Colonization of Lutzomyia shannoni (Diptera: Psychodidae) utilizing an artificial blood feeding technique



Laboratory colonization of hematophagous insects must include an efficient method of blood feeding, preferably by artificial means. Strict rules for obtaining animal use permits, extensive animal maintenance costs, and indirect anesthesia effects on animal health warrant the development of an artificial membrane feeding technique for sand fly colonization in laboratories. An attempt was made to colonize Lutzomyia shannoni using an artificial blood feeding membrane to replace the use of live animals commonly used for sand fly blood-feeding purposes. Lutzomyia shannoni readily fed through a pig intestine membrane exposed at an angle of 45°. However, it did not feed through a chicken skin membrane. Olfactory attractants were unable to improve blood-feeding efficiency. Plaster of Paris was the most suitable oviposition substrate. Female L. shannoni adults laid no eggs on moist sand substrate. Sand fly adults held in groups of ten or more laid higher numbers of eggs than did individually maintained sand flies. Inclusion of the L. longipalpis oviposition hormone dodecanoic acid or the presence of previously laid eggs did not stimulate L. shannoni oviposition. The average L. shannoni egg, larval, and pupal duration were 9.3, 36.7, and 17.8 days, respectively. The addition of a 20% sugar solution improved adult female longevity. Females survived longer (14.8 days) than males (11.9 days). Lutzomyia shannoni was successfully colonized in the laboratory for up to four generations using this artificial membrane technique.


Phlebotomine sand flies (Diptera: Psychodidae) are important vectors of human and animal pathogens including Leishmania, Bartonella, vesicular stomatitis virus, and phlebovirus (Young and Duncan 1994). Lutzomyia shannoni (Dyar) is a proven vector of vesicular stomatitis virus and is suspected to be capable of vectoring the pathogens causing visceral leishmania and sand fly fever (Tesh 1988, Comer et al. 1991, Young and Duncan 1994). Both males and females feed on nectar and other plant juices, but females require a blood meal for follicle maturation. Lutzomyia shannoni feeds on a variety of mammals. Known blood meal sources include white-tailed deer, horses, donkeys, mules, cattle, swine, raccoons, cats, dogs, rodents, birds, and humans (Tesh et al. 1971, Young and Perkins 1984, Comer et al. 1994).

Sand fly colonization is a critical and limiting factor in research on vector potential, life cycle, pathogen transmission, and management studies. Successful colonies are indispensable in genetics, physiology, and behavioral studies (Endris et al. 1982). Several authors have emphasized the difficulty of establishing sand fly colonies (Killick-Kendrick 1978, Killick-Kendrick and Killick-Kendrick 1991). Sand flies demand special care for colony establishment and maintenance and the initiation of laboratory colonies is far more difficult than the maintenance of already-established colonies (Killick-Kendrick 1978, Young et al. 1981). Of the 800 known sand fly species, approximately 20 species have been successfully colonized (Killick-Kendrick et al. 1991). The degree of success varies with the species. While some species are reared easily, others, which readily develop to adults from eggs laid by wild-caught females, often feed with great reluctance in the laboratory on any animal, including their natural host (Hertig and McConnell 1963).

Some species, including Phlebotomus papatasi (Scopoli) and L. longipalpis (Lutz and Neiva), have adapted readily to laboratory conditions, but others have not when reared under similar conditions. Colonization of hematophagous insects must include an efficient blood feeding method, preferably by artificial means (Harre et al. 2001). Perkins1 reported the first successful laboratory colonization of L. shannoni through 12 generations feeding on live hamsters. Later, Ferro et al. (1998) reared laboratory-colonized L. shannoni on live animals for life cycle and life table studies. Currently, strict rules for obtaining animal use permits, extensive animal maintenance costs, and indirect anesthesia effects on animal health warrant the development of a successful artificial membrane-feeding technique. Artificial membrane-feeding techniques have been used successfully for various haematophagous insects, including some sand fly species (Ward et al. 1978, Galun 1979, Modi and Tesh 1983, Tesh and Modi 1984, Ghosh and Mukhopadhyay 1998, Harre et al. 2001). However, we found no documented reports of L. shannoni colonization using artificial membrane techniques. Therefore, the present studies were aimed at the laboratory colonization of L. shannoni using an artificial membrane-feeding technique.


