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Studies of Lutzomyia anthophora (Addis) (Diptera: Psychodidae) and other potential vectors of Rio Grande virus

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Studies of Lutzomyia anthophora (Addis) (Diptera: Psychodidae) and other potential vectors of Rio Grande virus
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Endris, Richard German, 1948-
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Animal nesting ( jstor )
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Blood ( jstor )
Eclosion ( jstor )
Eggs ( jstor )
Female animals ( jstor )
Larvae ( jstor )
Species ( jstor )
Suckling ( jstor )
Sugars ( jstor )

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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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STUDIES OF Lutzomyia anthophora (ADDIS) (DIPTERA:
PSYCHODIDAE) AND OTHER POTENTIAL VECTORS
OF RIO GRANDE VIRUS




By

RICHARD G. ENDRIS


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











UNIVERSITY OF FLORIDA


1982



























Copyright 1982

by

Richard G. Endris





























A man's reach should exceed his grasp.

--Robert Browning








FRONTISPIECE


Lutzomyia anthophora feeding on the ear of its native host,
the woodrat, Neotoma micropus.














ACKNOWLEDGEMENTS


The question of how to thank someone for his years of friend-

ship, guidance, and wise counsel remains enigmatic. During the course

of this program I have incurred many debts which I will hopefully re-

pay through contributions to the science.

The members of my committee I thank for their guidance and sup-

port are Dr. Harvey Cromroy, chairman, Dr. Jerry Butler, Dr. David

Young, Dr. Donald Hall, and Dr. Stephen Zam. Mrs. Adele Koehler

typed the manuscript. Special thanks are in order to

Dr. David Young for suggesting this project, for his constant

support, and for his selfless assistance in the sandfly

colonization efforts;

Dr. Jerry Butler for generously providing a laboratory and much

of the equipment used for this project;

Dr. Robert Tesh, Yale Arbovirus Research Unit, for his generous

sharing of time, equipment, and knowledge that made the

virus transmission experiments possible;

E. Ann Ellis for her long hours of instruction on electron

microscopy and histology;

Diana Simon and Debbie Boyd who facilitated the daily accomplish-

ment of much of this research;

Maj. Peter Perkins with whom many hours of camaraderie were

shared and with whom experimental ideas were generated;









Kristin Figura, Kay Warren, and Annie Moreland for their assistance

in virus purification and titration.

Dr. A.G.B. Fairchild for sharing his vast knowledge and years of

experience.

A grant from the Steffen Brown Foundation provided the opportunity

to study at Yale University in 1981.














TABLE OF CONTENTS


Page

ACKNOWLEDGEMENTS ........ ......................... v

LIST OF TABLES ......... .... .................... ...ix

LIST OF FIGURES .......... ......................... xi

ABSTRACT ........ ... ............................. xiii

GENERAL RATIONALE ....... ........................ .i...1

SECTION

TECHNIQUES FOR LABORATORY REARING OF SANDFLIES (DIPTERA:
PSYCHODIDAE) ....... .......................... 2

Introduction and Literature Review ...... ............ 2
General Techniques ......... .................... 3
Larval Rearing ......... ...................... 5
Sugar Feeding ......... ....................... 5
N-butyl Pthalate ......... ..................... 7
Adult Feeding ......... ....................... 7
Lid Cleaning .......... ....................... 8
Adult Feeding Cages ........ .................... 8
Field Collection, Feeding Containers ............. ....12
Individual Oviposition and Rearing Containers .......... 15
Aspirators ..... ... ........................ ... 15

II COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia anthophora (ADDIS)(DIPTERA: PSYCHODIDAE). .... 17

Introduction and Literature Review ... ............ ...17
Field Collections ..... ...................... ..17
Immature Behavior and Development ................ ....18
Time of Eclosion...... . .................. ...24
Mating ........ .......................... ...24
Female Age at First Feeding .... ................ ...27
Feeding: Hosts ......... ...................... 33
Feeding: Temperature Preference ... ............. ... 34
Feeding: Behavior ... ........ . .. ........... 36
Feeding: Lymph ......... ...................... 39
Refeeding ....... ......................... ...41
Peritrophic Sac Rupture ........................ 41
Productivity ....... ....................... ...42
Longevity ....... ......................... ...45










III COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia diabolica(HALL)(DIPTERA: PSYCHODIDAE) ......... 47

Introduction and Literature Review ... ............ ..47
Field Collection ..... ... ..................... 47
Feeding ........ .......................... 50
Mating ......................................... 51
Egg Hatch and Fertility ....... .................. 51

IV TRANSOVARIAN TRANSMISSION OF RIO GRANDE VIRUS BY
Lutzomyia anthophora(ADDIS)(DIPTERA: PSYCHODIDAE)..... 54

Introduction and Literature Review ... ............ ..54
Materials and Methods ..... ................... ...54
Results ...... ..... .......................... 57
Discussion ..... .. ........................ ...61

V RIO GRANDE VIRUS AND Triatoma gerstaeckeri (STAL)
(HEMIPTERA: REDUVIIDAE) ..... .................. 64

Introduction and Literature Review ... ............ ..64
Materials and Methods ..... ................... ...65
Results and Conclusions ....... .................. 65

VI PURIFICATION OF RIO GRANDE VIRUS ... ............. ...67

Introduction ..... .. ....................... ..67
Materials and Methods ..... ................... ...67
Results ......... .. ......................... 69
Discussion ..... .. ........................ ...69

VII A COMPARISON OF OOCYTE TOPOGRAPHY OF FIVE PHLEBOTOMINE
SANDFLIES (Lutzomyia) WITH THE SCANNING ELECTRON MICRO-
SCOPE (DIPTERA: C HODIDAE) .... ............... ...72
Introduction ..... .. ....................... ..72
Materials and Methods ..... ................... ...72
Results ...... ... .. .......................... 74
Discussion ..... .. ........................ ...78
VIII PHOTOGRAPHIC TECHNIQUES ....... .................. 79

IX SUMMARY ........ .......................... 82

BIBLIOGRAPHY ......... ........................... ..83

BIOGRAPHICAL SKETCH ..... .... ....................... 90


viii














LIST OF TABLES

Table Page

1-1. Dosage required to anesthetize animals for 30-60 min
with Ketamine hydrochloride (100 mg/ml) injected IM. . 9

2-1. Mean duration (days) of immature stages of L. antho-
phora at 90% RH and 4 constant temperature regimes:
20'C, 24C, 28C, and 32C in contrast to the obser-
vations of Addis (1945b) made at 28-29C. Larvae
reared on the diet of Young et al. (1981) .... ........ 22
2-2. Comparison of the effect of larval diet composition pre-
pared by the method of Young et al. (1981) on mean
duration of immature development time (egg-adult) of
L. anthophora at 20'C and 28C, 90% RH ........... .... 23

2-3. L. anthophora--Comparison of mean development time
Tdays) for males and females reared at 200C, 240C,
280C, and 32C, 90% RH ........ ................. 25

2-4. L. anthophora--Adult sugar feeding, frequency, age of
feeding (days), time required for digestion (days) at
20C and 28C, 90% RH ........ .................. 31

2-5. Comparison of effects of blood vs. blood and sugar as
an energy source for L. anthophora fed on Didelphis
marsupialis (opossum) ....... ................ .... 32

2-6. Temperature (C) of body regions of anesthetized and
non-anesthetized hosts for L. anthophora in laboratory
culture .......... ......................... 35

2-7. Fecundity, percent of bloodfed females that laid no
eggs, and preoviposition period (days) for 12 genera-
tions of L. anthophora reared at 24C and 280C,
90% RH ........ ... ......................... 44

2-8. Comparison of longevity (days) of L. anthophora males
and females fed on either distilled water or 30% honey
solution at 240C, 90% RH .... ................ .... 46

3-1. Lutzomyia diabolica--Fecundity, preoviposition period
(days), and mortality factors for 3 generations in
laboratory culture at 280C, 90% RH ............. ....53








Table Page

4-1. Growth of Rio Grande virus in L. anthophora after
intrathoracic inoculation .... ................. ...59
4-2. Presence of Rio Grande virus in (1) Neotoma micropus
and (2) Peromyscus leucopus bled daily for 7 days
after subcutaneous inoculation ...... .............. 60

7-1. Classification of 41 species of Neotropical phlebotomine
sandfly eggs based on oocyte topographic patterns ..... .. 73














LIST OF FIGURES


Figure Page
1-1. Schematic diagram of laboratory rearing techniques for
phlebotomine sandflies ...... .. .................. 4

1-2. L. anthophora feeding on an apple slice .... ......... 6

1-3. Sandfly feeding cage--A modified aquarium with plaster
of Paris bottom and back ...... ................. 10

1-4. Cylindrical adult feeding cage ................. ... 13
1-5. Field collection apparatus, feeding and rearing con-
tainers for phlebotomine sandflies ............... ...14

2-1. Habitat of Lutzomyia anthophora ... ............. ... 19

2-2. Nest of Neotoma micropus .... ................. ... 19

2-3. Multiwell plate with lid used for rearing individual
larvae ...... ... .. .......................... 21

2-4. Eclosion distribution of 127 L. anthophora males and
females from F3 generation in a laboratory colony at
24-C, 90% RH ...... ....................... ....25

2-5. L. anthophora male <24 hrs old with unrotated
genitalia ..... ... ........................ ... 26

2-6. L. anthophora--Temporal age distribution at feeding and
death of unfed females at 28C, 90% RH ............ ....28

2-7. L. anthophora males and females engorged on 30% honey
solution dyed with red, blue, and green food dye ..... ... 30

2-8. Time sequence (20 sec) of L. anthophora feeding on
hamster (Mesocricetus aureus) ear ..... ............ 37

2-9. L. anthophora with mouthparts stuck in the ear of
Peromysus leucopus (white-footed mouse) .... ......... 38

2-10. L. anthophora excreting clear fluid droplets while
feeding ..... ..... ......................... 38








Figure Page

2-11. L. anthophora engorged on serum or lymph ........... ..40

2-12. Dead female L. anthophora after peritrophic sac rupture 40

3-1. Habitat of Lutzomyia diabolica ..... .............. 49

3-2. L. diabolica feeding on human arm ... ............ ..49

6-1. Electron micrographs of purified Rio Grande virus
at 125,000X ..... ... ....................... ..70

7-1. Scanning electron micrographs of eggs of four sandfly
species. (1) Lutzomyia diabolica, (2) Lutzomyia
shannoni, (3) Lutzomyia vexator, (4) Lutzomyia
cruciata spp ....... ....................... ...75

7-2. Scanning electron micrographs of oocyte topography of
five species of sandfly ..... ................. ...76

8-1. Chamber for photographing hematophagous insects feeding
on humans and small mammals .... ............... ...80

8-2. Chamber for photographing small insects .. ......... ..80










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


STUDIES OF Lutzomyia anthophora (ADDIS) (DIPTERA:
PSYCHODIDAE) AND OTHER POTENTIAL VECTORS
OF RIO GRANDE VIRUS

By

Richard G. Endris

May, 1982

Chairman: Dr. Harvey Cromroy
Major Department: Entomology and Nematology

Simple colonization techniques for rearing large numbers of

phlebotomine sandflies were developed. Lutzomyia anthophora (Addis)

and Lutzomyia diabolica (Hall) were colonized in the laboratory for the

first time for 16+ and 7+ generations, respectively, thus permitting

quantitative investigation of their ability to transmit viruses and

leishmaniasis. Notes on field behavior of L. anthophora and L. diabolica

are presented with detailed laboratory studies on the biology of the

two species. Laboratory transovarian transmission of a Phlebovirus

was demonstrated for the first time with L. anthophora when 54.8% of

the F adult progeny from parents infected by intrathoracic inoculation

became infected. Attempts to transmit Rio Grande virus by the bite of

L. anthophora and Triatoma gerstaeckeri (Stal) were unsuccessful.


xiii














GENERAL RATIONALE


The primary goal of this research was to conclusively demonstrate

transovarian transmission of a Phlebovirus in a sandfly for the first

time. A virus (Rio Grande), non-pathogenic for humans, was selected

because it could be safely studied. The potential vector sandfly,

Lutzomyia anthophora, is not anthropophilic and therefore is a safe

subject for study.

Before transmission experiments could be undertaken it was neces-

sary to first develop sandfly rearing and colonization techniques. In

order to plan and execute transmission experiments some aspects of the

laboratory biology of L. anthophora had to be elucidated.

After transovarian transmission was demonstrated, I realized that

this alone could not account for the distribution of neutralizing anti-

body in various animals from south Texas in view of the fact that L.

anthophora apparently does not feed on all of them. In order to more

fully understand the ecology of Rio Grande virus, preliminary studies

of other hematophagous insects, L. diabolica and Triatoma gerstaeckeri,

were undertaken.

In a broad sense it must be acknowledged that no single mechanism

such as transovarian transmission can account for the maintenance of a

virus when the species of interest is sympatric with other hematophages.

Each insect species that feeds on the host must be studied to determine

its relative role in the maintenance of a pathogen.














SECTION I

TECHNIQUES FOR LABORATORY REARING OF SANDFLIES
(DIPTERA: PSYCHODIDAE)


Introduction and Literature Review


The difficulty of efficiently producing large numbers of sand-

flies in the laboratory has hindered studies on their biology and

vector competence for viral and parasitic diseases (Killick-Kendrick

1978). Despite significant contributions by several workers (Chaniotis

1967, 1975, Gemetchu 1976, Killick-Kendrick et al. 1973, Killick-

Kendrick et al. 1977, Ward 1977) several major problems remain. Some

of these include (1) larval mortality due to unknown factors, (2) ex-

cessive labor requirements for colony maintenance, and (3) death of

females at oviposition. Use of the techniques described here have con-

siderably reduced the first two difficulties and partly solved the

third.

Six of the 600 known phlebotomine species, e.g. P. argentipes

Annandale & Brunetti, P. papatasi (Scopoli), L. longipalpis (Lutz &

Neiva), L. sanquinaria (Fairchild & Hertig), L. gomezi (Nitzulescu),

and L. flaviscutellata (Mangabeira), have been reared for 10 genera-

tions or more (Killick-Kendrick 1978, Ward 1977). The following

species have been reared by the methods described here: L. anthophora

(Addis), 15 generations; L. shannoni (Dyar), 15 generations; L. vexator

vexator (Coquillett), 7 generations; L. diabolica (Hall), 5 generations;









L. cruciata spp, 23 generations; L. cayennensis (Floch & Abonnenc),

5 generations. In addition, 3 African phlebotomine species were

reared to the 4th generation using these methods (D. Young, personal

communication).



General Techniques


Figure 1-1 represents the generalized rearing method for sandflies.

An explanation of each step is as follows: Step 1. The plaster of

Paris in a rearing cage is saturated with H20 with no free water re-

maining; Step 2. An engorged female is gently "herded" into the vial.

A drop of 30% honey solution or other sugar source is then placed on

the screen top; Step 3. After 3+ days most females oviposit on the

plaster bottom. If the female survived oviposition she is released

back into the feeding cage. Screen lids are replaced with solid tops

that have small punctures to allow for gas exchange but limit dessi-

cation; Step 4. Since eggs held at 26C usually hatch 6-14 days after

oviposition, a small amount of larval diet is placed in the vial 4-5

days after the eggs are laid; Step 5. Larvae should be checked weekly

and moist medium added as required; Step 6. Adults are released into

the feeding cage daily by placing lidless vials containing pupae in

the feeding cage. Adults soon begin mating and feeding on sugar from

apple slices provided; Step 7. An anesthetized or restrained vertebrate

host is placed inside the cage after a prefeeding period that varies

in time according to species.











I
NOT TO SCALE


I


SLEEVE REMOVED


-0


3



,1


Schematic diagram of laboratory rearing techniques for phlebotomine sandflies.


M..


Figure 1-1.


4000*


81 I








Larval Rearing


Later instar larvae are more tolerant of moisture variation than

earlier instar larvae. When larval medium (Young et al. 1981) is added

to the rearing vials it must be slightly moistened. Larvae can be

reared under conditions of >80% RH but 90-95% is preferable. Even at

this high humidity secondary fungal growth is uncommon, presumably be-

cause the first Rhizopus sp. bloom either exhausts an essential nutrient

or produces a fungal growth inhibitor. It is a primary colonist in

fungal succession and reduces proteins to amino acids and carbohydrates

to simple sugars. After the medium has completely dried, it is re-

moistened. Even then, there is little fungal growth, the hyphae are

not abundant enough to entangle the caudal setae of the larvae.

Mites frequently seen in larval vials have not been observed

attacking healthy larvae but they will feed on weak or dead larvae and

adults. Boiling water poured into vials before reuse will kill any

mites present. Autoclaving larval medium infested by mites for 5 min

at 15 psi will kill mites without apparent damage to the medium.



Sugar Feeding

Ready (1979) provided evidence that sugar feeding is important for

sandfly egg production. Adults were provided sugar ad libidum throughout

their lifetime by two methods.

Adult males and non-bloodfed females were provided sugar from thin

apple slices (<3 mm) leaned against the sides of the feeding cages

(Figure 1-2). Thicker apple slices tend to mold more rapidly than






































Figure 1-2. L. anthophora feeding on an apple slice.









thin ones which tend to dry. A tangential section of each apple slice

should be removed to produce a flat edge so that the slice will not

roll and crush flies. Fresh slices are added daily. Rhizopus sp. is

the mold that usually grows on "old" apple slices and it can be used as

inoculum for larval medium.

Bloodfed females are provided sugar in the oviposition cages by

placing a small drop of 30% honey solution or 50% Karo syrup solution

on the screen lid. A 30% honey solution was used initially in an

effort to produce a facsimile of nectar but a Karo syrup solution pro-

vided equivalent results. If fungal or bacterial growth become ap-

parent in the sugar solution the lid should be changed and a new drop

added. Refrigeration of stock sugar solutions at 3C greatly increases

their shelf life.



N-butyl Pthalate

Clear vinyl suction cups were used to suspend apple slices from

the top of the feeding cage. This practice was quickly abandoned after

adult mortality approaching 100% was associated with the vinyl use.

N-butyl pthalate, an elasticizer used in vinyl, is volatile at room

temperature and is highly toxic to sandflies and mosquitoes (David

Carlson, biochemist, personal communication). The compound has been

used as an insect repellant (The Merck Index 1976).



Adult Feeding

Two general methods for feeding adult females on an animals were

used. The first method is to hold a 7 dram vial or 120 ml specimen








container of flies against an animal's ear or nose allowing the flies

to feed through the screen top. This method is particularly useful for

feeding flies on leishmaniallesions. A mesh size of 18/cm is required

for small species such as L. anthophora and L. diabolica. A larger

mesh size of 10/cm is adequate to contain larger species such as

L. shannoni.

The second method is to place an anesthetized or restrained animal

in the feeding cage. Anesthesia dosage rates are given in Table 1-1.

Anesthesia was administered with a 1 ml Tuberculin syringe and a 26 or

27 gauge needle. Animals in poor condition require less anesthesia.

An insufficient dose will sometimes produce hyperactive behavior.



Lid Cleaning


Screen tops that have been used for sugar feeding are cleaned by

soaking in 5% Chlorox solution for 30 min, rinsing 2x in tap water for

30 min, and air dried. Tops can be reused many times until screening

material breaks or glue becomes brittle and non-adhesive. Use of a

more concentrated Chlorox solution or a longer soaking time results

in rapid deterioration of screen material and glue greatly reducing

the number of times lids can be reused.



Adult Feeding Cages


The most successful adult feeding cages developed were constructed

from 4 (26 x 20.5 x 16.5 cm), 6 (31.0 x 20.5 x 16.5 cm), and 12

(36 x 25.5 x 21.5 cm) liter aquariums (Figure 1-3).










Table 1-1. Dosage required to anesthetize animals for 30-60 min with
Ketamine hydrochloride (100 mg/ml) injected IM.


Animal Dosage (ml)


Mouse 0.05/adult

Squirrel 0.1/200 gm

Woodrat 0.2/adult

Opossum 0.25-0.30/2 kg

Rabbit 0.3-0.4 mg/kg














































ONE-THIRD SCALE


Figure 1-3. Sandfly feeding cage--A modified aquarium with plaster of Paris bottom and back.





-11-


The bottom and one side of the aquarium were covered with a 1 cm

layer of plaster of Paris. After the bottom has been poured and allowed

to dry it is imperative that it be saturated with water before the side

layer of plaster is poured. This will prevent the formation of un-

workable lumps at the junction of the two pours due to immediate des-

sication of the wet plaster by the dry layer. After the bottom and one

side have been poured the bottom should be rewet and the upper corners

filled in to a maximum depth of 2 cm. This allows for easy viewing of

flies and easy recovery of flies with an aspirator.

Front panels for the cages are constructed of 64 mm (1/4") Plexi-

glas. Screens of 18 mesh/cm of "Saran" (Chickopee Co., Cornelia, GA)

vinylidene polymer plastic are installed on the front panel. Experience

has shown that this is necessary because otherwise, excessive conden-

sation in the chamber will form when animals are left in for sandfly

feeding. The Saran screen is attached with epoxy cement. Care must

be exercised that epoxy components are not out of date and are well

mixed; otherwise the glue will remain sticky and trap the flies. Other

glues tried, i.e., contact cement, Elmer's glue, silicone, and super-

glues, do not adhere well to the Plexiglas. The minimum screen areas
2
are 78, 130, and 214 cm respectively.

A 50 cm sleeve of 15.3 cm (6") surgical stockinet (Johnson &

Johnson) is attached to the front panel by compression between the panel

and a Plexiglas frame (2.5 cm wide). This is secured with 8 (10/24 x

1") brass screws with flat washers and wingnuts. The brass screws and

flat washers are glued inside the front panel with epoxy glue in order

to facilitate changing of the sleeve which should be secured with tape

while the frame is being installed. The completed front panel is





-12-


attached to the cage with a thick layer of silicone glue that can be

easily cut away for repairs. It is essential to fill all small crevice,

with plaster of Paris or silicone to prevent adult sandflies from hiding

there and being difficult to recover.

A cylindrical adult feeding cage (Figure 1-4) was constructed from

a cylindrical (23 x 13 cm) glass fixture cover (Appleton Co. V-51).

Four centimeters of plaster of Paris were poured in the end and 1.5 cm

(tapered to the front) were poured on a side of cylinder by the method

described. The frame was constructed of 3 (18 cm x 18 cm x 64 mm)

Plexiglas plates and 18 cm (10/24) threaded steel rod. Relative posi-

tions of the plates is maintained by placing nuts and washers on both

sides of the sheets which are attached to the glass by a bead of silicone

glue. A 50 cm stockinet sleeve is secured to the front by compression

between 2 plates as with the rectangular feeding chamber. The advan-

tages of the cage include small size and ease of manufacture. Dis-

advantages are difficulty seeing through the glass, condensation due

to animal respiration, and difficulty in manipulating vials inside the

chamber.


Field Collection, Feeding Containers


The 120 ml specimen containers (Pharmaseal Laboratories, Glendale,

CA 91201) are modified for use as field collection and feeding con-

tainers (Figure 1-5).

Field collection containers are constructed as follows. Two

centimeters of plaster of Paris are poured in the bottom of the

containers; then a 2 cm entry port is cut in the container side

by heating a #15 cork borer then pushing it through the plastic




-13-


Figure 1-4. Cylindrical adult feeding cage.





-14-


PLASTER
--- OF PARIS





120 ML SPECIMEN CONTAINER

FULL SCALE


SCREEN LID



SOLID LID


7 DR VIAL

FULL SCALE


ASPIRATOR


ONE-THIRD SCALE


Figure 1-5.


Field collection apparatus, feeding and rearing containers
for phlebotomine sandflies.


tni





-15-


which should be done in a well ventilated area to avoid noxious fumes.

The edges of the hole are then filed smooth and pieces of latex surgi-

cal glove are glued to both sides with contact cement. If prepowdered

surgical gloves are used they must be washed in a 70% ethanol solution

to remove the powder to insure adhesion. Perpendicular cuts are made

in the respective latex pieces producing a fly-proof opening for the

insertion of an aspirator. Screen lids should be used on the containers

when used for collecting vials. These containers can also be used for

feeding flies. Screen lids are prepared by cutting a 4 cm hole in the

plastic top and attaching the desired mesh screen with contact cement.



Individual Oviposition and Rearing Containers


These containers are made by pouring 1 cm of plaster of Paris in

the bottom of a 7 dram plastic snap cap vial (Fisher Scientific Co.,

Pittsburgh, PA). When used for rearing containers the plastic tops

are punctured to facilitate limited gas exchange. When used as feeding

containers the lids are cut out with a #12 cork borer (1.5 cm hole)

and covered with the desired mesh screen that is secured with contact

cement. When the vials are inverted the plaster occasionally slides

down crushing the insects. This can be prevented by pushing a hot pin

through the plastic into'the plaster then cutting off the excess.



