University of Montana ScholarWorks at University of Montana
Graduate Student Theses, Dissertations, & Professional Papers Graduate School
2006
Microbes Stoneflies and Fish: ophicT Interactions in Aquati Ecosystems
Sandra Adams The University of Montana
Follow this and additional works at: https://scholarworks.umt.edu/etd Let us know how access to this document benefits ou.y
Recommended Citation Adams, Sandra, "Microbes Stoneflies and Fish: ophicT Interactions in Aquati Ecosystems" (2006). Graduate Student Theses, Dissertations, & Professional Papers. 9301. https://scholarworks.umt.edu/etd/9301
This Thesis is brought to you for free and open access by the Graduate School at ScholarWorks at University of Montana. It has been accepted for inclusion in Graduate Student Theses, Dissertations, & Professional Papers by an authorized administrator of ScholarWorks at University of Montana. For more information, please contact [email protected]. Maureen and Mike MANSFIELD LIBRARY
The University of Montana
Permission is granted by the author to reproduce this material in its entirety, provided that this material is used for scholarly purposes and is properly cited in published works and reports.
**Please check "Yes" or "No" and provide signature**
Yes, I grant permission X _____
No, I do not grant permission ______
Author's Signature:
Date:
Any copying for commercial purposes or financial gain may be undertaken only with the author's explicit consent.
8/98
MICROBES, STONEFLIES, AND FISH: TROPHIC INTERACTIONS IN
AQUATIC ECOSYSTEMS
by
Sandra Adams
B.S. Washington State University, 2001
Presented in partial fulfillment of the requirements
for the degree of
Master of Science
The University of Montana
July 2006
Approved by:
Chairperson
Dean, Graduate School
c { - l - O f c Date UMI Number: EP72610
All rights reserved
INFORMATION TO ALL USERS The quality of this reproduction is dependent upon the quality of the copy submitted.
In the unlikely event that the author did not send a complete manuscript and there are missing pages, these will be noted. Also,- if material had to be removed, a note will indicate the deletion.
Dissertation P«b:hsri*ng
UMI EP72610 Published by ProQuest LLC (2015). Copyright in the Dissertation held by the Author. Microform Edition © ProQuest LLC. All rights reserved. This work is protected against unauthorized copying under Title 17, United States Code
ProQuest LLC. 789 East Eisenhower Parkway P.O. Box 1346 Ann Arbor, Ml 48106- 1346 Adams, Sandra M.S., July 2006 Microbial Ecology
Microbes, Stoneflies, and Fish: Trophic Interactions in Aquatic Ecosystems
Chairperson: Dr. William Holben
Specific interactions that occur between stoneflies and microbial communities in freshwater ecosystems are largely unknown. However, it is generally accepted that early instar nymphs feed on microbial biofilm, either by direct grazing or by the consumption of leaf litter. Molecular techniques were used to compare microbial communities contained in the hindguts of stoneflies and the sediment of the Nyack floodplain, West Glacier, MT. Data obtained through a molecular survey of both the sediment environment (4,312 sequences) and the hindgut contents of Isocapnia sp. and Paraperla sp. nymphs (120 sequences) revealed that a diverse microbial assemblage is present in the floodplain. It was discovered that a known fish pathogen, Aeromonas salmonicida subsp. salmonicida, was in high abundance in the hindgut contents of stoneflies (28% of sequences recovered) as compared to the frequency in the sediment (0.02%). Techniques were then developed to assay attraction and preferential feeding behavior to further understand the interactions of individual stonefly nymphs toward bacterial isolates. Bacterial isolates were chosen based on their abundance in the hindguts and sediment and carbon to nitrogen ratios (C:N) The results indicate that stoneflies are attracted to and preferentially fed on A. salmonicida. In addition, bacterial isolates that had C:N similar to that of stonefly nymphs were preferred. Because of the abundance of sequences found in the hindguts of stonefly nymphs and the preferential feeding behavior exhibited towardA. salmonicida, I then investigated whether stonefly nymphs can act as a reservoir and vector for A. salmonicida transfer to fish. Rainbow trout juveniles were presented with stonefly nymphs fed a type strain of A. salmonicida, stonefly nymphs harboring a natural population of A. salmonicida or, a commercial fish diet to determine the vectoring capacity of the nymphs. Both stonefly treatments resulted in expression of disease in the trout, and stonefly nymphs harboring the “natural” pathogen appeared to be more effective at transmitting disease than stoneflies fed the commercial type strain of A. salmonicida. Acknowledgments
I would like to thank my committee members, Dr. Frank Rosenzweig, Dr. Lisa Eby, Dr. Diana Six, Dr. Jim Gannon, and Dr. Bill Holben for all of their help and support. I would like to especially thank Dr. Holben for the space and freedom to study trophic interactions in a way that was not predicted or planned at the beginning of the project. I thank Nathan Gordon and Dr. Gannon for providing bacterial strains used in Chapters 2 and 3 .1 would also like to thank Alejandra Valenzuela and Dr. Gannon for C:N data included in Chapter 2. In addition, I would like to thank George Kirsch, The Jocko Fish Hatchery, and M.A. Stevenson for the trout and help in husbandry techniques used in Chapter 3. In addition, I would like to thank my lab mates for all of their help and availability to bounce ideas off of. Special thanks are in order to Linda Schimmelpfennig (for her help and expertise) and Tara Westlie (for her willingness to help with everything from codes to enrolling me in the chocolate program). Finally, I would like to thank my family for their support and understanding. Without the assistance I received from my husband in the field, lab, and the kitchen, this work would have been completed. And without the understanding of my children, that sometimes mommy just needs to write, this thesis would not have been finished. Table of Contents Abstract...... ii
Acknowledgements...... iii
Table of Contents...... iv
List of Tables...... v
List of Figures...... vi
Introduction...... 1
References...... 5
Chapter I Molecular Analysis of a Hyporheic Ecosystem: Investigation of Microbial Communities in the Sediment and Hindguts of Stonefly Nymphs...... 7 Abstract...... 7 Introduction...... :8 Materials and Methods...... 11 Results and Discussion...... 13 Conclusion...... 16 References...... 33
Chapter 2 Attraction and Feeding Behavior of Plecoptera Nymphs to Different Bacteria Isolates ...... 37 Abstract...... 37 Introduction...... 37 Materials and Methods...... 41 Results and Discussion...... 44 Conclusion...... 45 References...... 51
Chapter 3: The Deadly Meal: Bacterial pathogen transmission to rainbow trout via stoneflies...... 56 Summary...... 56 Conclusion...... 62 Methods...... 6!2 References...... 68
Summary ...... 72 References...... 74
iv List of Tables
Table 1: Phylogenie relatedness of sediment dwelling microbial community..! 1
Table 2: Clones obtained from stonefly hindguts...... 31
Table 1: Carbon and nitrogen ratios of organisms...... 50
v List of Figures
Chapter 1: Figure 1: Rank abundance curve of sediment microbial community...... 19
Figure 2: Rank/abundance curve of sequences obtained from the hindgut contents of stoneflies...... 20
Chapter 2: Figure 1: Example of nymph feeding behavior...... 47
Figure 2: Paraperla species attraction...... 48
Figure 3: Insect feeding behavior...... 49
Chapter 3: Figure 1: Classic symptom...... 66
Figure 2: Stonefly feeding behavior towards bacterial isolates...... 67 Introduction
Prokaryotes and macroinvertebrates are considered to play key roles in the cycling of nutrients in surface waters (Huryn and Wallace 2000, Feris et al. 2003,
Alexandre et al. 2005). Microorganisms exist primarily as biofilms in aquatic environments (Costertonet al. 1994). These biofilms represent the majority of microbial activity in surface waters (Weitereet al. 2005). Production of microbial biomass can impact production of macroinvertebrates (Huryn and Wallace 2000). In turn, the productivity of aquatic macroinvertebrates is limited by the productivity of lower trophic levels. Further, the productivity of macroinvertebrates can impact higher trophic levels such as fish, beavers, and bears (Huryn and Wallace 2000). Understanding the mechanisms that drive specific interactions between macroinvertebrates and the microbial loop in riverine systems is necessary to gain insight into how nutrients flow through trophic levels.
The relationships that exist between plecopteran nymphs and microorganisms in aquatic environments are only poorly understood. However, it is generally accepted that early instars of many stonefly species consume microbial biofilms in river sediments
(Cummin and Merritt 1996). Understanding trophic interactions between stonefly nymphs and microbial populations is necessary to gain insight in the mechanisms that control the flow and turnover of nutrients (e.g. carbon, nitrogen, and phosphorous).
However, the biology of microorganisms makes understanding specific mechanisms that drive these interactions difficult. Many are fastidious, difficult to enumerate, and not easily maintained in the laboratory (Schloss and Handelsman 2004).
1 To overcome these difficulties, I combined traditional insect attraction techniques and molecular methods in three studies designed to understand specific interactions between stonefly and microbial populations in a pristine floodplain. In the first study,
“Molecular Analysis of a Hyporheic Ecosystem: Investigation of Microbial Communities in the Sediment and Hindguts of Stonefly Nymphs”, I hypothesized that the two microbial communities would contain similar members and that the hindgut-dwelling communities would represent a subset of the sediment community.
To test this hypothesis, I compared the microbial communities inhabiting the sediment of the Nyack floodplain, West Glacier, MT with the microbial communities found in the hindguts of stoneflies also dwelling in the region. I analyzed a massive phylogenetic survey of the open sediment environment that was created using generally- conserved primers for a region of the 16S rRNA gene. To determine the extent to which the sediment microbial community was represented in the hindguts of stoneflies, I performed a focused 16s rRNA survey of that compartment. The sediment survey revealed that a diverse microbial assemblage is present in the Nyack floodplain. The results of the stonefly hindgut sequencing showed that a diverse microbial assemblage is also present in this compartment. However, there was only one sequence that was found in both the sediment and the hindgut. The sequence that was present in both samples had an Sab of 1 000 (indicating an exact match) to that of a known fish pathogen, Aeromonas salmonicida subsp.salmonicida . This sequence was encountered in relatively high abundance in the hindgut contents of stoneflies (33% of individuals sampled) compared to the frequency in the sediment (0.02%).
2 Based on the results obtained from the first study, I hypothesized that stonefly nymphs would be attracted to and preferentially feed upon Aeromonas salmonicida subsp.salmonicida. For the study, “Attraction and Feeding Behavior of Plecopteran
Nymphs to Different Bacterial Isolates”, an aquatic Y-olfactometer was developed to determine choice preference of Paraperla sp.toward three bacterial isolates in head-to- head competitions. In addition, a choice test arena was developed to quantify feeding behavior ofIsocapnia sp.when presented with different microorganisms in discrete biofilms. The results indicated that these stoneflies were attracted to, and preferentially fed on A. salmonicida significantly more than most other isolates.
Aeromonas salmonicida is a known pathogen, capable of causing mortality in salmonid fish. Because A. salmonicida sequences were abundant in the hindguts of stonefly nymphs and because stoneflies were attracted to A. salmonicida, I hypothesized that stonefly nymphs would act as a reservoir and disease vector for transmission of this pathogen. Viable A. salmonicida were isolated from both hindgut and frass from nymphs fed A. salmonicida, indicating that stoneflies could act as a transmission vector for this bacterium. To test whether stoneflies transmit the pathogen to fish, I presented juvenile rainbow trout (Oncorhynchus mykiss) with stonefly nymphs fed a type strain of A. salmonicida, stonefly nymphs harboring A. salmonicida at natural frequencies, or commercial fish diet. Both stonefly treatments resulted in disease expression and stonefly nymphs harboring the “natural” pathogen appeared to be more effective at transmitting disease than stoneflies fed the type strain of A. salmonicida: In addition, natural strains, contained in the hindguts, appeared to be more virulent than the type
strain.
3 Collectively, the findings of these three studies (phylogenetic survey, preferred feeding, and disease transmission) suggest for the first time that stonefly nymphs and the fish pathogen Aeromonas salmonicida subsp.salmonicida are closely associated in freshwater environments and that the nymphs are capable of vectoring this pathogen to host fish. This is significant because, while it is well documented that A. salmonicida is often found in low abundance in open aquatic environments (Hineyet al 1992, Austin
1997, Smith et al. 2003), it has never been previously suggested that plecopteran nymphs can act as a pathogen reservoir or transmission vector to susceptible fish.
4 References
1. Alexandre, G., Greer-Phillips, S., and Zhulin, I.B. 2004. Ecological role of energy
taxis in microorganisms. FEMS Micro Rev. 28:113-126.
2. Austin, B. 1997. Progress in understanding the fish pathogen Aeromonas
salmonicida. Mar Biotechnol 15: 131-134.
3. Costerton, J.W., Lewandowski, Z., DeBeer, D., Caldwell, D., Korber, D., and
James, G. 1994. Biofilms, the customized microniche. Journal o f Bacteriology.
176: 2137-2142.
4. Cummins, K. W., and R. W. Merritt. 1996. Ecology and Distribution of Aquatic
Insects. Pages 74-86 in K. W. Cummins and R. W. Merritt, editors. Aquatic
Insects of North America. Kendall/Hunt Publishing Company, Dubuque.
5. Feris, K.P., Ramsey, P.W., Frazar, C., Rillig, M., Moore, J.N., Gannon, J.E., and
Holben, W.E. 2004. Seasonal dynamics of shallow-hyporheic-zone microbial
community structure along a heavy-metal contamination gradient. Appl Environ
Microbio. 70:2323-2331.
6. Hiney, M., D. Dawson, D. Heery, P. R. Smith, F. Gannon, and R. Powell. 1992.
DNA Probe for Aeromonas salmonicida. Appl Environ Microbiol 58:1039-1042.
7. Huryn, A. D., and Wallace, J. B. 2000. Life history and production of stream
insects. Annu Rev Entomol 45:83-110.
8. Smith, P., Hiney, M., Gilroy, D. Padley, D., Gallagher, T., Cotter, D., and Powell,
R. 2003. Confusion generated by the validation of the application of an enzyme
linked immunosorbent assay and a polymerase chain reaction method for the
detection of Aeromonas salmonicida in field samples. Aquaculture 218:67-79.
5 9. Schloss, P.D., and Handelsman. J, 2004. Status of the microbial census.
Microbiol. Mol. Biol. Rev. 68:686-691.
10. Weitere, M., Bergfeld, T., Rice, S. A., Matz, C., and Kjelleberg, S. 2005. Grazing
resistance of Pseudomonas aeruginosa biofilms depends on type of protective
mechanism, developmental stage and protozoan feeding mode. Environmental
Microbiology. 7: 1593-1601.
