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Survey of Bacterial and Fungal Associates of Black/Hybrid Imported Fire Ants from Mounds in Mississippi
Richard Baird, Sandra Woolfolk, and C.E. Watson

Southeastern Naturalist, Volume 6, Number 4 (2007): 615–632

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2007 SOUTHEASTERN NATURALIST 6(4):615–632 Survey of Bacterial and Fungal Associates of Black/Hybrid Imported Fire Ants from Mounds in Mississippi Richard Bairdl,*, Sandra Woolfolkl, and C.E. Watson2 Abstract - The assemblage of bacteria and fungi from black (Solenopsis richteri)/ hybrid imported fire ant (BIFA/HIFA) mounds were obtained from four counties in northeast Mississippi. These locations were selected due to high concentrations of BIFA/HIFA that were free from red imported fire ants (RIFA). Mound samples were obtained during October, November, and December in 2003 and January 2004. Patterns of species composition and diversity (species richness) were evaluated from mound soil, mound plant debris, and ant bodies. A total of 5742 isolates consisting of 58 bacterial and 35 fungal taxa were obtained. The most common bacteria identified included Chryseobacterium indolegenes, Stenotrophomonas maltophilia, Actinomadura yumaensis, and Arcanobacterium haemolyticum. Approximately 66% of the fungi cultured belonged to the artificial assemblage Fungi Imperfecti, including Curvularia geniculata, Penicillium spp., Nigrospora sphaerica, and Monoacrosporium leptosporium. The insect pathogen Beauveria bassiana was obtained from mound soil, mound plant debris, and ant bodies, with the greatest percentage from ant bodies. Species richness for bacteria and fungi were both highest from mound soil at 53 and 30 taxa, respectively, and lowest, with 8 bacteria and 25 fungal taxa, from mound plant debris. Species diversity for bacteria was also highest from mound soil, and highest for fungi from ant bodies than the other two isolation conditions. Evenness values for bacteria (0.72–0.80) and fungi (0.74–0.77) during each sampling date had moderate to high relative abundance (1.0 = highest level possible), indicating similarity of taxa among bacteria and among fungi from the four sampling dates. Coefficient of community values comparing sampling dates for bacteria and fungi were greatest between the first and last sampling date (October and January). Temperatures during those dates ranged from 14.4 ºC to 28.9 ºC in October and -2.8 ºC to 10.0 ºC in January. As a continuation of this research, cultures of the different bacteria and fungi obtained in this study are currently being evaluated for their potential as biological control agents of BIFA/HIFA and RIFA that occur in Mississippi. Introduction Imported fire ants (IFA), Solenopsis spp. are difficult insect pests to control and have become major pests throughout the southeastern United States (Vinson and Sorensen 1986). The IFAs are aggressive and effective at foraging, will mobilize rapidly, and sting relentlessly when their mounds are disturbed (Gilbert 1998, Vinson 1994). The black imported fire ant (BIFA), Solenopsis richteri Forel, which came from the Parana and Uruguay Rivers in Argentina and Uruguay, respectively (Vinson and Sorensen 1986), was first introduced 1Department of Entomology and Plant Pathology, Box 9655, Mississippi State University, Mississippi State, MS 39762. 2Oklahoma Agricultural Experiment Station, 139 Ag Hall, Stillwater, OK, 74078. Corresponding author - 616 Southeastern Naturalist Vol. 6, No. 4 into the United States around 1919 near the port at Mobile, AL (Green 1967). In the 1930s, the red imported fire ant (RIFA), Solenopsis invicta Buren was first identified near Mobile (Buren et al. 1974, Lofgren 1986, Lofgren et al. 1975) and rapidly established and spread in all directions. There are currently four species of fire ants in Mississippi, two native and two imported (IFA) species as previously discussed. In addition, a hybrid ant (HIFA), which is a cross between the two IFAs, occurs in the state (Streett et al. 2006). In Mississippi, RIFA are rapidly moving northward along an east to west corridor south of SR 82 (Anonymous 2000, Jarratt and Harris 2001). The RIFA and HIFA are displacing the BIFA in the northern counties and may soon become the only fire ant species in the state (Anonymous 2000, Streett et al. 2006). Therefore, data on soil microbial associates of BIFA mounds must be collected before RIFA and HIFA displace or eliminate them from Mississippi and the southeast. Baseline data of microbial associates such as bacteria and fungi from BIFA mounds may show a unique or distinct microbial community from those that normally occur in RIFA mounds. Species within the microbial population may be beneficial or pathogenic and could be important for potential