natural breeding places of phlebotomine sandflies

REV IEW ART ICLE

Natural breeding places of phlebotomine sandflies

M. D. FELICIANGELIUniversidad de Carabobo, Facultad de Ciencias de la Salud, Centro Nacional de Referencia de Flebotomos,

BIOMED, Nucleo Aragua, Maracay, Venezuela

Abstract. Methods of finding larvae and pupae of phlebotomine sandflies(Diptera: Psychodidae) are described and the known types of breeding sites usedby sandflies are listed. Three ways of detecting sandfly breeding places are the useof emergence traps placed over potential sources to catch newly emerged adultsandflies; flotation of larvae and pupae from soil, etc., and desiccation of media todrive out the larvae. Even so, remarkably little information is available on theecology of the developmental stages of sandflies, despite their importance as vectorsof Leishmania, Bartonella and phleboviruses affecting humans and other vertebratesin warmers parts of the world. Regarding the proven or suspected vectors ofleishmaniases, information on breeding sites is available for only 15 out of 29 speciesof sandflies involved in the Old World and 12 out of 44 species of sandflies involvedin the Americas, representing �3% of the known species of Phlebotominae.Ecotopes occupied by immature phlebotomines are usually organically rich

moist soils, such as the rain forest floor (Lutzomyia intermedia, Lu. umbratilis,Lu. whitmani in the Amazon; Lu. gomezi, Lu. panamensis, Lu. trapidoi in Panama),or contaminated soil of animal shelters (Lu. longipalpis s.l. in South America,Phlebotomus argentipes in India; P. chinensis in China; P. ariasi, P. perfiliewi,P. perniciosus in Europe). Developmental stages of some species (P. langeroni andP. martini in Africa; P. papatasi in Eurasia; Lu. longipalpis s.l. in South America),have been found in a wide range of ecotopes, and many species of sandflies employrodent burrows as breeding sites, although the importance of this niche is unclear.Larvae of some phlebotomines have been found in what appear to be specializedniches such as Lu. ovallesi on buttress roots of trees in Panama; P. celiae in termitehills in Kenya; P. longipes and P. pedifer in caves and among rocks in East Africa.Old World species found as immatures in the earthen floor of human habitationsincludeP. argentipes, P. chinensis, P. martini andP. papatasi. Muchmore informationon sandfly breeding sites is required to facilitate their control by source reduction.

Key words. Leishmania, Lutzomyia, Phlebotomus, Sergentomyia, emergencetraps, flotation methods, leishmaniasis vectors, sandfly breeding sites, sandflyecology, source reduction, vector control.

Introduction

Since the review by Killick-Kendrick (1990), important

advances have been made in understanding the biology of

phlebotomine sandflies and their vector roles for bartonel-

losis (Birtles, 2001), flaviviruses, orbiviruses, phleboviruses

and vesiculoviruses (Comer & Tesh, 1991; Ashford, 2001) as

well as leishmaniases (WHO, 1990; Killick-Kendrick, 1999),

but knowledge of sandfly breeding sites remains scanty.

Searching for developmental stages of sandflies in their

natural biotopes is difficult, tedious and has proved to be

remarkably unproductive (Deane & Deane, 1957; Killick-

Kendrick, 1987, 1999). The deficit of information on

sources of sandflies prevents us avoiding such sites and

disallows the targeting of control measures against the

Correspondence: Dr M. Dora Feliciangeli, Centro Nacional de

Referencia de Flebotomos (CNRFV), BIOMED, Facultad de

Ciencias de la Salud, Universidad de Carabobo, Apdo. 4873,

Nucleo Aragua, Maracay, Venezuela. E-mail: [email protected]

Medical and Veterinary Entomology (2004) 18, 71–80

# 2004 The Royal Entomological Society 71

preimaginal stages of sandflies. Hence the only feasible

countermeasures (Alexander & Maroli, 2003) depend on

adult sandfly control and personal protection.

The finding of a sandfly larva by Grassi (1907), in a cellar

in Rome, led to the description of a new species: Phleboto-

mus mascittiiGrassi 1908. This is regarded as the first report

of an immature stage of any phlebotomine sandfly in

nature. In the New World, the first findings of phlebotomine

breeding sites were at the base of a tree in Brazil, where

Ferreira et al. (1938) found four larvae, and Pifano (1941)

found a dozen larvae in a wall of a house in Venezuela.