Aspirators


Aspirators for collection and transfer of adults are constructed

of thick wall latex tubing (10 mm ID x 15 mm OD x 60 cm) and thickwall

Pyrexoglass tubing (12 mm ID x 15 mm OD x 30 cm) (Figure 1-5). The





-16-


latex tubing and glass are attached by a piece of hard plastic tubing

(9 mm OD x 5 cm) covered with nylon organdy cloth on one end and

secured with contact cement (Roberts Consolidated Industries, City of

Industry, CA). The screened end is inserted into the glass tubingwhere it is

held by friction and the latex tubing is pushed over the other end.

The latex/glass junction is securely taped so that the end of the plastic

tube is visible in the glass tubing.

Thickwall latex tubing is used for flexibility and to prevent

kinks from occluding the passageway. Pyrex glass is used instead of

plastic because plastic scratches easily making identification of speci-

mens difficult. The inside diameter of any aspirator used for phlebo-

tomine sandflies should be at least 10 mm because smaller diameters

at the same suction pressure result in much higher intake velocities

that cause damage to the flies.














SECTION II

COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia anthophora (ADDIS)(DIPTERA: PSYCHODIDAE)



Introduction and Literature Review


Lutzomyia anthophora was first collected while feeding on rabbits

in Uvalde, Uvalde Co., Texas (Addis 1945a). Subsequently it was re-

ported from NE Mexico (Fairchild and Hertig 1956), SW Mexico (Vargas

1952), and SE Texas (Young 1972). Young (1972) reported finding

L. anthophora in the nest of the plains woodrat, Neotoma micropus,

with which it appeared to enjoy a close host-parasite relationship.

Calisher et al. (1977) again reported the association of L. anthophora

and Neotoma nests when suggesting that Rio Grande virus could be main-

tained in the woodrat population by transovarian transmission in this

sandfly.

Addis (1945b) described the immature stages and the life cycle

of L. anthophora after rearing 72 flies from egg-adult at 28-29C.

In this section detailed investigations of the colonization and biology

of L. anthophora are reported.



Field Collections


Sandflies used to start the colony were collected by R.G. Endris

and D.G. Young with the assistance of G.B. Fairchild and R.N. Johnson

in the area of E and NE of Brownsville, Texas, from Neotoma nests in


-17-





-18-


May and June 1980. Vegetation was characterized by grasses, low grow-

ing shrubs, mesquite, and acacia trees common to xeric regions (Figure

2-1). Climatic conditions were quite dry at the time of collection;

however, a rainy season occurs in August and September.

Johnson (1966) described the structure of woodrat nests in detail.

The nests (Figure 2-2) were carefully disassembled and sandflies were

collected with tube aspirators when seen hopping on the sticks. Flies

were also recovered from under boards covering rodent burrows in a

refuse dump. In both sites the soil was powder dry and the flies'

moisture source remains an enigma. Aspirators and field collection

containers have been described in Section I as well as methods for

feeding freshly caught flies on hamsters.

As woodrats attempted to escape from their nests they were captured

as a blood source for flies. Other woodrats and white-footed mice,

Peromyscus leucopus, which also occupy woodrat dens, were trapped in

Sherman traps.


Immature Behavior and Development


The rearing methods for establishing and maintaining the colony

are described in Section I and by Young et al. (1981). Johnson

and Hertig (1961) and Hanson (1968) grouped Neotropical phlebotomine

larvae into two behavioral groups, i.e., those that burrow into the

larval medium and those that are surface feeders. This behavior

indicates where larvae may be found in nature, i.e., on the soil sur-

face or burrowing beneath it. In the laboratory L. anthophora larvae

exhibited no distinct behavioral preference and the degree of larval





-19-


Figure 2-1.


Habitat of Lutzomyia anthophora.


Nest of Neotoma micropus.


Figure 2-2.





-20-


burrowing appeared dependent on moisture content of the medium and the

stage of development.

Larval emergence, behavior, and pupation were consistent with the

observations of Chaniotis (1967), Johnson and Hertig (1961), and

Gemetchu (1976). In contrast to the observation of Killick-Kendrick

et al. (1977) with L. longipalpis, no cannabalism was observed among

4th instar larvae when starved. No effort was made to discover the

larval habitat in nature although it is presumably in or under the

woodrat nest (Young 1972).

Larval development rates at 4 different temperatures (20C, 24C,

28C, and 32C) were determined by rearing individual larvae in wells

of microtitre plates that were 1/3 filled with plaster of Paris.

Initially microtitre plates with 96 wells were used but the wells

proved too small for 4th instar larvae. Twenty-four well microtitre

plates were satisfactory. Attempts to use the lids designed for the

multiwell plates were not successful because they do not seal well and

larvae moved between wells. Lids (9 cm x 13 cm x 5 mm) made from

Plexiglas and secured with elastic bands solved this problem (Figure

2-3). After several days in chambers at 90% relative humidity (RH) it

was necessary to add drops of H20 in each well until the plaster

appeared damp.

Development times for immatures are presented in Table 2-1. The

difference between the results of Addis (1945b) shown in Table 2-1

and those obtained in this study at 28C are probably attributable to

differences in larval diet.

Three larval diets at 28% and two at 20C were compared to determine

the effect of diet on immature development time (Table 2-2). The diet pre-

pared with Purina Rabbit Chow #5315 is that reported by Young et al. (1981).





-21-


Figure 2-3.


Multiwell plate with lid used for rearing individual
larvae.









Table 2-1.


Mean duration (days) of immature stages of L. anthophora at 90% RH and 4 constant temperature
regimes: 20'C, 240C, 28C, and 32C in contrast to the observations of Addis (1945b) made at
28-29C. Larvae reared on the diet of Young et al. (1981).


Larval Instars
Temperature Total
(0C) Egg 1 2 3 4 Pupa (egg-adult)

20 15.60.5 12.73.3 8.52.5 9.94.7 22.06.9 16.83.3 83.39.3
(59)* (52) (47) (42) (35) (19) (19)

24 10.00.6 6.30.8 5.00.8 5.20.3 11.3+1.0 11.50.4 49.52.0
(174) (165) (162) (160) (133) (130) (130)
28 8.00.1 3.90.8 3.11.1 8.31.0 8.72.2 8.00.8 36.12.9
(41) (39) (39) (39) (36) (36) (36)
28-29 10.5 28.4 8.7 49
(Addis) (72)
32 6.70.6 3.71.3 3.80.8 3.91.2 8.82.3 6.70.8 33.5+3.7
(104) (93) (92) (92) (90) (88) (88)


( ) indicates number of individuals surviving each stage.





-23-


Table 2-2.


Comparison of the effect of larval diet composition pre-
pared by the method of Young et al. (1981) on mean duration
of immature development time (egg-adult) of L. anthophora
at 200C and 28C, 90% RH.


Diet Component
Temperature Rabbit Chow Horse Chow Laboratory Chow
(0C) Purina #5315 Purina #3501 Number (?)

20 83.39.3 99.66.0
n = 19 n = 18

28 36.12.9 38.54.2 43.54.1
n = 36 n = 34 n = 60





-24-


Time of Eclosion


It is a general observation that the males of many insect species

begin eclosion before the females in order to be reproductively mature

when the females emerge. I noted this to be the case with L. anthophora

(Table 2-3), because mean development time from egg-adult was about 2

days less for males than females at temperatures above 20'C. This is

noteworthy because males are not reproductively competent until 24 hrs

post eclosion.

In order to demonstrate this phenomenon the sex and time of eclo-

sion for all individuals from a cohort of the F3 generation were recorded.

The eclosion distribution of males and females is presented in Figure 2-4

and demonstrates the veracity of this observation. Sex ratios were 1:1.



Mating


Male genitalia rotate (Figure 2-5) about 12-24 hrs post eclosion

after which they were observed mating. Females were seen mating within

hours after eclosion and before, after, and during feeding. Mating

frequency was not determined for either sex although males do mate more

than once per lifetime. Copulation occurred regardless of nutritional

state or photoperiod.

L. anthophora males demonstrated the "characteristic epigamic

pattern" described by Chaniotis (1967). Mating usually lasted 2-5 min.

Based on the criterion of egg fertility more than 85% of the females

that laid eggs had successfully mated. In two generations studied the

percentage of females laying infertile eggs was 13.7% and 16.7% in the

F8 and the F15 generations, respectively.






-25-


Table 2-3.


L. anthophora--Comparison of mean development time (days)
for males and females reared at 20', 24C, 280C, and
32C, 90% RH.


Temperature ('C)

Sex 20 24 28 32

Female 83.37.0 50.73.6 40.63.7 34.64.0
n = 10 n = 66 n = 73 n = 36
Male 83.311.7 48.24.1 37.53.3 32.73.3
n = 9 n = 63 n = 57 n = 52

Difference 0.0 2.5 3.1 1.9


L1~


0 MALES
0 FEMALES


i1~ PP. .P


NUMBER OF DAYS IN ECLOSION PERIOD


Figure 2-4.


Eclosion distribution of 127 L. anthophora males and
females from F3 generation in a laboratory colony at
24-C, 90% RH.


20-

V)
I-
g15-
LiL
0

M 10-
D
z


10





-26-


Figure 2-5. L. anthophora male <24 hrs old with unrotated genitalia.





-27-


Female Age at First Feeding


To determine when L. anthophora females were physiologically ready

to take a blood meal, all adults from the F1, generation that eclosed

each day were held in the cylindrical feeding chamber. Each day an

anesthetized mouse was placed in the chamber for 60 min until all the

females in each group either fed or died. Of 244 females, 92 (60.5%)

fed within 1-7 days. The mean age at feeding was 3.7 days and the median

age was 3.5 days. Addis (1945b) noted that females fed 2-4 days post

eclosion. The temporal distribution of female age at feeding and the

age of death for those flies that did not feed is presented in Figure

(2-6).

The sugar feeding habits of hematophagous diptera are well known

and most previous attempts to colonize sandflies included the provision

of 30% sucrose solutions for the adults. Chaniotis (1974) and Ready

(1979) investigated sugar feeding in phlebotomine sandflies. Chaniotis

(1974) studied sandfly preference for various sugars and determined

that sugar concentration had no effect on fly longevity. Ready (1979)

found higher egg production among bloodfed females fed on sucrose solu-

tion vs. those fed on water. Killick-Kendrick (1979) suggested that

the presence or absence of sugar in the gut may have profound and

largely undetermined effects on the ability of sandflies to transmit

leishmaniasis.

An experiment was conducted to determine what percentage of adults

will ingest honey solutions, at what age, frequency of sugar feeding,

and the time required to digest each meal. Individual pupae were placed

in 7 dram feeding vials. The day of eclosion a small drop of 30% honey






























-J
ct

0

IO
a, C
z10-













0 2 4 6 8 10 12
NUMBER OF DAYS POST ECLOSION








Figure 2-6. L. anthophora--Temporal age distribution at feeding and death of unfed females
at 28oC, 90% RH.





-29-


solution mixed with either red, blue, or green food dye (C.F. Sauer Co.,

Richmond, VA) was placed on the screen lid. After a sandfly ingested

the solution the dye was clearly discernible in the distended crop with

or without the aid of a microscope (Figure 2-7). Yellow dye was not

used because it could not be seen. When an individual digested the

sugar solution a small colored droplet was excreted and no color re-

mained in the abdomen. Flies would not refeed until the previous meal

had been completely digested.

Results for sugar feeding experiments conducted at 20'C and 28C

are shown in Table 2-4. The reduction in number of flies feeding more

than 1x at 28C indicates the release of individuals back into the

breeding colony rather than mortality. Of the flies offered honey

solution at 28C, 93.2% (68/73) fed within 24 hrs after eclosion.

Of 23 flies held until death at 280C, 100% fed 1x, 30.4% fed 2x, 30.4%

fed 3x, 34.5% fed 4x, and 4% fed 5x.

The number of sugar fed adults held at 200C declined rapidly due

to poor survival. Digestion of the first and second sugar meal at 20C

requires more time than at 28C.

Results of a second experiment to determine the effect of sugar

feeding on percentage of females feeding on blood, fecundity, preovi-

position period, mating (fertility of eggs), and mortality factors are

presented in Table 2-5. Sugar feeding enhanced productivity for all

parameters measured.

The source of sugars for L. anthophora in nature is unknown. Many wood-

rat nests are located around clumps of prickly-pear cactus (Opuntia lind-

heimeri), a succulent that woodrats feed on in their nests. Sandflies

may obtain sugar from partly eaten cactus in the woodrat nest. To test





-30-


Figure 2-7. L. anthophora males and females engorged on 30% honey
solution dyed with red, blue, and green food dye.










Table 2-4.


L. anthophora--Adult sugar feeding, frequency, age of
digestion (days) at 20'C and 28C, 90% RH.


feeding (days), time required for


Tempera- Age at # Days to # Days to # Days to # Days to # Days to # Days to # Days to # Days to
ture I Feed Digest 2' Feed Digest 30 Feed Digest 40 Feed Digest 50 Feed
(0C) 10 Meal 20 Meal 30 Meal 40 Meal

280 1.30.8 1.40.8 1.41.1 1.30.5 1.50.7 1.60.9 1.81.1 1.30.5 1.0
n = 68 n = 48 n = 29 n = 25 n = 17 n = 16 n = 10 n = 6 n= 1

200 1.91.0 2.21.2 1.20.5 1.80.8 1.20.4 1.0 1.0
n = 17 n = 10 n = 8 n = 6 n = 2 n = 2 n = 1











Table 2-5.


Comparison of effects of blood vs. blood and sugar as an energy source for L. anthophora fed
on Didelphi.s marsupialis (opossum).


Energy % Females % Bloodfed Mean Number Preoviposition % Fertility % Females % Females
Source Fed on Females Eggs/Female Period (Days) of Females Peritrophic Fed on
Blood Laying Laying Eggs Sac Rupture Serum
No Eggs

Blood 40.8 39.6 17.68.4 4.51.2 65.2 11.7 1.7
(147)* (60)
Blood + 49.7 19.8 28.013.2 6.21.8 83.3 4.9 0.7
Sugar (282) (116)


) indicates number in sample.





-33-


this hypothesis a piece of Opuntia was sliced and placed in a feeding

cage with L. anthophora. Specimens were observed feeding on the plant

juice but not as avidly as on apple slices.


Feeding Hosts


L. anthophora has been reported to feed on rodents and lagomorphs

(Addis 1945a, Young 1972, and Calisher 1977). L. anthophora fed readily

on the following anesthetized animals introduced into the feeding cage:

Neotoma micropus (woodrat; Figure 2-7), Peromyscus leucopus (white-

footed mouse), Mesocricetus auretus (Syrian hamster), Sciurus carolinen-

sis (grey squirrel), Mus musculus (white mouse; Figure 2-8), Cavia por-

cellus (guinea pig), Oryctolagus cuniculus (domestic rabbit), and

Didelphis marsupialis (opossum). The preferred feeding site was the

nearly hairless portions of the ears. Fewer than 5% of the sandflies

fed on the feet or among the vibrissae on the nose.

It is notable that L. anthophora would not feed on suckling mice

when restrained with a cloth net on a tongue depressor. Attempts to

feed the sandflies on the poikilotherms, Gopherus polyphemus (gopher

tortoise) and Anolis carolinensis (anole), were unsuccessful.

Flies held in feeding cages on the ears of Canis familiaris (dog),

and Ovis aries (sheep), did not feed. Efforts to feed the flies on Homo

sapiens (human) were not successful. L. anthophora did feed on the

ear of a 2 day old calf (Bos taurus) and on a shaved chick (Gallus

gallus) restrained in a feeding cage.

When the sandflies did not feed on the preferred host they were

subsequently offered either a.n opossum, woodrat, hamster, or mouse.





-34-


This was done to verify that the imagos were ready to feed. In each

instance flies that refused the first host fed upon the second host.

It is apparent from the various species of mammals fed on that

L. anthophora is a more opportunistic feeder than was previously

recognized.


Feeding: Temperature Preference


On each of the host animals used several body regions were rela-

tively hairless, i.e., the ears, nose, tail, and feet. To determine if

relative temperature of the body regions influenced sandfly feeding site

preference the skin temperature of these areas was measured with a

BAT-4 Laboratory Thermometer (Bailey Instrument Co.) and a thermistor

(Table 2-6). The instrument was calibrated to human body temperature

of 37.2-37.4C.

More than 95% of the sandflies fed on the ears where the skin

temperature range was from 27.7C to 37.8C. Although the temperatures

of the tail, foot, and nose were within this range little feeding oc-

curred. Dermal temperature does not seem to be a determining factor

in fly feeding site preference.

Chaniotis (1975) reported that suckling mice were the least

satisfactory source of blood meals for L. trapidoi (Fairchild and

Hertig), a species which feeds on a wide variety of mammalian hosts.

Gemetchu (1976) reported that P. longipes (Parrot and Martin) which

normally feeds on humans would not feed on suckling mice. Although

L. anthophora feeds readily on adults of all rodent species offered,

it would not feed on suckling mice (SM). When the SM were placed in

the feeding cage their body temperature rapidly declined. In an effort









Table 2-6. Temperature (C) of body regions of anesthetized and non-anesthetized hosts for L. anthophora
in laboratory culture.


Animal

Neotoma micropus
(anesthetized)

Neotoma micropus
(non-anesthetized)

Mesocricetus auretus
(anesthetized)

Mesocricetus auretus
(non-anesthetized

Oryctolagus cuniculus
(non-anesthetized)


Tip

31.5


31.3


27.7


32.0


35.0


Ear

Middle

31.1


32.3


30.3


32.9


36.8


Base

34.2


37.1


34.1


37.2


Tail

Tip Base

32.4 32.4


31.3 36.8


33.2 35.0


34.2 35.6


Rear Foot

33.1





35 .*4


36.2


Nose

29.7


30.0


27.8


27.7


31.4


Back

37.0


37.8


36.9


36.9


31.5





-36-


to induce sandfly feeding the SM were placed on a cotton pad on a

variable temperature plate. Flies were released into a specimen con-

tainer over the SM. During a 6 hr period the temperature was raised

from 26C (ambient) to 40'C then returned to 26C. Of 30, 3 day old

adults none fed. A thermistor was taped to the SM to insure the skin

temperature was the activation source for the heater.

The reason for the failure of L. anthophora to feed on suckling

mice remains unknown.

Part of the stimulus for inducing feeding seems not to be the

temperature of the host but rather the differential between host tem-

perature and ambient temperature.



Feeding: Behavior


Within 2-5 min after initiating feeding,L. anthophora females

fed to repletion (Figure 2-8). The time required for feeding did not

change significantly due to host differences.

However, in a few instances it was noted that the flies failed to

withdraw their mouthparts from the ear of the host. This phenomenon

was observed only with Neotoma and Peromyscus which had been fed upon

repeatedly (Figure 2-9). It may be due to the development of a host

immune response to sandfly salivary products which prevented the fly

from withdrawing its proboscis. This phenomenon has been the subject

of considerable investigation with Ornithodorus coriaceus (Theresa

Haslett and Michel Laviopierre, personal communication, 1981).

Diuresis to reduce excess water and concentrate the blood meal has

been observed in L. anthophora (Figure 2-10) as the excretion of clear





-37-


Figure 2-8.


Time sequence (20 sec) of L. anthophora feeding on
hamster (Mesocricetus auretus) ear.





-38-


Figure 2-9. L. anthophora with mouthparts stuck in the ear of
TPeroryscus leucopus), white-footed mouse.


Figure 2-10.


L. anthophora excreting clear fluid droplets while
feeding.





-39-


fluid droplets from the anus while feeding. Chaniotis (1967), Gemetchu

(1976), and others have also observed this phenomenon in Phlebotomines.



Feeding: Lymph


Regardless of the host, 3.2-5.6% of the sandflies in a given cohort

engorged with a clear fluid that is presumably serum or lymph (Figure

2-11). This behavior indicates that some individuals may have the

ability to filter erythrocytes from the blood while feeding or else

they may feed by chance from lymphatic capillaries. The latter explana-

tion is feasible since sandflies are telmophages (Lewis 1975) and lymphatic

capillaries are numerous in the dermis. Ready (1978) noted that L.

longipalpis (Lutz & Neiva) was a non-selective feeder and would feed

to engorgement on isotonic saline and whole blood with equal avidity.

This may also be true of L. anthophora. Of the 20 serum fed individuals

studied in 3 generations 93% did not lay eggs and the maximum number of

eggs laid per female was 12 of those that did. In contrast the mean

egg production for bloodfed females from the same generations was 30.5.

The erythrocyte blood fraction apparently contains nutrients essential

for egg production. Ready (1979) found that the concentration of pro-

tein ingested had a significant direct relationship to the number of

oocytes produced and that the red cell fraction was more important than

plasma for egg production. The production of 12 oocytes or less by

lymph fed L. anthophora contrasts with the conclusion of Adler and

Theodor (1926) that plasma alone was essential for P. papatasi to pro-

duce eggs.





-40-


Figure 2-11.


Figure 2-12.


L. anthophora engorged on serum or lymph.


Dead female L. anthophora after peritrophic sac
rupture.





-41-


Refeeding


Investigations on the vector capability of sandflies have been

hampered by the failure of females to survive oviposition. Conse-

quently, demonstration of transmission of Leishmania and Phleboviruses

by bite has been difficult to establish (Killick-Kendrick 1979). Kil-

lick-Kendrick (1979) and Johnson and Hertig (1961) discussed those

species which feed more than once in the laboratory.

In the F6 and Flo generations of L. anthophora maintained in

laboratory culture, the individuals that laid eggs within a 24 hr period

were released into a feeding cage and offered an anesthetized host for

a 30-60 min period daily. Of the flies in those respective generations

19.1% and 16.1% took a second blood meal. Four individuals fed 3x in

the F6 generation. These results are consistent with the observations

of Schmidt and Schmidt (1965) on P. papatasi.

No experiments were conducted to discover an optimum oviposition

site. If this were determined perhaps a much higher percentage of

blood fed females would oviposit, survive oviposition, refeed, and

repeat the gonotrophic cycle. Most flies that died prior to oviposi-

tion retained eggs in the abdomen. It is unlikely that such high

mortality occurs at oviposition in wild populations.



Peritrophic Sac Rupture


The term "membrane" when used to describe the lattice-like sac

which surrounds the blood meal in hematophagous insects is a biological

misnomer since it is not a trilaminate phopholipid/protein membrane.

I suggest adoption of the term "peritrophic sac." Romoser and Rothman





-42-


(1973), Romoser (1974), Romoser and Cady (1975) described the lattice-

like sac in mosquito larvae, pupae, and adults. The similar structure

of the sandfly peritrophic membrane was investigated in detail by Gemetchu

(1974). He states that it is formed within 30 min after a blood meal is

taken and breaks up about 3 days later.

A phenomenon that occurred consistently in each generation was

the apparent rupture of the peritrophic sac and the midgut epithelium.

Blood from the gut penetrated all parts of the insect including the

thorax, legs, and antennae (Figure 2-12). The specimens that were killed

by this phenomenon possibly died as a result of changes in the hemolymph

osmoticum and the release of digestive enzymes. In 3 generations (F8,

F9, Flo) the frequency of "peritrophic sac rupture" occurred in 9.9%,

6.6%, and 8.7% of the bloodfed sandflies, respectively. Death of the

flies followed feeding in 1-4 days with 75% dead in less than 24 hr.

This phenomenon is not limited to L. anthophora since I have also

observed it in Ornithodorus turicata, Ornithodorus dugesi, Triatoma

gerstaeckeri, Triatoma sanguisuga, Triatoma neotomae, and Lutzomyia

diabolica. The mechanism of gut integrity disruption remains an enigma

and does not seem related to the mammalian blood source.


Productivity


Egg production for 12 of 15 generations of L. anthophora reared

to date are presented in Table 2-7. The preoviposition period indi-

cated in the table represents the time from blood meal ingestion to

egg laying, i.e., the period required for egg development. Since

sandflies often extrude 1-3 infertile eggs at death only those females





-43-


which laid four or more eggs were considered to have oviposited. No

autogeny was observed with this species nor could it be induced by feed-

ing mated females only water or sugar solutions. Egg laying was usually

completed in less than 24 hr but could require 2-3 days. Additional

blood meals are required for females to lay subsequent batches of eggs.

The first 3 generations were held at 24C and subsequent genera-

tions were held at 28C with no apparent effect upon egg production. Vary-

ing photoperiods also had no apparent effect on egg production or fertil-

ity. The effect of blood meal source on fecundity was not investigated.

