. Egg development was

asynchronous in blood fed females (Magnarelli 1984).

Most species of sand flies are gonotrophically

concordant and require only one blood meal for complete

development of a clutch of eggs. Certain sand flies, like

Ph. papatasi, take blood at any stage of follicular

development (Schmidt & Schmidt 1965; Killick-Kendrick 1979;

Magnarelli et al. 1984). Multiple feeding during one

gonotrophic cycle increases the chances of parasite

transmission by infected sand flies (Killick-Kendrick 1979).

The effect of a second blood meal on the rate of development

of parasites ingested with the first blood meal is unknown.

Adler and Theodor (1926) suggested that multiple feeding was

a phenomenon which occurred only in the earlier stages of

egg development, but Christensen and Herrer (1980) observed

a significant number of gravid Lutzomvia species, attracted

to animal baited traps. This observation led Magnarelli et

al. (1984) to conclude that sand flies would even take a

blood meal just prior to oviposition if a host was present.

In anautogenous species of sand flies, such as Lu.

longipalpis, fecundity is correlated with the size and

composition of the blood meal. Composition of the blood

meal differs depending upon the mammalian host species. On

the other hand, in autogenous species fecundity is directly

related to the larval nutrition (Magnarelli et al. 1984).

Different Methods of Age Grading Sand Flies

Although the following age grading techniques have been

used successufuly for mosquitoes, some of these such as

presence of parasitic water mites (Corbet 1963; Mullen

1975), appearance of scaling of the wings (Perry 1912),

tergal pigmentation patterns (Linley & Baverman 1984),

presence of meconium in the midgut (Detinova 1962), presence

of green colored fat in the haemocoel, eggs retained in the

ovaries, presence of greatly dilated, translucent internal

oviduct, size of ampullae, qualitative changes in the shape

of the ampullae, and tracheation of ovaries and midgut are

not effective for sand flies.

Presence of daily growth layers. Many exopterygote

insect species such as grasshoppers, flies, and mosquitoes

deposit cuticular growth layers on their exoskeleton.

Similarly, these are deposited as bands on internal muscle

apodemes of thoracic phragmata in endopterygotes. These

layers have been used to age grade populations in the field

(Neville 1963a, 1963b, 1963c, 1983; Schlein & Gratz 1972;

Tyndale-Biscoe & Kitching 1974; Ellison & Hampton 1982; Yual

& Schlein 1986).

The growth layers were initially described as

fluorescent layers in the resilin of locust-rubber like

cuticle (Neville 1963a, 1963b). They appear as dark and

light bands under phase contrast microscopy or crossed


polaroid light filters (Neville 1963c). The endocuticle

deposited during the day has its microfibrils oriented in

one direction, whereas that deposited during the night is

helicoidal. Thus, night deposited cuticle appears dark and

day deposited layer is light in color. A consecutive pair

of cuticular layers (appearing as a dark a light band by

crossed polaroids light) represents 24 hrs growth in

grasshoppers (Neville 1963c). The growth layers are easy to

observe in insects with melanized cuticles, but special

staining methods are required in insects with transparent

apodemes (Schlein & Gratz 1973; Schlein 1975; Schlein 1979a,

1979b; Yual & Schlein 1986; Alexander 1988).

In Glossina one day error was possible in counting of

the growth layers. The width of growth layers decreases in

older flies but is easy to count at 40X magnification

(Schlein 1979a). The banding is dependent on the

fluctuations in the daily temperature and cuticular growth

is known to cease completely at low temperature (Neville

1965; Schlein 1979b). Banding is most clear when the daily

temperature is below the threshold of cuticular growth of an

insect (Tyndale-Biscoe & Kitching 1974). In most insects

cuticular growth layers are formed immediately after

emergence and if the anomalies are taken into account, this

method is satisfactory for determining age of both sexes.


This method was successfully applied for age grading

wild caught sand flies (Yual & Schlein 1987; Alexander


Accessory glands. Granular accessory gland secretion

was first noted by Adler and Theodor (1935) in Phlebotomus

perniciosus (Newstead). The accessory gland secretion was

secreted after blood feeding and some of it was retained in

accessory glands after oviposition. The eggs are coated

with it during oviposition and adhere them to the

oviposition substrate (Ward 1985).

