Cutaneous leishmaniasis is a neglected, vector-borne parasitic disease and is responsible for persistent, often disfiguring lesions and other associated complications. Leishmania, causing zoonotic cutaneous leishmaniasis (ZCL) in the Old World are mainly transmitted by the predominant sand fly vector, Phlebotomus papatasi. To date, there is no efficient control measure or vaccine available for this widespread insect-borne infectious disease.
A survey was carried out to study the abundance of different natural gut flora in P. papatasi, with the long-term goal of generating a paratransgenic sand fly that can potentially block the development of Leishmania in the sand fly gut, thereby preventing transmission of leishmania in endemic disease foci. Sand flies, in particular, P. papatasi were captured from different habitats of various parts of the world. Gut microbes were cultured and identified using 16S ribosomal DNA analysis and a phylogenetic tree was constructed. We found variation in the species and abundance of gut flora in flies collected from different habitats. However, a few Gram-positive, nonpathogenic bacteria including Bacillus flexus and B. pumilus were common in most of the sites examined.
Our results indicate that there is a wide range of variation of aerobic gut flora inhabiting sand fly guts, which possibly reflect the ecological condition of the habitat where the fly breeds. Also, some species of bacteria (B. pumilus, and B. flexus) were found from most of the habitats. Important from an applied perspective of dissemination, our results support a link between oviposition induction and adult gut flora.
Citation: Mukhopadhyay J, Braig HR, Rowton ED, Ghosh K (2012) Naturally Occurring Culturable Aerobic Gut Flora of Adult Phlebotomus papatasi, Vector of Leishmania major in the Old World. PLoS ONE 7(5): e35748. https://doi.org/10.1371/journal.pone.0035748
Editor: Markus M. Heimesaat, Charité - Campus Benjamin Franklin, Germany
Received: October 12, 2011; Accepted: March 23, 2012; Published: May 22, 2012
Copyright: © 2012 Mukhopadhyay et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This study was funded by the Department of Defense through the In-House Laboratory Independent Research program at the Walter Reed Army Institute of Research. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Phlebotomine sand flies (Diptera: Psychodidae) are important vectors of leishmaniasis, Carrión's disease or bartonellosis, and a variety of arboviral diseases , , . Not only are novel viruses currently being discovered in sand flies, but also different reservoirs are being identified for pathogens and parasites of human diseases, transmitted by sand flies. The distribution areas of sand flies and the diseases they transmit are also expanding. New viral diseases of humans transmitted by sand flies are being reported as well , , , . From a public health viewpoint, however, their greatest importance is as vectors of leishmania , , . The genus Phlebotomus in the Old World, and Lutzomyia in the New World, include most of the important vectors of human leishmaniasis. This disease complex is widely distributed in tropical and subtropical regions of the Americas, Africa, southern Europe, and central Asia. It is estimated that some 350 million people in the world are at risk of acquiring leishmaniasis and that approximately 12 million people are currently infected , . After malaria, leishmaniasis is the second most important vector-borne parasitic disease and a leading cause of death. There are 500,000 annual new cases of visceral leishmaniasis (VL) or kala-azar in the world and about one-half of them are in India. Bihar, the most affected state in India witnesses almost 90% of the new cases of VL each year with a 10% mortality rate , .
Cutaneous leishmaniasis is more prevalent throughout the world and causes disfiguration and other associated complications. Cutaneous leishmaniasis and Zoonotic cutaneous leishmaniasis (ZCL), caused by L. tropica and L. major, respectively, are widely distributed in Turkey, Egypt, Israel, Saudi Arabia and the northern part of India, where mainly P. sergenti and P. papatasi have been incriminated as the vectors . The Afro-Asian vector of ZCL, P. papatasi is widely distributed and is the type species of the genus. The distribution of P. papatasi coincides with the distribution of ZCL in most parts of the old world and shows little population differentiation between peridomestic sites and borrows of wild rodents , . Despite the demonstrated public health importance, relatively little attempt has been undertaken to block the transmission of this disease by this insect vector.
Information on breeding sites of P. papatasi is available from several countries , . In India, immature stages of P. papatasi have been consistently recovered from cattle sheds and human dwellings in urban areas , . In rural areas, they have been found in various habitats: unused poultry houses made of bricks and clay, manure heaps, caves, embankments, dried-up cesspits and latrines . In Egypt, breeding sites of P. papatasi have been found in a similar range of ecotopes . Rabbit holes in peri-domestic areas serve as breeding sites, which reduce the indoor abundance of P. papatasi in Tunisia . In the Central Asian Republics of the former Soviet Union and neighboring states, burrows of the desert gerbil (Rhombomys opimus) are recognized as breeding sites . Caves and dense vegetation of valleys are important in the Judean desert . Dog shelters are important as breeding sites in peri-urban areas of southern Italy . The ease with which P. papatasi adapt to an urban environment can be illustrated with the collection of sand flies near the bed and in the bathroom on the second floor in a house in a big city and in another highly urbanized area in southern Italy .
Currently, insecticide application at breeding sites is the method of choice for the vector control vis-à-vis control of disease. This control effort targets adult sand flies to bring down populations in order to reduce transmission. Application of insecticide may be limited due to adverse effects on the environment, human health, and the emergence of insecticide resistance in sand flies , .
Sand flies spend a major part of their life as eggs, larvae and pupae in soil. During the immature stages, they are exposed to a variety of different soil microbes that are available for ingestion. In fact, gravid P. papatasi choose oviposition sites by presence of frass and certain soil bacteria , . Consequently, it is expected that the sand fly gut harbors a variety of microbial flora. The information on the distribution of the gut flora in feral sand fly populations, especially P. papatasi, across different habitat is still lacking. There are a few reports available on other species: from colonized P. duboscqi , , from natural population of L. longipalpis , and from feral population of P. argentipes . A very preliminary study on PCR fingerprinting of the gut flora from Moroccan P. papatasi flies identified just two bacteria , . There is also a small report on the distribution of gut flora from P. papatasi collected in Egypt , . Adler and Theodor suggested as early as 1929 that the presence of microbes in the midgut might interfere with Leishmania infection . Later, Schlein et al. saw a reduction of infection rate of L. major in P. papatasi under the influence of yeasts and bacteria . There is little doubt that the developing Leishmania in a sand fly gut is exposed to gut flora . In an attempt to develop a strategy to block the transmission of leishmania, which has been demonstrated for some other vector-borne disease pathogens , we searched for nonpathogenic gut flora that could be genetically manipulated to release an anti-leishmanial substance and then be reintroduced into the sand fly gut through larval breeding habitats. The long-term objective would be to block or partially disrupt the metacyclogenesis of Leishmania sp. by the released product of the recombinant bacterium and thereby render the sand fly incapable of transmitting the disease. This will help to prevent further epidemic outbreak of leishmaniasis. A similar approach has been successfully applied in the development of paratransgenic Rhodnius prolixus, a vector of Chagas disease in Central America, with the help of genetically transformed Rhodococcus rhodnii . A paratransgenic strategy has also been applied to Glossina morsitans, the vector of African sleeping sickness , . Additionally, a viral paratransgenic approach has been used to generate a transgenic Anopheles gambie, a vector of malaria . A paratransgenic control strategy has also been applied to the glossy-winged sharpshooter with the help of genetically marked Alcaligenes sp. , . The use of paratransgenesis is explored in the brine shrimp Artemia as a model for controlling infectious diseases in mariculture and in an increasing number of insect groups such as fleas and termites , , . In mosquitoes, symbiotic yeasts are discussed for control purposes . This is only a short step to consider other eukaryotic symbionts of arthropods , , .
