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Structural differences in gut bacteria communities in developmental stages of natural populations of Lutzomyia evansi from Colombia's Caribbean coast



Lutzomyia evansi, a phlebotomine insect endemic to Colombia’s Caribbean coast, is considered to be the main vector of visceral and cutaneous leishmaniasis in the region. Although insects of this species can harbor pathogenic and non-pathogenic microorganisms in their intestinal microbiota, there is little information available about the diversity of gut bacteria present in Lutzomyia evansi. In this study, conventional microbiological methods and molecular tools were used to assess the composition of bacterial communities associated with Lutzomyia evansi guts in immature and adult stages of natural populations from the department of Sucre (Caribbean coast of Colombia).


Sand flies were collected from two locations (peri-urban and jungle biotype) in the Department of Sucre (Caribbean coast of Colombia). A total of 752 Lutzomyia evansi intestines were dissected. In this study, 125 bacterial strains were isolated from different culture media (LB Agar, MacConkey Agar). Different methods were used for bacterial identification, including ribosomal intergenic spacer analysis (RISA) and analysis of the 16S rRNA and gyrB gene sequences. The genetic profiles of the bacterial populations were generated and temporal temperature gradient gel electrophoresis (TTGE) was used to compare them with total gut DNA. We also used PCR and DNA sequence analysis to determine the presence of Wolbachia endosymbiont bacteria and Leishmania parasites.


The culture-dependent technique showed that the dominant intestinal bacteria isolated belong to Acinetobacter, Enterobacter, Pseudomonas, Ochrobactrum, Shinella and Paenibacillus in the larval stage; Lysobacter, Microbacterium, Streptomyces, Bacillus and Rummeliibacillus in the pupal stage; and Staphylococcus, Streptomyces, Brevibacterium, Acinetobacter, Enterobacter and Pantoea in the adult stage. Statistical analysis revealed significant differences between the fingerprint patterns of the PCR-TTGE bands in bacterial communities from immature and adult stages. Additionally, differences were found in bacterial community structure in fed females, unfed females, males and larvae. The intestinal bacteria detected by PCR-TTGE were Enterobacter cloacae and Bacillus thuringiensis, which were present in different life stages of Lu. evansi, and Burkholderia cenocepacia and Bacillus gibsonii, which were detected only in the larval stage. Wolbachia and Leishmania were not detected in gut samples of Lutzomyia evansi.


The analyses conducted using microbiological and molecular approaches indicated significant variations in the bacterial communities associated with the gut of Lu. evansi, depending on the developmental stage and food source. We propose that these elements affect microbial diversity in L. evansi guts and may in turn influence pathogen transmission to humans bitten by this insect.


Insects from the genus Lutzomyia França, 1924 (subfamily Phlebotominae) are of great interest in biological studies because of their ability to transmit parasites of the genus Leishmania Ross, 1903 [1] to vertebrates, as well as other pathogens, including the bacterium Bartonella baciliformis and viruses from the families Bunyaviridae, Reoviridae and Rhabdoviridae [24].

The potential of Lutzomyia spp. to transmit parasites can be attributed primarily to the hematophagic habits of females, which are the only ones that ingest blood as a necessary source of protein for the maturation of their eggs [5]. Various studies on feeding preferences suggest that phlebotomine insects have eclectic feeding habits and can take blood from different vertebrates [5, 6]. There are also autogenous species that use protein stocks acquired during the larval stage to complete their first gonadotropic cycle [7]. For survival and movement, sap, nectar or aphid secretions are the most-described carbohydrate sources [8].

As described above, phlebotomine insects need a varied diet and have the ability to digest complex molecules throughout the different stages of their development [9, 10]. The different bacterial communities these insects acquire from the environment and via their eating habits are involved in the plant material digestion process and promote the availability of iron from erythrocytes and essential amino acids and vitamins from the B complex [11, 12].

The metabolic importance of bacterial communities associated with the guts of phlebotomine insects has led to the emergence of several new questions related to the parasite-vector interaction. The most basic and essential of these include what bacterial communities are present in the gut and how they interact with parasites [13]. Other questions relate to the location of bacteria in the intestine and other organs of physiological relevance [13], the identity of bacteria that remain during metamorphosis [14], the ability of bacteria to express recombinant molecules that display antitrypanosomal or antiviral activity [15], and the existence of intestinal bacteria with insecticidal properties or the ability to alter insect life-cycles [16].

In America, bacterial diversity analysis has only been recorded for two species: Lutzomyia longipalpis and Lu. cruzi (adult stages from colonies and natural populations in Brazil and Colombia) [1720]. The gut bacteria isolates registered for these species mainly belong to the genera Serratia, Enterobacter, Acinetobacter and Pseudomonas [19]. Microbiota associated with immature stages in natural breeding sites have not yet been evaluated, due to several difficulties related to their recovery and identification [21]. The large number of Lutzomyia spp. in Colombia serves as a motivation for exploring gut bacteria in species that have been found naturally infected with Leishmania.

Around 163 species from the subfamily Phlebotominae are registered in Colombia. Fourteen of these have been reported as vectors for different Leishmania spp. [22]; however Lu. longipalpis is the only species whose midgut microbiota have been described [19, 20]. Lutzomyia evansi is the most abundant on Colombia’s Caribbean coast; this species transmits Leishmania infantum chagasi and Leishmania braziliensis [23]. Studies have been conducted in order to understand the role of Lu. evansi as a vector of Leishmania infantum (vector competence, life-cycle, insecticide resistance, genetic structure, blood intake and bionomics) [24]. However, the composition of bacterial communities present in the gut and the changes in the microbiota associated with food sources, developmental stages, and the presence of parasite infections, have not been studied in this species. Among the gut microbiota, molecular detection and identification of endosymbionts such as Wolbachia is important due to their potential use for decreasing the population density of Lutzomyia species and interfering with parasite multiplication and thus, with Leishmania transmission [13, 16, 17].

The several feeding events of natural populations of Lu. evansi and their interaction with the environment (including multiple hosts), allows species to acquire a complex and dynamic microbiome. In recent years, a polyphasic approach incorporating classical microbiological techniques and culture-independent methods based on direct analysis of DNA (or RNA) cultures has gained popularity. This approach has had reliable and effective results with regard to detecting and identifying microorganisms, leading to major advances in the understanding of this complex microbial ecosystem [13, 16, 19]. For the culture-independent approach, fingerprinting methods represent a complete alternative for the study of intestinal microbiota. These methods include temperature gradient gel electrophoresis (TTGE), which is a powerful genetic tool for establishing distinctive profiles of microbial communities among different samples [25, 26].

The culture-dependent techniques, allow access to cultivable bacteria that may have anti-leishmanicidal and entomopathogenic activity. It is a platform bioprospecting, for this reason it is necessary to include bacterial isolation as a first phase. TTGE allows a quick screenshot of the abundance and presence of different bacterial communities. Therefore, these two approaches should be complementary.

The aim of our study was to investigate the diversity of bacterial communities associated with the guts of two natural populations (jungle and peri-urban biotype) of Lu. evansi in different developmental stages (adult and immature), through culture-dependent and culture-independent methods. This species was abundant in the endemic foci of visceral leishmaniasis on Colombia’s Caribbean coast and representative of the sand fly diversity in this area.


Collection, processing and identification of sand flies

Sand flies were collected from two locations in the Department of Sucre (Caribbean coast of Colombia). The first location was associated with a peri-urban biotype near the Ovejas municipality (75°13'W, 9°31'N; 277 metres above sea level, masl), classified as a tropical dry forest ecosystem [27]. The second location corresponded to a jungle biotype at the “Primates” Wildlife Experimental Station (09°31'48.0"N, 75°21'4.3"W, 220 masl) in the municipality of Colosó, located within the Protected Forest Reserve “Serranía de Coraza”. This nature reserve is classified as a transitional ecosystem of premontane dry forest to tropical dry forest [27]. Adult stage samples were collected from Ovejas and Colosó during July and October 2013 and January 2014, while immature stages were searched for in Ovejas during October and November 2013.

The adult specimens were collected from the two locations using Shannon-type extra-domiciliary white light traps that remained active between 18:00 and 22:00 h. The adult sand flies collected were transported live to the laboratory in entomological cages to obtain the gut and morphological structures (male genitalia and female spermathecae) for taxonomic identification of Lu. evansi using a taxonomic key [28].

Immature stages of Lu. evansi (larvae and pupae) were isolated from substrates extracted from potential breeding sites, usually at the base of Cordia dentata shrubs (common name: Uvito) associated with the urban environment of Ovejas (results not shown). The search was performed with a stereomicroscope, using the direct view technique [21]. Immature specimens were immediately processed to obtain the gut; the rest of each specimen (integument and head) was stored at -20 °C for DNA extraction [29]. The barcode region of the cytochrome c oxidase I gene [30] was amplified and high quality sequences were obtained in order to estimate the taxonomic identity of the immature specimens by comparing them with adult specimen sequences from the same locality and others reported in the NCBI GenBank database (results not shown).

