Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Supplemental material Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Mosi, L. Articles by Small, P. L. C. Search for Related Content PubMed PubMed Citation Articles by Mosi, L. Articles by Small, P. L. C. Agricola Articles by Mosi, L. Articles by Small, P. L. C.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, November 2008, p. 7036-7042, Vol. 74, No. 22
0099-2240/08/$08.00+0     doi:10.1128/AEM.01234-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Persistent Association of Mycobacterium ulcerans with West African Predaceous Insects of the Family Belostomatidae ^,

Lydia Mosi,¹ Heather Williamson,¹ John R. Wallace,² Richard W. Merritt,³ and P. L. C. Small^1*

Department of Microbiology, University of Tennessee, Knoxville, Tennessee 37996,¹ Department of Biology, Millersville University, Millersville, Pennsylvania 17551,² Department of Entomology, Michigan State University, East Lansing, Michigan 48824-1115³

Received 3 June 2008/ Accepted 22 September 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
A number of studies have suggested that Mycobacterium ulcerans, the etiological agent of Buruli ulcer, may be transmitted to humans by insect bites. M. ulcerans has been isolated from a predaceous aquatic insect, and PCR detection of M. ulcerans DNA in aquatic environments suggests that the organism is widely distributed within many invertebrate taxa and functional feeding groups. Thus, M. ulcerans may be concentrated through different trophic links. However, the specific environmental niche of M. ulcerans and route of transmission to humans remain a mystery. In this study, a biologically relevant infection model in which M. ulcerans-infected mosquito larvae were fed to a species of predaceous hemiptera (African Belostomatidae) was used to demonstrate the persistent colonization of M. ulcerans and subsequent transmission of bacteria to naïve prey. The association of M. ulcerans with specific anatomical compartments showed that M. ulcerans accumulates preferentially on the exoskeleton. In contrast, few organisms were found in dissected guts or salivary glands. No difference was found between the ability of wild-type M. ulcerans and an M. ulcerans isogenic mycolactone-negative mutant to colonize belostomatids. These data show that African belostomatids can successfully be colonized by M. ulcerans and support the trophic transfer of M. ulcerans within the environment.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mycobacterium ulcerans disease (Buruli ulcer) continues to be one of the most debilitating cutaneous diseases in West Africa. Although the distribution of the disease is global and affects people of all ages, the burden of disease is most severe in West Africa, where Buruli ulcer is an emerging disease. In West Africa, cases typically occur among rural, economically deprived populations (11, 31). M. ulcerans is an environmental pathogen; however, the method of transmission from the environment to humans remains elusive (5, 11, 32). Person-to-person transmission of Buruli ulcer is extremely rare, and a large body of evidence implicates exposure to slow moving or stagnant water as the most universally defined risk factor for infection (23, 25). A striking characteristic of the disease in all regions is its discontinuous focal distribution. Villages where the disease is endemic and those where it is not endemic may be found within a few kilometers of each other along a waterway (23, 25, 26).

The absence of the disease in arid parts of the world strongly suggests that environmental constraints may limit the distribution of disease. In addition, there is a strong association of the bacterium within aquatic ecosystems. The possibility of the bacterium being concentrated through trophic links and ultimately delivered to an unsuspecting host, via a vector or some other unknown route, has been suggested by several investigators (6, 14). A major advance in understanding transmission occurred with the detection of M. ulcerans DNA in predaceous insects (Naucoridae and Belostomatidae), leading to the hypothesis that insects may be involved in the transmission (20). Naucorids and belostomatids are aquatic hemiptera that exploit a wide range of prey, including snails, fish, anuran larvae, and other terrestrial and aquatic insects (29). Both insect groups are found worldwide near vegetative areas of stagnant water bodies. Although they do not feed on humans, they can bite if they are disturbed and for this reason are called "toe biters." Subsequent work from Ghana and Cote d'Ivoire have confirmed the presence of M. ulcerans DNA in a large number of aquatic invertebrate and vertebrate taxa (6, 18, 20, 21). There have been numerous unsuccessful attempts to culture the bacteria from the environment due to competition from other, faster-growing environmental bacteria. A major breakthrough occurred with the successful culture of M. ulcerans from a water strider (Gerridae) collected in Benin by Francoise Portaels et al. (22). None of the predaceous hempitera that were found to have positive PCR results for M. ulcerans DNA in Africa are hematophagous, and the percentage of biting hemiptera is often quite low in areas where the disease is endemic (2).

In addition to extensive field work on the occurrence of M. ulcerans in natural environments, the potential role of biting hemiptera as vectors of M. ulcerans has been extensively explored in a set of elegant laboratory studies conducted by Laurent Marsollier et al. (13, 15, 16). In these studies, diptera larvae (Phormia terrae-novae) were injected with suspensions of M. ulcerans and subsequently fed to Naucoris cimicoides, collected from swamps in France (13). Marsollier and colleagues showed from these studies that M. ulcerans successfully colonized the insect over a period exceeding 90 days, causing no growth impairment or death of the insect. More interestingly, they showed through microscopy that whereas wild-type bacteria could be detected on the raptorial arms of naucorids, the majority of organisms were localized in the salivary glands of the insect. Finally, Marsollier et al. (15) also presented data supporting the role of the M. ulcerans cytotoxic macrolide toxin, mycolactone, in colonization of the insect.

