Lutzomyia longipalpis: an update on this sand fly vector

Abstract

Lutzomyia longipalpis is the most important vector of Leishmania infantum, the etiological agent of visceral leishmaniasis (VL) in the New World. It is a permissive vector susceptible to infection with several Leishmania species. One of the advantages that favors the study of this sand fly is the possibility of colonization in the laboratory. For this reason, several researchers around the world use this species as a model for different subjects including biology, insecticides testing, host-parasite interaction, physiology, genetics, proteomics, molecular biology, and saliva among others. In 2003, we published our first review (Soares & Turco 2003) on this vector covering several aspects of Lu. longipalpis. This current review summarizes what has been published between 2003-2020. During this period, modern approaches were incorporated following the development of more advanced and sensitive techniques to assess this sand fly.

Key words
Lutzomyia longipalpis; sand flies; vector biology; interaction

INTRODUCTION

Lutzomyia longipalpis sensu lato Lutz & Neiva, 1912 is considered the main vector of Leishmania infantum Nicole, 1908 in the American continent (Lainson & Rangel 2005). This species is widely distributed, occurring in diverse ecological niches, such as dry habitats, humid forests but especially in urban and rural areas, where it has successfully established and spread itself (Ximenes et al. 2000, Souza et al. 2009b, Brazil 2013, Rodrigues et al. 2014, Dvorak et al. 2018).

Several components are involved in the urbanization and dispersion of Lu. longipalpis including climatic, environmental and sociocultural factors. This topic has been deeply reviewed by Salomón et al. (2015). Furthermore, the occurrence and the likely geographical distribution of this sand fly in Brazil has been predicted and modeled using geographic information systems and remote sensing (Andrade-Filho et al. 2017).

A great deal of information about Lu. longipalpis has already been reviewed by Soares & Turco (2003), therefore, here we discuss updates throughout the last decades on this sand fly vector, focusing on the information generated from 2003 to early 2020.

Lutzomyia Longipalpis SPECIES COMPLEX AND SEX PHEROMONES

Understanding the evolutionary history of Lu. longipalpis, as well as how geographical barriers and, more recently, anthropogenic environmental changes and activities have contributed to the evolution of sibling species continues to remain a challenge. Combined analyses using molecular markers and behavioral traits such as love songs and pheromones strongly suggest that Lu. longipalpis is a complex species, with distinct population structures as well as reproductively isolated populations (Arrivillaga et al. 2003, 2009, Hodgkinson et al. 2003, Bottecchia et al. 2004, Watts et al. 2005, Balbino et al. 2006, Bauzer et al. 2007, Araki et al. 2009). Details on the current status of the Lu. longipalpis species complex have been reviewed by Souza et al. (2017), especially regarding the historical overview, behavioral traits, courtship song and genetic characteristics of the group. Therefore, factors involved in mating have undoubtedly played (and continue to play) a significant role in maintaining reproductive isolation among the different sibling species.

The most current data on genetic diversity using several molecular markers of Lu. longipalpis indicate the presence of two clades: the first one is composed by Brazilian and Argentinian haplogroups and the second clade includes populations from Central America and northern South America (Guatemala, Honduras, Costa Rica, Colombia and Venezuela) (Pech-May et al. 2018). However, even belonging to the same clade, Argentinian and Brazilian populations present distinct genetic polymorphisms (Araki et al. 2009), resulting in separated populations (sub-clades) using a more refined analysis. A complex population structure of Lu. longipalpis from Brazil has been presented on a geographical scale by Casaril et al. (2019). The presence of geographical barriers may also contribute to divergence and the speciation process that seems to be occurring within the species complex. Genetic studies of Lu. longipalpis provide information about the heterogeneity of vector capacity/competence and vector susceptibility to insecticides as will be discussed later.

Based on the geographical distribution and pheromone types, it is predicted that (S)-9-methylgermacrene-B (9MGB) is the ancestral chemotype in Lu. longipalpis across South America, followed by subsequent speciation to either diterpenes (1S,3S,7R)-3-methyl-α-himachalene (3MαH) or cembrene (CEMB-1 or CEMB-2). To date, populations that produce diterpenes has been found only in Brazilian populations. All pheromone typed species in South and Central America, excluding Brazil, were 9MGB (Spiegel et al. 2016). Within the sibling species of Lu. longipalpis complex, Lutzomyia pseudolongipalpis from Venezuela produces 3MαH (Hamilton et al. 2005, Watts et al. 2005) and Lutzomyia cruzi from Corumbá, Mato Grosso do Sul state, Brazil, produces 9MGB (Vigoder et al. 2010). There is no information about the pheromone type produced by Lutzomyia gaminarai, a species endemic in the southern region of Brazil, occurring in the States of Paraná and Rio Grande do Sul (Galati 2018).

Several aspects on Lu. longipalpis complex still remains an open field to the investigators. Although most studies have focused on the genetic structure of the sibling species, description of pheromones and love songs, the female of Lu. gaminarai has not yet been formally described. Moreover, until today it is not clear how many species or incipient species within the Lu. longipalpis-complex exist in Brazil, or even which species is the original type, since the specimens used to describe this sand fly (Lutz & Neiva 1912) no longer exist. Furthermore, few specimens have been collected in Benjamin Constant, Minas Gerais State, Brazil, the type locality of Lu. longipalpis (Brazil et al. 2006), becoming one of the biggest bottlenecks on sand fly study, given the difficulty to establish a new species-type for the complex and consequently the description of sibling species.

Within the Brazilian populations of Lu. longipalpis, Araki et al. (2009) have proposed to segregate the species complex into two groups: the first one, more homogeneous, representing a single species in which males produce burst-type copulation songs and CEMB-1 pheromones; the other group, more heterogeneous, probably represents incipient species that produce different combinations between pulse-type songs (five patterns of pulse-type) and pheromones such as 9MGB, 3MαH, CEMB-1 and CEMB-2, totaling at least six sibling species (Vigoder et al. 2015). Genetic evidence suggests that introgressive hybridization has been a crucial phenomenon of the recent speciation process that occurs within the Lu. longipalpis complex (Araki et al. 2013). Microsatellite data have shown limited genetic flow and introgression between Lu. longipalpis and Lu. cruzi in which the divergence level was similar to that observed among Brazilian populations of Lu. longipalpis (Vigoder et al. 2010, Lins et al. 2012, Santos et al. 2013). However, data from 12S rDNA sequencing did not differentiate Lu. longipalpis from Lu. cruzi (Corumbá), suggesting that the speciation process is recent or still occurring (Ribolla et al. 2016). Nevertheless, more genetic data is needed to confirm the occurrence of the recent speciation process between Lu. longipalpis and Lu. cruzi. Moreover, introgression patterns in the genome seem to have a relevant effect on transmission dynamics of Leishmania parasites. Therefore, exploring these aspects on Lu. longipalpis complex may be a good way to understand the vectorial capacity of the sibling species (Araki et al. 2013). Distinct genetic composition of populations from Espírito Santo, Brazil, seems to affect their susceptibility to Leishmania or even the capability to transmit the pathogen in an anthroponotic environment by the low adaptability of Lu. longipalpis to this environment (Rocha et al. 2011). Although a lot of papers have focused on establishing the genetic and pheromone variations in the Lu. longipalpis species complex, there is still a gap in how those variations affect interaction with Le. infantum. It would be extremely important to address the vectorial competence of a given Lu. longipalpis population. Although it is a permissive vector, intra-populations variability may result in a lack of interaction between the parasite and the vector. This was the case of allopatric populations of Nyssomyia umbratilis collected in the south and north of Negro River in the Amazon (Soares et al. 2018). In this paper, using the in vitro system, the authors observed that the south population was refractory to interaction with Le. guyanensis. However, we do not know if such a phenomenon would occur in Lu. longipalpis and this would be a very interesting direction of further molecular and biochemical studies.

The sympatric populations from Sobral, State of Ceará, Brazil, have been deeply studied, focusing on the genetic, evolutionary and epidemiologic significance of the one-pair-of-spots (S1) and two-pairs-of-spots (S2) male phenotypes of Lu. longipalpis. Lins et al. (2008) have identified a clear difference between these populations using the paralytic (para) gene as well as an association of the para and the resistance to pyrethroid insecticides. Further studies on genetic polymorphisms in period gene (per) have also suggested the presence of two sibling species in Sobral (Costa-Júnior et al. 2015). This data added crucial information about these reproductive isolated populations, suggesting the importance of premating barriers in Lu. longipalpis sibling species speciation (Maingon et al. 2003). Additionally, genetic divergence in the cacophony gene (cac) showed that S2 population is more related to Natal population (both produce burst type and CEMB-1) whereas S1 (pulse type 3 and 9MGB) was closer to Jacobina (pulse type 1 and 3MαH) and Lapinha (pulse type 2 and 9MGB). The genetic diversity observed in S1 and S2 may also reflect distinct physiological and behavioral aspects for both populations. However, until today, there is a lack of information on host-parasite interaction comparing sympatric populations as S1 and S2. The genetic divergence between these populations may affect the interaction with Le. infantum. Although few variations have been observed, both males and females from the S2 population seem to initiate their crepuscular activity a little earlier than S1 (Rivas et al. 2008). However, more studies are needed to confirm distinct patterns of hourly activity as well as other differences in biological behavior between these populations. Besides their circadian rhythms, also the pheromones and patches were shown to affect bionomic aspects of Lu. longipalpis. Populations that produce homosesquiterpene (C16), such as sand flies from Jacobina (3MαH), Lapinha and Sobral one spot (1S) (both 9MGB) seems to be more easily adapted to the colonization conditions than the population whose males produces diterpenes (CEMB-1) such as the sand flies from Natal and Sobral two spots (2S) (Souza et al. 2009a). Since colonization is an important aspect that hinders sand fly studies, a better knowledge of those pheromones could also help to choose a more productive colony.

Finally, although most of the studies focused on patches occurred in Sobral, those phenotypes were also detected in other states. For example, S2 male phenotype was found in Jaíba (Minas Gerais State), Estrela de Alagoas (Alagoas State), Raposa and Codó (Maranhão State) (Araki et al. 2009). Consistent with the studies in Sobral, Silva et al. (2011) have shown genetic polymorphisms between Raposa and Codó sympatric populations, suggesting a clear segregation related to spot phenotypes (Lins et al. 2008, Costa-Júnior et al. 2015).

In conclusion, a large number of papers published before 2003 have addressed the pheromones and the genetic aspects of the Lu. longipalpis complex. Since 2003 those numbers have decreased, probably due to the acceptance of the species complex idea. How such variations affect the interaction with Le. infantum is an open field still needed to be explored by the investigators.

Lutzomyia Longipalpis CONTROL

Although nowadays it is still difficult to control the sand flies vector populations, important tools have been arising to improve the strategies, especially for VL control. The first problem is the difficulty to find the larval stages in the environment (Casanova 2001, Sangiorgi et al. 2012). In a field evaluation using an adulticide-larvicide mixture (100 mg of permethrin and 2 mg/m2 of pyriproxyfen), a significant decrease in the number of Lu. longipalpis was reported for at least two weeks (Juan et al. 2016). However, further studies are needed to evaluate the persistence of the residual effect of pyriproxyfen in controlling Lu. longipalpis larvae. For this reason, most of the studies have focused on the adult stages. Volatile compounds based on male pheromones and kairomones have demonstrated a good efficacy if used combined with automatic light traps improving catch rates, especially for Lu. longipalpis. Furthermore, synthetic pheromones can feasibly improve the efficacy of sand fly control programs when used alongside insecticides. This combined strategy attracts and kills both sexes, preventing host-seeking females from transmitting Le. infantum and males from establishing alternative aggregation sites elsewhere (Bray et al. 2009). A decrease in the number of sand flies attracted usually occurs as a consequence of insecticide treatments, however, the application of synthetic pheromones into insecticide-sprayed experimental sheds seems to prevent and reverse it, improving the catch rates of Lu. longipalpis (Bray et al. 2010). The number of pheromone-lures seems to have an influence on the effectiveness of this strategy to attract sand flies. Bell et al. (2018) have shown that increasing the number of lures results in an upward trend in the number of sand flies that are caught in the field, especially males. Kairomones have been extensively used to attract hematophagous insects, such as mosquitoes and tse tse flies, however, there are few studies focusing on sand fly attraction. The compounds 1-octanol, a volatile component of bovine and human breath, and 1-nonanol, a volatile from cattle urine, elicited the highest attractiveness response in Lu. longipalpis adults in a dose-dependent manner (Magalhães-Junior et al. 2014). However, these alcohols have been identified at small levels in human breath or skin odors, which may justify the lack of interest in their potential role as an attractant for sand flies (Magalhães-Junior et al. 2014).

Although the chemical attraction has been the newest tool in this field, sand fly control programmes still often rely on spraying potential resting sites (intra or peridomiciliary sites) with residual insecticides, especially pyrethroids as lambda-cyhalothrin (Feliciangeli et al. 2003, Camargo-Neves et al. 2007a), deltamethrin (Santini et al. 2010), alpha cypermethrin (Pessoa et al. 2015) and permethrin (Alexander et al. 2009), with varying effectiveness. However, spraying also requires training to be conducted effectively, in order to ensure that the correct concentration of insecticide is applied, minimizing exposure to sub-lethal amounts which might promote the onset of resistance to several compounds. Denlinger et al. (2015) have quantified the insecticide susceptibility in laboratory-reared Lu. longipalpis to ten insecticides, comprising four chemical classes: pyrethroid, organophosphate, carbamate and organochlorine. The organophosphate insecticides caused delayed mortality in the sand fly population, while carbamate caused mortality faster. Both insecticides classes have similar modes of action, and, despite the differences in killing rates for carbamates and organophosphates, Lu. longipalpis are most susceptible to bendiocarb and propoxur carbamates as well as to the organophosphate fenitrothion (Denlinger et al. 2015). Furthermore, the doses for each insecticide have been determined using the CDC bottle bioassay to assess Lu. longipalpis resistance, providing starting points to test on field populations (Denlinger et al. 2016).

The use of insecticide-impregnated nets has also been used as a complementary tool for sand fly control, especially Lu. longipalpis. The entomological efficacy of 25% deltamethrin EC insecticide-treated bednets has been evaluated by Courtenay et al. (2007), in a crossover field study in Amazon Brazil (Marajó Island, State of Pará). Compared with untreated nets, the insecticide ones increased the barrier effect of the nets by 39%, reduced human landing rates by 80% and increased the 24 hours mortality rate (Courtenay et al. 2007). The lambda-cyhalothrin seems to have a short residual effect, whose efficacy declined to 74% after six months. On the other hand, permethrin-impregnated nets maintained its effectiveness close to 100% lethality 24 hours post exposure for at least a year under laboratory conditions (Bray & Hamilton 2013). However, those conditions may not be possible in the field. Trials using a variety of indoor and outdoor surfaces are needed to confirm the effectiveness of this netting-treatment protocol in the field, especially close to animal shelters.