Lutzomyia evansi guts

Processed adult sand flies confirmed as Lu. evansi were categorized into three groups and represented by source as follows: fed females (Colosó coded as FFC; Ovejas coded as FFO), unfed females (Colosó coded as UFC; Ovejas coded as UFO) and males (Colosó coded as MC; Ovejas coded as MV) (Fig. 1). Isolation of sand fly guts was conducted in a sterile environment. Prior to gut dissection, adult and immature specimens were washed with 50 μl of 1× PBS (Phosphate Buffered Saline) and Tween 20, centrifuged at 3000× g for 5 min, and submerged in a 70 % ethanol wash for 2 min to remove excess microvilli, dust and exogenous bacteria; the latter were removed. After the head was removed, the gut was microdissected and isolated aseptically with stilettos on a sterile glass slide under a stereoscope in 1× PBS buffer. Intestinal homogenate was obtained by lysis of the epithelium, only when the gut was placed in sterile eppendorf tubes, to ensure the absence of external bacteria. This procedure was developed according to previously described protocols on intestinal microbiota sand flies [14, 17, 18, 20]. The rest of the dissected insect body was mounted on a slide for morphological identification.

Fig. 1
figure 1

Dissection and obtaining of guts. A1: male; A2: gut of a male; B1: unfed female; B2: gut of an unfed female; C1: fed female; C2: gut of fed female; D1: fourth-instar larvae recovered in natural breeding sites; D2: gut of a fourth-instar larva recovered in natural breeding sites; E1: pupa recovered in natural breeding sites; E2: gut of a pupa recovered in natural breeding sites

Gut pools were formed according to locality and physiological stage; these were re-suspended in 100 μl of sterile PBS. Gut groups (n = 50) were macerated and then vortexed for 30 s to break the intestinal wall and obtain intestinal homogenate. The guts of larvae (Ovejas, L4) and pupae (Ovejas, PP) (Fig. 1), were processed individually prior to taxonomic identification. Each intestinal homogenate was preserved and processed cold. Half was used for conventional microbiological methods of cultivable fraction (CF) and the remainder was used for the culture-independent molecular approach of the total intestinal bacteria fraction (TF). Given the lack of knowledge on the biology of immature stages and the lack of taxonomic keys for these forms, in this study the number of samples of immature specimens was lower and processed individually with respect to adult stages. To date no article of intestinal microbiota has included immature stages isolated from natural breeding sites.

Culture-dependent assays

Isolation of bacteria and colony-forming units

In this study, cultivable and aerobic midgut bacteria were isolated. We used methods and culture media described in other studies on intestinal microbiota in sand flies. Gut homogenates were cultured using serial dilutions for surface plating on LB Agar (Merck, Bogota, Colombia) and MacConkey Agar (Merck, Bogota, Colombia) [31]. The plates were incubated aerobically at 33 °C for 24 to 48 h, as described in previous studies [14, 17, 18, 20]. LB Agar was chosen as a non-selective medium to promote growth of a diversity of microbes including nutritionally fastidious bacteria. MacConkey agar is a selective and differential culture medium for bacteria designed to selectively isolate Gram-negative and enteric (normally found in the intestinal tract) bacilli and differentiate them based on lactose fermentation. A test tube containing LB (Merck) broth opened near the dissection area constituted our sterility control during the dissection process. The total number of colony-forming units (CFU) was determined for each homogenate. The isolates selected for molecular identification were purified; they were then characterized macroscopically based on colony characteristics and microscopically by Gram staining. All isolates were cryopreserved in 20 % glycerol at -80 °C. Pearson Chi-Square tests based on the colony counting was performed using SPSS software ver. 18 to detect statistical differences in bacterial populations isolated from Lu. evansi.

PCR amplification of the internal transcribed spacer (ITS) between the 23S and 16S ribosomal gene, 16S rRNA gene and gyrB gene

For total DNA extraction, each colony was incubated for 10 min at 95 °C in 100 μl of sterile Tris-EDTA 1X to generate cell lysis. This was followed by centrifugation at 10,000 rpm for 5 min, in order to obtain the DNA in the supernatant for use as a template in the PCR assays (colony PCR Protocol) [32]. The template DNA of all isolates was initially assessed by amplifying the ITS region using the primers L1 (5'-CAA GGC ATC CAC CGT-3') and G1 (5'-GAA GTC GTA ACA AGG-3') [32] as previously described [33]. GelCompar II software (Applied Maths Biosystems, Belgium) was used to build a dendrogram with the banding patterns of the ITS region. A distance matrix was calculated using the Dice coefficient [34] and cluster analysis was performed using the unweighted arithmetic average algorithm (UPGMA) [35].

In order to properly represent the bacterial diversity associated with guts of adult and immature Lu. evansi, similarity of ≥ 70 % between ITS standards was established as a criterion for selecting bacterial isolates to be considered for the subsequent molecular identification assays. The DNA isolated from the selected colonies was used to amplify the 16S rRNA gene (1.5 kbp), using the primers Eubac 27 F (5'-AGA GTT TGA TCC TGG CTC AG-3') and 1492R (5'-GGT TAC CTT GTT ACG ACT T-3' [32] as previously described [36]. The gyrB gene (encodes the B-subunit of DNA gyrase, a type II DNA topoisomerase) was amplified from the same template DNA of some isolates [37]. The primers UP1 (5'-GAA GTC ATC ATG ACC GTT CTG CAY GCN GGN GGN AAR TTY GA-3') and UP-2r (5'-AGC AGG GTA CGG ATG TGC GAG CCR TCN ACR TCN GCR TCN GTC AT-3') were used with a reaction mixture and a thermal profile designed to amplify two conserved regions of approximately 1.26 kbp of the gyrB gene [38]. Positive and negative controls (DNA from pure cultures of Escherichia coli or Bacillus cereus and ultrapure water) were routinely included in all PCR reactions for both genes.

PCR amplicons of partial 16S rRNA and the gyrB gene were verified by visualization in 1 % agarose gels and purified using Wizard PCR Preps (Promega, Medellin, Colombia) and the double-stranded DNA was sequenced in both directions using the ABI PRISM 3700 DNA analyzer service (Applied Biosystems) of Macrogen Company Inc. in Korea. The nucleotide sequences of the 16S rRNA and gyrB genes reported in this study were submitted to the NCBI GenBank database (Tables 2 and 4).

Bacterial identity and phylogenetic relationships

The sequences of the 16S rRNA and gyrB genes were edited using Bioedit v7.2.5 [39], in order to obtain consensus sequences for each isolate. Subsequently, they were compared with GenBank and RDP reference sequences, using BLASTN (National Center for Biotechnology Information; to confirm the identity and the taxonomic identity of the nucleotide fragment obtained, with ≥ 97 % similarity values. The ClustalW algorithm built in MEGA 5 [40] was used to align sequences of the 16S rRNA and gyrB genes with reference sequences of bacterial species found in GenBank as a result of the similarity search. The sequence matrix of delimited fragments of the 16S rRNA and gyrB genes was used to generate neighbor-joining dendrograms [41], using the DNA correction distances of the Kimura 2-parameter model [42] to provide a graphical representation of the clustering patterns between species. Verification and/or validation of recombination events and the presence of chimeras were carried out with RDP4 software [43] to ensure the accuracy of the nucleotide variability with respect to the sequences previously reported in the GenBank database.

Culture-independent assays

Temporal temperature gradient gel electrophoresis (PCR-TTGE)

TTGE protocol was used to monitor the dynamic changes in the intestinal microbe population in adult and immature specimens. Total gut DNA was extracted using the Ultra CleanTM Soil DNA Isolation Kit (MO BIO Laboratories, Inc., California, USA), according to the manufacturer’s instructions. Final DNA aliquots were quantified using an ND-100 Nanodrop Thermo Scientific spectrophotometer (Thermo Fisher Scientific Inc, Massachusetts, USA). The quality of genomic DNA was analyzed on 1 % agarose gels, followed by EZ-visionTM DNA 6X STAIN (AMRESCO). DNA samples were subjected to 16S rRNA gene amplification (V3–V6 variable regions) to obtain a 566 bp fragment with primers specific to the conserved domains. These were 341 F (5'-CCT GCA GGA GGC AGC AG-3'), with an extra GC termination at the 5' end, and 907R (5'-CCC TGA CGT GTT ATT CAA TTC Y-3') [44]. PCR reaction conditions were implemented as described previously [45].

All PCR products were checked for amplification by electrophoresis on 1 % (w/v) agarose gels. PCR products were concentrated with Concentrator (Eppendorf 5301) and dissolved in 20 μl of double-distilled H2O. Approximately 600 ng of amplified DNA were loaded into each well. TTGE was performed using the DCode TM Universal mutation detection system (Bio-Rad Laboratories, Hercules, CA, USA) with a 6 % (w/v) polyacrylamide-7 M urea gel and a 1.25× TAE running buffer (40 mM Tris base, 20 mM sodium acetate, 1 mM EDTA) at a constant voltage of 55 V for 15 h. The initial temperature was 66 °C, the final temperature was 69 °C, and a ramp of 0.2 °C h-1 was applied. Additionally, each gel contained at least two marker lanes consisting of a 100 bp ladder (Fermentas, Burlington, USA) and another consisting of 16S fragments of two reference bacterial strains (Enterobacter cloacae: KU134778 and Acinetobacter colcaeceticus: KU134748).