Although these studies were of significance, it is difficult to determine their relevance to transmission of Buruli ulcer in Africa for a number of reasons. First, the species of Naucoridae used was from France, not Africa where Buruli ulcer is endemic. Second, the bacterial strain used for most of these studies is a member of the "ancestral" lineage of M. ulcerans rather than a member of the "classical" lineage associated with severe Buruli ulcer in Africa and Malaysia (12).

To investigate the ability of M. ulcerans to colonize aquatic African hemiptera, we infected adult belostomatids collected in Ghana with a "classical" isolate of M. ulcerans. We also employed a mycolactone-negative mutant of M. ulcerans to determine the role of mycolactone in insect colonization. We showed that belostomatids can be persistently colonized by both mycolactone-producing and mycolactone-negative M. ulcerans; however, we obtained no evidence of replication within internal insect organs. We showed extensive colonization of the exoskeleton and showed that M. ulcerans is transmitted to prey via feeding. In our infection model, we demonstrated the significance of trophic-level transfer of M. ulcerans in the environment in which naturally infected mosquito larvae successfully passed M. ulcerans up the food chain. These results provide a useful model for beginning to understand the ecology of M. ulcerans in West Africa.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Bacterial strains and growth conditions.
The strains used in this study were a green fluorescent protein (GFP)-labeled strain, M. ulcerans 1615 (1615 GFP), M. ulcerans 1615::TN118 GFP (mycolactone negative), and M. ulcerans 01G897. The strain M. ulcerans 1615 GFP (ATCC 35840) is a well-characterized Malaysian human isolate with physical and biochemical properties very similar to those of the sequenced strain M. ulcerans Agy99 from Ghana (7). Transposon mutagenesis (27) was used to generate the mycolactone-negative mutant strain 1615::TN118 with an insertion in the FABH gene (MUP045). This strain produces neither the core nor the side chain of mycolactone, is not cytotoxic, and is avirulent. Both strains were fluorescently labeled by introduction of a GFP gene using the phage-integrating vector psm5 provided by Valdivia et al. (30). By using this method, the GFP gene is inserted into the chromosome of M. ulcerans 1615 GFP in the phage attachment site (att) and has no effect on the virulence of the bacterium. M. ulcerans 01G897 is a French Guyanan human isolate (4). M. ulcerans was grown to mid-log phase in Middlebrook 7H9 (M7H9) medium supplemented with 10% oleic acid-albumin-dextrose enrichment (OADC; Difco) and incubated at 32°C.

Inoculum preparation.
The total number of bacteria used for the infections was determined via CFU. One loop-full of bacteria growing at exponential phase was emulsified in 0.01% sodium dodecyl sulfate, and clumps were broken by being passed through a 25-gauge needle. One hundred microliters of the resulting suspension was plated on M7H9 agar medium supplemented with 10% OADC (Difco) and incubated at 32°C for 6 weeks to determine CFU.

Aquatic insects.
Adult belostomatids (Appasus sp. [Diplonychus sp.]), 1- to 3-cm long were collected from aquatic sampling sites in Ga district, Ghana. Insects were housed individually in deep petri dishes filled with double-distilled water and maintained under a 12-h light and dark photoperiod at 28°C. Insects were fed either midge larvae (Chironomidae) or blowfly larvae (Diptera: Calliphoridae) (Phormia regina) every other day, and the housing water was changed at the same time.

Mosquito (Ochlerotatus triseriatus) egg rafts were obtained from Michigan State University. The egg rafts were submerged in double-distilled water so that they could hatch into larvae in about 3 days. Larvae were maintained on powdered fish food each day until they developed into third instars, when they were fed bacteria and used for infection. Because some belostomatids were collected from areas of endemicity for Buruli ulcer and M. ulcerans DNA has been detected in a small number of belostomatids from West Africa, 110 belostomatids were analyzed by using microscopy and PCR for the presence of M. ulcerans DNA. All insects tested were PCR negative for M. ulcerans and therefore were used as negative controls.

Experimental infection of insects.
Mosquito larvae infected with M. ulcerans 1615 GFP via feeding were used as the primary prey for adult belostomatids in these studies. Infected prey were prepared by starving the naïve larvae (Ochlerotatus triseriatus) for 24 h and then transferring them to a fresh container containing 10⁶/ml M. ulcerans bacteria in 10 ml of double-distilled water, where larvae were allowed to feed on fluorescently tagged M. ulcerans for 24 h. Five representative larvae were removed and analyzed for the presence of M. ulcerans by using light (acid-fast stain) and fluorescent microscopy. At this time point, the guts of virtually all larvae were packed with M. ulcerans (see Fig. S1 in the supplemental material). Three infected larvae were then fed to each belostomatid that had previously been starved for 7 days. There were 36 adult belostomatids per bacterial strain used. Twenty-four hours after infection, each insect was transferred to a new petri dish and maintained on chironomid (Diptera: Chironomidae) midge larvae for the duration of the study period. The water was changed each time insects were fed.