A distinguished feature in Lu. longipalpis populations is their ability to respond differently to the action of pyrethroids and organophosphates. For example, Montes Claros sand flies were most susceptible to malathion, fenithorion and deltamethrin, while those from Lapinha were most susceptible to cialotrhin, malathion and permethrin in laboratory conditions (Alexander et al. 2009). Moreover, a significant reduction in the susceptibility to the insecticides reinforced the importance of developing tools for detecting resistance (Alexander et al. 2009). Since the efficacy of insecticides differ within Lu. longipalpis populations, the combined use of insecticides may be a better strategy for the sand fly control. In this context, the repellent efficacy of a spot-on topical combination of fipronil and permethrin has been evaluated in dogs (Cutolo et al. 2018). A significant repellent effect against Lu. longipalpis as soon as it was applied on the dogs and high protection rates for 28 days has been shown. However, due to the short anti-feeding effect, regular application in dogs may hinder its protective effect in VL-endemic areas (Cutolo et al. 2018). Likewise, the 4% deltamethrin-impregnated canine collar (ICC) has not presented a long-lasting effect compared with spot-on topical repellents; however, the ICC is currently being considered as a relevant tool for VL control (Albuquerque e Silva et al. 2018). The ICC tends to reduce the prevalence of canine VL, in two basic ways: 1) reducing the blood feeding by the vector and, 2) reducing the vector population, mediated by repellent and insecticidal action of deltamethrin (Coura et al. 2019). The use of ICC reduced the number of Lu. longipalpis captured in an interventional area in Montes Claros, State of Minas Gerais (14% of reduction) and Fortaleza, State of Ceará (60% of reduction). Moreover, a 40% decrease in canine VL prevalence has been reported in both municipalities (Albuquerque e Silva et al. 2018). The anti-feeding effect of ICC has also been reported in Europe for Phlebotomus perniciosus, the vector of Le. infantum (Maroli et al. 2001, Manzillo et al. 2006, Ferroglio et al. 2008). Until today, there are few studies in Brazil that evaluate the efficacy of this strategy in the field. Longer follow-up studies on how ICC affects vector population and its impact on VL cases are needed. Considering the importance of protecting dogs from sand fly bites, it would be interesting to evaluate the potential role of mass use of ICC as a strategy to reduce canine visceral leishmaniasis incidence. However, the short-lasting effect, the need to frequently replace the ICC, and local symptoms in dogs, are still problems to be solved.

Although insecticide-based control measures are available for sand flies, there is still an urgent need for novel and alternative methods that do not affect or are less harmful to the environment. In this context, biological control could represent an important initiative for future studies. The combined use of chemical insecticides and selective pathogens may increase the efficiency of insect control. In this way, a possible alternative to current strategies may be the biological control of the vector using the entomopathogenic fungi Beauveria bassiana. Amóra et al. (2009) report that Lu. longipalpis eggs infected with this fungus reduced the hatching to 59%, suggesting a pathogenic potential on both larvae and adults. Moreover, Metarhizium anisopliae var. acridum, another entomopathogenic fungal, was harmful to sand flies in the adult stage (Amóra et al. 2010). Even in the laboratory, the studies on entomopathogenic fungi are very scarce. This reinforces the need for more studies on the impact-cost of such organisms while applying them in the field for controlling sand flies.

Several studies have investigated the use of plants to control vector-borne diseases. Plants from the Meliaceae family (Azadirachta indica) have been deeply studied due to their effects against many insects, especially those of agricultural importance. However, few studies have focused on sand flies. Few ovicidal and larvicidal effects have been reported even in high concentration of A. indica oil when Lu. longipalpis eggs and larvae were treated in laboratory conditions (Maciel et al. 2010). On the other hand, the triterpenoid azadirachtin seems to block the metamorphosis when added to larval food of Lu. longipalpis (Andrade-Coelho et al. 2006). Studies have also showed that A. indica and Melia azedarach fruit and leaves in natura significantly increased larval mortality in comparison to untreated insects (Andrade-Coelho et al. 2009). Azadirachtin also seems to affect Lu. longipalpis oviposition and may increase the mortality in adults, indicating that azadirachtin may be a potent sterilizer that could be used against the development of Lu. longipalpis populations (Andrade-Coelho et al. 2014).

In conclusion, there are few field studies that have evaluated the impact of biological controls against sand fly vectors. Although distinct classes of insecticides are available, sand fly resistance has been reported in Brazil and other endemic countries (Surendran et al. 2005, Lins et al. 2008, Hassan et al. 2012). Thus, while studies on sand fly control are extremely relevant, other strategies than chemical control are necessary.

FOOD SOURCE IDENTIFICATION

During the past decades, several studies have been performed to identify the blood source of engorged females of potential and proven vectors such as Lu. longipalpis. Initially, the precipitin test was the most common technique to identify blood meal (Dias et al. 2003, Camargo-Neves et al. 2007b, Missawa et al. 2008) and ELISA (Marassá et al. 2006, Afonso et al. 2012). However, those techniques have some limitations, such as the need to know the previous local fauna and consequently obtain the specific antisera. Further, molecular methods (PCR and DNA sequencing) gradually replaced those techniques, improving blood meal identification by using CytB as universal primers (Sant’Anna et al. 2008, Soares et al. 2014, Carvalho et al. 2017b).

Lutzomyia longipalpis has broad-range feeding habits due to their adaptation to different habitats in both intradomiciliary and peridomiciliary sites. Several authors have reported that this vector fed on dogs, cats, pigs, cattles, horses, chickens and synanthropic vertebrates (rats and opossums). With the exception of chicken, most of the aforementioned hosts are potential reservoirs of Leishmania (Dias et al. 2003, Marassá et al. 2006, Camargo-Neves et al. 2007b, Missawa et al. 2008, Sant’Anna et al. 2008, Afonso et al. 2012, Soares et al. 2014, Carvalho et al. 2017b). Although chickens are refractory to Leishmania infection, Sant’Anna et al. (2010) have shown that, this vertebrate provides valuable blood sources to support the Lu. longipalpis population in peridomestic sites. The quality of chicken blood supports the development of transmissible Leishmania infections in Lu. longipalpis (Sant’Anna et al. 2010).

Besides blood, both females and males feed on plant-derived sugar meals as a source of energy. Sugary solutions such as nectar or honeydew (secreted by plant-sucking homopteran insects) and phloem sap are ingested by sand flies by probing plant tissues with their mouthparts. Many studies have addressed Lu. longipalpis plants preference. DNAs from Anacardiaceae, Meliaceae and Fabaceae families have been detected in the sand flies (Lima et al. 2016). More recently, the source of sand fly plant meals based on next generation sequencing (NGS) of chloroplast DNA gene ribulose bisphosphate carboxylase large chain (rbcL) was assessed. Interestingly, the predilection of several sand fly species such as Lu. longipalpis for feeding on Cannabis sativa, a presumably illegal plant in some countries, was found (Abbasi et al. 2018). However, there is still a lack of knowledge on how specific sugars from plants may affect Leishmania development in sand flies. It is already known that besides functioning as a source of energy, sugars may also be used by Leishmania during its establishment in the midgut.

MIDGUT PHYSIOLOGY

The sand fly gut is divided into three main regions: the foregut, the midgut, and the hindgut. The cardia separates the foregut from the midgut and the pyloric valve separates the midgut from the hindgut (Bates 2008). Most studies have focused on host-parasite interaction of suprapylarian Leishmania species (Assis et al. 2012). This development is restricted to the portion of the gut anterior to the pylorus, mainly in the thoracic and abdominal midgut (Lainson & Shaw 1987). This is different from Viannia species whose development occurs in the hindgut prior to migration to anterior parts. On the other hand, the mode of the gut development is poorly recognized by the subgenera Mundinia and Sauroleishmania (Espinosa et al. 2018). For this reason, more studies on how species from these subgenera behave in their respective vectors are needed. In this context, an early study (Luz et al. 1967) reported a suprapylarian development for Le. enriettii in Pintomyia monticola, the suspected vector. However, this species, together with Le. orientalis did not developed very well in Lu. longipalpis (Seblova et al. 2015b, Chanmol et al. 2019). Thus, studies with their suspected vectors (phlebotomine sand flies and/or ceratopogonids) can help to clarify this subject and are fertile fields for entomologists.

Molecular studies have contributed to understanding the events that occur during the establishment of Leishmania infection in sand flies (Ramalho-Ortigão et al. 2010). Leishmania molecules such as LPG (Pimenta et al. 1994, Svárovská et al. 2010), which binds to the sand fly midgut galectin receptor PpGalec (Kamhawi et al. 2004), sand fly digestive enzymes (Borovsky & Schlein 1987, Schlein & Jacobson 1998, Sant’Anna et al. 2009, Telleria et al. 2010) and the peritrophic matrix (PM) (Pimenta et al. 1997) contribute to the success of the infection. The PM is a chitinous structure that envelopes the bloodmeal along the entire midgut, separating the ingested food from the midgut epithelium. In most sand flies, this structure is formed between 12-24 h after blood ingestion and degraded after 72h, when digestion is completed (Secundino et al. 2005, Sádlová & Volf 2009). For more information about the Lu. longipalpis PM structure, composition, degradation and synthesis kinetics see Secundino et al. (2005). The authors also have described a midgut muscle network of Lu. longipalpis.

The PM degradation after blood digestion requires the activity of chitinases, which cleave the chitin microfibril components of the matrix. Although Leishmania chitinase is believed to take part in the escape of the parasite from the PM, it is likely that a sand fly-derived chitinase may also be involved. Ramalho-Ortigão & Traub-Csekö (2003) have isolated and characterized a cDNA encoding a chitinase (Llchit1) from midgut of Lu. longipalpis. Messenger RNA expression indicates that this gene is induced upon blood feeding and reaches a peak at approximately 72h post blood meal, presuming that this sand fly chitinase has a function in PM degradation (Ramalho-Ortigão et al. 2005). Besides that, Ortigão-Farias et al. (2018) have shown that alternative splicing generates chitinases with different domain structures. LlChit1A is present in adult females post blood meal, L4 larvae and pre-pupae, whereas LlChit1B and LlChit1C are found in L4 larvae and disappear just before pupation.

Serine proteases (trypsins and chymotrypsins) are the most abundant digestive enzymes in the midgut of sand flies. In addition to blood digestion, those proteases have been implicated in Le. infantum establishment in their respective insect vector, appearing to be detrimental to parasite survival during the first 48 hours prior to their increase after this period (Freitas et al. 2012). However, the same detrimental effect has not occurred in sand flies infected with L. major and L. donovani, suggesting that Leishmania mortality is not caused directly by sand fly proteases, but from toxic products of blood meal digestion. (Pruzinova et al. 2018). More studies are needed to better understand the effect of proteases on Leishmania establishment within sand flies. The opposite data may indicate distinct effects of proteases in Leishmania species. Sant’Anna et al. (2009) have reported that Leishmania mexicana was able to downregulate the trypsin secretion in Lu. longipalpis to its own advantage, promoting their establishment in the midgut. Likewise, a decrease of trypsin enzymatic activity in Lu. longipalpis infected by Le. infantum has been reported (Telleria et al. 2010). Lutzomyia longipalpis trypsin 1 gene knockdown through dsRNA microinjections into the thorax of females, seems to enhance the survival of Le. mexicana in comparison with mock-injected controls. Altogether, those data reinforce the inverse relationship between the expression and production of trypsin and the establishment of Leishmania in the sand fly midgut (Sant’Anna et al. 2009). Telleria et al. (2007) have identified and characterized two cDNAs, Lltryp1 and Lltryp2, coding for trypsin-like proteins in Lu. longipalpis. Lltryp1 expression remains undetected until blood feeding and reaches a peak at 12h post-blood meal, returning to pre-blood meal levels after 72h. Lltryp2, on the other hand, is constitutively expressed at high levels in the non-blood fed female but is reduced upon blood feeding. At the end of the digestive cycle, Lltryp2 regains its pre-blood meal levels (Telleria et al. 2007). The pattern of trypsin expression in Lu. longipalpis differs from the results obtained for the Old-World species Phlebotomus papatasi (Ramalho-Ortigão & Traub-Csekö 2003). However, there is still lack of information on how proteases from Ph. perniciosus and from other natural vectors affect the development of Le. infantum. The transcriptome analysis has demonstrated that L. infantum infection can reduce the transcript abundance of trypsin PperTryp3 in the midgut of Ph. perniciosus (Dostálová et al. 2011). Although Lu. longipalpis and Ph. perniciosus may sustain infection with Le. infantum, the kinetics of proteases in those vectors in parallel are yet to be determined.

Studies on midgut pH as well as the mechanisms involved in pH control are extremely relevant, since Leishmania develops exclusively in the sand fly gut and the digestive processes are essentially enzymatic (Bates & Rogers 2004). There are three known mechanisms involved in the process of controlling gut pH. The first involves the loss of CO₂ from ingested blood and the transport of different ions through the plasmatic membrane of the enterocytes (Santos et al. 2008). Other physiological processes related to the alkalinization of the abdominal midgut involves the presence of blood in the abdominal midgut composed by proteins and amino acids. Those components cause midgut endocrine cells to release alkalinizing hormones, increasing gut pH favoring blood digestion (Santos et al. 2011). The third mechanism act involves Proton-Assisted Amino Acid Transporter (LuloPATs), removing H+ ions from the gut lumen into the cytoplasm of the enterocytes (Nepomuceno et al. 2020). However, alkalinization of the lumen may occur by the entry of some amino acids into the cytoplasm of enterocytes triggering a luminal alkalinization mechanism independent of LuloPATs (Nepomuceno et al. 2020). Some reports along the decades have shown the influence of Leishmania on sand fly physiology and such behavior most likely evolved to favor the development and transmission of the parasite. Leishmania infantum is able to reduce the alkalinization in the vector midgut, decreasing the activity of proteases like trypsin, resulting in a decreased supply of amino acids to the enterocytes favoring the development of the parasites during digestion (Santos et al. 2014).

There are few studies regarding midgut physiology of Lu. longipalpis larvae. The anatomy of the digestive tube of Lu. longipalpis larvae as well as the pH along the midgut have been described in Vale et al. (2007). The carbohydrases α-amylase, present in the anterior midgut and probably involved in the digestion of glycogen; α-glucosidase, that completes the digestion of glycogen in the posterior midgut, and a membrane bound trehalase, that probably acts in the digestion of trehalose, seems to be the most abundant within the midgut of the larvae (Moraes et al. 2012, Vale et al. 2012). The expression pattern of glycoside hydrolase genes in Lu. longipalpis larvae have been described by Moraes et al. (2014), where the catabolism of microbial carbohydrates in insects generally involves β-1,3-glucanases, chitinases and digestive lysozymes. This is interesting because Le. infantum LPG possess terminal β-1,3-glucoses that could be cleaved by those enzymes and perhaps contribute to the sand fly midgut sugar milieu (Soares et al. 2002, Coelho-Finamore et al. 2011).

Early studies of Elnaiem have already focused on the effect of a second blood meal in the development of Lu. longipalpis (Elnaiem et al. 1992, 1994). Nowadays, most of the studies are interested in how a second bloodmeal affects Leishmania development. In this context, the effects of sequential blood meals on longevity, protein digestion, trypsin activity and Leishmania development within Lu. longipalpis midgut have been recently evaluated. The mortality of blood-fed females increases after a second blood meal as compared to sugar-fed females and the trypsin activity was lower during the second gonotrophic cycle (Moraes et al. 2018). The authors have not observed difference in the population size of Leishmania in the gut with sequential blood meals. However, Serafim et al. (2018) have reported that sequential blood meals promoted Leishmania replication and reversed metacyclogenesis to a leptomonad-like stage, the retroleptomonad promastigote, enhancing the Lu. longipalpis infectivity. Needless to say, this paper was a landmark study, bringing new information on parasite development after a second blood meal.