The number of females which did not take a blood meal was studied

in the F15 generation and observed to represent 50.3% of the total

number eclosed. Those females plus the 19.8% blood fed females which

did not lay eggs indicate that 58.5% of the total number of females in

a given generation are non-productive. Despite the number of non-

productive females, colony numbers could easily be increased to yield

as many insects as required for experimentation.

A phenomenon that was consistently observed in each generation

was the tendency for those females eclosing in the first half of the

generation cycle to lay the majority of the eggs and for many of those

emerging in the second half of the generation cycle to die without

ovi positing.

An attempt to maintain adults at 320C proved unsatisfactory since

of 36 females produced at that temperature, 11 fed on a mouse (69.4%

did not feed), 4/11 (36.4%) laid eggs, and 3/11 (27.3%) fed on serum (no

eggs). The mean number of eggs per female was 13.8 9.8. The number

of eggs laid by adults held at 320C was greatly reduced compared to

those held at 240C and 280C (Table 2-7).









Table 2-7. Fecundity, percent of bloodfed females that laid no eggs, and preoviposition period (days)
for 12 generations of L. anthophora reared at 24C and 280C, 90% RH.


Fecundity % Bloodfed Preoviposition Period (Days)
Females Laying
Generation n x S Maximum # No Eggs x S Range

F1 25 42.0 20.5 71 28.0 5.2 1.4 3-11

F2 99 34.7 21.6 79 53.5 5.5 1.3 4-7

F3 171 31.7 16.9 64 66.2 7.1 2.0 4-16

F4 80 39.5 19.0 73 53.3 5.3 1.1 4-8

F5 61 43.2 13.1 65 43.3 6.4 1.8 4-10

F6 94 40.6 23.2 107 50.0 6.2 1.3 4-10

F7 124 36.1 14.3 68 65.3 5.6 0.8 4-6

F8 121 32.2 18.3 75 45.6 4.5 1.7 2-9

F9 61 32.9 16.5 63 27.9 4.8 1.0 4-8

F10 92 32.5 17.8 73 32.6 5.8 1.5 2-9

F11 190 26.1 13.1 56 21.8 6.2 1.5 3-11

F15 125 28.0 13.2 56 19.8 6.2 1.8 3-11

Overall 1203 35.0 17.3 70.8 38.5 5.7 1.3 2-11
Means






-45-


Longevity


Adult longevity for males and non-bloodfed females was determined

by holding sandflies individually and placing either distilled water or

30% honey solution on the screen lid. Results are presented in Table

2-8. Sugar-fed adults lived 40-45% longer than those fed on distilled

water only.

These results agree with the work of Nayar and Sauerman (1975a,b)

and Edmund Davis (personal communication, 1981) who showed that sugar

feeding increased longevity of several mosquito species and Culicoides

mississippiensis (Hoffman), respectively.





-46-


Table 2-8.


Comparison of longevity (days)
fed on either distilled water
24C, 90% RH.


of L. anthophoramales and females
or 30% honey solution at


30% Honey Solution Distilled H20

Sex x S x S

Female 10.4 2.9 7.7 1.6
(31)* (18)
Male 10.1 3.1 7.1 1.5
(32) (16)

Total 10.2 3.0 7.4 1.6
(63) (34)


( ) indicates number of individuals tested.













SECTION III

COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia diabolica (HALL) (DIPTERA: PSYCHODIDAE)


Introduction and Literature Review


Lutzomyia diabolica (Hall 1936) has long been recognized as a pest

in South Central Texas (Parman 1919, Lindquist 1936) where it bites

humans in and near human dwellings. The status of the species was ques-

tioned by Disney (1968) but has been recently resolved by Young and

Perkins (1982). Lindquist (1936) studied the life cycle of the species,

described the immature stages but did not establish a laboratory colony.

Addis (1945a) made an unsuccessful attempt at colonization. Parman

(1919) described the bite on humans in detail and suggested that L.

diabolica may be a vector of a transient febrile human illness.

Several cases of autochthonous leishmaniasis have been recorded

from Texas (Shaw et al. 1976, Simpson et al. 1968, Stewart and Pilcher

1945, and Anderson et al. 1980). Since L. diabolica is the only known

man-biting sandfly in the region it is highly suspect as a potential

vector.


Field Collection

Although L. diabolica was first taken from Uvalde, Texas, it is

widely distributed in Northern Mexico (Najera 1971) and Texas (P.V.

Perkins and D.G. Young, personal communication, 1982). For establishment


-47-





-48-


of a laboratory colony, I collected specimens in July 1981 at Garner State

Park, a site located approximately 50 km N of Uvalde, Uvalde Co., Texas,

in the Frio river valley near the eastern edge of the Edwards Plateau.

The habitat is characterized by open grassland interspersed with oak,

acacia, and cedar trees surrounded by rocky hills (Figure 3-1).

All specimens were taken with a tube aspirator and held in field

collection vials. The first specimen was taken while feeding on a clerk

in the park office at 1800 hrs July 7, 1981. No sandflies were captured

with four CDC Light traps set 1 m above ground level in protected areas

along 0.5 km of the Frio river for 1 night, and within 20 m of the build-

ings from which sandflies were taken for a second night. Other workers

have taken L. diabolica in CDC Light traps (D.G. Young, personal

communication).

Meteorological conditions for the night of 7 July 1981 were 100%

overcast with intermittent rain, 25C, with winds gusting to 15 knots.

I noticed L. diabolica feeding on my arms at 2200 hrs while sitting near

a light in an open shower/latrine building in an open area with a few

adjacent acacia trees. About 40 females were aspirated from the walls

of the building between 2300-0300 hrs with the majority taken on the

leeward side. The flies were strong fliers and aggressive biters.

The following day I searched 12 latrine/shower buildings in the

park, 7 in open, grassy areas and 5 near the river under large trees.

Those buildings near the river yielded no sandflies. The other 7

buildings yielded about 100 females and 5 males resting on the walls.

No flies were observed flying or feeding during the day. The night of
8 July was humid, 23C, partly cloudy with 1/4 moon, and winds gusting

to 15 knots. Collections made between 2100 hrs (onset of darkness)

and 0500 hrs produced about 60 females. About 50 flies were captured





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Figure 3-1.


Habitat of Lutzomyia diabolica.


Figure 3-2. L. diabolica feeding on a human arm.





-50-


9 July in the same buildings as on the 8th but in lesser numbers.

L. diabolica appears to be strongly attracted to light especially on

warm, humid, overcast nights.

Approximately 10% of the 250 females collected were engorged with

blood. All of the unfed flies were allowed to feed on my forearms

through the screen lid of the container on 9, 10 July. More than 50%

avidly took a blood meal.

For air shipment the plaster in the collecting containers was

dampened and vials were packed in a sealed plastic container with wet

towels. When received 8 hrs later 90% of the females had died after

being trapped in condensation on the sides of the containers. There-

fore, specimens shipped by air should be shipped in vials with the

plaster dry and wrapped in slightly damp towels.



Feeding


Although Lindquist (1936) reported L. diabolica feeding on humans

between 2000 hrs and 2400 hrs it will feed during all hours of darkness.

When a cage of flies is held against the skin of a host, females will

feed irrespective of light conditions.

When exposed to anesthetized hosts in a feeding cage or while

holding a feeding container against the host skin, L. diabolica fed

on the following animals: Homo sapiens (human; Figure 3-2), Canis

familiaris (dog), Neotoma micropus (woodrat), Mesocricetus auretis

(Syrian hamster), Sciurus carolinensis (grey squirrel), Oryctolagus

cuniculus (domestic rabbit), Didelphis marsupialis (opossun), Bos

taurus (calf), and Equus caballus (horse). Only one unsuccessful





-51-


attempt was made to feed flies on Ovis aries (sheep). L. diabolica

feeds not only on the ears of mammalian hosts as does L. anthophora

but also on the nose, around the eyes, or any other hairless or nearly

hairless areas.

Females feed within 24 hrs of eclosion. Feeding behavior is con-

sistent with observations of Lindquist (1936).

Two of 10 females in a feeding cage took a bloodmeal from the

inguinal region of a dog that had been infected with Leishmania donovani

infantum at least 10 months earlier. Five days after feeding each fe-

male had 150-200 promastigotes in the midgut.



Mating


Mating was observed under a wide range of light conditions and

before, after, and during feeding.



Egg Hatch and Fertility


No autogeny was observed in this species. Data on fecundity, pre-

oviposition period, and mortality factors are presented in Table 3-1.

First instar larvae do not exhibit synchronous egg hatching in

contrast to L. anthophora in which all the eggs of a single batch will

hatch within a 2 day period regardless of temperature. As many as 70%

of eggs laid by a single L. diabolica female often fail to hatch within

a 30 day period whereas nearly all the eggs laid by a single L. anthophora

female will hatch. The mechanism of this "partial fertility" phenomenon

of some L. diabolica eggs remains a mystery. Lindquist (1936) noted

an apparent diapause in the egg stage of L. diabolica from October to





-52-


March. The failure of eggs to hatch when laid by a single female was

observed from July to December at 200C, 24C, 28C, and 30C.

Development of individual immatures at various temperatures was

not studied because of insufficient numbers resulting from "partial

fertility."

I have observed that the egg-adult development time of L. diabolica

is 3-6 days less than the 36 days required for L. anthophora at 28C.












Table 3-1.


Lutzomyia diabolica--Fecundity, preoviposition period (days), and mortality factors for 3
generations in laboratory culture at 280C, 90% RH.


% Bloodfed Fecundity Preoviposition % Females % Females
Genera- Females Laying Period (Days) Peritrophic Fed on
tion n No Eggs x S Maximum # x S Range Sac Rupture Serum

1 27 59.3 39.6 11.2 58 5.7 1.7 3-8 11.1 0.0

2 21 28.6 32.0 14.0 51 5.5 1.6 3-9 4.8 4.8

3 51 58.8 36.2 16.0 64 6.2 2.3 3-12














SECTION IV

TRANSOVARIAN TRANSMISSION OF RIO GRANDE VIRUS BY
Lutzomyia anthophora(ADDIS)(DIPTERA: PSYCHODIDAE)



Introduction and Literature Review


The genus Phlebovirus of the family Bunyaviridae (Bishop et al.

1980) includes more than 40 viruses (R.B. Tesh, personal communication)

distributed over 5 continents (Berge 1975, Karabatsos 1978). Calisher

et al. (1977) described Rio Grande virus from isolates made from wood-

rats, Neotoma micropus, collected near Brownsville, Texas, in 1973-

1974. L. anthophora was suspected of transmitting this virus because

of its intimate association with woodrats (Young 1972), the high anti-

body prevalence (46.3%) of the woodrats (Calisher et al. 1977), and the

fact that sandflies transmit other related phleboviruses. Transovarian

transmission of phleboviruses by sandflies has been suggested as a

mechanism of viral survival (TeshandChaniotis 1975). In the present

study experiments were undertaken to demonstrate transmission of Rio

Grande virus by L. anthophora transovarially and by bite.



Materials and Methods


Sandflies


The L. anthophora used in these experiments were from the F7

generation of a closed colony started from stock collected E and NE


-54-





-55-


of Brownsville, Texas, in May and June 1980 (Section II). Flies used

in the experiments were held at 25C, 80% RH, and a 14:10, light:dark

photoperiod regime.


Virus

Rio Grande virus (Strain TBM4-719) was kindly supplied by

Dr. Robert Tesh, Yale Arbovirus Research Unit (YARU).


Infection of Sandflies


One hundred twenty, 1-4 day old female sandflies, anesthetized

with CO2 and held on ice, were injected intrathoracically with 105

PFU*/ml virus in phosphate-buffered saline by the method of Rosen and

Guebler (1974). Maintenance medium (88% Leibowitz medium, 10% tryptose-

phosphate broth, 2% heat inactivated fetal calf serum, 1% penicillin

100 units/ml-streptomycin 100 pg/ml) was changed every 4 days. Tubes

were examined at regular intervals. After 14 days incubation at 37C

those cultures showing viral cytopathic effect (Tesh et al. 1974) were

recorded as positive and discarded. Virus titers were calculated by

the method of Reed and Muench (1938). Blood samples were diluted and

inoculated into Vero cell tube cultures as above.

Fluid medium from half the positive tubes was tested by complement

fixation (Hawkes 1979) to confirm the presence of Rio Grande Virus (RGV)

antigen.

Eggs laid by infected flies were reared to adults then tested in

the manner described except that a single dilution (1:5) of sandfly

suspension was cultured.


*PFU: plaque forming unit.





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Suckling mice (Suisse variety) from two litters were inoculated

with 0.2 ml 105 PFU/ml RGV. One litter was inoculated subcutaneously

and the other was inoculated intracerebrally. One mouse from each

treatment was bled daily from the carotid artery. All blood samples,

0.1 ml, were diluted with 1.0 ml PBS with 0.05% gelatin and frozen at

-70'C for titration.



Virus Assay


Individual flies were triturated in 1.0 ml of dilutent in a sterile

2 ml Ten Broeck tissue grinder. The diluent was phosphate-buffered

saline, pH 7.2, containing 0.5% gelatin and 30% heat inactivated bovine

serum. Sandfly suspensions were centrifuged at 10,000 rpm for 30 min.

The supernatant was prepared in serial ten-fold dilutions from 10-1 to

106. Four tube cultures of Vero cells were then inoculated with 0.1 ml

of each dilution and incubated. Daily samples of 5 infected flies

were frozen at -70'C for virus titration to determine the growth of

Rio Grande virus (RGV) in the sandflies. Surviving females were offered

an anesthetized hamster daily for a 60 min period of feeding. After

feeding, engorged females were held at 25C, 80% RH, in individual

oviposition containers.

Fourth instar larvae and pupae were inoculated with RGV intra-

abdominally by the same method as the adults.

Suckling mouse blood was tested only at 1:10 dilution.

Infection of Rodents


To determine the fate of RGV in the natural hosts two,white-footed

mice, Peromyscus leucopus, and one female Neotoma micropus derived from





-57-


stock collected near Brownsville, Texas, were inoculated subcutaneously

on the lower ventral abdomen with 104 PFU/ml RGV. Daily blood samples

were collected from each animal from the retro-orbital capillaries for

7 days.



Results


The growth of RGV in adult female sandflies is shown in Table 4-1.

No lag phase occurred during viral replication in the sandfly. The RGV

titer in the sandfly increased from Day 1 to Day 7 post inoculation,

then declined slightly to an equilibrium that persisted for the life

of the fly. The persistent virus titer in the sandfly, 4.2-4.8 TCID505

is similar to those obtained by Jennings and Boorman (1980) with Pacui

virus in L. longipalpis and slightly higher than those found by Tesh

(1975) when he found 8 phleboviruses replicated in Ae. albopictus and

C. fatigans.

Ten 4th instar larvae inoculated with RGV pupated 2 days after

inoculation. Within 2 days after pupation all were dead based on the

criteria of a) obvious deformity or b) no movement in response to being

touched. Ten pupae similarly infected with RGV died within 2 days

after inoculation.

The development of RGV in woodrats and white-footed mice is shown

in Table 4-2. Although circulating titers were not determined for the

infected animals it is clear that the viremia was transient, lasting

only 1-2 days and was probably of a low titer.

Attempts to demonstrate transmission of RGV by the bite of in-

fected L. anthophora on uninfected suckling mice were unsuccessful


*Tissue culture infective dose.





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because the sandflies refused to feed on the mice (although they sub-

sequently fed on a hamster).

The following mortality was observed after 120 female sandflies

were inoculated with RGV: Day 1(18/120, Day 2 (26/102), and Day 3

(7/94). This mortality is attributable to the inoculation procedure.

Twenty-two (22) infected sandflies fed on a hamster and 6 of the

22 (27.3%) survived to oviposit. Based on observations on other genera-

tions of L. anthophora the expected survival would have been 61.5%. The

inoculation procedure may have caused this reduction. Mean egg pro-

duction for the 6 flies that oviposited was 28.3 (range 12-47). The

mean number of eggs produced does not seem affected by inoculation

procedures.

From 170 eggs laid by infected females, 62 adult F1 progeny were

produced with a sex ratio of 1:1. The transovarian transmission (TOT)

rate was 54.8% (34 of 62 adults were infected with RGV). The infec-

tion rates for males and females was similar, 15/30 (50.0%) and 19/32

(59.3%), respectively. Filial infection rates for the F1 progeny were

not calculated because of insufficient numbers. Each of the 6 in-

fected parents produced 1 or more infected progeny. One adult female

survived the F2 generation and laid eggs but was not infected.

The suckling mice inoculated intracerebrally (IC) or subcutaneously

(SQ) with RGV died at 4 and 3 days, respectively. Daily blood samples

from a mouse in each group were all positive for Rio Grande virus

titrated at 1:10 dilution. The rapid kill rate for the group infected

SQ was suspicious because of possible mouse colony contamination with

mouse hepatitis virus and the surprising fact that they died before

those mice inoculated IC. The experiment was repeated with the same

results.










Table 4-1. Growth of Rio Grande virus in L. anthophora after intrathoracic inoculation.

Day Post-Inoculation Number/Number Range of Titers Mean Titer in
Infected/Sampled in Infected Flies* Infected Flies*
0 (immediately after
inoculation) 5/5 <100"4-101 100.6

1 5/5 100.7-1017 101.3
2 5/5 101.7-1034 02.5

3 5/5 101.7 103.7 102.6

4 4/5 1029_ 103.1 103.1

5 4/5 103.4_105.0 104.1

6

7 5/5 104.3-105.7 105.0
8 2/2 104.0-1043 104.2

9 2/2 104. 5 10 4.5

10 1/2 104.8 104.8

*Tissue culture infectious dose50 per insect.





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Table 4-2.


Presence of Rio Grande virus in (1) Neotoma micropus and
(2) Peromyscus leucopus bled daily for 7 days after sub-
cutaneous inoculation.


Days Post Inoculation

Animal 1 2 3 4 5 6 7

Neotoma micropus 0 + + 0 0 0 0

Peromyscus leucopus
(a) 0 0 + 0 0 0 0

(b) 0 + + 0 0 0 0





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Discussion

The existence of a transovarian transmission cycle for the

Phlebotomus group viruses has long been suspected (Doerr et al. 1909,

Whittingham 1924). In the absence of virus isolations, Russian workers

(Moshkovski 1937, Petricheva and Alymov 1938) demonstrated transmission

of an etiologic agent for Papataci fever through the eggs of P. papataci

and demonstrated overwintering of the agent in F1 larvae of the same

species. The recovery of phleboviruses from wild caught male sandflies

by several workers (Schmidt et al. 1971, Aitken et al. 1975, Tesh et al.

1974, 1977) lends further evidence to the existence of transovarian

transmission in this group. Transovarian transmission of another virus

serogroup transmitted by sandflies, Vesicular Stomatis Virus, has been

demonstrated by Tesh et al. (1972).

Despite overwhelming evidence indicating that Papataci fever virus

and other phleboviruses are transmitted by phlebotomine sandflies

(Schmidt 1971, Tesh 1975), no quantitative laboratory studies have been

conducted to demonstrate transmission by bite or by transovarial means.

The lack of controlled experiments has been partially a function of the

difficulty encountered in rearing sandflies.

In this series of experiments we were unable to demonstrate

transmission by bite due to the lack of susceptible laboratory animals,

by the fact that L. anthophora refused to feed on suckling mice, and

a lack of knowledge on virus dynamics in a rodent host. Therefore,

infection was established by means of intrathoracic injection. Sand-

flies infected by injection transmitted Rio Grande virus to 54.8% of

their progeny. This is the first demonstration of transovarian trans-

mission of a Phlebovirus by sandflies.





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Calisher et al. (1977) found neutralizing antibody to Rio Grande

virus in woodrats, opossums, gopher tortoises, horses, several species

of small rodents, birds, and a horned toad. I was unable to induce

L. anthophora to feed on gopher tortoises, or horses. This suggests

that L. anthophora probably is not the only arthropod vector of Rio

Grande virus. Other hematophagous arthropods that I recovered from

the woodrat nests were Ornithodorus dugesi (often identified as

0. tulaje), Triatoma gerstaeckeri, Triatoma sanguisuga, and Triatoma

neotomae. Johnson (1966) also reported several species of fleas and

Ixodidae from the woodrat nests. It is highly likely that mosquitoes

also use the nests for resting sites. Some of these other arthropods

are catholic in their feeding behavior and could possibly transmit

Rio Grande virus to animals not fed on by L. anthophora, and may be

capable of transovarian transmission.

The ecology of Rio Grande virus remains to be thoroughly studied.

Although Neotoma micropus and Peromyscus leucopus are susceptible to

infection by subcutaneous inoculation the infection is transient.

McLean et al. (1982) confirmed the results in Table 4-2 in that the

viremia in woodrats is short-lived, 2.5 days, and of low titer, mean

3.65 loglo PFU/ml in Vero cell culture. As yet, oral infection of

L. anthophora has not been demonstrated nor is the minimum infective

oral dose known. McLean et al. (1982) also determined that nearly all

the woodrats developed neutralizing antibody thus becoming refractory

to infection for life. This indicates that only young woodrats are

likely to be susceptible to infection after maternal antibody is no

longer present and they could only serve as amplifying hosts for 2.5

days during their lifetime. The high transovarian transmission rate





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obtained with L. anthophora could account for maintenance of RGV in

nature by the stabilization mechanism discussed by Tesh and Shroyer

(1980).

At Uvalde, Texas, Parman (1919) reported an epidemic of a mild

febrile illness (102-1040F, 3 days duration) concurrent with large popu-

lations of L. diabolica, a man-biting species also known to feed on

woodrats, opossums, cattle, dogs, and horses. Parman (1919) thought

the possible association of the sandfly numbers and the epidemic was

a suspicious coincidence requiring investigation. The symptoms de-

scribed by Parman (1919) are consistent with those known to occur after

infection by phleboviruses (Bartonnelli and Tesh 1976, Tesh et al.

1977).

From this study I must conclude that L. anthophora is probably

important in maintaining Rio Grande virus in the woodrat population but

may not solely account for its transmission. In order to more com-

pletely understand the ecology of Rio Grande virus detailed field and

laboratory investigations of the vector potential of other arthropods

must be undertaken.














SECTION V

RIO GRANDE VIRUS AND Triatoma gerstaeckeri
(STAL) (HEMIPTERA: REDUVIIDAE)



Introduction and Literature Review


The hematophagous Hemipterans, the Cimicidae and the Triatominae,

have been considered ideal potential vectors of arboviruses because of

the large blood meal ingested, their relative longevity, and their

cosmopolitan feeding habits. Kitselman and Grundman (1940) reported

isolating Western Equine Encephalitis (WEE) virus from Triatoma sangui-

suga taken in a Kansas pasture where animals had died of the disease in

previous years. Mangiafico et al. (1968) found that 2 species of

Triatome, R. prolixus and T. infestans, would harbor WEE virus 14-20

days when unpunctured. When punctured to simulate cannabalistic feed-

ing virus survived 98 days and one bug transmitted the virus by bite.

Justines and Sousa (1977) obtained similar results with punctured bugs

and bugs infected with Trypanosoma cruzi. Hayes et al. (1977) found the

cliff swallow bug, Cimicidae, to be capable of overwintering and trans-

mitting Ft. Morgan virus (Calisher et al. 1980).

In view of the findings noted and realizing that Triatoma ger-

staeckeri was known to feed on all of the animals (Lent and Wygodzinsky

1979), in which Calisher et al. (1977) had found neutralizing antibody

to Rio Grande virus, the vector potential of T. gerstaeckeri was in-

vestigated. Thurman (1945) and Pippin (1970) reported finding


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T. gerstaeckeri infected with T. cruzi in Neotoma nests. Pippin (1970)

noted 30.1% of the bugs found in the nests were infected.



Materials and Methods

Virus

Rio Grande virus (strain TMB4-719) was kindly supplied by Dr.

Robert Tesh, Yale Arbovirus Research Unit. Additional quantities of

virus were prepared by passage through suckling mouse brain.



Triatomes


A colony of T. gerstaeckeri was started from specimens collected

near Brownsville, Texas, in June, 1980, and augmented with specimens

collected near Lake Medina, San Antonio, Texas, in July, 1981. First

and second instar nymphs from the colony were used in the transmission

experiments.

One hundred forty (140) 2-3 week old first instar nymphs were fed

on 3 suckling mice that had been given Rio Grande virus by intracerebral

inoculation 4 days earlier. Three other mice from the same litter died

on Day 5 post inoculation.

Eight, 16, and 24 days after the initial feeding 80 first instar

nymphs that had fed on viremic mice fed on 6 unexposed suckling mice.

