Establishment and maintenance of sand fly colonies

Abstract

Sand flies used to have a reputation for being difficult and labour-intensive to breed. Here we summarize our experience with establishment and maintenance of sand fly colonies and their use for infective experiments: techniques for collection and handling wild-caught females, rearing larvae and adults and experimental infections of sand flies by Leishmania using membrane feeding. In addition, we compare major life cycle parameters between various colonies maintained under standard laboratory conditions. The sand fly rearing is tricky but some species can be reared in large numbers with a minimum of space and equipment. Initiation of new colonies from endemic sites is a prerequisite for accurate studies on parasite-vector interaction but it is more difficult step than routine maintenance of colonies already established in laboratory for many generations. Laboratory colonies are essential for studies on sand fly biology and sand fly-Leishmania interaction. A variety of techniques has been described for rearing these insects in laboratory conditions (reviewed by Hertig and Johnson 1961, Killick-Kendrick and Killick-Kendrick 1991, Modi 1997). A detailed bibliography on this subject is given by Killick-Kendrick et al. (1991). Of about 700 known sand fly species, fewer than five dozen have been colonized in the laboratory so far, but only a few of them have been successfully reared continuously in large numbers to provide sufficient material for experimental work. Such work is essential for understanding sand fly physiology, epidemiology of leishmaniases, and parasite-vector-host relationships. Sand fly species and even colonies of different geographical origin vary in many biological and physiological parameters, including saliva composition (Warburg et al. 1994, Volf et al. 2000). There are also significant differences in susceptibility for Leishmania parasites. Certain species, for example P. papatasi and P. sergenti, are termed “restricted” (Kamhawi 2006) or “specific” (Myskova et al. 2007) vectors, as they allow the maturation and transmission of a single Leishmania species. The others tested so far seem to be permissive as multiple Leishmania species are able to mature within their gut (for review see Kamhawi 2006, Volf and Myskova 2007). However, for the majority of sand fly species, there are no data on susceptibility to Leishmania yet. Therefore, any new colony is a treasure and we like to encourage medical entomologists to colonize sand flies from nature. The aim of this report is to summarize our experiences with the establishment and maintenance of sand fly colonies and their use in infective experiments: techniques for the collection and handling of wild-caught females, rearing larvae and adults, and experimental infections of sand flies by Leishmania using membrane feeding. In addition, we summarize data obtained by long-term maintenance of sand fly colonies in our laboratory and compare major life cycle parameters among various colonies under standard conditions. Sand fly adults of both sexes can be collected by several methods, either while resting during the day or foraging at night. Sampling from resting sites, like walls of houses, animal dwellings, caves, and tree holes, can be done by several forms of mouth aspirator; similarly, active catches with an aspirator can be done from bait animals or on human bait. For a review of sampling methods see Alexander (2000). The dimensions of each component of an aspirator can be varied according to preferences of the collector. Sand flies are fragile and therefore the body of the aspirator should be wider than the opening. On the other side, aspirators are closed with a piece of fine mesh gauze or sinter filter. The plastic aspirators are light and strong but could damage sand flies because of static electricity. Therefore, the glass ones could be recommended for most studies in the laboratory. However, they are not commercially available and must be made by a glass blower. Working with a large number of sand flies, we favor a reservoir type, with the body wider than the entrance (Figure 1a). It consists of a tube sealed at one end by rubber, except for a narrow tube through which the sand flies enter. This makes it difficult for the insects to escape and helps capture more specimens without the necessity to transfer them into a cage or other type of container. Emptying the reservoir aspirator may involve dismantling it within a cage. Equipment used for sand fly maintenance. a: two different types of glass aspirators and the glass feeder used for membrane feeding. b: boxes with larval pots and cages wrapped in plastic bags are maintained in incubator. c: two cabinets (one with front panel fitted) for fermentation of larval food. Up side down position of plastic trays ensures aerobic conditions during fermentation process. d: detail of larval food during composting process. Long, hair-like mycelia are evidence of optimal fermentation conditions. e: larval pots with a layer of plaster of Paris on sides and bottom are closed on the top with a snap cap and fine gauze. Pots are maintained in plastic boxes with a bottom filled with layer of fine sand. f: nylon cloth cages of different size are suspended on the steel frame (the larger cage here is enveloped in a plastic bag). One potential problem is that sand flies shed large numbers of hairs which are inhaled by the