6 Chapter 1
Molecular Analysis of a Hyporheic Ecosystem: Investigation of Microbial
Communities in the Sediment and Hindguts of Stonefly Nymphs
Abstract
Microorganisms are thought to be efficient at nutrient uptake and thus form the energy base of aquatic ecosystems (Ellis et al. 1998, Craft et al. 2002, Knapp et al. 2005). In hyporheic ecosystems, microbial production has been suggested to be driven by dissolved organic nutrients and may be orders of magnitude greater than in the stream channel (Stanford and Ward 1993, Ellis et al. 1998, Craft et al. 2002). The Nyack floodplain, West Glacier, MT is a pristine hyporheic zone, in which at least 70 taxa of invertebrates comprise an extensive food web (Stanford and Ward 1988, Stanford and Ward 1993, Craft et al. 2002). Plecopteran nymphs (stoneflies) are found in abundance in the groundwater surrounding the streambed and are thought to be important in both the groundwater and surface water food webs (Stanford and Ward 1993). Detailed knowledge of trophic interactions is required in order to understand the mechanisms that control flow and nutrient turnover in aquatic ecosystems (Huryn and Wallace 2000). Microorganisms play key roles in these processes and are assumed to be able to quickly assimilate dissolved organic nutrients (Knappet al. 2005). Because many microorganisms are difficult to enumerate, culture, and identify, understanding specific mechanisms that drive trophic interactions between microbes and other levels is difficult. Our approach to gaining a greater understanding of the processes that drive the flow of nutrients in aquatic ecosystems was to investigate and compare microbial communities inhabiting the sediment and hindguts of stonefly nymphs hyporheic ecosystem. Molecular techniques were utilized to assess prokaryotic diversity in the floodplain sediment and stonefly hindgut communities in the Nyack floodplain. For this survey, we recovered 4,312 partial 16s rRNA gene sequences using generally-conserved 16s rRNA gene primers. A focused survey of the stonefly hindguts using the same primer set led to the recovery of 120 sequences from 15 individual insects. Although the sediment survey was more extensive than hindgut survey, both revealed that a diverse microbial assemblage is present in the Nyack floodplain. Similar phylotypes were found in both environments. Sequence diversity was high, however, and only one sequence was shared between all samples. In addition, the predominance of individual phylotypes within the insect hindgut contents contrasted the much more even distribution across the diversity that was observed in the sediment at large.
7 Introduction
In riverine ecosystems, microorganisms comprise the majority of biomass and are responsible for cycling nutrients (Paerl et al. 2003). Microbial biomass is, in turn, available to protozoan and macroinvertebrate biofilm grazers and collectors as a nutrition source. Macroinvertebrates then become prey to larger animals (e.g. fish and birds)
(Stewart and Harper1996). The “microbial loop” is thus the energy “base” for much of the biota found inhabiting aquatic environments (Stanford and Ward 1993, Craftet al.
2002).
The Nyack floodplain, West Glacier, MT is a physically and ecologically complex environment (Stanford and Ward 1988, Stanford and Ward 1993, Craftet al.
2002). The hyporheic zone present in the subsurface of this region allows for extensive mixing of ground and surface water (Stanford and Ward 1993, Craft et al. 2002). In part because of this feature, the ecosystem is attributed with being a pristine habitat that contains a diverse macroinvertebrate assemblage, with hundreds of organisms being collected up to 3 km from the river channel (Craft et al. 2002). Included in this
ecosystem are two genera of plecopteran nymphs (stoneflies), Isocapnia sp. and
Pamper la sp., which spend much of their life cycle in the hyporheic zone (Stanford and
Ward 1993, Stewart and Harper 1996, Craft et al. 2002).
Stonefly nymphs are of particular interest in riverine ecosystems because they are
one of the must abundant Orders present and contribute to invertebrate secondary production (Stewart and Harper 1996, Huryn and Wallace 2000). Generally, stoneflies require cold, clean, and well oxygenated waters. Thus, they are frequently used as a biological indicator of stream health (Stewart and Harper 1996). Because of their
8 abundance, stoneflies are considered to be a major food source for many species of fish
(Stanford and Ward 1993, Stewart and Harper 1996, Huryn and Wallace 2000).
Stoneflies in the Nyack floodplain have also been suggested to be an important predator in hyporheic food webs (Stanford and Ward 1993). Although feeding may become more specialized in later instars, it is generally accepted that early developmental stages are omnivorous and consume large amounts of bacteria either by direct grazing or indirectly through the consumption of leaf litter and other particulate matter colonized by microbial biofilms (Cummin and Merritt 1996, Stewart and Harper 1996).
Understanding stonefly-bacterial interactions in aquatic environments is complicated by the inherent difficulties of studying microorganisms in natural settings.
They are small, difficult to identify visually, and otherwise difficult to isolate and study as evidenced by the well known gap between culture-dependent and culture-independent assessments of biomass and diversity employed in microbial ecology (Roszak and
Colwell 1987). In general, traditional techniques used in assessing microbial diversity in ecosystems, such as culture-based techniques, or respirometry, underestimate microbial diversity and abundance (Roszak and Colwell 1987, Streit and Schmitz 2004).
Consequently, molecular techniques have gained popularity for enumerating and characterizing the seemingly uncountable bacteria in many environments (Holben and
Harris 1995, Hughes et al. 2001, Streit and Schmitz 2004). However, even if all of the microorganisms could be identified and quantified in aquatic ecosystems, this does not provide a sufficient platform for understanding how microorganisms support or interact with macroinvertebrates in these environments.
9 The current study attempts to elucidate fundamental interactions and relationships between microbial communities and stoneflies in a pristine aquatic ecosystem. Using molecular techniques, we compared the diversity of microbial communities present in the hyporheic zone of the Nyack floodplain with those within the hindguts of plecopteran insects dwelling there. Using generally-conserved 16S rRNA gene primers (“universal primers”) we compared the indicated phylotypes obtained from hyporheic sediments to microbial communities inhabiting the hindguts of two stonefly genera commonly found in this same environment(Isocapnia sp. and Paraperla sp.) We hypothesized that the two microbial communities would contain similar members, and that the hindgut- dwelling communities would reflect an overlapping subset of the sediment community because stoneflies live in close proximity with the sediment and presumably consume microbial members of the sediment, or excrete microorganisms into the environment.
Remarkably, only one sequence was found in both the sediment and the hindguts by these surveys. Presumably this reflects the very high diversity present n the sediment community, where 2,521 of 4,312 total sequences were unique (i.e. were observed only once). This high degree of uniqueness suggests that full coverage of the diversity present in the community and that, even though 4,441 sequences were already examined,
essentially any new sequences examined (from sediment or sediment insect hindgut) was likely to be unique.
The one sequence that was common between sediment and hindgut had an Sab
score of 1.000 (i.e. was identical to) to Aeromonas salomincida subsp.salmonicida. In
addition, the predominance of individual phylotypes within the hindgut contents
exceeded the even distribution of microbial populations present in the sediment. Two
10 different mechanisms may account for these findings: 1) the hindgut compartment allows for bioamplification of select microbial populations; 2) the microbial populations consumed during the last biofilm meal are reflected in the hindgut survey, suggesting selective grazing by stoneflies on biofilms of preferred microbial populations that are patchily distributed throughout the floodplain.
Materials and Methods
Study Site The study was carried out in the Nyack floodplain near West Glacier, MT
(48°50’N, -113 °98’E). This site has a complex hydrology in that terrestrial surface water and groundwater intermingle extensively in this region. The physical structure of the environment is dynamic, changing seasonally (Stanford and Ward 1988, Stanford and
Ward 1993). It is considered unique because it is a relatively pristine regional floodplain system (Stanford et al. 1992, Craft et al. 2002).
Sediment Microbial Survey Sediment was collected from the Nyack floodplain in the fall
of 2004. Three samples were taken from three gravel bars in or near the river bed by
excavation down to the saturated zone. Total bacterial community DNA was extracted
from the three samples as previously described (Apajalahtiet al. 1998). GC fractionation
of this DNA was then performed as previously described (performed by Dr. Holben)
Holben et al., 2004, Holben and Harris 1995). This procedure reduces the complexity of the microbial community DNA by distributing it across several fractions based on the
%G+C content of the DNA of the component populations of the community (Holben and
Harris 1995). This work and the subsequent cloning and DNA sequence determination
were performed by Alimetrics Ltd (Helsinki, Finland), a commercial firm engaged in
11 molecular microbial ecology analyses. Briefly, DNA was subjected to equilibrium density centrifugation for fractionation based on %G+C content using the DNA binding dye bisbenzimadazole in conjunction with a CsCh solution buffered with 5 mM Tris (pH
8.0). Sixteen fractions were collected from each sample and partial 16S rRNA gene sequences representing the total microbial community from each fraction were amplified with the generally-conserved primers 536f (5’CAGCMGCCGCGGTAATWC3’) and
907r (5 ’CCGTCAATTCMTTTRAGTTT3’) (Holben et al. 2002). At least 61 clones were sequenced from each fraction. In all, 4,312 sequences were obtained from these efforts.
Recovery and analysis of insect hindgut microbial community DNA. A molecular survey of populations based on partial 16S rRNA gene sequences were performed on 7 individualIsocapnia Banks (Capniidae) and 9 individualParaperla Banks
(Chloroperlidae) stonefly nymphs. Stoneflies were obtained from the hyporheic zone of the Nyack floodplain in fall 2004, along the same gravel bars as the sediment sampling, by pumping and sieving water from previously established groundwater wells (Craftet al. 2002). Insects were surface sterilized with 95% ethanol prior to dissection. Hindguts were surgically removed and DNA extraction and purification was performed on the hindgut contents using the Power Soil Kit (MoBio, Solana Beach, CA). Purified DNA was subjected to PCR amplification using the same general primers 536f and 907r as
described above. Random cloning and phylogenetic analysis were performed as
described previously (Holbenet al. 2002). In all, 120 sequences were analyzed to
determine the predominant bacterial population(s) present in the insect hindguts.
DNA Sequence analysis and phylogenetic relationships of 16s rRNA. Sequence data was aligned and analyzed using BioNumerics v4.5 software. Data were analyzed with
12 rank/abundance curves and phylogenetic relationships were established for both sediment- and hindgut-dwelling microbial populations using the Ribosomal Database
Base Project (RDPII) website (http://rdp.cme.msu.edu/) (Maidaket al. 2001).
Phylogenetic relationships are presented based on their Sab score. Sab scores reflect how well the sequence being analyzed matched other sequences contained in the RDPII database (-180,000 sequences). An Sab score of 1.000 indicates an identical match to another sequence in the database. As the Sab score decreases, it indicates that the sequences being compared are less related to known sequences in the database. Where scores lower than 1.000 are obtained, there is still valuable information obtained through consideration of the most closely related known organisms. Although somewhat arbitrary, it is generally accepted that scores less than 0.900 should not be considered the same species (Holben et al. 2002).
Results and Discussion
Sediment Microbial Survey. A 16 rRNA gene sequence database was created
containing 4,312 sequences obtained by %GC fractionation and cloning of total community DNA from hyporheic sediment samples. The rank/abundance curve
generated from these data reveals that indeterminately large numbers of unique sequences
are present in the sediment of the Nyack floodplain (Fig 1). This is evidenced by the presence of 2,521unique sequences in the collection of 4,312 (58% of total) sequences
obtained when requiring 100% similarity to group sequences. As expected, sequence
homology continues to increase as less similarity is required, and at 95% similarity, 892
unique groups were present.
13 The DNA sequences were also subjected to phylogenetic analysis and compared to known phylotypes in the RDPII database. The indicated closest matches to known organisms showed a diverse distribution throughout current prokaryotic classification
(Table 1). The majority (86.7% of total) of 16S rRNA gene sequences were most closely related to as yet uncultured bacteria (Table 1). Interestingly, 75% of the uncultured sequences recovered are most closely related to uncultured phylotypes from terrestrial environments. The most abundant known phylotype detected (0.46% of total, average
Sab“0.991) was related to Microbacterium laevaniformans, which has also been isolated from soil (Accession number AF535159).
The results of the phylogenetic survey also indicated a wide array of microbial
“lifestyles” found in the sediment of the Nyack floodplain. For example, many of the taxa are capable of oxidizing or reducing many different compounds. Twelve of the sequences most closely matched Rhodoferax ferrireducens (averageSab=0.997), which is capable of acetate oxidization (AF435948). Five sequences obtained most closely matched Geobacter hephaestius, an organism capable of iron reduction (average
Sab“0.893, (AY737507)).The species Agrobacterium sanguineum was encountered 4 times and is able to degrade and biphenyl and dibenzofuran with an averageSab=0.960
(AB062105). In addition, many of the phylotypes obtained have previously been isolated
from the digestive tracts of animals. For example, Ruminococcus sp.sequences (2 recovered with an averageSab=0.900) is reported to be a butyrate-producing bacterium
from the human colon (AY305321).
The findings of the sediment molecular survey suggest that we have obtained
many sequences that have not previously been observed. Many of the Nyack sequences
14 matched phylotypes contained in the RDPII database with an Sab of less than 0.900. For example, 1,210 sequences are most closely related to an uncultured bacterium from uranium mining waste piles (AJ532724)with an average Sab=0.852. The phylotype
“uncultured Actinobacterium species (AY289386)”was encountered 366 times with an
averageSab of 0.881. These results indicate that the sediment of Nyack floodplain contains many novel microorganisms.
Stonefly Hindgut Microbial Community Analysis. Rank/abundance analysis
from this sequence pool reveals that the diversity in the16 rRNA gene sequences in the
insect hindguts is high (Fig. 2). This is evidenced by the presence of78 unique
sequences in the collection of 120 sequences (65% of total) obtained when requiring
100% similarity to group sequences. As expected, sequence homology continues to
increase as less similarity is required, and at 95% similarity, 28 unique groups were present.
The sequences obtained from the hindgut contents of both stonefly species
(Isocapnia sp. and Paraperla sp.) showed similar identities and distribution. Twenty-five percent and thirty-two percent of sequences for Isocapnia sp. and Paraperla sp.,
respectively, were most closely related to as yet uncultured bacterial species (Table 2).
Microbial phylotypes detected in the hindguts of both stonefly taxa included Acidovorax
spp., Massilia timonae, and Pedobacter spp. Perhaps the most striking result of the hindgut survey was the relatively high abundance of sequences identical (Sab=1.000) to
Aeromonas salmonicida subsp.salmonicida, a known fish pathogen (Austin 1997).
Thirty-three percent of the Paraperla sp. individuals contained at lease oneA.
salmonicida sequence with an Sab=1.000, which represents 14.8% of the total sequences
15 analyzed. Moreover, a total of 16.7% of the hindgut sequences were most closely related to A. salmonicida with Sab score of at least 0.950, suggesting that different strains of this pathogen may be present locally. Remarkably, this phylotype was only encountered only once in the survey of the total sediment microbial community (0.02% of total), indicating that it is in low abundance in the sediment environment relative to its abundance in the insect hindgut (16.7% of total sequences from hindgut).
Conclusion
The predominance of individual phylotypes within the insects exceeded the much more even distribution of microbial populations in the open sediment environment. Two mechanisms may underlie this pattern: i) bioamplification of select populations within the hindgut that exhibit animal-to-animal, or biogeographic variability; ii) selective grazing by animals on specific microbial populations in the form of biofilms or microcolonies as their last meal before sampling. Additional investigation will be required to reveal the relative contribution of these and/or other, mechanisms to understand the distribution patterns observed in this survey.
Of the most frequently encountered groups of bacteria in both the sediments and hindgut compartments, two are highly related to relatively well-known groups such as the pseudomonads and the Flavobacteria. This is perhaps not surprising since each of these
groups includes multiple examples of well-known and readily culturable species that
frequent both terrestrial and aquatic environments (Prescottet al. 1996). However, the
majority of sequences from the two microbial surveys represented uncultured phylotypes
from related environments such as glacial fields and coldwater lakes. This was not
16 unexpected, as there is a massive and still-growing body of work that supports the view that close to 99% of microorganisms have yet to be cultured and characterized.
