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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STUDIES OF Lutzomyia anthophora (ADDIS) (DIPTERA:






Copyright 1982


Richard G. Endris

A man's reach should exceed his grasp.

--Robert Browning


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


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


A grant from the Steffen Brown Foundation provided the opportunity

to study at Yale University in 1981.



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

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

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

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

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


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

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

Lutzomyia diabolica(HALL)(DIPTERA: PSYCHODIDAE) ......... 47

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

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


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

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



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


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:


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.



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.



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


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


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.







Schematic diagram of laboratory rearing techniques for phlebotomine sandflies.


Figure 1-1.


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


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


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
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


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


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


Figure 1-4. Cylindrical adult feeding cage.











Figure 1-5.

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



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 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


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.


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


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



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


Figure 2-1.

Habitat of Lutzomyia anthophora.

Nest of Neotoma micropus.

Figure 2-2.


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).


Figure 2-3.

Multiwell plate with lid used for rearing individual

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.


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


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.


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.


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



i1~ PP. .P


Figure 2-4.

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



M 10-



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


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


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


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



a, C

0 2 4 6 8 10 12

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


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


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.


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.


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


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.


Neotoma micropus

Neotoma micropus

Mesocricetus auretus

Mesocricetus auretus

Oryctolagus cuniculus




















Tip Base

32.4 32.4

31.3 36.8

33.2 35.0

34.2 35.6

Rear Foot


35 .*4















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


Figure 2-8.

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


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


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.


Figure 2-11.

Figure 2-12.

L. anthophora engorged on serum or lymph.

Dead female L. anthophora after peritrophic sac



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


(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.


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


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



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.


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.


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


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



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


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


Figure 3-1.

Habitat of Lutzomyia diabolica.

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


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.


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


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 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


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


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


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


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

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



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.


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)


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.


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


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.


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.


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


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


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


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.


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



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.


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


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

nature by the stabilization mechanism discussed by Tesh and Shroyer


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.


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.


RIO GRANDE VIRUS AND Triatoma gerstaeckeri

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



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


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.


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


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


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.




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


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.



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.


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.


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).


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


Figure 6-1.

Electron micrographs of purified Rio Grande virus at


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.



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:


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

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


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.


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


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.


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.


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.


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.


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




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-


The second type of photographic chamber (Figure 8-2) was con-

structed from a spectrophotometric cuvette (Wallace & Tiernan, Co.,



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

Chamber for photographing small insects.

Figure 8-2.


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.



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


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.



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