Containers used to hold adult sand flies for blood-feeding were made from transparent plastic specimen containers. A 2 to 3 cm diameter hole was made on the bottom and the sidewalls of the containers using a heated brass cork borer. The bottoms of the containers were filled with 2 to 3 cm of Plaster of Paris (POP) paste and dried to adhere to the container (Figure 1a). The side hole was covered with a fine nylon mesh (400 mesh, 37µm) utilizing silicon glue. The center of the lid was cut out and covered with cheesecloth, allowing exposure of the flies to the blood feeder.

Figure 1a.

Rearing equipment used for Lutzomyia shannoni colonization; a) a holding container, b) an oviposition container, and c) a Plaster of Paris resting platform.

Oviposition containers were similar to the holding containers (Figure 1b). However these containers had a l-cm sidewall hole. The hole was not screened, thereby allowing the release of adults into the container. A 2-cm hole was cut in the center of the lid and covered with nylon mesh utilizing silicon glue for provision of a sugar source. Another 3–4 cm hole was cut at the bottom of the container for moistening the POP through a moistened filter paper.

A 30×15×15 cm glass aquarium turned on its side was used to maintain and sort blood-fed female flies. The aquarium bottom and one of the 15×30 cm sidewalls of aquarium were covered with POP. A tubular cloth sleeve was affixed to a plastic plate by means of adhesive that was subsequently glued to the aquarium opening. This design allowed for sand fly transfer while preventing their escape (Perkins 1982). A moistened cylindrical platform made from POP was used as resting place for sand flies (Figure 1c). Its utilization reduced mold growth on the aquarium floor. A mouth aspirator with a HEPA filter (Model 612; John W. Hock, Gainesville, FL) was used to collect and release the sand flies into the containers and aquariums.

The glass blood feeder (Perpetual Systems Corp, Rockville, MD) consisted of an inverse funnel surrounded by a chamber through which water flowed via a thermostatic circulator (LKS, Model 2219 Minitemp II, Brommer, Sweden) (Figure 2a). Defibrinated bovine blood was obtained from a local slaughterhouse. Pig intestine (The SausageMaker Inc., Buffalo, NY) was utilized as the feeding membrane. Prior to use, the pig intestine was immersed in distilled water for 15 min to remove salts and other impurities. The large open end of the blood-feeder was covered with the pig intestine creating the feeding membrane (Figure 2b). A rubber band was used to fix the membrane to the feeder. The membrane was allowed to air dry for 30 min. The membrane was stretched sufficiently to be tight and firmly attached to the feeder prior to drying and subsequent placement on the feeding stand. Bovine blood was added to the feeder through the top opening using a needleless syringe. The feeder was connected to the thermostatic circulator by means of rubber tubing, with water circulating at a temperature between 28 and 32° C. The incubator temperature was also increased to 28°C during the feeding process to avoid condensation. The thermostatic circulator was switched off if condensation was observed.

Figure 2a.

Blood-feeding equipment and set-up used for membrane feeding of Lutzomyia shannoni a) a glass blood feeder prior to membrane attachment, and b) an operational glass blood feeder outfitted with a porcine intestine membrane and exposed to L. shannoni.

The circulator was allowed to run for at least for 10 min to warm the blood prior to exposing the sand flies to the feeder. The blood feeder was clamped on a stand and positioned at an angle of 45°. Sand flies were exposed to the feeder through the screened hole in the lid of the earlier described blood-feeding container.

Distilled water and table sugar were used to prepare a 20% sugar solution. A fresh sugar solution was prepared before each sand fly feeding. The larval food was prepared from rabbit feces and alfalfa pellets and allowed to fully age as per Modi and Tesh (1983).