This review gathers the scattered reports of methods used

effectively to detect sandfly breeding sites, or lack of them,

and summarizes the limited information available on the

places from which phlebotomine larvae and pupae have

been recovered. In particular, the known breeding sites of

proven or suspected vectors of Leishmanias in the Old

World (Table 1) and the NewWorld (Table 2) are described,

together with experiences of control efforts aimed at sandfly

immature stages.

Methods of searching for immature stages ofphlebotomines

Besides the occasional finding of sandfly larvae and pupae

in nature, four main methods have been employed to search

systematically for sandfly developmental stages in potential

ecotopes.

1 Direct visual searches of materials (e.g. fungi, leaf litter,

soils) sampled from potential habitats and examined

under a stereomicroscope.

2 Soil extraction, using flotation with saturated solutions

of sugar or salt, or desiccation to drive out the larvae.

3 Soil incubation, i.e. waiting for adult sandflies to emerge.

4 Use of emergence traps (cages) and sticky traps (oil

papers) over potential breeding sites of sandflies.

During the early decades of the 20th Century, in the quest

for sandfly breeding places, substantial efforts were made to

find phlebotomine larvae and pupae by directly searching

various types of natural habitats in several countries, i.e.

Malta (Marrett, 1910, 1913, 1915; Newstead, 1912; Witting-

ham & Rook, 1923), India (Howlett, 1913; Mitter, 1919) and

Sudan (King, 1913, 1914) as well as Italy. Although more

difficult and time-consuming than the other methods,

Dhiman et al. (1983) still preferred direct searches for recove-

ring immature stages of Phlebotomus argentipes from soil

litter in human dwellings and cattle sheds in Bihar, India.

The flotation method introduced in India by McCombie-

Young et al. (1926) was claimed to hasten the extraction of

sandfly larvae from soil, but required no less effort, nor

were striking results achieved. As an example of the low

but precious yield of phlebotomine immatures obtained by

flotation, Petrishcheva & Izyumskaya (1941) recovered 61

larvae and 91 pupae from 6 tons of soil processed in Sebas-

topol, Crimea, U.S.S.R.

Hanson (1961, 1968) modified McCombie-Young’s

method by using a combination of flotation with a satur-

ated sugar solution and washing the material through brass

gauze sieves (screening-flotation technique): this led to col-

lection of 2258 larvae of phlebotomines from the forest

floor in Panama during 4 years of intensive work. Unfortu-

nately, only 27% could be reared to the adult stage and

identified, mostly as Lutzomyia longipalpis (Table 2).

Soil extraction using the technique of MacFadyen (1961),

based on larval escape from desiccation with warming, was

employed by Seyedi Rashti & Nadim (1972) to recover

Phlebotomus papatasi larvae in Iran. Also with this method,

Killick-Kendrick (1987) processed 130 kg of negative sam-

ples from a goat cellar in Cevennes, France. Afterwards he

tested this technique in the laboratory, demonstrating its

efficiency for recovering good proportions of young larvae

from soil samples seeded with laboratory-reared phleboto-

mine larvae.

The soil incubation technique for rearing-out phleboto-

mine immatures to the adult stage for species identification

has been widely employed in China (Y.J. Leng, personal

communication). Mutinga & Kamau (1986) considered this

method to be better than any other for their searches in

Kenya.

A wide range of trap designs have been employed to

capture adult sandflies emerging from sites regarded as

suitable for development of immatures, i.e. the ‘armadilha’

(1.8m high� 1.2m wide) used by Deane & Deane (1957)

and ground photo-collectors covering 1m2 (Penny & Arias,

1982) in the Brazilian Amazon; traps of 0.5m2 area for

sampling of the forest floor by Rutledge & Ellenwood

(1975a) in Panama; small plastic bowls (0.1m2) of Bettini

et al. (1986), recently modified by Casanova (2001), and the

simple polyvinyl chloride (PVC) pipe and couplings of dif-

ferent sizes employed by Ferro et al. (1997) in Colombia, cut

so that the couplings would face available light. Emergence

traps allow estimates of population density from the

observed productivity of breeding sites, expressed as

adults/area/time (Southwood, 1966).

