- Email: [email protected]
Colonization of the Guts of Germ-Free Desert Locusts,Schistocerca Gregaria,by the BacteriumPantoea agglomerans
JOBNAME: 67#2 96 PAGE: 1 SESS: 22 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu JOURNAL OF INVERTEBRATE PATHOLOGY ARTICLE NO.
67, 11–14 (1996)
0002
Colonization of the Guts of Germ-Free Desert Locusts, Schistocerca Gregaria, by the Bacterium Pantoea agglomerans R. J. DILLON*
AND
A. K. CHARNLEY†
*Department of Entomology, Natural History Museum, Cromwell Road, London, SW7 5BD, United Kingdom; and †Microbial Pathogenicity Group, School of Biology and Biochemistry, University of Bath, Claverton Down, Bath, Avon, BA2- 7AY, United Kingdom Received May 24, 1995; accepted August 2, 1995
unknown. Knowledge of these processes and the mechanisms controlling them is important for an understanding of the colonization resistance of the gut toward potentially entomopathogenic microorganisms. The objective of this study was to follow the colonization of the bacterium Pantoea (4Enterobacter) agglomeransas a monoassociate in the gut of germ-free desert locusts, S. gregaria. P. agglomerans is a prominent member of the hindgut flora of the locust (Hunt and Charnley, 1981) and has been implicated in the production of antifungal phenols present in the guts of conventional but not germ-free insects (Dillon and Charnley, 1986a, 1988). This work formed a prelude to an investigation of the production of toxic phenols in monoassociated locusts which is reported elsewhere (Dillon and Charnley, 1995).
The aim of this study was to observe the colonization of the guts of germ-free desert locusts, Schistocerca gregaria, by Pantoea agglomerans, the dominant bacterial species found in the mature adult. P. agglomerans was established as a monoculture. During the first generation of the monoassociated locusts the bacterial population remained at a low level despite daily inoculations from Days 32–46. By the second generation, without additional inoculations of P. agglomerans, some insects had a large monoflora. However, there was a wide variation, indicating that the bacteria had successfully colonized some insects but not others. In contrast colonization of conventionally reared insects was rapid and occurred within days of hatching. The possible reasons for the difficulty in establishing P. agglomerans in the guts of germ-free locusts are discussed. © 1996 Academic Press, Inc. KEY WORDS: locusts; Schistocerca gregaria; Pantoea agglomerans.
MATERIALS AND METHODS
Insect Rearing Conventional and germ-free desert locusts were reared as described previously (Charnley et al., 1985). Germ-free hatchlings were produced for the monoassociation experiments by surface sterilizing the eggs with peracetic acid. The insects were placed in a stainless steel cage within a sterile flexible plastic isolator inflated under positive pressure with a constant flow of filter-sterilized air. A double door system facilitated the entrance of additional food and water. Insects were reared on a sterile diet of freeze-dried bran containing a vitamin supplement (administered dry) and freezedried grass, soaked in water. Control insects were reared in a nonsterile environment (in the same constant temperature room as the stock culture of locusts) on the same diet as the germ-free insects. They are termed ‘‘conventional insects.’’