Polyacrylamide gels were stained with SafeView™ DNA stain (Applied Biological Materials, Richmond, Canada) and imaged with a digital scanner. Banding patterns were analyzed through Pearson correlation and Complete Linkage clustering [46] using GelCompar II software (Applied Biosystems Maths, Massachusetts, USA) [47], in order to detect differences in diversity and abundance profiles among the gut samples. Bands of interest were excised and the DNA was eluted in 60 μl of ultrapure water. Five microliters of the DNA elution were re-amplified using the same primers, 341 F (without the GC tail) and 907R. Bands that were successfully re-amplified were sent for sequencing (Macrogen, Korea). Further analyses were carried out using the methods and procedures described above for the isolate sequences. A cluster analysis was constructed with the Dice coefficient, using an unweighted pair-group average with Euclidean distance estimations (GelCompar II software, Applied Maths Biosystems).

Detection of Wolbachia bacteria and Leishmania parasite

The intestinal DNA of female pools collected in Ovejas and Colosó was also searched for Wolbachia endosymbiont bacteria and the Leishmania parasite. The specific primers wsp81F (5'-TGG TCC AAT AAG TGA TGA AGA AAC-3') and wsp691R (5'-AAA AAT TAA ACG CTA CTC CA-3') were used to detect Wolbachia. These primers amplify a partial fragment (590–632 bp) of the gene that encodes Wolbachia main surface protein (WSP) [48]. The reaction mixture and working conditions used for detecting Wolbachia included a 2 μl DNA (30 ng) sample in a final reaction volume of 20 μl [49]. A PCR positive control was included, which consisted of DNA obtained from 10 Aedes (Stegomyia) aegypti larvae (L4) infected with a Wolbachia reference strain (Group A, wMel strain) under insectary laboratory conditions.

To detect Leishmania infection, the specific primers HSP70-F25 (5'-GGA CGC CGG CAC GAT TKC T-3') and HSP70-R617 (5'-CGA AGA AGT CCG ATA CGA GGG A-3') were used, as described by Fraga et al. [50]. These primers amplify a partial fragment (593 bp) of the HSP-70 N gene (cytoplasmic Heat Shock Protein 70).

Data analysis

A descriptive analysis was performed to determine possible differences in the gut bacterial load (colony forming units) in adult (fed females, unfed females, males) and immature stages (larvae, pupae) of Lu. evansi. The XLSTAT 3.04 ( program was used to generate a simple correspondence analysis between bacterial isolates and the TTGE profiles, Lu. evansi stages and sample origin, in order to distinguish possible trends or associations. Finally, a presence/absence matrix was used to perform an analysis of similarity (ANOSIM) based on the Bray-Curtis index, using PAST software, version 2.17 [51]. The analysis of similarity was used to examine the statistical significance of differences between the TTGE profiles.


Lutzomyia evansi guts

A total of 752 Lu. evansi intestines were dissected and processed form 31 pools, each containing 13–50 adult intestines. Analysis of individual forms was performed for immature stages (Table 1). We obtained taxonomic identifications of adults from both localities by using dichotomous keys [28], and confirmed these with the COI mitochondrial marker. Immature stages were also identified with the COI mitochondrial marker (results not shown).

Table 1 Description of samples of adult and immature stages of Lutzomyia evansi collected and processed for analysis of bacterial flora associated with the gut

Cultured bacterial community of Lu. evansi

In this study, 125 bacterial strains were isolated from different culture media (LB Agar, MacConkey Agar) (Table 1). The results showed significant differences (Chi-square test: χ 2 = 11.35, df = 7, P < 0.004) between stages in terms of the CFU count, which was highest for larvae in LB agar (CFU/intestine = 348.4 × 10-3), and lowest for pupae in both culture media (CFU /intestine = 1.25 × 10-3) (Fig. 2). In adults, the bacterial load was lower in male populations from Colosó (CFU/intestine = 8.8 × 10-3) than in the pools of fed females (CFU/intestine = 80.3 × 10-3–72.5 × 10-3) (Chi-square test: χ 2 = 150.64, df = 3, P < 0.001) and unfed females (CFU/intestine = 56.8 × 10-3–115 × 10-3) (Chi-square test: χ 2 = 215.75, df = 8, P < 0.001) (Fig. 2). After morphological (Gram stain) and molecular characterization of the isolates based on RISA pattern analysis revealed differences of more than 30 %, we found that 72.5 % were Gram-negative and 27.4 % were Gram-positive (mainly pupae). Fifty-one presumably different isolates were selected for 16S rDNA sequencing; 26 isolates from this group were selected for additional gyrB gene sequence analysis.

Fig. 2
figure 2

Bacterial counts (CFU) in Lu. evansi guts, obtained in media LB agar and MacConkey. Abbreviations: FFO, fed females from Ovejas; FFC, fed females from Coloso; UFO, unfed females from Ovejas; UFC, unfed females from Coloso; MC, males from Coloso; MV, males from Ovejas; L4, larvae from Ovejas; PP, pupae from Ovejas

Identification of bacterial isolates using 16S rDNA sequences

Three different predominant phyla (Proteobacteria, Actinobacteria and Firmicutes) with high similarity percentages (97–100 %) were isolated based on the similarity analysis of the related sequences from RDP and BLAST (NCBI database) (Tables 2 and 3 and Figs. 3 and 4).

Table 2 Closest phylogenetic identification of bacteria isolated from the digestive tracts of adult Lutzomyia evansi, according to their similarity with 16S rRNA gene sequences in the GenBank and RDP II databases
Table 3 Closest phylogenetic identification of the 16S rDNA sequences of bacteria isolated from the digestive tracts of immature Lutzomyia evansi from the Ovejas locality, according to their similarity with with sequences registered in the GenBank and RDP II databases
Fig. 3
figure 3

Neighbor-joining dendrogram for partial 16S rDNA sequences of bacteria obtained from adult stages of Lutzomyia evansi collected in the municipalities of Ovejas and Colosó (Sucre Department, Colombia), based on the Kimura 2-parameter model. Numbers at the nodes represent bootstrap values. The robustness of the phylogeny was tested by bootstrap analysis using 1000 iterations. Abbreviations: FFO, fed females from Ovejas; FFC, fed females from Coloso; UFO, unfed females from Ovejas; UFC, unfed females from Coloso; MC, males from Coloso; MV, males from Ovejas

Fig. 4
figure 4

Neighbor-joining dendrogram for partial 16S rDNA sequences of bacteria obtained from immature stages of Lutzomyia evansi collected in the Ovejas municipality (Sucre Department, Colombia), based on the Kimura 2-parameter model. Numbers at the nodes represent bootstrap values. The robustness of the phylogeny was tested by bootstrap analysis using 1000 iterations. Abbreviations: L4, larvae from Ovejas; PP, pupae from Ovejas

The genera Enterobacter, Pseudomonas and Acinetobacter were more frequent in adults (Table 2). The highest number of bacterial genera was found in the larval stage (n = 8), with the Ochrobactrum spp. being most prominent. The genera Lysobacter, Microbacterium, Bacillus, Streptomyces and Rummeliibacillus were identified in the pupal stage (Table 3). The diversity of aerobic bacterial flora in Lu. evansi from the two localities, urban and wild environments, was represented by 27 phylotypes at the species level (Tables 2 and 3).

Bacterial communities in unfed females from both locations had the species Acinetobacter calcoaceticus, Pseudomona putida and E. cloacae in common (Table 2). For fed females, the species E. aerogenes was prominent, and the presence of A. calcoaceticus was also observed (Table 2). In males, a slight similarity with the bacterial communities in the fed females was observed, with P. putida and E. aerogenes present (Table 2). Two species, Bacillus anthracis and Lysobacter soli, were only found in the guts of Lu. evansi larvae and pupae (Table 3). Ochrobactrum anthropi appeared more frequently in larvae. The analyses indicated significant variations in the bacterial species, depending on the developmental stage and origin of the insects (Tables 2 and 3).

Neighbor-joining cluster analysis of 16S rRNA and gyrB gene sequences from bacterial isolates

Molecular identification using 16S rDNA sequences from Lu. evansi isolates showed a high level of correspondence with sequences deposited in BlastN at species level and among different taxonomic divisions (Figs. 3 and 4). The analysis showed that most clusters had bootstrap values ranging between 96 and 100 % for bacterial isolates from Lu. evansi guts from different stages, geographical locations and types of natural environment (wild and urban). This was observed in the P. putida and A. calcoaceticus clusters in adults (Fig. 3) and in the L. soli and B. anthracis clusters in immature specimens (Fig. 4).

For some isolates, the identity was confirmed with the gyrB gene sequences due to difficulties that have been reported with some genera, including Enterebacter and Pseudomonas, particularly for environmental samples [38]. In our study, the gyrB molecular marker worked correctly with Pseudomonas species and other genera. This result is consistent with the results of the neighbor-joining analysis of the gyrB gene sequence, showing clusters and similar bootstrap values (99–100 %) for most of the bacterial isolates (A. calcoaceticus, Pantoea ananatis, O. anthropi, P. putida, P. aeruginosa and P. otitidis) that were also identified with the 16S rRNA gene sequence (Fig. 5).