Despite controversies over soil ecology (Andre et al.,

2002), the use of radiography (Villani & Gould, 1986; Vil-

lani et al., 1989) to investigate the behaviour of scarab grubs

in soil may point to ways of observing radio-labelled phle-

botomine immatures under semi-natural conditions in their

breeding sites.

Breeding sites of phlebotomine sandflies

Hanson (1968) stated that the location of breeding sites of

phlebotomine sandflies was often merely assumed, because

of the proximity of adult resting places. By investigation, he

detected immature stages of phlebotomines in only 53% of

the suspected breeding places he explored. Killick-Kendrick

(1987) suggested that, as for other Diptera, sandflies would

not lay eggs indiscriminately: they must recognize suitable

sites for larval development. Subsequent papers on sandlfy

oviposition attractants/stimulants (El Naiem &Ward, 1992)

72 M. D. Feliciangeli

# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80

Table1.Studiesonbreedingsitesofproven

orsuspectedvectors

ofleishmaniasesin

theOld

World:speciesandhabitats

accordingto

Killick-K

endrick

(1999).Clinicalform

s:ACL,

anthroponoticcutaneous;CL,cutaneous;DCL,diffuse

cutaneous;PKDL,post

kala-azardermal;VL,visceral;ZCL,zoonoticcutaneous.Methods.DS,directsearching;ET,em

ergence

traps(number

ofstickypapers:nights);FS,flotationwithsugarsolution;SD,soildesiccation;SI,soilincubation;Sites:AB,anim

alburrow

(#Apodem

us;yrodent);AS,domesticanim

al

shelter;CG,cellarwithgoats;CS,cattle

shelter;GO,groundoutdoors;HH,humanhabitations;HS,horsestables;RD,refuse

dump;FR,floorofroom;PS,poultry

shed;SL,soiland

litter;SP,stonepile;SS,sheepshelter;TM,term

itehill;TR,tree-hole;various(ztree-holesandroots,stonecracks,latrine,walls;**cave,latrine,under

bridge).Habitats:D,domestic;E,

peridomestic;S,silvatic.References:ECB/C

AMS,EastChinaBranch

oftheChineseAcadem

yofMedicalSciences;ECKI,EastChinaKala-azarInstitute.No,number

obtained:A,adults;

P,pupae;

L,larvae;

þpresent.

Parasite

Clinical

form

(s)

Vectors

*proven

Country:area

Method

Noþ/samples(%

):

materialweight

Sites

Habitat

No.

flies

No.

spp.

References

Le.

donovani

VL,PKDL

P.argentipes*

India:Assam

FS

–HH,HS

D,E

þ–

Shorttet

al.,1930,1932;

Smithet

al.,1936

(inHanson,1961)

India

FS

4/75(5%)

CS

E1P

–Pandya&Niyogi,1980

India

DS

10/102(10%)

CS,HH

D,E

2A,6P,50L

3Dhim

anet

al.,1983

India

FS

15/131(12%)

HH

D,E

11L,8P

–Ghosh

&Bhattacharya1991

SI

38/157(24%)

CS

D,E

38A

–Ghosh

&Bhattacharya1991

ET

–CS

D,E

69A

–Ghosh

&Bhattacharya1991

VL

P.martini*

Kenya

SI

109/150(73%):1859kg

TM

S41A

17

Mutingaet

al.,1989

SI

80/114(70%):1602kg

AB

S19A

14

Mutingaet

al.,1989

SI

25/54(42%):230kg

HH

D5A

9Mutingaet

al.,1989

SI

25/44(57%):172kg

TR

S2A

10

Mutingaet

al.,1989

VL

P.celiae*

Kenya

SI

109/150(73%):1858kg

TM

S1A

–Mutingaet

al.,1989

SI

25/44(57%):172kg

TR

S1A

–Mutingaet

al.,1989

Le.

infantum

CL,VL

P.ariasi*

France:Cevennes

ET(120)

–RD

E5A

1Killick-K

endrick,1987

–CG

E1A

–Killick-K

endrick,1987

SD

1/6

(17%)