Salivary proteins

In general, female sand flies, except autogenic species, need to ingest blood for egg development and sugar for energy metabolism. Saliva is essential in both types of feeding, playing different roles since it contains sets of enzymes for blood and sugar feeding, as α-amylase (Cavalcante et al. 2006). Early studies by Volf have shown the effect of salivary gland proteins in Old World sand flies, in which the composition of sand fly saliva depend not only on sex, but also on the physiological state of the female (Volf et al. 2000). The salivary protein composition of Lu. longipalpis also depends on age and diet (Prates et al. 2008). The protein content from unfed sand flies increased 94% from the first to the fifth day after emergence and such variation can be related to the synthesis of important enzymes for meal ingestion and initial digestion (Prates et al. 2008). A kinetic of protein content in salivary glands seems to occur after the blood meal, in which a depletion of total protein content has been observed with gradual increase in subsequent days, returning to similar basal values (Prates et al. 2008). The findings of Volf for Old World sand flies and the further records of Prates for New World ones could be generalized for sand flies worldwide, in view of the salivary content appears to follow the same pattern in several sand fly species.

Blood-feeding causes tissue damage creating a hemorrhagic pool resulting from probing and destruction of small capillaries. In this environment Leishmania and saliva interact with different host cells including peripheral blood and resident cells in the skin (Vasconcelos et al. 2014). It has been well documented that sand fly saliva possesses an array of potent pharmacological components, such as anticoagulants, anti-platelet, vasodilators, immunomodulators and anti-inflammatory molecules. For more details about the inflammatory role of Lu. longipalpis saliva in leishmaniasis see Prates et al. (2012).

To know the effect of these molecules, most studies on sand fly saliva have used experimental animals (mice), and to a lesser extent human cell. Salivary gland homogenates (SGH) of Lu. longipalpis induce an increase of IL-6, IL-8 and IL-12p40 and inhibits TNF-α and IL-10 production by human monocytes. SGH have also influenced the expression of cell surface molecules such as MHC class II, CD80 and CD86 on antigen-presenting cells, except on dendritic cells, representing a critical point for the development of a protective Tcell response (Costa et al. 2004). Moreover, SGH seem to increase the IL-17 expression in human peripheral blood mononuclear cells (Teixeira et al. 2018). Human volunteers exposed to laboratory-reared Lu. longipalpis bites developed both humoral and cell-mediated immune response against sand fly saliva, presenting increased frequency of CD4+CD25+ and CD8+CD25+ T cells as well as IFN-γ and IL-10 synthesis (Vinhas et al. 2007) and moreover, inducing heme oxygenase-1 expression at bite site (Luz et al. 2018). These studies confirm powerful immunomodulatory properties of saliva and help clarify how Leishmania takes advantage of them during the bite.

BALB/c mice exposed to repeated Lu. longipalpis bites have developed a diffuse inflammatory infiltrate characterized by neutrophils, eosinophils, and macrophages when challenged with SGH (Silva et al. 2005). Antibodies anti-saliva have also been detected in exposed mice, that presented significant increase of IgG and IgG1, but not IgG2a or IgG2b, suggesting a predominant Th2 response with a putative role for immune complexes in cell recruitment (Silva et al. 2005). Lu. longipalpis saliva is also capable of inducing neutrophil and macrophage recruitment and of modulating their function (Silva et al. 2005, Teixeira et al. 2005, Araújo-Santos et al. 2010, Prates et al. 2011, Carregaro et al. 2013). Neutrophil and macrophage activity seem to be impaired in the presence of saliva resulting in cell apoptosis, production of PGE2 and LTB4 promoting increased parasite survival (Monteiro et al. 2005, Araújo-Santos et al. 2010, Prates et al. 2011). Lutzomyia longipalpis saliva enhances Le. amazonensis infection affecting the macrophage function by upregulation of IL-10 and downregulation of NO production (Norsworthy et al. 2004). The same regulation pattern of immune response has been described in BALB/c mice experimentally infected with Le. major, in which a considerable increase of IL-10 and IFN-γ was detected, inducing preferentially type-2 cytokines and the sequential migration of neutrophils, eosinophils, and CD4+ CD45RB^low cells (Monteiro et al. 2007). However, Laurenti et al. (2009) have reported that SGH from wild-caught Lu. longipalpis have determined lower production of IL-4 and IL-10 but higher IL-12 levels in C57BL/6 compared with laboratory-reared SGH. These findings may indicate a probable bias by using SGH from laboratory-colonized sand flies instead of wild-caught vector SGH. In addition, it indicates differences on immune response of the most used experimental models for studies concerning saliva effects (Laurenti et al 2009). However, it is important to note that sand flies also inject their microbiota together with the salivary content, and the presence of distinct bacteria within laboratory-colonized sand flies compared to wild-caught ones, can also influence the immune response. The presence of SGH from Lu. longipalpis was able to differentially modulate the course of the lesion and macrophage differentiation in Cavia porcellus caused by avirulent and virulent Le. enriettii strains (Pinheiro et al. 2018). Several basic studies, especially those that used needle models, were very important for understanding the Leishmania infection. However, there is an urgent need that from now on, transmission needle studies use saliva at least from a colonized sand fly vector. Although, for obvious reasons, it is not possible to use the natural pairs depending on the Leishmania species, most of the properties of the saliva of different sand flies share similar effects.

Most of the studies above have used SGH, but it seems that the search for specific molecules has been the target for by several groups. Consistent with this observation, the structure and function of LJM11 has been described by Xu et al. (2011). A protective immunity driving a strong Th1 type immune response was observed in immunized C57BL/6 mice infected with Le. major (Xu et al. 2011) and in BALB/c mice infected with Le. braziliensis (Cunha et al. 2018). Immunization with salivary protein LJM19 induced protection in hamsters challenged with Le. braziliensis (Tavares et al. 2011). The presence of smaller lesion sizes as well as reduced parasite burdens both at lesion sites and in the draining lymph nodes, was associated with a significant decrease in the expression levels of IL-10 and TGF-β and increased IFN-γ expression have been reported (Tavares et al. 2011). Both LJM17 and LJL143-immunized dogs have presented a mixed (Th1/Th2) immune response and moreover, increased IFN-γ production (Abbehusen et al. 2018), providing immune responses qualitatively similar to those previously obtained by Collin et al. (2009). Although knowing specifically the activity of a given molecule, the use of several antigens that do not exhibit antagonistic properties could help the development of more potent saliva-based vaccines.

Valenzuela et al. (2004) have isolated and identified the most abundant secreted proteins from the salivary glands of Lu. longipalpis using massive cDNA sequencing, proteomics and customized computational biology approaches. However, several proteins coded by their corresponding salivary gland transcripts remain without a defined function until today (Valenzuela et al. 2004, Anderson et al. 2006). Likewise, some biological functions described in the salivary gland have not been associated with a specific protein. For example, the anticoagulant of Lu. longipalpis remained elusive for decades until Collin et al. (2012) describe Lufaxin (Lutzomyia longipalpis Factor Xa inhibitor). This recombinant protein has potent and specific anticoagulant activity toward FXa, impairing protease-activated receptor 2 activation and, consequently inhibiting the inflammation and thrombosis in C57BL/6 mice. New insights of recombinant hyaluronidase (LuloHya) and Lutzomyia NET destroying protein (Lundep), the proteins responsible for the hyaluronidase and endonuclease activities have been described (Chagas et al. 2014, Martin-Martin et al. 2018). Lundep seems to increase Le. major survival, destroy neutrophil traps and inhibits XIIa contact activation in human plasma. The relationship between Leishmania parasites and sand flies hyaluronidase was first described by Volfova et al. (2008). The authors have shown that co-inoculation of parasites with hyaluronidase enhances Leishmania infection. Altogether, those data indicate that saliva is an endless subject and several factors are still to be defined and how to block those molecules is an open field for alternative tools against transmission.

Lutzomyia longipalpis is able to feed on several mammal and bird species (Afonso et al. 2012). For this reason, an arsenal of complement inhibitors is needed to protect this species. In this context, Lu. longipalpis saliva was able to inhibit the serum complement activation from a wide range of vertebrates, including dogs, guinea pigs and rats (Mendes-Sousa et al. 2013). Studies involving the human complement inhibition by Lu. longipalpis saliva have shown at least two inhibitors of the classical pathway in this species. The first is a Salivary Anti-complement from Lu. longipalpis (SALO) (Ferreira et al. 2016), considered a leishmaniasis vaccine candidate (Asojo et al. 2017) and the second, a soluble intestinal inhibitor (Saab et al. 2020).

One of the most studied salivary peptides is the potent vasodilator maxadilan (MAX). MAX also seems immuno-modulate the host immune response. MAX treatment reduced the surface expression of CD80 on CD11c⁺ dendritic cells and resulted in a concomitant increase in CD86 expression on a subpopulation of these cells. Moreover, MAX seemed to upregulate the cytokines associated with a type-2 response (IL-10, IL-6, and TGF-β) and downregulated type-1 cytokines (IL-12p70 and TNF-α), NO and CCR7. This enhanced parasite survival in the vertebrate host in the early stages of infection (Brodie et al. 2007, Wheat et al. 2008). MAX was also able to drive plasma leakage via PAC1–CXCR1/2-pathway (Svensjö et al. 2009, 2012). A protective effect against Le. major infection in murine models has also been reported for MAX (Wheat et al. 2017).

Anti-saliva antibodies can be used to assess exposure of humans and other Leishmania hosts to sand fly bites (Rohousova et al. 2005, Bahia et al. 2007, Vinhas et al. 2007, Hostomska et al. 2008, Fraga et al. 2016). These anti-saliva antibodies seem to be species-specific as shown by Volf & Rohousova (2001) and Rohousova et al. (2005). The antibodies of hosts bitten by Old-World sand flies did not cross-react with Lu. longipalpis SGH. Therefore, this specificity of anti-saliva antibodies enables to measure/estimate the exposure to a particular species. Also, the protective effect of immunization by saliva have been species-specific as shown by Thiakaki et al. (2005): mice have been protected against co-inoculation of Leishmania with Lu. longipalpis saliva only if they were preimmunized by SGL of Lu. longipalpis but not if preimmunized by SGL of Phlebotomus species. Nine recombinant salivary proteins were developed and tested for immunogenicity and specificity in mammalian hosts (Teixeira et al. 2010). The recombinant proteins LJM17 and LJM11, both belonging to the insect “yellow” family of proteins, were potential markers of exposure to sand fly bite (Souza et al. 2010). LJM17 was recognized by human, dog, and fox sera and LJM11 by humans and dogs. Notably, LJM17 and LJM11 were specifically recognized by humans exposed to Lu. longipalpis but not by individuals exposed to Nyssomyia intermedia (Teixeira et al. 2010). A recent paper has shown that one of the salivary proteins of Ny. intermedia, LinB-13, could be a useful marker for the development of a more severe cutaneous leishmaniasis (Carvalho et al. 2017a). This study opens the possibility that similar mechanisms could also happen in the viscerotropic Leishmania species transmitted by Lu. longipalpis, especially in canine infection, that is a very susceptible host compared to humans.

HOST-PATHOGEN INTERACTIONS

Laboratory studies on sand fly competence to Leishmania parasites suggest that the sand flies fall into two groups. Several species are termed specific/restricted vectors that support the development of one Leishmania species. On the other hand, permissive vectors are susceptible to various Leishmania parasites (Volf & Myskova 2007, Dostálová & Volf 2012). The presence of the permissive vector Lu. longipalpis in Latin America was crucial for the establishment of L. infantum from Mediterranean to this continent (Volf & Myskova 2007). Another factor that seems to affect the establishment of Leishmania in sand flies is the temperature. Leishmania infantum and Le. braziliensis have developed well in Lu. longipalpis at 20 and 26 degrees C, while Le. peruviana, a mountain species, developed well in sand fly females kept at 20 degrees C (Hlavacova et al. 2013). Previous studies have suggested that for ‘specific’ vectors, successful parasite development is mediated by parasite surface glycoconjugates and sand fly lectins. However, Myšková et al. (2007) have shown that interactions involving ‘permissive’ vectors, as Lu. longipalpis utilize other molecules of the midgut epithelium as a parasite ligand. The Helix pomatia agglutinin (HPA), a lectin specific for terminal N-acetyl-galactosamine (GalNAc) present on O-linked glycoconjugates, bound to midgut proteins from permissive but not from specific vectors (Myšková et al. 2007). The characterization of O-linked glycoconjugate of Lu. longipalpis has revealed the presence of mucin-like properties, GPI-anchored in the membrane of enterocytes and localized it on the luminal side of the midgut (Myšková et al. 2016).

As Leishmania undergo metacyclogenesis and acquire infectivity within the sand fly gut, they secrete a unique class of serine-rich proteophosphoglycans (PPGs); which condense to form a gel in which the parasites are embedded (Rogers & Bates 2007). PPGs are synthesized by all species of Leishmania in vitro and the promastigote secretory gel (PSG) has been observed in all Leishmania-sand fly combinations examined to date. The Le. infantum PPGs regurgitated by the bite of Lu. longipalpis promote parasite establishment in mouse skin and skin-distant tissues, reinforcing PSG as an important part of Le. infantum transmission and visceral infection (Rogers et al. 2010). The binding of Leishmania promastigotes to the midgut epithelium is regarded as an essential part of the lifecycle in the sand fly vector, enabling the parasites to persist beyond the initial blood meal phase and establish the infection. Wilson et al. (2010) have shown that Leishmania gut binding is strictly stage-dependent and is a property of those forms found in the middle phase of development (nectomonad and leptomonad forms) but is absent in the early blood meal and final stages (procyclic and metacyclic forms). Furthermore, the adhesion is affected by glycoconjugates on Leishmania surface, especially LPG and gp63 (Jecna et al. 2013).

Significant advances have been made in exploring Leishmania-vector interactions throughout the last two decades, especially on permissiveness of Lu. longipalpis. The development of Le. infantum from establishment of infection to metacyclogenesis as well as the transmission dynamics by the bite to BALB/c mice and golden hamster have been described (Maia et al. 2011, Freitas et al. 2012, Secundino et al. 2012). For the first time Ph. perniciosus and Lu. longipalpis have been co-infected with transgenic promastigotes of Le. donovani strains carrying hygromycin or neomycin resistance genes (Sadlova et al. 2011). Seblova et al. (2015a) have tested the development of Le. infantum/Leishmania donovani natural hybrid (CUK strain) in Lu. longipalpis and the biological behavior appeared similar to what has been observed in the natural vector Phlebotomus tobbi. The phenotype impact of miltefosine-resistant Le. infantum has been evaluated on Lu. longipalpis showing a significant reduction in sand fly infection, stomodeal valve colonization and differentiation into metacyclic forms compared to the isogenic parent susceptible strain (Bockstal et al. 2019). Paromomycin-resistant Le. infantum (MHOM/FR/96/LEM3323-cl4) has behaved similar to those WT, in terms of infection and parasite location within Lu. longipalpis, and are able to colonize the stomodeal valve with metacyclic forms (Hendrickx et al. 2020). However, the mechanisms underlying drug-resistance phenotype during infection in the sand fly are yet to be determined.

In laboratory conditions Lu. longipalpis supports infection of other Leishmania species, besides Le. infantum. However, aflagellated Le. amazonensis promastigotes (Ld ARL-3A-Q70L-overexpressing) did not survive in experimentally infected Lu. longipalpis, in contrast to untransfected or native Ld ARL-3A overexpressing cells (Cuvillier et al. 2003). The role of Leishmania flagellar proteins in establishment of the parasite in the vector have been recently explored by Beneke et al. (2019). In mixed infections of the permissive sand fly Lu. longipalpis, paralyzed promastigotes and uncoordinated swimmers of Le. mexicana were severely diminished in the sand fly after the blood digestion. Furthermore, the parasites have not reached the anterior regions of the midgut, suggesting that L. mexicana needs directional motility for successful colonization of sand flies (Beneke et al. 2019). The relationship between the zinc protease gp63 and the parasite development in the sand fly vector has been evaluated (Hajmová et al. 2004). Leishmania amazonensis gp63-downregulated have presented a weak development especially in the early phase of infection, indicating that gp63 may protect promastigotes from degradation by the midgut digestive enzymes, favoring parasite survival. More recently, trying to understand the concomitant roles of gp63 and LPG, Soares et al. (2017) evaluated those two glycoconjugates using the midgut in vitro system (Pimenta et al. 1992) and LL5 cells. Both glycoconjugates were equally responsible for inhibiting parasite attachment in those models reinforcing their importance for interaction with the invertebrate host.