INTRODUCTION
In common with many other insects (Brooks, 1963; Lysenko, 1981), microbial colonization of the locust gut appears to be due to fortuitous contamination from the insect’s surroundings and its food (Hunt and Charnley, 1981). Thus the gut flora is reflective of the environment. There have been numerous investigations of the bacterial flora of insect guts. Some studies have been from an environmental health perspective viz. possible transmission of human pathogens by insects (e.g., Greenberg, 1960; Greenberg et al., 1970), others have catalogued the incidence of entomopathogens (see review by Lysenko (1981)). The contribution of gut microbiota to nutrition (e.g., Hagen, 1966) and disease suppression (e.g., Dillon and Charnley, 1986a,b, 1988, 1991) have also been a focus for attention. However, much of the literature describes what may be termed the climax or mature community within the gut. The process of colonization and succession of the gut microflora leading to the climax community are practically
Administration of the Bacterial Inoculum P. agglomerans (originally isolated from the gut of S. gregaria in our laboratory) was grown from a single 11 0022-2011/96 $18.00 Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
JOBNAME: 67#2 96 PAGE: 2 SESS: 21 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu 12
DILLON AND CHARNLEY
colony in nutrient broth (Unipath Ltd.) for 7 hr at 30°C. After being centrifuged (3000g for 15 min) and washed in quarter strength Ringer (Unipath Ltd.) the bacteria were suspended in Ringer to a concentration of 1 × 107 ml−1 (determination by viable count). The suspension was introduced into the isolator in a surface-sterilized Universal bottle and mixed with the freeze-dried grass prior to feeding the hatchlings. Inoculated grass was given initially to 6-day-old insects and then daily for 14 days after the insects became adults. No inoculations were made to the food given the conventional insects. Enumeration of Bacteria in the Locust Gut Locusts were selected randomly prior to the morning feed when the gut was empty and kept at 4°C for 15 min prior to handling to prevent regurgitation. The cervical membrane and ventral nerve cord were cut and the whole gut was removed from the body after dissection under aseptic conditions. The gut was weighed and homogenized in Ringer solution for 2 min in a 1-ml-volume glass conical tube with a tapered teflon homogenizer to obtain a uniform suspension free of pieces of gut tissue. Serial dilutions of the homogenate were inoculated onto duplicate nutrient agar (Unipath Ltd.) plates. The inoculated agar was incubated aerobically for 24–36 hr at 30°C, then the colonies were counted and number of colony-forming units were expressed per milligrams wet weight of locust gut. Homogenates from two guts were used to obtain the results on the first day of sampling for each point on the graphs (Figs. 1a and 1b). The remaining results were derived from one locust gut (duplicate counts) for each individual graph point. Identification of Bacteria P. agglomerans was identified by the API 20E system (bioMérieux UK Ltd., Hampshire, UK). Bacteria
were isolated from locust guts on a number of media under aerobic and anaerobic conditions (Charnley et al., 1985) and identified to ensure that a monoculture of P. agglomerans had been established. RESULTS
A sterile isolator was set up with germ-free first instar larvae and the diet was inoculated with P. agglomerans. Only P. agglomerans was detected in the isolator and its insects during the course of the experiment. Although some of the insects had bacteria in their guts during the larval stages, bacteria were not regularly detected in gut samples up to the molt to adult (Fig. 1a). The reason for the low level of colonization is not obvious. Samples from the food dish and water fountain confirmed that P. agglomerans was present consistently in a viable form in the cage. It seemed possible that under natural conditions the resident flora in the conventional locust gut may require regular inocula to maintain numbers, and this would be provided under natural conditions by P. agglomerans in the epiflora of the plant food. In an attempt to mimic the natural situation the insects were fed daily from the time when most of the population had molted to adult (Day 32) for 14 days with axenic grass diet inoculated with P. agglomerans. Isolations from the guts of locusts on Day 53 (7 days after the end of the period of daily inoculation) revealed populations of 606 ± 414 cfu mg−1 of P. agglomerans. This was much greater than at Day 32, prior to the start of the daily inoculation, when there were populations of <2 cfu mg−1, but significantly lower than in conventional insects at a similar stage which had total bacterial populations of 34,967.6 ± 15,161.7 cfu mg−1 (Table 1). Furthermore, the gut bacterial population of conventional locusts was established very quickly (within a few days of hatching) and
FIG. 1. (a) Bacterial colonization of the guts of germ-free locusts over time (given as the age of the insects in days). Each point represents the concentration of bacteria (Pantoea agglomerans) in one insect as colony forming units (cfu) per milligram wet weight of gut, apart from the first day of sampling when two insects were used per data point. The line indicates median total bacterial population levels. The insects became adult around Day 32 (arrow). Diet was inoculated with bacteria and fed to 6-day-old insects. Additional inoculations were made daily from Days 32–46. (b) Colonization of the digestive tract of conventional locusts by bacteria. Details as in a except that no inoculations were carried out.