After 1 week the suckling mice showed no apparent signs of viral infection.



Results and Conclusions

The failure to infect 6 suckling mice fed on by 80 nymphs that had

fed on viremic mice 8, 16, or 24 days earlier indicates that T. gerstaeckeri





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may not be capable of transmitting Rio Grande virus. However, the

additional experiments should be conducted before the Triatominae are

proven to be incompetent vectors of phleboviruses. These should in-

clude fluorescent antibody localization of viral antigen to determine

its fate in the insect.














SECTION VI

PURIFICATION OF RIO GRANDE VIRUS



Introduction


Since Calisher et al. (1977) first characterized Rio Grande virus

little other descriptive work has been performed. Because of its

ecological relationship to other members of the group, Rio Grande virus

was placed in the genus Phlebovirus of the family Bunyaviridae by

Bishop et al. (1980). In this section a purification scheme for the

virus is described and electron micrographs of the virus are presented.



Materials and Methods


Virus


Rio Grande virus (Strain TMB4-719) was kindly supplied by Dr.

Robert Tesh, Yale Arbovirus Research Unit. Additional quantities of

virus were produced by intracerebral inoculation of 2-4 day old suck-

ling mice with 0.01-0.02 ml stock virus. Four days after inoculation

the mouse brains were harvested and triturated in 2 ml Ten Broeck

tissue.grinders with 1x sterile phosphate buffered saline (PBS), pH 7.4.



Infection of Cells and Virus Purification


The purification scheme detailed below was based on several others

previously used for arboviruses (Kaariainen et al. 1969, Obijeski et al.


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-68-


1976, Clewley et al. 1977). Four 150 cm2 tissue culture flasks of Vero

cells in a confluent monolayer were inoculated with 6 ml mouse brain

suspension and allowed to adsorb for 1 hr at 370C. Cultures were then

overlaid with 50 ml minimum essential medium (MEM) and incubated at

37C. After 24 hrs the supernatant was poured off, 75 ml MEM was over-

laid, and the cells were incubated for an additional 6 days at 37C

after which >90% of the cells were destroyed. Supernatants were col-

lected, frozen to -700C, thawed to 37C, then clarified by low-speed

centrifugation at 40C for 30 min at 8,000 g in a Sorvall RCB-2 centri-

fuge to remove cell debris.

Virus was recovered from the clarified supernatant by precipita-

tion in a 7% polyethylene glycol/0.4 M NaCl solution stirred for 4 hr

at 4C followed by centrifugation at 10,000 g for 20 min. The pellet

was resuspended in 4 ml TSE buffer (0.01 M Tris hydrochloride buffer,

pH 7.5, containing 0.1 M NaCl and 0.002 M EDTA) and loaded over a com-

bination equilibrium: viscosity gradient of potassium tartrate (McCrea

et al. 1961) and glycerol (KT-GLY).

Two 10 ml KT-GLY gradients were made with a Bethesda Research

Products Gradient Former and an LKB peristaltic pump. Fourteen

milliliters of 50% (w/w) potassium tartrate in TSE buffer was loaded

into the inside chamber and 16 ml of 30% (w/w) glycerol was loaded into

the outside chamber (Obijeski et al. 1974, Barzilai et al. 1972).

Virus suspensions loaded onto KT-GLY gradients were centrifuged
in an SW 41 rotor at 4C for 8 hr at 40,000 g. Three nearly inseparable

visible bands were produced. The virus fraction was collected at 254 nm

(RNA absorbance peak) using an ISCO gradient column fractionator and

flow densitometer.





-69-


Electron Microscopy


A drop of virus suspension was placed on a Formvar carbon coated

grid and allowed to dry for I min before the excess fluid was wicked

off with filter paper. The grid was then negatively stained for 45 sec

with 2%, pH 6.8, phosphotungstic acid with KOH, using 50 vig/ml

Bacitracin as a spreading agent (Gregory and Pirie 1973). Specimens

were examined at 75 KV, 50,O00x and 100,O00x in a Hitachi 600 Trans-

mission Electron Microscope.



Results


Electron micrographs (Figure 6-1) show that the virion is spherical

and possesses an envelope bearing small spikes.

The virion is 71 nm in diameter as determined using a reference

catalase crystal. This is within the size range of 60-90 nm that is

characteristic of the Bunyaviridae (Bishop et al. 1980).



Discussion


The size and morphology of the Rio Grande virion is consistent

with those described for the genus Phlebovirus (Bishop et al. 1980).

An additional purification step of centrifuging the virus suspen-

sion in a 20-70% (w/v) sucrose gradient at 40C and 35,000 g for 4 hr was

not used since it was possible to recover the virus after the equilibrium-

density centrifugation in potassium tartrate.

In order to verify that the virions shown in the electron micro-

graphs retained infectivity, 1 ml of virus suspension was adsorbed onto





-70-


Figure 6-1.


Electron micrographs of purified Rio Grande virus at
125,000x.





-71-


a confluent monolayer of Vero cells in a 75 cm3 tissue culture flask

which was then overlaid with 50 ml minimum essential medium. After

6 days incubation at 37C nearly 100% of the cells were destroyed.














SECTION VII
A COMPARISON OF OOCYTE TOPOGRAPHY OF FIVE PHLEBOTOMINE SANDFLIES
(Lutzomyia) WITH THE SCANNING ELECTRON MICROSCOPE
(DIPTERA: PSYCHODIDAE)



Introduction

The egg surface structure of 19 neotropical Phlebotomine species

has been described (Zimmerman et al. 1977, Ward and Ready 1975) using

the scanning electron microscope (SEM). Ward and Ready (1975) noted

three species-specific topographic patterns, i.e., polygonal, parallel

ridging, and volcano-like. Several authors (Chaniotis and Anderson

1964, Addis 1945,Lindquist 1936, Barreto 1941, and Sherlock 1957a,b, 1963)

described and figured the eggs of 17 neotropical sandfly species using

light microscopy. After examining the literature cited and eggs from

the 5 species described herein we propose adding another category to

the patterns of Ward and Ready (1977); that is, parallel ridges connected

or parallel ridges unconnected.

The classification of eggs of 41 species of New World sandflies

according to the proposed scheme is presented in Table 7-1.



Materials and Methods

Eggs were obtained from females reared in laboratory colonies.

The preparation method of eggs for SEM based on the work of Quattlebaum

and Carner (1980) is as follows:


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Table 7-1. Classification of 41 species of Neotropical phlebotomine sandfly eggs based on oocyte
topographic patterns.

Topographic Pattern

Parallel Ridges Parallel Ridges
Describer Polygon (connected) (unconnected) Volcano-Like

Endris L. texana L. cruciata spp. L. diabolica
et al. L. vexator L. anthophora
L. shannoni
Sherlock L. lenti L. renei
L. bahiensis

Zimerman L. sanguinaria
et al. L. trapidoi
L. ylephilator
L. gomezi
Chaniotis L. vexator
occidentalis

Ward and L. antanesi L. longipalpis L. flaviscutellata
Ready L. yuilli L. complexa
L. nsp. 260.43 L. lainsoni
L. nsp. 260.44 L. carrerai
L. dendrophyla L. davisi
L. gomesi L. paraensis

Barreto L. guimaraisi L. pestanai L. lanei
L. pessoai L. arthuri L. whitmani
L. fischeri L. intermedia L. alphabetica
L. limai
L. monticola





-74-


1. Eggs were placed on a filter paper disc in a 1 cm deep plastic

container cut from a plastic film cannister.

2. The plastic container was floated in a 50 ml Tri-pour poly-

styrene beaker containing 5 ml aqueous 1% OsO4

3. The paper lid was installed and the entire container was sealed

in Parafilm and held in an exhaust hood at room temperature

for 5 days.

4. After 5 days exposure to osmium vapor the inner container was

transferred to a covered petri dish for 24 hr to allow slow

drying of the eggs.

5. Eggs were attached to an SEM stub using either double-sided

tape or 0.1% aqueous hydrobromide polylysine (Polysciences,

Inc., Warrington, PA 18976), sputter-coated with approximately

300 A of gold in an Eiko Engineering IB-2 Ion Coater, and

examined in a Hitachi S-450 scanning electron microscope (SEM)

at 20 KV.

Eggs were measured in microns at lOOx with a compound microscope

and an ocular micrometer. Intact fresh eggs or recently hatched eggs

were acceptable whereas old eggs or infertile eggs usually collapsed

making accurate measurement difficult. Eggs to be measured were placed

on a microscope slide in Histocon. In each sample the eggs were pro-

duced by 5-10 females.



Results


The SEM micrographs of the sandfly eggs are shown in Figures 7-1

and 7-2. Descriptions of eggs of each species are as follows. Measurements





-75-


Figure 7-1.


Scanning electron micrographs of eggs of four sandfly
species. (1) Lutzomyia diabolica, (2) Lutzomyia shannoni,
(3) Lutzomyia vexator, (4) Lutzomyia cruciata spp.





-76-


Figure 7-2. Scanning electron micrographs of oocyte topography of
five species of sandfly. (5) Lutzomyia diabolica, (6)
Lutzomyia shannoni, (7) Lutzomyia vexator, (8) Lutzomyia
cruciata spp., (9) Lutzomyia anthophora. 7,000x.





-77-


given are the range, mean, and standard deviation for egg length and

width for each species.


Lutzomyia shannoni (Dyar, 1929), Florida specimens

Figure 7-1(2), 7-2(6)

Size: N = 102, L: 290-340 (330 10), W: 70-110 (90 10)

Exochorion: High, narrow longitudinal ridges connected by prominent

perpendicular ridges forming 4 and 5 sided polygons which are fre-

quently rectangular.


Lutzomyia diabolica (Young and Perkins 1982), Uvalde Co., Texas

Figure 7-1(1), 7-2(5)

Size: N = 47, L: 340-370 (350 10), W: 90-110 (100 10)

Exochorion: Surface topography is characterized by a series of dis-

continuous parallel longitudinal ridges that are not laterally connected.


Lutzomyia vexator (Coquillett 1907), Levy Co., Florida

Figure 7-1(3), 7-2(7)

Size: N = 193, L: 330-390 (380 10), W: 80-110 (100 10)

Exochorion: Surface topography consists of delicate parallel longi-

tudinal ridges with regular perpendicular connections that form polygons

which are nearly square. There are also occasional oblong cells.


Lutzomyia anthophora (Addis 1945), Cameron Co., Texas

Figure 7-2(9)

Size: N = 100, L: 330-370 (340 10), W: 80-100 (80 10)

Exochorion: Reticulation consists of weak parallel longitudinal ridges

with slight perpendicular connections at irregular intervals.





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Lutzomyia cruciata spp. (Young and Perkins 1982), Alachua Co., Florida

Figure 7-1(4), 7-2(8)

Size: N = 61, L: 320-370 (340 10), W: 80-120 (100 10)

Exochorion: Wide, flat, parallel longitudinal ridges with occasional

weaker connecting ridges which are not usually perpendicular to the

longitudinal ridges.



Discussion

Several techniques were tried for preserving the eggs to prevent

collapse under vacuum in the SEM column. The method used here yielded

the best results when fertile eggs were used.

A "standard" EM fixation procedure using 1% OsO4 as a fixative

followed by 5% aqueous acrolein, dehydration in dimethoxypropane and

acetone, then critical point drying with Freon as a transition solvent

proved unsuccessful because most specimens collapsed in the SEM.

Lyypholization and critical point drying of eggs without fixation

were also unsuccessful.

A technique which was not used but one which may be promising is

freeze drying.

The size variation of eggs laid by individual females was deter-

mined by measuring 10 eggs from each of 10 L. vexator females. The

variation in egg length and width between females ranged from 10-50

microns and from 10-30 microns, respectively. As a result of broad

intraspecific variation it is not possible to separate the eggs of

different sandfly species by size. Therefore the surface sculpturing

is the only characteristic of the egg that can be used for species

determination.














SECTION VIII

PHOTOGRAPHIC TECHNIQUES


Quality photographs of living sandflies, Culicoides, and other

small insects have been notably lacking from the literature due to

the difficulty of producing them. Part of the difficulty involved in

photographing these insects is containing them. Two types of spe-

cialized containers were developed in order to photograph sandflies.

The first container (Figure 8-1) was constructed from a 40 liter

aquarium. Two sides were replaced with 2 mm Plexiglas. Three access

ports (19 x 19 cm) were cut in the sides and fitted with Plexiglas

compression frames (2 cm wide x 64 mm thick). Sleeves 50 cm in length

made of 15 cm surgical stockinet (Johnson & Johnson) were secured

around the ports by the compression frames which were attached with

8 brass screws (10/24 x 1") and wing nuts. The screws and flat washers

wereglued in place with epoxy glue for ease of attaching the sleeves.

Three sleeves are required, 1 for the camera, 1 for manipulating

specimens, and 1 for the host arm. A Plexiglas insert was attached to

the top frame of the aquarium with silicone glue thus making a re-

movable top. The back and bottom of the aquarium is covered with a

1 cm layer of plaster of Paris to provide a light reflective back-

ground.

The second type of photographic chamber (Figure 8-2) was con-

structed from a spectrophotometric cuvette (Wallace & Tiernan, Co.,


-79-




-80-


Figure 8-1. Chamber for photographing hematophagous insects feeding
on humans and small mammals.


Chamber for photographing small insects.


Figure 8-2.





-81-


Belleville, NJ) that is 7.5 x 2.5 x 1.5 cm. It was covered on the

bottom and ends with a 1 cm layer of white polyethylene foam (Ward's

Scientific Co., Rochester, NY). A Kodak Neutral Test card 90% re-

flectance on the white side and 18% on the gray side was used for a

background behind the cuvette chamber.

A third type of photographic chamber used occasionally was the

rectangular adult feeding cage.

All of the sandfly photographs presented in this manuscript were

photographed with a 200 mm Medical Nikkor lens at 3x magnification, F45.

This lens was used because it has a built-in ring flash and provides

8 cm working distance at 3x magnification. The camera used was a Nikon

F2 Photomic with type "C" focusing screen and cable release. Because

the lens is of a fixed focal length it was necessary to mount the

camera on a Slik 2-axis focusing rail on a tripod in order to achieve

reproducible results.

Fujichrome film, ASA 100, was used for all the photographs. Black

and white prints were all made from color slides using Ilford XP400

ultra fine grain film for an internegative.














SECTION IX

SUMMARY


During the four year course of this study the following objectives

were achieved:

1. Methods for laboratory culture of phlebotomine sandflies were

developed. Use of these culture techniques will permit detailed quanti-

tative study of the vector competence of sandflies for viral and other

parasitic diseases. To date twelve species have been reared in closed

colony for as many as 25 generations using these techniques.

2. Detailed studies on the laboratory biology of two species of

Neotropical phlebotomine sandfly, Lutzomyia anthophora and Lutzomyia

diabolica were conducted over 16 and 7 generations, respectively.

3. Transovarian transmission of a Phlebovirus by a sandfly was

conclusively demonstrated for the first time. Nearly 55% of L.

anthophora females injected with Rio Grande virus transmitted it to their

progeny.

4. The oocyte surface sculpturing of 5 species of Neotropical

sandflies were described for the first time with the scanning electron

microscope. The topographic patterns can be used to identify these

sandfly species.

5. A method for purification of Rio Grande virus was developed.

The virus was purified and photographed in a transmission electron

microscope. This is one of the first recorded purifications and char-

acterizations of a Phlebovirus.


-82-














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Full Text
STUDIES OF Lutzomyia anthophora (ADDIS) (DIPTERA:
PSYCHODIDAE) AND OTHER POTENTIAL VECTORS
OF RIO GRANDE VIRUS
By
RICHARD G. ENDRIS
A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF
THE UNIVERSITY OF FLORIDA
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE
DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA


Abstract of Dissertation Presented to the Graduate Council
of the University of Florida in Partial Fulfillment of the Requirements
for the Degree of Doctor of Philosophy
STUDIES OF Lutzomyia anthophora (ADDIS) (DIPTERA:
PSYCHODIDAE) AND OTHER POTENTIAL VECTORS
OF RIO GRANDE VIRUS
By
Richard G. Endris
May, 1982
Chairman: Dr. Harvey Cromroy
Major Department: Entomology and Nematology
Simple colonization techniques for rearing large numbers of
phlebotomine sandflies were developed. Lutzomyia anthophora (Addis)
and Lutzomyia diablica (Hall) were colonized in the laboratory for the
first time for 16+ and 7+ generations, respectively, thus permitting
quantitative investigation of their ability to transmit viruses and
leishmaniasis. Notes on field behavior of j_. anthophora and L_. diabolica
are presented with detailed laboratory studies on the biology of the
two species. Laboratory transovarian transmission of a Phlebovirus
was demonstrated for the first time with j_. anthophora when 54.8% of
the F^ adult progeny from parents infected by intrathoracic inoculation
became infected. Attempts to transmit Rio Grande virus by the bite of
J_. anthophora and Triatoma gerstaeckeri (Stal) were unsuccessful.
xi i i


-80-
Figure 8-1. Chamber for photographing hematophagous insects feeding
on humans and small mammals.
Figure 8-2. Chamber for photographing small insects.


Table 2-7. Fecundity, percent of bloodfed females that laid no eggs, and preoviposition period (days)
for 12 generations of L.. anthophora reared at 24C and 28C, 90% RH.
Generation
n
Fecundity
% Bloodfed
Females Laying
No Eggs
Preoviposition Period (Days)
X
S
Maximum #
X
S
Range
Fi
25
42.0
20.5
71
28.0
5.2
1.4
3-11
F2
99
34.7
21.6
79
53.5
5.5
1.3
4-7
F3
171
31.7
16.9
64
66.2
7.1
2.0
4-16
F4
80
39.5
19.0
73
53.3
5.3
1.1
4-8
F5
61
43.2
13.1
65
43.3
6.4
1.8
4-10
F6
94
40.6
23.2
107
50.0
6.2
1.3
4-10
F7
124
36.1
14.3
68
65.3
5.6
0.8
4-6
F8
121
32.2
18.3
75
45.6
4.5
1.7
2-9
F9
61
32.9
16.5
63
27.9
4.8
1.0
4-8
F10
92
32.5
17.8
73
32.6
5.8
1.5
2-9
F11
190
26.1
13.1
56
21.8
6.2
1.5
3-11
in
H
Ll_
125
28.0
13.2
56
19.8
6.2
1.8
3-11
Overa!1
Mpan<;
1203
35.0
17.3
70.8
38.5
5.7
1.3
2-11


Kristin Figura, Kay Warren, and Annie Moreland for their assistance
in virus purification and titration.
Dr. A.G.B. Fairchild for sharing his vast knowledge and years of
experience.
A grant from the Steffen Brown Foundation provided the opportunity
to study at Yale University in 1981.
VI


Table 7-1. Classification of 41 species of Neotropical phlebotomine sandfly eggs based on oocyte
topographic patterns.
Topographic Pattern
Describer
Polygon
Parallel Ridges
(connected)
Parallel Ridges
(unconnected)
Volcano-Like
Endris
et al.
L. texana
L. vexator
L. shannoni
L. cruciata spp.
L. anthophora
L. diablica
Sherlock
L. lenti
L. bahiensis
L. renei
Zimmerman
et al.
L. sanguinaria
L. trapidoi
L. ylephilator
L. qomezi
Chaniotis
L. vexator
occidental is
Ward and
Ready
L. antanesi
L. yuilli
L^. nsp. 260.43
j_. nsp. 260.44
L. dendroph.yla
L. qomesi
L. lonqipalpis
L. flaviscutellata
L. complexa
L. lainsoni
L. carrerai
L. davisi
L. paraensis
Barreto
L. quimaraisi
L. pessoai
L. fischeri
L. limai
L. monticola
L. pestanai
L. arthuri
L. intermedia
L. lanei
L. whitmani
L. alphabet!'ca


BIOGRAPHICAL SKETCH
On July 31, 1948, Richard G. Endris entered the world at the rural
farming community of Pana, Illinois. After twelve years residence there
his family moved to Somerville, NJ, where he completed primary school
and graduated from Somerville High School in 1966. After graduating
from Rutgers University, New Brunswick, NJ, with a degree in animal
science he enrolled at the University of Florida for a master's degree
in entomology which he completed in 1972. The subsequent three years
were spent in the U.S. Army as a medical entomologist with assignments
to the 82nd Airborne Division and 8th Army Korea. In 1976 he left the
Army to work as a health consultant to Fluor Corporation in Asia for a
year before enrolling in a Ph.D. program in entomology at the University
of Florida in 1977.
-90-


-66-
may not be capable of transmitting Rio Grande virus. However, the
additional experiments should be conducted before the Triatominae are
proven to be incompetent vectors of phleboviruses. These should in
clude fluorescent antibody localization of viral antigen to determine
its fate in the insect.


-55-
of Brownsville, Texas, in May and June 1980 (Section II). Flies used
in the experiments were held at 25C, 80% RH, and a 14:10, light:dark
photoperiod regime.
Virus
Rio Grande virus (Strain TBM4-719) was kindly supplied by
Dr. Robert Tesh, Yale Arbovirus Research Unit (YARU).
Infection of Sandflies
One hundred twenty, 1-4 day old female sandflies, anesthetized
5
with CO2 and held on ice, were injected intrathoracically with 10
PFU*/ml virus in phosphate-buffered saline by the method of Rosen and
Guebler (1974). Maintenance medium (88% Leibowitz medium, 10% tryptose-
phosphate broth, 2% heat inactivated fetal calf serum, 1% penicillin
100 units/ml-streptomycin 100 pg/ml) was changed every 4 days. Tubes
were examined at regular intervals. After 14 days incubation at 37C
those cultures showing viral cytopathic effect (Tesh et al. 1974) were
recorded as positive and discarded. Virus titers were calculated by
the method of Reed and Muench (1938). Blood samples were diluted and
inoculated into Vero cell tube cultures as above.
Fluid medium from half the positive tubes was tested by complement
fixation (Hawkes 1979) to confirm the presence of Rio Grande Virus (RGV)
antigen.
Eggs laid by infected flies were reared to adults then tested in
the manner described except that a single dilution (1:5) of sandfly
suspension was cultured.
*PFU: plaque forming unit.