Detection of M. ulcerans in insect tissues.
For infection studies, 36 insects were used for each M. ulcerans strain tested. At 1 day, 30 days, and 60 days postinfection (p.i.), 12 insects were sacrificed for analysis. The 24-h time point was chosen to determine the rate of infection, whereas the later time points were chosen to detect viability, colonization, and multiplication of the bacteria within the insects. At each time point, individual belostomatids' internal organs were carefully removed, and the salivary gland, gut, head, thorax, and forearms (Fig. 1) were homogenized in 200 µl of 1 M Tris-HCl buffer [pH 7.5]. For quantification of the bacteria, four 10-fold dilutions were made of each anatomical section, and smears were made for acid-fast staining and fluorescent microscopy. Acid-fast bacilli (AFB) were viewed with a light microscope (Olympus BX51/BX52). Statistical analysis on the detectable bacterial load per insect anatomical section and the type of bacterial strain were conducted by using the Stata Mann-Whitney two-sample rank-sum test. Wet mounts of each section were viewed using a fluorescent microscope (Nikon Eclipse E400) equipped with a standard epifluorescent attachment filter set for the detection of the fluorescently labeled bacteria. Although AFB microscopy provided better visualization of M. ulcerans morphology, the presence of fluorescently labeled M. ulcerans was required for scoring a belostomatid positive for M. ulcerans as determined by microscopy. For scanning electron microscopy, infected insects were vacuum dried, sputter coated with gold using a SPI-Module sputter coater for 10 s, and mounted on carbon-coated metal stubs. Imaging was performed on a Zeiss 1525 field emission scanning electron microscope equipped with a GEMINI field emission column.

View larger version (54K): [in this window] [in a new window]

FIG. 1. Images of dissected belostomatid, showing the relative sizes of the following insect compartments assayed in this study: head (A), raptorial arms (B), thorax (C), salivary glands (D), and gut (E).

For recovery of viable bacteria from the infected insects, 100 µl of each insect section homogenate was decontaminated via the modified Petroff's method (34). Briefly, 150 µl of 4% NaOH was incubated with 100 µl of insect homogenate for 15 min, followed by a 15-min incubation with 800 µl of sterile saline of the recovered pellet. The resulting pellet, after centrifugation at 3,000 x g, was resuspended in 100 µl sterile saline and plated on M7H9 agar plates supplemented with 10% OADC supplement (Difco), chloramphenicol (20 mg/ml), and cycloheximide (20 mg/ml).

Transmission of M. ulcerans infection to blowfly larvae via feeding.
Twelve infected belostomatids removed at 1 day, 30 days, and 60 days p.i. were allowed to feed individually on a single blowfly larva (Phormia regina). Larval exuviae were collected immediately, homogenized as described above, and analyzed by using microscopy and PCR for the presence of M. ulcerans. As controls, uninfected belostomatids were also allowed to feed on larvae and were analyzed for the presence of M. ulcerans.

DNA extraction and PCR analysis.
DNA was extracted from insect and larval homogenates with the UltraClean soil DNA extraction kit (Mo Bio Laboratories) according to the manufacturer's instruction. The enoyl reductase (mlsA) gene was chosen to determine the presence of mycobacterial DNA in insect tissues as previously described (33). Five microliters of each DNA sample was amplified with the mlsA primer pair 5'-GAGATCGGTCCCGACGTCTAC-3' and 5'-GGCTTGACTCATGTCACGTAAG-3' in 50-µl PCR mixtures using the GoTaq polymerase buffer system (Promega). Each reaction mixture contained 36.7 µl double-distilled water, 5 µl GoTaq green master mix (400 µl of each deoxynucleoside triphosphate, 3 mM MgCl₂, blue and yellow dyes), 1 µM of forward and reverse primers, 1.5 U of GoTaq polymerase, and 5 µl of DNA template. Cycling was performed in a Mastercycler gradient thermal cycler (Eppendorf) as follows: 95°C for 5 min; 35 cycles of 95°C for 1 min, 55°C for 1 min, and 72°C for 1 min; and 72°C for 10 min. Nine microliters of each reaction mixture was analyzed on 1.5% agarose gels in 1x Tris-acetate-EDTA stained with 1 µg/ml ethidium bromide for visualization of amplicons.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mycobacterium ulcerans exhibits prolonged infection in African belostomatids.
We showed that throughout the 60-day study period, all of the belostomatids that had been infected with M. ulcerans 1615 GFP were shown to be positive for the bacteria as determined by either microscopy, PCR analysis, or both (see data in Table S1 in the supplemental material). Based on microscopy and PCR, there was a 100% infection rate as determined by the presence of M. ulcerans in 12/12 insect guts after 24 h (Table 1). This amount declined significantly (P = 0) to 9% at 30 days p.i., with only one gut being AFB positive, and rose slightly to 30% at 60 days. This increase was not statistically significant (P = 0.8357). There was a low infection rate of the salivary glands assayed at all time points, with a mean of 12% insects shown to be positive for bacteria as determined by microscopy for M. ulcerans. By using PCR analysis, however, the average rate of infectivity over time was much higher (63%). The rates of infection in the head, raptorial arms, and thorax followed similar patterns. The largest number of M. ulcerans-positive insects for these three anatomical sections averaged 88% and occurred at 30 days p.i. This increase between 1 day p.i. and 30 days p.i. was statistically significant, with P values of 0.05, 0.0003, and 0.0004, respectively. Between 30 days and 60 days, even though there was a decrease in the number of insects that were positive for bacteria on the head, raptorial arms, and thorax, the difference was not significant. There was some discrepancy between PCR and microscopy results for the percentage of insects positive for M. ulcerans (Tables 1 and 2; see Tables S1 and S2 in the supplemental material). There were significantly more insects shown to be positive as determined by PCR analysis in the head, raptorial arms, and salivary glands at 1 day p.i. (P = 0.02, 0.001, and 0, respectively). By using microscopy, at 30 days p.i. there were significantly more insects shown to be positive for bacteria on the head, raptorial arms, and thorax (P = 0.0006, 0.02, and 0.0006, respectively). Contrary to the decrease in the number of insects that were positive for M. ulcerans at 60 days p.i. in the gut, there was a significant increase determined by PCR analysis (P = 0.002). All insect sections that were shown to be positive for bacteria by bright field microscopy were also positive for GFP bacilli, thus strengthening the observation. A total of three insects died prior to the day 30 and day 60 time points; however, the deaths were not attributed to any effect of M. ulcerans but rather to natural causes. A similar death rate occurred among the uninfected insects.