JOBNAME: 67#2 96 PAGE: 3 SESS: 22 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu BACTERIAL COLONIZATION OF THE LOCUST GUT
TABLE 1 Viable Bacterial Counts in the Gut of Gnotobiotic and Conventional Locusts Microbial association a
Monoassociation Monoassociation (2nd generation) Conventionalb
Age of insect (days)
Mean total viable counts ± SE CFU mg−1
N
53
606.7 ± 414.0
8
53 50
10419.0 ± 3947.0 34967.6 ± 15161.7
12 8
a Monoassociated, first generation, additional inoculations of Pantoea agglomerans made to the food during the first 14 days of adult life (see Fig. 1a). b Conventional, locusts with a normal gut flora (see Fig. 1b). Counts made from nutrient agar cultures incubated at 30°C for 24– 36 hr. Analysis of Variance: F 4 4.26, P 4 0.025. Multiple range analysis showed that the mean count from conventional locusts was significantly different from the count recorded for the generation 1 and 2 monoassociated insects. Generation 1 and 2 means were not significantly different.
remained consistently high throughout nymphal and early adult life (Fig. 1b). The large standard error associated with these results is a reflection of the wide variation in the bacterial population recorded in locusts and grasshoppers (see e.g., Bucher, 1959; Hunt and Charnley, 1981). Ten of the monoassociated insects were allowed to reproduce and their progeny were left to develop to adulthood in the isolator. There were no further inoculations of P. agglomerans and the isolator was maintained under otherwise germ-free conditions. Isolations from second generation monoassociated insects revealed large bacterial populations with a mean of 10,419 cfu mg−1 at 53 days (see Table 1 for summary), but these were still significantly lower than populations found in conventional insects. P. agglomerans was the sole organism isolated from the guts of second generation monoassociated insects. DISCUSSION
Inoculation of the diet of germ-free locusts with P. agglomerans, the predominant bacterial species found in the mature insect, did not result in the rapid colonization of the digestive tract. Studies on germ-free vertebrates show that bacteria often colonize in higher numbers when monoassociated due to the absence of interspecific competition (Coates and Fuller, 1977); this was not the case with first generation monoassociated locusts. Successful colonization is achieved when the bacterial population is stable in size over time without the need for periodic reintroduction by repeated oral doses (Freter, 1992). This clearly had not occurred in the first generation. Even daily inoculations of large numbers of bacteria failed to achieve a significant population of P. agglomerans. However, in the second generation without further inoculations,
13
some insects possessed substantial populations of P. agglomerans although the overall mean was not significantly different from the first generation. The presence of large populations in some of the second generation seems to imply ‘‘adaptation’’ on the part of the bacteria. There are examples in the vertebrate literature which support such a concept. Duval-Iflah et al. (1981) found that plasmid-free strains of Escherichia coli inhibited the establishment of plasmid-bearing strains in gnotobiotic mice (with a defined flora). The outcome, however, depended on the order of inoculation. The plasmid-free strain no longer exerted a barrier effect against the derivative plasmid-bearing strain when the former was inoculated 1 week after the latter. Maintaining the plasmid-free strain in monoassociation with mice resulted in adaptation of the bacteria to their host, since the ‘‘adapted’’ strain exerted the same barrier effect regardless of whether it was introduced before or after its plasmid-bearing derivative. Their ecological advantage disappeared when the adapted strain was cultured in broth. In the present case it is possible that successive in vitro transfers of the strain of P. agglomerans reduced its fitness for the locust gut which was restored by in vivo passage. The problem may have been exacerbated by the use of a single colony for the initial inoculum. The basis of this adaptation remains to be established. An alternative explanation for the colonization resistance of the germ-free gut toward the P. agglomerans is that ‘‘pioneer’’ species of bacteria are responsible for the sequential establishment