Table 2-6. Temperature (C) of body regions of anesthetized and non-anesthetized hosts for L. anthophora
in laboratory culture.
Ear Tail
Animal
Tip
Middle
Base
Tip
Base
Rear Foot
Nose
Back
Neotoma micropus
(anesthetized)
31.5
31.1
34.2
32.4
32.4
33.1
29.7
37.0
Neotoma micropus
(non-anesthetized)
31.3
32.3
37.1
' 31.3
36.8
30.0
37.8
Mesocricetus auretus
(anesthetized)
27.7
30.3
34.1
33.2
35.0
35.4

27.8
36.9
1
CJ
cn
i
Mesocricetus auretus
(non-anesthetized)
32.0
32.9
37.2
34.2
35.6
36.2
27.7
36.9
Oryctolaqus cuniculus
(non-anesthetized)
35.0
36.8
31.4
31.5


Table 2-4. J_. anthophora--Adult sugar feeding, frequency, age of feeding (days), time required for
digestion (days) at 20C and 28C, 90% RH.
Tempera
ture
(c)
Age at
1 Feed
# Days to
Digest
1 Meal
# Days to
2 Feed
# Days to
Digest
2 Meal
# Days to
3 Feed
# Days to
Digest
3 Meal
# Days to
4 Feed
# Days to
Digest
4 Meal
# Days to
5 Feed
28
1.30.8
1.4+0.8
1.41.1
1.30.5
1.50.7
1.6+0.9
1.81.1
1.30.5
1.0
n = 68
n = 48
n = 29
n = 25
n = 17
n = 16
n = 10
n = 6
n = 1
20
1.91.0
2.21.2
1.20.5
1.80.8
1.20.4
1.0
1.0
n = 17
n = 10
n = 8
n = 6
n = 2
n = 2
n = 1
I
OJ


-27-
Female Age at First Feeding
To determine when J_. anthophora females were physiologically ready
to take a blood meal, all adults from the F^ generation that eclosed
each day were held in the cylindrical feeding chamber. Each day an
anesthetized mouse was placed in the chamber for 60 min until all the
females in each group either fed or died. Of 244 females, 92 (60.5%)
fed within 1-7 days. The mean age at feeding was 3.7 days and the median
age was 3.5 days. Addis (1945b) noted that females fed 2-4 days post
eclosin. The temporal distribution of female age at feeding and the
age of death for those flies that did not feed is presented in Figure
(2-6).
The sugar feeding habits of hematophagous diptera are well known
and most previous attempts to colonize sandflies included the provision
of 30% sucrose solutions for the adults. Chaniotis (1974) and Ready
(1979) investigated sugar feeding in phlebotomine sandflies. Chaniotis
(1974) studied sandfly preference for various sugars and determined
that sugar concentration had no effect on fly longevity. Ready (1979)
found higher egg production among bloodfed females fed on sucrose solu
tion vs. those fed on water. Killick-Kendrick (1979) suggested that
the presence or absence of sugar in the gut may have profound and
largely undetermined effects on the ability of sandflies to transmit
leishmaniasis.
An experiment was conducted to determine what percentage of adults
will ingest honey solutions, at what age, frequency of sugar feeding,
and the time required to digest each meal. Individual pupae were placed
in 7 dram feeding vials. The day of eclosin a small drop of 30% honey


-7-
thin ones which tend to dry. A tangential section of each apple slice
should be removed to produce a flat edge so that the slice will not
roll and crush flies. Fresh slices are added daily. Rhizopus sp. is
the mold that usually grows on "old" apple slices and it can be used as
inoculum for larval medium.
Bloodfed females are provided sugar in the oviposition cages by
placing a small drop of 30% honey solution or 50% Karo syrup solution
on the screen lid. A 30% honey solution was used initially in an
effort to produce a facsimile of nectar but a Karo syrup solution pro
vided equivalent results. If fungal or bacterial growth become ap
parent in the sugar solution the lid should be changed and a new drop
added. Refrigeration of stock sugar solutions at 3C greatly increases
their shelf life.
N-butyl Pthalate
Clear vinyl suction cups were used to suspend apple slices from
the top of the feeding cage. This practice was quickly abandoned after
adult mortality approaching 100% was associated with the vinyl use.
N-butyl pthalate, an elasticizer used in vinyl, is volatile at room
temperature and is highly toxic to sandflies and mosquitoes (David
Carlson, biochemist, personal communication). The compound has been
used as an insect repellant (The Merck Index 1976).
Adult Feeding
Two general methods for feeding adult females on an animals were
used. The first method is to hold a 7 dram vial or 120 ml specimen



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-57-
stock collected near Brownsville, Texas, were inoculated subcutaneously
4
on the lower ventral abdomen with 10 PFU/ml RGV. Daily blood samples
were collected from each animal from the retro-orbital capillaries for
7 days.
Results
The growth of RGV in adult female sandflies is shown in Table 4-1.
No lag phase occurred during viral replication in the sandfly. The RGV
titer in the sandfly increased from Day 1 to Day 7 post inoculation,
then declined slightly to an equilibrium that persisted for the life
of the fly. The persistent virus titer in the sandfly, 4.2-4.8 TCID^g*
is similar to those obtained by Jennings and Boorman (1980) with Pacui
virus in _L. lonqipalpis and slightly higher than those found by Tesh
(1975) when he found 8 phleboviruses replicated in Ae. albopictus and
C. fatigans.
Ten 4th instar larvae inoculated with RGV pupated 2 days after
inoculation. Within 2 days after pupation all were dead based on the
criteria of a) obvious deformity or b) no movement in response to being
touched. Ten pupae similarly infected with RGV died within 2 days
after inoculation.
The development of RGV in woodrats and white-footed mice is shown
in Table 4-2. Although circulating titers were not determined for the
infected animals it is clear that the viremia was transient, lasting
only 1-2 days and was probably of a low titer.
Attempts to demonstrate transmission of RGV by the bite of in
fected J_. anthophora on uninfected suckling mice were unsuccessful
*Tissue culture infective dose.


-33-
this hypothesis a piece of Opuntia was sliced and placed in a feeding
cage with j_. anthophora. Specimens were observed feeding on the plant
juice but not as avidly as on apple slices.
Feeding Hosts
l. anthophora has been reported to feed on rodents and lagomorphs
(Addis 1945a, Young 1972, and Calisher 1977). J_. anthophora fed readily
on the following anesthetized animals introduced into the feeding cage:
Neotoma micropus (woodrat; Figure 2-7), Peromyscus leucopus (white
footed mouse), Mesocricetus auretus (Syrian hamster), Sciurus carolinen-
sis (grey squirrel), Mus musculus (white mouse; Figure 2-8), Cavia por-
cellus (guinea pig), Oryctolagus cuniculus (domestic rabbit), and
Didelphis marsupial is (opossum). The preferred feeding site was the
nearly hairless portions of the ears. Fewer than 5% of the sandflies
fed on the feet or among the vibrissae on the nose.
It is notable that l. anthophora would not feed on suckling mice
when restrained with a cloth net on a tongue depressor. Attempts to
feed the sandflies on the poikilotherms, Gopherus polyphemus (gopher
tortoise) and Anolis carolinensis (anole), were unsuccessful.
Flies held in feeding cages on the ears of Cam's familiaris (dog),
and Ovis aries (sheep), did not feed. Efforts to feed the flies on Homo
sapiens (human) were not successful. 1. anthophora did feed on the
ear of a 2 day old calf (Bos taurus) and on a shaved chick (Gallus
gallus) restrained in a feeding cage.
When the sandflies did not feed on the preferred host they were
subsequently offered either an opossum, woodrat, hamster, or mouse.


-14-
120 ML SPECIMEN CONTAINER
FULL SCALE
7 DR VIAL
FULL SCALE
ASPIRATOR
ONE-THIRD SCALE
Figure 1-5. Field collection apparatus, feeding and rearing containers
for phlebotomine sandflies.


-48-
of a laboratory colony, I collected specimens in July 1981 at Garner State
Park, a site located approximately 50 km N of Uvalde, Uvalde Co., Texas,
in the Frio river valley near the eastern edge of the Edwards Plateau.
The habitat is character!-zed by open grassland interspersed with oak,
acacia, and cedar trees surrounded by rocky hills (Figure 3-1).
All specimens were taken with a tube aspirator and held in field
collection vials. The first specimen was taken while feeding on a clerk
in the park office at 1800 hrs July 7, 1981. No sandflies were captured
with four CDC Light traps set 1 m above ground level in protected areas
along 0.5 km of the Frio river for 1 night, and within 20 m of the build
ings from which sandflies were taken for a second night. Other workers
have taken L_. diabolica in CDC Light traps (D.G. Young, personal
communication).
Meteorological conditions for the night of 7 July 1981 were 100%
overcast with intermittent rain, 25C, with winds gusting to 15 knots.
I noticed l. diabolica feeding on my arms at 2200 hrs while sitting near
a light in an open shower/latrine building in an open area with a few
adjacent acacia trees. About 40 females were aspirated from the walls
of the building between 2300-0300 hrs with the majority taken on the
leeward side. The flies were strong fliers and aggressive biters.
The following day I searched 12 latrine/shower buildings in the
park, 7 in open, grassy areas and 5 near the river under large trees.
Those buildings near the river yielded no sandflies. The other 7
buildings yielded about 100 females and 5 males resting on the walls.
No flies were observed flying or feeding during the day. The night of
8 July was humid, 23C, partly cloudy with 1/4 moon, and winds gusting
to 15 knots. Collections made between 2100 hrs (onset of darkness)
and 0500 hrs produced about 60 females. About 50 flies were captured


-23-
Table 2-2. Comparison of the effect of larval diet composition pre
pared by the method of Young et al. (1981) on mean duration
of immature development time (egg-adult) of L. anthophora
at 20C and 28C, 90% RH.-
Temperature
(C)
Diet Component
Rabbit Chow
Purina #5315
Horse Chow
Purina #3501
Laboratory Chow
Number (?)
20
83.39.3
99.66.0
n = 19
n = 18
28
36.U2.9
38.54.2
43.54.1
n = 36
n = 34
n = 60


GENERAL RATIONALE
The primary goal of this research was to conclusively demonstrate
transovarian transmission of a Phlebovirus in a sandfly for the first
time. A virus (Rio Grande), non-pathogenic for humans, was selected
because it could be safely studied. The potential vector sandfly,
Lutzomyia anthophora, is not anthropophilic and therefore is a safe
subject for study.
Before transmission experiments could be undertaken it was neces
sary to first develop sandfly rearing and colonization techniques. In
order to plan and execute transmission experiments some aspects of the
laboratory biology of l. anthophora had to be elucidated.
After transovarian transmission was demonstrated, I realized that
this alone could not account for the distribution of neutralizing anti
body in various animals from south Texas in view of the fact that L_.
anthophora apparently does not feed on all of them. In order to more
fully understand the ecology of Rio Grande virus, preliminary studies
of other hematophagous insects, J_. diabolica and Triatoma gerstaeckeri,
were undertaken.
In a broad sense it must be acknowledged that no single mechanism
such as transovarian transmission can account for the maintenance of a
virus when the species of interest is sympatric with other hematophages.
Each insect species that feeds on the host must be studied to determine
its relative role in the maintenance of a pathogen.
-1-


ACKNOWLEDGEMENTS
The question of how to thank someone for his years of friend
ship, guidance, and wise counsel remains enigmatic. During the course
of this program I have incurred many debts which I will hopefully re
pay through contributions to the science.
The members of my committee I thank for their guidance and sup
port are Dr. Harvey Cromroy, chairman, Dr. Jerry Butler, Dr. David
Young, Dr. Donald Hall, and Dr. Stephen Zam. Mrs. Adele Koehler
typed the manuscript. Special thanks are in order to
Dr. David Young for suggesting this project, for his constant
support, and for his selfless assistance in the sandfly
colonization efforts;
Dr. Jerry Butler for generously providing a laboratory and much
of the equipment used for this project;
Dr. Robert Tesh, Yale Arbovirus Research Unit, for his generous
sharing of time, equipment, and knowledge that made the
virus transmission experiments possible;
E. Ann Ellis for her long hours of instruction on electron
microscopy and histology;
Diana Simon and Debbie Boyd who facilitated the daily accomplish
ment of much of this research;
Maj. Peter Perkins with whom many hours of camaraderie were
shared and with whom experimental ideas were generated;
v


TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS v
LIST OF TABLES ix
LIST OF FIGURES xi
ABSTRACT xi i i
GENERAL RATIONALE 1
SECTION
I TECHNIQUES FOR LABORATORY REARING OF SANDFLIES (DIPTERA:
PSYCHODIDAE) 2
Introduction and Literature Review 2
General Techniques 3
Larval Rearing 5
Sugar Feeding 5
N-butyl Pthalate 7
Adult Feeding 7
Lid Cleaning 8
Adult Feeding Cages 8
Field Collection, Feeding Containers 12
Individual Oviposition and Rearing Containers 15
Aspirators 15
II COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia anthophora (ADDIS) (DIPTERA: PSYCHODIDAE) 17
Introduction and Literature Review 17
Field Collections 17
Immature Behavior and Development 18
Time of Eclosin . 24
Mating 24
Female Age at First Feeding 27
Feeding: Hosts 33
Feeding: Temperature Preference 34
Feeding: Behavior 36
Feeding: Lymph 39
Refeeding 41
Peri trophic Sac Rupture 41
Productivity 42
Longevity 45
vii


-61-
Discussion
The existence of a transovarian transmission cycle for the
Phlebotomus group viruses has long been suspected (Doerr et al. 1909,
Whittingham 1924). In the absence of virus isolations, Russian workers
(Moshkovski 1937, Petricheva and Alymov 1938) demonstrated transmission
of an etiologic agent for Papataci fever through the eggs of £_. papataci
and demonstrated overwintering of the agent in larvae of the same
species. The recovery of phleboviruses from wild caught male sandflies
by several workers (Schmidt et al. 1971, Aitken et al. 1975, Tesh et al.
1974, 1977) lends further evidence to the existence of transovarian
transmission in this group. Transovarian transmission of another virus
serogroup transmitted by sandflies, Vesicular Stomatis Virus, has been
demonstrated by Tesh et al. (1972).
Despite overwhelming evidence indicating that Papataci fever virus
and other phleboviruses are transmitted by phlebotomine sandflies
(Schmidt 1971, Tesh 1975), no quantitative laboratory studies have been
conducted to demonstrate transmission by bite or by transovarial means.
The lack of controlled experiments has been partially a function of the
difficulty encountered in rearing sandflies.
In this series of experiments we were unable to demonstrate
transmission by bite due to the lack of susceptible laboratory animals,
by the fact that L_. anthophora refused to feed on suckling mice, and
a lack of knowledge on virus dynamics in a rodent host. Therefore,
infection was established by means of intrathoracic injection. Sand
flies infected by injection transmitted Rio Grande virus to 54.8% of
their progeny. This is the first demonstration of transovarian trans
mission of a Phlebovirus by sandflies.


Table 2-5.
Comparison
on Di del phi.
of effects of
s marsupial is
blood vs. blood
(opossum).
and sugar as an
energy source
for L. anthophora
fed
Energy
Source
% Females
Fed on
Blood
l Bloodfed
Females
Laying
No Eggs
Mean Number
Eggs/Female
Preoviposition
Period (Days)
% Fertility
of Females
Laying Eggs
% Females
Peritrophic
Sac Rupture
% Females
Fed on
Serum
Blood
40.8
(147)*
39.6
(60)
17.68.4
4.51.2
65.2
11.7
1.7
Blood +
Sugar
49.7
(282)
19.8
(116)
28.013.2
6.21.8
83.3
4.9
0.7
( ) indicates number in sample.
-32-


-88-
Romoser, W.S., and M.E. Rothman. 1973. The presence of a peritrophic
membrane in pupal mosquitoes (Diptera: Culicidae). J. Med.
Entomol. 10(3):312-14.
Rosen, L., and D. Guebler. 1974. The use of mosquitoes to detect and
propagate dengue viruses. Am. J. Trop. Med. Hyg. 23(6):1153-60.
Schmidt, J.R., and M.L. Schmidt. 1965. Observations on the feeding
habits of Phlebotomus papatasi (Scopoli) under simulated natural
conditions. J. Med. Entomol. 2(3):225-30.
Schmidt, J.R., M.L. Schmidt, and M.I. Said. 1971. Phlebotomus fever
in Egypt: Isolation of Phlebotomus fever viruses from Phlebotomus
papataci. Am. J. Trop. Med. Hyg. 20(3):483-90.
Shaw, P.K., L.T. Quigg, D.S. Allain, D.D. Juranek, and E.R. Healy.
1976. Autochthonous dermal leishmaniasis in Texas. Am. J. Trop.
Med. Hyg. 25:788-96.
Sherlock, I.A. 1957(a). Sobre 0 Phlebotomus lenti Mangabeira, 1938
(Diptera: Psychodidae). Rev. Brasil. Biol. 17(1):77-88.
Sherlock, I.A. 1957(b). Sobre 0 Phlebotomus renei Martins, Falcao and
Silva, 1956 (Diptera: Psychodidae). Rev. Brasil. Biol. 17(4):
547-56.
Sherlock, I.A., and M. Carneiro. 1963. Descricao das faces imatures
do Phlebotomus bahiensis Mangabeira and Sherlock, 1961 (Diptera:
Psychodidae). Separata de Mems. Inst. Oswaldo Cruz 61(3):491-94.
Simpson, M.H., J.F. Mullins, and O.J. Stone. 1968. Disseminated aner
gic cutaneous leishmaniasis. Arch. Dermatol. 97:301-3.
Stewart, C.D., and J.F. Pilcher. 1945. American leishmaniasis: Report
of an autochthonous case. Arch. Dermatol. 51:124-28.
Tesh, R.B. 1975. Multiplication of Phlebotomus fever group arboviruses
in mosquitoes after intrathoracic inoculation. J. Med. Entomol.
12(1):1-4.
Tesh, R.B., and B.N. Chaniotis. 1975. Transovarial transmission of
viruses by phlebotomine sandflies. Ann. N.Y. Acad. Sci. 266:
125-34.
Tesh, R.B., B.N. Chaniotis, and K.M. Johnson. 1972. Vesicular stomatitis
virus (Indiana serotype): Transovarial transmission by Phlebotomine
sandflies. Science 175:1477-79.
Tesh, R.B., B.N. Chaniotis, P.H. Peralta, and K.M. Johnson. 1974.
Ecology of viruses isolated from Panamanian phlebotomine sand
flies. Am. J. Trop. Med. Hyg. 23(2):258-69.


I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dr. Harvey L. Cromroy, Chairman
Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dry Jerpi
Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
C>
Dr. David G. Young
Assistant Professor of Entomology
and Nematology


Figure 1-3. Sandfly feeding cage--A modified aquarium with plaster of Paris bottom and back.


-75-
Figure 7-1. Scanning electron micrographs of eggs of four sandfly
species. (1) Lutzomyia diablica, (2) Lutzomyia shannoni,
(3) Lutzomyia vexator, (4) Lutzomyia cruciata spp.


-50-
9 July in the same buildings as on the 8th but in lesser numbers.
j_. diablica appears to be strongly attracted to light especially on
warm, humid, overcast nights.
Approximately 10% of the 250 females collected were engorged with
blood. All of the unfed flies were allowed to feed on my forearms
through the screen lid of the container on 9, 10 July. More than 50%
avidly took a blood meal.
For air shipment the plaster in the collecting containers was
dampened and vials were packed in a sealed plastic container with wet
towels. When received 8 hrs later 90% of the females had died after
being trapped in condensation on the sides of the containers. There
fore, specimens shipped by air should be shipped in vials with the
plaster dry and wrapped in slightly damp towels.
Feeding
Although Lindquist (1936) reported l. diabolica feeding on humans
between 2000 hrs and 2400 hrs it will feed during all hours of darkness.
When a cage of flies is held against the skin of a host, females will
feed irrespective of light conditions.
When exposed to anesthetized hosts in a feeding cage or while
holding a feeding container against the host skin, J_. diabolica fed
on the following animals: Homo sapiens (human; Figure 3-2), Canis
familiaris (dog), Neotoma micropus (woodrat), Mesocricetus auretis
(Syrian hamster), Sciurus carolinensis (grey squirrel), Oryctolaqus
cuniculus (domestic rabbit), Pi del phis marsupial is (opossum), Bos
Taurus (calf), and Equus caballus (horse). Only one unsuccessful


-87-
Nayar, J.K., and D.M. Sauerman. 1975(a). The effects of nutrition on
survival and fecundity in Florida mosquitoes. Part 1. Utiliza
tion of sugar for survival. J. Med. Entomol. 12(1):92-8.
Nayar, J.K., and D.M. Sauerman. 1975(b). The effects of nutrition on
survival and fecundity in Florida mosquitoes. Part 2. Utiliza
tion of a blood meal for survival. J. Med. Entomol. 12(1):
99-103.
Obijeski, J.F., A.T. Marchenko, D.H.L. Bishop, B.W. Cann, and F.A.
Murphy. 1974. Comparative electrophoretic analysis of the virus
proteins of four Rhabdoviruses. J. Gen. Virol. 22:21-33.
Obijeski, J.F., D.H.L. Bishop, F.A. Murphy, and E.L. Palmer. 1976.
Structural proteins of La Crosse Virus. J. Virol. 19(3):985-97.
Parman, D.C. 1919. Notes on Phlebotomus species attacking man. J.
Econ. Entomol. 12:211-12.
Perfiliev, P.P. 1966. Fauna of USSR. Diptera: Phlebotomidae (sand
flies). Akademiya Nauk SSSR. Zoologicheskii Institut. (Israel
Programme for Scientific Translation, Jerusalem).
Petrischeva, P.A., and A.J. Alymov. 1938. On transovarial transmission
of virus of papataci fever by sandflies. Arch. Biol. Sci. 53:
138-44.
Pippin, W.F. 1970. The biology and sector capability of Triatoma
sanguisuga texana Usinger and Triatoma gerstaeckeri (Stal) com-
pared with Rhodnius prolixus (Stal) (Hemiptera: Triatominae).
J. Med. Entomol. 7(1):30-45.
Quattelbaum, E.C., and G.R. Garner. 1980. A technique for preparing
Beauveria spp. for scanning electron microscopy. Can. J. Bot.
58:1700-03.
Ready, P.D. 1978. The feeding habits of laboratory-bred Lutzomyia
lonqipalpis (Diptera: Psychodidae). J. Med. Entomol. 14(5):
545-52.
Ready, P.D. 1979. Factors affecting egg production of laboratory-
bred Lutzomyia Jon^ij^aljm (Diptera: Psychodidae). J. Med.
Entomol. 16(5):413-23.
Reed, L.J., and H. Muench. 1938. A simple method for estimating fifty
percent endpoints. Am. J. Hyg. 27:493-97.
Romoser, W.S. 1974. Peritrophic membranes in the midgut of pupal and
preblood meal adult mosquitoes. J. Med. Entomol. 11(4):397-402.
Romoser, W.S., and E. Cady. 1975. The formation and fate of the peri
trophic membrane in adult Culex nigripalpus (Diptera: Culicidae).
J. Med. Entomol. 12(3):371-78.


SECTION IX
SUMMARY
During the four year course of this study the following objectives
were achieved:
1. Methods for laboratory culture of phlebotomine sandflies were
developed. Use of these culture techniques will permit detailed quanti
tative study of the vector competence of sandflies for viral and other
parasitic diseases. To date twelve species have been reared in closed
colony for as many as 25 generations using these techniques.
2. Detailed studies on the laboratory biology of two species of
Neotropical phlebotomine sandfly, Lutzomyia anthophora and Lutzomyia
diablica were conducted over 16 and 7 generations, respectively.
3. Transovarian transmission of a Phlebovirus by a sandfly was
conclusively demonstrated for the first time. Nearly 55% of J_.
anthophora females injected with Rio Grande virus transmitted it to their
progeny.
4. The oocyte surface sculpturing of 5 species of Neotropical
sandflies were described for the first time with the scanning electron
microscope. The topographic patterns can be used to identify thse
sandfly species.
5. A method for purification of Rio Grande virus was developed.
The virus was purified and photographed in a transmission electron
microscope. This is one of the first recorded purifications and char
acterizations of a Phlebovirus.
-82-


Figure 2-3. Multiwell plate with lid used for rearing individual
larvae.


-89-
Tesh, R.B., E. Javadian, and N. Nadim. 1977. Studies on the epidemi
ology of sandfly fever in Iran. I. Virus isolates obtained from
Phlebotomus. Am. J. Trop. Med. Hyg. 26(2):288-97.
Tesh, R.B., and D.A. Shroyer. 1980. The mechanism of arbovirus
transovarial transmission in mosquitoes: San Angelo virus in
Aedes albopictus. Am. J. Trop. Med. Hyg. 29(6):1394-1404.
Thurman, D.C. 1945. The biology of Triatoma gerstaeckeri. J. Econ.
Entomol. 38(5):597-98.
Vargas, L. 1952. Nota Sobre Los Flebotomus de la Zona de Iguala,
Estado de Buerroro. Revista Invest. Clin. Mexico 4(1):47-53.
Ward, R.D. 1977. The colonization of Lutzomyia flaviscutellata
(Diptera: Psychodidae), a vector of Leishmania mexicana amazonensis
in Brazil. J. Med. Entomol. 14(4):469-76.
Ward, R.D., and P.A. Ready. 1975. Chorionic sculpturing in some sand
fly eggs. J. Ent. (A) 50(2):127-34.
Whittingham, H.E. 1924. Etiology of phlebotomus fever. J. State
Med. 32:461-69.
Young, D.G. 1972. Phlebotomine sandflies from Texas and Florida
(Diptera: Psychodidae). Fla. Entomol. 55(1):61-4.
Young, D.G., and P.V. Perkins. 1982. A review of the Lutzomyia sand
flies of North America (Diptera: Psychodidae). In press.
Young, D.G., P.V. Perkins, and R.G. Endris. 1981. A larval diet for
rearing Phlebotomine sand flies (Diptera: Psychodidae). J. Med.
Entomol. 18(5):446.
Zimmerman, J.H., H.D. Newson, G.R. Hooper, and H.A. Christensen. 1977.
A comparison of the egg surface structure of six anthropophilic
Phlebotomine sand flies (Lutzomyia) with the scanning electron
microscope (Diptera: Psychodidae). J. Med. Entomol. 13(4-5):574-79.


-9-
Table 1-1. Dosage required to anesthetize animals for 30-60 min with
Ketamine hydrochloride (100 mg/ml) injected IM.
Animal
Dosage (ml)
Mouse
0.05/adult
Squirrel
0.1/200 gm
Woodrat
0.2/adult
Opossum
0.25-0.30/2 kg
Rabbit
0.3-0.4 mg/kg


-52-
March. The failure of eggs to hatch when laid by a single female was
observed from July to December at 20C, 24C, 28C, and 30C.
Development of individual immatures at various temperatures was
not studied because of insufficient numbers resulting from "partial
frtility."
I have observed that the egg-adult development time of J_. diablica
is 3-6 days less than the 36 days required for L_. anthophora at 28C.