Fig. 5
figure 5

Neighbor-joining dendrogram for gyrB gene sequences of bacteria obtained from adult and immature stages of Lutzomyia evansi collected in the municipalities of Ovejas and Colosó (Sucre Department, Colombia), based on the Kimura 2-parameter model. Numbers in nodes represent bootstrap values. The robustness of the phylogeny was tested by bootstrap analysis using 1000 iterations. Abbreviations: FFO, fed females from Ovejas; FFC, fed females from Coloso; UFO, unfed females from Ovejas; UFC, unfed females from Coloso; MC, males from Coloso; MV, males from Ovejas; L4, larvae from Ovejas

PCR-TTGE and phylogenetic analysis of L. evansi intestinal bacteria

TTGE band profiles and analysis of partial 16S rDNA sequences indicative of different genera associated to adult and immature stages of Lu. evansi were compared with 16S rDNA sequence information for known bacteria retrieved from the GenBank database. The banding patterns were similar among fed females in both geographical locations, but indicated significant variations between the bacterial communities in unfed females and males (Fig. 6). The predominant TTGE bands for L. evansi intestinal bacteria indicated similarity values between 99 and 100 %, corresponding to E. cloacae and B. thuringiensis (Table 4), in adult and immature specimens (larvae and pupae). An uncultured bacterium sequence was detected only in samples from the Colosó region (Table 4). All the TTGE sequences obtained in this study have been registered in GenBank (Table 4). In the case of the larvae, we found a different banding pattern than that expressed in adults and pupae (Fig. 6). This pattern was related to the Burkholderia cenocepacia and Bacillus gibsonii bands. Partial 16S rDNA sequences of the bands were identified through comparison with the sequences available in the Genbank database and RDP II, with high similarity percentages found (97–100 %) (Table 4). The NJ topology generated comprised clusters with suitable bootstrap support (98–100 %), indicating that the phylogenetic affiliations were consistent (Fig. 7).

Fig. 6
figure 6

Dice-UPGMA clustering analysis of bacterial diversity obtained of PCR-TTGE Profiles of 16S rDNA amplification products (V3–V6 variable regions), through GelCompar II software. Abbreviations: L, larvae; MO, males from Ovejas; MC, males from Coloso; FFO, fed females from Ovejas; FFC, fed females from Coloso; UFO, unfed females from Ovejas; UFC, unfed females from Coloso. Numbers at the nodes represent the percent similarity between the profiles of banding in the samples analyzed, based on migration with respect to temperature melting of ribosomal amplicons, as well the intensity and thickness of the band that symbolizes the dominance of the microbial community

Table 4 Closest phylogenetic classification of bacteria found in the digestive tracts of adult and immature Lutzomyia evansi by TTGE analysis, according to their similarity with 16S rRNA gene sequences recorded in the GenBank and RDP II databases
Fig. 7
figure 7

Neighbor-joining dendrogram of 16S rDNA fragment sequences (~450 bp) obtained from the PCR-TGGE profiles, based on the Kimura 2-parameter model. Numbers in nodes represent bootstrap values. The robustness of the phylogeny was tested by bootstrap analysis using 1000 iterations. FFO, fed females from Ovejas; FFC, fed females from Coloso; UFO, unfed females from Ovejas; UFC, unfed females from Coloso; MC, males from Coloso; MV, males from Ovejas; LO, larvae from Ovejas; PO, pupae from Ovejas

Bacterial diversity of Lu. evansi guts associated with blood intake and developmental stage

In order to statistically analyze the TTGE bacterial profiles relative to the developmental and nutritional status of the insects, banding patterns were analyzed with Gel Compare II software. These revealed the formation of four groups of clearly differentiated bacterial communities (Fig. 6). Nevertheless, statistically significant differences were found between the banding patterns associated with the bacterial communities present in the guts of unfed females from different locations (one-way ANOSIM test: R = 0.8674, P = 0.0001); fed females from Colosó and unfed females from Ovejas (one-way ANOSIM test: R = 0.7651; P = 0.0001); and males from both locations (one-way ANOSIM test R = 0.8821, P = 0.0001). In immature specimens, there were no statistically significant variations among guts of larvae, but there were variations between the larvae and adult stages (one-way ANOSIM test: R = 0.8345, P = 0.001). This indicates that bacterial communities obtained by independent methods of cultivation, present variation depending on the origin and stages of Lu. evansi. A simple correspondence analysis explaining 71.47 % of the variation in the dataset and integrating the results obtained by both approaches (TGGE of the total fraction and the cultivable fraction), revealed various bacterial consortia specifically associated with the guts of larvae, pupae and unfed females (Colosó population) (Fig. 6). In contrast, the groups of males, fed females and unfed females (Ovejas population) analyzed, shared a bacterial species consortium, indicating greater proximity between the microbiota of males and fed females (Fig. 8). This suggests the complementary contribution of both approaches to the study of communities of bacteria.

Fig. 8
figure 8

Simple correspondence analysis of the intestinal bacterial flora identified with partial 16S rDNA sequences of isolates and fragment sequences (~450 bp) obtained from the PCR-TGGE profiles. Abbreviations: UFO, unfed females from Ovejas; UFC, unfed females from Coloso; FFO, fed females from Ovejas; FFC, fed females from Coloso; MC, males from Coloso; MO, males from Ovejas; PP, pupae; L4, larvae. The XLSTAT 3.04 ( program was used to generate a simple correspondence analysis. The dots symbolize the location of the group of bacteria (consortia) associated with intestines of Lu. evansi, using ellipses of black color

Bacterial diversity of Lu. evansi guts associated with the geographical origin of Lu. evansi

In general, the natural populations of Lu. evansi had the following bacterial genera in common: Pseudomonas, Acinetobacter, Enterobacter, Staphylococcus and Streptomyces. However, some differences are described according to the geographical provenance. In the jungle environment, the microbiota associated with the gut of Lutzomyia evansi, contained bacterial genera (Brevibacterium and Pantoea) that were not present in the urban environment. Other bacterial genera such as Lysobacter, Ochrobactrum, Shinella, Paenibacillus, Bacillus, Microbacterium and Rummeliibacillus, could be distinguished associated with substrates extracted from potential breeding sites (base of Cordia dentata shrubs) and from the urban environment of Ovejas. In conclusion, there is a greater number of bacterial communities associated with urban populations of Lu. evansi.

Detection of Wolbachia and Leishmania in guts of female Lu. evansi

Given that Wolbachia is very important in the identification of potential strains with the ability to interrupt the life-cycle of Lu. evansi or to block the development of parasites or viruses, and Leishmania is the most relevant parasite to control in Lu evansi, their presence in our insect samples was examined by PCR. Wolbachia and Leishmania were not detected or found to be amplified in the intestinal groups formed by female Lu. evansi in our conditions.


To our knowledge, this study represents the first report of bacteria associated with the guts of natural populations of adult and immature Lu. evansi present in both types of environment (urban and jungle biotypes) on the Caribbean coast of Colombia, based on a polyphasic approach including culture-dependent and culture-independent methods. The analyses permitted identification of bacterial species, with a percentage of microorganisms or cultivable phylotypes (3–5 %) obtained by the culture-dependent method under aerobic conditions [1720]. For this reason, and to increase the range of bacteria, we examined the microbial diversity associated with Lu. evansi guts in various development stages, using the PCR-TTGE technique and DNA sequence analysis [1720, 26, 5153]. We found significant variations in the bacterial communities associated with Lu. evansi guts, depending on the developmental stage (adult and immature stages), the food source (female blood intake) and location.

Many studies on intestinal microbiota do not include immature stages of sand flies because of the difficulties in finding at the natural breeding sites, the complexity of transporting them alive to the laboratory to obtain the guts, and the lack of taxonomic keys to identify them to species level [21, 54]. In this sense, it is necessary to highlight the usefulness of the COI gene barcode sequences, which enabled the taxonomic association of immature and adult specimens of Lu. evansi (data not shown) in a manner that was consistent with other studies in terms of intraspecific divergence percentages (0.002 to 0.031) and neighbor-joining cluster supports (98–100 %) [55, 56].

This approach provided information on unregistered bacterial communities belonging exclusively to these stages [21, 22, 53, 54]. The majority of bacterial species in immature specimens were Gram-positive and were associated with the soil, which is directly related to their habitat. However, most of these were transient bacteria and none were detected in the adult stages, presumably as a result of intestinal resorption [5759]. Bacterial load obtained by dependent cultivation methods was similar to that reported for adults states of sand flies, while the bacterial load of larvae was significantly higher and this can be justified by the wide range of bacteria that can exist in the soil or natural breeding sites [18, 20]. This indicates that the platform used to access the cultivable intestinal microbiota is adequate and reproducible, because of the high number of isolates that were identified later.

A wide representation of Gram-negative bacteria was detected in females, males and larvae belonging to the genera Pseudomonas, Acinetobacter, Enterobacter and Ochrobactrum, while in the pupal stages all bacteria were Gram-positive and included the genera Microbacterium, Streptomyces, Bacillus, Lysobacter and Rummeliibacillus. Studies of gut bacteria from wild and laboratory populations of Lu. longipalpis, Phlebotomus and different vector insects have shown a larger proportion of Gram-negative bacteria [19, 6062]. The dominance of Gram-negative bacteria may be associated with partial or complete inhibitory activity on the development of parasites (Plasmodium falciparum, Leishmania infantum chagasi) and the multiplication of viruses (Dengue) in the guts of some mosquito species [31].