There are differences in opinion regarding use of

accessory gland as indicator of age in wild caught sand

flies. Adler and Theodor (1957) considered the presence of

accessory gland secretion as indicator of old age. This

view was questioned by Garnham and Lewis (1959) who examined

field collected sand flies and found that a large proportion

of them contained accessory gland substance. They thought

that it was possible for nulliparous flies to develop the

accessory gland substance. The accessory glands of Ph.

papatasi also secrete accessory gland secretion after a

blood meal but not all females retain part of this substance

after oviposition (Dolmatova 1942). Some Panamanian sand

flies secrete the accessory gland fluid even before taking a

blood meal and are capable of discharging it at various

times (Johnson & Hertig 1961). Lewis (1965) noted

structural changes of the accessory glands in five species


of Lutzomyia from Bleize and concluded that this method was

unreliable. The accessory glands of Lu. flaviscutella

(Mang) and Lu. furcata (Mang) were also considered

unreliable indicators of parity (Ready et al. 1984).

The presence of accessory gland substance together with

small ovaries was considered a reliable indicator of old age

in some Kenyan sand flies (Lewis & Minter 1960). Soshina

(1951, reprint not seen referred from Detinova 1962) used

the accessory glands of Phlebotomus and age graded the

population by dividing it into four categories, 1)

nulliparous and newly emerged females, 2) females in their

first gonotrophic cycle, 3) one parous females and 4)

females which refed after their first oviposition.

The accessory glands were also used to study the

population trends, such as breeding seasons, appearance of

successive generations, and the mode of dry-season survival

of Sergentomvia garnhami (Minter 1964). Lewis et al. (1970)

found that in Lu. shanonni (Dyar), a higher proportion of

nulliparous females showed accessory glands secretions which

might be due to development of secretions in the glands

prior to development of the ovaries. They suggested that

some use of the accessory glands of Lu. ovallesi (Ortiz) and

Lu. panamensis (Shannon) could be made for the study of

general trends of their populations. Chaniotis and Anderson

(1967) considered the accessory glands of 3 California sand

fly species as very reliable indicators of their parous







'\ F 4- CALY




Figure 2-11.

Digramatic representation of the different
stages that are found during the formation of
dilatations. A = an ovarian follicle with a
sack stage that is present just after
oviposition; B = half contracted sack stage; C
= an ovariole of a 1-parous female showing one
dilatation; D = an ovariole of a 2-parous
female showing 2 dilatations; E = a
degenerated ovariole from a gravid female,
note the presence of yolk granules; An
ovariole from a nulliparous female, note the
presence of ovarian tissue or calyx attached
to the pedicle.



status. Only 3% of the 300 parous females lacked granular

material in their accessory glands.

Ovarian follicular dilatation. This method is

applicable to females of any age and stage of blood

digestion (Detinova 1962). Ovaries are dissected out in a

drop of physiological saline. An ovary is moved to the side

of the drop and the ovarian sheath is carefully removed at

places. In these areas the ovarioles are very loose and the

number of dilatations can be easily counted by stretching

the ovarian follicle. In females, recently oviposited, the

ovariole are in a sac-like stage (Fig. 2-11-A). The sac-

like stage contract with time and form a bead-like structure

(Polovoda 1949 reprint not seen referred from Detinova 1962)

(Figs. 2-11-B, 2-11-C & 2-11-D).

In each gonotrophic cycle of sand flies a certain

number of eggs degenerate. Most of the time the degenerated

eggs belong to the same ovariole. In multiparous females

which have recently oviposited number of dilatations can be

determined from the degenerated eggs (Fig. 2-11-E). The

number of dilatations found in the ovaries of a female give

an indication of her age. To estimate the female age in

different seasons, lengths of the gonotrophic cycles

determined at different temperature is required. This

method can be used to determine the vertical survivorship of

both healthy and infected field collected females in

different seasons of the year (Reisen et al. 1986).


Opinions vary concerning the applicability of

dilatation technique for age grading sand flies. The

ovaries of parous sand flies are very loose (Dolmatova

1942). Detinova (1962) considered the small size of sand

flies as the major obstacle for the dilatation technique.