Lutzomyia shannoni adults were collected from San Felasco Hammock Preserve State Park (29°45'N, 82°30'W), Gainesville, Alachua County, FL, using counter flow geometry traps (MM-X model, American Biophysics Corp., USA) as described in Mann et al. (2009). Briefly, the inlet of the traps was covered with a 1.2 × 1.2 mm mesh bag to exclude mosquitoes and other large insects from damaging collected sand flies. All traps were provided with an apple slice as a sugar source for captured sand flies. Following a 24-h capture period, traps were returned to the laboratory and placed in a −20° C freezer for 45 to 60 s, after which sand flies were removed, identified, and grouped for blood feeding. To immobilize sand flies and allow for proper identification, sand flies were held on glass dishes placed on crushed ice.

Up to 30 male and female sand flies were placed in the blood-feeding containers. The POP in feeding containers was moistened with up to 5 ml distilled water before the introduction of the insects. Flies were offered an apple slice as a sugar source through the side hole. The container was clamped in such a way that nylon mesh had direct contact with the feeder membrane (Figure 2b). Fresh blood and a new membrane were used at each feeding.

Flies were allowed to blood feed for at least 4 h. Thereafter, all flies were released into an aquarium. Those flies that had blood fed were aspirated and released into a similar aquarium containing two moistened cylindrical POP platforms. Platforms were placed in glass dishes that allowed sand flies to rest on a moistened surface. The aquarium POP was not moistened to avoid microbe growth. Flies were provided with both a 20% sugar solution on a moistened filter paper and an apple slice placed in a glass dish. Flies were allowed to remain in the aquarium for 4 days prior to transferring into oviposition containers.

Larval food was finely ground with a coffee grinder and oviposition containers were provisioned with 0.25 g of food (Modi and Tesh 1983). Blood-fed flies were transferred to the oviposition containers through the side hole using an aspirator. The hole was then closed with a rubber stopper. The flies were provided with a 20% sugar solution daily via a cotton wick placed on top of the nylon screen-covered lid. Dead flies were carefully removed from containers daily. Flies were allowed to oviposit in the containers until all the flies had died.

Containers with eggs were placed on moistened filter papers in glass dishes. The dishes were placed in trays containing 2% soapy water to enhance humidity and exclude ants. Filter papers were moistened periodically to maintain an adequate moisture level. Complete drying of a filter paper indicated too low a moisture level, while accumulation of moisture in the container indicated excessive moisture. Trays were kept in an incubator (I-36VL, Percival Scientific, Perry, IA) at 27° C, 85–90% RH and 8:16 (L:D) h photoperiod. Each container was provided with 1 to 2 g larval food periodically, depending upon the larval density and food consumption. Larvae congregated along walls if food was abundant, while low food levels resulted in larvae concentrating in clusters. The larvae were allowed to remain in the containers until adult emergence. Food supplementation was stopped after more than 90% of larvae pupated. Newly emerged adults were transferred into separate holding aquariums for 4 to 5 days before blood feeding.

Factors affecting specific sand fly colonization parameters

The effect of various factors on L. shannoni colonization efficiency was evaluated by altering several factors such as sugar concentration, oviposition substrates, adult sand fly density, and feeding and oviposition stimulants. Factors were evaluated on a laboratory-colonized F1 generation that was obtained by utilizing the above-mentioned colonization technique. All factors for subsequent experiments were evaluated on 20 adult female sand flies per replication, unless mentioned otherwise. There were five replications in every experiment.

Longevity of L. shannoni adults were evaluated under four sugar solution (2, 10, 20, or 30%) feeding regimes following blood feeding. Flies (as described previously) were held in oviposition containers and observations were recorded on adult female longevity until the death of all flies. Solutions were prepared by dissolving an appropriate amount of table sugar in 50 ml of distilled water. The sugar solutions were provided to adult sand fly females via a 1-cm cotton wick placed on the screened hole in the holding containers lid. The cotton wicks were changed daily and a stock solution was refreshed every 4–5 days to avoid fungal growth. The stock sugar solutions were held in the refrigerator at 4° C.