SECTION VII
A COMPARISON OF OOCYTE TOPOGRAPHY OF FIVE PHLEBOTOMINE SANDFLIES
(Lutzomyia) WITH THE SCANNING ELECTRON MICROSCOPE
(DIPTERA: PSYCHODIDAE)
Introduction
The egg surface structure of 19 neotropical Phlebotomine species
has been described (Zimmerman et al. 1977, Ward and Ready 1975) using
the scanning electron microscope (SEM). Ward and Ready (1975) noted
three species-specific topographic patterns, i.e., polygonal, parallel
ridging, and volcano-like. Several authors (Chaniotis and Anderson
1964, Addis 1945,Lindquist 1936, Barreto 1941, and Sherlock 1957a,b, 1963)
described and figured the eggs of 17 neotropical sandfly species using
light microscopy. After examining the literature cited and eggs from
the 5 species described herein we propose adding another category to
the patterns of Ward and Ready (1977); that is, parallel ridges connected
or parallel ridges unconnected.
The classification of eggs of 41 species of New World sandflies
according to the proposed scheme is presented in Table 7-1.
Materials and Methods
Eggs were obtained from females reared in laboratory colonies.
The preparation method of eggs for SEM based on the work of Quattlebaum
and Carner (1980) is as follows:
-72-


-86-
Killick-Kendrick, R., A.J. Leaney, and P.D. Ready. 1973. A laboratory
culture of Lutzomyia lonqipalpis. Trans. R. Trop. Med. Hyq.
67:434.
Kill ick-Kendrick, R. A.J. Leaney, and P.D. Ready. 1977. The estab
lishment, maintenance and productivity of a laboratory colony of
Lutzomyia lonqipalpis (Diptera: Psychodidae). J. Med. Entomol.
13(4-5):429-40.
Kitselman, C.M., and A.W. Grundmann. 1940. Equine encephalomyelitis
isolated from naturally infected Triatoma sanguisuga (Le Conte).
Kansas Agr. Exp. Sta. Tech. Bull. 50.
Lent, H., and P. Wygodzinsky. 1979. Revision of the Triatominae
(Hemiptera: Reduviidae), and their significance as vectors of
Chagas disease. Bulletin of the American Museum of Natural
History. Vol. 163, Art. 3. New York.
Lewis, D.J. 1975. Functional morphology of the mouth parts in New
World phlebotomine sandflies (Diptera: Psychodidae). Trans. R.
Entomol. Soc. Lond. 126(4):497-532.
Lewis, D.J. 1977. Phlebotomine sandfly research. Med. Entomol. Cont.
Symp. Roy. Soc. Trop. Med. Hyg., London 94-99.
Lindquist, A.W. 1936. Notes on the habits and biology of a sand fly,
Phlebotomus diabolicus Hall, in Southwestern Texas (Diptera:
Psychodidae). Proc. Entomol. Soc. Wash. 38(2):29-32.
Mangiafico, J.A., L. Whitman, and R.C. Wallis. 1968. Survival of
Western Equine Encephalitis virus in Triatominae. J. Med.
Entomol. 5(4):469-73.
McCrea, J.F., R.S. Epstein, and W.H. Barry. 1961. Use of potassium
tartrate for equilibrium density gradient centrifugation of animal
viruses. Nature 89(4760):220-21.
McLean, R.G., D.M. Szmyd, and C.H. Calisher. 1982. Experimental
studies of Rio Grande virus in rodent hosts. Am. J. Trop. Med.
Hyg. 31(3):in press.
The Merck Index. 1976. An Encyclopedia of Chemicals and Drugs.
Windholz, M. et al., eds. 9th ed. Merck & Co. Inc. Rahway, NJ.
Moshkovski, S.D., N.A. Diomina, V.D. Nossina, E.A. Pavlova, J.L. Liv-
chitz, H.J. Pels, and V.P. Roubtzova. 1937. Researches on sand
fly fever. Part VIIL Transmission of sandfly fever virus by
sandflies hatched from eggs laid by infected females. Med. Par
sito!. Parazit. Bolezn. 6:922-37.
Najera, A. Diaz. 1971. Presencia de Lutzomyia (Lutzomyia) diablica
(Hall, 1936) en Muzquiz, Coahuila, Mexico (Diptera: Psychodidae).
Revista de Investigacin en Salud Publica 31(2):62-6.


SECTION V
RIO GRANDE VIRUS AND Triatoma qerstaeckeri
(STAL) (HEMIPTERA: REDUVIIDAE)
Introduction and Literature Review
The hematophagous Hemipterans, the Cimicidae and the Triatominae,
have been considered ideal potential vectors of arboviruses because of
the large blood meal ingested, their relative longevity, and their
cosmopolitan feeding habits. Kitselman and Grundman (1940) reported
isolating Western Equine Encephalitis (WEE) virus from Triatoma sangui
suga taken in a Kansas pasture where animals had died of the disease in
previous years. Mangiafico et al. (1968) found that 2 species of
Triatome, jl. pro!ixus and T. infestans, would harbor WEE virus 14-20
days when unpunctured. When punctured to simulate cannabalistic feed
ing virus survived 98 days and one bug transmitted the virus by bite.
Justines and Sousa (1977) obtained similar results with punctured bugs
and bugs infected with Trypanosoma cruzi. Hayes et al. (1977) found the
cliff swallow bug, Cimicidae, to be capable of overwintering and trans
mitting Ft. Morgan virus (Calisher et al. 1980).
In view of the findings noted and realizing that Triatoma ger-
staeckeri was known to feed on all of the animals (Lent and Wygodzinsky
1979), in which Calisher et al. (1977) had found neutralizing antibody
to Rio Grande virus, the vector potential of T. qerstaeckeri was in
vestigated. Thurman (1945) and Pippin (1970) reported finding
-64-


-13-
Figure 1-4. Cylindrical adult feeding cage.


-45-
Longevity
Adult longevity for males and non-bloodfed females was determined
by holding sandflies individually and placing either distilled water or
30% honey solution on the screen lid. Results are presented in Table
2-8. Sugar-fed adults lived 40-45% longer than those fed on distilled
water only.
These results agree with the work of Nayar and Sauerman (1975a,b)
and Edmund Davis (personal communication, 1981) who showed that sugar
feeding increased longevity of several mosquito species and Culi coi des
mississippiensis (Hoffman), respectively.


-60-
Table 4-2. Presence of Rio Grande virus in (1) Neotoma micropus and
(2) Peromyscus leucopus bled daily for 7 days after sub
cutaneous inoculation.
Days Post Inoculation
Animal 1 234567
Neotoma micropus
Peromyscus leucopus
(a)
(b)
0
+ + 0 0
0 0
0
0 + 00
0 0
0
+ + 0 0
0 0


LIST OF TABLES
Table Page
1-1. Dosage required to anesthetize animals for 30-60 min
with Ketamine hydrochloride (100 mg/ml) injected IM. . 9
2-1. Mean duration (days) of immature stages of antho-
phora at 90% RH and 4 constant temperature regimes:
20C, 24C, 28C, and 32C in contrast to the obser
vations of Addis (1945b) made at 28-29C. Larvae
reared on the diet of Young et al. (1981) 22
2-2. Comparison of the effect of larval diet composition pre
pared by the method of Young et al. (1981) on mean
duration of immature development time (egg-adult) of
J_. anthophora at 20C and 28C, 90% RH 23
2-3. L. anthophoraComparison of mean development time
Tdays) for males and females reared at 20C, 24C,
28C, and 32C, 90% RH 25
2-4. 1. anthophoraAdul t sugar feeding, frequency, age of
feeding (days), time required for digestion (days) at
20C and 28C, 90% RH 31
2-5. Comparison of effects of blood vs. blood and sugar as
an energy source for L_. anthophora fed on Pidel phis '
marsupial is (opossum) 32
2-6. Temperature (C) of body regions of anesthetized and
non-anesthetized hosts for J_. anthophora in laboratory
culture 35
2-7. Fecundity, percent of bloodfed females that laid no
eggs, and preoviposition period (days) for 12 genera
tions of L. anthophora reared at 24C and 28C,
90% RH 44
2-8. Comparison of longevity (days) of J_. anthophora males
and females fed on either distilled water or 30% honey
solution at 24C, 90% RH 46
3-1. Lutzomyia diabolicaFecundity, preoviposition period
(days), and mortality factors for 3 generations in
laboratory culture at 28C, 90% RH 53
IX


-20-
burrowing appeared dependent on moisture content of the medium and the
stage of development.
Larval emergence, behavior, and pupation were consistent with the
observations of Chaniotis (1967), Johnson and Hertig (1961), and
Gemetchu (1976). In contrast to the observation of Killick-Kendrick
et al. (1977) with J_. longipalpis, no cannabalism was observed among
4th instar larvae when starved. No effort was made to discover the
larval habitat in nature although it is presumably in or under the
woodrat nest (Young 1972).
Larval development rates at 4 different temperatures (20C, 24C,
28C, and 32C) were determined by rearing individual larvae in wells
of microtitre plates that were 1/3 filled with plaster of Paris.
Initially microtitre plates with 96 wells were used but the wells
proved too small for 4th instar larvae. Twenty-four well microtitre
plates were satisfactory. Attempts to use the lids designed for the
multiwell plates were not successful because they do not seal well and
larvae moved between wells. Lids (9 cm x 13 cm x 5 rrni) made from
Plexiglas and secured with elastic bands solved this problem (Figure
2-3). After several days in chambers at 90% relative humidity (RH) it
was necessary to add drops of H^O in each well until the plaster
appeared damp.
Development times for immatures are presented in Table 2-1. The
difference between the results of Addis (1945b) shown in Table 2-1
and those obtained in this study at 28C are probably attributable to
differences in larval diet.
Three larval diets at 28C and two at 20C were compared to determine
the effect of diet on immature development time (Table 2-2). The diet pre
pared with Purina Rabbit Chow #5315 is that reported by Young et al. (1981).


Figure 2-
. L. anthophoraTemporal age distribution at feeding and death of unfed females
at 28C, 90% RH.
-28-


-69-
Electron Microscopy
A drop of virus suspension was placed on a Formvar carbon coated
grid and allowed to dry for 1 min before the excess fluid was wicked
off with filter paper. The grid was then negatively stained for 45 sec
with 2%, pH 6.8, phosphotungstic acid with KOH, using 50 yg/ml
Bacitracin as a spreading agent (Gregory and Pirie 1973). Specimens
were examined at 75 KV, 50,000x and 100,000x in a Hitachi 600 Trans
mission Electron Microscope.
Results
Electron micrographs (Figure 6-1) show that the virion is spherical
and possesses an envelope bearing small spikes.
The virion is 71 nm in diameter as determined using a reference
catalase crystal. This is within the size range of 60-90 nm that is
characteristic of the Bunyaviridae (Bishop et al. 1980).
Piscussion
The size and morphology of the Rio Grande virion is consistent
with those described for the genus Phiebovirus (Bishop et al. 1980).
An additional purification step of centrifuging the virus suspen
sion in a 20-70% (w/v) sucrose gradient at 4C and 35,000 g for 4 hr was
not used since it was possible to recover the virus after the equilibrium-
density centrifugation in potassium tartrate.
In order to verify that the virions shown in the electron micro
graphs retained infectivity, 1 ml of virus suspension was adsorbed onto


SECTION II
COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia anthophora (ADDIS)(DIPTERA: PSYCHODIDAE)
Introduction and Literature Review
Lutzomyia anthophora was first collected while feeding on rabbits
in Uvalde, Uvalde Co., Texas (Addis 1945a). Subsequently it was re
ported from NE Mexico (Fairchild and Hertig 1956), SW Mexico (Vargas
1952), and SE Texas (Young 1972). Young (1972) reported finding
l. anthophora in the nest of the plains woodrat, Neotoma micropus,
with which it appeared to enjoy a close host-parasite relationship.
Calisher et al. (1977) again reported the association of L,. anthophora
and Neotoma nests when suggesting that Rio Grande virus could be main
tained in the woodrat population by transovarian transmission in this
sandfly.
Addis (1945b) described the immature stages and the life cycle
of l. anthophora after rearing 72 flies from egg-adult at 28-29C.
In this section detailed investigations of the colonization and biology
of J_. anthophora are reported.
Field Collections
Sandflies used to start the colony were collected by R.G. Endris
and D.G. Young with the assistance of G.B. Fairchild and R.N. Johnson
in the area of E and NE of Brownsville, Texas, from Neotoma nests in
-17-


-62-
Calisher et al. (1977) found neutralizing antibody to Rio Grande
virus in woodrats, opossums, gopher tortoises, horses, several species
of small rodents, birds, and a horned toad. I was unable to induce
J_. anthophora to feed on gopher tortoises, or horses. This suggests
that J_. anthophora probably is not the only arthropod vector of Rio
Grande virus. Other hematophagous arthropods that I recovered from
the woodrat nests were Ornithodorus dugesi (often identified as
0. tulaje), Triatoma gerstaeckeri, Triatoma sanguisuga, and Triatoma
neotomae. Johnson (1966) also reported several species of fleas and
Ixodidae from the woodrat nests. It is highly likely that mosquitoes
also use the nests for resting sites. Some of these other arthropods
are catholic in their feeding behavior and could possibly transmit
Rio Grande virus to animals not fed on by l. anthophora, and may be
capable of transovarian transmission.
The ecology of Rio Grande virus remains to be thoroughly studied.
Although Neotoma micropus and Peromyscus leucopus are susceptible to
infection by subcutaneous inoculation the infection is transient.
McLean et al. (1982) confirmed the results in Table 4-2 in that the
viremia in woodrats is short-lived, 2.5 days, and of low titer, mean
3.65 iog^Q PFU/ml in Vero cell culture. As yet, oral infection of
J.. anthophora has not been demonstrated nor is the minimum infective
oral dose known. McLean et al. (1982) also determined that nearly all
the woodrats developed neutralizing antibody thus becoming refractory
to infection for life. This indicates that only young woodrats are
likely to be susceptible to infection after maternal antibody is no
longer present and they could only serve as amplifying hosts for 2.5
days during their lifetime. The high transovarian transmission rate


-56-
Suckling mice (Suisse variety) from two litters were inoculated
5
with 0.2 ml 10 PFU/ml RGV. One litter was inoculated subcutaneously
and the other was inoculated intracerebrally. One mouse from each
treatment was bled daily from the carotid artery. All blood samples,
0.1 ml, were diluted with 1.0 ml PBS with 0.05% gelatin and frozen at
-70C for titration.
Virus Assay
Individual flies were triturated in 1.0 ml of dilutent in a sterile
2 ml Ten Broeck tissue grinder. The diluent was phosphate-buffered
saline, pH 7.2, containing 0.5% gelatin and 30% heat inactivated bovine
serum. Sandfly suspensions were centrifuged at 10,000 rpm for 30 min.
The supernatant was prepared in serial ten-fold dilutions from 10"* to
10 Four tube cultures of Vero cells were then inoculated with 0.1 ml
of each dilution and incubated. Daily samples of 5 infected flies
were frozen at -70C for virus titration to determine the growth of
Rio Grande virus (RGV) in the sandflies. Surviving females were offered
an anesthetized hamster daily for a 60 min period of feeding. After
feeding, engorged females were held at 25C, 80% RH, in individual
oviposition containers.
Fourth instar larvae and pupae were inoculated with RGV intra-
abdominally by the same method as the adults.
Suckling mouse blood was tested only at 1:10 dilution.
Infection of Rodents
To determine the fate of RGV in the natural hosts two,white-footed
mice, Peromyscus leucopus, and one female Neotoma micropus derived from


SECTION VI
PURIFICATION OF RIO GRANDE VIRUS
Introduction
Since Calisher et al. (1977) first characterized Rio Grande virus
little other descriptive work has been performed. Because of its
ecological relationship to other members of the group, Rio Grande virus
was placed in the genus Phiebovirus of the family Bunyaviridae by
Bishop et al. (1980). In this section a purification scheme for the
virus is described and electron micrographs of the virus are presented.
Materials and Methods
Virus
Rio Grande virus (Strain TMB4-719) was kindly supplied by Dr.
Robert Tesh, Yale Arbovirus Research Unit. Additional quantities of
virus were produced by intracerebral inoculation of 2-4 day old suck
ling mice with 0.01-0.02 ml stock virus. Four days after inoculation
the mouse brains were harvested and triturated in 2 ml Ten Broeck
tissue .grinders with lx sterile phosphate buffered saline (PBS), pH 7.4
Infection of Cells and Virus Purification
The purification scheme detailed below was based on several others
previously used for arboviruses (Kaariainen et al. 1969, Obijeski et al
-67-


I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dr. Donald W. Hall
Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dr. Stephen G. ImJ
Associate Professor of Microbiology
and Cell Science
This dissertation was submitted to the Graduate Faculty of the College
of Agriculture and to the Graduate Council, and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
May 1982
Dean,
Dean for Graduate Studies and
Research


NUMBER OF ADULTS
-25-
Table 2-3. J_. anthophora--Comparison of mean development time (days)
for males and females reared at 20, 24C, 28C, and
32C, 90% RH.
Temperature
(c)
Sex
20
24
28
32
Female
83.37.0
n = 10
50.73.6
n = 66
40.63.7
n = 73
34.64.0
n = 36
Male
83.311.7
n = 9
48.24.1
n = 63
37.53.3
n = 57
32.73.3
n = 52
Difference
0.0
2.5
3.1
1.9
Figure 2-4. Eclosin distribution of 127 _L. anthophora males and
females from F3 generation in a laboratory colony at
24C, 90% RH.


A man's reach should exceed his grasp.
Robert Browning


Table 3-1. Lutzomyia diabolica--Fecundity, preoviposition period (days), and mortality factors for 3
generations in laboratory culture at 28C, 90% RH.
Genera
tion
n
% Bloodfed
Females Laying
No Eggs
Fecundity
Preoviposition
Period (Days)
% Females
Peri trophic
Sac Rupture
% Females
Fed on
Serum
X
S
Maximum #
X
S
Range
1
27
59.3
39.6
11.2
58
5.7
1.7
3-8
11.1
0.0
2
21
28.6
32.0
14.0
51
5.5
1.6
3-9
4.8
4.8
3
51
58.8
36.2
16.0
64
6.2
2.3
3-12


-42-
(1973), Romoser (1974), Romoser and Cady (1975) described the lattice
like sac in mosquito larvae, pupae, and adults. The similar structure
of the sandfly peri trophic membrane was investigated in detail by Gemetchu
(1974). He states that it is formed within 30 min after a blood meal is
taken and breaks up about 3 days later.
A phenomenon that occurred consistently in each generation was
the apparent rupture of the peritrophic sac and the midgut epithelium.
Blood from the gut penetrated all parts of the insect including the
thorax, legs, and antennae (Figure 2-12). The specimens that were killed
by this phenomenon possibly died as a result of changes in the hemolymph
osmoticum and the release of digestive enzymes. In 3 generations (Fg,
Fg, F Q) the frequency of "peritrophic sac rupture" occurred in 9.9%,
6.6%, and 8.7% of the bloodfed sandflies, respectively. Death of the
flies followed feeding in 1-4 days with 75% dead in less than 24 hr.
This phenomenon is not limited to J_. anthophora since I have also
observed it in Ornithodorus turicata, Ornithodorus dugesi, Triatoma
gerstaeckeri, Triatoma sanguisuga, Triatoma neotomae, and Lutzomyia
diabolica. The mechanism of gut integrity disruption remains an enigma
and does not seem related to the mammalian blood source.
Productivity
Egg production for 12 of 15 generations of _L. anthophora reared
to date are presented in Table 2-7. The preoviposition period indi
cated in the table represents the time from blood meal ingestion to
egg laying, i.e., the period required for egg development. Since
sandflies often extrude 1-3 infertile eggs at death only those females


-84-
Calisher, C.H., T.P. Monath, D.J. Muth, J.S. Lazuick, D.W. Trent, D.B.
Francy, G.E. Kemp, and F.W. Chandler. 1980. Characterization of
Fort Morgan virus, an alphavirus of the western equine encephalitis
complex in an unusual ecosystem. Am. J. Trop. Med. Hyg. 29(6):
1428-40.
Chaniotis, B.N. 1967. The biology of California phlebotomus (Diptera:
Psychodidae) under laboratory conditions. J. Med. Entomol. 4(2):
221-33.
Chaniotis, B.N. 1974. Sugar feeding behavior of Lutzomyia trapidoi
(Diptera: Psychodidae) under experimental conditions. J. Med.
Entomol. 11(1):73-9.
Chaniotis, B.N. 1975. A new method for rearing Lutzomyia trapidoi
(Diptera: Psychodidae), with observations on its development and
behavior in the laboratory. J. Med. Entomol. 12(2):183-88.
Chaniotis, B.N., and J.R. Anderson. 1964. Notes on the morphology
and laboratory rearing of Phlebotomus vexator occidental is
(Diptera: Psychodidae). Pan-Pacific Entomologist 40(1):27-32.
Clewley, J., J. Gentsch, and D.H.L. Bishop. 1977. Three unique viral
RNA species of Snowshoe Hare and La Crosse Bunyaviruses. J. Virol.
22(2):459-68.
Disney, R.H.L. 1968. Evidence that Luztomyia diablica Hall is con-
specific with l. cruciata Coquillett (Diptera: Psychodidae). J.
Med. Entomol. 5(2):267-68.
Doerr, R., K. Franz, and S. Taursig. 1909. Das pappatacifieber.
Deutike. Leipzig.
Fairchild, G.B., and M. Hertig. 1956. Notes on the Phlebotomus of
Panama (Diptera: Psychodidae). XII. The group Anthophorous,
with descriptions of four new species from Panama and Mexico.
Ann. Entomol. Soc. Amer. 49(4):307-12.
Fine, P.E.M. 1975. Vectors and vertical transmission: An epidemiologic
perspective. Ann. N.Y. Acad. Sci. 166:173-94.
Gemetchu, T. 1974. The morphology and fine structure of the midgut
and peri trophic membrane of the adult female, Phlebotomus lonqipes
Parrot and Martin (Diptera: Psychodidae). Ann. Trop. Med. &
Parasitol. 68(1):111-24.
Gemetchu, T. 1976. The biology of a laboratory colony of Phlebotomus
lonqipes Parrot and Martin (Diptera: Psychodidae). J. Med.
Entomol. 12(6):661-71.
Gregory, D.W., and B.J.S. Pirie. 1973. Wetting agents for biological
electron microscopy. 1. General considerations and negative
staining. J. Microsc. 99(3):261-65.


-51-
attempt was made to feed flies on Ovis aries (sheep). J_. diabolica
feeds not only on the ears of mammalian hosts as does JL. anthophora
but also on the nose, around the eyes, or any other hairless or nearly
hairless areas.
Females feed within 24 hrs of eclosin. Feeding behavior is con
sistent with observations of Lindquist (1936).
Two of 10 females in a feeding cage took a bloodmeal from the
inguinal region of a dog that had been infected with Leishmania donovani
infantum at least 10 months earlier. Five days after feeding each fe
male had 150-200 promastigotes in the midgut.
Mating
Mating was observed under a wide range of light conditions and
before, after, and during feeding.
Egg Hatch and Fertility
No autogeny was observed in this species. Data on fecundity, pre-
oviposition period, and mortality factors are presented in Table 3-1.
First instar larvae do not exhibit synchronous egg hatching in
contrast to l. anthophora in which all the eggs of a single batch will
hatch within a 2 day period regardless of temperature. As many as 70%
of eggs laid by a single J_. diablica female often fail to hatch within
a 30 day period whereas nearly all the eggs laid by a single L. anthophora
female will hatch. The mechanism of this "partial fertility" phenomenon
of some J_. diablica eggs remains a mystery. Lindquist (1936) noted
an apparent diapause in the egg stage of _L. diabol ica from October to


-43-
which laid four or more eggs were considered to have oviposited. No
autogeny was observed with this species nor could it be induced by feed
ing mated females only water or sugar solutions. Egg laying was usually
completed in less than 24 hr but could require 2-3 days. Additional
blood meals are required for females to lay subsequent batches of eggs.
The first 3 generations were held at 24C and subsequent genera
tions were held at 28C with no apparent effect upon egg production. Vary
ing photoperiods also had no apparent effect on egg production or fertil
ity. The effect of blood meal source on fecundity was not investigated.
The number of females which did not take a blood meal was studied
in the generation and observed to represent 50.3% of the total
number eclosed. Those females plus the 19.8% blood fed females which
did not lay eggs indicate that 58.5% of the total number of females in
a given generation are non-productive. Despite the number of non
productive females, colony numbers could easily be increased to yield
as many insects as required for experimentation.
A phenomenon that was consistently observed in each generation
was the tendency for those females eclosing in the first half of the
generation cycle to lay the majority of the eggs and for many of those
emerging in the second half of the generation cycle to die without
ovipositing.
An attempt to maintain adults at 32C proved unsatisfactory since
of 36 females produced at that temperature, 11 fed on a mouse (69.4%
did not feed), 4/11 (36.4%) laid eggs, and 3/11 (27.3%) fed on serum (no
eggs). The mean number of eggs per female was 13.8 9.8. The number
of eggs laid by adults held at 32C was greatly reduced compared to
those held at 24C and 28C (Table 2-7).