Lewis and Minter (1960) applied the dilatation technique to

determine the parity of unfed flies from Kenya. They

concluded that it was possible to determine the parity of

females by this technique. They found a good correlation in

the parous females and presence of the accessory gland

secretion. On the other hand Lewis (1965) found a certain

degree of error in determining parity of wild caught

Lutzomyia species by this technique.

Similarly, Ready et al. (1984) applied Polvodova's

method to assess the age of the laboratory reared Lu.

flaviscutellata (Mangabeira) and Lu. furcata (Mangabeira).

Results of their blind test showed that this method was good

for simply classifying females into nulliparous and parous


The dilatation technique was successfully applied to

age grade Lu. ylephiletor (Fairchild & Hertig). The average

age of was greater for females collected in biting

collections than those collected from resting sites. This

fact was supported by large number of flagellate infections

found in females from biting collections (Johnson et al.

1963). Field collected Ph. papatasi from Iran showed only


one-parous and nulliparous females. No multiparous female

were caught (Magnarelli et al. 1984). Chaniotis and

Anderson (1967) considered age grading by dilatations as an

impracticle and time consuming technique. On the other

hand, Wilkes and Rioux (1980) found dilatation technique

very useful. They utilized this technique to compare the

age structure of Ph. ariasi (Tonnoir) collected from the

walls of a farm house and in light traps. A significant

difference was found between ages of flies collected by the

two methods. They concluded that for epidemiological

studies, light trap collections were better. About 1/3 of

the females in light trap collection were one-parous and few

were two and three parous. The collections from the wall of

the farm house included only one female with one dilatation

and rest of 105 females were nulliparous.

Application of the dilatation technique requires some

caution because of false dilatations formation in

nulliparous Culex mosquitoes (Yajima 1970; Oda et al. 1978;

Nayar & Knight 1981; Reisen et al. 1986). In Culex

tritaeniorhynchus (Giles) the tracheolar coiling method gave

a better indication of nulliparity and was recommended for

use on one ovary and the dilatation technique on the other

ovary (Reisen et al. 1986). In laboratory control

experiments, the greatest difficulty was encountered in

distinguishing 2 parous gravid females; it was always much

easier to find the degenerated eggs in the field collected


mosquitoes as compared to the laboratory collected controls

(Reisen et al. 1986). During laboratory control studies,

experimental conditions prevented the females from refeeding

on the night of oviposition and might had contributed to the

discrepancy (Reisen et al. 1986). Gillies and Wilkes (1965)

and Spencer (1979) also cautioned against errors associated

with formation of dilatation in laboratory experiments. No

studies are available describing formation of false

dilatation in sand flies.

Hormonal Control of Insect Reproduction and Role of Juvenile
Hormone III in Adult Insects

Juvenile hormones (JH) are synthesized and released in

insects from the corpora allata. In larval instars of

insects JH and ecdysone act concurrently. Ecdysone triggers

the biosynthetic pathways essential for molting. During

morphogenesis JH act in concert with ecdysone. Juvenile

status in insects depends upon continuous presence of JH,

that act on the cells and prevent them from maturing. In

immature insects JH manifests a multiplicity of

morphogenetic effects. It control the kind of cuticle the

epidermal cells secrete in response to ecdysone. Juvenile

hormones also influence the development of internal organs

like central nervous system, gonads, and midgut, where they

deter maturation and metamorphosis (Sehnal 1968). Juvenile

hormones also hinder the maturation of the imaginal discs


which are the primordia for many adult integumentary

structures in endopterygote insects.

In nature, adult mosquitoes are ready to feed on nectar

after their cuticle hardens. An active search for sugar

feeding begins later due to locomotor activity and receptor

sensitivity (Meola & Readio 1988). Mosquitoes take blood to

utilize the proteins for egg maturation. Host seeking does

not depend on JH in Ae. aegypti and is apparently related to

maturation of antennal chemosensory afferent neurons

involved in detection of lactic acid (Bowen & Davis 1989).

In contrast, in Cx. pipiens, JH has been reported to control

the initiation of biting behavior (Meola & Petralia 1980;

Meola & Readio 1987).

