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
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80
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
74 M. D. Feliciangeli
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80
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*
Brazil:CentralAmazon
ET(243)
55A
Lu.paraensis
Brazil:CentralAmazon
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
Breeding places of phlebotomine sandflies 75
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80
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
76 M. D. Feliciangeli
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80
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.
Breeding places of phlebotomine sandflies 77
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80
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
80 M. D. Feliciangeli
# 2004 The Royal Entomological Society, Medical and Veterinary Entomology, 18, 71–80