collector and can cause respiratory problems. Also, resting sites involve dark and humid microhabitats with potential danger of fungal and bacterial infections, such as histoplasmosis or Q-fever caused by Coxiella burnetti. Therefore, some authors introduced a filter to prevent the inhalation of dust, hair, and microorganisms (Warburg 1989) or introduced the condom barrier (Tang 1996). Hand-held power-operated aspirators are also commercially available (see for example, Higgs and Beaty 1996) and we advise they be used in the field for sand fly collections from crevices, stables, or even from traps. Light traps are used extensively in field studies of sand flies. The most widely used type, battery-operated light-suction traps of the CDC type, can be left overnight to collect sand flies. The distance at which these traps are attractive to sand flies appears to be not great; Killick-Kendrick et al. (1985) estimated a maximum distance of only 2 m for P. ariasi. However, in resting or breeding places like stables, cellars, caves, or the entrance of burrows, sand fly population densities are high which results in good catches. As with mosquitoes, trapping methods using animal baits or carbon dioxide preferentially sample host-seeking females. Those involving light tend to yield a higher proportion of gravid females and males. However, in some sand fly species, like Lutzomyia longipalpis or P. argentipes, males are readily attracted to the hosts as mating sites (Lane 1993). It is advisable to collect traps early in the morning and take out the sand flies with an aspirator as soon as possible before they start to suffer from heat and desiccation. In addition to sand flies, the light trap also collects various other insects and spiders, some of which may eat the sand flies or damage them. Similarly, overcrowding and leaving sand flies too long in the aspirator increases their mortality. Initiation of new colonies is a more difficult step than the routine maintenance of colonies already established in the laboratory for many generations. Therefore, it is advisable to get expertise by first maintaining an “easy” colony like P. papatasi or L. longipalpis. Sand flies immobilized by cold or CO2 are sorted under the stereomicroscope according to sex and physiological state and gravid females intended for the establishment of colonies are transferred into plaster-lined plastic pots used for larval rearing (see below). When this is not possible, blood-fed females visible to the naked eye could be selected and transferred from the traps using mouth aspirators. The size of the pot should consider the number of gravid females and prior to the placement of the sand flies the pot must be moistened with water. Pots are kept in plastic rectangular boxes with tight-fitting lids. High humidity is ensured by a layer of moistened filter paper or tissue in the bottom of the box. Females are provided with a sugar source such as a fresh apple slice or a cotton wool pad saturated in 10–50% sucrose solution or honey. For most species, the optimum temperature is between 24–28° C. Pots in boxes are also useful for the transporting of flies, but cooler conditions (15–20° C) during transportation is recommended. Non-gravid and unengorged sand flies are first transferred into a suspended nylon cloth cage used for rearing the adults (see below) and offered a blood meal. Cages are enveloped in plastic bags, again with a piece of damp cotton wool or tissue on a Petri dish. Females can be fed on a variety of vertebrates, with the choice of animal dependent on its local availability and the species of sand fly. L. longipalpis or P. papatasi are opportunistic and readily feed on mice, while the others, like Larroussius or most Adlerius species prefer hamsters or rabbits. Anesthetized mice or hamsters are placed into the cage for about 1 h. The best results we obtained using anesthetization by combination of ketamin and xylazine (10 mg and 2 mg per 100 g body weight, respectively). The presence of sand fly males may improve the engorgement rate. Since females prefer feeding in the dark, the cage is covered with a dark tissue during feeding. After feeding, females are left undisturbed for 12–24 h, mainly for complete formation of the peritrophic matrix, which makes their gut less fragile. Details on formation of the peritrophic matrix and differences between various sand fly species are summarized by Sadlova and Volf (2009). After laying the eggs, females die or could be killed for species identification. Pots with eggs are then transported to the laboratory. Catches often consist of mixed species and therefore the first laboratory generation must be sorted according to species. This step is time-consuming and laborious but in favorable conditions it can be done even with wild-caught sand flies before transportation. The technique we used is adapted from Killick-Kendrick and Killick-Kendrick (1991). Gravid females are tubed individually in small glass vials with a piece of folded filter paper inside. The mouth of the vial is closed with gauze and a plastic cap with a hole in the center. A tiny piece of cotton wool soaked in sucrose solution is put on the gauze and changed daily. The vials are kept in containers lined with filter paper or tissue moistened by distilled water. Females are encouraged to lay eggs by moistening the folded filter paper inside the vials with few drops of distilled water using the syringe with needle. With