Newly emerged female mosquitoes become reproductively

competent due to a chain of events that starts by the

release of JH from their corpora allata. Juvenile hormone

stimulates the primary follicles of newly emerged females to

grow in size to a previtellogenic resting stage (Lea 1963;

Gwadz & Spielman 1973). The level of JH remains constant

in newly emerged and sugar fed Cx. pipens; but in Ae.

aegypti, the JH level is high during the first two days

after emergence and then gradually declines. This may be

due to feed back inhibition by resting stage ovaries

(Rossignol et al. 1981; Shapiro et al. 1986). The latter

scientists (1986) observed that in Ae. aegypti JH levels

dropped three days after emergence with no corresponding


increase in esterase levels. They suggested that formation

of resting stage follicles in 3-day-old mosquitoes might

inhibit hormone production by the corpora allata.

Implantation of both preresting and resting stage donor

ovaries into newly emerged females inhibited growth of the

follicles of the host ovaries. However, implantation of

resting stage ovaries only inhibited growth of the

preresting stage host ovaries (Meola & Readio 1988).

Absence of JH due to allatectomy shortly after emergence of

Ae. aeqypti and Cx. pipiens inhibited the growth of primary

follicles to the resting stage (Gwadz & Spielman 1973; Lea


Application of synthetic JH stimulated the growth of

primary follicles (Spielman 1974). Once the resting or

previtellogenic stage of the follicle was achieved and the

follicle was 50-75 pm in length; no further growth of

follicles occurred (Meola & Readio 1988). The follicles of

S. calcitrans did not develop when females were

allatectomized within 1 hr of emergence, regardless if they

were provided a sugar or a blood meal. Normal follicular

growth is achieved only after implanting of corpora allata

(Moobola & Cupp 1978).

A humoral factor released from the brain was thought to

be essential for mosquito egg development (Clements 1956;

Gillet 1956). Release of this factor was suggested to occur

after blood feeding due to the distention of midgut wall.


Midgut distension stimulated the neurosecretory cells of the

brain which secreted the brain hormone. The brain hormone

directly or indirectly activated the corpora allata. The

corpora allata secreted JH which initiated vitellogenesis

(Larson & Bodenstein 1959). The above hypothesis was

disputed by Bellamy and Bracken (1971), who showed that in

Cx. pipiens, only certain proteins initiated egg development

instead of sustained stretching of midgut receptors. A

brain hormone "egg development neurosecretory hormone"

(EDNH) from the median neurosecretory cells (MNC) was also

released after eclosion (Lea 1963, 1969, 1972; Meola & Lea

1972) and was stored in the corpus cardiacum (CC). EDNH was

released into the hemolymph after blood feeding. Its

release is triggered by corpus cardiacum stimulating factor

(CCSF) (Borovsky 1982).

Yolk is deposited in sugar fed mosquitoes of several

species either by injecting or feeding them high

concentrations of 20-OH-ecdysone (Spielman et al. 1971).

Spielman et al. (1971) considered the effect a

pharmacological artifact. During the same period it was

shown that in Ae. aeqypti vitellogenin was synthesized by

the fat body cells (Hagedorn & Judson 1972). Presence of

the ovaries was necessary for vitellogenin production. Fat

bodies of sugar fed females synthesized vitellogenin during

in vitro incubation with ovaries of blood fed females

(Fallon et al. 1974). This led Hagedorn (1974) to suggest


that vitellogenin stimulating hormone (VHS) was secreted by

the ovaries. Later, Schlaeger et al. (1974) showed that

ecdysteroid levels were higher in blood fed females than in

sugar fed females. In vitro incubation of fat bodies in the

presence of 105 M 20-OH-ecdysone also resulted in the

production of vitellogenin in levels comparable to those

found in blood fed females (Hagedorn 1974). However,

Borovsky and Van Handle (1979) were not able to stimulate

fat bodies in vitro with 20-OH-ecdysone. It was confirmed

that fat bodies removed from sugar fed females produced

smaller quantities of vitellogenin (5%) compared to fat

bodies removed from blood fed females (Borovsky 1984).

Continuous presence of 20-OH-ecdysone secreted by the

ovary was not necessary for the synthesis of vitellogenin.

