rearing mosquitoes

8/10/2019 Rearing Mosquitoes

1/124

Pubiished III.

,UlERICAS MXQU I-0 COY I-ROL ,ISSOC I_ITIO~, 15-C.


2/124

MaLlLLal

for

MOSQUITO REAKIKG AND EXPERIMENTAL TECHNIQUES

E~GESE J. GERBERC

Insect Control and Research. Inc.

IIII Sorth Roiling

Road, Baltimore, Jlcl. zrzz8

Bulletin 30. 3

Bulletin SO. 5

is a\.ailable from:

T. G.

R.\LLY,

Executive Secretnry, ;IMCX

P.O.

Box 278, S&m, California 93662

Price 53. j0


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CONTENTS

FOREWORD-KESUETH i_. KTISHT. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , , , . . , .

v

_1CKSOTYLEDGAIESTS

. . . . . . . . . . . . . . . . . . . . . . . . . . .

Ti

ITTRODL7CTIOT . . . . . . . . . . . . . .

xll

I. THE ITSECT_ \R1-

...........................................................

I

Construction of room>.

.....................................................

1

Temperature and humldlt equipment. . . . . . .

2

Lighting ,,............................................................... 3

SeiuritJ- and safet>- medrure>.

. .

-1. Faslllt> requirement< .

I. Building . . . . . . . .

2. Room layout . . .

. .

.

B. Labor&tory

and rearing prcxedures.

.....

I. Eggs ...........................

2.

Larl-at and

pupae.

................

3. _klults .........................

5

. .

5

C. SIiscellaneow requireme nts an d recom menda tion\

5

II. SI.U?XET_1SCE OF MOSQ L-IT0 COLOTIES. . .

;

Equipm ent for handling adult>. . . . . . . . . . . .

I.

;Ispxators or wition deliies.

. . .

2. cage> . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.Adult food . . . . . . . . . , . I. .

.......................

_irtificial feeding techniques. . .

.......................

Induced rndtmp . . . . . . .

....................... . .

Colleiting and handling eggs .. . . . . . . . . . . . . . . .

1;

Larl al rearing ttihnia~ueh dncl tquipnmt

...............

Counting larvae

...............................

Rearing tra -s ..................................

Handling larl ae ...............................

Equipm ent for feecling 1arTde.

Lxval toocf ancl ieedm:

....... ............

_ .

c ..............................

Handling.

counting. separating and sexing pupa e.

........

.

. . .

. .

Sterile

culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 j

I II . PROC EDURES FOR L_~BOR_~TO RYL- RE-ARISG OF SPECIFIC MOSQ UTOkS. .

. .

. . . .

. . . .

. . . .

. .

. . .

. . . .

. . . .

.

.

. .

.......

-

....... _ij

....... ;_j

....... qj

.......

46

.......

ir

.......

45

.......

50

. .

. . *

. . .

. .

. . . . . . . .

.

.

. . .

. . .

. . .

. .

. . . . .

.

.

. .

P~rJi.Op/lOi.,7

. . . . . . . . . . . . . . . . -. . . +. e .. . . . . . - . . . . . . . . . . . . . . . . . . . . . . . . . . . .

50

i


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Pnge

Et.etnupodites

. . . . . . . . . . . . . . . . . . . . . .

. , .

. * . ,

j2

dedes

. . . . . .

. .

. . .

a....,.....

. . . . . . . . . . . . . . . .

*... >3

Aiwigere~

. . . . . . . . 8 . . . . , . . . . .

.,...*.....

. .

. . . .

64

Hcienzngogm . ...,,..*.. . *. . . . . . .a

. . . .

65

Opifes . . . . .

. . . . . *

. ..I.......

. ..a .a a....

. . . .

66

Clllijetn . . . .

. . . . . .

a.. . . . . . . . .

,....*.....

* . . . . * .

68

Crrles

. . . . . . . .

. . . . . . . . . . .

. . .

.a. . . . .

. . . . I . .

69

Deirtocerite_i

. . . . . . . . . . . . . , . . . , . . . . . . . .

* . , .

78

APP END IX A. List of mosq uitoes and mosquito colonies maintained b>- laboratories, . . . . . . . . ig

.IPPEN DIX B. Tables of measu res and equivalents.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . , , . 85

APP END IX C. Relative humidity tables.

. . . . . . . . . . . . . . . . . . . . . , . . . . . . . , . . . . . . . . . . . . , .

89

REFE RENC ES . . . . . . . . . . . . . . . . ..~...~............................................ gI

ii


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LIST OF ILLUSTRATIONS

FIG. I.-Interior of insectary at V-alter R eed _k-m - Institute of Resea rch--US. _km y.

FIG. Z.-Interior or insectary at Insect Control k Rese arch . Inc.-R. Stacks. Baltimore

FIG. 3.--Mo rlan s

colony cage..-CDC.

.....................................

FIG. J.-Gerbergs collapsible cage-R. C. Ruehl. Jr.: Cornell C s E Co.

...........

FIG. 5.-Barraud cage-P. S. Barraud.

.......................................

FIG. 6.-Gillett cag e-J . D. Gillett

...........................................

FIG. 7.-Plastic cage-L7 .S. *Arm .............................................

FIG. 8.--hicCra -s escape-proof colon , cage -CDC .

..........................

FIG. 9.-Pollard cage -D. G. Pollard.

........................................

FIG. I o.-_knimal restrainers in

more Sun . . . . . . . . .

mass feeding of

. . . . . . . . . .

mosquitoes on

. . .

Sun

.

. . .

.

.

. .

chicks-R. Stacks.

. . . . . . . . . . . . .

Pnge

.Co\-er I

. . .

I

. .

8

. . .

9

. . .

IO

Balti-

. . . . . 12

FIG. II.--Apparatus for mem bran e feeding of mosqu itoes--U.S. ;irmy.

. . . . . . . . . . . . .

14

FIG. I z.--_\ppara tus for mem bran e feeding. detail--US. -Arm >-.

. . . . . . . . . . . . . . .

I 5

FIG. r3.-*kliquot container for mosquito larvae-R. Stacks. Baltimore Sun.

........

FIG. I _I.--\Vhite enam el tra s for rearing larvae-L.S. _Lrmy .

...................

FIG. IT.----SYhite en amel dishes for rearm g larv ae-K.S.D._L.

.................... .

. .

I9

. . 20

. . .

?I

FIG. 16.-Construc tion of gall-anized metal rearing tra --Morlan. . . . . . . . . . . . . . . . . . .

21

FIG. r;.-Mass rearing of msoqu itoes in

galvanized tray s-R. Stacks. Baltimore Sun. . . . . . 22


6/124

All Metal Collapsible Cage

(See Fig. 4, Page 9, This Bulletin)

The Standard Cage in Major Laboratories

All aluminum construction with plastic ham mock

Metal construction eliminates mold and contamination

Hinged for easy collapsing and storage

Cages easily disassem bled for cleaning

Escap e-proof feeding ham moc k enables e xposu re of animal for blood feeding with-

out mosquito escapage

Sucrose solutions on cotton balls can be fed mosquitos through hammock ; sugar

does not com e in contact with metal parts.

Surgical stockinet sleeve provides easy access inside cage

Econom ical in cost - efficient in operation

Available in three

convenient sizes

Gl 12 x 12 x 12

G2 24 x 24 x 24

G3 177/8x 18~ 22-1116

For further information and prices write-

LOW-COST AUTOMATIC HUMlDlFlCATION

Whe ther for mosqu ito-rearing or experimen tal conditions, there

is a Standard Humidifier available to provide accura te, auto-

matic humidification for your needs . Famo us Standard units

are engineere d for long, trouble-free service an d easy installa-

tion, and are priced surprisingly low. All units are fully

guaranteed.

Wfite for 4-page

free brochure

a

Model 42L

Humidifier

State de tails of your

project for promp t

guaranteed

recom-

mendation


7/124


8/124

On behalf of the members of the American Mosquito Control Association and

of mosquito workers everywhere, I first thankfully add this fine publication to my

library of mosquito literature and secondly extend sincerest thanks to its autho r

for his devotion to its preparation.

KENNETH

L.

KNIGHT,

Head

Department of Entomology

N. C. State University

Raleigh, N. C. 27607;

Chairman, Editorial Board of

Mosquito News, Journal of the

American Mosquito Control Association

vi


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DEDICATION

This manu al is dedicated to Mrs. Helen Louise Trembley Durkee,

who devoted a large part of her time and effort to producing the

original B ulletin 3, entitled Mosquito Cu lture Techn iques and Experi-

mental Procedures. Her work has served for many years as a guide

to entom ologists and other w orkers in the field of culicidology.

ACKNOWLEDGMENTS

The preparation of the present manual was originally intended to be the work of a

committee. However, committees perform best when they can meet, discussand work

together. Because distance and individual research problems made effective commit-

tee work very difficult and subject to long delays, I decided to tackle the problem

alone, with whatever local assistancecould be obtained. I was fortunate in having

Capt. James W. Gentry, U.S.A. (Ret.), join our staff in 1967. He spent many eve-

nings gathering data. Dr. Ronald A. Ward, at the Walter Reed Army Institute of

Research, was extremely helpful in securing literature and locating mosquito rearing

laboratories. This, then is the working Committee to whom I offer my sincere thanks.

Dr. Ross H. Arnett, Jr. kindly reviewed this manuscript and used his grammati-

cally minded red pencil with justifiable abandon. My sincere thanks and appreciation

are offered-any errors remaining are mine, not his.

To the many other culicidologists that assisted n one way or another, I give my

grateful acknowledgment for help and advice.