engorged females the filter paper inside vials is not moistened until after defecation when yellow, brown or black spots appear on filter paper or inner surface of vial. Vials are checked for oviposition twice a day and females that laid eggs are removed (either dead or still alive). The terminal segments of females should be detached and placed in a drop of saline for immediate identification under the microscope (preferably by phase-contrast) or mounted in a drop of Berlese's fluid and identified within a few days. The vial, labeled with date and identity of female, is kept for 2–4 days after the date of oviposition as during early period of embryonic development the eggs should not be handled. Then, eggs of the same species are transferred to plaster-lined pots using a fine brush or by washing with distilled water. Again, the diameter of the pot used depends on the number of eggs. Colonies are maintained either in a room with controlled temperature or in an incubator at 25 to 28° C. Most colonies in our laboratory are maintained at 25–26° C. Photoperiod is probably not a crucial factor but 14:10 (light:dark) is used in many laboratories. High humidity (70–95%) is ensured by wrapping cages in plastic bags with wet tissue or cotton wool inside (Figure 1b), the surrounding humidity in the insectary or incubator (60–70%) can be ensured by commercially available humidifiers. For larval rearing we used different types of pots from a solid plastic with a big hole cut in the bottom and walls roughened with sand paper. The bottom is filled with a 1 cm thick layer of white plaster of Paris and, when set, a layer is on the ensures the humidity in the pot and a resting surface without water The pot is closed with a fine gauze and or cap the of which is cut out (Figure The gauze should be fine to prevent the escape of Females are introduced through a small hole in the gauze using a aspirator and the hole is then with a cotton wool use different types of the one for to gravid females, and the one for gravid females. This to about 1 of the bottom per of the pot is mainly for larval like L. in pots while the others, like P. prefer conditions. 25–26° eggs are laid to days blood The of the cycle is mainly by the of blood which within species and Killick-Kendrick and also depends on such as temperature and Volf 2007) and blood source at al. et al. In our L. longipalpis laid eggs on days blood while P. P. and P. laid first eggs day blood from eggs after to days. The day before the and are visible in the as a dark and a dark before is a small of food is in several spots the eggs. food is a of and and are in an but the must be first The could be at for several The of of and is in a layer than inside plastic trays (for example, and saturated with distilled water. with food are in an down position at room temperature in cabinets for as a reservoir filled with water to humidity are placed in a to prevent and out into the laboratory. The front panel of the is and the is under aerobic conditions for to is controlled a and CO2 or other are during of the fermentation (Figure the food is in from the and in an It should be of fungal but of mycelia and of from can be at for several or at for several For some species, like Lutzomyia the food could be by Most species that we prefer food. The of larval food is a factor during early larval For the the food must be a fine This could be ensured by fine gauze the mouth of a vial with food. Pots with larvae are maintained in plastic boxes with a bottom filled with about 1 cm of fine sand with distilled water (Figure For maintenance of sand is than filter paper because of its higher water could be as it is and before the sand layer a fungal on the bottom of the container. Pots are checked at a and and the food is according to the number of larvae and their food to fungal its to and The of food given to larvae and the maintained are during larval larvae their them more this they not any food. The larval period for about (see Lutzomyia P. and P. the larval on days after while P. larvae for at days. differences between colonies of the same species in the geographical In a P. colony from the of larval development days while for colony from it days. differences in P. papatasi colonies from and The period for about to with the by eye and are visible in to In some in from larvae may For this it is difficult to P. or P. in the laboratory. However, we even with colonies from with with P. from and P. from In the P. some larvae for to In P. a long for the significant proportion of larvae and in some pots the generation one The first P. adults between days and a only after a long on days In colonies maintained for many at standard the generation is In the L. longipalpis colony we maintained continuously 1991, life cycle parameters the same in and that the size of the colony to females per In such generation is the first of problems. It be caused by larval or other conditions. The colony from and in a generation than the same colony a adults are from pots into nylon cloth cages with a steel frame (Figure For the maintenance of larger cages cm or are cages prevent damage of adults to into and are for sexes feed on sugar prefer a brown sugar but white sugar or could be used as sugar solution in distilled water is in a and in for several A small piece of cotton wool soaked with a sugar solution is offered on a Petri and changed at a solution is changed as it is more to fungal