Fewer Gram-positive bacteria were found in the gut of Lu. evansi. The mechanisms of action of the toxins produced by Gram-positive bacteria associated with insect guts are related to alkaline pH, and the responses from the insects’ immune systems are varied [15, 6062]. These bacteria are characterized by their ability to adapt to different environmental niches and insect intestinal systems [17], which could explain their association with immature stages (Table 3).

We suggest that differences in the microbiota associated with the gut of Lu. evansi may be the result of the fauna and flora in the two collection sites, as also of the influence of soil composition in Ovejas [17, 20, 57]. The anthropophilic habit, the need to ingest blood (blood-sucking habit) and get various sources of carbohydrates (phytophage habit), allows Lu. evansi to obtain a variety of bacteria from the skin of humans and animals as well as from plants. In unfed Lu. evansi females from both locations, the main species identified were A. calcoaceticus, P. putida and E. cloacae. Acinetobacter calcoaceticus is considered a human pathogen [63, 64], and although the presence of this specific species has not been previously reported in sand flies, studies on mosquitoes (Anopheles, Aedes and Culex) and sand flies (P. argentipes and Lu. longipalpis) have detected the presence of species of the genus Acinetobacter [20, 53, 65, 66]. This suggests that the genus Acinetobacter is dominant in vector insects and is considered a “core genus” in the guts of natural Anopheles populations [65].

Pseudomona putida isolates were discovered both in the guts of unfed Lu. evansi females and in males. Similar results were found in the guts of a natural population of Lu. longipalpis collected from a visceral leishmaniasis-endemic area in Brazil (Jacobina, State of Bahia) [20]. Pseudomona putida is associated with fermented foods and plants, has good potential for genetic transformation, and is contained within the group of non-pathogenic bacteria that easily adapt to the physiological and immunological conditions present in the mosquito midgut [18]. The fact that P. putida was present in both locations and in different stages (unfed females and males) could be associated with the food source [18].

Regarding the presence of the genus Pseudomonas in Lu. evansi guts, it is necessary to highlight the presence of P. aeruginosa and P. otitidis isolates in the guts of male and larval populations collected in Colosó and Ovejas, respectively. These two species are opportunistic human pathogens that appear in immunocompromised patients, and their presence have been recorded in other insects (Table 2) [20, 6570]. Pseudomona aeruginosa have the capacity to synthesize antiparasitic molecules (prodiogiosine, cytotoxic metalloproteinase, haemolysins, antibiotics and hemagglutinin) in insect guts [16, 71].

The discovery of E. cloacae isolates in the guts of unfed Lu. evansi females is consistent with previously recorded findings in sand flies and mosquitoes (Ph. argentipes, Lu. longipalpis, Ph. papatasi, Ph. sergenti, An. albimanus and An. stephensi) [31, 53, 72]. Gouveia et al. [20] suggested that E. cloacae does not influence parasite establishment in sand fly guts in leishmaniasis areas. However, other studies have shown that Enterobacter spp. (Enterobacter sp. strain Eng Z) can make insects more resistant to infections from other bacteria and can partially or fully inhibit ooquinete, sporozoite and oocyst formation in Anopheles [71, 72]. Other species of Enterobacter associated with Lu. evansi guts and considered pathogenic, including E. cancerogenus, E. ludwiggii, E. aerogenes and E. hormaechei, were detected in other insect vectors (Table 2) [20, 52, 57, 65].

There was a strong correlation between Lu. evansi gut bacteria and the potential source of bacterial infection, which is associated with the behavior and/or eating habits of this species (Fig. 6). The presence of B. linens, S. epidermidis, S. sciuri and S. agnetis is associated with the guts of fed and unfed females. These bacteria are important because of their close relationship with the skin and mucosa in both humans and animals [26, 7375], which constitute the main source of blood intake for Lu. evansi females, due to their anthropophilic and/or zoophilic characteristics [5].

A wide variety of bacteria with entomopathogenic or genetic potential were isolated mainly in the guts of Lu. evansi larvae and pupae (O. anthropi, L. soli, P. illinoisensi, B. anthracis and S. zoogloeoides, among others) collected from natural breeding sites in the Ovejas municipality (Table 3). Previous studies have indicated that these bacteria are associated with plant roots, decaying plant material, straw and forage, which are components of natural microhabitats where immature sand flies develop [21]. Recent studies suggest that Ochrobactrum species reduce Leishmania establishment in the gut and negatively impact Lu. longipalpis survival [76].

The study of microbiota for paratransgenic insect generation also involves the detection and isolation of symbionts like Wolbachia, and the determination of their influence on the vector insect’s life-cycle and on parasite or virus obstruction [77]. The culture-dependent and culture-independent approaches we used failed to identify these symbionts, possibly because these intracellular species have little representation in the gut and are difficult to isolate in conventional culture media [78]. However, it is important to emphasize the detection of the symbiont Pa. ananatis, which is related to plants and soils and is classified as entomopathogenic [31].

The culture-independent approach permitted differentiation of the structure of gut bacterial communities in developmental stages of natural populations of Lu. evansi. The most important results include the discovery of variations depending on the nutritional status of females and the wide variety of bacteria associated with immature stages. This technique has only been previously evaluated for the analysis of bacterial communities in P. dubosqui and P. papatasi sand flies [26]; the findings indicated low bacterial community diversity dominated by Microbacterium in wild adult stages and Chloroflexi in laboratory-reared immature stages [26].

The presence and abundance of E. cloacae was noteworthy mainly in males and females from both populations and in pupae from Colosó. These results suggest that E. cloacae is a resident population associated with Lu. evansi, with possible transstadial transmission. Other bacteria identified through the TTGE technique include Burkholderia cenocepacia, Bacillus thuringiensis, Bacillus gibsonii and uncultured bacteria, which were not detected by the culture-dependent approach.

Burkholderia cenocepacia was found only in Lu. evansi larvae from the Ovejas locality; this bacterial species belongs to the symbiont group with high genetic diversity (strains) associated with insects. Depending on the strain, it may have adverse physiological functions (growth retardation, mortality of nymphs or immature stages and/or sterility), positive physiological functions (fitness or protection of the insect against entomopathogenic fungi), or metabolic functions (nitrogen fixation ability) [79, 80]. Members of Burkholderia are strictly associated with soil and are commonly found in plant roots, adjacent areas and other moist environments. This explains the presence of Bu. cenocepacia in Lu. evansi larvae, which develop in this type of substrate [79].

The soil bacteria B. thuringiensis and B. gibsonii are of great importance in the biocontrol of agricultural pests and biotechnological applications [81], and can greatly affect the intestinal bacterial community of the host insect [81, 82]. These two species have not been reported among bacterial flora associated with sand fly guts.

Studies of gut microbiota in sand flies have mainly used the 16S molecular marker via direct sequencing of amplicons from isolates differentiated by morphology, cloned amplicons from total DNA, TTGE/DGGE techniques, and next-generation sequencing strategies [17, 19, 26, 31, 57]. However, the occurrence of multiple copies of the 16S gene in some bacteria and the sequence heterogeneity between these copies may complicate ecological interpretation by causing confusion about the taxonomic identity of closely related species, as in the genera Enterobacter and Pseudomonas [83]. Furthermore, it has been shown that 16S gene heterogeneity is typical in bacteria isolated from natural environments [84].

The gyrB gene, on the other hand, exists as a single copy in bacterial genomes thus allowing a more accurate differentiation between species and/or populations, with some taxonomic conflicts reported in genera such as Pseudomonas, Bacillus and Enterobacteriaceae, lactic acid bacteria and mycobacteria [85]. However, the results were not as good for other species. Low bootstrap values (89 %) were found for the cluster related to E. hormaechei and E. cloace, and no taxonomic definition was found for the isolates L4_Isolate 199 and L4_Isolate 188, which were identified by analysis of the16S rDNA gene as Shinella zoogloeoides and L. soli. This result may be attributed to the absence of gyrB sequences in GenBank for these species; however, there is some relationship with Xanthomonas campestris, a phylogenetically related species.

We indicate that due to the environmental plasticity of Lu. evansi, cultivation-based techniques have been unable to accurately capture the true diversity within microbial communities [86]. To address the lack of information for species that are recalcitrant to cultivation, the recent use of “culture-independent methods” based on DNA sequencing technology advances (next-generation sequencing-NGS) have established distinctive solid profiles of microbial communities from different samples [26]. Further studies would be needed to determine whether the observed changes in bacterial communities associated with the different developmental stages and blood sources of the insects could influence the development and transmission of the Leishmania parasite, using Illumina-based 16S rRNA gene sequencing. This could be achieved by utilizing next-generation sequencing methods for an in-depth comparative analysis of the different feeding stages and/or parasitic infection in Lu. evansi.

In this study, Leishmania was not detected. Other researchers have reported differences in the sensitivity of different molecular markers and conventional tests (PCR, RFLP, isozyme patterns, hybridization with DNA probes) for the detection, diagnosis and identification of Leishmania species; and they propose that exploring the possibility of viewing promastigotes by the dissection of digestive tracts and the implementation of more variants of PCR with genus-specific primers would be beneficial [6, 50]. However the prevalence of natural infections with Leishmania in sand flies is low and the process of simultaneous identification of Leishmania and gut microbiota can be complicated.