EUGENE J. GERBERG

vii


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INTRODUCTION

The need for information on mosquito rearing has been intensified by the increased

interest in mo squito biology, control, chemoltherapy of mosquito borne diseases, and

in other investigations in which mosquitoes are used as experimental animals. Since

the publication of Helen Louise Trembleys Mosquito Culture Techniques and

Experimental Procedures

in 1955 , published literature in this field has increased like

salt marsh mosquitoes after a high tide.

The acquisition elf knowledge concerning proper con trol of mosquitoes and mos-

quito borne diseasesrequires that studies be made of the biology, p hysiology, anatomy,

genetics, taxonomy and ecology of the insect. Insecticide resistance, biological, chemi-

cal, and integrated control must be investigated.

Research on mosquito genetics

may have far-reaching consequences in the control of m osquitoes and m osquito-borne

diseases. Th e use of m osquitoes as screening agents for pesticides or chemothera-

peutic compounds requires large numbers of mosquitoes. All phases of mosquito

research usually use individual or quantity rearings of mosquitoes.

Mass rearing of

mosquitoes for control purposes may become, in the near future, as common as mass

production of pesticides.

The purpose of this man ual is to provide information on the rearing of mosquitoes.

A thorough search of the literature p rovided many useful references. The great

amou nt of literature made this no easy task. Undoubted ly, articles were overlooked,

but it is hoped that no serious omissions have occurred. An apology is offered to

those authors so neglected.

Author credit-lines omitted in the text are included in

the list of references at the end of the rearing description for the species.

For some

species, particularly those reared in the authors laboratory, the technique is described

in more detail. Often this sam e technique will apply for related species as well.

No method for rearing mo squitoes is guaranteed. Perhaps the most important

requirem ent for successful rearing o f mosquitoes is attention to detail. Mosquito

rearing to be successful requires attention 24 h ours a day, 7 days a week.

The basic

rules are: avoid over-crowding mosquitoes, and overfeeding larvae; a void pesticide

contamination, observe temperature and humidity requirements, standardize rearing

methods and avoid nonstandardized food.

Observance of these principles should

produce uniform animals.

The following guide to mosquito insectary construction and rearing practices is

presented step by step with no elaboration of theory. Much of this theory m ay be

yearned by reference to the literature cited.

.

I.111


11/124


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the room.

Whenever possible use coved wall floor junctures and caulk and seal the

cracks and crevices n the walls.

The best insectariesare windowless with low ceilings

(not more than 8 also painted white or light colored. If possible, use recessed

lighting fixtures that are flush with the ceiling.

Smooth floors without a floor covering are recommended. A light colored epoxy

floor paint can be used to improve the appearance. The doors of the rearing area are

best constructed of metal and painted white or light colored. Weatherstripped door

frames and jambs are necessary for a tight fit. Overlapping marquisette curtains

hung on the inside of the door frame provide additional mosquito barriers (see sec-

tion on mosquito security).

Hot and cold running water and a sink with ample draining surfaces are required

in the rearing room. A water blending tank r installed above the sink facilitates

the filling of trays, separation of pupae and other tasks that require water to be a

certain temperature. Several water outlets around the room will avoid the clutter of

hoseson the floor.

Movable tables, benches, chairs, storage cabinets, etc., are preferable, since, in

general, permanently installed furniture prevents new arrangements to meet changing

conditions. Some laboratories have all furniture mounted on casters or wheels so

that it can be readily moved.

TEMPERATURE AND HUMIDITY EQUIPMENT

Temperature and humidity controls are probably the most important factors in the

successful earing of mosquitoes. The simplest method of providing regulated heat

and humidity is the use of a small electric light bulb and a wet towel draped over

the cage. Th

e most elaborate is a complex environmental chamber with programmed

electronic controls of temperature, humidity, and photoperiod.

The size of the

insectary will regulate the type of temperature and humidity control system required.

The proper design of equipment for the production, control, and recording of tem-

perature and humidity may require the services of an engineer.

If a large installation is planned, it is better to have

2

smaller heating and cooling

units than one large one.

For example if it is determined that 6 tons of air condition-

ing is

required, use two S-ton units rather than one 6-ton unit.

Install the heating

and air conditioning system of the peripheral area so that if all the inner area systems

fail, the peripheral system can be shunted to the inner area. When there is a possi-

bility of contamination from pesticides from adjoining laboratories, install air condi-

tioning equipment to insure that the outside air intake does not pick up contaminated

exhaust air.

Automatic thermostat controls for the heating and air conditioning

system are best.

Humidity regulation is usually provided by comparatively small commercially avail-

able units.2

If a source of clean compressed air is available, the air supply and water

supply can be connected and regulated by solenoids and humidistats. The water is

forced out through fine nozzles installed in a hose and produces a fine mist. Steam

units are available which will provide warm moisture. Various improvised humidi-

1 Sarco Co., Inc., Allentown, Pa. Blender Type DB.

2 Walton Laboratories, Inc., Union, N.J., Standard Engineering Works, Pawtucket, R.I., Bete Fog

Nozzle, Inc., Greenfield, Massachusetts.

2


13/124

fying methods have been described, including such methods as using wet towels,

blowing a fan through layers of wet excelsior or gauze; steam from boiling water or

from radiators; wet sand; chemicals in water.

In determining the type of humidifier unit that should be used in a rearing room,

certain factors must be taken into consideration:

(I) The relative humidity desired and what variation would be permissible; (2)

Room size-length, width and height; (3) A

verage temperature; (4) Heat factors,

i.e., number of people or animals in the room; number of lights and wattage, other

sources of heat, and heat loss; (5) Construction of room-walls, floors, ceiling, parti-

tions, windows; (6) Openings to other rooms; (7) Exhaust fans-capacity. As in

temperature control, it is better to have two smaller units than one large one.

Use a recording hygrothermograph

for a permanent, continuous record of tem-

perature and humidity conditions.

Humidity can be checked with a sling psychrom-

eter, a bulb psychrometer, or a Honeywell hygrometer with electronic sensors which

will show the relative humidity at the exact location desired without disturbing air

movements. Although it is recognized that the critical factor is the rate of evapora-

tion, usually an indication of the relative humidity is sufficient for successful earings.

A maximum-minimum thermometer will indicate the total range of temperature

change for any desired period.

References-

Barlow (1961); Bertram and Gordon (1939); Buxton (1931); Flitters (1964); Haufe (1964);

MacPhee and Patterson (1958)

;

P

rovost et al. (1965); Wharton and Kniille (1966); Winston

and Bates (1960).

LIGHTING

The photoperiod and light intensity affect the development of the various stages

in the life cycle of the mosquito and are the subject of a great many papers (see

references). In the insectary, a cycle of 14 hours of light and IO hours of darkness

appears to allow the best and most uniform development, without interfering with

normal working hours of the staff.

Some speciesmay require a crepuscular period to swarm and mate. The automa-

tion system described by Levin, Kugler and Barnett (1958) coordinates the numerous

pieces of electrical equipment that regulate temperature, humidity, and light. This

type of equipment can be readily modified to dim lights gradually for dusk effect,

turn off all lights for darkness, then turn up lights gradually for dawn and turn on

full lighting for daylight.

The standard fluorescent lights usually supply adequate light for daylight.

Some

especially made fluorescent lamps generate light at wavelengths approaching day-

light, when required by exacting experimental work.

Incandescent lights connected

to a variable transformer can be used for gradually diminishing or increasing light

intensity. A bl

ue or blue-green light stimulates swarming of some species (Bates

949; TheOdOr and ParSOnS 1945).

Commercial light meters are used to measure light intensity. Evans (1961) de-

scribes a portable visual photometer for measuring very low light intensities. The

1 Bendix Corp., Environmental Science Division,

Baltimore,Maryland,21204-Hygrotllermograph

Model NO. 594,

Belfort Instrument Company, Baltimore, Maryland 21202.

3


14/124

scale of a regular Weston light meter used for photograph y can be converted e asily

to foot-candles by multiplying the reading by

2.

References- I

Bates (1941)

;

Belton et al. (1967) ; Brennan and Harwood (1935)

;

Callot (1965)

;

Eldridge

(1963); Evans (1961); Flitters (1964); Fowler et ctl. (1958); Haufe (1962); Hubert et al.

(1954); Jobling (rg35), (1937); Killough and Weidhaas (1963); Levin et al. (1958); Nielsen

and Haeger (1955); Omardeen (1958); Parker and Rozeboom (1960); Seaton and Lumsden

(1941) ; Tate and Vincent (1932) ; Theodor and Parsons (1945).

SECURITY AND SAFETY MEASURES

The escape of possible insect vectors of disease, like the release of pathogenic

organisms cannot be tolerated.

The responsibility of the entomologist rearing mos-

quitoes is similar to the responsibility of the bacteriologist cultivating pathogenic

bacteria. Certain routine sec urity measu res are practiced by laboratories involved in

the rearing of medically important arthropods.

The Comm ittee l on Experimental Use of A&s aegypti of The En tomolog ical So-

ciety of America has developed a set of recomm endations on mosquito security and

surveillance that serves as a guide for mosq uito rearing laboratories. The follolwing

recomm endations closely follow, but are not an exact copy of the ESA report.

A.

FACILITY REQUIREMENTS

I.

Building

Window less construction, if possible, is preferred. If window s exist, keep locked.

A sign perman ently fastened near the lock indicating that the windows always should

be locked, will help assu re this protection.

Glass brick can som etimes be used to

replace windows.

Screened windows norm ally are not to be used, but if for some

specific reason they are required, double screens with

22

mesh screening and an addi-

tional outside protective screen of heavy hardw are cloth will provide the required

security.

2. Room layout

a. When the rearing room and laboratory are not one and the same, then the

rearing room should be an annex of the laboratory room and open into it,

If another

door exists or is required for safety reasons it shou ld be mark ed for emergency use

only .