Five known mosquito host-seeking attractants, namely red mix (1-octen-3-ol and 1-hexen-3-ol in acetone, as described in Mann et al. 2009), rosalva (Butler 2006), hexane extract (Blackwell et al. 1996), ocular solution (J.F. Butler, personal communication), and host urine (Nigam and Ward 1991) were evaluated for improving L. shannoni blood-feeding efficiency on the membrane feeder. In this study, bobcat urine was utilized. The stimulant was applied directly on the feeding membrane at 0.1 ml/7.06 cm2 (feeder area). The feeder was allowed to dry for 30 min before fly introduction. A blood feeder without a stimulant was used as a control. Sand flies were allowed access to the membrane for 4 h. The observations scored were the number of fully engorged females after 4 h of blood feeding and were expressed as a percentage of the total number of sand flies exposed. Sand flies that died due to static or handling were excluded from analysis. A female was considered engorged if its abdomen was more than half-filled with blood.

The effect of substrate on sand fly oviposition was evaluated by placing 20 blood fed adult L. shannoni females in the oviposition containers as described previously. For these experiments, the containers were outfitted with one of six substrates, namely, smooth-surfaced POP (SSPOP), irregular-surfaced POP (ISPOP), animal odor-containing field sand (AOFS), Play Sand (PS), and the combinations of SSPOP + AOFS and SSPOP + PS. The ISPOP was made by engraving lines in the plaster with a sharp knife. The animal odor-containing field sand was collected from the University of Florida dairy research unit (Alachua, FL). Play Sand (Quickrete, Atlanta, GA) was obtained from a local retailer. The two POP + sand substrates comprised POP and sand in a 1:1 ratio. Appropriate moisture was maintained in all the substrates by placing a moist filter paper at the base of the containers. The moist filter paper moistened the POP through absorption. The filter paper was remoistened and replaced when needed. The adult sand flies were maintained on these substrates utilizing the technique described previously until all the sand flies died in the containers. The total number of eggs laid per female were counted on each substrate.

Lutzomyia longipalpis oviposition hormone, ammonium laurate (Dougherty et al. 1994), Lutzomyia larval food (Nieves et al. 1997), uneclosed live L. shannoni eggs (Dougherty et al. 1994), and the mosquito and sand fly host-seeking attractants red mix and rosalva were evaluated for improving L. shannoni oviposition efficiency. Ten mg of each chemical or larval food was applied either directly onto an oviposition substrate or by dissolving the stimulant in 0.1 ml acetone. Ammonium laurate and larval food were applied directly on the POP while red mix and rosalva were applied following their dilution in acetone. Acetone was allowed to evaporate entirely from the substrate before exposing the females to the stimulants. The L. shannoni uneclosed egg treatment consisted of POP substrate with ten, two to four day-old live eggs per container. Containers with only POP served as a control. The number of eggs laid per female was scored after all females in a container had died.

Sand fly density effects on oviposition efficiency was evaluated by placing 1, 5, 10, and 20 L. shannoni adults per oviposition container. The standard oviposition containers described earlier were used for this experiment. Flies were held in oviposition containers with POP as the oviposition substrate. The containers were scored for number of eggs laid per female after all the sand flies in a container had died.

Duration of life stages

Duration of laboratory-reared F1 L. shannoni life stages were recorded from 20 randomly chosen rearing containers. The rearing containers were inspected daily and observations were made on egg, larval, pupal, and adult duration. Larval duration is reported as a total duration for all larval instars. First instar larvae were scored for time-in-stage. Thereafter, larval duration could not be scored separately by instar due to the difficulty in observing the 2nd through 4th stage larvae that remained hidden under the food substrate. Therefore, total duration in the larval stage was determined when flies had pupated.