CG

E6L

–Killick-K

endrick,1987

DS

–AB#

S0

–Killick-K

endrick,1987

DS

77kg

SL

S0L

–Killick-K

endrick,1987

SD

130kg

SL

S0L

–Killick-K

endrick,1987

P.perfiliew

i*Italy:Tuscany

ET

–AS

E19A

2Pozioet

al.,1983

Italy:Sardinia

ET

18/25(72%)

SS

S83A

2Bettini,1989

Italy:Sardinia

ET

18/25(72%)

AS

S3731A

3Bettini,1989

P.langeroni*

Egypt:

ET(11)

–Various

E115A

–Dohaet

al.,1990

ElAgamy,Alexandria

–SP

39A

–Dohaet

al.,1990

–AS

23A

–Dohaet

al.,1990

–ABy

1A

–Dohaet

al.,1990

–RD

3A

–Dohaet

al.,1990

P.tobbi

Greece

–Wells

þ–

Biocca&Costantini,1986

P.chinensis

China:Jiangsu

DS

20%

FR

Dþ

–CAMS

1952(unpubl.)

Jiangsu

DS

10%

AS

Eþ

–ECB/C

AMS&ECKI1954

(unpubl.)

Shandong

DS

54/2808(2%)

Variousz

Eþ

–ECB/C

AMS&ECKI1954

(unpubl.)

Jianchang,Liaoning,

1/4

(25%)

ABy

Dþ

–Leng,1956(pers.com.)

Suizhong

–Nicolescu&Bilbie,1980

Breeding places of phlebotomine sandflies 73


Table

1.Continued.

Parasite

Clinical

form

(s)

Vectors

*proven

Country:area

Method

Noþ/samples(%

):

materialweight

Sites

Habitat

No.

flies

No.

spp.

References

Le.

tropica

ACL

P.sergenti*

Romania

ET

–Rocks

S1A

1Nicolescu&Bılbıe,1980

ZCL

P.guggisbergi*

Kenya

SI

–Caves

S0

–Mutinga,1996

Le.

major

ZCL

P.duboscqi*

Senegal

SI

1.3

kg

ABy

S4A

1Dedet

etal.,1982

Kenya

SI

80/114(70%):1602kg

ABy

E2A

17

Mutingaet

al.,1989

DCL,ZCL

P.papatasi*

India

FS

FR,GO

D,E

þ–

McC

ombie-Y

ounget

al.,1926

Assam

FS

191/423(45%)

CS,HH

E5L

–Smithet

al.,1936

India

FS

4/75(5%)

CS,HH

E5L

–Pandya&Niyogi,1980

India:Bilhar

DS

10/102(10%)

CS,HH

D,E

4L

3Dhim

anet

al.,1983

CentralAsia

ET

ABy

Eþ

–Perfil’ev,1968

CentralAsia

ET,FS

ABy ,CS

D,S

–Artem

ievet

al.,1972

Iran

SD

7/226(3%)

E–

SeyediRashti&Nadim

,1972

Egypt:Alexandria

ET(11)

Various

EA115

2Dohaet

al.,1990

SP

EA28

–Dohaet

al.,1990

AS

DA18

–Dohaet

al.,1990

ABy

EA2

–Dohaet

al.,1990

RP

EA1

–Dohaet

al.,1990

Egypt

ET

ABy

Eþ

–Morsyet

al.,1993

ET

PS

Eþ

–Morsyet

al.,1993

Le.

aethiopica

ZCL,DCL

P.longipes*

Ethiopia

FS

43.5

kg

Various**

E15L13P

2Foster,1972

P.pedifer*

Kenya

SI

Caves

E–

Mutinga&Odhiambo,1986



Table2.StudiesonbreedingsitesofLutzomyia

spp.,proven

orsuspectedvectors

ofleishmaniasesin

theNew

World(speciesaccordingto

Killick-K

endrick,1999).A,adultsandflies;CL,

cutaneousleishmaniasis;ET,em

ergence

traps(number

oftrap-nights);L,sandflylarvae;

MCL,muco-cutaneousleishmaniasis;VL,visceralleishmaniasis.

Parasite

Clinical

form

(s)

Vectors

*proven

Country:area

Method

Noþ/samples(%

):

materialweight

Sites:proportionþ

orrate

ofproduction

No.

flies

No.

spp.