Plan exits from the laboratory roo m o r rearing room (if not an annex to the

laboratory room ) through an ante-room , i.e. a small vestibule with entry or exit doors

or through another small room.

A com pletely empty ante-room is best, with white

or light colored walls and ceiling.

Post signs to be sure one door is closed before the

other is opened.

Protect the entrance to the ante-room from the laboratory with

some protective barrier such as close weave nylon netting or an air curtain.a

b. Paint the walls and ceiling of the laboratory, rearing room, ante-room and other

work areas with a light shade of paint (gloss ename l, epoxy, etc.), preferably white.

Light colored ceramic tile may be substituted for paint.

1

Committee members: G. Craig, Jr., E. J. Gerberg, C. Judson, D. W. Micks, M. W. Provost, I-I.

F. &hoof, C. N. Smith.

2 Ecco Aire, Inc., New Castle, Pa.; Dynaforce Corp., Queens Village, N.Y. 11428; Penn Ventilator

Co., Inc., Philadelphia, Pa., 19140; Neico Products, Inc., San Francisco, Calif. 94124.

4


15/124

c. Screen all floor drains even though they enter a closed system. It is best to

drain into a tank or dry well in which insecticide is regularly introduced.

d. Screen or filter all air vents to the outside.

B. LABORATORY ND REARING PROCEDURES

I. Eggs

(a) Store eggs in a container in the rearing room.

(b) Treat surplus unhatched eggs with hot water or chemicals to kill them. If

possibleburn the old egg papers.

2. L.uruae and Pupae

a. Larvae

( I) In laboratories that do not maintain

containers with tight fitting lids to

adults. In mass-rearing laboratories,

several larval rearing racks are housed

b. Pupae

( I) Collect pupae routinely, so that no

containers.

T-day-week operations, rear larvae in

prevent the escape of any emerging

normally on a T-day-week operation,

in enclosures nside the rearing rooms.

adults emerge in the larval rearing

(2) Place pupae in special containers inside cages. Provide each container with

a lid so that the container can be closed after the pupae have emerged. It

should be removed from the cage through a sleeve.

(3) Treat unneeded larvae, pupae, and discarded rearing water by heating or

with chemicals so that only dead waste materials enter the sewage system.

3. Adults

a. Use sturdy cages, preferably of metal, fitted with a surgical stockinette sleeve.

When used properly the tight-fitting sleeve will allow manipulation of objects within

the cage. Th

e mosquitoes obtain their blood meals by feeding on the host through

the screening of a nylon mesh hammock.

This is preferable to introducing the verte-

brate host animal into the cage.

b. Keep the rearing room neat and clean.

Remove excess sugar pads, fruits, etc.

c. Hosts for blood feeding.

(I) Provide animal rooms separate from the rearing room. Check host animals

to be sure they are free of mosquitoes.

d. Transfer of adults.

All insects ransported from one secure laboratory to another must be housed within

a screened, securely closed container.

C. MISCELLANEOUS EQUIREMENTSAND RECOMMENDATIONS

I. Train employees to handle mosquitoes and make them aware of the potential

hazard of escapees.

Loose mosquitoes endanger the entire project. Even one

loose mosquito must be killed or caught immediately.

A sign in the rearing room

indicating the name of the individual responsible for the mosquito security helps

to maintain vigilance.

Replace careless workers or technicians with competent,

conscientious personnel.

2. Do not remove any living stage of the insect from security conditions without

taking proper precautions.

5


16/124

3. Place surveillance oviposition-traps (Fay and Perry, 1965 ; Fay and Eliason, 1 966)

in the laboratory, rearing rooms and adjoining rooms. Numb er each trap and record

on a map. Keep a record of the weekly check.

Place additional surveillance oviposi-

tion-traps outside within 25 feet of the laboratory bu ilding, at approx imately r5o-foot

intervals. Rem ove all potential breeding containers from the surveillance area. (For

detailed information on preparation of surveillance oviposition-traps, and selection of

sites, see Ovipos ition trap reference handbook , CD C, Aedes aegyptt Handboo k series

No. 6, 1967.)

4. Personnel

Limit access o rearing and laboratory facilities to authorized personnel who under-

stand the need to retain all live s tages under security conditions.

.References-

Bertram

(1958) ; Fay

and Eliason (1966);

Fay

and Perry (1965); Hocking,

B. (1960)

;

Pratt

and Jakob (1967).


17/124


18/124

this is done the significance of the DR S can be ascertained for any species of m os-

quitoes. Preliminary research indicates that a DR S of 1.8 gives good results.

Many cages for holding and/or mating mosquitoes have been described. McKiel

(1957 ) describes a cage 30 wide x

20

long x

20

high (RS= 12,90 0 sq. cm.) for

4000 Aedes aegypti adults.

This would provide a DR S of 3.2. Weathersby (1962 )

used a cage 18x18~18 for

Aedes togoi

and

Armigeres

sp. Blakeslee et

al., (1962)

used a cage 24x24~24 for Culex erythrothorQx. Kitzmiller and Micks (1954) cage

is 12X12x12

and 24X24X24

cages for Culex

pipiens.

Hayes and Morlan ( 1957)

used a similar size cage for Aedes

triseriatus.

Williams ( 196 2) employed a 12x9s x

14 cage for the same species.

None of these authorrs mention the density per cage.

McLintocks (1952 ) cage is 12x12~2 2 (RS= 6812 )

f or 1500-2000 Culiseta inornata.

This gives a DR S of approximately 3.4-4.5. Casanges et al. (1949 ) describe a

22x.36x24 cage (RS-17 957) f

or

3

000-5000 Anopheles quadrimaculatus, or a DRS

of 3.6-6. Aedes aegypti have been kept in many types of cages.

Christophers (1960)

descriibes a numb er of cages for this species including one type 8 2x8 2x8 1/z and

another 14x8~/~~12.

Liles et aZ. (1960) employed a cage 12x8~9 for IOO A. aegypti.

Wall is (1954) cage is 36x36x30

for 10-350 female A. aegypti and concludes that

the density of ovipositing females did not increase the percentage of eggs produced.

Roth (1948 ) used an 11x11~15 cage for his work on aegypti.

Morlan et al. (1963)

indicate that their colony cages (Fig. 3)

consist of approximately 10,00 0 adults in

MOSQUITO COLONY CAGE

ASSEMBLY

FIG. 3.-Mm-hs colony cage.

8


19/124

FIG. q.-C &be rgs collapsible cag e.

a

22x22~22

cage, (RS-12,487 ) or a DRS of 1.2. Thus, there is no standardized

size, type or construction material or population density.

A standard size cage may be of value for rearings for research purposes.

To this

end a light weight aluminum cage has been developed 1 that is now widely used

(Fig. 4). This cage is constructed of pressed-form .032 inch alum inum frame 7s

thick by 3/4

wide and is screened with 18 x 22 mesh aluminum screening. The

screening is held in place by a thin rubbe r sp line, for easy replacem ent of screening

when necessary.

The floor of the cage is constructed of either .090 or .03 2 inch

aluminum sheeting depending upon the model of the cage. A hamm ock constructed

of a fine mesh nylon is an integral part of the top of the cag e.

Anesthetized animals

such as guinea pigs with shaved backs can be used for blood feeding.

Sucrose-soaked

cotton balls, fruit, and su gar, can be placed in the hamm ock for o ther types of feed-

ings. The

cage is fitted w ith a 24-inch long su rgical stockinette sleeve which fits

a 9l/2x97/s opening.

The size of the cages may be

12~12x12

(3ox3ox3ocm) or


20/124

; 1

I 1

I

I

:

I

;, I

,I

_--

----___

,._. _----

_ -q

BARRAUD

CAG E

Barraud,F?J.(1929)

FIG. 5.-Barraud cage.

GILLET CAGE

Gillet, J. G.(1962)

PIG. 6.-Gillett cage

:

-LJ

L

POL LARD

CAGE

,-

Pollard, D.G,(1960)

FIG, 8,-&kCrays escape-proof colony cage,

FIG. g.-Pollarcl cage.

IO


21/124

24x24~24~

(6ox6ox6ocm ) ( h

own in Fig. 4 as G-I, G-2, and G-3, respectively), or

other combinations.

All cages are collapsible for ease of storage and can readily be

cleaned by steam or hot water.

These cages have been successfully used with many

species of mosquitoes in most of the major mosquito laboratories.

Many kinds of cages for special purposes have been described. Perhaps one of

the most useful cages for field work is the Barraud Cage, Fig. 5, (Barraud, Igag),

a cage with a wire frame over which is hung netting. Lantern globe cages, made

from the glass globes of kerosene lanterns have been widely used (Fig. 6) (Trembley

195 5; Gillett 1962 ). Others have modified glass cylinders (Burgess and Young

rg46), glass jars (Eldridge and Gould 196 0) and plastic containers (Fig. 7) (Yolles

FIG.

7.-Plastic cage.

and Knigin 1943; Young and Burgess 1946; Barnett 1955; Gerberg et al.

1967;

Alger

1968 ). Escape proof cages have been described by Bar-Zeev and Ga lun (1960 )

and McC ray (Fig. 8) (1963 ), b u are not practical for mass rearings. Pollard (1960 )

describes a cage (Fig. 9) that minimizes the escape of mosquitoes and is still easy to

References-

Alger (1968); Barnett ( 1955) ; Barraud ( Igag) ; Bar-Zeev and Galun ( 1960) ; Blakeslee et al.