Females are offered a blood on an animal or twice a For most colonies we use mice, but feeding preferences of various species For example P. P. P. sergenti, P. and L. longipalpis colonies are maintained on only On the other Larroussius and P. and some Adlerius species and P. not feed readily on but prefer hamsters and of such colonies to an blood source is a long which may take many generations. a good experience with mice which are attractive even for colonies that to feed on mice P. females are left undisturbed in a large cage for and then transferred into a small cage for When they are to lay eggs (for most species five to days they are transferred to oviposition pots using an This the early of rearing pots by blood-fed females is laborious but it is in colonies only a small proportion of females take a blood meal. For large colonies with high feeding a is Females are left to in the first cage and then females, with are transferred into the rearing Such a is for Lutzomyia the feeding may and the dark of adults makes the of blood fed females wool soaked with sucrose is placed on top of the nylon of the rearing pot and changed at the of most females lay eggs and The dead females are removed with the few ones can be placed into the big cage the most are the when the colony is for laboratory conditions. this even females readily blood and larvae are not the generation is days than in the established In some colonies the P. papatasi and P. colonies from the larval development in the and On the other some such as P. from larval development for more than to generations. Colonies are in danger of by and by larvae from a larval food are to adults and may cause their et al. in the gut may also with experimental Leishmania infections et al. Pots and larval food are by most by are not in large can damage the colony the larval food and which larval have a good experience with the biological of by of the which are by are commercially available for biological of and are not even to sand fly transmission of infections could be or even by washing the eggs in a solution and or in the of used by et al. eggs are down with distilled water from a rearing pot to filter paper in a to a water is with the water on for to until is eggs are with with the water for by an of by the water on for 1 with the water is used for the eggs are with distilled water with the water on and down into a new this we colonies of L. longipalpis from the et al. the obtained in P. and P. by other species. should be to with these and of cages and pots is a prerequisite for fly interaction and transmission of Leishmania by sand flies by Kamhawi and For experimental of sand flies with two methods can be feeding on and membrane feeding. laboratory animals are mice of the or hamsters into the and into the The used of females is for the of flies to to days for most not while the ones than days for most have a after blood feeding. an the feeding of females that is However, the of blood-fed females that is as there are many that the One of them is a local in the of in the the it is useful to the only and other of the body with a thick sand flies are placed in plastic vials covered at one end with a nylon mesh and the of mice are the mesh with that the vial (Kamhawi et al. et al. females may feed on different sites of the with a different of parasites. The is used mainly in Leishmania species like L. L. or L. high could be the is sand flies on animals with or L. results in which are too for most experimental et al. 1994, et al. 2007). by of by et al. and high Leishmania by et al. in fed on For membrane feeding, or are mixed with blood and offered to flies using a glass This to the of in the infective blood meal. set, this and high of It also to the numbers which may be useful for studies of of glass can be used Higgs and Beaty but we prefer a one with a long body and on the top (Figure 1a). This type more difficult for a to many types of the best results obtained using from days et al. are not recommended for any sand fly species. are removed from the the and the is from the It is with and saline and then at C. before the membrane is in saline and laid the of the feeder with the surface A of the are suspended in the blood that the inner of the feeder under conditions. The feeder is a to the water with through the water at a temperature of one of during membrane feeding. The feeder is placed into the cage through the to prevent sand flies from and the cage in the or could be placed in a blood or or human blood from a blood can be For infections, the blood must be by at for to may from different of animals Leishmania or as from of or and Modi they can be in of with and before For on of Leishmania see for sand fly infections should from a and of to they can be and by of sand flies to Leishmania infections is and to conditions the infective should not per on the feeder are than on the However, the presence of males in the in the temperature of the cages by a dark cloth and by human may feeding The sugar may be also removed from the cage the day before to the feeding Females are left to feed for h, than the feeder is removed a not to any flies one day fed females are and for are kept at with a to the females are for Leishmania are immobilized on or in a are done in drops of but some prefer For on see Those not with should first on flies. have provided colonies and on their rearing when we to work on sand flies. to and and The by the of of the