In ovariectomized mosquitoes, accumulation of vitellogenin

caused feed back inhibition of vitellogenin synthesis

(Borovsky 1981a). To explain the above discrepancies,

Hagedorn (1983) suggested that fat bodies of sugar fed

females were nutritionally inferior than those of blood fed

females and there was a high turnover of 20-OH-ecdysone.

Later studies, showed that injection of physiological

amounts of 20-OH-ecdysone caused the separation of secondary

follicles; continuous infusion of physiological amounts of

20-OH-ecdysone did not bring about vitellogenin synthesis in

sugar fed females (Beckemeyer & Lea 1980; Lea 1982).


Injection of physiological amounts of 20-OH-ecdysone

into isolated abdomens of autogenous Ae. atropalpus

(Coquillet) initiated oogenesis. Egg development occurred

when the abdomens were treated first with small amounts of

JH (Fuchs et al. 1981). Similar results were obtained with

anautogenous females of Ae. aeqypti (Borovsky 1981b). Thus,

the presence of both JH and ecdysone is necessary for egg

development in blood fed mosquitoes. Borovsky et al. (1985)

showed for the first time that JH III was produced in blood

fed females, and vitellogenic ovaries induced egg

development in a nonvitellogenic host only if the

nonvitellogenic host was first treated with methoprene.

Vitellogenesis was induced in isolated abdomens by 20-OH-

ecdysone only if the abdomens were first treated with


Two stages of oocyte growth follow after ingestion of a

blood meal. The first stage, called the initiation stage,

when the oocyte grows from the resting stage to stage II or

early stage III (0-50 pm) (Gillet 1956). This stage is

independent of brain factors (Gillet 1956, 1957). The

second stage, or promotion stage, represents when the oocyte

grows from stage III to stage V (>50 450 upm) and requires

the release of EDNH 4-8 hrs after blood feeding (Lea 1972;

Gillet 1957). Greenplate et al. (1985) showed that EDNH is

released twice in Ae. aeqypti, once before, immediately

after blood meal and once 8-hrs after the blood meal. They


also suggested that the critical time of release of factors

stimulating ecdysteroid production occurs at 4-8 hrs.

Greenplate et al. (1985) suggested that the release of

ecdysteroid is controlled by EDNH and indicated that the

first release of EDNH immediately after blood feeding brings

ecdysteroid level to 50-60% of its maximum. Eight hrs

later, a second release of EDNH brings ecdysteroid to its

maximum level.

Release of JH prepares the primary follicles of

mosquitoes to become receptive to brain hormone (EDNH)

(Shapiro & Hagedorn 1982) and prepares the fat bodies for

20-OH-ecdysone (Hagedorn 1977). Ecdysone is synthesized by

the ovaries and is converted into 20-OH-ecdysone by the fat

bodies; it stimulates the fat bodies to produce vitellogenin

(Hagedorn et al. 1975). The level of JH declines rapidally

just after blood feeding in both Ae. aeqypti and Cx. pipens

(Shapiro et al. 1986; Readio et al. 1988).

Different changes occur in the morphology of the

follicular epithelium soon after ingestion of a blood meal.

In mosquitoes such as Ae. aeqypti, pits or wells form in the

surface of the oocyte. The pits later pinch off from the

follicular membrane and carry yolk proteins into the oocyte

and fuse to form large yolk bodies. Thus, the transport of

yolk proteins occurs by pinocytosis (Roth & Porter 1964).

Follicle cells of Rhodnius prolixus lose fluid and

shrink as a result of the action of juvenile hormone.


Large spaces appear between adjacent follicle cells

resulting in a direct contact between the vitellogenin in

the haemolymph and the oocyte surface (Pratt & Davey 1972).

Similarly, in cockroaches, the presence of JH induces

appearance of intercellular spaces in the follicular

epithelium of the ovary (Bell & Barth 1971). In Leucophea

maderae (L), JH also stimulates DNA synthesis in the

follicular epithelium of the terminal follicle (Koeppe &

Wellman 1980).

In Ae. aegypti, the secondary follicle separates from

the primary follicle within 20-hrs after blood meal and is

presumably regulated by a rise in the titer of 20-OH-

ecdysone (Beckemeyer & Lea 1980).