Here we examine the presence and distribution of different aerobic gut microbes of P. papatasi, the major vector of ZCL, in different habitats of various geographical parts of the world.
Materials and Methods
Collection of field samples
A large number of live sand flies were collected from India, Turkey and Tunisia. Sand flies were captured mainly from human dwellings, sheep sheds, chick pens, rabbit holes and mixed dwellings using light traps, or an aspirator and a flash light. Oral informed consent was obtained from head of households for indoor aspiration of sand flies and/or property owners for shed and outdoor collections that may have included light traps operated overnight. An explanation, in the local language, of the purposes for the collection, how the specimens would be used, the collection methods and any effects the collecting might have on the residents and/or their animals was provided before consent was obtained. Consents were listed in a written log kept by the collectors. Collected sand flies were released in containers with a plaster of Paris bottom. The containers were placed in individual plastic bags with moist cotton to provide necessary humidity for transportation to the laboratory.
Laboratory colony (control)
We used a laboratory colony of P. papatasi originated from field-collected samples from North Sinai, Egypt (PPNS). The colony is maintained at WRAIR following the method of Modi and Rowton .
Preparation of the media
Both liquid and solid agar based sterile media were prepared for the gut bacterial culture. Brain Heart Infusion (BHI) agar plates were prepared following the manufacturer's protocol (BD Biosciences; Cat. # 241830) and liquid culture broth was prepared using Terrific Broth Base (Invitrogen, Cat. # 22711-022).
Isolation and preservation of bacterial flora
Field collected sand flies were identified following the description by Lewis . Only female P. papatasi was selected for the isolation of gut flora. In a sterile hood, each sand fly was rinsed in 70% ethanol for two minutes, followed by three quick rinses of sterile 1× PBS. Then the fly gut was dissected out and homogenized in about 60 µl of sterile 1× PBS in a sterile microfuge tube. Forty micro liter of the fly sample homogenate was quickly plated on BHI-agar plate, previously labeled with sand fly origin and number. The plates were subsequently placed in a 33±1°C incubator overnight. The remainder of the homogenate was cryopreserved in a −70°C freezer.
Selection and culture of clones
After overnight incubation, two to six colonies (depending on the number of colonies obtained from each fly) were picked up using a sterile toothpick and two copies of each colony were cultured in liquid media. The bacterial cultures were allowed to grow overnight in a shaker at 250 rpm at 33±1°C. One culture was used for isolation of DNA while the other was cryopreserved (using 17% sterile glycerol) in a freezer at −70°C. A relatively high incubation temperature was selected for the isolation of the flora because we are mostly interested in generating a recombinant bacterial flora that can grow well and withstand a higher temperature when spread in natural breeding places in a tropical climate.
DNA extraction, PCR amplification and identification of the bacteria
Genomic DNA was isolated from individual cultures, using DNeasy Blood & Tissue Kit (Qiagen, Cat. #69581). Two sets of primers were used for amplification of the 16S rDNA: a) 533F- 5′-GT TGC CAG CAG CCG CGG TAA-3′ and 1541R- 5′-AAG GAG GTG WTC CAR CC-3′ , ; and b) 8F-I 5′-AGA GTT TGA TYM TGG CTC AI-3′and 907R-I 5′-CCG TCA ATT CMT TTG AGT TI-3′ . PCR reactions were carried out in a 25 µl reaction mixture containing 25–50 ng of template DNA, 1× PCR buffer (with 2.5 mM MgCl2, 0.2–1 µM of each primer, 0.2 mM dNTPs) and 1 Unit of Taq DNA polymerase (Promega, Cat. #M186). The PCR machine was programmed for the following amplification protocol: one cycle at 95°C for four min; 35 cycles for: 95°C (60 sec), 52°C (60 sec) and 72°C (90 sec) and the final extension step of one cycle at 72°C for six minutes. One non-template control was used for each run. PCR products were detected by agarose gel electrophoresis and purified with QIAquick gel extraction kit (Qiagen, Cat#28704). Nucleotide sequence for each amplicon was determined by using BigDye Terminator v3.1Cycle Sequencing Kit (Applied Biosystems, Cat# 4337455), and 1 U of one of the primers used during PCR amplification. Sequences were blasted and compared with the available sequences at the GenBank database. Isolates were recognized as the same species when their 16S rDNA sequences shared ≥97% homology with complete 16S rDNA.
Data collection: After identification, the results were tabulated to show the relative abundance of different species of bacteria isolated from sand flies, collected from different locations and habitats.
The sequences were manipulated in programs SeaView version 4 and MEGA version 5.05 , . Alignment of the sequences was based on the secondary structure of their RNAs with the alignment program SINA in the ARB software package using the silva comprehensive ribosomal RNA database version 108 . The alignment was checked by hand. The best evolutionary model among 88 options for the analysis of the alignment was chosen with the help of the program jModeltest version 2 , : GTR+I+Γ. Bayesian analysis was performed with the program Mr Bayes version 3.2 . The analysis was carried out with two independent runs with four chains each for 1,000,000 generations of which the first 25% were dismissed. An average standard deviation of split frequencies of 0.0075 was reached, at which point the maximum Potential Scale Reduction Factor for parameter values was 1.002 suggesting conversion. The harmonic mean of the log likelihood of the resulting trees was – 9,612.0. The tree was drawn with the help of the program Figtree version 1.3.1.