-36-
to induce sandfly feeding the SM were placed on a cotton pad on a
variable temperature plate. Flies were released into a specimen con
tainer over the SM. During a 6 hr period the temperature was raised
from 26C (ambient) to 40C then returned to 26C. Of 30, 3 day old
adults none fed. A thermistor was taped to the SM to insure the skin
temperature was the activation source for the heater.
The reason for the failure of L_. anthophora to feed on suckling
mice remains unknown.
Part of the stimulus for inducing feeding seems not to be the
temperature of the host but rather the differential between host tem
perature and ambient temperature.
Feeding: Behavior
Within 2-5 min after initiating feeding,]., anthophora females
fed to repletion (Figure 2-8). The time required for feeding did not
change significantly due to host differences.
However, in a few instances it was noted that the flies failed to
withdraw their mouthparts from the ear of the host. This phenomenon
was observed only with Neotoma and Peromyscus which had been fed upon
repeatedly (Figure 2-9). It may be due to the development of a host
immune response to sandfly salivary products which prevented the fly
from withdrawing its proboscis. This phenomenon has been the subject
of considerable investigation with Ornithodorus coriaceus (Theresa
Haslett and Michel Laviopierre, personal communication, 1981).
Diuresis to reduce excess water and concentrate the blood meal has
been observed in L^. anthophora (Figure 2-10) as the excretion of clear


Figure Page
2-11. 1. anthophora engorged on serum or lymph 40
2-12. Dead female j_. anthophora after peri trophic sac rupture 40
3-1. Habitat of Lutzomyia diablica 49
3-2. L^. diablica feeding on human arm 49
6-1. Electron micrographs of purified Rio Grande virus
at 125.000X 70
7-1. Scanning electron micrographs of eggs of four sandfly
species. (1) Lutzomyia diablica, (2) Lutzomyia
shannoni, (3) Lutzomyia vexator, (4) Lutzomyia
cruciata spp 75
7-2. Scanning electron micrographs of oocyte topography of
five species of sandfly 76
8-1. Chamber for photographing hematophagous insects feeding
on humans and small mammals 80
8-2. Chamber for photographing small insects 80


Figure 2-5. JL. anthophora male <24 hrs old with unrotated genitalia.


SECTION I
TECHNIQUES FOR LABORATORY REARING OF SANDFLIES
(DIPTERA: PSYCHODIDAE)
Introduction and Literature Review
The difficulty of efficiently producing large numbers of sand
flies in the laboratory has hindered studies on their biology and
vector competence for viral and parasitic diseases (Killick-Kendrick
1978). Despite significant contributions by several workers (Chaniotis
1967, 1975, Gemetchu 1976, Killick-Kendrick et al. 1973, Killick-
Kendrick et al. 1977, Ward 1977) several major problems remain. Some
of these include (1) larval mortality due to unknown factors, (2) ex
cessive labor requirements for colony maintenance, and (3) death of
females at oviposition. Use of the techniques described here have con
siderably reduced the first two difficulties and partly solved the
third.
Six of the 600 known phlebotomine species, e.g. £_. argentipes
Annandale & Brunetti, P_. papatasi (Scopoli), J_. longipalpis (Lutz &
Neiva), L_. sanguinaria (Fairchild & Hertig), l. gomezi (Nitzulescu),
and L_. flaviscutellata (Mangabeira), have been reared for 10 genera
tions or more (Killick-Kendrick 1978, Ward 1977). The following
species have been reared by the methods described here: J_. anthophora
(Addis), 15 generations; j_. shannoni (Dyar), 15 generations; J_. vexator
vexator (Coquillett), 7 generations; l. diablica (Hall), 5 generations
-2-


-63-
obtained with l. anthophora could account for maintenance of RGV in
nature by the stabilization mechanism discussed by Tesh and Shroyer
(1980).
At Uvalde, Texas, Pamian (1919) reported an epidemic of a mild
febrile illness (102-104F, 3 days duration) concurrent with large popu
lations of L. diabolica, a man-biting species also known to feed on
woodrats, opossums, cattle, dogs, and horses. Parman (1919) thought
the possible association of the sandfly numbers and the epidemic was
a suspicious coincidence requiring investigation. The symptoms de
scribed by Parman (1919) are consistent with those known to occur after
infection by phieboviruses (Bartonnelli and Tesh 1976, Tesh et al.
1977).
From this study I must conclude that anthophora is probably
important in maintaining Rio Grande virus in the woodrat population but
may not solely account for its transmission. In order to more com
pletely understand the ecology of Rio Grande virus detailed field and
laboratory investigations of the vector potential of other arthropods
must be undertaken.


Table 4-1. Growth of Rio Grande virus in l. anthophora after intrathoracic inoculation.
Day Post-Inoculation
Number/Number
Infected/Sampled
Range of Titers
in Infected Flies*
Mean Titer in
Infected Flies*
0 (immediately after
inoculation)
5/5
<10-4-101-1
10 0" 6
1
5/5
iH
o
tH
o
o
r-H
io1-3
2
5/5
101* 7-io3*4
ig2.5
3
5/5
101* 7-103* 7
IO2*6
4
4/5
102* 9-103*1
103-1
5
4/5
103-4-105*0
104-1
6
-
-
-
7
5/5
104*3-105-7
ig5.0
8
2/2
104-104*3
104-2
9
2/2
104-5
IO4-5
10
1/2
104-8
1048
*Tissue culture infectious dose^g per insect.


-37-
Figure 2-8. Time sequence (20 sec) of L. anthophora feeding on
hamster (Mesocricetus auretus) ear.


Table 2-1. Mean duration (days) of immature stages of l_. anthophora at 90% RH and 4 constant temperature
regimes: 20C, 24C, 28C, and 32C in contrast to the observations of Addis (1945b) made at
28-29C. Larvae reared on the diet of Young et al. (1981).
Temperature
(C)
Larval
Instars
Total
(egg-adult)
Egg
1
2
3
4
Pupa
20
15.60.5
(59)*
12.73.3
(52)
8.52.5
(47)
9.94.7
(42)
22.016.9
(35)
16.813.3
(19)
83.319.3
(19)
24
10.00.6
(174)
6.30.8
(165)
5.00.8
(162)
5.20.3
(160)
11.3H.0
(133)
11.510.4
(130)
49.512.0
(130)
28
8.00.1
(41)
3.90.8
(39)
3.11.1
(39)
8.31.0
(39)
8.712.2
(36)
8.010.8
(36)
36.U2.9
(36)
28-29
(Addis)
10.5
(72)
28.4
8.7
49
32
6.70.6
(104)
3.71.3
(93)
3.80.8
(92)
3.9H.2
(92)
8.812.3
(90)
6.710.8
(88)
33.513.7
(88)
( ) indicates number of individuals surviving each stage.


I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dr. Donald W. Hall
Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.
Dr. Stephen G. ImJ
Associate Professor of Microbiology
and Cell Science
This dissertation was submitted to the Graduate Faculty of the College
of Agriculture and to the Graduate Council, and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
May 1982
Dean,
Dean for Graduate Studies and
Research


BIBLIOGRAPHY
Addis, C.J. 1945(a). Phlebotomus (Dampfomyia) anthophorus, n.sp., and
Phlebotomus diabolicus Hall from Texas (Diptera: Psychodidae).
J. Parsito!. 31(2):119-27.
Addis, C.J. 1945(b). Laboratory rearing and life cycle of Phlebotomus
(Dampfomyia) anthophorus Addis (Diptera: Psychodidae). J.
Parsito!. 31(5):319-21.
Adler, S.,andO. Theodor. 1926. The mouthparts, alimentary tract
and salivary apparatus of the female in Phlebotomus papatasi.
Ann. Trop. Med. Parasitol. 20:109-42.
Aitken, T.H.G., J.P. Woodall, A.H.P. De Andrade, G. Bensabath, and
R.E. Shope. 1975. Pacui virus, Phlebotomine flies, and small
mammals in Brazil--An epidemiological study. Am. J. Trop. Med.
Hyg. 24(2):358-68.
Anderson, D.C., R.G. Buckner, B.L. Glenn, and D.W. MacVean. 1980.
Endemic canine leishmaniasis. Vet. Pathol. 17:94-6.
Barreto, M.P. 1941. Morphologia dos ovos, das larvas e das pupas de
alguns Flebotomus de Sao Paulo. Anais Fac. Med. Univ. S. Paulo
17:357-427.
Bartonelli, P.J.,andR.B. Tesh. 1976. Clinical and serological
responses of volunteers infected with Phlebotomus fever virus
(Sicilian type). Am. J. Trop. Med. Hyg. 25(3):456-62.
Barzilai, R., B.H. Lazarus, and N. Goldblum. 1972. Viscosity-density
gradient purification of foot-and-mouth disease virus. Archiv fur
die gesamte Virusforschung 36:141-46.
Bishop, D.H.L., C.H. Calisher, J. Casals, M.P. Chumakov, S. Ya.
Gaidamovich, C. Hamnoun, D.K. Luov, I.D. Marshall, N. Oker-Blom,
R.F. Petterson, J.S. Porterfield, P.K. Russell, R.E. Shope, and
E.G. Westaway. 1980. Bunyaviridae. Intervirol. 14:125-43.
Calisher, C.H., R.G. McLean, G.C. Smith, D.M. Szmyd, D.J. Muth, and
J.S. Lazuick. 1977. Rio-Grande--A new phlebotomus fever group
virus from South Texas. Am. J. Trop. Med. Hyg. 26(5):997-1002.
-83-


-18-
May and June 1980. Vegetation was characterized by grasses, low grow
ing shrubs, mesquite, and acacia trees common to xeric regions (Figure
2-1). Climatic conditions were quite dry at the time of collection;
however, a rainy season occurs in August and September.
Johnson (1966) described the structure of woodrat nests in detail.
The nests (Figure 2-2) were carefully disassembled and sandflies were
collected with tube aspirators when seen hopping on the sticks. Flies
were also recovered from under boards covering rodent burrows in a
refuse dump. In both sites the soil was powder dry and the flies'
moisture source remains an enigma. Aspirators and field collection
containers have been described in Section I as well as methods for
feeding freshly caught flies on hamsters.
As woodrats attempted to escape from their nests they were captured
as a blood source for flies. Other woodrats and white-footed mice,
Peromyscus leucopus, which also occupy woodrat dens, were trapped in
Sherman traps.
Immature Behavior and Development
The rearing methods for establishing and maintaining the colony
are described in Section I and by Young et al. (1981). Johnson
and Hertig (1961) and Hanson (1968) grouped Neotropical" phlebotomine
larvae into two behavioral groups, i.e., those that burrow into the
larval medium and those that are surface feeders. This behavior
indicates where larvae may be found in nature, i.e., on the soil sur
face or burrowing beneath it. In the laboratory l. anthophora larvae
exhibited no distinct behavioral preference and the degree of larval


-19-
Figure 2-1. Habitat of Lutzomyia anthophora.
Figure 2-2. Nest of Neotoma micropus.


-11-
The bottom and one side of the aquarium were covered with a 1 cm
layer of plaster of Paris. After the bottom has been poured and allowed
to dry it is imperative that it be saturated with water before the side
layer of plaster is poured. This will prevent the formation of un
workable lumps at the junction of the two pours due to immediate des-
sication of the wet plaster by the dry layer. After the bottom and one
side have been poured the bottom should be rewet and the upper corners
filled in to a maximum depth of 2 cm. This allows for easy viewing of
flies and easy recovery of flies with an aspirator.
Front panels for the cages are constructed of 64 mm (1/4") Plexi-
glas Screens of 18 mesh/cm of "Saran" (Chickopee Co., Cornelia, GA)
vinylidene polymer plastic are installed on the front panel. Experience
has shown that this is necessary because otherwise, excessive conden
sation in the chamber will form when animals are left in for sandfly
feeding. The Saran screen is attached with epoxy cement. Care must
be exercised that epoxy components are not out of date and are well
mixed; otherwise the glue will remain sticky and trap the flies. Other
glues tried, i.e., contact cement, Elmer's glue silicone, and super-

glue do not adhere well to the Plexiglas. The minimum screen areas
2
are 78, 130, and 214 cm respectively.
A 50 cm sleeve of 15.3 cm (6") surgical stockinet (Johnson &
Johnson) is attached to the front panel by compression between the panel
and a Plexiglas frame (2.5 cm wide). This is secured with 8 (10/24 x
1") brass screws with flat washers and wingnuts. The brass screws and
flat washers are glued inside the front panel with epoxy glue in order
to facilitate changing of the sleeve which should be secured with tape
while the frame is being installed. The completed front panel is


-85-
Hall, D.G. 1936. Phlebotomus (Brumptomyia) diabolicus, a new species
of biting gnat from Texas (Diptera: Psychodidae). Proc. Entomol.
Soc. Wash. 38(2):27-9.
Hanson, W.J. 1968. The immature stages of the subfamily Phlebotominae
in Panama (Diptera: Psychodidae). Ph.D. Thesis, Univ. Kansas.
104 p. Univ. Microfilms, Ann Arbor, Mich. (Diss. Abstr. 68-171,390)
Hawkes, R.A. 1979. General principles underlying laboratory diagnosis
of viral infection. Lennette, E.H., and N.J. Schmidt, eds.
Diagnostic Procedures for Viral, Rickettsial, and Chlamydial In
fections. 5th ed. American Public Health Association. Washing
ton, D.C.
Hayes, R.O., D.B. Francy, J.S. Lazuick, G.C. Smith, and E.P.J. Gibbs.
1977. Role of the cliff swallow bug Oeciacus vi cari us in the
natural cycle of a Western Equine Encephalitis-related Alpha-
virus. J. Med. Entomol. 14(3):257-62.
Jennings, M., and J. Boorman. 1980. The susceptibility of Lutzomyia
longipalpis (Lutz and Neival), Diptera, Psychodidae, to artificial
infection with three viruses of the Phlebotomus fever group. Ann.
Trop. Med. Hyg. 74(4):455-62.
Johnson, D.W. 1966. A Population of Woodrats (Neotoma micropus) in
Southern Texas.. Gerald G. Raun, ed. Bulletin of the Texas
Memorial Museum. Bulletin 16. Welder Wildlife Foundation Con
tribution, Series B, No. 4. Texas Memorial Museum, 24th & Trinity.
Austin, Texas.
Johnson, P.T., and M. Hertig. 1961. The rearing of Phlebotomus sand
flies (Diptera: Psychodidae). II. Development and behavior of
Panamanian sandflies in laboratory culture. Ann. Entomol. Soc.
of Amer. 54(6):764-76.
Justines, G., and O.E. Sousa. 1977. Survival of Arboviruses in try
panosome-infected Triatomines. Am. J. Trop. Med. Hyg. 26(2):
326-28.
Kaariainen, L., K. Simons, and C.-H. von Bonsdorff. 1969. Studies in
subviral components of Semliki Forest Virus. Ann. Med. exp. Fenn.
47:235-48.
Karabatsos, N., ed. 1978. Supplement to the International Catalogue
of Arboviruses including certain other viruses of vertebrates.
Am. J. Trop. Med. Hyg. 27(2):415-17.
Ki11ick-Kendrick, R. 1978. Recent advances and outstanding problems
in the biology of phlebotomine sandflies. Acta Tropica 35:297-313.
Killick-Kendrick, R. 1979. Biology of Leishmania in phlebotomine
sandflies. Lumsden, W.H.R., and D.A. Evans, eds. Biology of the
Kinetoplastida. Vol. 2. Academic Press. London/New York.


-49-
Figure 3-1. Habitat of Lutzomyia diablica.
Figure 3-2. J_. diablica feeding on a human arm.


-74-
1. Eggs were placed on a filter paper disc in a 1 cm deep plastic
container cut from a plastic film cannister.
2. The plastic container was floated in a 50 ml Tri-pour poly
styrene beaker containing 5 ml aqueous 1% OsO^
3. The paper lid was installed and the entire container was sealed
in Parafilm and held in an exhaust hood at room temperature
for 5 days.
4. After 5 days exposure to osmium vapor the inner container was
transferred to a covered petri dish for 24 hr to allow slow
drying of the eggs.
5. Eggs were attached to an SEM stub using either double-sided
tape or 0.1% aqueous hydrobromide polylysine (Polysciences,
Inc., Warrington, PA 18976), sputter-coated with approximately
300 A of gold in an Eiko Engineering IB-2 Ion Coater, and
examined in a Hitachi S-450 scanning electron microscope (SEM)
at 20 KV.
Eggs were measured in microns at lOOx with a compound microscope
and an ocular micrometer. Intact fresh eggs or recently hatched eggs
were acceptable whereas old eggs or infertile eggs usually collapsed
making accurate measurement difficult. Eggs to be measured were placed
on a microscope slide in Histocon. In each sample the eggs were pro
duced by 5-10 females.
Results
The SEM micrographs of the sandfly eggs are shown in Figures 7-1
and 7-2. Descriptions of eggs of each species are as follows. Measurements


-3-
].. cruciata spp, 23 generations; L_. cayennensis (Floch & Abonnenc),
5 generations. In addition, 3 African phlebotomine species were
reared to the 4th generation using these methods (D. Young, personal
communication).
General Techniques
Figure 1-1 represents the generalized rearing method for sandflies.
An explanation of each step is as follows: Step 1. The plaster of
Paris in a rearing cage is saturated with H^O with no free water re
maining; Step 2. An engorged female is gently "herded" into the vial.
A drop of 30% honey solution or other sugar source is then placed on
the screen top; Step 3. After 3+ days most females oviposit on the
plaster bottom. If the female survived oviposition she is released
back into the feeding cage. Screen lids are replaced with solid tops
that have small punctures to allow for gas exchange but limit dessi-
cation; Step 4. Since eggs held at 26C usually hatch 6-14 days after
oviposition, a small amount of larval diet is placed in the vial 4-5
days after the eggs are laid; Step 5. Larvae should be checked weekly
and moist medium added as required; Step 6. Adults are released into
the feeding cage daily by placing lidless vials containing pupae in
the feeding cage. Adults soon begin mating and feeding on sugar from
apple slices provided; Step 7. An anesthetized or restrained vertebrate
host is placed inside the cage after a prefeeding period that varies
in time according to species.


-6-
Figure 1-2. J_. anthophora feeding on an apple slice.


-41-
Refeeding
Investigations on the vector capability of sandflies have been
hampered by the failure of females to survive oviposition. Conse
quently, demonstration of transmission of Leishmania and Phleboviruses
by bite has been difficult to establish (Kil1ick-Kendrick 1979). Kil-
1ick-Kendrick (1979) and Johnson and Hertig (1961) discussed those
species which feed more than once in the laboratory.
In the Fg and F^g generations of L. anthophora maintained in
laboratory culture, the individuals that laid eggs within a 24 hr period
were released into a feeding cage and offered an anesthetized host for
a 30-60 min period daily. Of the flies in those respective generations
19.1% and 16.1% took a second blood meal. Four individuals fed 3x in
the Fg generation. These results are consistent with the observations
of Schmidt and Schmidt (1965) on P_. papatasi.
No experiments were conducted to discover an optimum oviposition
site. If this were determined perhaps a much higher percentage of
blood fed females would oviposit, survive oviposition, refeed, and
repeat the gonotrophic cycle. Most flies that died prior to oviposi
tion retained eggs in the abdomen. It is unlikely that such high
mortality occurs at oviposition in wild populations.
Peri trophic Sac Rupture
The term "membrane" when used to describe the lattice-like sac
which surrounds the blood meal in hematophagous insects is a biological
misnomer since it is not a trilaminate phopholipi d/protein membrand.
I suggest adoption of the term "peritrophic sac." Romoser and Rothman


-46-
Table 2-8. Comparison of longevity (days) of L.. anthophore mal es and females
fed on either distilled water or 30% honey solution at
24C, 90% RH.
30% Honey Solution
Distilled
h2o
Sex
X
S
X
S
Female
10.4
(31)'
2.9
k
7.7
(18)
1.6
Male
10.1
(32)
3.1
7.1
(16)
1.5
Total
10.2
(63)
3.0
7.4
(34)
1.6

( ) indicates
number of
individuals tested.


-71-
3
a confluent monolayer of Vero cells in a 75 cm tissue culture flask
which was then overlaid with 50 ml minimum essential medium. After
6 days incubation at 37C nearly 100% of the cells were destroyed.


-77-
given are the range, mean, and standard deviation for egg length and
width for each species.
Lutzomyia shannoni (Dyar, 1929), Florida specimens
Figure 7-1(2), 7-2(6)
Size: N = 102, L: 290-340 (330 10), W: 70-110 (90 10)
Exochorion: High, narrow longitudinal ridges connected by prominent
perpendicular ridges forming 4 and 5 sided polygons which are fre
quently rectangular.
Lutzomyia diablica (Young and Perkins 1982), Uvalde Co., Texas
Figure 7-1(1), 7-2(5)
Size: N = 47, L: 340-370 (350 10), W: 90-110 (100 10)
Exochorion: Surface topography is characterized by a series of dis
continuous parallel longitudinal ridges that are not laterally connected.
Lutzomyia vexator (Coquillett 1907), Levy Co., Florida
Figure 7-1(3), 7-2(7)
Size: N = 193, L: 330-390 (380 10), W: 80-110 (100 10)
Exochorion: Surface topography consists of delicate parallel longi
tudinal ridges with regular perpendicular connections that form polygons
which are nearly square. There are also occasional oblong cells.
Lutzomyia anthophora (Addis 1945), Cameron Co., Texas
Figure 7-2(9)
Size: N = 100, L: 330-370 (340 10), W: 80-100 (80 10)
Exochorion: Reticulation consists of weak parallel longitudinal ridges
with slight perpendicular connections at irregular intervals.


SECTION IV
TRANSOVAR IAN TRANSMISSION OF RIO GRANDE VIRUS BY
Lutzomyia anthophora (ADDIS) (DIPTERA: PSYCHODIDAE)
Introduction and Literature Review
The genus Phlebovirus of the family Bunyaviridae (Bishop et al.
1980) includes more than 40 viruses (R.B. Tesh, personal communication)
distributed over 5 continents (Berge 1975, Karabatsos 1978). Calisher
et al. (1977) described Rio Grande virus from isolates made from wood-
rats, Neotoma micropus, collected near Brownsville, Texas, in 1973-
1974. J_. anthophora was suspected of transmitting this virus because
of its intimate association with woodrats (Young 1972), the high anti
body prevalence (46.3%) of the woodrats (Calisher et al. 1977), and the
fact that sandflies transmit other related phleboviruses. Transovarian
transmission of phleboviruses by sandflies has been suggested as a
mechanism of viral survival (Teshand Chaniotis 1975). In the present
study experiments were undertaken to demonstrate transmission of Rio
Grande virus by j_. anthophora transovarially and by bite.
Materials and Methods
Sandflies
The J_. anthophora used in these experiments were from the F-j
generation of a closed colony started from stock collected E and NE
-54-


-76-
Figure 7-2. Scanning electron micrographs of oocyte topography of
five species of sandfly. (5) Lutzomyia diablica, (6)
Lutzomyia shannon!, (7) Lutzomyia vexator, (8) Lutzomyia
cruciata spp., (9) Lutzomyia anthophora. 7,000x.


-81-
Belleville, NJ) that is 7.5 x 2.5 x 1.5 cm. It was covered on the
bottom and ends with a 1 cm layer of white polyethylene foam (Ward's
Scientific Co., Rochester, NY). A Kodak Neutral Test card 90% re
flectance on the white side and 18% on the gray side was used for a
background behind the cuvette chamber.
A third type of photographic chamber used occasionally was the
rectangular adult feeding cage.
All of the sandfly photographs presented in this manuscript were
photographed with a 200 mm Medical Nikkor lens at 3x magnification, F45.
This lens was used because it has a built-in ring flash and provides
8 cm working distance at 3x magnification. The camera used was a Nikon
F2 Photomic with type "C" focusing screen and cable release. Because
the lens is of a fixed focal length it was necessary to mount the
camera on a SIik 2-axis focusing rail on a tripod in order to achieve
reproducible results.
Fujichrome film, ASA 100, was used for all the photographs. Black
and white prints were all made from color slides using Ilford XP400
ultra fine grain film for an internegative.