A more surprising and more interesting observation was that only one sequence was obtained from both the sediments and the hindguts. This sequence matched the phylotype Aeromonas salmonicida subsp.salmonicida, which was encountered in high numbers in the hindgut contents, but was only encountered once in the entire sediment
survey of over 4,000 sequences. This raises the possibility that a known fish pathogen is
selectively grazed and/or bioamplified by plecopteran insects and subsequently
transmitted to fish hosts under favorable conditions, a possibility that we explore in a
separate study (Adams et al., submitted)
The data presented here, along with our prior studies (Adams et al. 2006a, Adams
et al. 2006b), collectively suggest that a relationship exists between stonefly nymphs and microbial communities inhabiting the sediments of the Nyack floodplain. We suggest
that plecopteran nymphs receive a nutritional benefit from selective grazing of patchily
distributed microbial biofilms (Adamset al. 2006), and that certain bacterial populations
receive the benefit of residence in a hindgut habitat that allows for biological and
physiological processes to occur in a controlled environment. For example, the genus
Acidovorax was found in both stonefly species’ hindguts. The potential benefit to the
microorganism in this case might be containment in an environment conducive to
denitrification, a trait for which Acidovorax are known. The potential benefit to the
animal may be access to nutrients that they do not obtain from other food sources. Many
terrestrial insects receive added nutritional benefits from the microorganisms inhabiting
the hindgut (Santo Domingo et al. 1998).
17 The frequency with which A. salmonicida was encountered in Paraperla sp. nymphs is of particular interest. To date, there is no known invertebrate reservoir or vector for this important fish pathogen and these findings suggest the intriguing possibility that stoneflies might somehow be involved.
18 Figure 1 Rank/abundance curve of sediment microbial community phylotypes.As depicted by this graph, there is great diversity in the sequences obtained from the microbial community of the Nyack flood plain. Approximately, 560 sequences were present at least twice. However, 2,512 were encountered only once. Because the curve flattens out and never reaches zero (i.e. appears to be asymptotic), it can be assumed that there many unique microbial 16s rRNA sequences present in the floodplain.
'T3
io o G
C/3 £ 0 Di-< 1 1 210 419 628 837 1046 1255 1464 1673 1882 2091 2300 2509 2718 2927 3136 3345 355^ Sequence ID 19 Figure 2 Rank/abundance curve ofphylotypes obtained from the hindgut contents of stoneflies. Sequence data obtained from both Paraperla spand Isocapnia sp. were pooled for this analysis. The curve is similar to that of the sediment survey and it appears that there are a large number of microbial 16s rRNA sequences present in the hindgut although numerical predominance of several populations is more apparent here than for the sediment community. 7 6 a> aS 5 2 o G • aw - i o 1 =tfc 0 1 4 7 10 13 16 19 22 25 28 31 34 37 40 43 46 49 52 55 58 61 64 67 70 73 76 79 82 85 88 91 94 Sequence ID 20 Table 1 Phylogenetic relatedness o f sediment dwelling microbial populations was determined for 4,312 clones. The RDP II database was searched for the best match to all phylotypes. The table below contains information for phylotypes that were encountered. Average Sab scores for sequences that matched the phylotype are listed. How Avg % of Phylotype notes (Accession Many Sab Best Match to All Phylotypes total number) uranium mining waste piles 1210 0.852 uncultured bacterium 28.06 (AJ5 32724) 366 0.881 uncultured actinobacterium 8.49 soil (AY289386) 354 0.878 uncultured actinobacterium 8.21 actinobacterium (AY395336) 263 0.904 uncultured beta proteobacterium 6.10 (AKYG1130) 221 0.891 uncultured alpha proteobacterium 5.13 Cave deposits (AY695717) alpha proteobacterium 220 0.893 uncultured alpha proteobacterium 5.10 (AY9219) 187 0.825 uncultured soil bacterium 4.34 soil bacterium (AJ390482) 160 0.823 uncultured delta proteobacterium 3.71 delta proteobacterium (AY9219) 117 0.880 uncultured Chloroflexi bacterium 2.71 (AKYG1672) 110 0.898 bacterium Ellin325 2.55 soil bacteria (AF498707) 104 0.860 uncultured Holophaga sp 2.41 (AJ519367) gamma proteobacterium 96 0.870 uncultured gamma proteobacterium 2.23 (AY92186) uncultured Green Bay Green Bay ferromanganous ferromanganous micronodule micronodule bacterium 83 0.951 bacterium 1.92 (AY612302) wastewater treatment plant 75 0.801 uncultured planctomycete 1.74 (BX294744) arsenic-resistant proteobacterium 60 0.981 beta proteobacterium 1.39 (AY612302) 59 0.895 Bacteria 1.37 (Z95708) 49 0.856 uncultured sludge bacterium 1.14 sludge bacterium uranium mining waste piles and 41 0.786 uncultured proteobacterium 0.95 mill tailings (AJ534624) Nullarbor caves, Australia 40 0.729 unidentified bacterium 0.93 (AF317771) 38 0.720 uncultured verrucomicrobium 0.88 (AJ401115) butyrate-producing bacterium 37 0.986 butyrate-producing bacterium 0.86 (AJ270484) human colonic 34 0.971 uncultured bacterium 0.79 sample(AJ408980) uncultured Geobacteraceae 33 0.981 bacterium 0.77 pasture soil (AY395) 21 Table 1 con’t biphenyl-polluted soil 30 0.794 uncultured eubacterium 0.70 (AJ292600) uncultured Rubrobacteridae Rubrobacteridae bacterium 30 0.917 bacterium 0.70 (AY39) forest soil bacterium 26 0.842 uncultured forest soil bacterium 0.60 (AY612302) uncultured Gemmatimonadetes Gemmatimonadetes bacterium 25 0.898 bacterium 0.58 (AKYG781) Isolated from Manufactured Gas 20 0.991 Microbacterium laevaniformans 0.46 Plant Soil (AF535159) uncultured candidate division SPAM 19 0.707 bacterium 0.44 (AKYG) humic-reducing bacteria from a diversity of environments 18 0.889 Geobacter sp. 0.42 (AF019932) sulfate-reducing bacterium 17 0.930 sulfate-reducing bacterium 0.39 (AJ389622) rhizobia occurring on shrubby 14 0.989 Bradyrhizobium genosp 0.32 legumes (Z94818) bacterium degrading methyloxylated aromatic 14 0.824 Holophaga foetida 0.32 compounds (X77215) 14 0.939 delta proteobacterium 0.32 archaeological wood human intestinal bacteria 13 0.972 Bacteroides sp 0.30 (DQ100446) Microbial diversity of spacecraft 13 0.992 Micrococcus luteus 0.30 assembly facilities (AY 167858) 13 0.996 Phenanthrene-degrading bacterium 0.30 soil (AY177358) phenanthrene-degrading 13 0.996 Phenanthrene-degrading bacterium 0.30 bacterium (AY297795) 12 0.949 Faecalibacterium prausnitzii 0.28 butyrate producing (AJ270469) 12 0.914 Nocardioides sp. 0.28 atrazine-degrading (DQ485407) 12 0.997 Rhodoferax ferrireducens 0.28 oxidizes acetate (AF435948) 12 0.958 Antarctic bacterium 0.28 (AJ440984) 12 0.966 uncultivated soil bacterium clone 0.28 soil bacterium clone (AF013550) River T aff Epithilium 11 0.935 Flavobacterium sp 0.26 (AF493654 ) 11 0.932 Hyphomicrobium sp 0.26 Soil Bacteria (AF408954) 11 0.837 uncultured Bacteroidetes bacterium 0.26 Bacteroidetes bacterium (AY92) 10 0.961 Marmoricola sp 0.23 Soil Bacteria(AF408953) Alpha- 10 0.810 Nordella oligomobilis 0.23 Proteobacterium( AF37088) 22 T a b le 1 c o n ’t 10 0.954 alpha proteobacterium 0.23 (AF235998) 10 0.657 uncultured sponge symbiont 0.23 sponge symbiont (AF186413) 9 0.975 Bradyrhizobium japonicum 0.21 (PAUC37f) 9 0.966 Bradyrhizobium sp 0.21 (AF435948) 9 0.982 Mycobacterium buckleii 0.21 (AF000225) 9 0.758 Nitrosospira sp. 0.21 ammonia-oxidizer (DQ497443) 9 0.856 Nocardioides plantarum 0.21 (AF005008) 9 0.964 Ruminococcus bromii 0.21 (X85099) 9 1.000 Staphylococcus aureus 0.21 (X68417) coal-tar-waste-contaminated 9 0.922 uncultured Nitrospira sp. 0.21 aquifer waters (AF351231) 8 0.974 Pseudomonas sp. 0.19 Arctic sea ice(AF468334) rRNA group II Pseudomonas 8 0.961 Ralstonia pickettii 0.19 (X67042) hexane degrading biofilters 8 0.918 Sphingomonas sp. 0.19 (AJ555475) human oral bacterium 8 0.967 human oral bacterium 0.19 (AF202012) model drinking water biofilms 7 0.913 Caulobacter sp 0.16 (AY957895) 7 0.979 Eubacterium ramulus 0.16 (AJ011522) 7 0.766 agricultural soil bacterium 0.16 (AJ252614) Arctic sea ice bacterium 7 0.908 Arctic sea ice bacterium 0.16 (AF468386) eubacterium from Cecidotrioza eubacterium from Cecidotrioza 7 0.591 sozanica 0.16 sozanica (AF286124) Aquificales bacterium 7 0.272 uncultured Aquificales bacterium 0.16 (AF445658) uncultured Comamonadaceae 7 0.970 bacterium 0.16 (AF52300) 7 0.578 uncultured Gram-positive bacterium 0.16 (AF4457) 7 0.654 uncultured thermal soil bacterium 0.16 thermal soil bacterium (AF391) wastewater treatment 6 0.978 Flavobacterium granulensis 0.14 plant(AB 180738) bottled mineral 6 0.824 Herminiimonas fonticola 0.14 water( A Y676462) 6 0.980 Shewanella putrefaciens 0.14 (X82133) bacteria growing in natural 6 1.000 uncultured Aquabacterium sp. 0.14 mineral water (AF523008) Planktonic Microbial 6 1.000 uncultured Aquabacterium sp. 0.14 Community (AF523008) PCE-contaminated site 5 0.996 Acinetobacter junii 0.12 (AF529331) 23 Table 1 con’t 5 0.819 Anaeromyxobacter dehalogenans 0.12 aryl-halorespiring (AF3 82400) phytopathogenic bacteria 5 0.844 Bdellovibrio sp 0.12 (AF148940) 5 0.893 Geobacter hephaestius 0.12 iron-reducing (AY737507) Argentinean dry fermented 5 0.983 Pediococcus acidilactici 0.12 sausages(AY865646) 5 0.962 Psychrobacter phenylpyruvicus 0.12 (AF005192) 5 0.992 Sphingomonas melonis 0.12 (AB055863) Acidobacteria bacterium 5 0.587 Verrucomicrobium spinosum 0.12 (AY921986) Endosymbionts of Paramecium 5 0.662 endosymbiont of Acanthamoeba sp. 0.12 spp. (AY102614) 5 0.711 uncultured Chlamydiales bacterium 0.12 (AF364560) biphenyl and dibenzofuran 4 0.960 Agrobacterium sanguineum 0.09 degrading(AB062105) 4 0.880 Bactero ides fragilis 0.09 (X83946) Desulfonatronovibrio sulfate-reducing 4 0.301 hydrogenovorans 0.09 bacterium(E71X99236) human subgingival 4 0.989 Dialister sp 0.09 plaque(AF287787) 4 1.000 Mesorhizobium sp 0.09 cold-adapted (AF282922) 4 0.943 Moraxella osloensis 0.09 phosphorus removal (Y15855) 4 0.839 Planctomyces sp. 0.09 (X81955) 4 0.990 Propionibacterium acnes 0.09 (AB108480) 4 0.967 Pseudomonas saccharophila 0.09 ultrapure water (AF368755) 4 0.991 Variovorax sp 0.09 phenol-stimulated (AB051688) 4 0.912 Caulobacter sp. 0.09 (AJ227774) human subgingival plaque 4 0.989 Dialister sp. oral clone 0.09 (AF287787) 4 0.568 Legionella-like amoebal pathogen 0.09 (AY741401) uncultured Banisveld landfill Banisveld landfill bacterium 4 0.685 bacterium 0.09 (AY0) deep-well injection site Tomsk- 4 0.977 uncultured Geothrix sp. 0.09 7, Siberia (AJ583203) uncultured Xiphinematobacteriaceae 4 0.905 bacterium 0.09 ginseng field (EB10) Arctic sea ice bacterium 3 0.983 Acinetobacter sp. 0.07 (AF468386) Myxobacteria 3 0.802 Archangium gephyra 0.07 (AJ233913) 3 1.000 Arthrobacter ramosus 0.07 (X80742) Human Intestinal Bacteroides 3 0.993 Bacteroides uniformis 0.07 (AB050110) 3 0.970 Brevibacterium casei 0.07 Brevibacterium (AJ251418) 24 Table 1 con’t freshwater lake 3 0.961 Chryseobacterium daecheongense 0.07 sediment(AJ457206) H2-utilizing bacteria 3 0.944 Desulfomicrobium hypogeium 0.07 (AF132738) sulfate-reducing bacteria 3 0.604 Desulfonema ishimotonii 0.07 (U45992) Mongolian soda lake (AY298904) 3 0.637 Ectothiorhodosinus mongolicum 0.07 Greenland glacier ice core 3 0.975 Eubacterium rectale 0.07 AY169428 turfgrass rhizosphere soil 3 0.959 Flavobacterium degerlachei 0.07 (DQ335110) 3 0.807 Mesorhizobium thiogangeticum 0.07 sulfur-oxidizing (AJ864462) 3 0.927 Methylobacterium aquaticum 0.07 drinking water (AJ635304) plant-nematode 3 0.937 Microbacterium sp 0.07 Microbacteriaceae(AB042073) Sphingomonads 3 0.977 Novosphingobium sp. 0.07 (AJ000920) 3 0.913 Novosphingobium stygium 0.07 deep-sea sediments (AB025013) 3 0.997 Ralstonia insidiosa 0.07 (AJ539233) 3 0.961 Rhodoferax antarcticus 0.07 Lake Fryxell (AY609198) 3 0.941 Rubrivivax gelatinosus 0.07 purple bacterium ( ABO 16167) sugarcane-Sesbania cannabina 3 0.922 Sinorhizobium sp. 0.07 rotation fields (AF285965) Carbon Monoxide oxidizer 3 0.789 Stappia sp. 0.07 (AY307928) 3 1.000 Stenotrophomonas sp. 0.07 cooked vegetables (AY259519) 3 0.975 uncultured Bradyrhizobium sp. 0.07 (AY599) 3 0.734 uncultured Clostridiaceae bacterium 0.07 Dugong (A B 218350) 3 0.702 uncultured Crater Lake bacterium 0.07 Crater Lake bacterium (AF3167) uncultured Holophaga/Acidobacterium 3 0.807 Holophaga/Acidobacterium 0.07 (AJ241) uncultured low G+C Gram-positive low G+C Gram-positive 3 0.769 bacterium 0.07 bacterium (4C28d-4) 3 0.768 uncultured rumen bacterium 0.07 rumen bacterium (AB034125) uncultured sulfate-reducing sulfate-reducing bacterium 3 0.926 bacterium 0.07 (AJ389622) uncultured Termite group 1 Termite group 1 bacterium 3 0.707 bacterium 0.07 (AB08) 2 0.983 Acidovorax sp. 0.05 (AY212579) 2 1.000 Actinomyces naeslundii 0.05 tropical molasses grass 2 0.679 Azospirillum sp 0.05 (DQ022959) 2 0.970 Bacteroides ovatus 0.05 (AF139524) 25 Table 1 con’t laboratory mouse liver 2 1.000 Brachybacterium muris 0.05 (AJ537574) commercial inoculants 2 0.971 Bradyrhizobium elkanii 0.05 (AY904762) 2 0.869 Chondromyces lanuginosus 0.05 (AJ233939). Cold-Adapted Bacteria 2 0.955 Cytophaga sp 0.05 (AF260716) Anaerobic benzene oxidation 2 0.918 Dechloromonas sp 0.05 (AY032611) Dao red wine (Portugal) 2 0.981 Gluconobacter oxydans 0.05 (AY206688) 2 0.540 Methylocaldum sp 0.05 landfill cover soil (AF215632) fed-batch garbage composters 2 0.824 Micrococcus sp. 0.05 (AB188213) Soil Bacteria 2 0.909 Nocardioidaceae str 0.05 (AF408943) protease-producing bacteria 2 0.911 Oxalobacter sp 0.05 from antarctic soil (AJ496038) 2 0.580 Parachlamydia acanthamoebae 0.05 Obligate intracellular (Y07556) initiates catabolism 2 0.886 Parvibaculum lavamentivorans 0.05 (AY387398) 2 0.974 Phascolarctobacterium faecium 0.05 (X72867) 2 0.914 Phormidium sp 0.05 (AB 183566) 2 0.696 Pirellula staleyi 0.05 Pirellula staleyi (AF399914) soil column system 2 0.974 Rhizobium sp. 0.05 (AY177365) H2/C02-using acetogenic 2 0.990 Ruminococcus gnavus 0.05 bacterium (X94967) butyrate-producing bacteria from 2 0.900 Ruminococcus sp. 0.05 the human colon (AY305321) human subgingival crevice 2 0.776 Streptococcus salivarius 0.05 (AF202012) human subgingival crevice 2 0.981 Streptococcus sp. 0.05 (AF202012) 2 0.458 Thermodesulfovibrio sp 0.05 sulfate-reducers (AB021302) 2 0.986 Zea mays 0.05 chloroplast (X86563) gamma proteobacterium 2 0.792 gamma proteobacterium 0.05 (AB 174845) bacteria from soil and freshwater 2 0.737 Gemmata-like str. 0.05 (JW10-3f) Gram-positive bacteria 2 0.847 Gram-positive bacteria SOGA 31 0.05 (A.T7.44807) Gram-positive bacteria 2 0.746 uncultured Clostridia bacterium 0.05 (AY370633) 2 0.984 uncultured Corynebacterium sp. 0.05 Prostatitis (AF115945) 26 Table 1 con’t 2 0.263 uncultured Deinococci bacterium 0.05 (AF51396) uncultured Sphingobacteriales 2 0.897 bacterium; SF54; AJ6 0.05 Sphingobacteriales bacterium contaminated boreal 2 0.848 unidentified eubacterium 0.05 groundwater (J009714) paper mill waste water 1 0.767 Acetanaerobacterium elongatum 0.02 (AY518589) 1 0.925 Acidimicrobidae bacterium 0.02 soil bacteria (AY673309) Delftia acidovorans 1 0.949 Acidovorax temperans 0.02 (AF078766) 1 1.000 Acinetobacter johnsonii 0.02 Eikelbooms Type (X89775) 1 1.000 Aeromonas salmonicida 0.02 Fish pathogen (X74680) 1 0.963 Afipia genosp. 