A total of 107 P. papatasi field samples were dissected, of which 43 were collected from Tunisia, 31 originated from Turkey and 33 from India. Of the samples collected, 103 guts were cultured (two guts did not produce any colonies and two others were contaminated during preparation, Table 1). Forty-three female P. papatasi from one of our laboratory colonies originating from Egypt (PPNS) were used as control. The number of colonies generated from each fly gut varied widely. From some flies, there were as few as three colonies, while in others there were as many as 153 colonies (Figure 1). Two to six colonies from each sand fly gut were selected for further processing and identification of the flora.
The diversity of flora among P. papatasi populations, collected from several habitats in three different countries and a laboratory colony is shown in Figure 2. It is evident that there is more variation of the gut flora in the flies collected from animal dwellings of Tunisia and Turkey than in the samples captured from human dwellings in India.
In P. papatasi samples from Tunisia, Bacillus flexus was the most dominant bacterium irrespective of the collection habitat. Two other bacteria, B. pumilus and B. megaterium were also quite common. The flora from P. papatasi samples collected in Turkey was diversified with a clear dominance (31%) of B. pumilus. Other bacteria, including B. clausii, B. cereus, B. subtilis and Brevibacillus brevis were also present but at lower frequencies. Aerobic gut microbes in female P. papatasi collected from human dwellings of Patna, India, showed less diversity compared to the other two sites; the majority of them (54%) were B. pumilus. Four other species were also present in the captured samples but with much lower frequencies (Figure 2). The colonized sand flies from Egypt also showed a relative abundance of B. pumilus (30%) with few other microbes. With the exception of two species, Enterobacter aerogenes (Enterobacteriaceae, Proteobacteria) and Plantibacter flavus (Microbacteriaceae, Actinobacteria), all other bacteria belong to the families Bacillaceae and Paenibacillaceae (Figure 3).
The present study is the most comprehensive evaluation of the distribution of intestinal flora of P. papatasi to date, as it describes the abundance of the bacterial gut flora from different habitats of three different countries. Our results show that P. papatasi harbor a wide selection of gut bacteria. Roughly, half of the detected bacteria are described for the first time from sand flies and some are described for the first time from insects (Table 2). The diversity of microbes from different habitats strongly suggests that the sand fly gut–microbial association is dependent on microbes in the environment in which those sand flies breed and live.
The gut flora in sand flies collected from sheep sheds and rabbit holes in Tunisia and Turkey showed more diversity than other groups. However, no significant differences in the distribution of the microbes were observed in the gut of sand flies collected from sheep shed or rabbit holes in Tunisia. Among the predominant flora observed from the flies collected from these two habitats, B. cereus is a potential human pathogenic bacterium . The same is true for En. aerogenes, which has also been found to cause infections . An interesting case is B.circulans because in the older Russian literature, it was mentioned together with B. mycoides as an entomopathogen of the gut of larval fleas . Later, it was also recognized as a gut pathogen of mosquito larvae . More recently, B. circulans was investigated as a potential probiotic for juvenile rohu in freshwater fish aquaculture . Among the bacterial flora, B. megaterium and B. flexus are reported not only as non-pathogenic but also having some beneficial effect as probiotics , .
The sand flies in Tunisia and Turkey were collected from animal shelters including sheep sheds, rabbit holes and poultry pens. Usually, the soil in and around these areas is contaminated by the excreta of the animals and other environmental contaminants making the soil a fertile medium for the growth of coprophilic bacteria. This contamination could explain the diversity of the bacterial flora found in the sand fly gut collected from these habitats. The diversity may be accentuated in places where animal shelters are in close proximity to agricultural land where the use of biofertilizers add more microbes to the nearby animal shelters. One example of this is the presence of B. megaterium, which have been found to have a good growth enhancement effect and yield, and have been used as a biofertilizer , .
The diversity of bacterial population is somewhat restricted in the sand flies captured from India. Although other bacteria are present in less frequency, B. pumilus is the predominant bacterium found in the sand flies from Patna, India. Here, the majority of P. papatasi were obtained from human dwellings which is consistent with the anthropophilic nature of P. papatasi , . Blood-fed sand flies use the loose soil in the dark corners inside the mud houses as the most favorable place for oviposition . Larvae are only exposed to the microbes inside the mud-house but not to the excreta of animals and other environmental contaminants. This may explain the lower diversity of the gut flora isolated from sand flies captured from human dwellings.
An unexpected result is that the Egypt lab colony seems to show a higher or similar diversity of bacterial flora as samples originated from any of the natural habitats. This observation might be explained by the fact that the sand fly larvae are maintained in the laboratory on a diet composed of rabbit chow and rabbit feces, which are additional resources of gut flora and might have contributed to the bacterial diversity. Blood-fed females defecate gut bacteria along with the remains of the blood meal. Sand fly larvae are coprophagous. Therefore some gut bacteria are vertically transmitted to the next generation.
Bacillus pumilus, one of the most dominant bacteria of all the populations, is a Gram-positive, aerobic, rod-shaped, soil-dwelling bacterium. Like other Bacillus species, the spores produced by B. pumilus are more resistant than vegetative cells to heat, desiccation, UV radiation, γ-radiation, H2O2, and starvation. This species has been found in extreme environments such as the interior of Sonoran desert basalt and the Mars Odyssey spacecraft , . The presence of B. pumilus in higher numbers in sand flies collected from human dwellings might be significant from the microbiological point of view as it has been shown that B. pumilus exhibits strong antifungal and antiviral activity , . Schlein et al. postulated that some gut bacteria might help to destroy fungi, thereby indirectly helping the development of Leishmania in the sand fly gut . It is not clear at this point if B. pumilus is engaged in antifungal activity in the sand fly gut at all or if it acts together with other closely related Bacillus species or in combination with other gut factors to make the sand flies mycosis free. A fungi-free gut may help Leishmania survive which would make sand flies a more competent vector.
In the present study, a large number of Bacillus species was identified from P. papatasi. A preliminary study reported a different profile of bacteria. Species of Enterobacter and Cronobacter were isolated in greatest abundance from P. papatasi from Egypt by Dillon and others . The authors emphasized that they used a rather selective medium and culture conditions. However, in a previous study on P. argentipes from India, we found a higher abundance of Enterobacteriaceae .
For the New World sand flies, Oliveira et al. found a high percentage of Staphylococcus sp. (28%) and B. thuringiensis (18%) in Lutz. longipalpis samples collected from Lapinha cave, Brazil . They also recorded a relatively low percentage of En. cloacae (9%). We believe that these variations in the abundance of different bacteria from feral populations of sand flies are due to the ecological setting of their breeding habitat and species related.