References

Le.i.chagasi

VL

Lu.longipalpiss.l.*

Brazil:Ceara

Salt-flotation

12/241(5%)

anim

alcorrals

12A

1Deane&Deane,

1957

2/15(13%)

amongrocks

7A

3Deane&Deane,

1957

6/50(12%)

Colombia:Cundinamarca

ET(232)

Pig-pens

11A

3Ferro

etal.,1997

(5flies/100m

2)

ET(25)

tree

trunks

3A

3Ferro

etal.,1997

(1.63flies/100m

2)

ET(288)

rocks

1A

3Ferro

etal.,1997

(1.42flies/100m

2)

Le.

panamensis

CL,MCL

Lu.gomezi

PanamaCanalZone

Screen-flotation

88/370(24%)

forest

floor

600A,2258L

15

Hanson,1961,1968

Lu.panamensis

deadleaves

8A

Lu.ylephiletor

2A

Lu.trapidoi*

12A

Lu.trapidoi*

Directexam

forest

floor

39A

Lu.ovallesi*

buttressed

roots

24A

Hanson,1961,

1968;Rutledge

&Mosser,1972

Lu.trapidoi*

ET(2061)

leaflitter

(24.4

flies/100m

2/day)

307A

6Rutledge

&Ellenwood,1975b,c

Lu.panamensis

90A

Lu.gomezi

33A

Lu.ylephiletor

28A

CL,MCL

Lu.umbratilis*

Brazil:CentralAmazon

ET(243)

forest

floor

40A

15

Arias&Freitas,1982

(4.1-14s/100m

2/day)

Le.

guyanensis

CL,MCL

Lu.anduzei*


ET(243)

55A

Lu.paraensis


ET(243)

2A

Le.

braziliensis

CL,MCL

Lu.interm

edia

s.l.

Brazil:SaoPaulo

Soilincubation

1/1

pig-shed

12A

1Forattini,1954

Lu.interm

edia

s.l.

1/1

forest

floor

1A

1

Lu.pessoai

Soilincubation

1/1

forest

floor

1A

1

Lu.interm

edia

s.l.

Brazil:EspiritoSanto

ET

Soil

6A

3Vieiraet

al.,2000

Lu.whitmani*

Brazil:SaoPaolo

ET

forest

floor

7A

3Casanova,2001

Lu.interm

edia

s.l.

ET

forest

floor

39A

(32.6-38.7

s/100m

2/day)

Lu.interm

edia

s.l.

ET

forest

floor

25A

(24s/100m

2/day)

Lu.pessoai

ET

2A



produced by the accessory glands (Dougherty et al., 1992) and

the response of gravid females to compounds emitted by faeces

of chicken or rabbit (Dougherty et al., 1995), as well as to soil

bacteria isolated from natural breeding sites in India (Radjame

et al., 1997), support such a hypothesis.

Tables 1–3 list the sites in domestic, peridomestic and

silvatic habitats from which sandfly pre-imaginal stages

have sometimes been recovered. However, based on the

frequency of the collections and the abundance of speci-

mens caught, only a few of them might be considered as

stable breeding sites. Human dwellings and cattle sheds in

India and gerbil burrows in Central Asia (Perfil’ev, 1968)

may be regarded as stable breeding sites of Phlebotomus

papatasi. Developmental stages of P. duboscqi were

recovered from burrows of the giant African cane rat,

Cricetomys gambianus, by Dedet et al. (1982) and from

unidentified rodent burrows in Kenya (Mutinga et al.,

1986), Phlebotomus martini was found to breed mainly in

animal burrows, whereas termite hills were regarded as

secondary breeding sites in Kenya (Mutinga et al., 1989).

Research on the breeding sites of sandflyvectors of leishmaniases

Studies on potential breeding sites of proven and suspected

vectors of leishmaniases are summarized in Table 1 for the

Old World and Table 2 for the New World, in relation to

the Leishmania parasites they transmit, the locality, meth-

ods used and data on the work effort, if available. The

source of samples studied and the habitat (domestic, peri-

domestic or silvatic) are given. Results are expressed as the

proportions of positive samples among those processed, the

amount of material screened and/or the numbers of positive

and total emergence traps used. The number and stage(s) of

specimens recovered (larvae or pupae) and the number of

species found are also reported with the source reference.