of bacteria in the conventional insect. Pioneer Clostridium species released a growth promotor in gnotobiotic mice which permitted the development of a strain of E. coli (Ducluzeau et al., 1986). Similarly cellulolytic bacteria were dependent on the prior establishment by other bacteria in gnotobiotic lambs (Fonty et al., 1983). Experiments to establish a gut flora in germ-free locusts using several species of bacteria might reveal whether P. agglomerans requires the presence of other species for rapid colonization. ACKNOWLEDGMENT The work was supported by Research Grant AG 86/35 from the Agricultural and Food Research Council. REFERENCES Brooks, M. A. 1963. The microorganisms of healthy insects. In ‘‘Insect Pathology’’ (E. A. Steinhaus, Ed.), Vol. 1., pp. 215–250. Academic Press, New York. Bucher, G. E. 1959. Bacteria of grasshoppers of Western Canada: III. Frequency of occurrence, pathogenicity. J. Insect Pathol. 1, 391– 405. Charnley, A. K., Hunt, J., and Dillon, R. J. 1985. The germ free culture of desert locusts Schistocerca gregaria. J. Insect Physiol. 31, 477–485. Coates, M. E., and Fuller, R. 1977. The gnotobiotic animal in the study of gut microbiology. In ‘‘Microbial Ecology of the Gut’’
JOBNAME: 67#2 96 PAGE: 4 SESS: 22 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu 14
DILLON AND CHARNLEY
(R. T. J. Clarke and T. Bauchop, Eds.), pp. 311–346. Academic Press, New York. Dillon, R. J., and Charnley, A. K. 1986a. Inhibition of Metarhizium anisopliae by the gut bacterial flora of the desert locust Schistocerca gregaria: Evidence for an antifungal toxin. J. Invertebr. Pathol. 47, 350–360. Dillon, R. J., and Charnley, A. K. 1986b. Invasion of the pathogenic fungus Metarhizium anisopliae through the guts of germ-free locusts, Schistocerca gregaria. Mycopathologia 96, 59–66. Dillon, R. J., and Charnley, A. K. 1988. Inhibition of Metarhizium anisopliae by the gut bacterial flora of the desert locust: Characterisation of antifungal toxins. Can. J. Microbiol. 34, 1075–1082. Dillon, R. J., and Charnley, A. K. 1991. The fate of fungal spores in the insect gut. In ‘‘The Fungal Spore in Disease Initiation in Plants and Animals’’. (G. T. Cole and H. C. Hoch, Eds.), pp. 129–156. Plenum, New York. Dillon, R. J., and Charnley, A. K. 1995. Chemical barriers to gut infection in the desert locust: In vivo production of antimicrobial phenols associated with the bacterium Pantoea agglomerans. J. Invertebr. Pathol. 66, 72–75. Ducluzeau, R., Ladiré, M., and Raibaud, P. 1986. Implantation d’un mutant de Escherichia coli exigeant en acide diaminopimélique dans le tube digestif de souris gnotoxéniques. Ann. Microbiol. (Inst. Pasteur) 137A, 79–87. Duval-Iflah, Y., Raibaud, P., and Rousseau, M. 1981. Antagonisms among isogenic strains of Escherichia coli in the digestive tracts of gnotobiotic mice. Infect. Immun. 34, 957–969. Fonty, G., Gouet, P., Jouany, J. P., and Senaud, J. 1983. Ecological
factors determining establishment of cellulolytic bacteria and protozoa in the rumen of meroxenic lambs. J. Gen. Microbiol. 129, 213–223. Freter, R. 1992. Factors affecting the microecology of the gut. In ‘‘Probiotics: The Scientific Basis’’ (R. Fuller, Ed.), pp. 111–145. Chapman and Hall, London. Greenberg, B. 1960. Host-contaminant biology of Muscoid Flies: 1. Bacterial survival in the pre-adult stages and adults of four species of blow flies. J. Insect. Pathol. 2, 44–54. Greenberg, B., Kowalski, J. A., and Klowden, M. J. 1970. Factors affecting the transmission of Salmonella by flies: Natural resistance to colonization and bacterial interference. Infect. Immun. 2, 800–809. Hagen, K. S. 1966. Dependence of the olive fly, Dacus oleae, larvae on symbiosis with Pseudomonas savastanoi for the utilization of olive. Nature 209, 423–424. Hunt, J., and Charnley, A. K. 1981. Abundance and distribution of the gut flora of the desert locust, Schistocerca gregaria. J. Invertebr. Pathol. 38, 378–385. Lysenko, O. 1981. Principles of pathogenesis of insect bacterial diseases as exemplified by the nonsporeforming bacteria. In ‘‘Pathogenesis of Invertebrate Microbial Diseases’’ (E. W. Davidson, ed.), pp. 163–188. Allanheld Osmun, Monclair. Lysenko, O. 1985. Non-sporeforming bacteria pathogenic to insects: Incidence and mechanisms. Ann. Rev. Microbiol. 39, 673–695. Tanada, Y., and Kaya, H. J. 1993. “Insect Pathology.” Academic Press, San Diego.