Finally, further studies may include the study of bacterial communities under anaerobic conditions; there is a possibility of finding a higher diversity. Oxygenation of insect guts varies from fully aerobic to anaerobic, with anaerobic conditions more common in insects that have enlarged gut compartments and robust gut communities, such as sand flies [87].


This study may serve as a starting point for understanding particular aspects of the interaction and dynamic changes in gut bacterial communities in the developmental stages of Lu. evansi from Colombia’s Caribbean coast. TTGE profiles and culture-dependent methods are efficient and complementary. These methods work together to record and increase the range of bacterial communities detected in Lu. evansi guts. The dominant bacterial communities associated with Lu. evansi belong to the genera Enterobacter, Pseudomonas, Acinetobacter and Ochrobactrum. Additionally, the large number of bacterial species reported may have biological and genetic potential for anti-tripanosomal or biological control assays in leishmaniasis vectors.



Cytochrome c oxidase subunit I gene


Temporal temperature gradient gel electrophoresis

gyrB :

Gene that encodes the subunit B of DNA gyrase, a type II DNA topoisomerase






Fed females from Ovejas


Fed females from Coloso


Unfed Females from Ovejas


Unfed Females from Coloso


Males from Coloso


Fourth-instar larvae




  1. Amora S, Bevilaqua C, Feijo F, Alves N, Maciel M. Control de phlebotomine (Diptera: Psychodidae) leishmaniasis vectors. Neotrop Entomol. 2009;38(3):303–10.

    Article  CAS  PubMed  Google Scholar 

  2. Gomes T, Collin N, Reynoso D, Jochim R, Oliveira F, Seitz A, et al. Discovery of markers of exposure specific to bites of Lutzomyia longipalpis, the vector of Leishmania infantum chagasi in Latin America. PLoS Negl Trop Dis. 2010;4(3):e638. doi:10.1371/journal.pntd.0000638.

    Article  PubMed  PubMed Central  Google Scholar 

  3. Brazil R. The dispersion of Lutzomyia longipalpis in urban areas. Rev Soc Bras Med Trop. 2013;46(3):263–4.

    Article  PubMed  Google Scholar 

  4. Acevedo M, Arrivillaga J. Ecoepidemiology of phlebovirus (Bunyaviridae, Phlebovirus) transmitted by phlebotomine (Psychodidae, Phlebotominae). Bol Dir Malariol Sanea Amb. 2008;48:2–16.

  5. Haouas N, Pesson B, Boudabous R, Dedet JP, Babba H, Ravel C. Development of a molecular tool for the identification of Leishmania reservoir hosts by blood meal analysis in the insect vectors. Am J Trop Med Hyg. 2007;77:1054–9.

    CAS  PubMed  Google Scholar 

  6. Cochero S, Anaya Y, Díaz-Olmos Y, Paternina-Gómez M, Luna A, Paternina L, et al. Infección natural de Lutzomyia cayennensis cayennensis con parásitos tripanosomatídeos (Kinetoplastida: Trypanosomatidae) en Los Montes de María, Colombia. Rev Cubana Med Trop. 2007;59(1):35–9.

    PubMed  Google Scholar 

  7. Ferro C, Morales A. Flebótomos de Colombia: Estudios realizados por el laboratorio de Entomología 1965-1997. In: Toro G, Hernández CA, Raad J, (Eds). Instituto Nacional de Salud 1917-1997 Una historia, un compromiso; 1998. p. 219-33.

  8. Tang Y, Ward R. Sugar feeding and fluid destination control in the phlebotomine sandfly Lutzomyia longipalpis (Diptera: Psychodidae). Med Vet Entomol. 1998;12(1):13–9.

    Article  CAS  PubMed  Google Scholar 

  9. Minard G, Mavingui P, Valiente C, Minard et al. Diversity and function of bacterial microbiota in the mosquito holobiont. Parasit Vectors. 2013;6:146.

  10. Janson E, Stireman J, Singer M, Abbot P. Phytophagous insect-microbe mutualisms and adaptive evolutionary diversification. The society for the study of evolution. Evolution. 2007;62(5):997–1012.

    Article  Google Scholar 

  11. Raffa K, Adams A, Broderick N, Boone C, Cardoza Y, Delalibera I, et al. Symbionts of invasive insects: characterization, ecological roles, and relation to invasive potential and management strategies. In: USDA. Research Forum on Invasive Species. Madison: Department of Entomology, University of Wisconsin-Madison; 2008. p. 61–2.

    Google Scholar 

  12. De Gaio A, Gusmão D, Santos A, Berbert M, Pimenta P, Lemos F. Contribution of midgut bacteria to blood digestion and egg production in Aedes aegypti (Diptera: Culicidae) (L.). Parasite Vectors. 2011;4:105.

    Article  Google Scholar 

  13. Shanchez-Contreras M, Vlisidou I. The diversity of insect-bacteria interactions and its applications for disease control. Biotechnol Genet Eng. 2008;25:203–44.

    Article  Google Scholar 

  14. Volf P, Kiewegová A, Nemec A. Bacterial colonisation in the gut of Phlebotomus dubosqi (Diptera: Psychodidae): transtadial passage and the role of female diet. Folia Parasitol. 2002;49(1):73–7.

    Article  PubMed  Google Scholar 

  15. Eleftherianos I, Atri J, Accetta J, Castillo J. Endosymbiotic bacteria in insects: guardians of the immune system? Front Physiol. 2013;15:4–46. doi:10.3389/fphys.2013.00046.

    Google Scholar 

  16. Azambuja P, Garcia E, Ratcliffe N. Gut microbiota and parasite transmission by insect vectors. Trends Parasitol. 2005;21(12):568–72.

    Article  PubMed  Google Scholar 

  17. Sant’anna M, Darby A, Brazil R, Montoya J, Dillon V, et al. Investigation of the bacterial communities associated with females of Lutzomyia sand fly species from South America. PLoS One. 2012;7(8):e42531. doi:10.1371/journal.pone.0042531.

    Article  PubMed  PubMed Central  Google Scholar 

  18. Oliveira S, Moraes B, Goncalves C, Giordano C, D’Almeida J, et al. Prevalence of microbiota in the digestive tract of wild females of Lutzomyia longipalpis (Diptera: Psychodidae). Rev Soc Bras Med Trop. 2000;33:319–22.

    CAS  PubMed  Google Scholar 

  19. Mccarthy C, Diambra L, Rivera R. Metagenomic analysis of taxa associated with Lutzomyia longipalpis, vector of visceral leishmaniasis, using an unbiased high-throughput approach. PLoS Negl Trop Dis. 2011;5(9):e1304. doi:10.1371/journal.pntd.0001304.

    Article  PubMed  PubMed Central  Google Scholar 

  20. Gouveia C, AsensI M, Zahner V, Rangel E, Oliveira S. Study on the bacterial midgut microbiota associated to different Brazilian populations of Lutzomyia longipalpis (Lutz & Neiva) (Diptera: Psychodidae). Neotrop Entomol. 2008;37:597–601.

    Article  PubMed  Google Scholar 

  21. Vivero R, Torres-Gutierrez C, Bejarano E, Cadena H, Estrada L, Florez F, et al. Study on natural breeding sites of sand flies (Diptera: Phlebotominae) in areas of Leishmania transmission in Colombia. Parasit Vectors. 2015;8(116). doi: 10.1186/s13071-015-0711-y.

  22. Vivero R, Ortega E, Aparicio Y, Torres C, Muskus C, Bejarano E. Flebotominos adultos e inmaduros (Diptera: Psychodidae): registros para el Caribe colombiano. Malariol Salud Amb. 2013;3(2):1–10.

    Google Scholar 

  23. Travi B, Adler G, Lozano M, Cadena H, Montoya-Lerma J. Impact of habitat degradation on Phlebotominae (Diptera: Psychodidae) of tropical dry forests in Northern Colombia. J Med Entomol. 2002;39:451–6.

    Article  PubMed  Google Scholar 

  24. González C, Cabrera O, Munstermann L, Ferro C. Distribución de los vectores de Leishmania infantum (Kinetoplastida: Trypanosomatidae) en Colombia. Biomedica. 2006;26(1):64–72.

    Article  PubMed  Google Scholar 

  25. Vargas A, Vivero R, Uribe S, Moreno C, Cadavid G. Interacción de microbiotas bacterianas e insectos hospederos. MEFLG. 2012;4(3):13–22. ISSN 2027–4378.

    Google Scholar 

  26. Guernaoui S, Garcia D, Gazanion E, Boumezzough A, Pesson B, Fontenille D, et al. Bacterial flora as indicated by PCR-temperature gradient gel electrophoresis (TTGE) of 16S rDNA gene fragments from isolated guts of phlebotomine sand flies (Diptera: Psychodidae). J Vector Ecol. 2011;36(1):S144–7.