Statistical analysis

Analysis of Variance (ANOVA) was performed to evaluate oviposition substrate, feeding stimulants, sugar levels, and oviposition stimulants. If differences among means were present, then the means were separated using Tukey's mean separation test with (α= 0.05). All the data were log (n+1) transformed. Statistical analysis was carried out using SAS software version 9.1 (SAS 2003).


Lutzomyia shannoni was successfully colonized in the laboratory to the fourth generation using an artificial membrane blood feeding technique. Sugar levels yielded significant differences among the treatments for adult female longevity (F3,16= 11.30; p=0.0003) (Table 1). Sand flies showed the shortest survival at the 2% sugar level. Increasing the sugar concentration up to 20% significantly improved longevity with survival ranging from 6–18 days at this concentration. However, increasing the sugar concentration up to 30% did not further increase longevity.

Table 1.  Influence of sugar concentrations on Lutzomyia shannoni survival.
 Longevity (days)
Sugar level (%)Mean±SERange
  1. Means followed by different letters are statistically different at p<0.05. n=20, ANOVA: F3,16= 11.30; p=0.0003.

26.84 ± 0.45a5–10
108.92 ± 0.46b5–14
2013.53 ±0.68c6–18
3012.62 ±0.55c5–18

Although visual observations indicated that higher numbers of sand flies were attracted to the stimulant-treated blood feeders, the stimulants did not increase feeding efficiency over the controls. Numerically, the highest feeding efficiency (45.9%) was achieved with hexane extract compared to 39.2% with no stimulant.

The type of material used in the oviposition container influenced the total number of eggs laid in each container (F5,24=61.70; p<0.0001) (Table 2). Up to 12.2 eggs per female were laid on ISPOP. However, there were no significant differences between ISPOP, SSPOP, AOFS, and SPOP+AOFS. No eggs were laid in containers containing PS or SSPOP+PS. None of the tested oviposition stimulants resulted in significantly more oviposition activity. Observed oviposition ranged from 7.1 to 12.4 eggs per female.

Table 2.  Mean (SE) number of Lutzomyia shannoni eggs laid on six oviposition substrates.
SubstrateNumber of eggs laid per female
  1. SSPOP: Smooth surfaced Plaster of Paris, ISPOP: Irregular surfaced Plaster of Paris, AOFS: Animal odor-containing field sand, PS: Play sand.

  2. Means followed by different letters are statistically different at p<0.05. n=30, ANOVA: F5,24=61.70; p<0.0001.

  ISPOP11.2 ±1.23a

As the number of L. shannoni females per cage increased, the number of eggs laid also increased. When ten (11.20 ± 0.83) or 20 (11.72 ± 0.48) sand flies were held together in oviposition containers, significantly (F3,16=34.50; p<0.0001) higher numbers of eggs were laid than when one (2.40 ± 0.67) or five (6.08 ± 0.65) sand flies were held in a container (Table 3).

Table 3.  Effect of adult sand fly density on Lutzomyia shannoni oviposition.
Number of sand flies perNumber of eggs laid per
  1. Means followed by different letters are statistically different at p<0.05. n=20, ANOVA: F3,16=34.50; p<0.0001.


Average L. shannoni egg duration was 9.3 days with a range of 6 to 14 days (Table 4). The average 1st instar duration was 7.9 days with a range of 5–11 days. Visual observations indicated that the larval length increased with each progressive instar. All instars, with the exception of 1st instar, had four caudal setae. The average total of the 2nd through 4th instar larval durations was 28.8 days. Diapause or quiescence was also observed in larval instars, thereby extending the larval period from 28 to 47 days. The pupal period ranged from 11 to 26 days with an average of 17.8 days. Adults emerged in the morning hours and males emerged up to two days before females. Females (14.8 days) survived longer than the males (11.9 days), and the male and female adult lifespan duration ranged between 7 to 13 and 8 to 18 days, respectively. The total egg to adult development period ranged from 64 to 80 days.