(1962)

;

Burgess and Young (1946); Casanges et al. (1949); Christophers (1960); Eastwood

and Jamieson (1965); Eldridge and Gould (1960) ; Gerberg et al. (1967) ; Gillett (1962) ;

Hayes and Morlan (1957); Kitzmiller and Micks (1954); Lennox (1959); Liles et al. (1960);

McCray (1963); McKiel (1957); McLintock (1952); Morlan et al. (1963); Pollard (1960);

Reynolds (1960); Roth (1948); Trembley (1955); Wallis (1954); Weathersby (1962);

Williams (1962); Yolles and Knigin (1943);

Youngand Burgess1946).


22/124

ADULT FOOD

Although the female mosquito usually must ingest a blood meal for ovarian devel-

opment, adu lts of both sexes require carbohydrate foods in addition. Carbohy drates

are generally supplied as a sugar solution.

Although sucrose and g lucose in concen-

trations of from 37/o to 20% have been used,

10%

sucrose, made by dissolving

IOO

gms of ordinary household white sugar in one liter of water appears to provide

the best results.

Other forms of suga r, such as corn syru p, honey, various fruit

juices, raisins, apple slices, and bananas , have also been used.

FIG. Io.--Animal restrainers in use; mass feeding of mosquitoes on chicks.

Soake d cotton balls are the easiest method of providing the suga r solution to ad ult

mosquitoes. Th

e cotton balls are soaked in the

IO~~

sugar solution, the moisture is

squeezed out and the balls are then placed on the top of the cage. Usually 4 cotton

balls, changed daily, are sufficient for a 30x30~30 cm

(1x1~1

cage. The cotton balls

must be changed daily. Coluzzi (1964) used a tank and a side cup trough. The

liquid in the cup maintains its level and keeps a piece of filter paper wet continuously.

Porter et al. ( 196 1) describe a small solution feeder for adult mosquitoes m ade from

a ro-dram plastic vial in which is inserted a wick made of a small roll of blotting

paper.

The vial is filled with the feeding solution and the mosquitoes feed through

12


23/124

small holes punched through the wall of a plastic vial. Eliason (1963) used a tech-

nique of feeding mosquitoes on solid sugar.

Blood meals are provided by supplying animals that have been anesthetized or

restrained, and placed on the adult cage, rather than in the cage.

This prevents loss

of mosquitoes, hazards to the technician, and destruction of mosquitoes by the blood

donor.

Animals may be anesthetized by the use of I grain nembutal (see appendix)

administered intraperitoneally at the rate of

0.2

ml for each I lb (0.5 kg) of body

weight. Various types of restrainers are available commercially and are described

by Porter et al. (1961); Gerberg et al. (1967) (Fig. IO); Morlan et al. (1963);

Trembley (1955); Dunn (1932); Jones and Scheltema (1952).

The more commonly used sources of blood meals, other than a technicians bared

arm, are guinea pigs,

rabbits, mice, rats, hamsters, monkeys, and chicks.

Blood

meals may also be supplied by artificial means (see artificial feeding techniques).

Shave the animal prior to presentation to the mosquitoes.

Some prefer shaving or

clipping the hair from the back of animals, and others the underside. Animal clip-

pers have proved very useful for this purpose.

References-

Dimondet al. (1955); Dunn (1932); Eliason 1963); Gerberg t

al. (1967);

Greenberg

1951);

Knierim et al. (1955); Lang and Wallis

(1956);

L 1es

et

al. (1960); Morlan et

al.

(1963);

Porter et al. (1961); Roy (1936);

T rembley

1955); Woke (1937);

Woke et al. (1956).

ARTIFICIAL FEEDING TECHNIQUES

Xmembrane feeding technique is one approach for artificially providing a blood

meal to mosquitoes.

This technique, used with other haematophagous arthropods

prior to the successfulfeeding of mosquitoes, consists of a membrane-covered con-

tainer, such as a glass tube or test tube, containing the liquid mosquito food. The

container is placed so the membrane side is accessible o the mosquitoes for feeding.

The liquid should b

e several degrees warmer than the ambient temperature to induce

certain species to feed. Tarshis (1958)

review of artificial feeding techniques

includes the various types of membranes used, and describes various techniques and

materials.

Knowlton and Rowe (1935)

used animal mesentery bags to feed mos-

quitoes on

suspensionsof diseased (equine encephalitis) brains. Woke (1937) fed

A&es

aegypt;

on whole blood through rat skin membranes. Yoeli (1938) gave

Anop&des elutus blood through rabbit skin membranes. Bishopp and Gilchrist

(1944, 1946) induced feeding by Aedes aegypti on blood through chicken skins.

Greenberg (1949,

1951)

reports the feeding of Aedes aegypti on a variety of artificial

fluids through Baudruche (a bovine

intestinal preparation) capping membranes.

Eyles ( 1952)

used hog-gut sausage casings, and Trembley (1952) used Baudruche

capping membranes

and chicken-skin membranes.

Kartman (1953) fed Aedes

aegypti

blood through hog-gut sausage casings.

Bar-Zeev and Smith (1959) in

studying the action of repellents, gave A.

aegypti titrated blood through a membrane

made from the outermost layer of ox caecum.

Tarshis (1959) discusses ive different

1 Oster Mfg. Co., Milwaukee, Wise. Model A2 Clipper, No. 40 blade.


24/124

memb ranes and an apparatus for feeding Culex

tarsalis.

Collins et al. (1965 ) used

the Baud ruche (untreated) memb rane to feed Anopheles mosquitoes Plajmodium

falciparum

infected blood.

FIG. I I.-Apparatus for me mb rane feeding of mosq uitoes.

Rutledge et

al. (

1964)

d

escribe in detail their techniques for mem brane feeding

of mosquitoes, Figs. I I, 12.

The feeder is mad e of heat res istant glass or stainless

I4


25/124

Mosquito Membrane Feeder

Inlet

FIG. I x--Apparatus for

detail.

membrane feeding,

steel. Circulating warm water controls

the temperature of the blood. Accord-

ing to them, chick skin is superior to

the Baudruche membrane, and chick

blood is more acceptable than erythro-

cyte extract or serum.

Blood meals can be supplied by means

of preserved blood absorbed on a cotton

pad.

Russell (

193 I )

reports feeding

mosquitoes blood distributed over net-

ting.

Others (MacGregor and Lee

1929; Knowlton and Rowe 1935; Shiavi

and Franc0 1949; McL,intock 1952;

Dimond et al. 1955; Trembley 1952;

Liles et al. 1960) mix blood with honey,

sucrose or glycerine.

Knierim et al.

(1955) used frozen titrated beef blood plus 10% honey (or

without honey) heated

to 35-38 c.

References-

Bar-Zeev

and Smith (1959)

;

Behin (1967)

;

Bishop and Gilchrist (1944, 1946)

;

Collins (1963)

;

Collins et al. (1965, 1966); Co11

ns et al. (1964)

;

Collins et a/. (1964)

;

Dimond et al. (1955) ;

Eyles (1952) ; Galun (1965) ; Greenberg (1949) ; Grifliths and Gordon (1952) ; Jones (1956) ;

Kartman (1953) ;

Lang

and Wallis (1956) ;

Liles et al. (I

960) ; MacGregor (I

930) ; MacGregor

and Lee (1929); Mattingly (1964); McConnell (1956); Msangi (1956); Ogden (1961);

Ogunba (1967); R

ass (1956) ; Russell (1931) ; Rutledge et al. (1964) ; Schiavi and Franc0

(1949) ; Shambaugh (1954); St. J h n et d. (1930); Tarshis (1957, 1958, 1959); Trembley

(1952); Willis (1958); Woke (1937); Yoeli (1938).

INDUCED MATING

Aedine mosquitoes are normally difficult to maintain in the laboratory but Mc-

Daniel and Horsfall (1947)

used efficient procedures for stimulating copulation and

obtaining Aedes stimulans and Aedes vexans eggs. Horsfall and Taylor (1967)

modified the technique and used it successfully to mate over 40 species of mosquitoes.

Copulation is induced by manipulating an immobilized female into contact with a

decapitated male.

This technique consists of using at least 3 day old mosquitoes.

The female is anesthetized lightly with chloroform, and is picked up with a suction

pipette and manipulated to make contact with the male. The dorsum of the thorax

of living decapitated males is cemented with a non-toxic glue to a small piece of

glass slide. Contact between the two immobilized mosquitoes is accomplished

manually under a

20x

stereomicroscope.

The female is placed in the mating position

with the ventral side up and the head directed away from the male.

Apexes of the

two abdomens are then brought together, venters uppermost. The male is stimu-

lated by touching the claspers several times with the female. Insemination is usually

completed within a 5-50 second period.

Usually 3-4 males are thus prepared so

that if one male is unresponsive,

others are immediately available. Frizzi (1958)

used the induced copulation technique on Anoptieles macdipennis with variable


26/124

results, and Carvaglios (1961 ) using the technique without decapitation of males

maintained a colony of Anopheles Zabranchiae.

After mating, the females are transferred to a feeding cage for recovery from

anesthesia.

A blood meal is provided after 12 hours. Bake r et L$ (1962 ) obtained

IOO O/~ fficiency in single pair matings of A.

maculipennis

by induced copulation.

Anopheles punctipennis, A. quadrimaculatus, A. freeborni, and A. albimanus were

maintained in the laboratory by the above authors, using the induced copulation

technique.

Wheeler (1962 ) modified the technique by gluing cold-anesthetized mosquitoes to

the heads of insect pins.

The female-bearing pin is inserted into a cork glued to a

microscope slide so that the mosqu itos body axis is in a horizontal plane. The

male-bea ring pin is inserted into an adjustable pinned insect holder an d then

oriented until the genitalia meet. Ow Yang et al.

(1963)

simplified the technique

by using 3-6 day old males caught w ith a fine pipette attached to a suction device, and

pinned latera lly through the thorax using a minuten pin fixed into the end of a 6-

long soft wooden stick. The females are blood-fed 2 days after emergence, and are

collected in individu al tubes the next m orning.