For the known and suspected sandfly vectors of the

important leishmaniases (WHO, 1990), immature stages

have been recovered for at least 7 out of 10 proven vectors

of Leishmania donovani s.l and Le. infantum in the Old

World, as follows with their references: P. argentipes (Shortt

et al., 1930, 1936; Smith et al., 1936; Pandya & Niyogi, 1980;

Dhiman et al., 1983; Ghosh & Bhatthacharya, 1991),P. celiae

and P. martini (Mutinga & Kamau, 1986; Mutinga et al.,

1989), P. ariasi (Killick-Kendrick, 1987), P. perfilievi and

P. perniciosus (Pozio et al., 1980; Bettini et al., 1986; Bettini

& Melis, 1988) and P. langeroni (Doha et al., 1990).

Immature stages of P. tobbi, suspected vector of

Le. infantum around the eastern Mediterranean and Sicily,

were detected in wells on the island of Zakinthos, Greece

(Biocca & Costantini, 1986). According to Professor Yan

Jia Leng (personal communication), larvae and pupae of

P. chinensis have been detected in several ecotopes (Table 1)

during extensive work on leishmaniasis and its vectors by

the East China Branch of the Chinese Academy of Medical

Sciences (ECB/CAMS) and the East China Kala-azar

Institute (ECKI).

Only two species of sandfly, P. argentipes in India and

P. chinensis in China, have been encountered repeatedly in

soil samples in the earth floor of human dwellings and cow-

sheds, whereas only small proportions of the larvae and pupae

ofP. martini in Kenya have been found in the houses.Mutinga

&Kamau (1986) andMutinga et al. (1989) stated that themain

breeding sites of this species are animal burrows and termite

hills, which serve as incubators with well regulated internal

environments. Of the closely related P. celiae in Kenya, they

found only two males (from a tree-hole and a termite hill),

insufficient to reveal much about the breeding sites of this

species. In southern Ethiopia, however, Leishmania-infected

P. martini and P. celiae have been recovered from termite

hills, suggesting that termite galleries may provide an import-

ant ecotope for the epidemiology of Le. donovani, the aetio-

logical agent of kala-azar (Gebre-Michael & Lane, 1996).

Poor results were obtained from searching for the breed-

ing sites of P. ariasi in the Cevennes focus of Leishmania

infantum, causing canine and human visceral leishmaniasis

in France. However, Killick-Kendrick (1987) reached the

conclusion that the larval ecotopes of P. ariasi are probably

domestic and have a high content of organic matter.

Table 3. Summary of situations from which immature stages of

phlebotomine sandflies have been recovered

Habitat Ectotope

Domestic Abandoned buildings

Basements and cellars of houses

Cracks in mud floors and walls

Soil in human dwellings

Peridomestic Animal burrows

Animal shelters (cattle, pigs)

Caves

Chicken coops

Debris and soil cracks

Dry excreta of small domestic animals

Earth dyke

Embankments

Latrines

Rotted manure

Rubbish in the street

Soil at the base of old walls

Under stones

Wells

Silvatic Ant nests

Burrows of gerbil and other rodents

Burrows of other animals (unknown)

Caves

Cesspits, dry

Drains

Garbage

Hollow trees

Leaf litter on forest floor

Nests of terrestrial tortoises

Nests of birds

Rocks, between and under

Roots of large trees

Soil at base of trees

Soil under overhanging rocks

Termite hills



The findings of Bettini (1989) in Sardinia were, as he

wrote, ‘surely exceptional’. An abandoned cement structure

(area 25m2), used as a sheep shelter, was found to be

productive of three species of sandfly: Sergentomyia minuta

and two species of Phlebotomus (Bettini et al., 1986). After

sealing the shelter, they collected totals of 23 338 P. perfi-

liewi and 1309 P. perniciosus with six exit traps during 1983–

1985 (Bettini, 1989). Analysis of the substrate soil for tex-

ture, pH, CaCO3, organic matter and water content

showed no correlation with the number of sandflies that

emerged from the spots where soil samples were taken.

Apparently the developmental stages were associated

with a relatively stable, cool and humid environment pro-

tected from sunshine and rain, rich in clay and organic

nitrogen. Sampling from a similar site in Tuscany, however,

yielded very few specimens of these species (Pozio et al.,

1980). Immature stages of P. langeroni were found in Egypt

(Doha et al., 1990), mainly in rubbish from stone piles.