View this table: [in this window] [in a new window]

TABLE 1. Persistence of M. ulcerans 1615 GFP in specific anatomic compartments as determined by microscopy and PCRa

View this table: [in this window] [in a new window]

TABLE 2. Comparison between persistence of M. ulcerans 1615 GFP and M. ulcerans 1615::TN118 GFP in belostomatids

M. ulcerans increases in external compartments and decreases in internal organs over 60 days.
In order to quantitatively determine multiplication of M. ulcerans within the infected insects, dissected anatomical sections were homogenized, decontaminated by the modified Petroff's method, and cultured on M7H9 medium supplemented with OADC and 20 µg/ml each of chloramphenicol and cycloheximide. These efforts, however, were mostly frustrated by the overgrowth of faster-growing bacteria and fungi present in many insects. The salivary glands produced little bacterial growth and no isolates of M. ulcerans. For this reason, a semiquantitative method for obtaining evidence for the growth of M. ulcerans was employed based on detection and enumeration of acid-fast and GFP-positive bacilli present in dilutions of insect section homogenates. Results from these studies showed that the bacterial load on the exoskeleton (head, raptorial arms, and thorax) was greater than that in the internal organs (salivary glands and gut) at all time points (Fig. 2; see Fig. S2 in the supplemental material). Since the primary infection of the belostomatids was performed orally, it was expected that the insect guts would have a large amount of bacteria compared to other anatomical sites at early time points, and this phenomenon was indeed observed. However, bacterial density dropped significantly (P = 0.0003) in the gut by the 60-day time point. The mean bacterial density within the salivary gland remained below 10 detectable bacterial cells for all time points without any significant changes. In contrast, anatomical sections of the insect covered by a substantial exoskeleton showed a significant and steady increase in density of detectable bacteria over the 60-day study period with as many as 1,000 bacterial cells seen per viewing field at x100 magnification (P values of 0.05, 0.01, and 0.019 for head, raptorial arms, and thorax, respectively).

View larger version (37K): [in this window] [in a new window]

FIG. 2. Average number of M. ulcerans 1615 GFP and M. ulcerans 1615::TN118 GFP bacterial cells per the following insect sections: head (A), raptorial arms (B), thorax (C), salivary glands (D), and gut (E). AFB abundance as determined by the number of bacilli within 50 random viewing fields at x100 magnification computed from a most probable number direct smear preparation. y axis, number of bacterial cells; x axis, days p.i.; *, significant difference between M. ulcerans 1615 GFP (dark gray bars) and M. ulcerans 1615::TN118 GFP (light gray bars).

M. ulcerans is efficiently transmitted from infected belostomatids to blowfly larva through feeding.
Belostomatids feed by grabbing and immobilizing prey with their raptorial arms and sucking out prey contents with their stylet (28). In order to determine the ability of M. ulcerans-infected belostomatids to transmit M. ulcerans through grabbing and biting, blowfly larva exuviae, which were fed to infected belostomatids, were analyzed for the presence of M. ulcerans by using microscopy, PCR analysis, and culture (see Fig. S3 in the supplemental material). Each insect prior to being sacrificed at 1 day, 30 days, and 60 days p.i. was given a naïve blowfly larva. Larva exuviae collected at each time point were pooled for analysis. M. ulcerans could be detected by using microscopy (Fig. 3) and PCR analysis (data not shown) in all exuviae. Even 60 days after infection with M. ulcerans, belostomatids were still able to transmit M. ulcerans to naïve blowfly larvae through feeding. The attempt to culture back from the blowfly larva exuviae was unsuccessful even after decontamination of samples due to overgrowth of native bacteria. In this experiment, 100% infectivity of maggot exuviae was found as determined by both PCR and microscopy, indicating a direct transmission of bacteria from the insects to their prey.

View larger version (18K): [in this window] [in a new window]

FIG. 3. Presence of M. ulcerans in exuviae of maggots infected by belostomatids. Ziehl Neelsen stain (A and C) and observation under a GFP filer (B and D) of maggot exuviae collected from uninfected (A and B) and M. ulcerans-infected (C and D) belostomatids postfeeding. Magnification, x100. Arrows indicate clusters of acid-fast and fluorescent bacilli, respectively.