Parasites of the subgenus Leishmania (Mundinia) (Espinosa et al. 2018) are becoming increasingly important to human health, since some species have been reported to infect humans, such as Le. martiniquensis, Le. “Ghana strain”, and Le. orientalis (previously called “Le. siamensis”) (Pothirat et al. 2014, Chiewchanvit et al. 2015, Kwakye-Nuako et al. 2015, Jariyapan et al. 2018). The two other known species, Le. enrietti, have been found in guinea pigs (Cavia porcellus), and Le. macropodum (previously called “Le. sp. AM-2004”), have been found in red kangaroos and other macropods (Rose et al. 2004, Dougall et al. 2011, Barratt et al. 2017). Some authors have evaluated the biological behavior of Leishmania (Mundinia) parasites in permissive vectors, such as Lu. longipalpis in view of the uncertainty about the probable natural vector. Seblova et al. (2015b) have described that both Le. enrietti and Le. macropodum were able to develop late-stage infections in Culicoides sonorensis and Lu. longipalpis. However, Le. orientalis was able to establish infection in Cu. sonorensis midges but not in Lu. longipalpis (Chanmol et al. 2019), suggesting that the biting midges might be natural vectors of some Leishmania (Mundinia) species. This is of importance, because those insects were once not considered as vectors of Leishmaniasis. However, Cu. sonorensis achieved 5 out 6 criteria of Killick-Kendrick (1990) in the work of Dougall et al. (2011). Still, transmission is yet to be demonstrated for those Mundinia species (Paranaiba et al. 2017).

Insects cell lines have been used as a value tool to understand host-parasite interactions in vitro. There is two established Lu. longipalpis cell lines derived from embryonic tissues, LL5 (Tesh & Modi 1983) and Lulo (Rey et al. 2000). When LL5 cells were transfected with double stranded RNA (dsRNAs), they developed a nonspecific antiviral response (Pitaluga et al. 2008). Secreted molecules implicated in immune response in LL5 cell line have been described, such as phospholipid scramblase, an interferon-inducible protein and forskolin-binding protein, a member of the immunophilin family (Martins-da-Silva et al. 2018). A complex immune response in LL5 line cell has also been detected when challenged by different pathogens, as bacteria, yeast and Leishmania (Tinoco-Nunes et al. 2016). The Lulo cell line can be infected by Le. infantum (Bello et al. 2005) and moreover, other Leishmania species were also able to adhere to Lulo cells at different rates (Côrtes et al. 2011). The mechanisms involved in the adhesion of parasites to Lulo cells remains unclear. Côrtes et al. (2012) have described the participation of heparin binding proteins from the surface of Le. braziliensis promastigotes to Lulo cells, by their glycosaminoglycans, through heparan sulfate participation. However, lectin-like activity specific for heparin has been previously described by (Svobodová et al. 1997). Although the development of those cells could help to understand some aspects of the interaction of the parasites, there are few published papers using those models in the past years or replacement for in vivo studies have decreased their use along the years.

Finally, the presence of naturally infected sand fly by non-Leishmania trypanosomatids and other microorganisms have been reported throughout the last decades, reinforcing the role of these insects as multi-pathogens host (Shaw et al. 2003). Despite this fact, there is a lack of information about the biological behavior and infectivity of these pathogens in sand flies. Flagellates of Endotrypanum schaudinni were able to infect the abdominal midgut, pylorus, ileum, and rectal ampulla but a scarcity of infection has been observed near the stomodeal valve in Lu. longipalpis (Barbosa et al. 2006). Moreover, the presence of Le. guyanensis in a mixed infection has inhibited the development of Endotrypanum, suggesting the effect of selective pressures that have already been reported previously, among co-cultivated trypanosomatids (Barbosa et al. 2006). Also, Lu. longipalpis seems to be the host for gregarines, fungi and nematodes (Secundino et al. 2002, Matos et al. 2006, Caligiuri et al. 2014), but also the vector of other pathogens, including viruses and bacteria. Carvalho et al. (2018) have detected and isolated a putative new Phlebovirus (Viola Phlebovirus) from Lu. longipalpis in Brazil. Phylogenetic analysis revealed proximity with viruses causing disease in humans, rodents and isolated from sand flies belonging to phlebotomus fever serogroup. Moreover, the isolation of Viola virus in mammalian cells indicates that this virus is not an insect-specific virus and represents a novel species with unknown vertebrate host (Carvalho et al. 2018). In general, Lu. longipalpis was able to support the Bartonella bacilliformis infection and seems to be a user-friendly, live vector/host model system (Battisti et al. 2015). Rocha et al. (2018) have reported for the first time the occurrence of Wolbachia pipientis in a natural population of Lu. longipalpis from the State of Bahia, Brazil. Recently, the endosymbiont bacterium Wolbachia has been used as an alternative strategy to control vector-borne diseases, through the reduction or blocking of pathogen infections. However, Gonçalves et al. (2019) have shown that the Wolbachia introduction into Lu. longipalpis cell lines has not affected the infection with Le. infantum. Endosymbiotic bacteria present in sand flies, especially in the midgut, can affect their capacity to transmit Leishmania (Telleria et al. 2013). Moreover, the microbiota is able to differentially infect the larval digestive tract and regulate the immune response in Lu. longipalpis larvae (Heerman et al. 2015). Pires et al. (2017) have described the native microbiota of wild-caught Lu. longipalpis under distinct physiological conditions including a Leishmania-infected group. The amplicon oriented metagenomic profiling revealed five phyla (Actinobacteria, Bacteroidetes, Firmicutes, Proteobacteria and Spirochaetes), 64 bacterial genera and 46 families associated with wild-caught Lu. longipalpis (Pires et al. 2017). The gut microbiome of laboratory-reared Lu. longipalpis was recently shown to be essential for survival of the parasite (Kelly et al. 2017). The authors have shown that an antibiotic-mediated decrease in midgut microbiota impaired Le. infantum survival in the sand fly, inhibited parasite growth, and decreased differentiation to the infectious metacyclic form was observed (Kelly et al. 2017). Furthermore, when Lu. longipalpis was pre-fed with Pseudozyma, Asaia or Ochrobactrum, a reduced parasite survival rate has been observed by Sant’Anna et al. (2014). Still, more field-studies using such bacteria are important to establish their biological role as possible alternative control measures.

FINAL CONSIDERATIONS

The genome annotation of Lu. longipalpis is still underway and most of the omics approaches are very scarce. Dillon et al. (2006) analyzed expressed sequences tags (ESTs) of Lu. longipalpis to investigate the critical proteins underlying the host-parasite relationship and recently, an improved annotation of Lu. longipalpis genome has been published (Yang & Wu 2019). Besides that, a global approach for the identification of midgut ESTs via random, uni-directional sequencing of clones from cDNA libraries obtained using mRNAs extracted from midguts of Lu. longipalpis have been published (Jochim et al. 2008, Pitaluga et al. 2009). Moreover, transcriptome analysis of the salivary and pheromone glands as well as annotation of both female and male adults have brought important insights into the repertoire of molecules expressed in the vector (Oliveira et al. 2009, Azevedo et al. 2012, González-Caballero et al. 2013, McCarthy et al. 2013). It seems likely that in the next decade, these approaches, and perhaps more advanced ones will bring additional information of functional aspects on how molecular biology of Lu. longipalpis affects its interactions with vertebrate host and parasites. The establishment of VL in urban areas, where until recently, the disease did not occur, is closely related to the adaptation of the natural vector Lu. longipalpis to this environment. Several factors are involved in the difficulty to control VL such as the presence of sibling species in the Lu. longipalpis complex, as well as differences on vectorial capacity among populations. Moreover, the presence of another vector species has been reported in Brazil especially in absence of the main vector (de Carvalho et al. 2010, Dias et al. 2013, Guimarães et al. 2016; Rêgo et al. 2020). Studies on biological behavior of the vector, salivary components, gut physiology as well as host-parasite interaction represent a wide and important field to better understand several aspects involved in the transmission and establishment of Leishmania parasites in permissive vectors. Omics approaches are also added in this context, even in its initial phase, but providing tremendous opportunities for the research on sand flies and Leishmania species in the Americas.

ACKNOWLEDGMENTS

We thank Jason Memmott for English review of the manuscript.