Table 4.  Duration of Lutzomyia shannoni immature and adult stages under laboratory conditions.
StageMean ± SEM (days)Range (days)
1st instar7.9±0.595–11
2nd-4th stage36.7±2.1728–47
 Adult male11.9±1.067–13
 Adult female14.8±1.368–18
Total life cycle64–80

The sand flies fed readily on defibrinated bovine blood through pig intestine membrane. The angle of blood feeder when presented to sand flies also influenced the feeding behavior of adults. Holding containers in the vertical position with membranes horizontal to the flies resulted in very few (< 5%) engorged females as compared to more than 50% blood feedings in containers held at a 45° angle. Visual observations indicated that membranes on which sand flies had previously fed upon, when held at an angular position, were even more attractive to sand flies. However, considering the potential of bacterial and fungal infections on reused membranes we avoided this practice. Based on our evaluation of various factors, a periodic schedule for colonizing L. shannoni was developed (Table 5).

Table 5.  A periodic schedule developed for rearing Lutzomyia shannoni under laboratory conditions utilizing artificial blood feeding membrane technique.
1–4Blood feed sand flies and release into glass aquariums. Separate blood fed sand flies and maintain in a separate aquarium until day 4 while providing an apple slice and a sugar solution.
4–5Transfer to and maintain the blood fed sand flies in the oviposition containers with larval food supplementation.
5–15Maintain flies in the oviposition containers on sugar solution until day 15.
15–50Remove dead adult flies from the containers and add larval food as needed based on larval density until adult emergence. Maintain proper moisture and sufficient larval food in the containers. Stop adding food when all larvae have pupated.
50–70Maintain pupae without disturbing the containers.
65–90Examine for adult emergence. Maintain newly emerged adults in separate holding containers up to 3 days before blood feeding.

This technique allowed for uninterrupted colonization of L. shannoni for up to four consecutive generations. It was possible to produce eggs, larvae, and adults of known ages for experimental purposes and to facilitate further studies on this insect. However, the number of adults produced continued to decline with each generation.


To our knowledge, this is a first report that documents the successful colonization of L. shannoni utilizing artificial membrane to provide blood meals. Blood feeding sand flies in a laboratory setting is a difficult process. Reared adults have shown great reluctance to blood feed in a laboratory on any animal, including their natural host (Hertig and McConnell 1963). Chicken skin has been commonly used as a blood feeding membrane (Schlein et al. 1983, Ciufolini et al. 1991, Ghosh 1994, Hanafi et al. 1998, Rowton et al. 2008); however, we were unable to blood feed even wild caught sand flies through chicken skin membranes. This is somewhat surprising since chickens have been reported as a host of L. shannoni (Thatcher and Hertig 1966).

Condensation and static increased the risk of flies getting stuck to the feeder sides and subsequently dying. The use of high quality, hard plastic feeding containers that had no or lower static properties along with fluctuating the circulating temperature reduced both condensation and static, thereby improving sand fly feeding success.

It was important to use a completely mature larval diet (a diet in which all microbe growth had ceased) and to remove all dead adults from the containers prior to the addition of more food. Larval diet that had not fully matured resulted in mold growth inside the containers, which killed the younger larvae caught in the fungal growth. Proper moisture levels in the containers resulted in reduced mold growth. This occurrence also was reported by Rangel et al. (1985).

Irregular surfaces stimulated L. longipalpis oviposition (Nieves et al. 1997). However, no such thigmotropic response was observed for L. shannoni as it laid an equivalent number of eggs on regular and irregular surfaced POP. Lutzomyia shannoni laid no eggs on PS and PS-POP mixtures. However, it laid eggs on AOFS alone and in combination with POP indicating the role of physical and olfactory stimuli in L. shannoni oviposition. Several unknown factors appear to govern L. shannoni oviposition besides olfactory stimuli, as previously laid eggs, larval food, and oviposition hormone did not stimulate oviposition. Ammonium laurate, previously identified as a L. longipalpis oviposition stimulant (Dougherty et al. 1994), had no effect on L. shannoni oviposition. Casanova et al. (2006) recognized different sex pheromones within L. longipalpis populations in Brazil and perhaps L. shannoni utilizes an undetermined oviposition pheromone.