67, 11–14 (1996)
0002
Colonization of the Guts of Germ-Free Desert Locusts, Schistocerca Gregaria, by the Bacterium Pantoea agglomerans R. J. DILLON*
AND
A. K. CHARNLEY†
*Department of Entomology, Natural History Museum, Cromwell Road, London, SW7 5BD, United Kingdom; and †Microbial Pathogenicity Group, School of Biology and Biochemistry, University of Bath, Claverton Down, Bath, Avon, BA2- 7AY, United Kingdom Received May 24, 1995; accepted August 2, 1995
unknown. Knowledge of these processes and the mechanisms controlling them is important for an understanding of the colonization resistance of the gut toward potentially entomopathogenic microorganisms. The objective of this study was to follow the colonization of the bacterium Pantoea (4Enterobacter) agglomeransas a monoassociate in the gut of germ-free desert locusts, S. gregaria. P. agglomerans is a prominent member of the hindgut flora of the locust (Hunt and Charnley, 1981) and has been implicated in the production of antifungal phenols present in the guts of conventional but not germ-free insects (Dillon and Charnley, 1986a, 1988). This work formed a prelude to an investigation of the production of toxic phenols in monoassociated locusts which is reported elsewhere (Dillon and Charnley, 1995).
The aim of this study was to observe the colonization of the guts of germ-free desert locusts, Schistocerca gregaria, by Pantoea agglomerans, the dominant bacterial species found in the mature adult. P. agglomerans was established as a monoculture. During the first generation of the monoassociated locusts the bacterial population remained at a low level despite daily inoculations from Days 32–46. By the second generation, without additional inoculations of P. agglomerans, some insects had a large monoflora. However, there was a wide variation, indicating that the bacteria had successfully colonized some insects but not others. In contrast colonization of conventionally reared insects was rapid and occurred within days of hatching. The possible reasons for the difficulty in establishing P. agglomerans in the guts of germ-free locusts are discussed. © 1996 Academic Press, Inc. KEY WORDS: locusts; Schistocerca gregaria; Pantoea agglomerans.
MATERIALS AND METHODS
Insect Rearing Conventional and germ-free desert locusts were reared as described previously (Charnley et al., 1985). Germ-free hatchlings were produced for the monoassociation experiments by surface sterilizing the eggs with peracetic acid. The insects were placed in a stainless steel cage within a sterile flexible plastic isolator inflated under positive pressure with a constant flow of filter-sterilized air. A double door system facilitated the entrance of additional food and water. Insects were reared on a sterile diet of freeze-dried bran containing a vitamin supplement (administered dry) and freezedried grass, soaked in water. Control insects were reared in a nonsterile environment (in the same constant temperature room as the stock culture of locusts) on the same diet as the germ-free insects. They are termed ‘‘conventional insects.’’
INTRODUCTION
In common with many other insects (Brooks, 1963; Lysenko, 1981), microbial colonization of the locust gut appears to be due to fortuitous contamination from the insect’s surroundings and its food (Hunt and Charnley, 1981). Thus the gut flora is reflective of the environment. There have been numerous investigations of the bacterial flora of insect guts. Some studies have been from an environmental health perspective viz. possible transmission of human pathogens by insects (e.g., Greenberg, 1960; Greenberg et al., 1970), others have catalogued the incidence of entomopathogens (see review by Lysenko (1981)). The contribution of gut microbiota to nutrition (e.g., Hagen, 1966) and disease suppression (e.g., Dillon and Charnley, 1986a,b, 1988, 1991) have also been a focus for attention. However, much of the literature describes what may be termed the climax or mature community within the gut. The process of colonization and succession of the gut microflora leading to the climax community are practically
Administration of the Bacterial Inoculum P. agglomerans (originally isolated from the gut of S. gregaria in our laboratory) was grown from a single 11 0022-2011/96 $18.00 Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
JOBNAME: 67#2 96 PAGE: 2 SESS: 21 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu 12