In Cx. pipens, the synthesis of JH declines rapidly

after blood feeding; whereas in sugar fed females of the

same age it remains high. Rising ecdysteroid levels may act

on the corpora allata and suppress their activity (Rosignol

et al. 1981). However, Meola and Readio (1988) suggested

that a drop in the level of JH few hours after blood feeding

was faster than the increase of ecdysteroid. Shapiro et al

(1986) failed to find a direct correlation between JH

esterase activity and JH concentration, and they were unable

to explain the rapid decline in the level of JH after blood

feeding, without an increase in JH concentration.

In Ae. aeqypti, an increase in JH III biosynthesis

occurred from eclosion to 144-hrs after emergence (Borovsky


et al. 1989). When synthesis of JH III was followed in vivo

at different time intervals after blood feeding, an increase

in the synthesis of JH III was found within 2-hrs after

blood feeding, followed by a decline in JH III synthesis.

Juvenile hormone III concentrations were similar to those

found in newly closed females. The decline in JH III

synthesis was followed by a gradual increase which reached a

maximum 46 to 48-hrs after the blood meal. Minimum amount

of JH III was synthesized 120-hrs after the blood meal

(Borovsky et al. 1989).

Metabolites of JH III were followed in sugar fed

mosquitoes and in females at different times after blood

meal ingestion (Borovsky et al. 1989). These authors showed

that JH was metabolized first to JH-acid and then

immediately to JH-diol-acid by epoxide hydrase. JH-acid was

a better substrate then JH III for the enzyme epoxide

hydrase which hydrated it 24 times faster then JH III

(Borovsky et al. 1989).

In Cx. pipiens, the secondary follicles grow up to 43

mu in length within the first three days after a blood meal.

Further growth of the follicle, to 57 pm in length, requires

the presence of JH secreted from 4th to 8th day after the

ingestion of a blood meal (Meola & Readio 1988).

In An. quadrimaculatus, the secondary follicles grow up

to 85 pm long after ingestion of a blood meal. After

oviposition, the secondary follicles develop to the resting


stage that average 105 ipm long (Meola & Readio 1988). On

the other hand, previtellogenic growth of the secondary

follicles of An. stephensi is completed during the first

gonotrophic cycle, before the ovipostion of the primary

follicles (Redfern 1982).

Fat bodies of mosquitoes lose their ability to respond

to 20-OH-ecdysone 20-hours after the blood meal (Bohm et al.

1978). The fat bodies may require another burst of juvenile

hormone to produce a second batch of eggs (Hagedorn 1983).

These observations are also supported by the studies of

Readio and Meola (1985). Who allatectomized Cx. pipiens

females after their first oviposition, refed them, then

noted the length of their secondary follicles. Only a small

percentage of the mosquitoes refed and of those that fed,

very few had vitellogenic ovaries. They also measured the

length of secondary follicles of mosquitoes with intact

corpora allata. Their results indicated that JH became

available for the growth of secondary follicles 24 hrs after

a blood meal. The resting stage follicles were 70 pm in

length. The resting stage secondary follicles were

competent to complete a second gonotrophic cycle as soon as

the mosquito took another blood meal after oviposition. Egg

retention beyond the normal gontrophic cycle inhibited JH

synthesis by the corpora allata and JH synthesis was resumed

by the corpora allata after oviposition (Readio et al.



Thus, we can conclude that reproduction in

haematophagous insects consists of different behavioral and

physiological processes that are closely regulated and which

function in concert.



Earlier studies on the reproductive biology of male

sand flies dealt with their mode of copulation and the

anatomical aspects of mating (Sinton 1925; Christopher &

Barraud 1926; Sinton 1932; Hertig 1949; Sherlock & Carnerio

1964). Some information was also available about their

epigamic behavior and the time required to complete

insemination (Johnson & Hertig 1961; Chaniotis 1967; Ward

1977; Endris 1982). Sherlock & Carnerio (1964) described

similarities and differences in the morphology of the male

reproductive system in some sand flies and discussed the

importance of the male anatomy as a taxonomic tool.