REFERENCES

• ABBASI I ET AL. 2018. Plant-feeding phlebotomine sand flies, vectors of leishmaniasis, prefer Cannabis sativa. Proc Natl Acad Sci USA 115: 11790-11795.
• ABBEHUSEN MMC ET AL. 2018. Immunization of experimental dogs with salivary proteins from Lutzomyia longipalpis, using DNA and recombinant canarypox virus induces immune responses consistent with protection against Leishmania infantum. Front Immunol 9: 1-12.
• AFONSO MMDS, DUARTE R, MIRANDA JC, CARANHA L & RANGEL EF. 2012. Studies on the feeding habits of Lutzomyia (Lutzomyia) longipalpis (Lutz & Neiva, 1912) (Diptera: Psychodidae: Phlebotominae) populations from endemic areas of American Visceral Leishmaniasis in Northeastern Brazil. J Trop Med 2012.
• ALBUQUERQUE E SILVA R, ANDRADE AJ DE, QUINT BB, RAFFOUL GES, WERNECK GL, RANGEL EF & ROMERO GAS. 2018. Effectiveness of dog collars impregnated with 4% deltamethrin in controlling visceral leishmaniasis in Lutzomyia longipalpis (Diptera: Psychodidade: Phlebotominae) populations. Mem Inst Oswaldo Cruz 113: 1-9.
• ALEXANDER B, BARROS VC, SOUZA SF DE, BARROS SS, TEODORO LP, SOARES ZR, GONTIJO NF & REITHINGER R. 2009. Susceptibility to chemical insecticides of two Brazilian populations of the visceral leishmaniasis vector Lutzomyia longipalpis (Diptera: Psychodidae). Trop Med Int Heal 14: 1272-1277.
• AMÓRA SSA, BEVILAQUA CML, FEIJÓ FMC, PEREIRA RHMA, ALVES ND, FREIRE FA DE M, KAMIMURA MT, OLIVEIRA DM DE, LIMA EÁLA & ROCHA MFG. 2010. The effects of the fungus Metarhizium anisopliae var. acridum on different stages of Lutzomyia longipalpis (Diptera: Psychodidae). Acta Trop 113: 214-220.
• AMÓRA SSA, BEVILAQUA CML, FEIJÓ FMC, SILVA MA, PEREIRA RHMA, SILVA SC, ALVES ND, FREIRE FAM & OLIVEIRA DM. 2009. Evaluation of the fungus Beauveria bassiana (Deuteromycotina: Hyphomycetes), a potential biological control agent of Lutzomyia longipalpis (Diptera, Psychodidae). Biol Control 50: 329-335.
• ANDERSON JM, OLIVEIRA F, KAMHAWI S, MANS BJ, REYNOSO D, SEITZ AE, LAWYER P, GARFIELD M, PHAM MV & VALENZUELA JG. 2006. Comparative salivary gland transcriptomics of sandfly vectors of visceral leishmaniasis. BMC Genomics 7: 1-23.
• ANDRADE-COELHO CA, SOUZA NA, FEDER MD, SILVA CE, GARCIA ES, AZAMBUJA P, GONZALEZ MS & RANGEL EF. 2006. Effects of Azadirachtin on the development and mortality of Lutzomyia longipalpis larvae (Diptera: Psychodidae: Phlebotominae). J Med Entomol 43: 262-266.
• ANDRADE-COELHO CA, SOUZA NA, SILVA VC, SOUZA AA, GONZALEZ MS & RANGEL EF. 2014. Effects of Azadirachtin on the biology of Lutzomyia longipalpis (Diptera: Psychodidae: Phlebotominae) adult female, the main vector of American visceral leishmaniasis. J Med Entomol 51: 891-895.
• ANDRADE-COELHO CA, SOUZA NA, GOUVEIA C, SILVA VC, GONZALEZ MS & RANGEL EF. 2009. Effect of fruit and leaves of Meliaceae plants (Azadirachta indica and Melia azedarach) on the development of Lutzomyia longipalpis larvae (Diptera: Psychodidae: Phlebotominae) under experimental conditions. J Med Entomol 46: 1125-1130.
• ANDRADE-FILHO JD, SCHOLTE RGC, AMARAL ALG, SHIMABUKURO PHF, CARVALHO OS & CALDEIRA RL. 2017. Occurrence and probability maps of Lutzomyia longipalpis and Lutzomyia cruzi (Diptera: Psychodidae: Phlebotominae) in Brazil. J Med Entomol 54: 1430-1434.
• ARAKI AS, FERREIRA GEM, MAZZONI CJ, SOUZA NA, MACHADO RC, BRUNO RV & PEIXOTO AA. 2013. Multilocus analysis of divergence and introgression in sympatric and allopatric sibling species of the Lutzomyia longipalpis complex in Brazil. PLoS Negl Trop Dis 7: e2495.
• ARAKI AS, VIGODER FM, BAUZER LGSR, FERREIRA GEM, SOUZA NA, ARAÚJO IB, HAMILTON JGC, BRAZIL RP & PEIXOTO AA. 2009. Molecular and behavioral differentiation among Brazilian populations of Lutzomyia longipalpis (Diptera: Psychodidae: Phlebotominae). PLoS Negl Trop Dis 3: e365.
• ARAÚJO-SANTOS T ET AL. 2010. Lutzomyia longipalpis saliva triggers lipid body formation and prostaglandin E2 production in murine macrophages. PLoS Negl Trop Dis 4: e873.
• ARRIVILLAGA J, MUTEBI JP, PIÑANGO H, NORRIS D, ALEXANDER B, FELICIANGELI MD & LANZARO GC. 2003. The taxonomic status of genetically divergent populations of Lutzomyia longipalpis (Diptera: Psychodidae) based on the distribution of mitochondrial and isozyme variation. J Med Entomol 40: 615-627.
• ARRIVILLAGA J, SALERNO P & RANGEL Y. 2009. Aislamiento reproductivo asimétrico entre Lutzomyia pseudolongipalpis y Lutzomyia longipalpis (especie C2), vectores neotropicales de leishmaniasis visceral (Diptera: Pshychodidae). Rev Biol Trop 57: 23-31.
• ASOJO OA ET AL. 2017. Structure of SALO, a leishmaniasis vaccine candidate from the sand fly Lutzomyia longipalpis. PLoS Negl Trop Dis 11: 1-15.
• ASSIS RR, IBRAIM IC, NOGUEIRA PM, SOARES RP & TURCO SJ. 2012. Glycoconjugates in New World species of Leishmania: Polymorphisms in lipophosphoglycan and glycoinositolphospholipids and interaction with hosts. Biochim Biophys Acta Gen Subj 1820: 1354-1365.
• AZEVEDO RVDM, DIAS DBS, BRETÃS JAC, MAZZONI CJ, SOUZA NA, ALBANO RM, WAGNER G, DAVILA AMR & PEIXOTO AA. 2012. The transcriptome of Lutzomyia longipalpis (Diptera: Psychodidae) male reproductive organs. PLoS One 7: e34495.
• BAHIA D, GONTIJO NF, LEÓN IR, PERALES J, PEREIRA MH, OLIVEIRA G, CORRÊA-OLIVEIRA R & REIS AB. 2007. Antibodies from dogs with canine visceral leishmaniasis recognise two proteins from the saliva of Lutzomyia longipalpis. Parasitol Res 100: 449-454.
• BALBINO VQ, COUTINHO-ABREU IV, SONODA IV, MELO MA, ANDRADE PP, CASTRO JAF, REBÊLO JM, CARVALHO SMS & RAMALHO-ORTIGÃO JM. 2006. Genetic structure of natural populations of the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) from the Brazilian northeastern region. Acta Trop 98: 15-24.
• BARBOSA AF, OLIVEIRA SMP, BERTHO ÁL, FRANCO AMR & RANGEL EF. 2006. Single and concomitant experimental infections by Endotrypanum spp. and Leishmania (Viannia) guyanensis (Kinetoplastida: Trypanosomatidae) in the neotropical sand fly Lutzomyia longipalpis (Diptera: Psychodidae). Mem Inst Oswaldo Cruz 101: 851-856.
• BARRATT J, KAUFER A, PETERS B, CRAIG D, LAWRENCE A, ROBERTS T, LEE R, MCAULIFFE G, STARK D & ELLIS J. 2017. Isolation of novel Trypanosomatid, Zelonia australiensis sp. nov. (Kinetoplastida: Trypanosomatidae) provides support for a Gondwanan origin of dixenous parasitism in the Leishmaniinae. PLoS Negl Trop Dis 11: e0005215.
• BATES PA. 2008. Leishmania sand fly interaction: progress and challenges. Curr Opin Microbiol 11(4): 340-344.
• BATES PA & ROGERS ME. 2004. New insights into the developmental biology and transmission mechanisms of Leishmania. Curr Mol Med 4(6): 601-609.
• BATTISTI JM, LAWYER PG & MINNICK MF. 2015. Colonization of Lutzomyia verrucarum and Lutzomyia longipalpis sand flies (Diptera: Psychodidae) by Bartonella bacilliformis, the etiologic agent of Carrión’s disease. PLoS Negl Trop Dis 9: e0004128.
• BAUZER LGSR, SOUZA NA, MAINGON RDC & PEIXOTO AA. 2007. Lutzomyia longipalpis in Brazil: A complex or a single species? A mini-review. Mem Inst Oswaldo Cruz 102: 1-12.
• BELL MJ, SEDDA L, GONZALEZ MA, SOUZA CF, DILGER E, BRAZIL RP, COURTENAY O & HAMILTON JGC. 2018. Attraction of Lutzomyia longipalpis to synthetic sex-aggregation pheromone: Effect of release rate and proximity of adjacent pheromone sources. PLoS Negl Trop Dis 12: e0007007.
• BELLO FJ, MEJÍA AJ, PILAR-CORENA M, AYALA M, SARMIENTO L, ZUÑIGA C & PALAU MT. 2005. Experimental infection of Leishmania (L.) chagasi in a cell line derived from Lutzomyia longipalpis (Diptera:Psychodidae). Mem Inst Oswaldo Cruz 100: 518-525.
• BENEKE T ET AL. 2019. Genetic dissection of a Leishmania flagellar proteome demonstrates requirement for directional motility in sand fly infections. PLoS Pathog 15: e1007828.
• BOCKSTAL LV, SÁDLOVÁ J, SUAU HA, HENDRICKX S, MENESES C, KAMHAWI S, VOLF P, MAES LV & CALJON G. 2019. Impaired development of a miltefosine-resistant Leishmania infantum strain in the sand fly vectors Phlebotomus perniciosus and Lutzomyia longipalpis. Int J Parasitol Drugs Drug Resist 11: 1-7.
• BOROVSKY D & SCHLEIN Y. 1987. Trypsin and chymotrypsin-Iike enzymes of the sandfly Phlebotomus papatasi infected with Leishmania and their possible role in vector competence. Med Vet Entomol 1: 235-242.
• BOTTECCHIA M, OLIVEIRA SG, BAUZER LGSR, SOUZA NA, WARD RD, GARNER KJ, KYRIACOU CP & PEIXOTO AA. 2004. Genetic divergence in the cacophony IVS6 intron among five Brazilian populations of Lutzomyia longipalpis. J Mol Evol 58: 754-761.
• BRAY DP, ALVES GB, DORVAL ME, BRAZIL RP & HAMILTON JGC. 2010. Synthetic sex pheromone attracts the leishmaniasis vector Lutzomyia longipalpis to experimental chicken sheds treated with insecticide. Parasit Vectors 3: 1-11.
• BRAY DP, BANDI KK, BRAZIL RP, OLIVEIRA AG & HAMILTON JGC. 2009. Synthetic sex pheromone attracts the leishmaniasis vector Lutzomyia longipalpis (Diptera: Psychodidae) to traps in the field. J Med Entomol 46: 428-434.
• BRAY DP & HAMILTON JGC. 2013. Insecticide-impregnated netting as a potential tool for long-lasting control of the leishmaniasis vector Lutzomyia longipalpis in animal shelters. Parasit Vectors 6: 1-7.
• BRAZIL RP. 2013. The dispersion of Lutzomyia longipalpis in urban areas. Rev Soc Bras Med Trop 46: 263-264.
• BRAZIL RP, LANÇA-PASSOS W, FUZARI AA, FALCÃO AL & ANDRADE-FILHO JD. 2006. The peridomiciliar sand fly fauna (Diptera: Psychodidae) in areas of cutaneous leishmaniasis in Além Paraíba, Minas Gerais, Brazil. J Vec Ecol 31: 1-3.
• BRODIE TM, SMITH MC, MORRIS RV & TITUS RG. 2007. Immunomodulatory effects of the Lutzomyia longipalpis salivary gland protein Maxadilan on mouse macrophages. Infect Immun 75: 2359-2365.
• CALIGIURI LG, ACARDI SA, SANTINI MS, SALOMÓN OD & MCCARTHY CB. 2014. Polymerase chain reaction-based assay for the detection and identification of sand fly gregarines in Lutzomyia longipalpis, a vector of visceral leishmaniasis. J Vec Ecol 39: 83-93.
• CAMARGO-NEVES VLF, RODAS LAC, CABRAL G & PAULIQUÉVIS-JR C. 2007a. Avaliação da eficácia Lambdacialotrina para o controle de Lutzomyia longipalpis. BEPA Bol Epidem Paul 4: 04-11.
• CAMARGO-NEVES VLF, RODAS LAC & GOMES AC. 2007b. Evaluation of feeding habits of Lutzomyia longipalpis in the State of São Paulo. Bol Epidem Paul 4: 1-6.
• CARREGARO V, COSTA DL, BRODSKYN C, BARRAL AM, BARRAL-NETTO M, CUNHA FQ & SILVA JS. 2013. Dual effect of Lutzomyia longipalpis saliva on Leishmania braziliensis infection is mediated by distinct saliva-induced cellular recruitment into BALB/c mice ear. BMC Microbiol 13(1): 1-11.
• CARVALHO MR, VALENÇA HF, SILVA FJ, PITA-PEREIRA D, ARAÚJO-PEREIRA T, BRITTO C, BRAZIL RP & FILHO SPB. 2010. Natural Leishmania infantum infection in Migonemyia migonei (França, 1920) (Diptera: Psychodidae: Phlebotominae) the putative vector of visceral leishmaniasis in Pernambuco State, Brazil. Acta Trop 116: 108-110.
• CARVALHO AM, FUKUTANI KF, SHARMA R, CURVELO RP, MIRANDA JC, BARRAL A, CARVALHO EM, VALENZUELA JG, OLIVEIRA F & OLIVEIRA CI. 2017a. Seroconversion to Lutzomyia intermedia LinB-13 as a biomarker for developing cutaneous leishmaniasis. Sci Rep 7: 3149.
• CARVALHO MS, LARA-PINTO AZ, PINHEIRO A, RODRIGUES JSV, MELO FL, SILVA LA, RIBEIRO BM & DEZENGRINI-SLHESSARENKO R. 2018. Viola phlebovirus is a novel Phlebotomus fever serogroup member identified in Lutzomyia (Lutzomyia) longipalpis from Brazilian Pantanal. Parasit Vectors 11: 1-10.
• CARVALHO GML, RÊGO FD, TANURE A, SILVA ACP, DIAS TA, PAZ GF & ANDRADE-FILHO JD. 2017b. Bloodmeal identification in field-collected sand flies from Casa Branca, Brazil, using the cytochrome b PCR method. J Med Entomol 54(4): 1049-1054.
• CASANOVA C. 2001. A soil emergence trap for collections of phlebotomine sand flies. Mem Inst Oswaldo Cruz 96: 273-275.
• CASARIL AE, ALONSO DP, FRANCO KG, ALVAREZ MVN, BARRIOS SPG, FERNANDES WS, MOURA IJO, RODRIGUES ACM, RIBOLLA PEM & OLIVEIRA AG. 2019. Macrogeographic genetic structure of Lutzomyia longipalpis complex populations using Next Generation Sequencing. PLoS One 14: e0223277.
• CAVALCANTE RR, PEREIRA MH, FREITAS JM & GONTIJO NDF. 2006. Ingestion of saliva during carbohydrate feeding by Lutzomyia longipalpis (Diptera; Psychodidae). Mem Inst Oswaldo Cruz 101: 85-87.
• CHAGAS AC, OLIVEIRA F, DEBRABANT A, VALENZUELA JG, RIBEIRO JMC & CALVO E 2014. Lundep, a sand fly salivary endonuclease increases Leishmania parasite survival in neutrophils and inhibits XIIa contact activation in human plasma. PLoS Pathog 10: e1003923.
• CHANMOL W, JARIYAPAN N, SOMBOON P, BATES MD & BATES PA. 2019. Development of Leishmania orientalis in the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) and the biting midge Culicoides soronensis (Diptera: Ceratopogonidae). Acta Trop 199: 105157.
• CHIEWCHANVIT S, TOVANABUTRA N, JARIYAPAN N, BATES MD, MAHANUPAB P, CHUAMANOCHAN M, TANTIWORAWIT A & BATES PA. 2015. Chronic generalized fibrotic skin lesions from disseminated leishmaniasis caused by Leishmania martiniquensis in two patients from northern Thailand infected with HIV. Br J Dermatol 173: 663-670.
• COELHO-FINAMORE JM, FREITAS VC, ASSIS RR, MELO MN, NOVOZHILOVA N, SECUNDINO NFC, PIMENTA PF, TURCO SJ & SOARES RP. 2011. Leishmania infantum: Lipophosphoglycan intraspecific variation and interaction with vertebrate and invertebrate hosts. Int J Parasitol 41: 333-342.