The phylogenetic analysis shows strong support for all clades with the exception of B. badius. It is very reassuring that with the exception of B. megaterium, all species of our phylogenetic analysis using 16SRNA only showed similar relationship to a recent whole-genome phylogenetic analysis of the family Bacillaceae . Staphylococcus species often clusters in 16S phylogenies within clades of Bacillus species . Bacillus fusiformis of the literature cited here should be recognized as a Lysinibacillus species . Species that have been observed by Radjame et al. to induce oviposition behavior in gravid P. papatasi females do not form a strict clade but cluster in a bigger group among the species recovered in this study .
Since there is very little information on the symbiotic association of bacteria with sand flies, gut colonization of bacteria is believed to be dependent on the larval food and the breeding soil. The larvae acquire many soil microbes during their immature stages of development which are believed to survive during the transformation until the adult emergence as reported in P.duboscqi by Volf et al.  (unpublished observation, Ghosh) However, in nature, adult sand flies may also have the opportunity to ingest microorganism through contaminated sugar meal derived from leaves, fruits or aphid honeydew taken between blood meals. Some sand fly species, in particular P. papatasi, may ingest microorganism from the plant cuticle while sucking the plant juice . This explains some of the plant-associated bacteria found in our study.
Radjame et al. found that several soil bacteria significantly enhance the oviposition response of P. papatasi females . The most pronounced effect was observed with B. firmus (P 0.00001 in cattle sheds), followed by S. saprophyticus (0.0003 in termite mounts and 0.002 in human dwellings), and B. licheniformis (0.0007 in cattle sheds, 0.003 in termite mounts and 0.0091 in human dwellings). Importantly, B. pumilus also induced oviposition of sand flies in cattle sheds significantly . More studies are needed to find out the ability of Bacillus species to induce oviposition behavior under various conditions, especially in human dwellings. Of all the species considered, B. pumilus is particularly attractive because it has been recovered from all our study sites.
This study succeeded in identifying several candidate species for paratransgenesis in P. papatasi: B. flexus, B. pumilus, B. licheniformis, B. megaterium and B. subtilis. These bacteria are genetically tractable and trackable and are often used as probiotics. Most importantly, B. pumilus and B. licheniformis have been proposed as strong oviposition inducers for gravid P. papatasi . The latter fact identifies those bacteria as true symbionts and not merely as environmental contaminants, which might be crucial for the dissemination of the bacteria into sand fly populations.
The authors would like to thank Dr. Elyes Zhioua and Dr. Ifhem Chelbi for their help in collecting sand flies in Tunisia and to Tobin Rowland for his help with sand fly maintenance at WRAIR. The opinions or assertions contained herein are the private views of the authors and are not to be construed as official, or as reflecting true views of the Department of the Army or the Department of Defense.
Conceived and designed the experiments: KG JM HRB ER. Performed the experiments: JM KG. Analyzed the data: JM KG HRB. Contributed reagents/materials/analysis tools: JM KG ER HRB. Wrote the paper: JM KG HRB ER.
- 1. Killick-Kendrick R (2002) Phlebotomine sand flies: biology and control. In: Farrell JP, editor. Leishmania. Dordrecht: Kluwer Academic. pp. 33–43.
- 2. Lane RP (1993) Sandflies (Phlebotominae). In: Lane RP, Crosskey RW, editors. Medical Insects and Arachnids. London: Chapman & Hall. pp. 79–119.
- 3. Rutledge LC, Gupta RK (2009) Moth flies and sand flies (Psychodidae). Med Vet Entomol. 2nd ed. London: Academic Press. pp. 147–162.
- 4. Depaquit J, Grandadam M, Fouque F, Andry PE, Peyrefitte C (2010) Arthropod-borne viruses transmitted by Phlebotomine sandflies in Europe: a review. Eurosurveillance 15: 40–47.
- 5. Feldmann H (2011) Truly emerging-A new disease caused by a novel virus. N Engl J Med 364: 1561–1563.
- 6. Papa A, Velo E, Bino S (2011) A novel phlebovirus in Albanian sandflies. Clin Microbiol Infect 17: 585–587.
- 7. Yu X-J, Liang M-F, Zhang S-Y, Liu Y, Li J-D, et al. (2011) Fever with thrombocytopenia associated with a novel Bunyavirus in China. N Engl J Med 364: 1523–1532.
- 8. Jhingaran A, Chatterjee M, Madhubala R (2008) Leishmaniasis: Epidemiological trends and diagnosis. In: Myler PJ, Fasel N, editors. Leishmania: After the Genome. Norfolk: Caister Academic Press. pp. 1–14.
- 9. Pavli A, Maltezou HC (2010) Leishmaniasis, an emerging infection in travelers. Int J Infect Dis 14: E1032–E1039.
- 10. Ready PD (2010) Leishmaniasis emergence in Europe Eurosurveillance 15: e19505.
- 11. Desjeux P (2004) Leishmaniasis: current situation and new perspectives. Comp Immunol Microbiol Infect Dis 27: 305–318.
- 12. WHO (2011) Leishmaniasis. http://www.who.int/leishmaniasis/en/.
- 13. Mondal S, Bhattacharya P, Ali N (2010) Current diagnosis and treatment of visceral leishmaniasis Expert Rev Anti-Infect Ther 8: 919–944.
- 14. Sundar S, Mehta H, Suresh A, Singh SP, Rai M, et al. (2004) Amphotericin B treatment for Indian visceral leishmaniasis: Conventional versus lipid formulations. Clin Infect Dis 38: 377–383.
- 15. Killick-Kendrick R (1990) Phlebotomine vectors of the leishmaniases-a review. Med Vet Entomol 4: 1–24.
- 16. Parvizi P, Benlarbi M, Ready PD (2003) Mitochondrial and Wolbachia markers for the sandfly Phlebotomus papatasi: little population differentiation between peridomestic sites and gerbil burrows in Isfahan province, Iran. Med Vet Entomol 17: 351–362.
- 17. Tarallo VD, Dantas-Torres F, Lia RP, Otranto D (2010) Phlebotomine sand fly population dynamics in a leishmaniasis endemic peri-urban area in southern Italy. Acta Trop 116: 227–234.
- 18. Feliciangeli MD (2004) Natural breeding places of phlebotomine sandflies. Med Vet Entomol 18: 71–80.