Sugar is an essential food for adult phlebotomine sand flies. It is the only nutrient consumed by males and probably the commonest for females, even though they need vertebrate blood to produce eggs (Schlein and Muller 1995, Tang and Ward 1998, Schlein and Jacobson 1999, Muller and Schlein 2004). Not surprisingly, L. shannoni longevity increased as sugar concentrations increased. The majority of the females showing early mortality died during the egg-laying process. Killick-Kendrick et al. (1977) and Rangel et al. (1985, 1986, 1987) obtained higher female survival rates for ovipositing females by sugar feeding the females during and after the oviposition. Schlein and Jacobson (1999) and Souza et al. (1995) obtained similar results and suggested inclusion of sugar-containing foods in sand fly diets to increase survival rates in experimental studies.

Lutzomyia shannoni feeds on a variety of mammals, including cats (Tesh et al. 1971, Young and Perkins 1984). However, feeding stimulants reported to attract mosquitoes and sand flies (Butler 2006, Mann et al. 2009), including host urine (Nigam and Ward 1991) (bobcat), did not stimulate L. shannoni feeding. Similarly, the presence of sand fly eggs, oviposition stimulant hormone, and larval food did not stimulate oviposition. This suggests the role of several other factors in the blood feeding and oviposition process. However, Elnaeim and Ward (1992) reported oviposition stimulation with larval food for L. migonei (Franca). Trans beta farnesene stimulated L. longipalpis feeding, but not L. shannoni (Tesh et al. 1992).

Lutzomyia shannoni oviposition was directly influenced by cohabiting density, wherein egg density increased as the number of flies per container increased. However, this effect peaked with groups of ten or more adults. Similarly, an increase in egg density with increasing numbers of sand flies, up to the carrying capacity of 50 adults per container, has been reported for L. serrana (Damasceno and Arouck) (Santamaria et al. 2002). This effect suggests the presence of a pheromone or a compounding effect when density is increased. Future studies are needed to identify the presence or role of such a compound or crowding effect.

Our results for L. shannoni life history corroborated those of Ferro et al. (1998), however, we obtained longer adult survival than what has been previously reported for L. shannoni. Larval instar duration was longer for those instars emerging in winter, even though they were reared under similar laboratory conditions. We also observed apparent larval diapause in 4th instars. Previous research suggests that L. shannoni larval diapause appears to be facultative in response to poor rearing conditions such as diet quality and disturbances. Lawyer and Young (1991) associated 3rd and 4th stage larval diapause of L. diabolica (Hall) with adverse conditions such as excessive moisture, extreme temperatures, or poor diet. Facultative diapause in the 4th instar has been recognized as a type of dormancy in sand flies of the Palearctic region (Ready and Croset 1980, Killick-Kendrick and Killick-Kendrick 1987).

We demonstrated that L. shannoni can be successfully colonized and maintained in a laboratory setting utilizing an artificial feeding membrane technique. The increased experience in manipulating the membrane feeder tended to improve the feeding percentage. This artificial membrane feeding technique will not only reduce the need for animals, but it will also reduce the related costs, time, and supervision needed for the rearing of this species.


  • 1

    Perkins, P.V. 1982. The identification and distribution of phlebotomine sand flies in the United States with notes on the biology of two species from Florida (Diptera: Psychodidae) Ph.D. dissertation, University of Florida, Gainesville, FL.


We thank Uli Bernier and Daniel Kline for providing the RM attractant, San Felasco Hammock Preserve State Park, Gainesville, FL, U.S.A. for study sites. We also thank R. Hammel, L. Wood, J. Butler, P. Perkins, J. Pecora, J. Matta, S. Nunez, and E. Fitzgerald for their assistance with this study. Funding was provided by the U.S. Armed Forces Pest Management Board (W9113M-06-S-001) and from State Project FLA-ENY-04616.