The female is anesthetized with

ether and tipped out on her back onto a clean white table. Copu lation is achieved by

bringing the pinned male down at about a 45 angle to the female. When copula-

tion is affected the pair can be lifted together, with the fema le firmly clasped by the

inale, and gently dropped into a container, by removing the male from the pin.

The

mating procedure is carried ou t under a microscope at a room temperature of 26-28

C. and RH of about 80%. The authors were successful in maintaining a laboratory

colony of

Anopheles maculatus

by using this technique.

Coluzzi (1962) maintained Culex

impudicus

and C.

territans

by induced mating.

Hors fall (1964 ) states that species of Aedes, Anopheles, Culex, Culiseta and Psoro-

phora have been effectively insemina ted in his laboratory. He recomm ends that

adults of the same age be held for a 60-72 h ours premating interval at 80 + 7; RH .

They are fed on a carbohydrate diet (honey diluted I :I w ith water). Mating is

carried out in a room with a temperature of 26-28 C. and about 85% RH. One

technician prepares the males, the second prepares the females and joins the two in

copula, at the rate of 40 matings per hour.

The males are collected into individual

tubes and the open vial is inverted over a Buchn er funnel through which flows a

gentle stream of CQ2.

Immediately upon knockdow n the male is plucked from

the funne l with forceps and is glued ventral side upw ard to a glass slide (casein

glue is used).

Four or five males are placed in a row. Decapitation may precede

gluing, or delayed for longer surviva l until just prior to the act of copulation, w hich

will not take place unless the male is decapitated. The male survives longer if

decapitation is delayed.

Initiate the mating procedure only after full recovery of the

males (2-5 minutes).

The females are anesthetized with chloroform only to the

point of relaxation and are then picked up with a vacuu m p ipette made from a 5-mm

glass tube, bent at right angles and d rawn to an open tip small enough to apply to

the anterior slope of the mesonotum.

A vacuu m pum p with a bleeder tube or valve

is used. Th

f

emale is held ventral side uppermost and broug ht into contact with

the claspettes.

Manipulate the female so that her genital opening is brought proximal

to the phallosome of the male.

Proper positioning is prerequisite to bringing abo ut

complete union.

For

Aedes

an angle of about 120 between the .?bdomens will

16


27/124

permit proper union.

For Culex the angle must be nearer to 180 Horsfail blood-

feeds on the day following mating

(6-12

hours between time of anesthesia and

feeding).

Baker (1964) feeds th

e males, prior to copulation, with honey on a cotton ball,

and the females with water on a saturated sponge. Generally he provides a blood

meal for the female shortly before mating. Baker does not anesthetize the males

but collects them with an aspirator and softly blows them between layers of cotton

wool. This holds them until pinned laterally through the thorax using a dissecting

needle fixed into the end of wooden stick

7.5-15

cm (3-6) long. If it is advan-

tageous to keep the male alive, he can be held with a vacuum pipette.

Baker reports

success with Culex t~salis using this technique.

Esah and Scanlon ( 1966) estab-

lished a colony of Anopheles b. balabacensis by induced mating.

References-

Baker (1964)

;

Baker and Kitzmiller (I 961)

;

Baker et al. (1962)

;

Burcham (1957)

;

Caravaglios

(1961); Coluzzi (1962); Esah and Scanlon (1966); Frizzi (1958, 1959); Gopal and Wattal

(1962) ; Hayes, D. E. (1968) J Hayes, R. 0. (1953); Horsfall (1964); Horsfall and Taylor

(I 967) ; McDaniel and Horsfall ( 1957) ; Ow Yang et al. ( 1963) ; Wheeler (I 962) ; Wheeler and

Jones (1963).

COLLECTING AND HANDLING EGGS

The eggs of mosquitoes are usually deposited on the surface of the water, or on

surfaces that are subsequently inundated. To obtain mosquito eggs, a reasonable

substitute must be provided for the oviposition site.

A plastic container approximately rgxrox5cm (6~4x2) has proved quite successful

for anophelines.

A layer of cotton balls is placed in the bottom, covered with a

sheet of filter paper or paper toweling and then flooded with water so that the water

surface barely covers the filter paper (Gerberg et al. 1968). Some investigators use

bowls or pans of open water as oviposition sites. Giglioli (1947) used a white glazed

china finger bowl (II cm in diam. 7 cm in depth) half filled with water. Others

have lined the oviposition container with paper to prevent the anopheline eggs from

sticking to the sides of the container. Kepler et al. (1964) employed a water-filled

petri dish, the bottom lined with a strip of filter paper. Yoeli and Bone

(1967)

provided small cork rafts and protruding stones on which the females alight prior

to laying eggs.

Trembley (1955) reports that some Anopt2eZes preferred dark colored

containers or containers lined with darker paper toweling.

Species of the genus CuZex usually lay their rafts of eggs on open water, in pans,

bowls, petri dishes or other type containers. Krishnan (1964) claims that more egg

rafts were obtained in trays containing hay infusion than in those containing tap

water.

Jupp and Brown (1967)

confined individual gravid females in laying tubes

r in. in diameter containing a little water and a strip of filter paper down the side

which provided a suitable surface for the mosquito to rest on. The tubes are closed

with nylon gauze on which is placed a cotton pledget soaked in sugar solution.

Takahashi (1968) bt

ained eggs of C. portesi by placing an inverted clay flower

Fat, with an additional I

side entrance hole in a white enamel dish containing

water.

Inside the flower pot and standing in the water is a black-painted g ass

oviposition jar.

7


28/124

Laurence (1960) collects Mansonia eggs from floating discs of plastic-impregn ated

&aft paper in an oviposition chamber.

Usually aedine eggs can withstand desiccation and are collected on a moist sub,

strate.

The most comm on method for the collection of A. mgypti eggs is to use a

3

wide paper toweling, lining the interior of a pint ice cream carton o lr a glass jar.

Fill the carton with enough water to cover approximately I in. of the paper toweling.

The eggs are

conditioned by leaving the egg paper in the container for an addi-

tional 24 hours, after removing the co,ntainer from the cage.

The egg paper is then

air dried (at 80 F. and 80% RH) for

4

d

a

y

s and then stored for future use. Other

A&es species may be induced to lay eggs on various types of m oist surfaces.

Cotton

wrapped in cheesecloth, porous sponges, sphagnum moss wrapped in cheesecloth,

large-pored plastic materials, and other types of rough or porous materials have

been used as oviposition sites.

Reynolds (1960 ) describes an escape proof m osquito

egg harvester, for use under maximum mosquito security.

Anoplieles, Culex and

Culiseta eggs

are not readily stored and are usually hatched

in the water where laid. Cond itioned aedine eggs can be stored. To hatch these

eggs, place in water with a reduced oxygen content.

Many methods for deoxygena-

tion have been described, since the most uniform method of hatching is by imm ersing

the egg papers in deoxygenated water.

To make deoxygenated water, submerge

quart Mason jars in boiling water and cap under water.

The jars can then be stored

a t 27 C . ( 80 OF.).

When an egg paper is to be used, the jar is uncapped and the

egg paper is immediately placed in the w ater and the cap replaced.

Eclosion usually

begins in minutes.

Some egg papers can be used over again as all the eggs do not

always hatch after the first immersion.

Tho irgh it is possible to count aedine eggs on the egg paper, for mass rearing

purposes this is not feasible.

Instead an aliquot sam pling m ethod is used on the

newly hatched larvae.

References-

Burgess (1959); Elzinga (1961); Fay and Perry (1965); Giglioli (1947); Gjullin et

al. (1941);

Horsfall (1958); J d

son (I 960) ; Jupp and Brown (1967) ; Keppler et al. (1964) ; Krishnan

(I 964) ; Laurence (I 960) ; Lavoipierre (1953) ; Reynolds (I 960) ; Shute (1933) ; Takahashi

(1968); Trembley (1955); Williams (1962); Yoeli and BonC (1967).

LARVAL REARING TECHNIQUES AND EQUIPMENT

Counting Lmuae

1

Newly hatched larvae may be counted individually using a hand lens and a hand

counter.

For large rearing trays or mass rearing purposes, the aliquot sample method,

Fig. 13, (Gerberg et OZ. 196 8) is useful. Pour the first instar larvae into a gallon

battery jar and fill to 3500 ml of 27O C. (80 F.) water. The battery jar has 5

random ly placed outlets (at 5 levels).

A variable speed stirrer, mo unted on a ring

stand, is immersed in the water.

The stirrer is operated at approximately 250 rpm,

to suspend the larvae

uniformly in the water.

Twenty-five samples of I ml each are

withdrawn through the random outlets and the mean number of larvae per sample

(ml) is determined.

It is then possible to withdraw a given qu antity of water

containing a known number of larvae.

Morlan et al. ( 1963)

d

escribe and figure an autom atic dispenser for obtaining

18


29/124

Fig. r3.-Aliquot container for mosquito larvae.

large volumes of dispersed mosquito larvae. This equipment is made from a modi-

fied agitator type washing machine and a rain gaug e type of tripping bucket.

Bar-Zeev (19 62) describes a simple technique for obtaining standard numbers of

newly hatched mosquito larvae.

Rearing Trays

A wide variety of containers capable of holding water may be used as rearing trays.

Trembley (1955) suggests white enam el photographic developing trays 16xgf/2x2,


30/124

or containers of 3 quart capacity, 2-3 inches deep, preferably, but not necessarily,

opaque . Containers of these types are shown in Figs.

14

and 15. For mass rearing

of A&es aegypti, Morlan et al. (1963)

used a galvanized metal tra y 2x10~ 72.