DILLON AND CHARNLEY
colony in nutrient broth (Unipath Ltd.) for 7 hr at 30°C. After being centrifuged (3000g for 15 min) and washed in quarter strength Ringer (Unipath Ltd.) the bacteria were suspended in Ringer to a concentration of 1 × 107 ml−1 (determination by viable count). The suspension was introduced into the isolator in a surface-sterilized Universal bottle and mixed with the freeze-dried grass prior to feeding the hatchlings. Inoculated grass was given initially to 6-day-old insects and then daily for 14 days after the insects became adults. No inoculations were made to the food given the conventional insects. Enumeration of Bacteria in the Locust Gut Locusts were selected randomly prior to the morning feed when the gut was empty and kept at 4°C for 15 min prior to handling to prevent regurgitation. The cervical membrane and ventral nerve cord were cut and the whole gut was removed from the body after dissection under aseptic conditions. The gut was weighed and homogenized in Ringer solution for 2 min in a 1-ml-volume glass conical tube with a tapered teflon homogenizer to obtain a uniform suspension free of pieces of gut tissue. Serial dilutions of the homogenate were inoculated onto duplicate nutrient agar (Unipath Ltd.) plates. The inoculated agar was incubated aerobically for 24–36 hr at 30°C, then the colonies were counted and number of colony-forming units were expressed per milligrams wet weight of locust gut. Homogenates from two guts were used to obtain the results on the first day of sampling for each point on the graphs (Figs. 1a and 1b). The remaining results were derived from one locust gut (duplicate counts) for each individual graph point. Identification of Bacteria P. agglomerans was identified by the API 20E system (bioMérieux UK Ltd., Hampshire, UK). Bacteria
were isolated from locust guts on a number of media under aerobic and anaerobic conditions (Charnley et al., 1985) and identified to ensure that a monoculture of P. agglomerans had been established. RESULTS
A sterile isolator was set up with germ-free first instar larvae and the diet was inoculated with P. agglomerans. Only P. agglomerans was detected in the isolator and its insects during the course of the experiment. Although some of the insects had bacteria in their guts during the larval stages, bacteria were not regularly detected in gut samples up to the molt to adult (Fig. 1a). The reason for the low level of colonization is not obvious. Samples from the food dish and water fountain confirmed that P. agglomerans was present consistently in a viable form in the cage. It seemed possible that under natural conditions the resident flora in the conventional locust gut may require regular inocula to maintain numbers, and this would be provided under natural conditions by P. agglomerans in the epiflora of the plant food. In an attempt to mimic the natural situation the insects were fed daily from the time when most of the population had molted to adult (Day 32) for 14 days with axenic grass diet inoculated with P. agglomerans. Isolations from the guts of locusts on Day 53 (7 days after the end of the period of daily inoculation) revealed populations of 606 ± 414 cfu mg−1 of P. agglomerans. This was much greater than at Day 32, prior to the start of the daily inoculation, when there were populations of <2 cfu mg−1, but significantly lower than in conventional insects at a similar stage which had total bacterial populations of 34,967.6 ± 15,161.7 cfu mg−1 (Table 1). Furthermore, the gut bacterial population of conventional locusts was established very quickly (within a few days of hatching) and
FIG. 1. (a) Bacterial colonization of the guts of germ-free locusts over time (given as the age of the insects in days). Each point represents the concentration of bacteria (Pantoea agglomerans) in one insect as colony forming units (cfu) per milligram wet weight of gut, apart from the first day of sampling when two insects were used per data point. The line indicates median total bacterial population levels. The insects became adult around Day 32 (arrow). Diet was inoculated with bacteria and fed to 6-day-old insects. Additional inoculations were made daily from Days 32–46. (b) Colonization of the digestive tract of conventional locusts by bacteria. Details as in a except that no inoculations were carried out.