About the vectors of Le. tropica causing cutaneous

leishmaniasis, Mutinga & Odhiambo (1986) claimed that

P. guggisbergi breeds in Kenyan caves. In Romania, surveys

of potential sandfly biotopes at Dobrudja, using oil papers,

yielded only one P. sergenti (Nicolescu & Bılbıe, 1980),

although that species was formerly abundant in Canaraua

Fetii rocks (Duport et al., 1971). The decrease of P. sergenti

may be attributed to geological exploitation, spoiling the

sandfly breeding sites (Nicolescu & Bılbıe, 1980).

Regarding the most widespread form of cutaneous leish-

maniasis, caused by Leishmania major, the predominant

vector Phlebotomus papatasi is well known and apparently

not complex (Parvizi et al., 2003): information on breeding

sites of this sandfly is available from several countries

(Table 1). In India, immature stages of P. papatasi have

been recovered consistently from cattle sheds and human

dwellings in urban areas (McCombie-Young et al., 1926;

Smith et al., 1936; Pandya & Niyogi, 1980; Dhiman et al.,

1983). In rural areas they have been found in various

habitats: unused poultry houses made of bricks and clay,

manure heaps, caves, embankments, dried-up cesspits and

latrines (Sivagnaname & Amaldraj, 1997). In Egypt, breed-

ing sites of P. papatasi have been found in a similar range of

ecotopes (Artemiev et al., 1972; Doha et al., 1990). In the

Central Asian Republics of the former Soviet Union, bur-

rows of the desert gerbil (Rhombomys opimus) are recog-

nized as breeding sites of this sandfly species (Perfil’ev,

1968; Artemiev et al., 1972). Towards the equator, Dedet

et al. (1982) in Senegal and Mutinga et al. (1986) in Kenya

found P. duboscqi, the Afrotropical vector of L. major,

breeding in animal burrows.

Caves were implicated as probable breeding sites of

P. longipes and P. pedifer, proven vectors of Le. aethiopica,

studied by Foster (1972) in Ethiopia and by Mutinga &

Odhiambo (1986) in Kenya.

Far less is known about the breeding sites of phleboto-

mine sandfly vectors in the New World. Research on this

subject during a period of 20 years (1940–1960) yielded only

�60 specimens of immature phlebotomines (Hanson, 1961).

Among these, 19 were Lutzomyia longipalpis s.l., the main

vector of American visceral leishmaniasis, recovered by

Deane & Deane (1957) from animal corrals and among

rocks in Brazil. Those workers stressed the discrepancy

between the abundance of Lu. longipalpis adult sandflies,

which they said were ‘found everywhere’ in the State of

Ceara, Brazil, and the rarity of finding the pre-imaginal

stages. Similar results were obtained 40 years later

in Colombia, where more immatures were recovered from

animal sheds close to houses rather than from isolated

microhabitats (Ferro et al., 1997). No information is

available on the breeding sites of Lu. evansi, a proven vector

of Leishmania infantum chagasi in Colombia (Travi et al.,

1990) and Venezuela (Feliciangeli et al., 1999).

In undisturbed Neotropical forests, where vector sand-

flies are closely associated with the wild animal reservoirs of

Leishmania spp., pre-imaginal stages have been obtained for

10 out of 42 proven or suspected vectors of cutaneous

leishmaniasis (Killick-Kendrick, 1999). Notably, Lutzomyia

gomezi, Lu. ovallesi, Lu. panamensis, Lu. trapidoi and Lu.

ylephiletor were all caught emerging from the forest floor in

Panama (Hanson, 1968). Moreover, Rutledge & Ellenwood

(1975a,b) pointed out that plant–sandfly interactions lar-

gely determine the pattern of sandfly production in or on

the forest floor habitat; thus, Lu. trapidoi was associated

with large lianas (Ouruparia and Sabicea), whereas

Lu. panamensis and Lu. gomezi were associated with large

Anacardium trees. No plant association was established for

Lu. ovallesi, although its larvae were found between tree

buttresses (Hanson, 1968; Rutledge & Mosser, 1972). It was

inferred that Lu. ovallesi seeks out these sheltered places for

oviposition, as adults seldom use these habitats for daytime

resting places (Rutledge & Ellenwood, 1975a,b). This beha-

viour of Lu. ovallesi was also observed in north-central

Venezuela (Feliciangeli, 1987) where this species is the

main vector of cutaneous leishmaniasis (Feliciangeli &

Rabinovich, 1998).