Mycolactone is not required for the colonization of belostomatids.
Insects that were infected with the mycolactone-negative strain M. ulcerans 1615::TN118 GFP showed patterns of persistence within African belostomatids similar to those of M. ulcerans 1615 GFP bacteria (see Tables S2 and S3 in the supplemental material). As stated earlier, each infected insect was dissected into five anatomical sections, homogenized, and assayed for the presence of the bacteria. There was a 100% infectivity of insect guts 1 day p.i. as determined by both microscopy and PCR. This efficiency decreased steadily over time to 25% insects at 60 days p.i. as determined by microscopy. However, PCR analysis showed there was a significant increase in the number of insect guts that were positive for bacteria (P = 0.04). Similarly, the salivary glands of the insects were the least infected, and there were no significant differences in the number of positive results as determined by either microscopy or PCR. There were comparable numbers of insects that were positive for bacteria on the head, raptorial arms, and thorax as determined by either microscopy or PCR, and there was no significant difference between the two methods, with the exception of more positive results for bacteria on the head at 30 days and 60 days p.i., shown by microscopy.

More importantly, there were very few significant differences between the number of insects positive for M. ulcerans 1615 GFP and M. ulcerans 1615::TN118 GFP (mycolactone negative), as determined by either microscopy or PCR (see P values in Table 2). Similarly, the external surfaces of the insects showed the highest bacterial density over time. At 1 day p.i., there were significantly more insect raptorial arms positive for the mycolactone-negative strain than for the wild type (P = 0.0004). At 30 days p.i., there were significantly more insects with wild-type-infected raptorial arms and thoraces than there were mycolactone-negative-infected insects (P = 0.003 and 0.0001, respectively). There were no significant differences between M. ulcerans 1615 GFP and mycolactone-negative M. ulcerans 1615::TN118 GFP-infected insects 60 days p.i. In the salivary glands and guts, the bacterial density decreased significantly over the 60-day period, and there was no significant difference between this observation and that of M. ulcerans 1615 GFP-infected insects.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The hypothesis that M. ulcerans is a vectored pathogen which is transmitted to humans via the bite of predaceous water bugs has received considerable attention (2, 13, 17, 24). The interpretation of the data presented in earlier studies has been complicated by the following three major issues: (i) the insect species used were not African species, (ii) the primary M. ulcerans used was not representative of the classical M. ulcerans from Africa, and (iii) none of the studies have provided comprehensive quantitative data on the location of the bacterium within the insect. In this study, we have used an African insect species that has been found to have positive PCR results for M. ulcerans in nature and a classical lineage strain of M. ulcerans to determine the following: (i) whether M. ulcerans persistently colonizes or grows within African predaceous water bugs, (ii) how the bacterium is partitioned within external and internal body parts, (iii) whether mycolactone plays a role in insect infections, and (iv) whether M. ulcerans can be transmitted by water bugs to prey within a food chain (see Fig. S3 in the supplemental material).

Although, like Marsollier and colleagues, we show extensive colonization of the exoskeleton of belostomatids by M. ulcerans (Fig. 2 and 4), we do not have convincing evidence of bacterial growth and replication in the internal compartments of belostomatids. We found very low numbers of M. ulcerans in the salivary glands of belostomatids at all times, in contrast to the work of Marsollier et al. We initially thought that the differential findings of our laboratory and those of Marsollier and colleagues might be due to the use of different lineages of M. ulcerans. However, when tested under comparative conditions, we found no major differences between the ancestral and classical strains in their abilities to associate with belostomatids (data not shown).

View larger version (102K): [in this window] [in a new window]

FIG. 4. Scanning electron micrographs of uninfected (top panel) and M. ulcerans-infected (bottom panel) belostomatids. Shown are the stylet (A and D), the tip of a raptorial arm (B), setae of a raptorial arm (C and F); and a rostrum of an infected insect (E). Arrows indicate clusters of adherent bacteria 60 days p.i.

We also ruled out the possibility that the colonization of external compartments of belostomatids occurred as an artifact of the infection method. In previous studies, and in early work in our lab, M. ulcerans bacteria were injected into blowfly larvae. During this procedure, inoculum leaks out of the larva, producing a significant amount of surface contamination which could be transferred to the insect's raptorial arms when it grasps prey for feeding. We were able to avoid this problem by taking advantage of the fact that mosquito larvae readily ingest bacteria and in turn are consumed by predaceous water insects higher up the food chain. Nonetheless, in our work, as in the work of Marsollier and colleagues, considerable colonization of raptorial arms occurred (Fig. 2 and 4). Belostomatids use their arms for grooming their stylet as well as for grabbing prey (28). This behavior could also lead to colonization of the raptorial arms.

In accordance with Marsollier and colleagues, we showed that infected insects could transmit M. ulcerans via feeding; however, this transfer of bacteria is most likely to have occurred through contact with the heavily colonized raptorial arms and other external parts rather than the salivary glands.