Generally, however, more information has been obtained

about the reproductive system of female haematophagous

insects because of their role in the spread of disease.

Studies of female reproduction have also received attention

because of their control by hormones and the significant

morphological changes observed in the eggs after blood



Recently, insecticide resistance has evolved in many

haematophagous insects and has prompted a search for new

insect control methods. One such method is to release

sterile males that are produced by genetic manipulation and

chemosterilization. Such a technique, if ever devised for

sand fly control, would require a vast knowledge about their

mating behavior in nature. In particular, a study of

changes in the morphology of the male reproductive system

caused by aging and mating would be helpful in assessing the

mating competitiveness of laboratory reared males (Reisen et

al. 1981).

The objectives of the present investigation were: 1)

to investigate the epigamic behavior of male and female Lu.

anthophora; 2) to study the effect of age on the mating

competence of virgin females and males; 3) to study the

morphology of the male reproductive system and 4) to

evaluate changes in the male reproductive system caused by

increasing age and multiple matings.

Materials and Methods

Strain. Lu. anthophora (Addis), was originally

colonized from Texas (Addis 1945a, b). The present strain

was colonized from Texas 8 years ago (Endris et al. 1982).

Colony Maintenance. During the present study, the

colony of Lu. anthophora was reared in an environmental

chamber (Hotpack model 434310), maintained at 26 + 20C, 80-


85% relative humidity and with a 16:8 L:D photoperiod. The

rearing procedure followed Young et al. (1981) and Endris et

al. (1982).

Adults were maintained in 12x25.5x21.5 cm glass

aquarium cages lined with plaster of Paris on three sides

(Endris et al. 1982). Adults were provided with 10% sucrose

solution on cotton pledgets as well as fresh apple slices;

the latter was replaced daily and the former was changed

every third day. Young (3-day-old) females were blood fed

on an anesthetized hamster, immobilized with ketamine

hydrochloride (KetasetR) that was injected intramuscularly

(0.2 ml/adult hamster). Females were transferred 24 hrs

after blood feeding to 120 ml oviposition tubes, covered at

the bottom with plaster of Paris. Sugar solution was

constantly available to these females. Females oviposited 4

to 6-days after the blood meal and the eggs hatched 8 days

following oviposition.

Larvae were furnished with an aged 1:1 mixture (by

volume) of dry rabbit feces and Purina rabbit chow (complete

diet 5315R) as food (Young et al. 1981). Larvae were reared

in 120 ml containers, lined with plaster of Paris. A

maximum of 100 larvae was placed in each container and ample

food, along with a few drops of water, was added every other

day to avoid competition. Pupae were placed in another

container one day after their formation.

Investigation of Epiqamic Behavior

The behavioral patterns of males and females before,

during, and after mating were determined as follows. Ten 3

to 5-day-old virgin males were allowed to cohabit with 10

virgin females of the same age in an ice cream carton (1

pint) lined with plaster of Paris at the base and covered on

top with a fine mesh screen. The mating behavior was

observed for 3-hrs. The time required for complete

insemination of a female by a male was determined by

allowing one 5-day-old virgin male to cohabit with five

virgin females of the same age and noting the time required

for complete insemination. The observations were repeated 5

times. The epigamic behavior of both sexes, before and

during initiation of mating, was also noted. Similarly,

variable numbers of males and females were isolated in an

aquarium cage and the different epigamic behaviors of both

sexes observed before, during and after blood feeding.

The Effect of Age on Mating Competence

The effect of age on the mating competence of virgin

females and males of different ages was determined as


Females. Female pupae were sexed by checking the last

abdominal segment then isolated for emergence in separate


vials. After emergence, 250 females were kept in the

aquarium cages and were aged for 0, 1, 2, 3, and 4 days.

Similarly, 250 male pupae were sexed by examining their last

abdominal segment and the males were aged after emergence in

aquarium cages.