- 19. Killick-Kendrick R (1999) The biology and control of phlebotomine sand flies. Clin Dermatol 17: 279–289.
- 20. Dhiman RC, Shetty PS, Dhanda V (1983) Breeding habitats of phlebotomine sandflies in Bihar, India. Indian J Med Res 77: 29–32.
- 21. Pandya AP, Niyogi AK (1980) Ecological studies on immature stage phlebotomid sandflies in Gujarat. Indian J Med Res 72: 355–358.
- 22. Sivagnaname N, Amalraj DD (1997) Breeding habitats of vector sandflies and their control in India. J Commun Dis 29: 153–159.
- 23. Doha S, Kamal H, Shehata M, Helmy N, Kader MA, et al. (1980) The breeding habitats of Phlebotomus sand flies (Diptera: Psychodidae) in El Agamy, Alexandria, Egypt. J Egypt Soc Parasitol 20: 747–752.
- 24. Chelbi I, Kaabi B, Derbali M, Ahmed SBH, Dellagi K, et al. (2008) Zooprophylaxis: Impact of breeding rabbits around houses on reducing the indoor abundance of Phlebotomus papatasi. Vector-Borne Zoonotic Dis 8: 741–747.
- 25. Mueller GC, Kravchenko VD, Rybalov L, Schlein Y (2011) Characteristics of resting and breeding habitats of adult sand flies in the Judean desert. J Vector Ecol. pp. S195–S205.
- 26. Dantas-Torres F, Latrofa MS, Otranto D (2010) Occurrence and genetic variability of Phlebotomus papatasi in an urban area of southern Italy. Parasit Vectors 3: e77.
- 27. Alexander B, Maroli M (2003) Control of phlebotomine sandflies. Med Vet Entomol 17: 1–18.
- 28. Kishore K, Kumar V, Kesari S, Dinesh DS, Kumar AJ, et al. (2006) Vector control in leishmaniasis. Indian J Med Res 123: 467–472.
- 29. Radjame K, Srinivasan R, Dhanda V (1997) Oviposition response of phlebotomid sandfly Phlebotomus papatasi to soil bacteria isolated from natural breeding habitats. Indian J Exp Biol 35: 59–61.
- 30. Wasserberg G, Rowton ED (2011) Sub-additive effect of conspecific eggs and frass on oviposition rate of Lutzomyia longipalpis and Phlebotomus papatasi. J Vector Ecol. pp. S138–S143.
- 31. Volf P, Kiewegova A, Nemec A (2002) Bacterial colonisation in the gut of Phlebotomus duboscqi (Diptera: Psychodidae): transtadial passage and the role of female diet. Folia Parasitol 49: 73–77.
- 32. Guernaoui S, Garcia D, Gazanion E, Ouhdouch Y, Boumezzough A, et al. (2011) Bacterial flora as indicated by PCR-temperature gradient gel electrophoresis (TGGE) of 16S rDNA gene fragments from isolated guts of phlebotomine sand flies (Diptera: Psychodidae). J Vector Ecol. pp. S144–S147.
- 33. Gouveia C, Asensi MD, Zahner V, Rangel EF, de Oliveira SMP (2008) Study on the bacterial midgut microbiota associated to different Brazilian populations of Lutzomyia longipalpis (Lutz & Neiva) (Diptera: Psychodidae). Neotrop Entomol 37: 597–601.
- 34. Hillesland H, Read A, Subhadra B, Hurwitz I, McKelvey R, et al. (2008) 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 79: 881–886.
- 35. Dillon RJ, ElKordy E, Shehata M, Lane RP (1996) The prevalence of a microbiota in the digestive tract of Phlebotomus papatasi. Ann Trop Med Parasitol 90: 669–673.
- 36. Adler S, Theodor O (1929) Attempts to transmit Leishmania tropica by bite: the transmision of L. tropica by Phlebotomus sergenti. Ann Trop Med Parasitol 23: 1–18.
- 37. Schlein Y, Polacheck I, Yuva lB (1985) Mycoses, bacterial infections and antibacterial activity in sandflies (Psychodidae) and their possible role in the transmission of leishmaniasis. Parasitology 90: 57–66.
- 38. Azambuja P, Garcia ES, Ratcliffe NA (2005) Gut microbiota and parasite transmission by insect vectors. Trends Parasitol 21: 568–572.
- 39. Coutinho-Abreu IV, Zhu KY, Ramalho-Ortigao M (2010) Transgenesis and paratransgenesis to control insect-borne diseases: current status and future challenges. Parasitol Int 59: 1–8.
- 40. Fieck A, Hurwitz I, Kang AS, Durvasula R (2010) Trypanosoma cruzi: Synergistic cytotoxicity of multiple amphipathic anti-microbial peptides to T. cruzi and potential bacterial hosts. Exp Parasitol 125: 42–347.
- 41. Aksoy S, Weiss B, Attardo G (2008) Paratransgenesis applied for control of tsetse transmitted sleeping sickness. Adv Exp Med Biol 627: 35–48.
- 42. Pontes M H, Dale C (2011) Lambda Red-mediated genetic modification of the insect endosymbiont Sodalis glossinidius. Appl Environ Microbiol 77: 1918–1920.
- 43. Ren X, Hoiczyk E, Rasgon JL (2008) Viral paratransgenesis in the malaria vector Anopheles gambiae. PLoS Pathog 4: e1000135.
- 44. Bextine B, Lauzon C, Potter S, Lampe D, Miller TA (2004) Delivery of a genetically marked Alcaligenes sp. to the glossy-winged sharpshooter for use in a paratransgenic control strategy. Curr Microbiol 48: 327–331.
- 45. Ramirez JL, Perring TM, Miller TA (2008) Fate of a genetically modified bacterium in foregut of glassy-winged sharpshooter (Hemiptera: Cicadellidae). J Econ Entomol 101: 1519–1525.
- 46. Husseneder C, Collier RE, Bourtzis K, Miller TA (2009) Paratransgenesis in termites. In: Bourtzis K, Miller TA, editors. Symbiosis. Boca Raton: CRC Press. pp. 361–376.
- 47. Subhadra B, Hurwitz I, Fieck A, Rao DVS, Rao GS, et al. (2010) Development of paratransgenic Artemia as a platform for control of infectious diseases in shrimp mariculture. J Appl Microbiol 108: 831–840.
- 48. Erickson DL, Anderson NE, Cromar LM, Jolley A (2009) Bacterial communities associated with flea vectors of plague. J Med Entomol 46: 1532–1536.