We do not find that the mycolactone toxin plays a significant role in the ability of M. ulcerans to persist within insects because both toxin-positive and isogenic toxin-negative strains persist equally. The evidence for the impaired ability of mycolactone-negative strains to colonize French naucorids rested on a 10-fold difference between wild-type and mycolactone-negative strains, which was minimal considering the length of the experiment (15).

Despite the fact that we collected insects three times during field trips to Ghana, we never obtained sufficient numbers of naucorids to produce statistically sound data. We did, however, conduct limited studies with the small number of naucorids we were able to collect. Results from these studies were similar to those using belostomatids in that we did not see any evidence of replication in the salivary gland, but the numbers of insects and time points were too few for statistical analysis.

The presence of the numerous and complex microbial flora found in belostomatids, especially in the gut, made it impossible to obtain any M. ulcerans CFU. Despite the fact that we detected few other bacteria in the salivary glands, we did not obtain a culture of M. ulcerans. The long-term instability of GFP under neutral pH suggests that the presence of fluorescent M. ulcerans in belostomatids is consistent with the presence of viable organisms. We did show long-term colonization of belostomatids by M. ulcerans using direct smear microscopy and PCR.

With respect to methodology, we did not always find a concordance between microscopy and PCR results. Similar results have been reported in other studies (1, 3, 8, 9, 10, 19). The direct smear method of detecting acid-fast bacilli has a sensitivity range of between 40 to 85% and a specificity of 67 to 100% (4, 8). The degree of sensitivity of PCR for the detection of pathogens within specimens has rendered it the method of choice for most studies (19). Where false positivity has been suspected with PCR, microscopy has been used to confirm the accuracy of positive PCR results in some cases (1, 9, 10). Generally, the power of detection of the bacterium of interest is increased by the use of both methods.

Our findings support the hypothesis that predaceous aquatic insects may play an important role in maintaining M. ulcerans within food webs in the aquatic environment (2). In this respect, external contamination of insect skeletons could also play a role because we have observed tadpoles grazing on the surfaces of predaceous water bugs in microcosm environments (H. Williamson, unpublished data). We also cannot rule out the possibility that belostomatids may be involved in mechanical transmission of M. ulcerans.

Finally, this work suggests that M. ulcerans can live as a commensal on belostomatids (2, 33). It is surprising that neither mosquito larvae nor belostomatids suffer developmental or behavioral defects as a result of M. ulcerans infection. Likewise, M. ulcerans appears to survive on the exoskeleton of belostomatids and within the guts of mosquito larvae for long periods of time. This close association between bacterium and insect, which is neither detrimental nor beneficial, is the hallmark of commensalism in nature.

 
We thank Ryan Kimbirauskas, Mollie Mcintosh, and Charles Quaye for assistance with insect collection in the field. We are grateful to Lindsay Hartsell and Marina Sligh for assistance with the maintenance of insects in the lab. We thank Lalita Ramakrishnana for the kind gift of the strain M. ulcerans 1615 GFP and the mycolactone-negative strain M. ulcerans 1615::TN118 GFP. We thank Juan Luis Jurat-Fuentes for technical assistance with insect dissection. We also thank John Dunlap for technical assistance with scanning electron microscopy analysis.

This research was supported in part by the NIH National Institute of Allergy and Infectious Diseases Grant Award 1 R03 AI026719-01A1 and the Division of International Training and Research Fogarty International Center, Ecology of Emerging Infectious Disease (EID) Grant Award 1 R01 TW007550 on Buruli ulcer disease ecology and disease transmission.

The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

 
* Corresponding author. Mailing address: Department of Microbiology, 409 Walters Life Sciences, University of Tennessee, Knoxville, TN 37996-0845. Phone: (865) 974-4042. Fax: (865) 974-4007. E-mail: psmall{at}utk.edu

Published ahead of print on 3 October 2008.