0.02 (U87783) dibenzothiophene-degrading 1 0.963 Agrobacterium tumefaciens 0.02 bacterium (AY364329) 1 0.793 Amaricoccus macauensis 0.02 sludge biomass (U88042) oxalotrophic bacteria 1 0.799 Ammoniphilus oxalaticus 0.02 (Y14578) 1 0.777 Anaerosinus glycerini 0.02 (AJ010960) Berlin drinking water system 1 0.952 Aquabacterium parvum 0.02 (AF035052) 1 0.827 Aquamonas fontana 0.02 (AB 120964) 1 0.970 Arthrobacter oxydans 0.02 (X83408) psychrophilic isolates 1 0.983 Arthrobacter psychrolactophilus 0.02 (AF134180) electricity-generating 1 0.950 Azonexus fungiphilus 0.02 (AJ630292) 1 0.942 Arthrobacter sp 0.02 (AJ810894) Anaerobic degradation 1 0.986 Azoarcus sp 0.02 (Y13222) spacecraft assembly facility 1 0.989 Bacillus nealsonii 0.02 (AF234863) Drentse A grasslands 1 0.713 Bacillus novalis 0.02 (AJ542512) Human Intestinal Bacteroides 1 1.000 Bacteroides vulgatus 0.02 (AB050111) 1 0.983 Bifidobacterium animalis 0.02 (AB050136) 1 0.938 Bifidobacterium breve 0.02 rat cecal content (AF491832) Salmon and coalfish 1 0.904 Brochothrix thermosphacta 0.02 (AY543017) 1 1.000 Bacillus sp 0.02 (AF519171) 1 1.000 Blastococcus sp 0.02 Soil Bacteria AY234675 1 0.735 Bosea sp 0.02 hexane degrading AJ313022 1 0.397 Caedibacter taeniospiralis 0.02 AY102612 27 1 0.329 Candidatus Carsonella ruddii 0.02 psyllids AF243137 1 0.963 Caulobacter fusiformis 0.02 NO INFO AJ22775 1 0.912 Chelatococcus asaccharovorans 0.02 AJ871433 Table 1 con’t Antarctic microbial mat 1 0.798 Clostridium bowmanii 0.02 CB0506120 1 0.798 Clostridium colinum 0.02 Rodent and Rabbit DQ352812 1 0.933 Clostridium leptum 0.02 AJ305238 Methanogenic degradation 1 0.944 Clostridium sp. 0.02 Y15985 dune soil beta-subclass 1 0.924 Collimonas fungivorans 0.02 Proteobacteria AJ310395 1 0.969 Comamonadaceae bacterium 0.02 freshwater plankton AJ556799 Gut Of Folsomia Candida 1 1.000 Comamonas sp. 0.02 AJ002803 1 0.611 Crossiella equi 0.02 equine placentas AF245017 1 0.876 Curvibacter gracilis 0.02 well water AB 109889 1 0.719 Dehalobacterium formicoaceticum 0.02 anaerobic bacteria X86690 1 0.766 Denitromonas aromaticus 0.02 aromatic-degrading AB049763 1 0.969 Desulfomicrobium sp 0.02 H2-utilizing bacteria AF132738 anaerobic dehalogenating 1 0.457 Desulfomonile limimaris 0.02 AF230531 sulfate-reducing bacteria 0.945 Desulfotalea sp 0.02 AJ318381 Human faeces 1 1.000 Eggerthella sp 0.02 AF304434 Diarrhea stool specimens 1 0.967 Escherichia coli 0.02 Z83205 1 0.776 Flexibacter aggregans 0.02 AB078038 reduction of iron and other 1 0.624 Geobacter metallireducens 0.02 metals L07834 marine environment 1 0.693 Haliangium ochraceum 0.02 AB016470 1 0.711 Hyphomicrobium denitrificans 0.02 Y14308 Soil Bacteria 1 0.840 Hyphomicrobium zavarzinii 0.02 AF408954 Lake Sediments of Ardley 1 0.349 Janthinobacterium sp 0.02 Island, Antarctica AJ5 51147 1 0.981 Klebsiella pneumoniae 0.02 Y17656 1 0.945 Lactobacillus gasseri 0.02 AF519171 1 0.601 Lactobacillus paracasei subsp 0.02 AB237509 human oral cavity 1 0.983 Lactobacillus sp. 0.02 AY349382 28 Table 1 con’t cultivated mushroom Agaricus 1 0.850 Lechevalieria aerocolonigenes 0.02 bisporus AW324527 1 0.593 Legionella beliardensis 0.02 Tags of Fruits AT000119 Chaco Arido region (Argentina) 1 0.941 Mesorhizobium chacoense 0.02 AJ278249 1 0.763 Mesorhizobium loti 0.02 leguminous plants D 14514 1 0.908 Methylobacter psychrophilus 0.02 AF152597 methane-oxidizing bacterium 1 0.728 Methylobacter sp. 0.02 AJ414655 Wadden Sea 1 0.625 Micrococcaceae bacterium 0.02 AY370619 acid in the cell wall 1 0.890 Nakamurella multipartita 0.02 Y08541 1 0.934 Nocardia sp. 0.02 AF430064 1 0.909 Odontella sinensis 0.02 Chloroplast Genome Z67753 1 0.911 Paenibacillus durus 0.02 X77846 1 0.931 Pelobacter propionicus 0.02 X70954 anaerobic granular sludge 1 0.353 Pelotomaculum sp. 0.02 (AB159557) Bacterial diversity in a nonsaline alkaline environment 1 1.000 Phenylobacterium sp 0.02 (AJ717391) 1 0.330 Phytophthora infestans 0.02 West Virginia (U17009) 1 0.601 Pseudomonas alcaligenes 0.02 (AF390747) 1 0.961 Pseudomonas tolaasii 0.02 (AF348507) 1 0.961 Pseudonocardia yunnanensis 0.02 (D85472) soil crusts from Colorado 1 0.913 Ramlibacter tataouinensis 0.02 (AJ871248) 1 0.304 Reclinomonas americana 0.02 (DQ187322) 1 0.929 Rhodopseudomonas rhenobacensis 0.02 (AB087719) Baer Soda Lake in Inner 1 0.909 Salinicoccus alkaliphilus 0.02 Mongolia (AF275710) isolated from the Russian space 1 0.986 Sphingomonas yabuuchiae 0.02 laboratory (AB071955) soil crusts from Colorado 1 0.848 Spirosoma escalantus 0.02 (AJ871248) biodeteriorated wall paintings 1 0.972 Staphylococcus sp 0.02 (AJ276810) BTEX degrading isolates and in 1 0.815 Stenotrophomonas maltophilia 0.02 subsurface soils (AY512626) isolated from petroleum sludge 1 0.919 Terrahaemophilus aromaticivorans 0.02 (AB098612) Great Artesian Basin of 1 0.489 Thermaerobacter subterraneus 0.02 Australia (AF34566) 29 Table 1 con’t 1 0.956 Veillonella dispar 0.02 (AY995770) Marine Actinomycetes 1 0.879 Verrucosispora gifhornensis 0.02 (DQ416204) Caedibacter taeniospiralis; 1 0.397 AY102612 0.02 (AY 102612) 1 0.329 Candidatus Carsonella ruddii; 0.02 Psyllid (AF243137) 1 0.963 Caulobacter fusiformis (T) 0.02 (AJ22775) endosymbiont of Scyphophorus endosymbiont of Scyphophorus 1 0.329 yuccae 0.02 yuccae (AY 126637) endosymbiont of unidentified scaly endosymbiont of unidentified 0.620 snail 0.02 scaly snail (AY310506) 1 0.951 marine bacterium P wp0225 0.02 marine bacterium (AY 188939) sulfur-oxidizing bacterium 1 0.693 sulfur-oxidizing bacterium OAII2 0.02 (AF170423) anaerobic bacterium 1 0.696 uncultured anaerobic bacterium 0.02 (AY953202) natural mineral water l 0.835 uncultured Caulobacter sp. 0.02 (AF523036) 1 0.744 uncultured Chlorobi bacterium 0.02 (AJ428453) cold-seep sediments in Japan 1 0.614 uncultured Cytophaga sp. 0.02 Trench (AB 189358) Sulfate-Reducing Bacterial 1 0.448 uncultured Desulforhopalus sp. 0.02 (AY177796) uncultured hydrocarbon seep hydrocarbon seep bacterium 1 0.717 bacterium 0.02 (GCA112) marine environmental 1 0.750 uncultured Pirellula clone 0.02 (AF029076) sludge treated cassava starch 1 0.864 uncultured Pseudomonas sp. 0.02 wastewater (AY693823) uncultured rape rhizosphere rape rhizosphere bacterium 1 0.915 bacterium 0.02 (AJ29) 1 0.969 uncultured Streptococcus sp. 0.02 Oral Microflora (DQ016723) 1 0.489 uncultured Thermacetogenium sp.; 0.02 (DQ0976) 1 0.608 unidentified proteobacterium 0.02 marine coastal picoplankton 1 0.743 unidentified rumen bacterium 0.02 rumen bacteria (AF018564) 30 Table 2Phylotypes obtained from stonefly hindguts were submitted to the RDPII database and the best match to all phylotypes is presented. Average Sab scores for sequences that matched the phylotype listed Biological Notes (Prescott Sequence Match to Avg. Number of % of et al. 1996, unless otherwise Insect all Phylotypes ^ab sequences total noted) denitrifying, common in Isocapnia Pseudomonas sp. 0.944 15 28.8 aquatic environments Acidovorax sp. 0.808 12 23.1 Denitrifying Uncultured 0.875 13 25.0 common in aquatic ecosystems, aerobic Flavobacterium sp. 0.954 8 15.4 respiration aquatic, aerobic (Shivaji et Pedobacter sp. 0.869 4 7.7 al. 2005) Nonfermentative aerobic Massilia timonae 0.892 4 7.7 (Lindquist et al. 2003) Pseudomonas fluorescens (T) 1.000 2 3.8 aquatic Antrobacter spp. 0.650 1 1.9 Atrazine-degrading Cytophaga sp. 0.618 1 1.9 Paraperla Uncultured 0.787 17 31.5 Aeromonas salmonicida (T) 0.986 9 16.7 fish pathogen Acidovorax sp. 0.915 9 16.7 Denitrifying 31 Table 2 Aeromonas con’t salmonicida (T) 1.000 8 14.8 fish pathogen Antrobacter spp. 0.856 3 5.6 endosymbiont of leech Rickettsia sp. 0.849 3 5.6 (Kikuchi et al. 2002) Micrococcus sp. 0.813 2 3.7 Alkalophiles denitrifying, common in Pseudomonas sp. 0.949 2 3.7 aquatic environments generally anaerobic Streptococccus sp. 0.685 2 3.7 respiration aquatic sediments Thermoincola sp. 0.563 2 3.7 (DQ394909) Aquamonas Fontana 0.890 1 1.9 Aquatic habitats Nonfermentative aerobic Massilia timonae 0.938 1 1.9 (Lindquist et al. 2003) aquatic, aerobic (Shivaji et Pedobacter sp. 0.922 1 1.9 al. 2005) 32 References 1. Adams, S.A., Adams, A.S., and Holben. In preparation, 2006a. The deadly meal: Bacterial pathogen transmission to rainbow trout via stoneflies. 2. Adams, S.A., Gordon, N., Valenzuela, A. Gannon, J., Holdben, W. In preparation, 2006b. Attraction and feeding behavior of Plecopteran nymphs to different bacterial isolates. 3. Apajalahti, J. H., L. K. Sarkilahti, B. R. Maki, J. P. Heikkinen, P. H. Nurminen, and W. E. Holben. 1998. Effective recovery of bacterial DNA and percent- guanine-plus-cytosine-based analysis of community structure in the gastrointestinal tract of broiler chickens. Appl Environ Microbio. 64:4084—4088. 4. Austin, B. 1997. Progress in understanding the fish pathogen Aeromonas salmonicida. Mar Biotechnol. 15: 131-134. 5. Craft, J.A., Stanford, J., and Pusch, M. 2002. Microbial respiration within a floodplain aquifer of a large scale gravel-bed river.Freshwater Biology. 47:251- 261. 6. Cummin, K. W., and R. W. Merritt. 1996. Ecology and Distribution of Aquatic Insects. Pages 74-86 in K. W. Cummins and R. W. Merritt, editors. Aquatic Insects of North America. Kendall/Hunt Publishing Company, Dubuque. 7. Duncan, S.H., Hold, G.L., Harmsen, H.J.M., Stewart, C.S. and Flint, H.J. 2002. Growth requirements and fermentation products of Fusobacterium prausnitzii, and a proposal to reclassify it as Faecalibacterium prausnitzii gen. nov., comb, nov.Int JSyst EvolMicrobiol. 52: 2141-2146. 33 8. Ellis, B., Stanford, J., and Ward, J. 1998. Microbial assemblages and production in alluvial aquifers of the Flathead River, Montana, USA.J N Am Benthol Soc. 17:382-402. 9. Feris, K. P., Ramsey, P.W., Frazar, C., Rillig, M., Moore, J.N., Gannon, J.E., and Holben, W.E. 2004. Seasonal dynamics of shallow-hyporheic-zone microbial community structure along a heavy-metal contamination gradient. Appl Environ Microbio. 70:2323-2331. 10. Holben, W. E., and D. Harris. 1995. DNA-based monitoring of total bacterial community structure in environmental samples.Mol Ecol. 4:627-631. 11. Holben, W.E., Williams, P., Saarinen, Sarkilahti, M., and Apajalhti, J.H.A. 2002. Phylogenetic Analysis of Intestinal Microflora Indicates a Novel Mycoplasma Phylotype in Farmed and Wild Salmon. Microb Ecol. 44:175-185. 12. Hughes, J., Hellmann, J., Ricketts, T., Bohannan, B. 2001. Counting the uncountable: Statistical approaches to estimating microbial diversity.Appl Environ Microbiol. 67:4399-4406. 13. Huryn, A. D., and Wallace, J. B. 2000. Life history and production of stream insects. Annu Rev Entomol. 45:83-110. 14. Finneran, K., Johnsen, C., and Lovley, D. 2003. Rhodoferax ferrireducens sp. nov., a psychrotolerant, facultatively anaerobic bacterium that oxidizes acetate with the reduction of Fe(III). IntJSyst Evol Microbiol. 53: 669-673. 15. La Scola B., Barrassi L., Raoult D. 2004. A novel alpha-Proteobacterium, Nordella oligomobilis gen. nov., sp. nov., isolated by using amoebal co-cultures. Research in Microbiology. 155: 47-51 34 16. Liesack W., Bak F., Kreft J.U., Stackebrandt E.1994. Holophaga foetida gen. nov., sp. nov., a new, homoacetogenic bacterium degrading methoxylated aromatic compounds. Arch Microbiol. 162:85-90. 17. Lindquist D, Murrill D, Burran WP, Winans G, Janda JM, Probert W. 2003. Characteristics of Massilia timonae and Massilia timonae-likQ isolates from human patients, with an emended description of the species. J Clin Microbiol. 41:192-196. 18. Kikuchi Y, Sameshima S, Kitade O, Kojima J, Fukatsu T. 2002. Novel Clade of Rickettsia spp. from Leeches. Appl Environ Microbiol. 68: 999-1004. 19. Knapp, C., Caquet, T., Hanson, M., Lagadic, L., and Graham, D. 2005. Response of water column microbial communities to sudden exposure to deltamethrin in aquatic mesocosms. FEMS Micrbiol Ecol. 54:157-165. 20. Maidak, B.L., Cole, J.R., Lilbum, T.G., Parker, C.T., Saxman, P.R., Farris, R.J., Garrity, G.M., Olsen, G.J., Schmidt, T.M., and Tiedje, J.M. 2001. The RDP-II (Ribosomal Database Project).Nucleic Acid Research. 29:173-174. 21. Paerl, H., Dyble, J., Moisander, P., Noble, R., Piehler, M., Pinckney, J., Stepp, T., Twomey, L., Valdes, L. 2003. Microbial indicators of aquatic ecosystem change: current applications to eutrophication studies. FEMS Micrbiol Ecol. 46:233-246. 22. Prescott, L.M., Harley, J.P., and Klein, D.A. 1996. Microbiology. In L.M. Pescott, J.P. Harley, and D.A. Klein eds. Microbiology. WCB Publishers. Dubuque, IA. 23. Roszak, D.B. and Colwell, R.R. 1987. Survival strategies of bacteria in the natural environment.Microbiological Reviews. 51:365-379. 35 24. Santo Domingo, J.W, Kaufman, M.G., Klug, M.J., and Tiedje, J.M. 1998. Characterization of the Cricket Hindgut Microbiota with Fluorescently Labeled rRNA-Targeted Oligonucleotide Probes. Appl Environ Microbiol. 64:752-755. 25. Shivaji S., Chaturvedi P., Reddy G.S., and Suresh K. 2005. Pedobacter himalayensis sp. nov., from the Hamta glacier located in the Himalayan mountain ranges of India. Int J Syst Evol Microbiol. 55:1083-1088. 26. Stanford, J., and Ward, J. 1988. The hyporheic habitat of river ecosystems. Nature. 