• COLLIN N ET AL. 2012. Lufaxin, a Novel Factor Xa Inhibitor from the salivary gland of the sand fly Lutzomyia longipalpis, blocks PAR2 activation and inhibits inflammation and thrombosis in vivo. Arter Thromb Vasc Biol 32: 2185-2198.
• COLLIN N, GOMES R, TEIXEIRA C, CHENG L, LAUGHINGHOUSE A, WARD JM, ELNAIEM DE, FISCHER L, VALENZUELA JG & KAMHAWI S. 2009. Sand fly salivary proteins induce strong cellular immunity in a natural reservoir of visceral leishmaniasis with adverse consequences for Leishmania. PLoS Pathog 5: e1000441.
• CÔRTES LMDC ET AL. 2012. Participation of heparin binding proteins from the surface of Leishmania (Viannia) braziliensis promastigotes in the adhesion of parasites to Lutzomyia longipalpis cells (Lulo) in vitro. Parasit Vectors 5: 1-10.
• CÔRTES LMDC ET AL. 2011. Lulo cell line derived from Lutzomyia longipalpis (Diptera: Psychodidae): A novel model to assay Leishmania spp. and vector interaction. Parasit Vectors 4: 3-7.
• COSTA-JÚNIOR CRL ET AL. 2015. Genetic structuring and fixed polymorphisms in the gene period among natural populations of Lutzomyia longipalpis in Brazil. Parasit Vectors 8: 1-9.
• COSTA DJ ET AL. 2004. Lutzomyia longipalpis salivary gland homogenate impairs cytokine production and costimulatory molecule expression on human monocytes and dendritic cells. Infect Immun 72: 1298-1305.
• COURA FM, OLIVEIRA F, LEME P, ALVES S, BARACT R, ARAUJO D, PIMENTA A & BICALHO V. 2019. Evaluation of the antifeeding and insecticidal effects of a deltamethrin- impregnated collar on Lutzomyia longipalpis. Acta Vet Bras. 13: 192-197.
• COURTENAY O, GILLINGWATER K, GOMES PAF, GARCEZ LM & DAVIES CR. 2007. Deltamethrin-impregnated bednets reduce human landing rates of sandfly vector Lutzomyia longipalpis in Amazon households. Med Vet Entomol 21: 168-176.
• CUNHA JM, ABBEHUSEN M, SUAREZ M, VALENZUELA J, TEIXEIRA CR & BRODSKYN CI. 2018. Immunization with LJM11 salivary protein protects against infection with Leishmania braziliensis in the presence of Lutzomyia longipalpis saliva. Acta Trop 177: 164-170.
• CUTOLO AA, GALVIS-OVALLOS F, SOUZA NEVES E, SILVA FO, CHESTER ST & FANKHAUSER B. 2018. Repellent efficacy of a new combination of fipronil and permethrin against Lutzomyia longipalpis. Parasit Vectors 11: 4-9.
• CUVILLIER A, MIRANDA JC, AMBIT A, BARRAL A & MERLIN G. 2003. Abortive infection of Lutzomyia longipalpis insect vectors by aflagellated LdARL-3A-Q70L overexpressing Leishmania amazonensis parasites. Cell Microbiol 5: 717-728.
• DENLINGER DS, CRESWELL JA, ANDERSON JL, REESE CK & BERNHARDT SA. 2016. Diagnostic doses and times for Phlebotomus papatasi and Lutzomyia longipalpis sand flies (Diptera: Psychodidae: Phlebotominae) using the CDC bottle bioassay to assess insecticide resistance. Parasit Vectors 9: 1-11.
• DENLINGER DS, LOZANO-FUENTES S, LAWYER PG, BLACK WC & BERNHARDT SA 2015. Assessing insecticide susceptibility of laboratory Lutzomyia longipalpis and Phlebotomus papatasi sand flies (Diptera: Psychodidae: Phlebotominae). J Med Entomol 52: 1003-1012.
• DIAS FOP, LOROSA ES & REBÊLO JMM. 2003. Fonte alimentar sanguínea e a peridomiciliação de Lutzomyia longipalpis (Lutz & Neiva, 1912) (Psychodidae, Phlebotominae). Cad Saude Publica 19: 1373-1380.
• DIAS ES, MICHALSKY ÉM, NASCIMENTO JC, FERREIRA EC, LOPES JV & FORTES-DIAS CL. 2013. Detection of Leishmania infantum, the etiological agent of visceral leishmaniasis, in Lutzomyia neivai, a putative vector of cutaneous leishmaniasis. J Vector Ecol 38: 193-196.
• DILLON RJ ET AL. 2006. Analysis of ESTs from Lutzomyia longipalpis sand flies and their contribution toward understanding the insect-parasite relationship. Genomics 88: 831-840.
• DOSTÁLOVÁ A & VOLF P. 2012. Leishmania development in sand flies: parasite-vector interactions overview. Parasit Vectors 5(1): 1-12.
• DOSTÁLOVÁ A, VOTÝPKA J, FAVREAU AJ, BARBIAN KD, VOLF P, VALENZUELA JG & JOCHIM RC. 2011. The midgut transcriptome of Phlebotomus (Larroussius) perniciosus, a vector of Leishmania infantum: Comparison of sugar fed and blood fed sand flies. BMC Genomics 12: 1-21.
• DOUGALL AM, ALEXANDER B, HOLT DC, HARRIS T, SULTAN AH, BATES PA, ROSE K & WALTON SF. 2011. Evidence incriminating midges (Diptera: Ceratopogonidae) as potential vectors of Leishmania in Australia. Int J Parasitol 41: 571-579.
• DVORAK V, SHAW J & VOLF P. 2018. Parasite biology: The vectors. In: Bruschi F & Gradoni L (Eds), Leishmaniases Old Neglected Trop Dis, Springer International Publishing, p. 31-77.
• ELNAIEM DA, MORTON I, BRAZIL RP & WARD RD. 1992. Field and laboratory evidence for multiple bloodfeeding by Lutzomyia longipalpis (Diptera: Psychodidae). Med Vet Entomol 6: 173-174.
• ELNAIEM DA, WARD RD & YOUNG PE. 1994. Development of Leishmania chagasi (Kinetoplastida: Trypanosomatidae) in the second blood-meal of its vector Lutzomyia longipalpis (Diptera: Psychodidae). Parasitol Res 80: 414-419.
• ESPINOSA OA, SERRANO MG, CAMARGO EP, TEIXEIRA MMG & SHAW JJ. 2018. An appraisal of the taxonomy and nomenclature of trypanosomatids presently classified as Leishmania and Endotrypanum. Parasitology 145: 430-442.
• FELICIANGELI MD, MAZZARRI MB, BLAS SS & ZERPA O. 2003. Control trial of Lutzomyia longipalpis s.l. in the Island of Margarita, Venezuela. Trop Med Int Heal 8: 1131-1136.
• FERREIRA VP ET AL. 2016. SALO, a novel classical pathway complement inhibitor from saliva of the sand fly Lutzomyia longipalpis. Scientific Reports 6(1): 1-13.
• FERROGLIO E, POGGI M & TRISCIUOGLIO A. 2008. Evaluation of 65% permethrin spot-on and deltamethrin-impregnated collars for canine Leishmania infantum infection prevention. Zoonoses Public Health 55: 145-148.
• FRAGA TL, FERNANDES MF, PONTES ERJC, LEVAY APS, ALMEIDA-DA-CUNHA EB, FRANÇA ADO & DORVAL MEC. 2016. Antissaliva antibodies of Lutzomyia longipalpis in area of visceral leishmaniasis. Pediatr Infect Dis J 35: 805-807.
• FREITAS VC, PARREIRAS KP, DUARTE APM, SECUNDINO NFC & PIMENTA PFP. 2012. Development of Leishmania (Leishmania) infantum chagasi in its natural sandfly vector Lutzomyia longipalpis. Am J Trop Med Hyg 86: 606-612.
• GALATI EAB. 2018. Phlebotominae (Diptera, Psychodidae): Classification, morphology and terminology of adults and identification of American Taxa. In: Brazilian Sand Flies Biol Taxon Med Importance Control, Springer International Publishing, p. 9-212.
• GONÇALVES DS, ITURBE-ORMAETXE I, MARTINS-DA-SILVA A, TELLERIA EL, ROCHA MN, TRAUB-CSEKÖ YM, O’NEILL SL, SANT’ANNA MRV & MOREIRA LA. 2019. Wolbachia introduction into Lutzomyia longipalpis (Diptera: Psychodidae) cell lines and its effects on immune-related gene expression and interaction with Leishmania infantum. Parasit Vectors 12: 1-13.
• GONZÁLEZ-CABALLERO N, VALENZUELA JG, RIBEIRO JMC, CUERVO P & BRAZIL RP. 2013. Transcriptome exploration of the sex pheromone gland of Lutzomyia longipalpis (Diptera: Psychodidae: Phlebotominae). Parasit Vectors 6: 1-16.
• GUIMARÃES VCFV, PRUZINOVA K, SADLOVA J, VOLFOVA V, MYŠKOVÁ J, FILHO SPB & VOLF P. 2016. Lutzomyia migonei is a permissive vector competent for Leishmania infantum. Parasit Vectors 9: 159.
• HAJMOVÁ M, CHANG KP, KOLLI B & VOLF P. 2004. Down-regulation of gp63 in Leishmania amazonensis reduces its early development in Lutzomyia longipalpis. Microbes Infect 6: 646-649.
• HAMILTON JGC, MAINGON RDC, ALEXANDER B, WARD RD & BRAZIL RP. 2005. Analysis of the sex pheromone extract of individual male Lutzomyia longipalpis sandflies from six regions in Brazil. Med Vet Entomol 19: 480-488.
• HASSAN MM, WIDAA SO, OSMAN OM, NUMIARY MSM, IBRAHIM MA & ABUSHAMA HM. 2012. Insecticide resistance in the sand fly, Phlebotomus papatasi from Khartoum State, Sudan. Parasit Vectors 5: 46.
• HEERMAN M, WENG JL, HURWITZ I, DURVASULA R & RAMALHO-ORTIGÃO JM. 2015. Bacterial infection and immune responses in Lutzomyia longipalpis sand fly larvae midgut. PLoS Negl Trop Dis 9: e0003923.
• HENDRICKX S, VAN-BOCKSTAL L, ASLAN H, SADLOVA J, MAES L, VOLF P & CALJON G. 2020. Transmission potential of paromomycin-resistant Leishmania infantum and Leishmania donovani. J Antimicrob Chemother 75: 951-957.
• HLAVACOVA J, VOTYPKA J & VOLF P. 2013. The effect of temperature on Leishmania (Kinetoplastida: Trypanosomatidae) development in sand flies. J Med Entomol 50(5): 955-958.
• HODGKINSON VH, BIRUNGI J, QUINTANA M, DEITZE R & MUNSTERMANN LE. 2003. Mitochondrial cytochrome B variation in populations of the visceral leishmaniasis vector Lutzomyia longipalpis across eastern Brazil. Am J Trop Med Hyg 69: 386-392.
• HOSTOMSKA J, ROHOUSOVA I, VOLFOVA V, STANNECK D, MENCKE N & VOLF P. 2008. Kinetics of canine antibody response to saliva of the sand fly Lutzomyia longipalpis. Vector Borne Zoonotic Dis 8: 443-450.
• JARIYAPAN N, DAROONTUM T, JAIWONG K, CHANMOL W, INTAKHAN N, SOR-SUWAN S, SIRIYASATIEN P, SOMBOON P, BATES MD & BATES PA. 2018. Leishmania (Mundinia) orientalis n. sp. (Trypanosomatidae), a parasite from Thailand responsible for localised cutaneous leishmaniasis. Parasit Vectors 11: 351.
• JECNA L, DOSTALOVA A, WILSON R, SEBLOVA V, CHANG KP, BATES PA & VOLF P. 2013. The role of surface glycoconjugates in Leishmania midgut attachment examined by competitive binding assays and experimental development in sand flies. Parasitology 140: 1026-1032.
• JOCHIM RC, TEIXEIRA CR, LAUGHINGHOUSE A, MU J, OLIVEIRA F, GOMES RB, ELNAIEM DE & VALENZUELA JG. 2008. The midgut transcriptome of Lutzomyia longipalpis: Comparative analysis of cDNA libraries from sugar-fed, blood-fed, post-digested and Leishmania infantum chagasi-infected sand flies. BMC Genomics 9: 1-24.
• JUAN LW, LUCIA A, ALZOGARAY RLA, STEINHORST II, LÓPEZ K, PETTERSEN M, BUSSE J & ZERBA EN. 2016. Field evaluation of a new strategy to control Lutzomyia longipalpis, based on simultaneous application of an adulticide-larvicide mixture. J Am Mosq Control Assoc 32: 224-229.
• KAMHAWI S, RAMALHO-ORTIGÃO JM, VAN MP, KUMAR S, LAWYER PG, TURCO SJ, BARILLAS-MURY C, SACKS DL & VALENZUELA JG. 2004. A role for insect galectins in parasite survival. Cell 119: 329-341.
• KELLY PH, BAHR SM, SERAFIM TD, AJAMI NJ, PETROSINO JF, MENESES C, KIRBY JR, VALENZUELA JG, KAMHAWI S & WILSON ME. 2017. The gut microbiome of the vector Lutzomyia longipalpis is essential for survival of Leishmania infantum. MBio 8: e01121-16.
• KILLICK-KENDRICK R. 1990. Phlebotomine vectors of the leishmaniases: a review. Med Vet Entomol 4: 1-24.
• KWAKYE-NUAKO G ET AL. 2015. First isolation of a new species of Leishmania responsible for human cutaneous leishmaniasis in Ghana and classification in the Leishmania enriettii complex. Int J Parasitol 45: 679-684.
• LAINSON R & RANGEL EF. 2005. Lutzomyia longipalpis and the eco-epidemiology of American visceral leishmaniasis, with particular reference to Brazil: a review. Mem Inst Oswaldo Cruz 100: 811-827.
• LAINSON R & SHAW J. 1987. Evolution, classification and geographical distribution. In: Peters W & Killick-Kendrick R (Eds), Leishmaniases Biol Med, Academic Press Inc., Orlando, p. 1-120.
• LAURENTI MD, MATTA VLR, PERNICHELLI T, SECUNDINO NFC, PINTO LC, CORBETT CEP & PIMENTA PPF. 2009. Effects of salivary gland homogenate from wild-caught and laboratory-reared Lutzomyia longipalpis on the evolution and immunomodulation of Leishmania (Leishmania) amazonensis infection. Scand J Immunol 70: 389-395.
• LIMA LHGDM, MESQUITA MR, SKRIP L, SOUZA-FREITAS MT, SILVA VC, KIRSTEIN OD, ABBASI I, WARBURG A, BALBINO VDQ & COSTA CHN. 2016. DNA barcode for the identification of the sand fly Lutzomyia longipalpis plant feeding preferences in a tropical urban environment. Sci Rep 6: 29742.
• LINS RMMA, SOUZA NA, BRAZIL RP, MAINGON RDC & PEIXOTO AA. 2012. Fixed differences in the paralytic gene define two lineages within the Lutzomyia longipalpis complex producing different types of courtship songs. PLoS One 7: e44323.
• LINS RMMA, SOUZA NA & PEIXOTO AA. 2008. Genetic divergence between two sympatric species of the Lutzomyia longipalpis complex in the paralytic gene, a locus associated with insecticide resistance and lovesong production. Mem Inst Oswaldo Cruz 103: 736-740.
• LUTZ A & NEIVA A. 1912. Contribuição para o conhecimento das espécies do gênero Phlebotomus existentes no Brasil. Mem Inst Oswaldo Cruz 4: 84-95.
• LUZ NF ET AL. 2018. Lutzomyia longipalpis saliva induces heme oxygenase-1 expression at bite sites. Front Immunol 9: 1-10.
• LUZ E, GIOVANNONI M & BORBA A. 1967. Infecção de Lutzomyia monticola por Leishmania enriettii. An Fac Med Univ Fed Paraná 9: 121-128.
• MACIEL MV, MORAIS SM, BEVILAQUA CML, SILVA RA, BARROS RS, SOUSA RN, SOUSA LC, MACHADO LKA, BRITO ES & SOUZA-NETO MA. 2010. Atividade inseticida in vitro do óleo de sementes de nim sobre Lutzomyia longipalpis (Diptera: Psychodidae). Rev Bras Parasitol Vet 19: 7-11.
• MAGALHÃES-JUNIOR JT, BARROUIN-MELO SM, CORRÊA AG, ROCHA SILVA FB, MACHADO VE, GOVONE JS & PINTO MC. 2014. A laboratory evaluation of alcohols as attractants for the sandfly Lutzomyia longipalpis (Diptera:Psychodidae). Parasit Vectors 7: 2-6.
• MAIA C, SEBLOVA V, SADLOVA J, VOTYPKA J & VOLF P. 2011. Experimental transmission of Leishmania infantum by two major vectors: A comparison between a viscerotropic and a dermotropic strain. PLoS Negl Trop Dis 5: e1181.
• MAINGON RDC, WARD RD, HAMILTON JGC, NOYES HA, SOUZA N, KEMP SJ & WATTS PC. 2003. Genetic identification of two sibling species of Lutzomyia longipalpis (Diptera: Psychodidae) that produce distinct male sex pheromones in Sobral, Ceará State, Brazil. Mol Ecol 12: 1879-1894.