(Fig. 16). Th

e ra was coated with a thin layer of paraffin. Each tray had a j/4

y

copper-tube outlet to facilitate draining and was stacked on a metal rack mad e of

i/sx 3/4

3/4 I1

angle iron. Each

rack held 24 rearing trays. They used 7 liters of

water in each tray to rear 7000 larvae. This system provides for 1.4 larvae/sq. cm.

of surface area and 1.4 larvae/ml of water. Fay et al. (1963) reared

8000-14,000

larvae per tray (2~10x 72) with no significant difference of percentage of yield or

adult vitality.

In the IC R laboratories, mosquitoes are mass reared in paraffin-coated, galvanized

trays r38x7 6x5cm (54x20x2), (Fig. 17). Each tray is filled to a depth of 2cm

( g) with water and approximately 15,000 larvae are introduced. Gahan (1967)

suggests 100 -250 Amp/&es larvae for a polyethylene pan 12 in diameter.

For small scale rearings, white polyethylene pan s 13I/4x1ox2 have been used very

successfully.

Handling Larvae

If larvae have to be moved or transferred, Trem bley (195 5) suggests a 50 ml.

volumetric transfer pipette shortened to 8-10 and fitted with a rubber bulb. We

have found very useful a 12-18

length of rigid clear plastic tubing covered with a

FIG. Iq.-White enamel trays for rearing larvae.

20


31/124

FIG.

IS.-White enamel dishes for rearing larvae.

LARVAL REARING TRAY

28 GA. GALVANIZED

SHEET METAL

314. LCl 1.062 WALL)

COPPER TUBING -SILVER

SOLDER TUBING FLUsn

RIVET AT EACH CORNER

ROTE: MAKE ALL SEANS WATER TIGHT.

FIG. 16.-Construction of galvanized metal rearing tray.

21


32/124

FIG. IT.-Mass rearing of mosqu itoes in galvanized trays.

fine mesh cloth at one end and fitted into a rubber bulb. Also useful for picking

up larvae or pupae is a piece of 20-40 mesh stainless steel screening 27x75m m

(1x3)

and bent so that approximately

I

inch is at a 45 angle to the remaining 2

Equipment for Feeding Larvae

Food

can be premeasured into small containers and then dispensed over the water

surface.

Some technicians prefer a salt shaker and can judge the amount of food

22


33/124

by the number of shakes. Heal and Peregrin (1945) recommend sheet metal spoons.

Hunt and DaveY (1947) used reversed pen nibs. Morlan et al. (1963) used a

plastic spoon of appropriate capacity. Regardless of the method of measurement, it

is imperative that a rigid feeding schedule be followed and the type and amount of

food be standardized if uniform and consistent results are desired.

Refevences-

Bar-Zeev (1957 , 196 2); Fay et al. (1963); Galun (1967); Gerberg et al. (1968); Heal and

Peregrin (1945); Long and Breland (1956); Morlan et al. (1963); Soderstrom and Levitt

(1967); Trembley (1955).

LARVAL FOOD AND FEEDING

Mosquito larvae are fed everything from a highly refined chemical diet to a witchs

brew made from guinea pig droppings, mud, and the ground-up skins of water

boatmen. Perhaps one of the greatest reasons for variability in the results of rearing

mosquitoes is the lack of uniformity or standardization of larval food. An abundance

of literature refers to larval foods, both natural and chemically defined. Asahina

(1964)

provides an excellent review of food material and feeding procedures for

mosquito larvae.

Some of the early investigators (Bacot 1916) report that bacteria and yeasts are

essential items of diet.

MacGregor

(1915)

increased the bacterial and protozoan

population by the addition of guinea pig feces. Boyd et al. (1932) used hay infusion

to provide protozoan food for the larvae. Weyer (1934) added dried blood and

powdered calfs liver.

Crowell (1940) was probably the first to use commercial dog

biscuits as a complete and somewhat standardized larval food. Perhaps the most

widely used food and feeding schedule is the one developed at the Communicable

Disease Center (Morlan et al. 1963). The larvae are fed dog chow (5% crude fat)

ground to pass through a 40 mesh screen.

This food is primarily for A&es aegyptz

but will work for other speciesas well, with the following schedule:

Day o (day of hatching) o.zmg/larva

I

0.3mgJarva

2

o.qmg/larva

3-7

o.6mg/larva

Note that the addition of a live yeast mixture (0.7 gm/I liter of water/Io,ooo

larvae) on day 3 improves adult production.

Trembley (1955)

used the following procedure for Anopheles quadCmacuZatus,

using a 16x9 l/*x2

tray for 300 larvae.

12 hours before day o

25 mg powered dried brewers yeast

and 25 mg Bacto-brain heart infusion.

Day o

IOO

mg of

I :I

powdered yeast and

powdered dog food

Day

I

50 mg

Day 2-3

100 mg

Day 4-14

150 mg

Gerberg et al. (1968)

use the following schedule for Anopheles stephensi.

23


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The larval food consists of a 50:5 0 mixture of dog chow a nd pig liver pow der

ground to 40 mesh.

Day o-Day 2 .I 3 mg/larva

Day 3 .20 mg + .045 mg Fleischman ns

dried brewers yeast/larva

Day 4 .26 mg/larva

Day 5 .40 mg/larva

Day 6-9 .53 mg/larva

The nutritional requiremen ts of mos quito larvae were investigated by Trag er

(1935, 1937, 1953), de M 11

l on et al. ( 1945), Goldberg ( 1947), Goldberg and de

Meillon ( 1948), Lea et al. ( 1956), Lea and DeLong ( 1956), Singh and Brown

(1957) and Akov (1962).

References-

Akov (1962);

Asahina

(1964); Racot (1917);

Boyd et al. (1932): Buddington (1941);

Crowell (1940) ; de Meillon et al. (1945) ; Frost et al. (1936) ; Gerberg ez ai. (1968) : Golberg

(1947); Goldberg and de Meillon (1948); Hinman (1930, 1932, 1933); Lea et al. (1956);

Lea and DeLong (1956); MacGregor (1915); Morlan et al. (1963); Singh and Brown (1957);

Trager (1935, 1937, 1942, 1953); TrembleY (1955); WeYer (1934).

HANDLING , COUN TING, SEPARATING AND SEXING PUPAE

Pupae may be picked up by means of pipettes or by small lifters made of screening.

The comm ercial m edicine dropp er with the open end filed off to ma ke a wide m outh,

or Kom ps modification of a syringe (Trem bley 195 5) may be used. Plastic pipettes

have become popular.

The rubber bulb normally supplies adequate suction but

others have relied on the use of vacuum pumps or water-operated suction pumps

(Johnson 194 7) to increase the efficiency of operation.

Lifters can be mad e from small pieces of metal screening, 20-4 0 mesh, 25x75c m

(1x3). Individual pupae can be lifted by means of small wire loops.

Pupae may be counted by manually picking them out with a pipette or lifter.

They can also be counted vo lumetrically.

A volumetric counter can be made from

a piece of plastic tubing with an ID of 3/s-2, approxim ately 6 long with one

end covered with fine wire screening. A know n num ber of one size pupae are added

to the tube and the level of pupae is marked on the tube. From then on pupae

can be added to the m ark, the tube inverted and the pupae washed out by flushing.

Pup ae can be separated rapidly from larvae by the ice water techniques of Ram a-

krishnan et QZ. 1963), Weathersby ( 1963 ) and Hazard ( 1967). Larvae and pupae

are concentrated in a sieve and im mersed in ice water.

The larvae sink immediately

and the pupae float and are poured off.

Bar-Zeev an d Ga lun ( 1961 ) used a rather

unique method of separating larvae from pupae by magnetic means.

Iron dust is

added to the rearing trays and the culture exposed to a magnetic field. Only the

larvae, which have ingested the iron p articles, are held b ack by the mag netic field.

Fay and Morlan (1959) describe a mechanical device not only for separating larvae

from pupa e, but ca pable of sexing the pupae of some species.

The principle of sepa-

ration and sexing is based on size differences.

The culture of mixed larvae and

pupae is

polured through a funnel into a wedge-shaped space between two slightly

separated and adjustable glass plates.

The female pupae, being largest are retained

24


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in a line at the upper levels of the plates, the smaller male pupae are held below

the females and the even smaller larvae are retained towards the bottom. The

smallest larvae drain out. The authors claim the separation of IOOO pupae in 20

minutes. Grose et al. ( 1966) made improvements on Fay and Morlans design.

McCray (1961) d escribes a simple separating device made from an aluminum

sheet with stamped-out 0.039 slits. The larvae and male pupae wash through

and the female are retained. The author claims that 30,000 individuals can be

processed n 5 minutes. Gentry and Gerberg are developing a small separating and

sexing device that uses removable comb-like plates in a trough arrangement. The

plates are interchangeable and the proper size plate is inserted depending upon the

species to be separated and the size of the pupae and larvae.

The sorted pupae are placed in containers of convenient sizes and the containers

placed inside cages to await adult emergence. Various devices have been used to

aid the emergence of adults. Some workers use small floating objects, such as small

pieces of cork, (Trembley 1955, Christophers 1960) or twigs to assist the adults in

leaving the water.

Smith and Whitlaw (1963) d escribe an emergence disc of wide

mesh dacron cloth that is claimed to reduce the mortality of emerging adults.

References-

Bar-Zeev and Galun (1961); Christophers (1960); Fay and Morlan (1959); Gillett (1955) ;

Grose et al. (I 966) ; Jones (1957) ; Lowrie and Gubler (I 968) ;Hazard (I 967) ; Johnson (I 947)

McCray (1961); Ramakrishnan et

al.

(1964): Smith and Whitlaw (1963); Trem bley (1955);

Weathersby (1963).