335:64-66 27. Stanford, J. and Ward, J. 1993. An ecosystem perspective of alluvial rivers: connectivity and the hyporheic corridor. JNAm Benthol Soc. 12:48-60. 28. Streit, W. R. and Schmitz, R.A. 2004. Metagenomics-the key to the uncultured microbes. Current Opinion in Microbiology. 7:492-498. 29. Stewart, K.W and Haper, P.P. 1996. Plecoptera. Pages 217-266 in K. W. Cummins and R. W. Merritt, editors. Aquatic Insects of North America. Kendall/Hunt Publishing Company, Dubuque, IA. 30. Sullivan J.T, Brown S.D, Yocum RR, and Ronson C.W. 2001. The bio operon on the acquired symbiosis island of Mesorhizobium sp. strain R7A includes a novel gene involved in pimeloyl-CoA synthesis. Microbiology. 147:1315-1322. 31. Xie C.H. and Yokota A. 2006. Sphingomonas azotifigens sp. nov., a nitrogen- fixing bacterium isolated from the roots of Oryza sativa. Int J Syst Evol Microbiol. 56:889-93. 36 Chapter 2 Attraction and Feeding Behavior of Plecopteran Nymphs to Different Bacterial Isolates Abstract Early instars of many plecopteran (stonefly) species are believed to consume microbial biofilms in river sediments (Cummin and Merritt 1996, Stewart and Harper 1996). Results of our previous studies indicate that microbial diversity in the hindguts of stoneflies is uneven (see Chapter 1). We developed two different hypotheses to account for this observation: 1) the observed unevenness of microbial populations in the insect hindguts may simply reflect the last meal consumed by the individual insect; or, 2) that stonefly nymphs selectively graze on discrete microbial biofilms primarily comprised of specific microbial populations that are patchily distributed in the environment. To test these hypotheses, we created discrete microbial biofilms with frequently encountered individual hindgut phylotypes, namely, Pedobacter sp.II14, Pseudomonas sp.II5 and 173, and Aeromonas salmonicida subsp.salmonicida. In addition, two other microorganisms were chosen based on their nutritional (i.e. carbon to nitrogen; C:N ratio) content, Rhodococcus sp.lS9 (high C:N ratio), and Flavobacterium 1143sp. (low C:N ratio). We demonstrate that plecopteran nymphs are attracted to, and preferentially feed on specific microbial biofilms, and suggest that these feeding behaviors are correlated with the nutritional value of individual bacterial isolates. Introduction Macroinvertebrates are thought to impact microbial communities in freshwater ecosystems through biofilm grazing (Hakenkamp and Morin 2000). In freshwater environments, microorganisms are capable of quickly assimilating nutrients which are stored as microbial biomass and subsequently become available to other organisms in the ecosystem, such as biofilm grazers (Ellis et al. 1998, Craft et al. 2002 Feris et al. 2004, Knapp et al. 2005). Microorganisms being grazed upon may in turn receive the benefit of being transported to new habitats or brought into close proximity with other bacteria in 37 the animal hindgut facilitating processes such as horizontal gene transfer (Dillion and Dillion 2004). Predation on microbial biofilm has been shown to alter the freshwater prokaryotic microbial community (Jardillier et al. 2005). It has been suggested that many factors influence predation by mesofauna on bacteria in aquatic ecosystems. Selective grazing on biofilms by protozoan may be influenced by prokaryotic morphology, cell density, secondary metabolites, or nutritional content (Jardillier et al. 2005, Hahn and Ho fie 2001, Huws et al. 2005). Protozoan grazing has been shown to alter phenotypic characteristics of bacterial prey and prokaryotic community structure, potentially influencing nutrient turnover in freshwater ecosystems (Hakenkamp and Morin 2000, Matz and Kjelleberg 2005, Weitere et al. 2005). A growing body of research is accumulating in the scientific literature addressing specific interactions between protozoan grazers and microbial biofilms in freshwater ecosystems (e.g. Matz and Kjelleberg 2005). However, very little is known regarding the effects of aquatic insect grazing on microbial biofilms (Doi et al. 2005). It has been suggested that in coldwater environments, macroinvertebrate production is limited by nutrient or food limitation (Huryn and Wallace 2000). Macroinvertebrate production is an important trophic linkage, considering that up to 90% of macroinvertebrate production supports higher trophic levels (e.g. insect predators, fish, and birds) (Huryn and Wallace 2000). Previous research has shown that the biology of many aquatic macroinvertebrates is intimately related to bacteria (Huryn and Wallace 2000, Trexler et al. 2003, Dillion and Dillion 2004, Diaz and Trochine 2005, Frost et al.2005). For example, mosquito 38 oviposition response in aquatic ecosystems appears to be mediated by certain bacterial isolates (Trexler et al. 2003). It has also been shown that microorganisms play a key role in the ability of macroinvertebrate “shredders” to break down coarse particulate matter (Meyer 1994, Diaz and Trochine 2005). The Nyack floodplain, West Glacier, MT is a physically and ecologically complex environment comprised of multiple interbraided streambeds, channels, and gravel bars that often change shape, form and location seasonally (Stanford and Ward 1988, Stanford and Ward 1993, Craft et al. 2002). The hyporheic zone present in the subsurface of this region allows for extensive mixing of ground and surface water (Stanford and Ward 1993, Craft et al. 2002). This habitat complexity is considered to be a major driver of the broad ecological diversity that is associated with this habitat, which includes a diverse macroinvertebrate assemblage. Among these are two genera of plecopteran nymphs (stoneflies), Isocapnia sp. and Paraperla sp. that spend much of their life cycle in the hyporheic zone (Stanford and Ward 1993, Stewart and Harper 1996, Craft et al. 2002). Stonefly nymphs are of particular interest in riverine ecosystems because they are one of the most abundant insect orders present, particularly in the winter months (Stewart and Harper 1996). Although feeding may become more specialized in later instars, it is generally accepted that early developmental stages are omnivorous and consume large numbers amounts of bacteria either by direct grazing or by the consumption of leaf litter and other particulate matter colonized by microbial biofilms (Cummin and Merritt 1996, Stewart and Harper 1996). 39 Stonefly nymphs are not visual predators, but are generally believed to respond to tactile or chemotactile cues (Elliot 2004). To date, few studies have been performed on selective feeding by stoneflies. However, adult stoneflies have been found to intentionally ingest flowers and grains, and appear to prefer cyanolichens and cyanophyceae in the winter months (Tiemo De Figueroa and Sachez-Ortega 2000). Mature nymphs have also exhibited preferentially feeding patterns in the laboratory (Elliot 2004). For the current study, we hypothesized that early instar stonefly nymphs would be attracted to and preferentially feed on specific microbial populations when presented with a selection in an arena freely allowing choice. It is well established that many interactions observed between insects and their hosts are mediated by chemical cues (Fraenkel 1959, Schultz 1988, Coumoyer and Boivin 2004). Choice tests and attraction assays have been used extensively to investigate the ecology and behavior of many insects, including herbivores, predators, and parasites (Pszczolkowski et al. 2003, Raffa and Havill 2004). To test our hypothesis, we modified assays previously used to assess attraction between hosts and prey in terrestrial systems (Raffa and Havill 2002, Coumoyer and Boivin 2004) to gain insight into trophic interactions between stoneflies and microbial biofilms in aquatic environments. We created an aquatic Y-olfactometer to assess stonefly nymph attraction toward bacteria via chemical cues in an aqueous environment. We also developed an aquatic choice test arena to determine whether stonefly nymphs preferentially feed on specific populations of microorganisms. The results obtained in these assays demonstrate that plecopteran nymphs are attracted to, and preferentially feed on specific microbial 40 biofilms and we speculate that these feeding behaviors are correlated with the nutritional value of the individual bacterial isolates. Materials and Methods Insect collection. Early instar Isocapnia Banks nymphs were obtained from Beaver Creek and the surrounding hyporheic zone in the Nyack floodplain near West Glacier, Montana (48°50’N, -113°98’E) in the fall of 2004 by pumping a well already emplaced in a region of infiltration of channel water into the subsurface (Craft et al. 2002). Early instar Paraperla Banks nymphs were collected in the winter of 2005 from Petty Creek, near Alberton, MT (46°99’12”N, 114°44’73”) by standard kick-net technique (Cummin and Merritt 1996). Insects were identified through the use of standard taxonomic keys (Stewart and Harper 1996). Bacterial biofilms for behavioral studies. Six different bacterial isolates were chosen for attraction and feeding studies (Table 1). Five of these were from the Nyack floodplain and characterized to the genus level based on 16S rRNA gene phylogenetic affiliation as previously described (Adamset al. 2006). These isolates included a numerically dominant, novelPedobacter sp. 1114, two Pseudomonads (Pseudomonas sp. 115 and Pseudomonas sp. 173), a Flavobacterium sp. 1143, and a Rhodococcus sp. 189. The sixth bacterium assayed was the type strain Aeromonas salmonicida subsp. salmonicida (ATCC #33658; ATCC®, Manassas, VA). Previous work demonstrated that A. salmonicida is found in high frequency in stonefly hindguts contents (Adams et al. 2006), but the indigenous form of this strain proved recalcitrant to cultivation despite multiple attempts. Isolate bacterial biofilms were developed as previously described, (Adams et al. 2006). Briefly, individual filter disks (Polypropylene 0.45 um, 25 mm 41 diameter; Millipore, Billerica, MA) were inoculated with 25 ul of an overnight culture of a given isolate grown in LB Broth at 20°C. Inoculated filter disks were subsequently incubated on a LB-saturated pad of Whatman #1 filter paper (American Membrane Corporation, Ann Arbor, MI) for 48 h at 20°C. Excess medium was removed by two subsequent transfers to Whatman #1 filter paper saturated with filter sterilized Nyack river water. Negative control disks were treated in the same manner except that sterile LB was added to the filter disks instead of bacterial culture. Insect attraction. An aquatic Y- olfactometer was devised to assesParaperla spp. attraction to different microbial biofilms. Experiments were performed at 4°C under indirect red light. The Y-olfactometer consists of two small Petri dishes (right and left arm) connected to one large central Petri dish with ~2 inches of 18mm inner diameter plastic tubing sealed with all joints thoroughly sealed with silicon caulking. Individual biofilm disks or negative control disks (created as described previously) were randomly placed in the right or left-hand dishes. Fifty milliliters of filter-sterilized water was then added simultaneously to each arm to flood the system. The dishes were then pre incubated for 5 min at 4°C to allow diffusion of possible microbial attractants, after which an individual insect was placed in the middle of the flooded central dish and its attraction behavior monitored. Head-to-head competition choice tests were performed between the following microbial isolates: control disk (no biofilm) vs.A. salmonicida^ A. salmonicida vs.Rhodococcus sp. 189; A. salmonicida vs.Flavobacterium sp. 1143; and Flavobacterium sp. 1143 vs.Rhodococcus sp. 189. Individual nymphs were considered to have made a choice when the individual fully entered the tubing connected to the left or right arm. If the nymph did not enter 42 either arm within 5 min, the individual was considered to be non-responsive. Seventeen replications with unique individuals were performed for each trial, with the exception of the A. salmonicida vs. the control disk. Between trials, the Y-olfactometers were rinsed with 10% bleach, rinsed three times with sterilized water, and allowed to air dry. Insect behavior was statistically analyzed using the chi-square goodness-of-fit test. Preferential feeding behavior o f Isocapnia. Nymph sp feeding behavior toward biofilms of different microbial isolates was assessed using three isolates obtained from the Nyack flood plain: Pedobacter sp. 1114; Pseudomonas sp. 115; and Pseudomonas sp. 173, and also the type strain Aeromonas salmonicida subsp.salmonicida. Forty Isocapnia sp.nymphs, starved for 24 to 72 h, were introduced individually into separate 150 x 15 mm Petri dish feeding arenas under red light at 4°C as previously described (Adams et al. 2006). Briefly, each Petri dish contained 75ml of filter-sterilized Nyack river water with one each of the four bacterial isolate disks and one negative control disk evenly spaced in random order around the perimeter of the arena (Fig. 1). For each insect, time spent feeding on each biofilm disk was recorded for a period of 15 min. Signs of insect feeding were defined as a combination of conspicuous head and mouth-part movement on a biofilm surface. A representative example an insect nymph feeding trail is provided in Figure 2. Time spent feeding (in seconds) on each bacterial isolate biofilm was log-transformed and analyzed using multivariate ANOVA (MANOVA) to test for insect feeding preference. Fifteen of the forty insects did not exhibit feeding behavior and were excluded from the analysis. No insects exhibited feeding behavior toward the negative control disks, therefore only time spent feeding upon the four treatment disks having biofilm was included in the analysis. 