Supplemental material for this article may be found at http://aem.asm.org/.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Beige, J., J. Lokies, T. Schaberg, U. Finckh, M. Fischer, H. Mauch, H. Lode, B. Kohler, and A. Rolfs. 1995. Clinical evaluation of a Mycobacterium tuberculosis PCR assay. J. Clin. Microbiol. 33:90-95.[Abstract]
2
2. Benbow, M. E., H. Williamson, R. Kimbirauskas, M. D. McIntosh, R. Kolar, C. Quaye, F. Akpabey, D. Boakye, P. Small, and R. W. Merritt. 2008, posting date. Aquatic invertebrates as unlikely vectors of Buruli ulcer disease. Emerg. Infect. Dis. [Epub ahead of print.] doi:10.3201/eid1408.071503.
3
3. Chakravorty, S., M. Dudeja, M. Hanif, and J. S. Tyagi. 2005. Utility of universal sample processing methodology, combining smear microscopy, culture, and PCR, for diagnosis of pulmonary tuberculosis. J. Clin. Microbiol. 43:2703-2708.[Abstract/Free Full Text]
4
4. De Gentile, P. L., C. Mahaza, F. Rolland, B. Carbonnelle, J. L. Verret, and D. Chabasse. 1992. Cutaneous ulcer from Mycobacterium ulcerans. Apropos of 1 case in French Guiana. Bull. Soc. Pathol. Exot. 85:212-214. (In French.)[Medline]
5
5. Duker, A. A., F. Portaels, and M. Hale. 2006. Pathways of Mycobacterium ulcerans infection: a review. Environ. Int. 32:567-573.[CrossRef][Medline]
6
6. Eddyani, M., D. Ofori-Adjei, G. Teugels, D. De Weirdt, D. Boakye, W. M. Meyers, and F. Portaels. 2004. Potential role for fish in transmission of Mycobacterium ulcerans disease (Buruli ulcer): an environmental study. Appl. Environ. Microbiol. 70:5679-5681.[Abstract/Free Full Text]
7
7. George, K. M., D. Chatterjee, G. Gunawardana, D. Welty, J. Hayman, R. Lee, and P. L. Small. 1999. Mycolactone: a polyketide toxin from Mycobacterium ulcerans required for virulence. Science 283:854-857.[Abstract/Free Full Text]
8
8. Habeenzu, C., D. Lubasi, and A. F. Fleming. 1998. Improved sensitivity of direct microscopy for detection of acid-fast bacilli in sputum in developing countries. Trans. R. Soc. Trop. Med. Hyg. 92:415-416.[CrossRef][Medline]
9
9. Haldar, S., S. De Majumdar, S. Chakravorty, J. S. Tyagi, M. Bhalla, and M. K. Sen. 2005. Detection of acid-fast bacilli in postlysis debris of clinical specimens improves the reliability of PCR. J. Clin. Microbiol. 43:3580-3581.[Free Full Text]
10
10. Haldar, S., S. Chakravorty, M. Bhalla, S. De Majumdar, and J. S. Tyagi. 2007. Simplified detection of Mycobacterium tuberculosis in sputum using smear microscopy and PCR with molecular beacons. J. Med. Microbiol. 56:1356-1362.[Abstract/Free Full Text]
11
11. Johnson, P. D., T. Stinear, P. L. Small, G. Pluschke, R. W. Merritt, F. Portaels, K. Huygen, J. A. Hayman, and K. Asiedu. 2005. Buruli ulcer (M. ulcerans infection): new insights, new hope for disease control. PLoS Med. 2:e108.[CrossRef][Medline]
12
12. Käser, M., S. Rondini, M. Naegeli, T. Stinear, F. Portaels, U. Certa, and G. Pluschke. 2007. Evolution of two distinct phylogenetic lineages of the emerging human pathogen Mycobacterium ulcerans. BMC Evol. Biol. 7:177.[CrossRef][Medline]
13
13. Marsollier, L., R. Robert, J. Aubry, J. P. Saint Andre, H. Kouakou, P. Legras, A. L. Manceau, C. Mahaza, and B. Carbonnelle. 2002. Aquatic insects as a vector for Mycobacterium ulcerans. Appl. Environ. Microbiol. 68:4623-4628.[Abstract/Free Full Text]
14
14. Marsollier, L., T. Severin, J. Aubry, R. W. Merritt, J. P. Saint Andre, P. Legras, A. L. Manceau, A. Chauty, B. Carbonnelle, and S. T. Cole. 2004. Aquatic snails, passive hosts of Mycobacterium ulcerans. Appl. Environ. Microbiol. 70:6296-6298.[Abstract/Free Full Text]
15
15. Marsollier, L., J. Aubry, E. Coutanceau, J. P. Andre, P. L. Small, G. Milon, P. Legras, S. Guadagnini, B. Carbonnelle, and S. T. Cole. 2005. Colonization of the salivary glands of Naucoris cimicoides by Mycobacterium ulcerans requires host plasmatocytes and a macrolide toxin, mycolactone. Cell. Microbiol. 7:935-943.[CrossRef][Medline]
16
16. Marsollier, L., J. P. Andre, W. Frigui, G. Reysset, G. Milon, B. Carbonnelle, J. Aubry, and S. T. Cole. 2007. Early trafficking events of Mycobacterium ulcerans within Naucoris cimicoides. Cell. Microbiol. 9:347-355.[CrossRef][Medline]
17
17. Merritt, R. W., M. E. Benbow, and P. L. C. Small. 2005. Unraveling an emerging disease associated with disturbed aquatic environments: the case of Buruli ulcer. Front. Ecol. Environ. 3:323-331.[CrossRef]
18
18. Mitchell, P. J., I. V. Jerrett, and K. J. Slee. 1984. Skin ulcers caused by Mycobacterium ulcerans in koalas near Bairnsdale, Australia. Pathology 16:256-260.[CrossRef][Medline]
19