• MANZILLO VF, OLIVA G, PAGANO A, MANNA L, MAROLI M & GRADONI L. 2006. Deltamethrin-impregnated collars for the control of canine leishmaniasis: Evaluation of the protective effect and influence on the clinical outcome of Leishmania infection in kennelled stray dogs. Vet Parasitol 142: 142-145.
• MARASSÁ AM, CONSALES CA, GALATI EAB & NUNES VLB. 2006. Identificação do sangue ingerido por Lutzomyia (Lutzomyia) longipalpis (Lutz & Neiva, 1912) e Lutzomyia (Lutzomyia) almerioi (Galati & Nunes, 1999) pela técnica imunoenzimática do ELISA de captura, no sistema avidina-biotina. Rev Soc Bras Med Trop 39: 183-186.
• MAROLI M, MIZZONI V, SIRAGUSA C, D’ORAZI A & GRADONI L. 2001. Evidence for an impact on the incidence of canine leishmaniasis by the mass use of deltamethrin-impregnated dog collars in southern Italy. Med Vet Entomol 15: 358-363.
• MARTIN-MARTIN I ET AL. 2018. Immunity to LuloHya and Lundep, the salivary spreading factors from Lutzomyia longipalpis, protects against Leishmania major infection. PLoS Pathog 14: e1007006.
• MARTINS-DA-SILVA A, TELLERIA EL, BATISTA M, MARCHINI FK, TRAUB-CSEKÖ YM & TEMPONE AJ. 2018. Identification of secreted proteins involved in nonspecific dsRNA-mediated Lutzomyia longipalpis LL5 cell antiviral response. Viruses 10(1): 43.
• MATOS E, MENDONÇA I & AZEVEDO C. 2006. Vavraia lutzomyiae n. sp. (Phylum Microspora) infecting the sandfly Lutzomyia longipalpis (Psychodidae, Phlebotominae), a vector of human visceral leishmaniasis. Eur J Protistol 42: 21-28.
• MCCARTHY CB, SANTINI MS, PIMENTA PFP & DIAMBRA LA. 2013. First comparative transcriptomic analysis of wild adult male and female Lutzomyia longipalpis, vector of visceral leishmaniasis. PLoS One 8: e58645.
• MENDES-SOUSA AF, NASCIMENTO AAS, QUEIROZ DC, VALE VF, FUJIWARA RT, ARAÚJO RN, PEREIRA MH & GONTIJO NF. 2013. Different host complement systems and their interactions with saliva from Lutzomyia longipalpis (Diptera, Psychodidae) and Leishmania infantum promastigotes. PLoS One 8: e79787.
• MISSAWA NA, LOROSA ES & DIAS ES. 2008. Preferência alimentar de Lutzomyia longipalpis (Lutz & Neiva, 1912) em área de transmissão de leishmaniose visceral em Mato Grosso. Rev Soc Bras Med Trop 41: 365-368.
• MONTEIRO MC, LIMA HC, SOUZA AAA, TITUS RG, ROMÃO PRT & CUNHA FDQ. 2007. Effect of Lutzomyia longipalpis salivary gland extracts on leukocyte migration induced by Leishmania major. Am J Trop Med Hyg 76: 88-94.
• MONTEIRO MC, NOGUEIRA LG, ALMEIDA SOUZA AA, RIBEIRO JMC, SILVA JS & CUNHA FQ. 2005. Effect of salivary gland extract of Leishmania vector, Lutzomyia longipalpis, on leukocyte migration in OVA-induced immune peritonitis. Eur J Immunol 35: 2424-2433.
• MORAES CS, AGUIAR-MARTINS K, COSTA SG, BATES PA, DILLON RJ & GENTA FA. 2018. Second blood meal by female Lutzomyia longipalpis: enhancement by oviposition and its effects on digestion, longevity, and Leishmania infection. Biomed Res Int 2018: 2472508.
• MORAES CS, DIAZ-ALBITER HM, FARIA M DO V, SANT’ANNA MRV, DILLON RJ & GENTA FA. 2014. Expression pattern of glycoside hydrolase genes in Lutzomyia longipalpis reveals key enzymes involved in larval digestion. Front Physiol 5: 276.
• MORAES CS, LUCENA SA, MOREIRA BHS, BRAZIL RP, GONTIJO NF & GENTA FA. 2012. Relationship between digestive enzymes and food habit of Lutzomyia longipalpis (Diptera: Psychodidae) larvae: Characterization of carbohydrases and digestion of microorganisms. J Insect Physiol 58: 1136-1145.
• MYŠKOVÁ J, DOSTÁLOVÁ A, PĚNIČKOVÁ L, HALADA P, BATES PA & VOLF P. 2016. Characterization of a midgut mucin-like glycoconjugate of Lutzomyia longipalpis with a potential role in Leishmania attachment. Parasit Vectors 9: 1-10.
• MYŠKOVÁ J, SVOBODOVA M, BEVERLEY SM & VOLF P. 2007. A lipophosphoglycan-independent development of Leishmania in permissive sand flies. Microbes Infect 9: 317-324.
• NEPOMUCENO DB, PAIM RMM, ARAÚJO RN, PEREIRA MH, PESSOA GCDÁ, KOERICH LB, SANT’ANNA MRV & GONTIJO NF 2020. The role of LuloPAT amino acid/proton symporters in midgut alkalinization in the sandfly Lutzomyia longipalpis (Diptera - Psychodidae). J Insect Physiol 120: 103973.
• NORSWORTHY NB, SUN J, ELNAIEM D, LANZARO G & SOONG L. 2004. Sand fly saliva enhances Leishmania amazonensis infection by modulating interleukin-10 Production. Infect Immun 72: 1240-1247.
• OLIVEIRA F, JOCHIM RC, VALENZUELA JG & KAMHAWI S. 2009. Sand flies, Leishmania, and transcriptome-borne solutions. Parasitol Int 58: 1-5.
• ORTIGÃO-FARIAS JR, DI-BLASI T, TELLERIA EL, ANDORINHO AC, LEMOS-SILVA T, RAMALHO-ORTIGÃO JM, TEMPONE AJ & TRAUB-CSEKÖ YM. 2018. Alternative splicing originates different domain structure organization of Lutzomyia longipalpis chitinases. Mem Inst Oswaldo Cruz 113: 96-101.
• PARANAIBA LF, PINHEIRO LJ, TORRECILHAS AC, MACEDO DH, MENEZES-NETO A, TAFURI WL & SOARES RP. 2017. Leishmania enriettii (Muniz & Medina, 1948): A highly diverse parasite is here to stay. PLoS Pathog 13: e1006303.
• PECH-MAY A, RAMSEY JM, GONZÁLEZ ITTIG RE, GIULIANI M, BERROZPE P, QUINTANA MG & SALOMÓN OD. 2018. Genetic diversity, phylogeography and molecular clock of the Lutzomyia longipalpis complex (Diptera: Psychodidae). PLoS Negl Trop Dis 12: e0006614.
• PESSOA GCD, LOPES JV, ROCHA MF, PINHEIRO LC, ROSA ACL, MICHALSKY ÉM & DIAS ES. 2015. Baseline susceptibility to alpha-cypermethrin in Lutzomyia longipalpis (Lutz & Neiva, 1912) from Lapinha Cave (Brazil). Parasit Vectors 8.
• PIMENTA PFP, MODI GB, PEREIRA ST, SHAHABUDDIN M & SACKS DL. 1997. A novel role for the peritrophic matrix in protecting Leishmania from the hydrolytic activities of the sand fly midgut. Parasitology 115: 359-369.
• PIMENTA PFP, SARAIVA EMB, ROWTON E, MODI GB, GARRAWAY LA, BEVERLEY SM, TURCO SJ & SACKS DL. 1994. Evidence that the vectorial competence of phlebotomine sand flies for different species of Leishmania is controlled by structural polymorphisms in the surface lipophosphoglycan. Proc Natl Acad Sci USA 91: 9155-9159.
• PIMENTA PFP, TURCO SJ, MCCONVILLE MJ, LAWYER PG, PERKINS PV & SACKS DL. 1992. Stage-specific adhesion of Leishmania promastigotes to the sandfly midgut. Science 256: 1812-1815.
• PINHEIRO LJ, PARANAÍBA LF, ALVES AF, PARREIRAS PM, GONTIJO NF, SOARES RP & TAFURI WL. 2018. Salivary gland extract modulates the infection of two Leishmania enriettii strains by interfering with macrophage differentiation in the model of Cavia porcellus. Front Microbiol 9: 969.
• PIRES ACAM, VILLEGAS LEM, CAMPOLINA TB, ORFANÓ AS, PIMENTA PFP & SECUNDINO NFC. 2017. Bacterial diversity of wild-caught Lutzomyia longipalpis (a vector of zoonotic visceral leishmaniasis in Brazil) under distinct physiological conditions by metagenomics analysis. Parasit Vectors 10: 1-9.
• PITALUGA AN, BETEILLE V, LOBO AR, ORTIGÃO-FARIAS JR, DÁVILA AMR, SOUZA AA, RAMALHO-ORTIGÃO JM & TRAUB-CSEKO YM. 2009. EST sequencing of blood-fed and Leishmania-infected midgut of Lutzomyia longipalpis, the principal visceral leishmaniasis vector in the Americas. Mol Genet Genomics 282: 307-317.
• PITALUGA AN, MASON PW & TRAUB-CSEKO YM. 2008. Non-specific antiviral response detected in RNA-treated cultured cells of the sandfly, Lutzomyia longipalpis. Dev Comp Immunol 32: 191-197.
• POTHIRAT T, TANTIWORAWIT A, CHAIWARITH R, JARIYAPAN N, WANNASAN A, SIRIYASATIEN P, SUPPARATPINYO K, BATES MD, KWAKYE-NUAKO G & BATES PA. 2014. First Isolation of Leishmania from Northern Thailand: Case Report, Identification as Leishmania martiniquensis and Phylogenetic Position within the Leishmania enriettii Complex. PLoS Negl Trop Dis 8: e3339.
• PRATES DB, ARAÚJO-SANTOS T, BRODSKYN C, BARRAL-NETTO M, BARRAL A & BORGES VM. 2012. New insights on the inflammatory role of Lutzomyia longipalpis saliva in leishmaniasis. J Parasitol Res 2012: 643029.
• PRATES DB ET AL. 2011. Lutzomyia longipalpis saliva drives apoptosis and enhances parasite burden in neutrophils. J Leukoc Biol 90: 575-582.
• PRATES DB, SANTOS LD, MIRANDA JC, SOUZA APA, PALMA MS, BARRAL-NETTO M & BARRAL A .2008. Changes in Amounts of Total Salivary Gland Proteins of Lutzomyia longipalpis (Diptera: Psychodidae) According to Age and Diet. J Med Entomol 45: 409-413.
• PRUZINOVA K, SADLOVA J, MYSKOVA J, LESTINOVA T, JANDA J & VOLF P. 2018. Leishmania mortality in sand fly blood meal is not species-specific and does not result from direct effect of proteinases. Parasites and Vectors 11: 37.
• RAMALHO-ORTIGÃO JM, KAMHAWI S, JOSHI MB, REYNOSO D, LAWYER PG, DWYER DM, SACKS DL & VALENZUELA JG. 2005. Characterization of a blood activated chitinolytic system in the midgut of the sand fly vectors Lutzomyia longipalpis and Phlebotomus papatasi. Insect Mol Biol 14: 703-712.
• RAMALHO-ORTIGÃO JM, SARAIVA EM & TRAUB-CSEKÖ YM. 2010. Sand Fly- Leishmania Interactions: Long Relationships are Not Necessarily Easy. Open Parasitol J 195-204.
• RAMALHO-ORTIGÃO JM & TRAUB-CSEKÖ YM. 2003. Molecular characterization of Llchit1, a midgut chitinase cDNA from the leishmaniasis vector Lutzomyia longipalpis. Insect Biochem Mol Biol 33: 279-287.
• RÊGO FD, SOUZA GD, DORNELLES LFP & ANDRADE FILHO JD. 2020. Ecology and Molecular Detection of Leishmania infantum Nicolle, 1908 (Kinetoplastida: Trypanosomatida) in Wild-Caught Sand Flies (Psychodidae: Phlebotominae) Collected in Porto Alegre, Rio Grande do Sul: A New Focus of Visceral Leishmaniasis in Brazil. J Med Entomol 56: 519-525.
• REY GJ, FERRO C & BELLO FJ. 2000. Establishment and Characterization of a New Continuous Cell Line from Lutzomyia longipalpis (Diptera: Psychodidae) and its Susceptibility to Infections with Arboviruses and Leishmania chagasi. Mem Inst Oswaldo Cruz 95: 103-110.
• RIBOLLA PEM ET AL. 2016. Leishmania infantum Genetic Diversity and Lutzomyia longipalpis Mitochondrial Haplotypes in Brazil. Biomed Res Int 2016.
• RIVAS GBS, SOUZA NA & PEIXOTO AA 2008. Analysis of the activity patterns of two sympatric sandfly siblings of the Lutzomyia longipalpis species complex from Brazil. Med Vet Entomol 22: 288-290.
• ROCHA L DE S, FALQUETO A, SANTOS CB DOS, GRIMALDI G & CUPOLILLO E. 2011. Possible Implication of the Genetic Composition of the Lutzomyia longipalpis (Diptera: Psychodidae) Populations in the Epidemiology of the Visceral Leishmaniasis. J Med Entomol 48: 1016-1022.
• ROCHA NO, LAMBERT SM, DIAS-LIMA AG, JULIÃO FS & SOUZA BMPS. 2018. Molecular detection of Wolbachia pipientis in natural populations of sandfly vectors of Leishmania infantum in endemic areas: first detection in Lutzomyia longipalpis. Med Vet Entomol 32: 111-114.
• RODRIGUES ACM, SILVA RA, MELO LM, LUCIANO MCS & BEVILAQUA CML. 2014. Epidemiological survey of Lutzomyia longipalpis infected by Leishmania infantum in an endemic area of Brazil. Rev Bras Parasitol Veterinária 23: 55-62.
• ROGERS ME & BATES PA. 2007. Leishmania manipulation of sand fly feeding behavior results in enhanced transmission. PLoS Pathog 3: e91.
• ROGERS ME, CORWARE K, MÜLLER I & BATES PA. 2010. Leishmania infantum proteophosphoglycans regurgitated by the bite of its natural sand fly vector, Lutzomyia longipalpis, promote parasite establishment in mouse skin and skin-distant tissues. Microbes Infect 12: 875-879.
• ROHOUSOVA I, OZENSOY S, OZBEL Y & VOLF P. 2005. Detection of species-specific antibody response of humans and mice bitten by sand flies. Parasitology 130: 493-499.
• ROSE K, CURTIS J, BALDWIN T, MATHIS A, KUMAR B, SAKTHIANANDESWAREN A, SPURCK T, LOW-CHOY J & HANDMAN E. 2004. Cutaneous leishmaniasis in red kangaroos: isolation and characterisation of the causative organisms. Int J Parasitol 34: 655-664.
• SAAB NAA ET AL. 2020. How Lutzomyia longipalpis deals with the complement system present in the ingested blood: The role of soluble inhibitors and the adsorption of factor H by midgut. J Insect Physiol 120: 103992.
• SÁDLOVÁ J & VOLF P. 2009. Peritrophic matrix of Phlebotomus duboscqi and its kinetics during Leishmania major development. Cell Tissue Res 337: 313-325.
• SADLOVA J, YEO M, SEBLOVA V, LEWIS MD, MAURICIO I, VOLF P & MILES MA. 2011. Visualisation of Leishmania donovani fluorescent hybrids during early stage development in the sand fly vector. PLoS One 6: e19851.
• SALOMÓN OD, FELICIANGELI MD, QUINTANA MG, AFONSO MMS & RANGEL EF. 2015. Lutzomyia longipalpis urbanisation and control. Mem Inst Oswaldo Cruz 110: 831-846.
• SANGIORGI B, MIRANDA DN, OLIVEIRA DF, SANTOS EP, GOMES FR, SANTOS EO, BARRAL A & MIRANDA JC. 2012. Natural breeding places for phlebotomine sand flies (Diptera: Psychodidae) in a semiarid region of Bahia state, Brazil. J Trop Med 2012: 124068.
• SANT’ANNA MR, DIAZ-ALBITER H, AGUIAR-MARTINS K, AL-SALEM WS, CAVALCANTE RR, DILLON VM, BATES PA, GENTA FA & DILLON RJ. 2014. Colonisation resistance in the sand fly gut: Leishmania protects Lutzomyia longipalpis from bacterial infection. Parasit Vectors 7: 329.
• SANT’ANNA MR, DIAZ-ALBITER H, MUBARAKI M, DILLON RJ & BATES PA. 2009. Inhibition of trypsin expression in Lutzomyia longipalpis using RNAi enhances the survival of Leishmania. Parasit Vectors 2: 1-10.
• SANT’ANNA MRV, JONES NG, HINDLEY JA, MENDES-SOUSA AF, DILLON RJ, CAVALCANTE RR, ALEXANDER B & BATES PA. 2008. Blood meal identification and parasite detection in laboratory-fed and field-captured Lutzomyia longipalpis by PCR using FTA databasing paper. Acta Trop 107: 230-237.
• SANT’ANNA MR, NASCIMENTO A, ALEXANDER B, DILGER E, CAVALCANTE RR, DIAZ-ALBITER HM, BATES PA & DILLON RJ. 2010. Chicken blood provides a suitable meal for the sand fly Lutzomyia longipalpis and does not inhibit Leishmania development in the gut. Parasit Vectors 3: 1-11.