JOBNAME: 67#2 96 PAGE: 3 SESS: 22 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu BACTERIAL COLONIZATION OF THE LOCUST GUT
TABLE 1 Viable Bacterial Counts in the Gut of Gnotobiotic and Conventional Locusts Microbial association a
Monoassociation Monoassociation (2nd generation) Conventionalb
Age of insect (days)
Mean total viable counts ± SE CFU mg−1
N
53
606.7 ± 414.0
8
53 50
10419.0 ± 3947.0 34967.6 ± 15161.7
12 8
a Monoassociated, first generation, additional inoculations of Pantoea agglomerans made to the food during the first 14 days of adult life (see Fig. 1a). b Conventional, locusts with a normal gut flora (see Fig. 1b). Counts made from nutrient agar cultures incubated at 30°C for 24– 36 hr. Analysis of Variance: F 4 4.26, P 4 0.025. Multiple range analysis showed that the mean count from conventional locusts was significantly different from the count recorded for the generation 1 and 2 monoassociated insects. Generation 1 and 2 means were not significantly different.
remained consistently high throughout nymphal and early adult life (Fig. 1b). The large standard error associated with these results is a reflection of the wide variation in the bacterial population recorded in locusts and grasshoppers (see e.g., Bucher, 1959; Hunt and Charnley, 1981). Ten of the monoassociated insects were allowed to reproduce and their progeny were left to develop to adulthood in the isolator. There were no further inoculations of P. agglomerans and the isolator was maintained under otherwise germ-free conditions. Isolations from second generation monoassociated insects revealed large bacterial populations with a mean of 10,419 cfu mg−1 at 53 days (see Table 1 for summary), but these were still significantly lower than populations found in conventional insects. P. agglomerans was the sole organism isolated from the guts of second generation monoassociated insects. DISCUSSION
Inoculation of the diet of germ-free locusts with P. agglomerans, the predominant bacterial species found in the mature insect, did not result in the rapid colonization of the digestive tract. Studies on germ-free vertebrates show that bacteria often colonize in higher numbers when monoassociated due to the absence of interspecific competition (Coates and Fuller, 1977); this was not the case with first generation monoassociated locusts. Successful colonization is achieved when the bacterial population is stable in size over time without the need for periodic reintroduction by repeated oral doses (Freter, 1992). This clearly had not occurred in the first generation. Even daily inoculations of large numbers of bacteria failed to achieve a significant population of P. agglomerans. However, in the second generation without further inoculations,
13
some insects possessed substantial populations of P. agglomerans although the overall mean was not significantly different from the first generation. The presence of large populations in some of the second generation seems to imply ‘‘adaptation’’ on the part of the bacteria. There are examples in the vertebrate literature which support such a concept. Duval-Iflah et al. (1981) found that plasmid-free strains of Escherichia coli inhibited the establishment of plasmid-bearing strains in gnotobiotic mice (with a defined flora). The outcome, however, depended on the order of inoculation. The plasmid-free strain no longer exerted a barrier effect against the derivative plasmid-bearing strain when the former was inoculated 1 week after the latter. Maintaining the plasmid-free strain in monoassociation with mice resulted in adaptation of the bacteria to their host, since the ‘‘adapted’’ strain exerted the same barrier effect regardless of whether it was introduced before or after its plasmid-bearing derivative. Their ecological advantage disappeared when the adapted strain was cultured in broth. In the present case it is possible that successive in vitro transfers of the strain of P. agglomerans reduced its fitness for the locust gut which was restored by in vivo passage. The problem may have been exacerbated by the use of a single colony for the initial inoculum. The basis of this adaptation remains to be established. An alternative explanation for the colonization resistance of the germ-free gut toward the P. agglomerans is that ‘‘pioneer’’ species of bacteria are responsible for the sequential establishment