19. Pizarro, J. C., D. E. Lucero, and L. Stevens. 2007. PCR reveals significantly higher rates of Trypanosoma cruzi infection than microscopy in the Chagas vector, Triatoma infestans: high rates found in Chuquisaca, Bolivia. BMC Infect. Dis. 7:66.[CrossRef][Medline]
20
20. Portaels, F., P. Elsen, A. Guimaraes-Peres, P. A. Fonteyne, and W. M. Meyers. 1999. Insects in the transmission of Mycobacterium ulcerans infection. Lancet 353:986.[CrossRef][Medline]
21
21. Portaels, F., K. Chemlal, P. Elsen, P. D. Johnson, J. A. Hayman, J. Hibble, R. Kirkwood, and W. M. Meyers. 2001. Mycobacterium ulcerans in wild animals. Rev. Sci. Tech. 20:252-264.[Medline]
22
22. Portaels, F., W. M. Meyers, A. Ablordey, A. G. Castro, K. Chemlal, P. de Rijk, P. Elsen, K. Fissette, A. G. Fraga, R. Lee, E. Mahrous, P. L. Small, P. Stragier, E. Torrado, A. Van Aerde, M. T. Silva, and J. Pedrosa. 2008. First cultivation and characterization of Mycobacterium ulcerans from the environment. PLoS Negl. Trop. Dis. 2:e178.[CrossRef]
23
23. Raghunathan, P. L., E. A. Whitney, K. Asamoa, Y. Stienstra, T. H. Taylor, Jr., G. K. Amofah, D. Ofori-Adjei, K. Dobos, J. Guarner, S. Martin, S. Pathak, E. Klutse, S. Etuaful, W. T. van der Graaf, T. S. van der Werf, C. H. King, J. W. Tappero, and D. A. Ashford. 2005. Risk factors for Buruli ulcer disease (Mycobacterium ulcerans infection): results from a case-control study in Ghana. Clin. Infect. Dis. 40:1445-1453.[CrossRef][Medline]
24
24. Silva, M. T., F. Portaels, and J. Pedrosa. 2007. Aquatic insects and Mycobacterium ulcerans: an association relevant to Buruli ulcer control? PLoS Med. 4:e63.[CrossRef][Medline]
25
25. Stienstra, Y., W. T. van der Graaf, G. J. te Meerman, T. H. The, L. F. de Leij, and T. S. van der Werf. 2001. Susceptibility to development of Mycobacterium ulcerans disease: review of possible risk factors. Trop. Med. Int. Health 6:554-562.[CrossRef][Medline]
26
26. Stinear, T., J. K. Davies, G. A. Jenkin, J. A. Hayman, F. Oppedisano, and P. D. Johnson. 2000. Identification of Mycobacterium ulcerans in the environment from regions in Southeast Australia in which it is endemic with sequence capture-PCR. Appl. Environ. Microbiol. 66:3206-3213.[Abstract/Free Full Text]
27
27. Stinear, T. P., A. Mve-Obiang, P. L. Small, W. Frigui, M. J. Pryor, R. Brosch, G. A. Jenkin, P. D. Johnson, J. K. Davies, R. E. Lee, S. Adusumilli, T. Garnier, S. F. Haydock, P. F. Leadlay, and S. T. Cole. 2004. Giant plasmid-encoded polyketide synthases produce the macrolide toxin of Mycobacterium ulcerans. Proc. Natl. Acad. Sci. USA 101:1345-1349.[Abstract/Free Full Text]
28
28. Swart, C. C., and B. E. Felgenhauer. 2003. Structure and function of the mouthparts and salivary gland complex of the giant waterbug, Belostoma lutarium (Stål) (Hemiptera: Belostomatidae). Ann. Entomol. Soc. Am. 96:870-882.[CrossRef]
29
29. Swart, C. C., L. E. Deaton, and B. E. Felgenhauer. 2006. The salivary gland and salivary enzymes of the giant waterbugs (Heteroptera; Belostomatidae). Comp. Biochem. Physiol. A 145:114-122.[CrossRef][Medline]
30
30. Valdivia, R. H., A. E. Hromockyj, D. Monack, L. Ramakrishnan, and S. Falkow. 1996. Applications for green fluorescent protein (GFP) in the study of host-pathogen interactions. Gene 173:47-52.[CrossRef][Medline]
31
31. van der Werf, T. S., W. T. van der Graaf, D. G. Groothuis, and A. J. Knell. 1989. Mycobacterium ulcerans infection in Ashanti region, Ghana. Trans. R. Soc. Trop. Med. Hyg. 83:410-413.[CrossRef][Medline]
32
32. van der Werf, T. S., Y. Stienstra, R. C. Johnson, R. Phillips, O. Adjei, B. Fleischer, M. H. Wansbrough-Jones, P. D. Johnson, F. Portaels, W. T. van der Graaf, and K. Asiedu. 2005. Mycobacterium ulcerans disease. Bull. World Health Organ. 83:785-791.[Medline]
33
33. Williamson, H. R., M. E. Benbow, K. D. Nguyen, D. C. Beachboard, R. K. Kimbirauskas, M. D. McIntosh, C. Quaye, E. O. Ampadu, D. Boakye, R. W. Merritt, and P. L. Small. 2008. Distribution of Mycobacterium ulcerans in Buruli ulcer endemic and non-endemic aquatic sites in Ghana. PLoS Negl. Trop. Dis. 2:e205.[CrossRef]
34
34. World Health Organization. 2001. Buruli ulcer: diagnosis of Mycobacterium ulcerans disease. World Health Organization, Geneva, Switzerland.

---

Applied and Environmental Microbiology, November 2008, p. 7036-7042, Vol. 74, No. 22
0099-2240/08/$08.00+0     doi:10.1128/AEM.01234-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Supplemental material Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Mosi, L. Articles by Small, P. L. C. Search for Related Content PubMed PubMed Citation Articles by Mosi, L. Articles by Small, P. L. C. Agricola Articles by Mosi, L. Articles by Small, P. L. C.