    Article  PubMed  Google Scholar 

  27. Hernández J, Hurtado A, Ortiz R, Walschburger T. Unidades biogeográficas de Colombia. In: Halffter G, editor. La Diversidad Biológica de Iberoamérica I. CYTED-B Programa iberoamericano de ciencia y tecnología para el desarrollo. Xalapa: Instituto de Ecología; 1992. p. 105–52.

    Google Scholar 

  28. Young G, Duncan M. Guide to identification and geographic distribution of Lutzomyia sand flies in Mexico, the West Indies, Central and South America (Diptera: Psychodidae). Mem Am Entomol Inst. 1994;54:1–881.

    Google Scholar 

  29. Golczer G, Arrivillaga J. Modificación de un protocolo estándar de extracción de ADN para flebotominos pequeños (Phlebotominae: Lutzomyia). Rev Colomb Entomol. 2008;34:199–202.

    CAS  Google Scholar 

  30. Hebert P, Cywinska A, Ball S, Dewaard J. Biological identifications through DNA barcodes. Proc R Soc Lond. 2003;270:313–21.

    Article  CAS  Google Scholar 

  31. Akhoundi M, Bakhtiari R, Guillard T, Baghaei A, Tolouei R, Sereno D, et al. Diversity of the bacterial and fungal microflora from the midgut and cuticle of phlebotomine sand flies collected in North-Western Iran. PLoS One. 2012;7(11):e50259.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Jansen P. Identifying the main soil bacterial taxa in libraries of 16S rRNA and 16 rRNA genes. Appli Environ Microbiol. 2006;72(3):1719–23.

    Article  Google Scholar 

  33. Espejo R, Feijo C, Romero J, Vásquez M. PAGE analysis of the heteroduplexes formed between PCR-amplified 16S rRNA genes: estimation of sequence similarity and rADN complexity. Microbiology. 1998;144:1611–7.

    Article  CAS  PubMed  Google Scholar 

  34. Neil L. Mathemathical model for studing genetic variation in terms of restriction endonuclease. Proc Natl Acad Sci USA. 1979;76:5269–73.

    Article  Google Scholar 

  35. Mohammadi A, Prasanna B. Analysis of genetic diversity in crop plants. Salient statistical tools and considerations. Crop Sci. 2003;43:1235–48.

    Article  Google Scholar 

  36. Moreno C, Moy F, Daniel T, Godfrey H, Cabello F. Molecular analysis of microbial communities identified in different developmental stages of Ixodes scapularis ticks from Westchester and Dutchess Counties, New York. Enviro Microbiol. 2006;8(5):761–72.

    Article  CAS  Google Scholar 

  37. Wang L, Lee F, Tai C, Kasai H. Comparison of gyrB gene sequences, 16S rRNA gene sequences and DNA-DNA hybridization in the Bacillus subtilis group. Int J Syst Evol Microbiol. 2007;57:1846–50.

    Article  CAS  PubMed  Google Scholar 

  38. Yamamoto S, Harayama S. PCR amplification and direct sequencing of gyrB genes with universal primers and their application to the detection and taxonomic analysis of Pseudomonas putida strains. Appli Environ Microbiol. 1995;61:1104–9.

    CAS  Google Scholar 

  39. Hall T. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser. 1999;41:95–8.

    CAS  Google Scholar 

  40. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. MEGA 5: Molecular Evolutionary Genetics Analysis using Maximum Likelihood, Evolutionary Distance, and Maximum Parsimony Methods. Mol Biol Evol. 2011;28:2731–9. doi:10.1093/molbev/msr121.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–25. doi:10.1016/j.ympev.2008.01.019. 2011.

    CAS  PubMed  Google Scholar 

  42. Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;15:111–20.

    Article  Google Scholar 

  43. Martin D, Williamson C, Posada D. 2RDP2: Recombination detection and analysis from sequence alignments. Bioinformatics. 2005;21(2):260–2. doi:10.1093/bioinformatics/bth490.

    Article  CAS  PubMed  Google Scholar 

  44. Muyzer G, Waal E, Uitterlinden G. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol. 1993;59:695–700.

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Gomez A, Yannarell A, Sims G, Cadavid-Restrepo G, Moreno C. Characterization of bacterial diversity at different depths in the Moravia Hill landfill site at Medellín, Colombia. Soil Biol Biochem. 2011;43(6):1275–84. doi:10.1016/j.soilbio.2011.02.018.

    Article  CAS  Google Scholar 

  46. Koeppel A, Perry E, Sikorski J, Krizanc D, Warner A, Ward D, et al. Identifying the fundamental units of bacterial diversity: a paradigm shift to incorporate ecology into bacterial systematics. Proc Natl Acad Sci USA. 2008;105:2504e2509.

    Article  Google Scholar 

  47. Rademaker J, De Bruijin F. Characterization and classification of microbes by rep-PCR genomic fingerprinting and computer assisted pattern analysis. In: Caetano-Anollés G, Gresshoff PM, editors. DNA Markers: Protocols, Aplications and Overviews. New York: J. Wiley & Sons; 1997. pp. 151–171.

  48. Braig H, Zhou W, Dobson S, O’Neill S. Cloning and characterization of a gene encoding the major Surface protein of the bacterial endosymbiont Wolbachia. J Bacteriol. 1998;180(9):2373–8.

    CAS  PubMed  PubMed Central  Google Scholar 

  49. Zhou W, Rousset F, O’Neill S. Phylogeny and PCR-based classication of Wolbachia strains using WSP gene sequences. Proc R Soc Lond B. 1998;265:509–15.

    Article  CAS  Google Scholar 

  50. Fraga J, Montalvo M, De Doncker S, Dujardin J, Van der Auwera G. Phylogeny of Leishmania species based on the heatshock protein 70 gene. Infect Genet Evol. 2012;10(2):238–45.

    Article  Google Scholar 

  51. Hammer O, Harper D, Ryan P. PAST: Paleontological statistics software package for education and data analysis. Palaeontologia Electronica. 2001; 4(1): 9 pp.

  52. Dillon R, Dillon V. The gut bacteria of insects: nonpathogenic interactions. Annu Rev Entomol. 2004;49:71–92.

    Article  CAS  PubMed  Google Scholar 

  53. Hillesland H, Read A, Subhadra B, Hurwitz I, McKelvey R, et al. Identification of aerobic gut bacteria from the Kala Azar vector, Phlebotomus argentipes: A platform for potential paratransgenic manipulation of sand flies. Am J Trop Med Hyg. 2008;79:881–6.

    PubMed  Google Scholar 

  54. Maleki N, Ali M, Hajikhani S, Saeidi Z, Ahmad A, Gerami M, Hasan M, et al. Aerobic microbial community of insectary population of Phlebotomus papatasi. J Arthropod Borne Dis. 2014;8(1):69–75.

    Google Scholar 

  55. Contreras-Gutiérrez MA, Vivero RJ, Vélez ID, Porter CH, Uribe S. DNA Barcoding for the identification of sand fly species (Diptera, Psychodidae, Phlebotominae) in Colombia. PLoS One. 2014;9(1):e85496. doi:10.1371/journal.pone.0085496.

    Article  PubMed  PubMed Central  Google Scholar 

  56. Contreras-Gutiérrez MA, Vivero RJ, Uribe S. DNA barcode: A tool for identification of Lutzomyia spp. From larvae. MEFLG. 2013;5(4):7–16.

    Google Scholar 

  57. Mukhopadhyay J, Braig H, Rowton E, Ghosh K. Naturally occurring culturable aerobic gut flora of adult Phlebotomus papatasi, Vector of Leishmania major in the old world. PLoS One. 2012;7(5):e35748. org/10.1371/journal.pone.0035748.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  58. Peterkova K, Robles M, Ramalho M, Zurek L. Significance of bacteria in oviposition and larval development of the sand fly Lutzomyia longipalpis. Parasit Vectors. 2012;5:145. doi:10.1186/1756-3305-5-145.

    Article  Google Scholar 

  59. Chaniotis BN. Sugar-feeding behavior of Lutzomyia trapidoi (Diptera: Psychodidae) under experimental conditions. J Med Entomol. 1974;11:73–9.

    Article  CAS  PubMed  Google Scholar 

  60. Hurwitz I, Hillesland H, Fieck A, Das P, Durvasula R. The paratransgenic sand fly: A platform for control of Leishmania transmission. Parasit Vectors. 2011;4:82.

    Article  PubMed  PubMed Central  Google Scholar 

  61. Rani A, Sharma A, Rajagopal R, Adak T, Bhatnagar R. Bacterial diversity analysis of larvae and adult midgut microflora using culture-dependent and culture-independent methods in lab-reared and field-collected Anopheles stephensi: an Asian malarial vector. BMC Microbiol. 2009;9:96.

    Article  PubMed  PubMed Central  Google Scholar 

  62. Broderick N, Raffa K, Handelsman J. Midgut bacteria required for Bacillus thuringiensis insecticidal activity. PNAS. 2006;103(41).

  63. Dillion J, Kordy E, Lane R. The prevalence of a microbiota in the digestive tract of Phlebotomus papatasi. Ann Trop Med Parasitol. 1996;90:669–73.

    Google Scholar 

  64. Doughari H, Ndakidemi P, Human I, Benade S. The ecology, biology and pathogenesis of Acinetobacter spp.: An overview. Microbes Environ. 2011;26:101–12.