-15-
which should be done in a well ventilated area to avoid noxious fumes.
The edges of the hole are then filed smooth and pieces of latex surgi
cal glove are glued to both sides with contact cement. If prepowdered
surgical gloves are used they must be washed in a 70% ethanol solution
to remove the powder to insure adhesion. Perpendicular cuts are made
in the respective latex pieces producing a fly-proof opening for the
insertion of an aspirator. Screen lids should be used on the containers
when used for collecting vials. These containers can also be used for
feeding flies. Screen lids are prepared by cutting a 4 cm hole in the
plastic top and attaching the desired mesh screen with contact cement.
Individual Oviposition and Rearing Containers
These containers are made by pouring 1 cm of plaster of Paris in
the bottom of a 7 dram plastic snap cap vial (Fisher Scientific Co.,
Pittsburgh, PA). When used for rearing containers the plastic tops
are punctured to facilitate limited gas exchange. When used as feeding
containers the lids are cut out with a #12 cork borer (1.5 cm hole)
and covered with the desired mesh screen that is secured with contact
cement. When the vials are inverted the plaster occasionally slides
down crushing the insects. This can be prevented by pushing a hot pin
through the plastic into'the plaster then cutting off the excess.
Aspirators
Aspirators for collection and transfer of adults are constructed
of thick wall latex tubing (10 mm ID x 15 mm 0D x 60 cm) and thickwall
Pyre/glass tubing (12 mm ID x 15 mm 0D x 30 cm) (Figure 1-5). The


-24-
Time of Eclosin
It is a general observation that the males of many insect species
begin eclosin before the females in order to be reproducin'vely mature
when the females emerge. I noted this to be the case with l. anthophora
(Table 2-3), because mean development time from egg-adult was about 2
days less for males than females at temperatures above 20C. This is
noteworthy because males are not reproductively competent until 24 hrs
post eclosin.
In order to demonstrate this phenomenon the sex and time of eclo
sin for all individuals from a cohort of the Fg generation were recorded.
The eclosin distribution of males and females is presented in Figure 2-4
and demonstrates the veracity of this observation. Sex ratios were 1:1.
Mating
Male genitalia rotate (Figure 2-5) about 12-24 hrs post eclosin
after which they were observed mating. Females were seen mating within
hours after eclosin and before, after, and during feeding. Mating
frequency was not determined for either sex although males do mate more
than once per lifetime. Copulation occurred regardless of nutritional
state or photoperiod.
1. anthophora males demonstrated the "characteristic epigamic
pattern" described by Chaniotis (1967). Mating usually lasted 2-5 min.
Based on the criterion of egg fertility more than 85% of the females
that laid eggs had successfully mated. In two generations studied the
percentage of females laying infertile eggs was 13.7% and 16.7% in the
Fg and the F^5 generations, respectively.


-8-
container of flies against an animal's ear or nose allowing the flies
to feed through the screen top. This method is particularly useful for
feeding flies on leishmanial lesions. A mesh size of 18/cm is required
for small species such as l. anthophora and l. diabolica. A larger
mesh size of 10/cm is adequate to contain larger species such as
l. shannoni.
The second method is to place an anesthetized or restrained animal
in the feeding cage. Anesthesia dosage rates are given in Table 1-1.
Anesthesia was administered with a 1 ml Tuberculin syringe and a 26 or
27 gauge needle. Animals in poor condition require less anesthesia.
An insufficient dose will sometimes produce hyperactive behavior.
Lid Cleaning
Screen tops that have been used for sugar feeding are cleaned by
soaking in 5% Chlorox solution for 30 min, rinsing 2x in tap water for
30 min, and air dried. Tops can be reused many times until screening
material breaks or glue becomes brittle and non-adhesive. Use of a
more concentrated Chlorox solution or a longer soaking time results
in rapid deterioration of screen material and glue greatly reducing
the number of times lids can be reused.
Adult Feeding Cages
The most successful adult feeding cages developed were constructed
from 4 (26 x 20.5 x 16.5 cm), 6 (31.0 x 20.5 x 16.5 cm), and 12
(36 x 25.5 x 21.5 cm) liter aquariums (Figure 1-3).


-5-
Larval Rearing
Later instar larvae are more tolerant of moisture variation than
earlier instar larvae. When larval medium (Young et al. 1981) is added
to the rearing vials it must be slightly moistened. Larvae can be
reared under conditions of >80% RH but 90-95% is preferable. Even at
this high humidity secondary fungal growth is uncommon, presumably be
cause the first Rhizopus sp. bloom either exhausts an essential nutrient
or produces a fungal growth inhibitor. It is a primary colonist in
fungal succession and reduces proteins to amino acids and carbohydrates
to simple sugars. After the medium has completely dried, it is re
moistened. Even then, there is little fungal growth, the hyphae are
not abundant enough to entangle the caudal setae of the larvae.
Mites frequently seen in larval vials have not been observed
attacking healthy larvae but they will feed on weak or dead larvae and
adults. Boiling water poured into vials before reuse will kill any
mites present. Autoclaving larval medium infested by mites for 5 min
at 15 psi will kill mites without apparent damage to the medium.
Sugar Feeding
Ready (1979) provided evidence that sugar feeding is important for
sandfly egg production. Adults were provided sugar ad libidum throughout
their lifetime by two methods.
Adult males and non-bloodfed females were provided sugar from thin
apple slices (<3 mm) leaned against the sides of the feeding cages
(Figure 1-2). Thicker apple slices tend to mold more rapidly than


-38-
Figure 2-9. L. anthophora with mouthparts stuck in the ear of
TPeromyscus leucopus), white-footed mouse.
Figure 2-10. L_. anthophora excreting clear fluid droplets while
feeding.


-39-
fluid droplets from the anus while feeding. Chaniotis (1967), Gemetchu
(1976), and others have also observed this phenomenon in Phlebotomines.
Feeding: Lymph
Regardless of the host, 3.2-5.6% of the sandflies in a given cohort
engorged with a clear fluid that is presumably serum or lymph (Figure
2-11). This behavior indicates that some individuals may have the
ability to filter erythrocytes from the blood while feeding or else
they may feed by chance from lymphatic capillaries. The latter explana
tion is feasible since sandflies are telmophages (Lewis 1975) and lymphatic
capillaries are numerous in the dermis. Ready (1978) noted that _L.
longipalpis (Lutz & Neiva) was a non-selective feeder and would feed
to engorgement on isotonic saline and whole blood with equal avidity.
This may also be true of]., anthophora. Of the 20 serum fed individuals
studied in 3 generations 93% did not lay eggs and the maximum number of
eggs laid per female was 12 of those that did. In contrast the mean
egg production for bloodfed females from the same generations was 30.5.
The erythrocyte blood fraction apparently contains nutrients essential
for egg production. Ready (1979) found that the concentration of pro
tein ingested had a significant direct relationship to the number of
oocytes produced and that the red cell fraction was more important than
plasma for egg production. The production of 12 oocytes or less by
lymph fed J_. anthophora contrasts with the conclusion of Adler and
Theodor (1926) that plasma alone was essential for _P. papatasi to pro
duce eggs.


-40-
Figure 2-11. J_. anthophora engorged on serum or lymph.
Figure 2-12. Dead female _L. anthophora after peritrophic sac
rupture.


Figure 1-1. Schematic diagram of laboratory rearing techniques for phlebotomine sandflies.


LIST OF FIGURES
Figure Page
1-1. Schematic diagram of laboratory rearing techniques for
phlebotomine sandflies 4
1-2. J_. anthophora feeding on an apple slice 6
1-3. Sandfly feeding cage--A modified aquarium with plaster
of Paris bottom and back 10
1-4. Cylindrical adult feeding cage 13
1-5. Field collection apparatus, feeding and rearing con
tainers for phlebotomine sandflies 14
2-1. Habitat of Lutzomyia anthophora 19
2-2. Nest of Neotoma micropus 19
2-3. Multiwell plate with lid used for rearing individual
larvae 21
2-4. Eclosin distribution of 127 J_. anthophora males and
females from F3 generation in a laboratory colony at
24 C, 90% RH 25
2-5. L.. anthophora male <24 hrs old with unrotated
genitalia 26
2-6. 1. anthophora--Temporal age distribution at feeding and
death of unfed females at 28C, 90% RH 28
2-7. j_. anthophora males and females engorged on 30% honey
solution dyed with red, blue, and green food dye 30
2-8. Time sequence (20 sec) of l. anthophora feeding on
hamster (Mesocricetus auretus) ear. . 37
2-9. 1. anthophora with mouthparts stuck in the ear of
Peromysus leucopus (white-footed mouse) 38
2-10. J_. anthophora excreting clear fluid droplets while
feeding 38
xi


-12-
attached to the cage with a thick layer of silicone glue that can be
easily cut away for repairs. It is essential to fill all small crevices
with plaster of Paris or silicone to prevent adult sandflies from hiding
there and being difficult to recover.
A cylindrical adult feeding cage (Figure 1-4) was constructed from
a cylindrical (23 x 13 cm) glass fixture cover (Appleton Co. V-51).
Four centimeters of plaster of Paris were poured in the end and 1.5 cm
(tapered to the front) were poured on a side of cylinder by the method
described. The frame was constructed of 3 (18 cm x 18 cm x 64 mm)
Plexiglas plates and 18 cm (10/24) threaded steel rod. Relative posi
tions of the plates is maintained by placing nuts and washers on both
sides of the sheets which are attached to the glass by a bead of silicone
glue. A 50 cm stockinet sleeve is secured to the front by compression
between 2 plates as with the rectangular feeding chamber. The advan
tages of the cage include small size and ease of manufacture. Dis
advantages are difficulty seeing through the glass, condensation due
to animal respiration, and difficulty in manipulating vials inside the
chamber.
Field Collection, Feeding Containers
The 120ml specimen containers (Pharmaseal Laboratories, Glendale,
CA 91201) are modified for use as field collection and feeding con
tainers (Figure 1-5).
Field collection containers are constructed as follows. Two
centimeters of plaster of Paris are poured in the bottom of the
containers; then a 2 cm entry port is cut in the container side
by heating a #15 cork borer then pushing it through the plastic


-29-
solution mixed with either red, blue, or green food dye (C.F. Sauer Co.,
Richmond, VA) was placed on the screen lid. After a sandfly ingested
the solution the dye was clearly discernible in the distended crop with
or without the aid of a microscope (Figure 2-7). Yellow dye was not
used because it could not be seen. When an individual digested the
sugar solution a small colored droplet was excreted and no color re
mained in the abdomen. Flies would not refeed until the previous meal
had been completely digested.
Results for sugar feeding experiments conducted at 20C and 28C
are shown in Table 2-4. The reduction in number of flies feeding more
than lx at 28C indicates the release of individuals back into the
breeding colony rather than mortality. Of the flies offered honey
solution at 28C, 93.2% (68/73) fed within 24 hrs after eclosin.
Of 23 flies held until death at 28C, 100% fed lx, 30.4% fed 2x, 30.4%
fed 3x, 34.5% fed 4x, and 4% fed 5x.
The number of sugar fed adults held at 20C declined rapidly due
to poor survival. Digestion of the first and second sugar meal at 20C
requires more time than at 28C.
Results of a second experiment to determine the effect of sugar
feeding on percentage of females feeding on blood, fecundity, preovi-
position period, mating (fertility of eggs), and mortality factors are
presented in Table 2-5. Sugar feeding enhanced productivity for all
parameters measured.
The source of sugars for j_. anthophora in nature is unknown. Many wood-
rat nests are located around clumps of prickly-pear cactus (Opuntia lind-
heimeri), a succulent that woodrats feed on in their nests. Sandflies
may obtain sugar from partly eaten cactus in the woodrat nest. To test


SECTION VIII
PHOTOGRAPHIC TECHNIQUES
Quality photographs of living sandflies, Culi coi des, and other
small insects have been notably lacking from the literature due to
the difficulty of producing them. Part of the difficulty involved in
photographing these insects is containing them. Two types of spe
cialized containers were developed in order to photograph sandflies.
The first container (Figure 8-1) was constructed from a 40 liter
aquarium. Two sides were replaced with 2 mm Plexiglas. Three access
ports (19 x 19 cm) were cut in the sides and fitted with Plexiglas
compression frames (2 cm wide x 64 mm thick). Sleeves 50 cm in length
made of 15 cm surgical stockinet (Johnson & Johnson) were secured
around the ports by the compression frames which were attached with
8 brass screws (10/24 x 1") and wing nuts. The screws and flat washers
wereglued in place with epoxy glue for ease of attaching the sleeves.
Three sleeves are required, 1 for the camera, 1 for manipulating
specimens, and 1 for the host arm. A Plexiglas insertwas attached to
the top frame of the aquarium with silicone glue thus making a re
movable top. The back and bottom of the aquarium is covered with a
1 cm layer of plaster of Paris to provide a light reflective back
ground.
The second type of photographic chamber (Figure 8-2) was con
structed from a spectrophotometric cuvette (Wallace & Tiernan, Co.,
-79-


Pae
IIICOLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia diabolica(HALL) (DIPTERA: PSYCHODIDAE) 47
Introduction and Literature Review 47
Field Collection 47
Feeding 50
Mating 51
Egg Hatch and Fertility 51
IVTRANSOVARIAN TRANSMISSION OF RIO GRANDE VIRUS BY
Lutzomyia anthophora(ADDIS) (DIPTERA: PSYCHODIDAE) 54
Introduction and Literature Review 54
Materials and Methods 54
Results 57
Discussion 61
V RIO GRANDE VIRUS AND Triatoma qerstaeckeri (STAL)
(HEMIPTERA: REDUVIIDAE). 64
Introduction and Literature Review 64
Materials and Methods 65
Results and Conclusions 65
VI PURIFICATION OF RIO GRANDE VIRUS 67
Introduction 67
Materials and Methods 67
Results 69
Discussion 69
VII A COMPARISON OF OOCYTE TOPOGRAPHY OF FIVE PHLEBOTOMINE
SANDFLIES (Lutzomyia) WITH THE SCANNING ELECTRON MICRO
SCOPE (DIPTERAT P5YCH0DI DAE) 72
Introduction 72
Materials and Methods 72
Results 74
Discussion 78
VIII PHOTOGRAPHIC TECHNIQUES 79
IX SUMMARY 82
BIBLIOGRAPHY 83
BIOGRAPHICAL SKETCH 90
vi 11


SECTION III
COLONIZATION AND LABORATORY BIOLOGY OF THE SANDFLY,
Lutzomyia diablica (HALL) (DIPTERA: PSYCHODIDAE)
Introduction and Literature Review
Lutzomyia diablica (Hall 1936) has long been recognized as a pest
in South Central Texas (Parman 1919, Lindquist 1936) where it bites
humans in and near human dwellings. The status of the species was ques
tioned by Disney (1968) but has been recently resolved by Young and
Perkins (1982). Lindquist (1936) studied the life cycle of the species,
described the immature stages but did not establish a laboratory colony.
Addis (1945a) made an unsuccessful attempt at colonization. Parman
(1919) described the bite on humans in detail and suggested that J_.
diablica may be a vector of a transient febrile human illness.
Several cases of autochthonous leishmaniasis have been recorded
from Texas (Shaw et al. 1976, Simpson et al. 1968, Stewart and Pilcher
1945, and Anderson et al. 1980). Since l. diabolica is the only known
man-biting sandfly in the region it is highly suspect as a potential
vector.
Field Collection
Although J_. diablica was first taken from Uvalde, Texas, it is
widely distributed in Northern Mexico (Najera 1971) and Texas (P.V.
Perkins and D.G. Young, personal communication, 1982). For establishment
-47-


-78-
Lutzomyia cruciata spp. (Young and Perkins 1982), Alachua Co., Florida
Figure 7-1(4), 7-2(8)
Size: N = 61, L: 320-370 (340 10), W: 80-120 (100 10)
Exochorion: Wide, flat, parallel longitudinal ridges with occasional
weaker connecting ridges which are not usually perpendicular to the
longitudinal ridges.
Piscussion
Several techniques were tried for preserving the eggs to prevent
collapse under vacuum in the SEM column. The method used here yielded
the best results when fertile eggs were used.
A "standard" EM fixation procedure using 1% 0s0^ as a fixative
followed by 5% aqueous acrolein, dehydration in dimethoxypropane and
w
acetone, then critical point drying with Freon as a transition solvent
proved unsuccessful because most specimens collapsed in the SEM.
Lyypholization and critical point drying of eggs without fixation
were also unsuccessful.
A technique which was not used but one which may be promising is
freeze drying.
The size variation of eggs laid by individual females was deter
mined by measuring 10 eggs from each of 10 JL. vexator females. The
variation in egg length and width between females ranged from 10-50
microns and from 10-30 microns, respectively. As a result of broad
intraspecific variation it is not possible to separate the eggs of
different sandfly species by size. Therefore the surface sculpturing
is the only characteristic of the egg that can be used for species
determination.


-34-
This was done to verify that the imagos were ready to feed. In each
instance flies that refused the first host fed upon the second host.
It is apparent from the various species of mammals fed on that
JL. anthophora is a more opportunistic feeder than was previously
recognized.
Feeding: Temperature Preference
On each of the host animals used several body regions were rela
tively hairless, i.e., the ears, nose, tail, and feet. To determine if
relative temperature of the body regions influenced sandfly feeding site
preference the skin temperature of these areas was measured with a
BAT-4 Laboratory Thermometer (Bailey Instrument Co.) and a thermistor
(Table 2-6). The instrument was calibrated to human body temperature
of 37.2-37.4C.
More than 95% of the sandflies fed on the ears where the skin
temperature range was from 27.7C to 37.8C. Although the temperatures
of the tail, foot, and nose were within this range little feeding oc
curred. Dermal temperature does not seem to be a determining factor
in fly feeding site preference.
Chaniotis (1975) reported that suckling mice were the least
satisfactory source of blood meals for J_. trapidoi (Fairchild and
Hertig), a species which feeds on a wide variety of mammalian hosts.
Gemetchu (1976) reported that _P. longipes (Parrot and Martin) which
normally feeds on humans would not feed on suckling mice. Although
L. anthophora feeds readily on adults of all rodent species offered,
it would not feed on suckling mice (SM). When the SM were placed in
the feeding cage their body temperature rapidly declined. In an effort


-16-
latex tubing and glass are attached by a piece of hard plastic tubing
(9 mm OD x 5 cm) covered with nylon organdy cloth on one end and
secured with contact cement (Roberts Consolidated Industries, City of
Industry, CA). The screened end is inserted into the glass tubing where it is
held by friction and the latex tubing is pushed over the other end.
The latex/glass junction is securely taped so that the end of the plastic
tube is visible in the glass tubing.
Thickwall latex tubing is used for flexibility and to prevent
kinks from occluding the passageway. Pyrex glass is used instead of
plastic because plastic scratches easily making identification of speci
mens difficult. The inside diameter of any aspirator used for phlebo-
tomine sandflies should be at least 10 mm because smaller diameters
at the same suction pressure result in much higher intake velocities
that cause damage to the flies.


Copyright 1982
by
Richard G.
Endris


-58-
because the sandflies refused to feed on the mice (although they sub
sequently fed on a hamster).
The following mortality was observed after 120 female sandflies
were inoculated with RGV: Day 1(18/120, Day 2 (26/102), and Day 3
(7/94). This mortality is attributable to the inoculation procedure.
Twenty-two (22) infected sandflies fed on a hamster and 6 of the
22 (27.3%) survived to oviposit. Based on observations on other genera
tions of l. anthophora the expected survival would have been 61.5%. The
inoculation procedure may have caused this reduction. Mean egg pro
duction for the 6 flies that oviposited was 28.3 (range 12-47). The
mean number of eggs produced does not seem affected by inoculation
procedures.
From 170 eggs laid by infected females, 62 adult progeny were
produced with a sex ratio of 1:1. The transovarian transmission (TOT)
rate was 54.8% (34 of 62 adults were infected with RGV). The infec
tion rates for males and females was similar, 15/30 (50.0%) and 19/32
(59.3%), respectively. Filial infection rates for the F^ progeny were
not calculated because of insufficient numbers. Each of the 6 in
fected parents produced 1 or more infected progeny. One adult female
survived the F^ generation and laid eggs but was not infected.
The suckling mice inoculated intracerebrally (IC) or subcutaneously
(SQ) with RGV died at 4 and 3 days, respectively. Daily blood samples
from a mouse in each group were all positive for Rio Grande virus
titrated at 1:10 dilution. The rapid kill rate for the group infected
SQ was suspicious because of possible mouse colony contamination with
mouse hepatitis virus and the surprising fact that they died before
those mice inoculated IC. The experiment was repeated with the same
results.


-70-
Figure 6-1. Electron micrographs of purified Rio Grande virus at
125,000x.


Tabl e Page
4-1. Growth of Rio Grande virus in J_. anthophora after
intrathoracic inoculation 59
4-2. Presence of Rio Grande virus in (1) Neotoma micropus
and (2) Peromyscus leucopus bled daily for 7 days
after subcutaneous inoculation 60
7-1. Classification of 41 species of Neotropical phlebotomine
sandfly eggs based on oocyte topographic patterns 73
x


-68-
2
1976, Clewley et al. 1977). Four 150 cm tissue culture flasks of Vero
cells in a confluent monolayer were inoculated with 6 ml mouse brain
suspension and allowed to adsorb for 1 hr at 37C. Cultures were then
overlaid with 50 ml minimum essential medium (MEM) and incubated at
37C. After 24 hrs the supernatant was poured off, 75 ml MEM was over
laid, and the cells were incubated for an additional 6 days at 37C
after which >90% of the cells were destroyed. Supernatants were col
lected, frozen to -70C, thawed to 37C, then clarified by low-speed
centrifugation at 4C for 30 min at 8,000 g in a Sorvall RCB-2 centri
fuge to remove cell debris.
Virus was recovered from the clarified supernatant by precipita
tion in a 7% polyethylene glycol/0.4 M NaCl solution stirred for 4 hr
at 4C followed by centrifugation at 10,000 g for 20 min. The pellet
was resuspended in 4 ml TSE buffer (0.01 M Tris hydrochloride buffer,
pH 7.5, containing 0.1 M NaCl and 0.002 M EDTA) and loaded over a com
bination equilibrium: viscosity gradient of potassium tartrate (McCrea
et al. 1961) and glycerol (KT-GLY).
Two 10 ml KT-GLY gradients were made with a Bethesda Research
Products Gradient Former and an LKB peristaltic pump. Fourteen
milliliters of 50% (w/w) potassium tartrate in TSE buffer was loaded
into the inside chamber and 16 ml of 30% (w/w) glycerol was loaded into
the outside chamber (Obijeski et al. 1974, Barzilai et al. 1972).
Virus suspensions loaded onto KT-GLY gradients were centrifuged
in an SW 41 rotor at 4C for 8 hr at 40,000 g. Three nearly inseparable
visible bands were produced. The virus fraction was collected at 254 nm
(RNA absorbance peak) using an ISC0 gradient column fractionator and
flow densitometer .


FRONTISPIECE
Lutzomyia anthophora feeding on the ear of its native host,
the woodrat, Neotoma micropus.


-65-
T. gerstaeckeri infected with T. cruzi in Neotoma nests. Pippin (1970)
noted 30.1% of the bugs found in the nests were infected.
Materials and Methods
Virus
Rio Grande virus (strain TMB4-719) was kindly supplied by Dr.
Robert Tesh, Yale Arbovirus Research Unit. Additional quantities of
virus were prepared by passage through suckling mouse brain.
Triatomes
A colony of T. gerstaeckeri was started from specimens collected
near Brownsville, Texas, in June, 1980, and augmented with specimens
collected near Lake Medina, San Antonio, Texas, in July, 1981. First
and second instar nymphs from the colony were used in the transmission
experiments.
One hundred forty (140) 2-3 week old first instar nymphs were fed
on 3 suckling mice that had been given Rio Grande virus by intracerebral
inoculation 4 days earlier. Three other mice from the same litter died
on Day 5 post inoculation.
Eight, 16, and 24 days after the initial feeding 80 first instar
nymphs that had fed on viremic mice fed on 6 unexposed suckling mice.
After 1 week the suckling mice showed no apparent signs of viral infection.
Results and Conclusions
The failure to infect 6 suckling mice fed on by 80 nymphs that had
fed on viremic mice 8, 16, or 24 days earlier indicates that T. gerstaeckeri


-30-
Figure 2-7. L. anthophora males and females engorged on 30% honey
solution dyed with red, blue, and green food dye.