43 Results and Discussion Stonefly nymphs were attracted to and preferentially fed on bacterial isolates that were previously found to be in high abundance in the hindgut compartment (see Chapter 1). Stonefly taxa demonstrated preference towards Aeromonas salmonicida subsp. salmonicida. Isocapnia preferentially fed on A. salmonicida significantly more than any other bacterial isolate. Paraperla was attracted to A. salmonicida significantly more than almost all other isolates, with the exception of Rhodococcus 5/7.189. The aquatic Y-olfactometer assessed relative rates of attraction of plecopteran nymphs toward different microbial biofilms or a negative control in pair-wise combinations. Paraperla sp. nymphs responded to bacteria tested in the aquatic Y-olfactometer at high rates (Fig. 3). The experiment that paired Rhodococcus 5/7.189 vs. Flavobacterium 5/7.1143 resulted in a nymph response rate of 82%. Nymphs responded positively and evenly to bothA. salmonicida and Rhodococcus 5/7.189 (82% response rate) in the head-to-head competition. Nymphs never showed a significant preference for Flavobacterium 5/7.1143 in any of the pairings (Fig. 4). These response rates are similar those obtained when a Y-olfactometer is used to assess terrestrial insect interactions (Pettersson 2001). The preferential feeding assay scored the amount of time that individual nymphs exhibited feeding when presented with a suite of different microbial biofilms. Isocapnia sp. nymphs responded to the feeding behavior assay at a rate of 62.5%. Insects preferred A. salmonicida to any other isolate (Fig 4), with the average individual feeding for 44 35.7±8.8 seconds. The nymphs also significantly preferred feeding on Pedobacter sp. Ill 4 over the twoPseudomonas sp. 173 and 115. Preliminary data from an ongoing study of C:N:P stoichiometry by A. Valenzuela leads us to suggest that the behaviors exhibited in the current study, by bothIsocapnia sp. and Paraperla sp. toward microbial biofilms appear to be correlated with nutrient content, as indicated by C:N ratio, of the isolates. Both early instar stonefly nymph species were most highly attracted to bacterial species with a C:N ratio close to or exceeding 5:1, which is most similar to their own (Table 1). The isolates with the highest C:N ratios, Rhodococcus sp. 189 (C:N = 7.8),A. salmonicida (C:N = 4.6), and Pedobacter sp. II14(C:N = 5.1) were chosen significantly more by nymphs than the other isolates (Figs. 3 and 4). Conclusions Results of our previous studies indicate that microbial diversity in the hindguts of stoneflies is limited relative to that observed in the surrounding native sediment (see Chapter 1). Aeromonas salmonicida subsp.salmonicida was found in relatively high abundance inParaperla sp. hindguts, yet was only indicated once in our 16S rRNA gene survey of the bulk sediment with >4,000 sequences analyzed, and was not encountered at all in Isocapnia sp.hindguts. However, both stonefly taxa responded positively to this bacterium. This phenomenon maybe explained by the nutritional requirements needed by immature insects to molt. Tyrosine is critical to the sclerotization and pigmentation of insect cuticle (Kramer and Hopkins 2005). A. salmonicida is known to secrete a highly pigmented secondary metabolite, containing tyrosine (Koppang et al 2000). 45 In addition, our data and those of A. Valenzuela suggest that plecopteran nymphs are attracted to, and preferentially feed on microbial biofilms that have C:N ratios of-5.0 and higher. These results are consistent with the feeding behaviors of other insect nymphs. For example, grasshopper nymphs preferred diets high in carbon compounds rather than protein in laboratory experiments (Chown and Nicolson 2004). It is suggested that nymphs require higher levels of carbon than adults in order to molt. As the nymph matures, and becomes an adult, the feeding behavior changes and protein is preferred, in order to produce offspring (Chown and Nicolson 2004). The data presented here intriguingly suggest that stonefly nymphs maybe capable of altering aquatic prokaryotic community structure in hyporheic and other aquatic environments through selective grazing. Both stonefly taxa were attracted to and preferred feeding upon the known fish pathogen Aeromonas salmonicida subsp. salmonicida over the other isolates. Controversy has surrounded the ability of this bacterium to survive in the open environment (Austin 1997). Potentially, a key to understanding A. salmonicida, its transmission, and the disease it can cause, can be discovered through the observation of selective feeding patterns by stonefly nymphs. 46 Figure 1 Example of Nymph feeding behavior.Isocapnia sp. Individuals were placed in the middle of the feeding arena portrayed in the figure and permitted to feed freely for 15 min on whichever isolates were preferred. Isolates included in the assay: Pseudomonas spp. 115 and 173, Pedobacter sp. II14, and Aeromonas salmonicida subsp. salmonicida. This example amply demonstrates the attraction and feeding preference toward of the insects toward individual isolates. The insect moved primarily between the novelPedobacter sp. II14 and the type strain of Aeromonas salmonicida subsp. salmonicida (see movements 2-10). In addition, this individual spent more cumulative time feeding on A. salmonicida than any other bacterial biofilm. Although all disks were visited, nymphs did not exhibit signs of feeding behavior toward the negative control disk. Move Time Isolate ment (sec) 1 173 4 2 1114 3 A. Salmonicida A. 3 17 salmonicida 4 1114 4 A. 5 15 salmonicida 6 115 5 A. 7 25 salmonicida 8 1114 13 173 9 173 6 A. 10 18 salmonicida 11 1114 8 47 Figure 2 Paraperla sp. attraction. Response of individual nymphs in the aquatic Y-olfactometer to pairs of microbial isolates. Experiment A tested nymph attraction to Aeromonas salmonicida subsp.salmonicida vs. a negative control. Experiment B tested A. salmonicida vs.Flavobacterium sp. 1143. Experiment C tested A salmonicida vs. Rhodococcus sp. 189. Experiment D tested Rhodococcus sp. 189 vs.Flavobacterium sp. 1143. Asterisks indicate a significant response to the treatment as determined by the goodness of fit chi-test (* P < 0.005). Exoeriment No response A * A. salmonicida 43% B A. salmonicida Flcn>obacterium i 35% sp. I 43 A. salmonicida Rhodococcus sp. 189 18% Flavob acterium sp. 18% D Rhodococcus sp. 189 1143 90 80 70 60 50 40 30 20 10 0 10 20 30 40 50 60 70 80 90 Percent Response 48 Figure 3 Insect preferential feeding behavior. Isocapnia sp. nymphs were significantly attracted to A. salmonicida over all other bacterial isolates available in the choice test assay (n = 25). Error bars represent one standard error (P<0.001) 4 5 A. salmonicida 4 0 c W) Pedobacter sp.II14 T 3 5 - a6X 3 0 - 2 5 - Pseudomonas sp.I73 Va a 20 a 0> E Pseudomonas sp.JI5 1 5 H a i ____ ex T ua 10 a>> < 5 ■ Biofilm 49 Table 1 Carbon and nitrogen (C:N) ratios of bacterial isolates and stoneflies. Samples 1-6 are bacterial isolates used in the insect attraction and feeding experiments. Samples 7 and 8 are the stonefly species used (Data courtesy of A. Valenzuela and J. Gannon). Organism C:N ratio 1. Pseudomonas 577.173 4.353 ± 0.062 2. Pseudomonas 577.115 4.303 ± 0.093 3. Pedobacter 577.114 5.120 ±0.009 4. Flavobacterium 57?.II43 4.270 ±0.017 5. Aeromonas salmonicida subsp. salmonicida 4.560 ± 0.793 6. Rhodococcus 577.189 7.837 ±T).150 5.7 ± 0.8 (Evans-White et 7. Isocapnia sp. al. 2005) 8. Paraperla sp. 5.322 ±0.408 50 References 1. Adams, S.A., Adams, A.S., and Holben. In preparation, 2006. The deadly meal: Bacterial pathogen transmission to rainbow trout via stoneflies. 2. Craft, J.A., Stanford, J., and Pusch, M. 2002. Microbial respiration within a floodplain aquifer of a large scale gravel-bed river.Freshwater Biology. 47:251- 261. 3. Coumoyer, M. and Boivin, G. 2004. Infochemical-mediated preference behavior of the parasitoid Microctonus hyperodae when searching for its adult weevil hosts. Entomologia Experimentalis et Applicata. 112: 117—124. 4. Chown, S.L. and Nicolson, S.W. 2004. Nutritional Physiology and Ecology. Pages 22-23 in S.L. Chown and S.W. Nicolson, editors. Insect Physiology and Ecology. Oxford University Press, Inc., New York. 5. Cummin, K. W., and R. W. Merritt. 1996. Ecology and Distribution of Aquatic Insects. Pages 74-86 in K. W. Cummins and R. W. Merritt, editors. Aquatic Insects of North America. Kendall/Hunt Publishing Company, Dubuque. 6. Diaz, V. and Trochine, C. The role of microorganisms in the diest of Verger CF limnophilus (Trichoptera: Limnephilidae) larvae in a Patagonian Andean temporary pond. Wetlands. 25: 125-127. 7. Dillion, R.J., and Dillion, V.M. 2004. The gut bacteria of insects: Nonpathogentic interactions. Annu Rev Entomol. 49:71-92. 8. Doi, H., Katano, I., and Kikuchi, E. 2005. The use of agal-mat habitats by aquatic insect grazers: effects of microalgal cues. Basic and Applied Ecology (in press). 51 9. Duncan, S.H., Hold, G.L., Harmsen, HStewart, C.S. and Flint, H.J. 2002. Growth requirements and fermentation products of Fusobacterium prausnitzii, and a proposal to reclassify it as Faecalibacterium prausnitzii gen. nov., comb, nov.Int JSyst Evol Microbiol. 52: 2141-2146. 10. Elliot, J. 2004. Prey switching in four species of carnivorous stoneflies. Freshwater Biology. 49:709-720. 11. Ellis, B., Stanford, J., and Ward, J. 1998. Microbial assemblages and production in alluvial aquifers of the Flathead River, Montana, USA.J N Am Benthol Soc. 17:382-402. 12. Evans-White, M.A., Stelzer, R.S., and Lamberti, G.A. 2005. Taxonomic and region patterns in benthic macroinvertebrate elemental composition in streams. Freshwater Biology. 50: 1786-1799. 13. Feris, K.P., Ramsey, P.W., Frazar, C., Rillig, M., Moore, J.N., Gannon, J.E., and Holben, W.E. 2004. Seasonal dynamics of shallow-hyporheic-zone microbial community structure along a heavy-metal contamination gradient. Appl Environ Microbiol. 70:2323-2331. 14. Fraenkel, G.S. 1959. The raison d’Etre of secondary plant substances. Science 129:1466-1470. 15. Frost, P.C., Cross, W.F., and Benstead, J.P. 2005. Ecological stoichiometry in freshwater benthic ecosystems: an introduction. Freshwater Biology. 50: 1781- 1785. 16. Hahn, M.W., and Ho fie, M.G. 2001. Grazing of protozoa and its effect on populations of aquatic bacteria. FEMS Micrbiol Ecol. 35:175-185. 52 17. Hakenkamp, C.C., and Morin, A. 2000. The importance of meiofauna to lotic ecosystem functioning. Freshwater Biology. 44: 165-175. 18. Huryn, A. D., and Wallace, J. B. 2000. Life history and production of stream insects. Annu Rev Entomol. 45:83-110. 19. Huws, S.A, McBain, A.J.., and Gilbert, P. 2005. Protozoan grazing and its impact upon population dynamics in biofilm communities. JApp Micro. 98: 238-244. 20. Jardillier, L., Boucher, D., Personnic, S., Jacquet, S., Thenot, A., Sargos, D., Amblard, C., and Debroas, D. 2005. Relative importance of nutrients and mortality factors on prokaryotic community composition in two lakes of different trophic status: Microcosm experiments. FEMS Micrbiol Ecol. 53:429-443. 21. Knapp, C., Caquet, T., Hanson, M., Lagadic, L., and Graham, D. 2005. Response of water column microbial communities to sudden exposure to deltamethrin in aquatic mesocosms. FEMS Micrbiol Ecol. 54:157-165. 22. Koppang, E.O., Fjolstad M., Melgard, B., Vigerust, M., and 0rum, H.S. 2000. Pigmentation (melanin) by Aeromonas salmonicida subsp salmonicida formed through the modification of tyrosine. J Fish Diseases. 23:39-48. 23. Kramer, K.J., and Hopkins, T.L. 2005. Tyrosine metabolism for insect cuticle tanning. Archives of Insect Biochemistry and Physiology. 6: 279-301. 24. Matz, C., and Kjelleberg, S. 2005. Off the hook-how bacteria survive protozoan grazing. Trends in Microbiology. 13:302-307. 25. Merritt, Cummins, and Resh. 1996. Design of aquatic insect studies: Collecting, sampling, and rearing procedures. Pages 74-86 in K. W. Cummins and R. W. 53 Merritt, editors. Aquatic Insects of North America. Kendall/Hunt Publishing Company, Dubuque. 26. Meyer, J.L. 1994. The microbial loop in flowing waters. Microbial Ecology. 27. Moriarty, D.W. 1997. The role of microorganisms in aquaculture ponds. Aquaculture. 151:333-349. 28. Pattersson, E.M. 2001.Volatile attractants for three Pteromalid parasitoids attacking concealed spruce bark beetles.Chemoecology. 11:89-95 29. Polis, G.A., and Strong, D.R. 1996. Food web complexity and community dynamics. Am Nat. 147:813-846. 30. Pszczolkowski M.A., Zahand. A, Bushman S.M, and Brown J.J. 2003. Effects of calcium and glutamate receptor agonists on leaf consumption by lepidopteran neonates. Pharmacology Biochemistry Behavior 74:389-942. 31. Raffa, K.P. and Havill, N.P. 2002. How many choices can your test animal compare effectively? Evaluating a critical assumption of behavioral preference tests. Oecologia 133:422-429. 32. Trexler, J., Apperson, C., Zurek, L., Gemeno, C., Schal., Kaufman, M., Walker, E., Watson, D., and Wallace, L. 2003. Role of bacteria in mediating the oviposition responses ofAedes albopictus (Diptera: Culicidae). J Med Entomol. 40:841-848. 33. Schultz, J.C. 1988. Many factors influence the evolution of herbivore diets, but plant chemistry is central. Ecology 69:896-897. 34. Stanford, J., and Ward, J. 1988. The hyporheic habitat of river ecosystems. Nature. 335:64-66 54 35. Stanford, J. and Ward, J. 1993. An ecosystem perspective of alluvial rivers: connectivity and the hyporheic corridor. JNAm Benthol Soc. 12:48-60. 36. Stewart, K.W and Haper, P.P. 1996. Plecoptera. Pages 217-266 in K. W. Cummins and R. W. Merritt, editors. Aquatic Insects of North America. Kendall/Hunt Publishing Company, Dubuque, IA. 37. Tiemo De Figueroa, J., and Sachez-Ortega, A. 2000. Imaginal feeding of twelve Nemouroidean stonefly species (Insecta, Plecoptera). Arthropod Biology. 93:251- 253. 38. Trexler, J.D., Apperson, C.S., Zurek, L., Gemeno, C., Schal., Kaufman, M., Walker, E., Watson, D.W., and Wallace, L. 2003. Role of bacteria in mediating the oviposition responses ofAdes albopictus (Diptera:Culicidae). Journal of Medical Entomology. 39. Weitere, M., Bergfeld, T., Rice, S., Matz, C., and Kjelleberg, S. 2005. Grazing resistance of Pseudomonas aeruginosa biofilms depends on type of protective mechanism, developmental stage and protozoan feeding mode. Environmental Microbiology. 7:1593-1601. 