of bacteria in the conventional insect. Pioneer Clostridium species released a growth promotor in gnotobiotic mice which permitted the development of a strain of E. coli (Ducluzeau et al., 1986). Similarly cellulolytic bacteria were dependent on the prior establishment by other bacteria in gnotobiotic lambs (Fonty et al., 1983). Experiments to establish a gut flora in germ-free locusts using several species of bacteria might reveal whether P. agglomerans requires the presence of other species for rapid colonization. ACKNOWLEDGMENT The work was supported by Research Grant AG 86/35 from the Agricultural and Food Research Council. REFERENCES Brooks, M. A. 1963. The microorganisms of healthy insects. In ‘‘Insect Pathology’’ (E. A. Steinhaus, Ed.), Vol. 1., pp. 215–250. Academic Press, New York. Bucher, G. E. 1959. Bacteria of grasshoppers of Western Canada: III. Frequency of occurrence, pathogenicity. J. Insect Pathol. 1, 391– 405. Charnley, A. K., Hunt, J., and Dillon, R. J. 1985. The germ free culture of desert locusts Schistocerca gregaria. J. Insect Physiol. 31, 477–485. Coates, M. E., and Fuller, R. 1977. The gnotobiotic animal in the study of gut microbiology. In ‘‘Microbial Ecology of the Gut’’
JOBNAME: 67#2 96 PAGE: 4 SESS: 22 OUTPUT: Fri May 3 13:52:39 1996 /xypage/worksmart/tsp000/0365/22pu 14
DILLON AND CHARNLEY
(R. T. J. Clarke and T. Bauchop, Eds.), pp. 311–346. Academic Press, New York. Dillon, R. J., and Charnley, A. K. 1986a. Inhibition of Metarhizium anisopliae by the gut bacterial flora of the desert locust Schistocerca gregaria: Evidence for an antifungal toxin. J. Invertebr. Pathol. 47, 350–360. Dillon, R. J., and Charnley, A. K. 1986b. Invasion of the pathogenic fungus Metarhizium anisopliae through the guts of germ-free locusts, Schistocerca gregaria. Mycopathologia 96, 59–66. Dillon, R. J., and Charnley, A. K. 1988. Inhibition of Metarhizium anisopliae by the gut bacterial flora of the desert locust: Characterisation of antifungal toxins. Can. J. Microbiol. 34, 1075–1082. Dillon, R. J., and Charnley, A. K. 1991. The fate of fungal spores in the insect gut. In ‘‘The Fungal Spore in Disease Initiation in Plants and Animals’’. (G. T. Cole and H. C. Hoch, Eds.), pp. 129–156. Plenum, New York. Dillon, R. J., and Charnley, A. K. 1995. Chemical barriers to gut infection in the desert locust: In vivo production of antimicrobial phenols associated with the bacterium Pantoea agglomerans. J. Invertebr. Pathol. 66, 72–75. Ducluzeau, R., Ladiré, M., and Raibaud, P. 1986. Implantation d’un mutant de Escherichia coli exigeant en acide diaminopimélique dans le tube digestif de souris gnotoxéniques. Ann. Microbiol. (Inst. Pasteur) 137A, 79–87. Duval-Iflah, Y., Raibaud, P., and Rousseau, M. 1981. Antagonisms among isogenic strains of Escherichia coli in the digestive tracts of gnotobiotic mice. Infect. Immun. 34, 957–969. Fonty, G., Gouet, P., Jouany, J. P., and Senaud, J. 1983. Ecological
factors determining establishment of cellulolytic bacteria and protozoa in the rumen of meroxenic lambs. J. Gen. Microbiol. 129, 213–223. Freter, R. 1992. Factors affecting the microecology of the gut. In ‘‘Probiotics: The Scientific Basis’’ (R. Fuller, Ed.), pp. 111–145. Chapman and Hall, London. Greenberg, B. 1960. Host-contaminant biology of Muscoid Flies: 1. Bacterial survival in the pre-adult stages and adults of four species of blow flies. J. Insect. Pathol. 2, 44–54. Greenberg, B., Kowalski, J. A., and Klowden, M. J. 1970. Factors affecting the transmission of Salmonella by flies: Natural resistance to colonization and bacterial interference. Infect. Immun. 2, 800–809. Hagen, K. S. 1966. Dependence of the olive fly, Dacus oleae, larvae on symbiosis with Pseudomonas savastanoi for the utilization of olive. Nature 209, 423–424. Hunt, J., and Charnley, A. K. 1981. Abundance and distribution of the gut flora of the desert locust, Schistocerca gregaria. J. Invertebr. Pathol. 38, 378–385. Lysenko, O. 1981. Principles of pathogenesis of insect bacterial diseases as exemplified by the nonsporeforming bacteria. In ‘‘Pathogenesis of Invertebrate Microbial Diseases’’ (E. W. Davidson, ed.), pp. 163–188. Allanheld Osmun, Monclair. Lysenko, O. 1985. Non-sporeforming bacteria pathogenic to insects: Incidence and mechanisms. Ann. Rev. Microbiol. 39, 673–695. Tanada, Y., and Kaya, H. J. 1993. “Insect Pathology.” Academic Press, San Diego.