    Article  PubMed  Google Scholar 

  65. Manguin S, Ngo C, Tainchum K, Juntarajumnong W, Chareonviriyaphap T, Michon A, et al. Bacterial biodiversity in midguts of Anopheles mosquitoes, malaria vectors in southeast Asia. In: Manguin S, editor. Anopheles mosquitoes - New insights into malaria vectors. Croatia: INTECH open science; 2013. p. 549–76. ISBN 978-953-51-1188-7.

    Chapter  Google Scholar 

  66. Pidiyar V, Jangid K, Patole M, Shouche Y. Studies on cultured and uncultured microbiota of wild Culex quinquefasciatus mosquito midgut based on 16 s ribosomal RNA gene analysis. Am J Trop Med Hyg. 2004;70(6):597–603.

    CAS  PubMed  Google Scholar 

  67. Malhotra J, Dua A, Saxena A, Sangwan N, Mukherjee U, Pandey N, et al. Genome sequence of Acinetobacter sp. strain HA, isolated from the gut of the polyphagous insect pest Helicoverpa armigera. J Bacteriol. 2012;194(18):5156. doi:10.1128/JB.01194-12.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Chavshin A, Oshaghi M, Vatandoost H, Pourmand M, Raeisi A, Terenius O. Isolation and identification of culturable bacteria from wild Anopheles culicifacies, a first step in a paratransgenesis approach. Parasit Vectors. 2014;7:419. doi:10.1186/1756-3305-7-419.

    Article  PubMed  PubMed Central  Google Scholar 

  69. Oliveira S, Morais B, Gonçalves C, Giordano-Dias C, Vilela M, Brazil R, et al. Microbiota do trato digestivo de fêmeas de Lutzomyia longipalpis (Lutz & Neiva, 1912) (Diptera: Psychodidae) provenientes de colônia alimentadas com sangue e com sangue e sacarose. Cad Saúde Pública Rio de Janeiro. 2001;17(1):229–32.

    Article  Google Scholar 

  70. Demaio J, Pumpuni C, Kent M, Beier J. The migut bacterial flora of wild Aedes triseriatus, Culex pipiens and Psorophora columbiae mosquitoes. Am J Trop Med Hyg. 1996;54:219.

    CAS  PubMed  Google Scholar 

  71. Dong Y, Manfredini F, Dimopoulos G. Implication of the mosquito midgut microbiota in the defense against malaria parasites. PLoS Pathog. 2009;5:e1000423.

    Article  PubMed  PubMed Central  Google Scholar 

  72. Husseneder C, Kenneth J. Genetically engineered termite gut bacteria (Enterobacter cloacae) deliver and spread foreign genes in termite colonies. Appl Microbiol Biotechnol. 2005;68:360–7.

    Article  CAS  PubMed  Google Scholar 

  73. Chakraborty U, Chakraborty B, Basnet M. Plant growth promotion and induction of resistance in Camellia sinensis by Bacillus megaterium. J Basic Microbiol. 2006;46:186–95.

    Article  CAS  PubMed  Google Scholar 

  74. Braks MA, Anderson RA, Knols BG. Infochemicals in mosquito host selection: human skin microflora and Plasmodium parasites. Parasitol Today. 1999;15:409–13.

    Article  CAS  PubMed  Google Scholar 

  75. Meijerink J, van Loon J. Sensitivities of antennal olfactory neurons of the malaria mosquito, Anopheles gambiae, to carboxylic acids. J Insect Physiol. 1999;45:365–73.

    Article  CAS  PubMed  Google Scholar 

  76. Sant’Anna M, Diaz H, Aguiar H, Waleed S, Salem A, Cavalcante R, et al. Colonisation resistance in the sand fly gut: Leishmania protects Lutzomyia longipalpis from bacterial infection. Parasit Vectors. 2014;7:329.

    Article  PubMed  PubMed Central  Google Scholar 

  77. Ngwa C, Ckner V, Abdelmohsen R, Scheuermayer M, Fischer R, Hentschel U, et al. 16S rRNA gene-based identification of Elizabethkingia meningoseptica (Flavobacteriales: Flavobacteriaceae) as a dominant midgut bacterium of the Asian malaria vector Anopheles stephensi (Dipteria: Culicidae) with antimicrobial activities. J Med Entomol. 2013;50(2):404–14.

    Article  CAS  PubMed  Google Scholar 

  78. Floate K, Kyei-Poku G, Coghlin P. Overview and relevance of Wolbachia bacteria in biocontrol research. Biocontrol Sci Technol. 2006;16(8):767–88.

    Article  Google Scholar 

  79. Kikuchi Y, Meng X, Fukatsu T. Gut symbiotic bacteria of the genus Burkholderia in the broad-headed bugs Riptortus clavatus and Leptocorisa chinensis (Heteroptera: Alydidae). Appl Environ Microbiol. 2005;71(7):4035–43. doi:10.1128/AEM.71.7.40354043.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  80. Estrada S, Bustillos P, Caballero J. Burkholderia, a genus rich in plant-associated nitrogen fixers with wide environmental and geographic distribution. Appl Environ Microbiol. 2001;67:2790–8.

    Article  Google Scholar 

  81. Sezen K, Isci S, Muratoglu H, Inan K, Demirbag Z. Identification and pathogenicity of bacteria from Gryllotalpa gryllotalpa L. (Orthoptera: Gryllotalpidae). Türk Biyo Müc Derg. 2013;4(2):89–108.

    Google Scholar 

  82. Tchioffo M, Bachar D, Abate L, Marie A, Nsango S, Shahbazkia H, et al. Midgut microbiota of the malaria mosquito vector Anopheles gambiae and interactions with Plasmodium falciparum infection. PLoS Pathog. 2012;8(5). doi: 10.1371/journal.ppat.1002742.

  83. Peixoto H, Coutinho D, Rumjanek N, Macrae A, Rosado A. Use of rpoB and 16S rRNA genes to analyse bacterial diversity of a tropical soil using PCR and DGGE R.S. Lett Appl Microbiol. 2002;35:316–20.

    Article  CAS  PubMed  Google Scholar 

  84. Dahllo F, Baillie H, Kjelleberg S. RpoB-based microbial community analysis avoids limitations inherent in 16S rRNA gene intraspecies heterogeneity. Appl Environ Microbiol. 2000;66:3376–80.

    Article  Google Scholar 

  85. Boua B, Fernández A, García C, Sáez-Nieto S, Valdezated S. Métodos de identificación bacteriana en el laboratorio de microbiología. Enferm Infecc Microbiol Clin. 2011;29(8):601–8.

    Article  Google Scholar 

  86. Degnan P, Ochman H. Illumina-based analysis of microbial community diversity. ISME J. 2012;6(1):183–94. doi:10.1038/ismej.2011.74.

    Article  CAS  PubMed  Google Scholar 

  87. Engel P, Moran N. The gut microbiota of insects - diversity in structure and function. FEMS Microbiol Rev. 2013;37:699–735.

    Article  CAS  PubMed  Google Scholar 

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The authors acknowledge the support that members (Horacio Cadena Peña and Andres Velez Mira) of the Programa de Estudio y Control de Enfermedades Tropicales (PECET) of the University of Antioquia and the Biomédicas Research Group of the University of Sucre in Sincelejo, Colombia provided for the entomological survey. We would like to thank the different communities we visited during our surveys in the municipalities of Ovejas and Colosó (department of Sucre) for giving us access to their facilities, providing hospitality and collaborating with fieldwork. We also acknowledge the support that Dr. Eduar E. Bejarano, Luis Gregorio Estrada and Edgar D. Ortega (Biomedical Research Group, University of Sucre) provided for the survey of insects in the two locations in the department of Sucre.


This study was funded by the Administrative Department of Science, Technology and Innovation (COLCIENCIAS code # 695-2014; Administrative Department of Science, Technology and Innovation; National Call 528 for doctoral studies in Colombia, 2011); Grupo de Microbiodiversidad y Bioprospección and Grupo de Investigación en Sistematica Molecular, Universidad Nacional de Colombia at Medellín. The funders had no role in the study design, data collection and analysis, decision to publish or preparation of the manuscript.

Availability of data and materials

The datasets supporting the conclusions of this article are included within the article. Sequences generated in the present study are submitted in the GenBank database under accession numbers: KU134739–KU134799.

Authors’ contributions

RJV: Designed the study, performed the experiments and field work, analyzed the data and contributed to writing the manuscript. CXMH, GECR and SUS: Designed the study, analyzed the data and contributed to writing the manuscript. NGR: Performed the experiments and analyzed the data. All authors read and approved the final version of the manuscript.

Competing interests

The authors declare that they have no competing interests.

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Not applicable.

Ethics approval and consent to participate

Sand fly collection was performed in accordance with the parameters of Colombian decree number 1376, which regulates specimen collection of biologically diverse wild species for non-commercial research. No specific permits were required for this study. Sand flies were collected on private property and permission was received from landowners prior to sampling.

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Correspondence to Rafael José Vivero.

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Vivero, R.J., Jaramillo, N.G., Cadavid-Restrepo, G. et al. Structural differences in gut bacteria communities in developmental stages of natural populations of Lutzomyia evansi from Colombia's Caribbean coast. Parasites Vectors 9, 496 (2016).

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