- 49. Ricci I, Mosca M, Valzano M, Damiani C, Scuppa P, et al. (2011) Different mosquito species host Wickerhamomyces anomalus (Pichia anomala): perspectives on vector-borne diseases symbiotic control. Antonie Van Leeuwenhoek 99: 43–50.
- 50. Lantova L, Ghosh K, Svobodova M, Braig HR, Rowton E, et al. (2010) The life cycle and host specificity of Psychodiella sergenti n. sp. and Ps. tobbi n. sp. (Protozoa: Apicomplexa) in sand flies Phlebotomus sergenti and Ph. tobbi (Diptera: Psychodidae). J Invertebr Pathol 105: 182–189.
- 51. Perotti MA, Braig HR (2011) Eukaryotic ectosymbionts of Acari. J Appl Entomol 135: 514–523.
- 52. Votypka J, Lantova L, Ghosh K, Braig H, Volf P (2009) Molecular characterization of gregarines from sand flies (Diptera: Psychodidae) and description of Psychodiella n. g. (Apicomplexa: Gregarinida). J Eukaryot Microbiol 56: 583–588.
- 53. Modi GB, Rowton ED (1999) Laboratory maintenance of phlebotomine sand flies. In: Maramorosch K, Mahmood F, editors. Maintenance of human, animal, and plant pathogen vectors. Enfield, NH: Science Publishers. pp. 109–121.
- 54. Lewis DJ (1978) Phlebotomine sand flies of the oriental region. Bull Brit Mus (Nat Hist) Entomol 37: 217–343.
- 55. Lane DJ (1991) 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M, editors. Nucleic Acid Techniques in Bacterial Systematics. Chichester: Wiley and Sons. pp. 115–175.
- 56. Reysenbach AL, Wickham GS, Pace NR (1994) Phylogenetic analysis of the hyperthermophilic pink filament community in Octopus Spring, Yellowstone National Park. Appl Environ Microbiol 60: 2113–2119.
- 57. Ben-Dov E, Shapiro OH, Siboni N, Kushmaro A (2006) Advantage of using inosine at the 3 termini of 16S rRNA gene universal primers for the study of microbial diversity. Appl Environ Microbiol 72: 6902–6906.
- 58. Gouy M, Guindon S, Gascuel O (2010) SeaView version 4: a multiplatform graphical user interface for sequence alignment and phylogenetic tree building. Mol Biol Evol 27: 221–224.
- 59. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. (2011) MEGA5: Molecular Evolutionary Genetics Analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol Biol Evol 28: 2731–2739.
- 60. Pruesse E, Quast C, Knittel K, Fuchs B, Ludwig W, et al. (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Res 35: 7188–7196.
- 61. Guindon S, Gascuel O (2003) A simple, fast and accurate method to estimate large phylogenies by maximum-likelihood. Syst Biol 52: 696–704.
- 62. Posada D (2008) jModelTest: Phylogenetic model averaging. Mol Biol Evol 25: 1253–1256.
- 63. Ronquist F, Huelsenbeck JP (2003) MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19: 1572–1574.
- 64. Tay B-Y, Lokesh BE, Lee C-Y, Sudesh K (2010) Polyhydroxyalkanoate (PHA) accumulating bacteria from the gut of higher termite Macrotermes carbonarius (Blattodea: Termitidae). World J Microbiol Biotechnol 26: 1015–1024.
- 65. Sanchez-Gonzalez M, Blanco-Gamez A, Escalante A, Valladares AG, Olvera C, et al. (2011) Isolation and characterization of new facultative alkaliphilic Bacillus flexus strains from maize processing waste water (nejayote). Lett Appl Microbiol 52: 413–419.
- 66. Singh RP, Mantri VA, Reddy CRK, Jha B (2011) Isolation of seaweed-associated bacteria and their morphogenesis-inducing capability in axenic cultures of the green alga Ulva fasciata. Aquat Biol 12: 13–21.
- 67. Patil PB, Zeng Y, Coursey T, Houston P, Miller I, et al. (2010) Isolation and characterization of a Nocardiopsis sp. from honeybee guts. FEMS Microbiol Lett 312: 110–118.
- 68. Cutting SM (2011) Bacillus probiotics. Food Microbiol 28: 214–220.
- 69. Mandiki SNM, Milla S, Wang N, Blanchard G, Djonkack T, et al. (2011) Effects of probiotic bacteria on growth parameters and immune defence in Eurasian perch Perca fluviatilis L. larvae under intensive culture conditions. Aquacult Res 42: 693–703.
- 70. Molina CA, Cana-Roca JF, Osuna A, Vilchez S (2010) Selection of a Bacillus pumilus strain highly active against Ceratitis capitata (Wiedemann) larvae. Appl Environ Microbiol 76: 1320–1327.
- 71. Pichinoty F (1984) [Description of the type strain of Bacillus badius]. Ann Microbiol (Paris) B135: 21–27.
- 72. Aksoy HM, Ozman-Sullivan SK (2008) Isolation of Bacillus megaterium from Aphis pomi (Homoptera: Aphididae) and assessment of its pathogenicity. J Plant Pathol 90: 449–452.
- 73. Swiecicka I (2008) Natural occurrence of Bacillus thuringiensis and Bacillus cereus in eukaryotic organisms: a case for symbiosis. Biocontrol Sci Technol 18: 221–239.
- 74. Chatterjee S, Ghosh TS, Das S (2010) Virulence of Bacillus cereus as natural facultative pathogen of Anopheles subpictus Grassi (Diptera: Culicidae) larvae in submerged rice-fields and shallow ponds. Afr J Biotechnol 9: 6983–6986.
- 75. Stenfors Arnesen LP, Fagerlund A, Granum PE (2008) From soil to gut: Bacillus cereus and its food poisoning toxins. FEMS Microbiol Rev 32: 579–606.
- 76. Toledo AV, Alippi AM, de Remes Lenicov AMM (2011) Growth inhibition of Beauveria bassiana by bacteria isolated from the cuticular surface of the corn leafhopper, Dalbulus maidis and the planthopper, Delphacodes kuscheli, two important vectors of maize pathogens. J Insect Sci 11: 1–13.
- 77. Reva ON, Smirnov VV, Pettersson B, Priest FG (2002) Bacillus endophyticus sp nov., isolated from the inner tissues of cotton plants (Gossypium sp.). Int J Syst Evol Microbiol 52: 101–107.
- 78. Sanders ME, Morelli L, Tompkins TA (2003) Sporeformers as human probiotics: Bacillus, Sporolactobacillus, and Brevibacillus. Compr Rev Food Sci Food Safety 2: 101–110.