STERILE CULTURE

Rearing of mosquitoes under aseptic conditions may be required for

mosquito

nutrition investigations for in vitro studies, and other types of research.

Eggs may be surface sterilized.

Techniques described by Atkin and Bacot (1917)

use 0.5% lysol; Barber (1927)

dripped 80%

alcohol for 2-3 minutes over A.

mgypti

eggsand Roubaud and Colas-Belcour (1929) surface sterilized eggs using

hydrogen

peroxide

and also mercuric chloride. Whites (193 I) solution, consisting of mercuric

chloride 0.25 gm, sodium chloride 6.5 gm, hydrochloric acid, 125 ml, ethyl alcohol

.

250 ml and distilled water 750 ml, was more effective against bacterial contamination

than against other types of contaminants.

Hinman (1932) disinfected eggs in hexyl-

resorcinol.

Trager (1935, 1937)

used 5% castile soap and then Whites solution.

Lea et ul. ( 1956)

and Jones and DeLong (1961) surface sterilized eggs by

washing

the eggs in

70% alcohol for 5 minutes, transferring eggs by means of flamed forceps

to a 0.87% sodium hypochlorite solution for 2 minutes, then to a second 70% alcohol

rinse for 4 minutes,

and then to a final wash in autoclaved distilled water. Mar-

tignoni and

Milstead (1960) sterilized A. aegypti eggs on paper by placing one-

month-old eggs in a petri dish containing an autoclaved IO:/, solution of benzalko-

nium chloride for 30-60 minutes,

then washing in sterile water, followed with

immersion for

15-30

minutes in 80/~ alcohol and then two more washings in sterile

water. Akov (1962) describes a method of obtaining sterile larvae of uniform age

by placing

the sterile eggs in a desiccator and reducing the pressure for 10-20 minutes.

Trager (1935, 1937); de Meillon et al. (I945), Lea et al. (I956), Lea and DeLong

(1956), Nayar (1966)~ Singh

and Brown (I957), Akov (1962) reared larvae on a

sterile chemical media.


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Pupae can be washed to surface sterilize them, and adults may be fed on sterile

diets.

Thou gh it may be superfluous to mention, it is important to see that all wo rk in

rearing aseptic anim als is condu cted under com pletely aseptic cond itions.

References-

Akov (1962, 1964) ; Atkin and Bacot (1917) ; Barber (1927) ; Boorman (1967) ; Butt and Keller

(1961); Hinman (1930, 1932); Jonesand DeLong (1961); Lea et al. (1956); Lea and DeLong

(1956); de M ei on1 et al. (1945)

;

Lichtenstein (1948)

;

Martignoni and Milstead (1960);

Nayar (1966)

;

Roubaud and Colas-Belcour (1929)

;

Singh, K. R. P. and Brown (1957) ;

Trager (1935,

1937);

White (1931).

26


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III. PROCEDURES FOR LABORATORY REARING OF

SPECIFIC MOSQUITOES

Information gleaned from literature has been compiled to present a rearing tech-

nique for individual species,or in some casesa genus.

Usually, for each genus there

is a detailed description for at least one species hat could serve as a guide for rearing

other members of the genus.

The arrangement of genera and nomenclature is based on Stone, Knight and Starke

(1959).

SYSTBMATIC

ARRANGEMENT OF THOSE GENERA IN

ARE AVAILABLE

Subfamily Anophelinae

Genus

Anopheles

Subfamily Toxorhynchitinae

Genus Toxorhynchites

Subfamily Culicinae

Tribe Sabethini

Genus

Thchoprosopon

Genus

Wyeomyia

Genus

Sabethes

Tribe Culicini

Genus

Coquiltettidia

Genus

Mansonia

Genus

Uranotaenia

Genus

Ohopodomyia

Genus

Psorophora

Genus

Eretmapodites

Genus

Aedes

Genus

Armigeres

Genus

Haemagogus

Genus Opifex

Genus

Culiseta

Genus Culex

Genus

Deinocerites

ANOPHELES

WHICH REARING TECHNIQUES

Eggs may be collected in a small container, usually r5cm (6) in diameter or

15xrox5cm

(6x4~2).

Cotton balls, or an absorbent cotton pad, are laid inside the

container and then barely covered with water.

Filter paper or paper toweling is

then laid over the cotton pads so that the water surface is even with the surface of


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coarse-ground mixture of equal parts of Kelloggs concentrate, wheat germ and live

yeast. At the 2nd instar they are transferred to clean pans with fresh water and food

(Keppler et al. 1964). Gilotra ( 1966)

reared 200-250 larvae in enamel pans

I I .5x7.5x2

containing I l/2 liters dechlorinated tap water, and fed on powdered

Purina laboratory chow supplemented with Fleischmanns yeast. The larval period

usually lasts

7-14

days when reared at 27 C. (80 F.).

Pupae

The

1936).

Ad&S

pupae are collected once a day and placed

The pupal stage lasts 30-33 hours.

in cups of fresh water (Rozeboom

The adults are maintained in cages goxgoxgocm (2.5x2.5x2.5. They are fed on a

5% honey solution or sugar solution. The females are provided a blood meal at

dusk. A rabbit is placed in the cage for 2 hours. The best temperature appears to

be 28'

C.

(SIF.) (D

owns and Arizmendi 1951). Adults are also maintained in

cages 6ox6ox6ocm

(2x2x2'),

and are fed on sugar water, and various fruits. The

females are provided with a blood meal (human arm) once a day. Cages r2xgxg

maintained at 65% RH are also used for adults. A guinea pig provides a blood meal

and adults are maintained on honey-saturated balls of cotton wool (Keppler et al.

1964). Adult longevity is approximately 30 days.

References-

Coluzzi (1964) ; Dow ns and Arlzmendi (1951); Gilotra (1966) ; Ikpplcr rt 4. (1964) ; Roze-

boom (1936).

Anopheles albitarsis

Lynch Arribalzaga

Referrnces-

Barrett0 and Coutinho (1943); Galvao and Grieco (~9~3) JGalvao et nl. (1944).

Anopheles at-gy&arsis

Robineau-Desvoidy

Refeyences-

Galvao et al.

(1944).

Anopheles axtecus Hoff mann

See methods used for A. quadrimaculatus and A. freeborni.

References-

Downs et

al. (1948);

Dow ns and Arizmendi ( 1 g5 I) ;

Wallis (I 955 ).

Anopheles ba.Zabacensi.s

Baisas

Eggs are collected on filter paper discs kept in contact with moist cotton pads.

The mean number of eggs per female is 163.

Eggs are removed from the filter

paper discs to enamel trays containing 2,500 ml of seasoned tap water over a layer

of sterilized stream sand.

The eggs hatch in 48-72 hours.


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Anopheles crucians Wiedemann

See A. quadrimacuZatus techniques.

References-

Boy1 (I

926) ; Boyd et al. (1935).

Anopheles culicifacies

Giles

Reference-

Jayewickreme ( 1952)

;

Pal ( 1945).

Anopheles darlingi Root

Eggs are collected on moistened filter paper (Freire and Faria 1947) or in white

glazed fingerbowls (diam.

I I

cm, depth 7 cm) half filled with water. The eggs

are allowed to hatch in these bowls. Eggs hatch in 24-48 hours at 27 C. (80 F.).

Larvae

The larvae are reared in containers 49x7oxIocm or in white enamel basins (28 cm

in diam, and 8 cm deep). App

roximately

IOO

larvae are placed in the basin.

The

larvae are fed on an infusion made by mixing toasted fish and toasted bread in pro-

portions of I :2 plus a trace of brewers yeast. This mixture is suspended in water

4-5 days at 27O C. (80 F.), and should have a pH of 7.o-7.8. Giglioli (1947)

feeds larvae on dried brewers yeast or poultry laying mash or combination of both.

No special attempt is made to regulate light, temperature (70-M F.) or relative

humidity (66-95% RH). Th

e water used is clean, fresh, with a pH of

larval stage ranges from 6-12 days.

6-7. The

Pupae

The pupal period is approximately 2 days. The pupae are collected

placed in containers in the adult cage.

daily and

Ad&s

The adults are kept in cages 4ox4ox5ocm in an insectary maintained at

and 80-90~/~ RH. The time required from egg to imago is IO days. The

__ _

26-2a" c.

adults are

fed hony on cotton pads.

Adult longevity is 35-45 days. A cage 3 m high with

a 1x1-m base is required for mating (Freire and Faria 1947).

Giglioli ( 1947) main-

tains adults in a cylindrical cage 60 cm high and 35 cm in diameter, consisting of F

cotton mosquito gauze sleeve stretched over a IO gauge wire frame. The adults

are supplied with cotton wads soaked in cane sugar solution and changed daily.

Blood meals (human) are provided daily for l/2 hour in the morning. A tempera-

ture of 80 F. and a relative humidity of 85% seems optimal.

The first oviposition

usually occurs 6-8 days after emergence.

References-

Bates (I

947) ;

Freire and Faria (I

947) ;

Giglioli (I 947).

31


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43/124

Larvae

The larval stage lasts from g-10 days.

Pupae

The pupal stage lasts for

I-I%

days.

Adults

Mating will occur in an insectary.

A minimum of

2

days after emergence is

required by adults before mating.

Eggs are laid

2

days after a blood meal. Females

will feed on man, and animals. Adults feed on fruit juices. Females will live for

51 days at 85% RH when provided with blood meals.

References-

J&m (1945); Perry (1946).

Anopheles fluuiatilis James

-

Eggs are collected in earthen pots, lined inside with mud and containing water.

The eggs are transferred to large enamel basins and dishes containing water. The

eggs are floated inside paraffined cork rings.

Larvae

The larvae are fed on hay infusion, plus a small quantity of a mixture of 2 parts

litmus milk and

I

part dehydrated blood serum.