• SANTINI MS, SALOMÓN OD, ACARDI SA, SANDOVAL EA & TARTAGLINO L. 2010. Comportamento e controle de Lutzomyia longipalpis em foco de leishmaniose visceral urbana na Argentina. Rev Inst Med Trop Sao Paulo 52: 187-191.
• SANTOS VC, ARAUJO RN, MACHADO LAD, PEREIRA MH & GONTIJO NF. 2008. The physiology of the midgut of Lutzomyia longipalpis (Lutz and Neiva 1912): PH in different physiological conditions and mechanisms involved in its control. J Exp Biol 211: 2792-2798.
• SANTOS VC, NUNES CA, PEREIRA MH & GONTIJO NF. 2011. Mechanisms of pH control in the midgut of Lutzomyia longipalpis: Roles for ingested molecules and hormones. J Exp Biol 214: 1411-1418.
• SANTOS MFC, RIBOLLA PEM, ALONSO DP, ANDRADE-FILHO JD, CASARIL AE, FERREIRA AMT, FERNANDES CES, BRAZIL RP & OLIVEIRA AG. 2013. Genetic structure of Lutzomyia longipalpis populations in Mato Grosso do Sul, Brazil, based on microsatellite markers. PLoS One 8: e74268.
• SANTOS VC ET AL. 2014. Host modulation by a parasite: How Leishmania infantum modifies the intestinal environment of Lutzomyia longipalpis to favor its development. PLoS One 9: e111241.
• SCHLEIN Y & JACOBSON RL. 1998. Resistance of Phlebotomus papatasi to infection with Leishmania donovani is modulated by components of the infective bloodmeal. Parasitology 117: 467-473.
• SEBLOVA V, MYŠKOVÁ J, HLAVACOVA J, VOTYPKA J, ANTONIOU M & VOLF P. 2015a. Natural hybrid of Leishmania infantum/L. donovani: Development in Phlebotomus tobbi, P. perniciosus and Lutzomyia longipalpis and comparison with non-hybrid strains differing in tissue tropism. Parasit Vectors 8: 1-8.
• SEBLOVA V, SADLOVA J, VOJTKOVA B, VOTYPKA J, CARPENTER S, BATES PA & VOLF P. 2015b. The biting midge Culicoides sonorensis (Diptera: Ceratopogonidae) is capable of developing late stage infections of Leishmania enriettii. PLoS Negl Trop Dis 9: e0004060.
• SECUNDINO NFC, ARAÚJO MSS, OLIVEIRA GHB, MASSARA CL, CARVALHO OS, LANFREDI RM & PIMENTA PFP. 2002. Preliminary description of a new entomoparasitic nematode infecting Lutzomyia longipalpis sand fly, the vector of visceral leishmaniasis in the new world. J Invertebr Pathol 80: 35-40.
• SECUNDINO NFC, EGER-MANGRICH I, BRAGA EM, SANTORO MM & PIMENTA PFP. 2005. Lutzomyia longipalpis peritrophic matrix: Formation, structure, and chemical composition. J Med Entomol 42: 928-938.
• SECUNDINO NFC, FREITAS VC, MONTEIRO CC, PIRES ACAM, DAVID BA & PIMENTA PFP. 2012. The transmission of Leishmania infantum chagasi by the bite of the Lutzomyia longipalpis to two different vertebrates. Parasit Vectors 5: 2-5.
• SERAFIM TD, COUTINHO-ABREU IV, OLIVEIRA F, MENESES C, KAMHAWI S & VALENZUELA JG. 2018. Sequential blood meals promote Leishmania replication and reverse metacyclogenesis augmenting vector infectivity. Nat Microbiol 3: 548-555.
• SHAW J, ROSA AT, SOUZA A & CRUZ AC. 2003. Transmissão de outros agentes: os flebotomíneos brasileiros como hospedeiros e vetores de determinadas espécies. In: Rangel EF & Lainson R (Eds), Flebotomíneos do Brasil, Rio de Janeiro, p. 337-351.
• SILVA F, GOMES R, PRATES DB, MIRANDA JC, ANDRADE B & BARRAL-NETTO M, BARRAL A. 2005. Inflammatory cell infiltration and high antibody production in BALB/c mice caused by natural exposure to Lutzomyia longipalpis bites. Am J Trop Med Hyg 72: 94-98.
• SILVA MH, NASCIMENTO MDSH, LEONARDO FS, REBELO JMM & PEREIRA SRF. 2011. Genetic differentiation in natural populations of Lutzomyia longipalpis (Lutz & Neiva) (Diptera: Psychodidae) with different phenotypic spot patterns on tergites in males. Neotrop Entomol 40: 501-506.
• SOARES RP, ALTOÉ ECF, ENNES-VIDAL V, COSTA SM DA, RANGEL EF, SOUZA NA, SILVA VC, VOLF P & D’AVILA-LEVY CM. 2017. In vitro inhibition of Leishmania attachment to sandfly midguts and LL-5 cells by divalent metal chelators, anti-gp63 and phosphoglycans. Protist 168: 326-334.
• SOARES RPP, MACEDO ME, ROPERT C, GONTIJO NF, ALMEIDA IC, GAZZINELLI RT, PIMENTA PFP & TURCO SJ. 2002. Leishmania chagasi: lipophosphoglycan characterization and binding to the midgut of the sand fly vector Lutzomyia longipalpis. Mol Biochem Parasitol 121: 213-224.
• SOARES RP, NOGUEIRA PM, SECUNDINO NFC, MARIALVA EF, RÍOS-VELÁSQUEZ CM & PESSOA FAC. 2018. Lutzomyia umbratilis from an area south of the Negro river is refractory to in vitro interaction with Leishmania guyanensis. Mem Inst Oswaldo Cruz 113: 202-205.
• SOARES VYR ET AL. 2014. Identification of blood meal sources of Lutzomyia longipalpis using polymerase chain reaction-restriction fragment length polymorphism analysis of the cytochrome B gene. Mem Inst Oswaldo Cruz 109: 379-383.
• SOARES RPP & TURCO SJ. 2003. Lutzomyia longipalpis (Diptera: Psychodidae: Phlebotominae): A Review. An Acad Bras Cienc 75: 301-330.
• SOUZA NA, ANDRADE-COELHO CA, SILVA VC, WARD RD & PEIXOTO AA. 2009a. Life cycle differences among Brazilian sandflies of the Lutzomyia longipalpis sibling species complex. Med Vet Entomol 23: 287-292.
• SOUZA AP ET AL. 2010. Using recombinant proteins from Lutzomyia longipalpis saliva to estimate human vector exposure in visceral leishmaniasis endemic areas. PLoS Negl Trop Dis 4: e649.
• SOUZA NA, BRAZIL RP & ARAKI AS. 2017. The current status of the Lutzomyia longipalpis (Diptera: Psychodidae: Phlebotominae) species complex. Mem Inst Oswaldo Cruz 112: 161-174.
• SOUZA GD, SANTOS E & ANDRADE-FILHO JD. 2009b. The first report of the main vector of visceral leishmaniasis in America, Lutzomyia longipalpis (Lutz & Neiva) (Diptera: Psychodidae: Phlebotominae), in the state of Rio Grande do Sul, Brazil. Mem Inst Oswaldo Cruz 104: 1181-1182.
• SPIEGEL CN, DIAS DBS, ARAKI AS, HAMILTON JGC, BRAZIL RP & JONES TM. 2016. The Lutzomyia longipalpis complex: a brief natural history of aggregation-sex pheromone communication. Parasit Vectors 9: 1-15.
• SURENDRAN SN, KARUNARATNE SHPP, ADAMSN Z, HEMINGWAY J & HAWKES NJ. 2005. Molecular and biochemical characterization of a sand fly population from Sri Lanka: evidence for insecticide resistance due to altered esterases and insensitive acetylcholinesterase. Bull Entomol Res 95: 371-380.
• SVÁROVSKÁ A, ANT TH, SEBLOVÁ V, JECNÁ L, BEVERLEY SM & VOLF P. 2010. Leishmania major glycosylation mutants require phosphoglycans (lpg2 -) but not lipophosphoglycan (lpg1-) for survival in permissive sand fly vectors. PLoS Negl Trop Dis 4: e580.
• SVENSJÖ E, SARAIVA EM, AMENDOLA RS, BARJA-FIDALGO C, BOZZA MT, LERNER EA, TEIXEIRA MM & SCHARFSTEIN J. 2012. Maxadilan, the Lutzomyia longipalpis vasodilator, drives plasma leakage via PAC1-CXCR1/2-pathway. Microvasc Res 83: 185-193.
• SVENSJÖ E, SARAIVA EM, BOZZA MT, OLIVEIRA SMP, LERNER EA & SCHARFSTEIN J. 2009. Salivary gland homogenates of Lutzomyia longipalpis and its vasodilatory peptide maxadilan cause plasma leakage via PAC1 receptor activation. J Vasc Res 46: 435-446.
• SVOBODOVÁ M, BATES PA & VOLF P. 1997. Detection of lectin activity in Leishmania promastigotes and amastigotes. Acta Trop 68: 23-35.
• TAVARES ET AL. 2011. Lutzomyia longipalpis saliva or salivary protein LJM19 protects against Leishmania braziliensis and the saliva of its vector, Lutzomyia intermedia. PLoS Negl Trop Dis 5: e1169.
• TEIXEIRA C ET AL. 2010. Discovery of markers of exposure specific to bites of Lutzomyia longipalpis, the vector of Leishmania infantum chagasi in Latin America. PLoS Negl Trop Dis 4: e638.
• TEIXEIRA CR, SANTOS CS, PRATES DB, SANTOS RT, ARAÚJO-SANTOS T, SOUZA-NETO SM, BORGES VM, BARRAL-NETTO M & BRODSKYN CI. 2018. Lutzomyia longipalpis saliva drives interleukin-17-induced neutrophil recruitment favoring Leishmania infantum infection. Front Microbiol 9: 881.
• TEIXEIRA CR ET AL. 2005. Saliva from Lutzomyia longipalpis induces CC chemokine ligand 2/monocyte chemoattractant protein-1 expression and macrophage recruitment. J Immunol 175: 8346-8353.
• TELLERIA EL, ARAÚJO APO, SECUNDINO NFC, D’AVILA-LEVY CM & TRAUB-CSEKÖ YM. 2010. Trypsin-like serine proteases in Lutzomyia longipalpis - expression, activity and possible modulation by Leishmania infantum chagasi. PLoS One 5: e10697.
• TELLERIA EL, PITALUGA AN, ORTIGÃO-FARIAS JR, ARAÚJO APO, RAMALHO-ORTIGÃO JM & TRAUB-CSEKO YM. 2007. Constitutive and blood meal-induced trypsin genes in Lutzomyia longipalpis. Arch Insect Biochem Physiol 66: 53-63.
• TELLERIA EL, SANT’ANNA MRV, ALKURBI MO, PITALUGA AN, DILLON RJ & TRAUB-CSEKÖ YM 2013. Bacterial feeding, Leishmania infection and distinct infection routes induce differential defensin expression in Lutzomyia longipalpis. Parasit Vectors 6: 2-9.
• TESH RB & MODI GB. 1983. Development of a continuous cell line from the sand fly Lutzomyia longipalpis (Diptera: Psychodidae), and its susceptibility to infection with arboviruses. J Med Entomol 20: 199-202.
• THIAKAKI M, ROHOUSOVA I, VOLFOVA V, VOLF P, CHANG KP & SOTERIADOU K. 2005. Sand fly specificity of saliva-mediated protective immunity in Leishmania amazonensis-BALB/c mouse model. Microbes Infect 7:760-766.
• TINOCO-NUNES B, TELLERIA EL, SILVA-NEVES M, MARQUES C, AZEVEDO-BRITO DA, PITALUGA NA & TRAUB-CSEKÖ YM. 2016. The sandfly Lutzomyia longipalpis LL5 embryonic cell line has active Toll and Imd pathways and shows immune responses to bacteria, yeast and Leishmania. Parasit Vectors 9: 1-11.
• VALE VF, MOREIRA BH, MORAES CS, PEREIRA MH, GENTA FA & GONTIJO NF. 2012. Carbohydrate digestion in Lutzomyia longipalpis’ larvae (Diptera - Psychodidae). J Insect Physiol 58: 1314-1324.
• VALE VF, PEREIRA MH & GONTIJO NF. 2007. Midgut pH profile and protein digestion in the larvae of Lutzomyia longipalpis (Diptera: Psychodidae). J Insect Physiol 53: 1151-1159.
• VALENZUELA JG, GARFIELD M, ROWTON ED & PHAM VM. 2004. Identification of the most abundant secreted proteins from the salivary glands of the sand fly Lutzomyia longipalpis, vector of Leishmania chagasi. J Exp Biol 207: 3717-3729.
• VASCONCELOS CO, COÊLHO ZCB, CHAVES CS, TEIXEIRA CR, POMPEU MML & TEIXEIRA MJ. 2014. Distinct cellular migration induced by Leishmania infantum chagasi and saliva from Lutzomyia longipalpis in a hemorrhagic pool model. Rev Inst Med Trop Sao Paulo 56: 21-27.
• VIGODER FM, ARAKI AS, BAUZER LGSR, SOUZA NA, BRAZIL RP & PEIXOTO AA. 2010. Lovesongs and period gene polymorphisms indicate Lutzomyia cruzi (Mangabeira, 1938) as a sibling species of the Lutzomyia longipalpis (Lutz and Neiva, 1912) complex. Infect Genet Evol 10: 734-739.
• VIGODER FM, SOUZA NA, BRAZIL RP, BRUNO RV., COSTA PL, RITCHIE MG, KLACZKO LB & PEIXOTO AA. 2015. Phenotypic differentiation in love song traits among sibling species of the Lutzomyia longipalpis complex in Brazil. Parasit Vectors 8: 1-14.
• VINHAS V, ANDRADE BB, PAES F, BOMURA A, CLARENCIO J, MIRANDA JC, BÁFICA A, BARRAL A & BARRAL-NETTO M. 2007. Human anti-saliva immune response following experimental exposure to the visceral leishmaniasis vector, Lutzomyia longipalpis. Eur J Immunol 37: 3111-3121.
• VOLF P & MYSKOVA J. 2007. Sand flies and Leishmania: specific versus permissive vectors. Trends Parasitol 23: 91-92.
• VOLF P & ROHOUSOVA I. 2001. Species-specific antigens in salivary glands of phlebotomine sandflies. Parasitology 122: 37-41.
• VOLF P, TESAROVA P & NOHYNKOVA E. 2000. Salivary proteins and glycoproteins in phlebotomine sandflies of various species, sex and age. Med Vet Entomol 14: 251-256.
• VOLFOVA V, HOSTOMSKA J, CERNY M, VOTYPKA J & VOLF P. 2008. Hyaluronidase of bloodsucking insects and its enhancing effect on Leishmania infection in mice. PLoS Negl Trop Dis 2: e294.
• WATTS PC, HAMILTON JGC, WARD RD, NOYES HA, SOUZA NA, KEMP SJ, FELICIANGELI MD, BRAZIL R & MAINGON RDC. 2005. Male sex pheromones and the phylogeographic structure of the Lutzomyia longipalpis species complex (Diptera: Psychodidae) from Brazil and Venezuela. Am J Trop Med Hyg 73: 734-743.
• WHEAT WH, ARTHUN EN, SPENCER JS, REGAN DP, TITUS RG & DOW SW. 2017. Immunization against full-length protein and peptides from the Lutzomyia longipalpis sand fly salivary component maxadilan protects against Leishmania major infection in a murine model. Vaccine 35: 6611-6619.
• WHEAT WH, PAUKEN KE, MORRIS RV & TITUS RG. 2008. Lutzomyia longipalpis salivary peptide Maxadilan alters murine dendritic cell expression of CD80/86, CCR7, and cytokine secretion and reprograms dendritic cell-mediated cytokine release from cultures containing allogeneic T cells. J Immunol 180: 8286-8298.
• WILSON R, BATES MD, DOSTALOVA A, JECNA L, DILLON RJ, VOLF P & BATES PA. 2010. Stage-specific adhesion of Leishmania promastigotes to sand fly midguts assessed using an improved comparative binding assay. PLoS Negl Trop Dis 4: e816.
• XIMENES MFFM, CASTELLÓN EG, SOUZA MF, FREITAS RA, PEARSON RD, WILSON ME & JERÔNIMO SMB. 2000. Distribution of phlebotomine sand flies (Diptera: Psychodidae) in the State of Rio Grande do Norte, Brazil. J Med Entomol 37: 162-169.
• XU X ET AL. 2011. Structure and function of a “yellow” protein from saliva of the sand fly Lutzomyia longipalpis that confers protective immunity against Leishmania major infection. J Biol Chem 286: 32383-32393.
• YANG Z &WU Y. 2019. Improved annotation of Lutzomyia longipalpis genome using bioinformatics analysis. PeerJ 7: e7862.

Publication Dates

• Publication in this collection
  30 Apr 2021
• Date of issue
  2021

History

• Received
  21 Feb 2020
• Accepted
  17 May 2020