- 79. Evans JD, Armstrong TN (2005) Inhibition of the American foulbrood bacterium, Paenibacillus larvae larvae, by bacteria isolated from honey bees. J Apicult Res 44: 168–171.
- 80. Lin C, Gan L, Chen Z-L (2010) Biodegradation of naphthalene by strain Bacillus fusiformis (BFN). J Hazard Mater 182: 771–777.
- 81. Zhao CQ, Zhang XY, Song D, Zheng LX, Chen WY (2010) Removal of chromium (VI) from tannery wastewater by immobilized Bacillus fusiformis. J Soc Leather Technol Chem 94: 21–25.
- 82. Ahmed I, Yokota A, Yamazoe A, Fujiwara T (2007) Proposal of Lysinibacillus boronitolerans gen. nov. sp. nov., and transfer of Bacillus fusiformis to Lysinibacillus fusiformis comb. nov. and Bacillus sphaericus to Lysinibacillus sphaericus comb. nov. Int J Syst Evol Microbiol 57: 1117–1125.
- 83. Raats D, Halpern M (2007) Oceanobacillus chironomi sp nov., a halotolerant and facultatively alkaliphilic species isolated from a chironomid egg mass. Int J Syst Evol Microbiol 57: 255–259.
- 84. Whon TW, Jung M-J, Roh SW, Nam Y-D, Park E-J, et al. (2010) Oceanobacillus kimchii sp nov isolated from a traditional Korean fermented food. J Microbiol 48: 862–866.
- 85. An S-Y, Asahara M, Goto K, Kasai H, Yokota A (2007) Terribacillus saccharophilus gen. nov., sp nov and Terribacillus halophilus sp nov., spore-forming bacteria isolated from field soil in Japan. Int J Syst Evol Microbiol 57: 51–55.
- 86. Tozlu E, Dadasoglu F, Kotan R, Tozlu G (2011) Insecticidal effects of some bacteria on Brachus dentipes Baudi (Coleoptera: Bruchidae). Fresenius Environ Bull 20: 918–923.
- 87. Chandel S, Allan EJ, Woodward S (2010) Biological control of Fusarium oxysporum f. sp. lycopersici on tomato by Brevibacillus brevis. J Phytopathol 158: 470–478.
- 88. Yildirim E, Karlidag H, Turan M, Dursun A, Goktepe F (2011) Growth, nutrient uptake, and yield promotion of broccoli by plant growth promoting rhizobacteria with manure. HortScience 46: 932–936.
- 89. Genersch E (2010) American foulbrood in honeybees and its causative agent, Paenibacillus larvae. J Invertebr Pathol. pp. S10–S19.
- 90. Govindasamy V, Senthilkumar M, Magheshwaran V, Kumar U, Bose P, et al. (2010) Bacillus and Paenibacillus spp.: Potential PGPR for sustainable agriculture. In: Maheshwari DK, editor. Plant Growth and Health Promoting Bacteria. New York: Springer. pp. 333–364.
- 91. Butler JF, Garcia-Maruniak A, Meek F, Maruniak JE (2010) Wild Florida house flies (Musca domestica) as carriers of pathogenic bacteria. Fla Entomol 93: 218–223.
- 92. Krishnamurthi S, Chakrabarti T, Stackebrand E (2009) Re-examination of the taxonomic position of Bacillus silvestris Rheims et al. 1999 and proposal to transfer it to Solibacillus gen. nov. as Solibacillus silvestris comb. nov. Int J Syst Evol Microbiol 59: 1054–1058.
- 93. Nada DV, Vukovic V, Nedic N (2010) Pathogenicity of some bacterial species isolated from the bee digestive tract. Acta Vet 60: 49–57.
- 94. Matsuura Y, Koga R, Nikoh N, Meng X-Y, Hanada S, et al. (2009) Huge symbiotic organs in giant scale insects of the genus Drosicha (Coccoidea: Monophlebidae) harbor flavobacterial and enterobacterial endosymbionts. Zoolog Sci 26: 448–456.
- 95. Chang E-P, Chiang D-H, Lin M-L, Chen T-L, Wang F-D, et al. (2009) Clinical characteristics and predictors of mortality in patients with Enterobacter aerogenes bacteremia. J Microbiol Immunol Infect 42: 329–335.
- 96. Hamilton JV, Lehane MJ, Braig HR (2003) Isolation of Enterobacter sakazakii from midgut of Stomoxys calcitrans. Lancet 9: 1355–1356.
- 97. Mramba F, Broce AB, Zurek L (2007) Vector competence of stable flies, Stomoxys calcitrans L. (Diptera: Muscidae) for Enterobacter sakazakii. J Vector Ecol 32: 134–139.
- 98. Healy B, Cooney S, O'Brien S, Iversen C, Whyte P, et al. (2010) Cronobacter (Enterobacter sakazakii): An opportunistic foodborne pathogen. Foodborne Pathog Dis 7: 339–350.
- 99. Caspi-Fluger A, Zchori-Fein E (2010) Do plants and insects share the same symbionts? Isr J Plant Sci 58: 113–119.
- 100. Nadarasah G, Stavrinides J (2011) Insects as alternative hosts for phytopathogenic bacteria. FEMS Microbiol Rev 35: 555–575.
- 101. Kelley ST, Dobler S (2011) Comparative analysis of microbial diversity in Longitarsus flea beetles (Coleoptera: Chrysomelidae). Genetica 139: 541–550.
- 102. Sanchez-Contreras M, Vlisidou I (2008) The diversity of insect-bacteria interactions and its applications for disease control. Biotechnol Genet Eng Rev 25: 203–243.
- 103. Weber DJ, Rutala WA, Sickbert-Bennett EE (2007) Outbreaks associated with contaminated antiseptics and disinfectants. Antimicrob Agents Chemother 51: 4217–4224.
- 104. Doughari HJ, Ndakidemi PA, Human IS, Benade S (2011) The ecology, biology and pathogenesis of Acinetobacter spp.: An overview. Microbes Environ 26: 101–112.
- 105. Davari B, Kalantar E, Zahirnia A, Moosa-Kazemi SH (2010) Frequency of resistance and susceptible bacteria isolated from houseflies. Iran J Arthropod-Borne Dis 4: 50–55.
- 106. Silby MW, Winstanley C, Godfrey SAC, Levy SB, Jackson RW (2011) Pseudomonas genomes: diverse and adaptable. FEMS Microbiol Rev 35: 652–680.