Dried brewers yeast may be sub-

stituted for the above.

The water in the basin is aerated vigorously once each day.

Relatively larger amounts of yeast are added to the rearing basins when the larvae

reach the fourth instar.

Pupae

The pupae are removed from the larval rearing basins, placed in a bowl of clean

water and placed inside the colony cage.

A few blades of grass are floated in the

bowl to provide a foothold for the emerging adults.

Adults

The colony cage measures

2x2~2

and is placed inside a larger cage with solid

wooden sides and glass top.

Humidity is maintained by suspending pieces of cloth

in a saturated solution of common salt. Adults are fed on

IO~

glucose on cotton

wool. A rabbit provides nightly blood meals. Mating takes place only in the

presence of a blue

light.

Egg laying occurs 48-72 hours after a blood meal. Len-

gevity is

17-18 days at 27 C. and 60-80*/~ RH.

References-

Mohan (1945); Pal (1943); Sin& and M&an (1951); Viswanathan et al. (1944).

Anopkeles ft-eeborni Aitken

Qxs

Eggs are collected in a glass crystallizing dish containing distilled water.

Approxi-

mately 100-200 eggs are laid per female. The eggs hatch in 3 days at 220 C.


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The larvae are reared in enameled iron pans 25cm (IO) in diameter in water

about r2.5-17mm (s-3/4) deep,

(A

rmstrong and Bransby-Williams

1961).

The

water is boiled, cooled stream water.

Approximately 150 larvae are added to each

pan. Powdered meat meal is provided as food. Powdered blood and yeast suspen-

sion used by Shute (1956)

seemed to produce heavy scum.

Wegesa ( 1964) com-

pares meat meal and yeast and finds adult emergence and survival rates significantly

increased by using meat meal.

A fine mesh tea strainer is used for applying the

food.

Larvae are reared in an enamel tray 35x3ox5cm at 26.5 C. and require

7-13

days to pupate.

Two hundred newly hatched larvae are placed in 2 liters of distilled

water. They are fed

0.2

g meat meal on alternate days.

Pupae

Pupae are collected daily and 300 placed in a 7.5cm (3) diameter aluminum pot

with a minimum of larval water.

Distilled water is added to bring the water level

within

r2.5-r7mm (l/2-3/4) of the top. Pupae are maintained at 26.5 C. & 0.5 OC.

Adult emergence is complete in about

2

days.

Eight hundred pupae are placed

in a cage.

Adults

Adults are kept for the first 7 days in cages 3ox3ox3ocm

(1x1~1') and then

transferred to larger stock cages 75x38x4ocm (30x15~16). The adults are main-

tained in an insulated, temperature controlled (26.5 C.) (70-90% RH) darkened

room.

The adults are maintained on sugar solution or corn syrup on cotton wool and

are offered a blood meal on the day after emergence, and on alternate days after

that. Rabbits are used as a source of blood meals. The back of the rat&it

is shaved

and the unanesthetized rabbit is placed in a box in the cage. Mating occurs at dusk

or in a darkened room or cage.

Maximum mating occurs 3-5 days after emergence

(Shute

1956).

Fertilized females take a blood meal more readily

than unfertilized

females.

Approximately 15.6/, survive 20 days (Wegesa 1964).

References-

Armstrong and Bransby-Williams (1961); Causey et al. (1943); Coluzzi (1964); Gillies

(1961) ; Goma (1959)

;

Haddow and Ssenkubuge (1962)

;

Jones,M. D. R. et al. (I

967)

; Mathis,

C. (1936); Mathis, M. (1935); Moores (1953); Muirhead-Thompson (1948); Philip (1930);

Shute, G.

T.

(1956); Wall (1953); Wegesa (1964).

Anopheles kymanus

Pallas

Anopheles jamesii Theobald

-

Eggs

are

collected in petri dishes 1% in diameter.

Larvae

Larvae are reared on a medium (pH 7.1)

consistingof mud water and tap \lrater.

Larvae are fed on brown bread powder.

Pupae

Pupation occurs IO days after oviposition.


46/124

Adz&s

The adults are maintained in Barraud cages 6x6~6. They are fed water-soaked,

split raisins, placed on top of the cage and covered with a pad of dam p cotton w ool.

The adults emerge in 24 hours.

Reference-

Jayewickreme1952).

Anopheles labranchiae Falleroni

Anopheles labranchiae subsp.

atroparvus

Van Thiel

Eggs are collected on damp filter paper. The filter pape r is placed on wet cotton

in small plastic cups or in petri dishes. Eggs are left on damp filter p aper for at least

24 hours but not more than 72 before hatching. The eggs are removed with the

filter paper for hatching purposes.

Larvae

Approximately 300-40 0 larvae are placed in a white enamel or polyethylene bowl

34cm (14) in diameter, containing water to a depth of approximately 3-4 cm (I s").

The water temperature should be about 27 C. (80 F.). Mu d or a piece of grass

turf is used in add ition to artificial food. The use of these variables leads to non-

standardized mosquitoes.

Ground dog biscuits, grain foods and liver powder are

used successfully.

Pupae

Pupae are usually hand picked though mechanical means of separation may be

used.

Adults

One thousand or more adults are maintained in a 2x2~2' cage. The pup ae are

placed in the cage in sm all containers and the containers covered or rem oved after

adult emergence.

The adults are maintained on 10% sucrose soaked cotton wool or

cotton balls. Restrained guinea pigs are laid on top of a cage, or a human arm is

inserted into the cage to provide a blood m eal.

References-

DpAlessandrot al. (1961); Bertram and Gordon (1939); Coluzzi (1964); Meller (1962);

Shute,P. G. (19 36).

Anopheles maculatus Theobald

Each female lays about 80-100 eggs on moist filter paper, in individual tubes or

in paper cups,

(Ow

Yang

et al. 1963).

At the ICR laboratories the fertilized females

are placed in 1~1x1

alum inum screened cages and lay eggs in plastic containers.

Larvae

Jayewickreme (1952) reared larvae in a medium consisting of 20 parts hay infu-

36


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sion to 80 parts tap water.

75 cc of hay infusion was added daily. Larvae were

reared in an earthenware dish with a diameter of 20 cm and a surface of 314.2 sq cm

containing 500 cc of media.

Larvae were fed on the brown bread powder after the

4th day.

Pupae

Maximum pupation occurred on the 13th day after oviposition.

Adults

/

Maximum adult emergence occurs on the 15th day after oviposition. The adults

are maintained in Barraud cages 6x6~6 and are fed water-soaked raisins placed on

top of the cage and covered with a pad of damp cotton wool.

Jayewickreme (1952)

does not mention mating or egg production. Ow Yang et

al.

(1963) were able to

maintain a colony by artificial mating.

Sexed pupae (by size) emerged in separate

cages,

1~1x1

They were maintained at 27 C. and 70-90% RH and fed on 5%

glucose solution. Two days after emergence the glucose was removed and a guinea

pig was placed in the cage overnight. The following morning engorged females were

collected in individual tubes.

Males 3-6 days old were used for the artificial mating

(see techniques section).

References-

Jayewickreme

(1952); Ow

Yang etd. (1963).

Anopheles pharoensis Theobald

Eggs are collected in petri dishes containing water. The embryonic period is 2-4

days, depending upon temperature. The percentage of egg fertility varies from

35-95% with an emergence of 70% (Theodor and Parsons 1945).

Larvae

The optimum density of larvae is roe-150 first stage larvae and 50 third or fourth

stage larvae in a bowl of 15 cm diameter.

The rearing technique of Theodor and

ParSOnS 1945)

consistsof placing IOO first stage larvae into the white enamel bowl

containing a layer of mud I cm in depth and about 4 cm of water. The larvae are

daily fed a few drops of bakers yeast in water.

At the second stage, they are fed

powdered biscuits,

sprinkled on the surface of the water-sometimes 3 or 4 times

a day. The larval period is I 1-12 days at 28-29 C. At 23 C. the larval period

is 23 days and at 27O C. about 18 days.

The pupae are removed every morning and placed in a bowl inside the cage.

The

pupal period lasts 3-4 days at

20~

C.

The adults are maintained in a large cage 6ox6oxIoocm.

Humidity is maintained

by covering the sides and front of the cage with a moist blanket. The adults are

fed a sugar solution on a soaked cotton pad or sponge. A rabbit was first used for

blood meals, but a marked reluctance to feed was observed. Human blood is

readily

37


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acceptable. Swarming and

white light is turned on.

mating takes

place in blue light and

ceases

when a

References-

Abdel-Malek et al. (1966)

;

de Meillon et al. (1963); Theodor and Parsons (1945).

Anopheles pseudopunctipennis Theobald

References-

Downs and Arizmendi

(1951);

Hardman

1947).

Anopheles punctipennis (Say)

Reference-

Hardm an (I 947).

Anopheles punctulatus Donitz

References-

Backhouse

(1937);

MacKerras and Lemerle

(1949).

Anopheles quadrimaculatus Say

-

Eggs are collected in a pan or bow l of water, or on damp filter paper or toweling.

Avoid desiccation of the eggs. Gah an (1967 ) states that whenever a surplus of eggs

is obtained, the excess supply may be wrapped in damp filter paper o r toweling and

may be stored in a refrigerator for IO days to 2 weeks. Eggs are usually deposited

at night. Rep lace the egg collecting container daily. Eggs are concentrated and

introduced to larval rearing trays or the eggs are left to hatch and then co unted by

the aliquot method and placed in the rearing trays. The eggs hatch in 30-40 hours.

Larvae

Trays of various sizes

and composition are used.

White enamel

(14x10~4")

or white

plastic trays are used with su

rearing mosquitoes

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