Four to five groups, each of 10 females, of each of the

above ages were allowed to cohabit in 1 pint ice cream

carton cages with 4-day-old males for one night (1800-0900

hrs). The following day, the spermathecae of all females

were dissected in normal saline (0.9 g NaCl/litre) and the

number of inseminated females was recorded. The amount of

sperm and male accessory gland substance (MAGS) transferred

by the males to the spermathecae of females was visually


Virgin males. Two hundred male pupae were allowed to

emerge in an aquarium cage. Males with terminalia rotated

0-450 were isolated into separate cages and provided 10%

sucrose solution and fresh apple slices as sugar. The males

were aged for 0, 1, 2, 3, 5 and 7 days. Single males of the

above ages were allowed females were allowed to cohabit in 1

pint ice cream cartons with 5 virgin 2 to 3-day-old females

overnight. The following morning, the spermathecae of all

females were dissected and number of inseminated females

recorded. Males were saved for later dissections to

determine the effect of age and mating on their reproductive




Figure 3-1.









The reproductive system of male Lutzomyia
anthophora. Distance between brackets
represents the parts of structures measured.


Effects of Age and Mating on the Morphology of Male
Reproductive System

The effect of age on the reproductive system of virgin

males was investigated as follows. Two hundred male pupae

were separated and the males were allowed to emerge. Two

hundred individuals with terminallia rotated 0-450 were

isolated in an aquarium cage and allowed to age for 0, 1, 2,

3, 5 and 7 days. At each age, 10 males were dissected;

different parts of their reproductive system were measured

with a calibrated ocular micrometer under a compound

microscope (Fig. 3-1).

To observe the reproductive system in mated males,

individuals of different ages (0 to 7-days) and matings (1

to 5) were obtained from the mating experiment already

discussed. These insects were dissected by the following

method and different parts of their reproductive system were

measured (Fig. 3-1).

Dissections and measurements. A single live male was

blown from a mouth aspirator into a petri dish containing

dilute detergent solution (dish washing soap). This

procedure removed the majority of scales from the male's

body. The male was immediately transferred into a drop of

normal saline and washed several times to remove the

detergent. Later, the male was transferred to another drop

of normal saline on a glass slide and dissected with fine


dissection needles under a microscope at 30 X. One needle

was placed on the last abdominal segment and the other on

the thorax, allowing excision of the whole reproductive

system by traction on the last abdominal segment.

The following parts (Fig. 3-1) of the male reproductive

system were measured without applying a cover slip (Mahmood

& Reisen 1982; Mahmood et al. 1986): 1) length and width of

testes; 2) length of portion containing the spermatocyst; 3)

length and width of sperm reservoir; 4) total number of

spermatocysts; 5) number of mature spermatocysts

(spermatocysts containing spermatozoa); 6) length and width

of accessory glands; 7) Width of seminal vesicles. Color

and the degree of repleteness of the accessory glands with

MAGS and sperms (if any) were also noted. Similarly, the

amount of sperm in the sperm reservoir and seminal vesicles

was also recorded photographically on a Zeiss microscope.


Mating Behavior of Sand Flies

Virgin 5-day-old males initiated their epigamic

behavior within 0-10 minutes after being placed in the cages

containing the virgin females. The epigamic behavior of the

two sexes consisted of the following steps.

1) The males reached the females following a zig zag

path, during which time they vigorously flapped their wings


and moved their last abdominal segment up and down and

sometimes sideways.

2) After reaching the female, most of the males either

faced the females or were parallel to her, facing in the

same direction.

3) When in contact, the males turned 1800 angle and

coupled tip to tip with the female their heads facing in the

opposite directions. Not all attempts made by the males

were successful. Sometimes the female disengaged herself

and flew away within seconds.

4) Some mating attempts lasted from a few seconds to 4

to 5-minutes. A successful mating was achieved when the it

lasted at least more than 10 minutes. After a successful

mating the female disengaged herself. The males continued

to flap their wings during copulation.

5) Many times virgin females moved near the mating

couple but only as close as 1-5 cm. The presence of another

female did not distract the mating couple.

6) The male attempted to copulate with a new female

within the first five minutes after the completion of first

mating. One male started to mate for the first time 11

minutes after it was placed in a cage with virgin females.

The first mating lasted for 18 minutes. Seven minutes after

the first mating, the male commenced following new females

and attempted to mate with at least four different females.

Although these females disengaged themselves within few

Figure 3-2. Spermathecae of a mated Lu. anthophora.