Arias & Freitas (1982) caught Lu. anduzei and Lu. umbra-

tilis, emerging from the ‘terra firme’ forest floor in the

central Amazon region of Brazil (Table 2). However, they

concluded that the open forest floor is not one of the major

breeding sites for these sandflies.

Breeding sites of Lu. whitmani and Lu. intermedia have

been detected more often in the peridomestic than the

silvatic habitat (Table 2), as demonstrated by Casanova

(2001) in rural areas of the Mogy Guacu River, Brazil.

This tendency may be linked with changing epidemiological

patterns in the transmission of American cutaneous leish-

maniasis (ACL), increasing risk factors being associated

with peri- and intradomiciliary habitats (Campbell-Lendrum

et al., 2001; Desjeux, 2001).

Controlling the immature stages of phlebotominesandflies

This topic was reviewed by Alexander & Maroli (2003). For

eco-epidemiological reasons, it is important to focus on

strengthening research in the detection of breeding places.



The first attempted control aimed at immature stages of

phlebotomine sandflies, using necrocene with crude oil

and kerosene mixture, was applied in India by Smith et al.

(1936), without much success. In the Central Asian Repub-

lics, destruction of burrows of the great gerbil (Psammomys)

effectively controlled P. papatasi (Faizulin et al., 1976),

whereas chemical control with organochlorine insecticides

did not give such a good result (Vioukov, 1987). Larval

habitat modification by plastering walls of human dwellings

and cattle sheds was quite effective to control P. argentipes,

the Indian vector of visceral leishmaniasis (Kumar et al.,

1995). However, there were problems with the high cost of

the product and its high viscosity, which made it difficult to

spray. More encouragingly, selective application of the bio-

pesticide Bacillus sphaericus (Bs) into the burrows of host

rodents was successful against P. papatasi (Pener et al.,

1996). The Bs toxin works as a stomach poison against

target insects, but it remains unclear whether the impact

was mostly on immature or adult sandflies. Spraying plants

with a sugar solution of B. sphaericus, for adult sandflies to

ingest and carry back to poison the larvae, reduced sandfly

populations emerging from animal burrows but not those

emerging from termitaria in Kenya (Robert et al., 1997).

As no stable silvatic breeding sites of ACL vectors have

yet been identified in the Americas, it remains impossible to

target the immature stages or to attempt sandfly source

reduction. Even so, some efforts to reduce sandfly popula-

tions were applied to tree-holes where they might affect

immature as well as adult sandflies (Floch, 1957; Chaniotis

et al., 1982; Ready et al., 1985). Perhaps entomopathogenic

fungi could be used against sandfly immatures in defined

areas; for example Beauveria bassiana is applied to control

the coffee berry borer Hypothenemus hampei in Colombian

coffee plantations. Unfortunately, a trial evaluation of

B. bassiana was unsuccessful for sandfly control (Reithinger

et al., 1997). Because of the increasingly close human–vector

associations, giving rise to greater risks of Leishmania trans-

mission (Dye, 1996), there is growing hope and scope for

identifying restricted peridomestic breeding sites of sand-

flies that might be amenable to control, by larviciding or

environmental modification to eliminate the source.

Acknowledgements

This review is dedicated to Dr Lawrence Quate, who, in

1964, processed 2500 kg of soil in Sudan to recover only a

single larva of Sergentomyia africana (Quate, 1964). With

the recent death of Dr Quate, we have lost one of the very

few expert taxonomists of the family Psychodidae.

I wish to thank Dr Philippe Desjeux for the encourage-

ment to publish this review and his helpful criticisms,

Professor Yan Jia Leng for kindly providing me informa-

tion on the breeding sites of vectors of leishmaniases in

China, Dr Michele Maroli of the Istituto Superiore di

Sanita, Roma, Italia, and Sinead Fitzpatrick, postgraduate

student at the London School of Hygiene and Tropical

Medicine, U.K., for kindly providing photocopies of papers

not available in Venezuela.

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Accepted 10 February 2004



natural breeding places of phlebotomine sandflies

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