- 107. Behrendt U, Ulrich A, Schumann P, Naumann D, Suzuki K (2002) Diversity of grass-associated Microbacteriaceae isolated from the phyllosphere and litter layer after mulching the sward; polyphasic characterization of Subtercola pratensis sp. nov., Curtobacterium herbarum sp. nov. and Plantibacter flavus gen. nov., sp. nov. Int J Syst Evol Microbiol 52: 1441–1454.
- 108. Indiragandhi P, Yoon C, Yang JO, Cho S, Sa TM, et al. (2010) Microbial communities in the developmental stages of B and Q biotypes of sweetpotato whitefly, Bemisia tabaci (Hemiptera: Aleyrodidae). J Korean Soc Appl Biol Chem 53: 605–617.
- 109. Gneiding K, Frodl R, Funke G (2008) Identities of Microbacterium spp. encountered in human clinical specimens. J Clin Microbiol 46: 3646–3652.
- 110. Hogg JC, Lehane MJ (2001) Microfloral diversity of cultured and wild strains of Psoroptes ovis infesting sheep. Parasitology 123: 441–446.
- 111. Cousin FJ, Mater DDG, Foligne B, Jan G (2011) Dairy Propionibacteria as human probiotics: A review of recent evidence. Dairy Sci Technol 91: 1–26.
- 112. Seo JK, Kim S-W, Kim MH, Upadhaya SD, Kam DK, et al. (2010) Direct-fed microbials for ruminant animals. Asian-Australas J Anim Sci 23: 1657–1667.
- 113. Jappe U (2003) Pathological mechanisms of acne with special emphasis on Propionibacterium acnes and related therapy. Acta Derm Venereol 83: 241–248.
- 114. Yamada T, Sekiguchi Y (2009) Cultivation of uncultured Chloroflexi subphyla: Significance and ecophysiology of formerly uncultured Chloroflexi ‘Subphylum I’ with natural and biotechnological relevance. Microbes Environ 24: 205–216.
- 115. Didelot X, Barker M, Falush D, Priest FG (2009) Evolution of pathogenicity in the Bacillus cereus group. Syst Appl Microbiol 32: 81–90.
- 116. De Gheldre Y, Maes N, Rost F, De Ryck R, Clevenbergh P, et al. (1997) Molecular epidemiology of an outbreak of multidrug-resistant Enterobacter aerogenes infections and in vivo emergence of imipenem resistance. J Clin Microbiol 35: 152–160.
- 117. Vasilev GI, Bazanova LP (1987) [The effect of the entomopathogenic bacteria-Bacillus mycoides Flugge and Bacillus circulans Jordan-on the gut microflora of larval fleas]. In: Cherepanov AI, editor. Ekologiya i geografiya chlenistonogikh Sibiri [Ecology and Geography of Arthropods of Siberia]. Novosibirsk: Nauka. pp. 208–210.
- 118. Darriet F, Hougard J-M (2002) An isolate of Bacillus circulans toxic to mosquito larvae. J Am Mosq Control Assoc 18: 65–67.
- 119. Ghosh K, Sen SK, Ray AK (2003) Supplementation of an isolated fish gut bacterium, Bacillus circulans, in formulated diets for rohu, Labeo rohita, fingerlings. Isr J Aquacult Bamidgeh 55: 13–21.
- 120. Chakraborty U, Chakraborty B, Basnet M (2006) Plant growth promotion and induction of resistance in Camellia sinensis by Bacillus megaterium. J Basic Microbiol 46: 186–195.
- 121. de Freitas JR, Banerjee MR, Germida JJ (1997) Phosphate-solubilizing rhizobacteria enhance the growth and yield but not phosphorus uptake of canola (Brassica napus L.). Biol Fertil Soils 24: 4358–4364.
- 122. Morsy TA, Aboul ERG, Abdelmawla MM, el Gozamy BM (1993) Counter immuno-electrophoresis, a modified technique for the identification of blood meals of sandflies collected from Qualyobia Governorate, Egypt. J Egypt Soc Parasitol 23: 109–132.
- 123. Namita M, Joshi V, Bansal SK (1991) Host preference pattern of phlebotomine sandflies of Bikaner city. Indian J Med Res 93: 328–329.
- 124. Singh R, Lal S, Saxena VK (2008) Breeding ecology of visceral leishmaniasis vector sandfly in Bihar state of India. Acta Trop 107: 117–120.
- 125. Benardini JN, Sawyer J, Venkateswaran K, Nicholson WL (2003) Spore, UV and acceleration resistance of endolithic Bacillus pumilus and Bacillus subtilis isolates obtained from Sonoran desert basalt: implications for lithopanspermia. Astrobiology 3: 709–717.
- 126. Kempf MJ, Chen F, Kern R, Venkateswaran K (2005) Recurrent isolation of hydrogen peroxide-resistant spores of Bacillus pumilus from a spacecraft assembly facility. Astrobiology 5: 391–405.
- 127. Bottone EJ, Peluso RW (2003) Production by Bacillus pumilus (MSH) of an antifungal compound that is active against Mucoraceae and Aspergillus species: preliminary report. J Med Microbiol 52: 69–74.
- 128. Naruse N, Tenmyo O, Kobaru S, Kamei H, Miyaki T, et al. (1990) Pumilacidin, a complex of new antiviral antibiotics. Production, isolation, chemical properties, structure and biological activity. J Antibiot (Tokyo) 43: 267–280.
- 129. Oliveira S M, Moraes BA, Gonçalves CA, Giordano-Dias CM, D'Almeida JM, et al. (2000) [Prevalence of microbiota in the digestive tract of wild females of Lutzomyia longipalpis Lutz & Neiva, 1912) (Diptera: Psychodidae)]. Rev Soc Bras Med Trop 33: 319–322.
- 130. Schmidt TR, Scott EJ, II , Dyer DW (2011) Whole-genome phylogenies of the family Bacillaceae and expansion of the sigma factor gene family in the Bacillus cereus species-group. BMC Genomics 12: e430.
- 131. Boehme K, Fernandez-No IC, Barros-Velazquez J, Gallardo JM, Canas B, et al. (2011) Rapid species identification of seafood spoilage and pathogenic Gram-positive bacteria by MALDI-TOF mass fingerprinting. Electrophoresis 32: 2951–2965.
- 132. Schlein Y, Jacobson RL (1994) Mortality of Leishmania major in Phlebotomus papatasi caused by plant feeding of the sand flies. Am J Trop Med Hyg 50: 20–27.