55 Chapter 3 The deadly meal: Bacterial pathogen transmission to rainbow trout via stoneflies Summary It has been predicted that the incidence of vector-borne disease will increase as temperatures rise due to global warming (Zell 2004). However, the increased incidence of vector-borne illness is difficult to assess, in part because of the complexity of “pathogen transmission dynamics” (Zell 2004). In addition, with bacterial strains becoming increasingly resistant to antimicrobial agents (Gootz 2006), the need to investigate and understand pathogen interactions with organisms other than susceptible hosts is timely. As a model of bacterial pathogen transfer to susceptible hosts via trophic interactions, we examined the relationship between the bacterium Aeromonas salmonicida subsp. salmonicida, stonefly nymphs (Plecoptera: Chloroperlidae), and rainbow trout (Oncorhynchus mykiss), an opportunistically susceptible host where A. salmonicida can cause the disease furunculosis. In this study, we investigated whether stonefly nymphs can act as a reservoir and vector forA. salmonicida transfer to fish. Feeding assays were conducted in which stonefly nymphs were presented with a suite of different bacterial isolates. Nymphs preferred to feed onA. salmonicida significantly more than all other bacteria presented, indicating specific attraction to this bacterium. In subsequent experiments, viableA. salmonicida were isolated from both hindguts and frass obtained from nymphs fedA. salmonicida, indicating that stoneflies may act as a 56 vector for transmission of this pathogen to its fish hosts. Finally, rainbow trout juveniles were presented with stonefly nymphs fed a type strain A.of salmonicida, stonefly nymphs naturally harboringA. salmonicida or, as a control, commercial fish diet to determine the vectoring capacity of the nymphs. Both stonefly treatments resulted in expression of disease in the trout, and stonefly nymphs naturally harboring the pathogen appeared to be even more effective at transmitting disease than stoneflies fed the commercial type strain ofA. salmonicida. The physiology and pathology of the fish pathogen Aeromonas salmonicida subsp.salmonicida has been studied extensively. However, its activity and transmission in natural environments have yet to be fully elucidated. This opportunistic pathogen is capable of causing the disease furunculosis (Fig 1), which can result in significant mortality in farm-raised salmonids and other freshwater and marine fish (Bemoth 1997, Smith 1997, Boyd et al 2003, Pirhonen et al. 2003, Ringo et al. 2004). A. salmonicida has been believed to gain entry into host fish through the gills, mouth, anus, or through an epidermal surface injury (Austin 1997). To date, there has been no reported vector forA. salmonicida transmission to fish, and it has been assumed that naive fish populations acquire infection through contact with infected fish (Ogut and Reno 2005). It has been suggested that a clearer understanding of how A. salmonicida is capable of surviving outside the fish host in natural freshwater environments is necessary, not only to gain insight into the mechanisms by which it is able to cause disease, but also to assess risk of the disease to native fish in open environments (Hiney etal. 2002). Aeromonas salmonicida was one the earliest fish pathogens studied and was isolated from symptomatic fish over a century ago (Gustafson et al. 1992, Bemoth 1997). 57 It was originally believed thatA. salmonicida was an obligate pathogen, incapable of surviving outside of a fish host (Gustafson etal. 1992, Morgan et al. 1993). It is now understood that this pathogen is free-living and capable of causing disease only in susceptible hosts, typically those living under stressful conditions such as high water temperature and crowding, which are often associated with fisheries and fish farming) (Pickering 1997). However, this bacterium is not numerically dominant in the environment and is often detected in aquatic sediments only in low numbers through the use of molecular techniques (Morgan et al. 1993, O’Brien et al. 1994, Austin 1997). As a consequence, the ecology of this pathogen and its role in freshwater ecosystems has been largely unstudied (Morgan et al. 1992, Gustafson et al. 1993, Enger 1997, and Smith 1997). Due, in part, to the difficulty in culturing viableA. salmonicida from sediment and river environments, there are numerous questions regarding its persistence in the environment and its mode of infection (Austin 1997). Control of this pathogen in fish hatcheries has typically relied heavily on the use of antimicrobial compounds and current research in this arena is focused on discovering novel therapeutic drugs (Bansemir etal. 2006). Stoneflies are regarded as one of the most numerous insect orders in freshwater ecosystems (Stewart and Stark 1988). Although the feeding behavior of stoneflies may become more specialized in latter instars, it is generally accepted that early instars graze on microbial bio films (Stewart and Stark 1996). Further, stonefly nymphs are considered to be one of the first and most abundant macroinvertebrates to colonize dead salmon in aquatic ecosystems (Chaloner et al.2002). Interestingly, diseased or dead fish are considered a major source of A. salmonicida in the wild (Enger et al. 1997). Indeed, 58 there are many opportunities for stonefly nymphs to interact with A. salmonicida in the environment. In the current study, we hypothesized that A. salmonicida utilizes stonefly nymphs as a host reservoir, which may facilitate amplification of the pathogen to infectious titers, thereby representing a mode of transmission for A. salmonicida to fish hosts. To investigate the relationship between stoneflies and microbial communities in aquatic ecosystems, we probed the hindgut contents of stoneflies from two freshwater ecosystems in Western Montana. First, we performed random cloning of partial 16S (prokaryotic) rRNA gene sequences on the hindgut contents of nymphs from the Nyack floodplain, West Glacier, MT. Thirty-three percent of the individuals tested were found to naturally harbor bacteria with a sequence identical to the known fish pathogen A. salmonicida. Additionally, DNA isolated from hindgut contents of individual stoneflies from Petty Creek, Alberton, MT were probed with A. salmonicida specific primers11,24 and were found to naturally harbor A. salmonicida at a frequency of 58%. By contrast, the results of an extensive 16S rRNA survey of the sediment microbial community inhabiting the Nyack floodplain showed that A. salmonicida was rare, being detected once out of >4,200 cloned sequences (data not shown). While molecular techniques showed that A. salmonicida was present in the sediment and hindguts of stoneflies in western Montana, attempts to isolate viableA. salmonicida from these environments were unsuccessful, presumably due to its fastidious nature (Austin 1997, Enger 1997). Therefore, a readily culturable type strain was utilized {Aeromonas salmonicida subsp. salmonicida ATCC #33658; ATCC®, Manassas, VA) was utilized in subsequent feeding and disease transmission experiments. 59 Despite the observation thatA. salmonicida was not numerically dominant the sediment from the Nyack floodplain, stonefly nymphs from the region preferentially fed on A. salmonicida when presented with a suite of 4 different bacterial isolates in feeding choice assays. In this experiment, 24 of 40 stoneflies responded to the behavior assay (a response rate of 60%) and preferentially fed on A. salmonicida more than all other bacteria tested (P<0.005, Fig. 2). In a subsequent experiment, thirteen stonefly nymphs were fed a diet consisting solely of A. salmonicida for five days. Viable Asalmonicida cells were obtained from all seven stonefly hindguts sampled, and also from all six frass collections made (data not shown). This suggests that stonefly nymphs may play a pivotal role as a reservoir maintaining and transporting populations ofA. salmonicida in the environment. Stonefly nymphs not only act as a reservoir for A. salmonicida, but are also able to act as a vector. We performed an experiment with three replicated treatment groups to test the hypothesis that stoneflies are capable of vectoring A. salmonicida to trout. These included: Group 1) trout fed stoneflies with a natural incidence (58%) of wild-type A. salmonicida; Group 2) trout fed stoneflies maintained on a diet consisting solely of the type strain A. salmonicida (positive control); and Group 3) trout fed commercial trout diet (negative control). Group 1 became symptomatic with furunculosis after two days of treatment, resulting in acute infection and ultimately death. Group 2, exhibited disease symptoms 4 days after initiation of treatment. After 1 week, Group 3 exhibited no outward signs of infection. Further all individuals in Group 3 tested negative in PCR- based assays of liver tissue with theA. salmonicida primers, while all individuals in Groups 1 and 2 tested positive. The observed symptoms of acute infection in both 60 stonefly treatments were consistent with induced infections where salmonid fish are fed a diet saturated with, injected with, or bathed in A. salmonicida (Siwicki et a l 1994, Vipond et al. 1998, Ogut and Reno 2005). Also consistent with other research findings (Stuber etal. 2003), the native strains ofA. salmonicida (harbored in the hindguts of the local stoneflies) were more virulent than the commercially available type strain used as positive control. Aeromonas salmonicida is known to harbor multiple plasmids, at least some of which are associated with the virulence, pathogenesis and symptoms of furunculosis (Stuber etal. 2003). While current methods of control of A. salmonicida infection rely on antimicrobial agents (Ellis 1999), it has become apparent that bacterial pathogens can readily develop resistance to chemotherapeutics through acquisition of resistance genes via horizontal gene transfer (Ochman and Moran 2001, Gootz 2006). This is highly salient to this system since insect hindguts have been shown to be “hot spots” for gene transfer between bacteria (Dillion and Dillion 2004), and genetic information can readily be transferred between strains ofA. salmonicida and possibly other species via plasmid exchange (Noonan and Trust 1995). Stonefly nymphs clearly play a role in the transportation of A. salmonicida in the aquatic environment. Confusion and controversy have always surroundedA. salmonicida and the disease furunculosis (Austin 1997, Bemoth 1997). To our knowledge, this is the first study to show that stoneflies are capable of vectoring A. salmonicida to trout. We suggest that studies like ours, which investigate multitrophic interactions, may not only lend insight into basic ecological processes, but may also aid in understanding the 61 strategies and mechanisms for survival and transmission of opportunistic pathogens in the environment. Conclusion While it is clear from our data that stoneflies can vector A. salmonicida, and thus furunculosis to salmonid fish, this disease is considered to be an opportunistic pathogen that causes disease only under certain conditions. In winter months, stoneflies are believed to constitute a large portion of the trout diet (Duffield and Nelson 1998). However, as the temperature of surface waters rises in summer months, stoneflies become less abundant. This suggests that when trout are most susceptible to the disease, i.e. warm water temperatures (Pickering 1997), the stonefly vectors are not as readily available as a food source. This may explain why large numbers of native fish do not succumb to furunculosis in the natural environment and it is largely confined to fish maintained under crowded, warm, artificial conditions. Importantly, as temperatures increase due to current global warming trends and environmental dynamics subsequently shift, the balance of these trophic interactions may be upset, ultimately affecting the dynamics of this and perhaps other diseases. Methods Early instar Paraperla frontalis Banks nymphs were obtained from the hyporheic zone in the Nyack floodplain, West Glacier, MT in fall 2004 by straining of water pumped from a groundwater well. Plecopteran nymphs were collected from Petty Creek, near Alberton, MT using standard kick-net technique in winter 2005. 62 Preferential feeding behavior of stonefly nymphs toward bacterial isolates was assessed using a suite of bacterial isolates: a Pedobactera sp.1114 and two Pseudomonas spp. (115 and 173) from the Nyack site, and the commercially available type strain Aeromonas salmonicida subsp.salmonicida (ATCC #33658, ATCC®, Manassas, VA). Microbial isolate biofilms were developed by inoculating individual filter disks (Polypropylene 0.45pm, 25mm diameter; Millipore, Billerica, MA) with 25 pi of an overnight culture grown in LB Broth (Bacto®, Detroit, M I) at 20°C, then incubated on a LB-saturated pad of Whatman filter paper (American Membrane Corporation, Ann Arbor, MI) for 48h at 20°C. Excess medium was removed by two subsequent transfers to filter-sterilized river water-saturated Whatman filter paper. Negative control disks were treated in the same manner except that sterile LB medium was substituted for bacterial culture. For the preferential feeding assay, forty stonefly nymphs were starved for 24h, then introduced individually into the center of separate 150 x 15mm Petri dishes under indirect red light at 4°C. Each Petri dish contained one of each of the 4 bacterial isolate biofilm disks and one negative control evenly spaced around the perimeter in random order and covered with 75ml of filter-sterilized river water to create an aqueous feeding arena. Each insect was allowed 15min to feed and feeding time was recorded only when the insect exhibited visual signs of feeding behavior (i.e. moving headparts). Recovery and analysis of insect hindgut microbial community DNA was performed on 9 stoneflies collected in fall 2004. Hindguts were removed and the contents subjected to random cloning as previously described (Holben etal. 2002). Ten clones from each insect were randomly selected for DNA sequence analysis to determine 63 the predominant bacterial population(s) present. Phylogenetic relationships to known bacteria were then established using the RDPII database (Maidak etal. 2001). In addition, hindgut contents from 12 individual stonefly nymphs collected from the Petty Creek site in winter 2005 were probed with A. salmonicida specific primers (Hiney et al. 1992, Gustafson et al. 2002, Hiney et al. 2002). Hindgut dissection and frass collection was performed on 13 nymphs from the Nyack site to determine whether A. salmonicida could pass through insects in viable form. To achieve this, sediment from the Nyack floodplain was sieved through wire mesh (2.54 x 5.08 cm), washed and autoclaved 3X. Sterile sediment (500 ml) was inoculated with 400ml of an overnight culture of A. salmonicida (~1 x 109 cells/ml) grown in TSB (Bacto®, Detroit, MI), then incubated at 20°C for 2 days. The media was drained away and the sediment placed in a uniform layer in 150 mm X 15 mm petri dishes with 75 ml of filter-sterilized river water. Stonefly nymphs were placed in individual dishes and maintained at 4°C for five days to feed on theA. salmonicida type strain biofilm. Eight nymphs were randomly chosen for hindgut dissection, while 7 were used for frass collection. Hindgut contents were spread on TSA plates and incubated overnight at 20°C. For frass collection, insects were surface-washed with sterile dH20, and transferred to sterile filters soaked with sterile river water. Frass was collected from the filters, plated onto TSA, and incubated overnight. To verify the identity of putativeA. salmonicida, molecular fingerprint analysis was performed using rep-PCR with the BOX1AR primer (Hulton et al. 1991). Hindgut or frass isolates producing fingerprints 64 >95% similar to the type strain using BioNumerics v4.5 software (Applied-Maths, Austin, TX) were presumed to beA. salmonicida. Vectoring o A. f salmonicida to trout via stoneflies was tested using juvenile (80 to lOOg) rainbow trout obtained from the Jocko Fish Hatchery, Arlee, MT in February 2006 (IACUC 038-05WHDBS-112905). This hatchery has no history of furunculosis. Two fish were randomly selected before start of the experiment and their liver and kidney tissues were probed with A. salmonicida primers to ensure that chronic A. salmonicida was not present in the fish population. Each treatment was performed in duplicate aquaria, each containing two individuals. For each treatment, animals were held in 10 gallon closed-system aquaria at 20°C and equilibrated for 72 hrs before the experiment began. Husbandry was the same for all treatments; tanks were skimmed daily and conditioned water was exchanged daily. The animals were fed ~1% of their body weight daily of the appropriate feed treatment; Group 1) Negative control diet (Rangens Fish Food, Buhl, ID); Group 2) Test insects; stonefly nymphs fed on sediments containing native microbial populations; Group 3) Positive control insects; stonefly nymphs fed on A. salmonicida type strain as described above. Fish were euthanized in 300 mg/L MS 222, TMS (Acros Organics, Geel, Belgium) after symptoms of disease (i.e. inability to swim and inflamed anus) were observed. Dissection occurred at the end of experiment and liver tissue was harvested for PCR-based analysis as already described. DNA extraction was performed on livers with Qiagen Blood and Cell Culture DNA Midi Kit (Qiagen, Valencia, CA) and probed with A. salmonicida primers (Gustafson et al. 2002, Hiney et al. 2002). 65 Figures Figure 1: Classic symptom of the disease furunculosis caused byAeromonas salmonicida subsp.salmonicida. The term furunculosis is derived form the furuncules or deep ulceration of tissue often observed in infected fish. Photograph by R. Cipriano. 66 Average time spent feeding (sec) 35 401 20 25 30 10 15 0 5” these were excluded from the analysis. the from excluded were these 2 Figure than any other bacterial isolate. Average time spent feeding on each each on feeding spent time Average isolate. bacterial other any than isolate is shown (n=24). Error bars represent one standard error. Time Time error. standard one represent bars Error (n=24). shown is isolate assay, but no insects fed upon the negative control disks and therefore therefore and disks control negative the upon fed the in insects no included but were assay, disks control Negative not analysis. were the in and included respond, not did were sixteen insects however, Forty assay, in included variance. the as bacteria with multivariate a (MANOVA), using design analyzed and log-transformed was (sec) feeding (Paraperla Pedobacter sp. Pedobacter Stonefly feeding behavior towards bacterial isolates. bacterial towards behavior feeding Stonefly sp.) feed significantly (P<0.001) more on on more (P<0.001) significantly feed sp.) b Pseudomonas sp. Pseudomonas II14 II14 Stonefly Feeding Behavior Feeding Stonefly ab T 67 P<0.005 Biofilm 115 salmonicida Aeromonas D seudomonas sp.ll?> Dseudomonas . salmonicida A. . „„ a Nymphs Nymphs References 1. Austin, B. 1997. Progress in understanding the fish pathogen Aeromonas salmonicida. Mar Biotechnol 15: 131-134. 2. Bansemir, A., Blume, M., Schroder, S., and Lindequist, U. 2006. Screening of cultivated seaweeds for antibacterial activity against fish pathogenic bacteria. Aquaculture 252: 79—84. 3. Bemoth, E. M. 1997. Furunculosis: The History of Disease and of Disease Research. Pages 1-20 in Bemoth, E.M., Ellis, A., Midtlyng, P.J., Oliver, G., and Smith, P., editor. Furunculosis: Multidisciplinary Fish Disease and Research. Academic Press, Inc, San Diego. 4. Boyd, J. Williams, J., Curtis, B., Kozera, C., Singh, R., and Reith, M. Three small, cryptic plasmids from Aeromonas salmonicida subsp.salmonicida A449. Plasmid 50: 131-144. 5. Chaloner, D.T., Wipfli, M.S., and Caouette, J.P. 2002. Mass loss and macroinvertebrate colonization of Pacific salmon carcasses in south-eastern Alaskan streams. Freshwater Biology 47: 263-273. 6. Dillion, R.J., and Dillion, V.M. 2004. The gut bacteria of insects: Nonpathogentic Interactions. Annu Rev Entomol 49:71-92. 7. Duffield, R.M., Nelson, C.H. 1998. Stoneflies (Plecoptera) in the diet of brook trout (Salvelinus frontinalis Mitchell) in Libby Creek, Wyoming, USA. Hydrobiologia 380: 59-65. 8. Ellis, A.E. 1999. Immunity to bacteria in fish. Fish Shellfish Immunol 9: 291-308. 68 9. Enger, 0 .1997. Survival and inactivationAeromonas of salmonicida outside the host-a most superficial way of life, in Bemoth, E.M., Ellis, A., Midtlyng, P.J., Oliver, G., and Smith, P., editor. Fumnculosis: Multidisciplinary Fish Disease and Research. Academic Press, Inc, San Diego. 10. Gootz, T.D. 2006. The forgotten Gram-negative bacilli: What genetic determinants are telling us about the spread of antibiotic resistance. Biochem Pharmacol 71: 1073-1084. 11. Gustafson, C.E., Thomas, C.J., and Trust, T.J. 1992. Detection of Aeromonas salmonicida from fish by using polyermase chain reaction amplification of the vimlence surface array protein gene. Appl Environ Microbiol 58: 3816-3825. 12. Hiney, M., D. Daawson, D. Heery, P. R. Smith, F. Gannon, and R.Powell. 1992. DNA Probe for Aeromonas salmonicida. Appl Environ Microbiol 58:1039-1042. 13. Hiney, M., C. Stanley, L. Martin, A. Cooney, and P. Smith. 2002. The influence of sediment type on the survival of colony forming ability ofAeromonas salmonicida in laboratory microcosms. Aquaculture 212:11-19. 14. Holben, W.E., Williams, P., Saarinen, Sarkilahti, M., and Apajalhti, J.H.A. 2002. Phylogenetic Analysis of Intestinal Micro flora Indicates a Novel Mycoplasma Phylotype in Farmed and Wild Salmon. Microb Ecol 44:175-185. 15. Hulton, C.S., Higgins, C.F. and Sharp, P.M. 1991. ERIC sequences: a novel family of repetitive elements in the genomes of Escherichia coli, Salmonella typhimurium and other enterobacteria. Mol Microbiol 5: 825-834. 69 16. Maidak, B.L., Cole, J.R., Lilbum, T.G., Parker, C.T., Saxman, P.R., Farris, R.J., Garrity, G.M., Olsen, G.J., Schmidt, T.M., and Tiedje, J.M. 2001. The RDP-II (Ribosomal Database Project).Nucleic Acid Research 29:173-174. 17. Morgan, J.A.W., Rhodes, G., and Pickup, R.W. 1993. Survival of nonculturable Aeromonas salmonicida in Lake water. Appl Environ Microbiol 59: 874-880. 18. Noonan, B., and Trust, T. 1995. The molecular biology of Aeromonas salmonicida. Annual Review of Fish Diseases 5:95-111. 19. O’Brien, D., Mooney, J., Ryan, D., Powell, E., Hiney, M., Smith, P.R., and Powell, R. 1994. Detection of Aeromonas salmonicida, causal agent of furunculosis in salmonid fish, from the tank effluent of hatchery-reared Atlantic salmon smolts. Appl Environ Microbiol 60: 3874-3877. 20. Ochman, H. and Moran, N.A. 2001. Genes lost and found: Evolution of bacterial pathogenesis and symbiosis. Science 292: 1096-1099. 21. Ogut, H. and Reno, P.W., 2005. Evaluation of an experimental Aeromonas salmonicida epidemic in Chinook salmon, Oncorhynchus tshawytscha (Walbaum).J Fish Dis 28: 263-269. 22. Pickering, A.D. 1997. Husbandry and stress. In Bemoth, E.M., Ellis, A., Midtlyng, P.J., Oliver, G., and Smith, P. editors Fumnculosis: Multidisciplinary fish disease research, eds Academic Press, San Diego, CA. 23. Pirhonen, J., Schreck, C. B ., Reno, P. W ., and Ogut, H. 2003. Effect of fasting on feed intake, growth and mortality of chinook salmon, Oncorhynchus tshawyscha, during an induced Aeromonas salmonicida epizootic. Aquaculture 216:31-38. 70 24. Ring 0 , E., Jutfelt, F., Kanapathippillai, P., Bakken, Y., Sundell, K., Glette, J., Mayhew, T. M., Myklebust, R. and Olsen, R. E.. 2004. Damaging effect of the fish pathogen Aeromonas salmonicida subsp.salmonicida on intestinal enterocytes of Atlantic salmon (Salmo salar L.). Cell Tissue Res 318:305-311. 25. Smith, P., Hiney, M., Gilroy, D. Padley, D., Gallagher, T., Cotter, D., and Powell, R. 2003. Confusion generated by the validation of the application of an enzyme linked immunosorbent assay and a polymerase chain reaction method for the detection of Aeromonas salmonicida in field samples. Aquaculture 218:67-79. 26. Stewart, K. W., and B. P. Stark. 1988. Nymphs of North American stonefly genera (Plecoptera). Entomological Society of America, College Park. 27. Stuber, K., Burr, S.E., Braun, M., Wahli, T., and Frey, J. 2003. Type III secretion genes in Aeromonas salmonicida subsp.salmonicida are located on a large thermolabile virulence plasmid.J Clin Microbiol 41: 3854-3856. 28. Siwicki, Anderson D.P., and Rumsey, G.L. 1994. Dietary intake of immunostimulants by rainbow trout affects non-specific immunity and protection against furunculosis. Vet Immunol Immunopathol 41:125-139. 29. Vipond, R., Bricknell, I.R., Durant, E., Bowden, T.J., Ellis, A.E., Smith, M., and MacIntyre, S. 1998. Defined deletion mutations demonstrate that the major secreted toxins are not essential for the virulence ofAeromonas salmonicida. Infect Immun 66:1990-1998. 30. Zell, R. 2004. Global climate change and emergence/re-emergence of infectious diseases. Int J Med Microbiol. 293: 16-24. 71 Summary Collectively, the data presented here suggests that an intimate relationship exists between stonefly nymphs and microbial communities inhabiting the sediments of the Nyack floodplain. Of particular interest is the relationship that nymphs have with the fish pathogen Aeromonas salmonicida subsp.salmonicida. Stonefly nymph hindguts, collected from two sites (Petty Creek, Alberton, MT and Nyack floodplain, West Glacier, MT), were found to naturally harbor the pathogen at relatively high rates (55% and 31.5% of individuals) as compared to the sediment (0.02% of sequences). Aeromonas salmonicida is known to harbor multiple plasmids, which are associated with the pathogenesis and symptoms of the disease that it is capable of causing (Stuberet al. 2003). This is relevant, since insect hindguts serve as “hot spots” for gene transfer amongst bacteria (Dillion and Dillion 2004) and genetic information can readily be transferred between strains ofA. salmonicida and possibly other species, (Noonan and Trust 1995) through the use of plasmid exchange. It has been predicted that the incidence of vector-borne disease will increase as temperatures rise due to global warming (Zell 2004). As a model for investigating “pathogen transmission dynamics”, I investigated trophic interactions of Aeromonas salmonicida subsp.salmonicida and stonefly nymphs. In this study, I investigated whether stonefly nymphs can act as a reservoir and vector for A. salmonicida transfer to fish. Feeding and attraction assays were conducted in which stonefly nymphs were presented with a suite of different bacterial isolates. Nymphs preferred to feed on A. salmonicida significantly more than all other bacteria presented. In subsequent experiments, viableA. salmonicida were isolated from both hindgut and frass obtained 72 from nymphs fed A. salmonicida. Finally, rainbow trout juveniles were presented with stonefly nymphs fed a type strain of A. salmonicida, stonefly nymphs naturally harboring A. salmonicida or, commercial fish diet to determine the vectoring capacity of the nymphs. Both stonefly treatments resulted in expression of disease in the trout, and stonefly nymphs naturally harboring the pathogen appeared to be more effective at transmitting disease than stoneflies fed the commercial type strain of A. salmonicida. The results presented here are novel; trophic interactions have not been previously shown to vectorA. salmonicida. The relationships between stonefly nymphs and A. salmonicida are of particular interest as A. salmonicida is often found in low abundance in aquatic environments through culture-based techniques (Austin 1997). Controversy has surrounded the ability of this bacterium to survive in the open environment. Potentially, stonefly nymphs act as a controlling mechanism for the abundance of A. salmonicida, and the disease it can cause. 73 References 1. Austin, B. 1997. Progress in understanding the fish pathogen Aeromonas salmonicida. Mar Biotechnol 15: 131-134. 2. Bemoth, E. M. 1997. Furunculosis: The History of Disease and of Disease Research. Pages 1-20 in Bemoth, E.M., Ellis, A., Midtlyng, P.J., Oliver, G., and Smith, P., editor. Furunculosis: Multidisciplinary Fish Disease and Research. Academic Press, Inc, San Diego. 3. Dillion, R.J., and Dillion, V.M. 2004. The gut bacteria of insects: Nonpathogentic Interactions. Annu Rev Entomol. 49:71-92. 4. Noonan, B., and Tmst, T. 1995. The molecular biology of Aeromonas salmonicida. Annual Review of Fish Diseases 5:95-111. 5. Zell, R. 2004. Global climate change and emergence/re-emergence of infectious diseases. Int J Med Microbiol. 293: 16-24. 74