biological control considerations of IFAs. Currently, several potential biological control agents are under study by scientists throughout the southern United States. Most of these biological control agents are of South America origin (Oi and Williams 2002, Pereira et al. 2002, Williams et al. 2003). Several methods are available for control of RIFAs, but are usually temporary, and the ants quickly reestablish in areas previously treated. These methods are not economical for large areas and can have negative impact to other organisms including beneficials (Williams et al. 2003). The potential of entomopathogenic fungi as biological control agents of IFAs has also been investigated. The two most common species are Beauveria bassiana (Balsamo) Vuillemin and Metarhizium anisopliae (Metschnikoff) Sorokin. In laboratory studies, a mortality rate of 90% had been observed with BIFA exposed to B. bassiana (Broome 1974, Broome et al. 1976). Pereira et al. (1993), and Stimac et al. (1993) observed that a Brazilian strain of the fungus provided approximately 80% control to IFA colonies during field investigations. Beauveria bassiana has also been formulated as baits (Barr and Dress 2003 Barr et al. 2003 Patterson et al. 1993 Williams et al. 2003). The IFAs retrieved bait resulting in an increased level of fungal infection to the IFAs (Williams et al. 2003). In one study, B. bassiana baits caused little mortality to workers, broods, or queens (Barr and Drees 2003). In another, inconsistent control with bait was believed to be caused by several factors including ant defenses, environmental conditions (Fuxa and Richter 2004), and possible strain variability within a different geographical region. Metarhizium anisopliae caused 100% mortality of 15 IFA queens after 5 days under controlled conditions (Sanchez-Peña 1992). No further studies were reported about the potential of the fungus to control IFAs. It is 2007 R. Baird, S. Woolfolk, and C.E. Watson 617 uncertain why no studies have been conducted on occurrence of B. bassiana and M. anisopliae on ants in the midsouth or Mississippi. Surveys to identify naturally occurring populations of these fungi would be important to determine their potential for control of IFAs in this region of the country. There has been very limited research in the United States on the survey of microorganisms associated with IFAs (Beckham et al. 1982, Jouvenaz et al. 1977, Zettler et al. 2002). While previous research has centered on fungi as biological control agents for fire ants, bacteria have not been evaluated. It is uncertain if the diversity and density of bacteria associated with IFA mounds form mutalistic, antagonistic, or parasitic relationships. Select species of bacteria associated with IFAs may be potential biological control agents, but almost no studies have been conducted to determine the potential of these microbes or other organisms (e.g., bacteria) for their control (Williams et al. 2003). More recently, viruses or virus-like organisms have been found to infect S. invicta (Valles et al. 2004, Valles and Strong 2005). Isolation and identification of microorganisms such as bacteria and fungi from IFA bodies, soil, and plant debris in the mounds may identify microbial taxa that can later be evaluated for long-term sustainable population reductions or control of IFAs. Because of the need to develop baseline data of associated bacteria and fungi of BIFA, a survey of mound microbes was conducted. This study compared the diversity and densities of culturable bacteria and fungi associated with IFA, soils, and plant debris within mounds collected from select northern counties in Mississippi. Materials and Methods Eight black/hybrid imported fire ant (BIFA/HIFA) mounds per month were randomly selected and sampled from each of four counties in northeast Mississippi from October to December 2002. Mounds were randomly sampled on 10 October 2002 throughout Oktibbeha County, 7–14 November in Clay County, 2–10 December in Lowndes County, and 4 January in Noxubee County. Each county was subdivided into four quadrants and each of those into two subquandrants. One mound per subquadrant was selected based on randomization from 1–20. For example, if the number 12 was selected, then the 12th mound observed within the subquadrant was sampled. At each location, soil from the lower third of each mound was collected using 3.8-L Ziploc® bags. In this portion of the mounds, IFAs and their broods are at a higher concentration (Vinson 1994, Woolfolk et al. 2006). The samples were returned to the laboratory in an ice cooler and stored up to 24 hrs at 4 °C. Each sample (mound) was processed by weighing 500 g for microbial assessment. In addition to the soil samples, ten BIFA/HIFAs were randomly collected from the samples and preserved in 10-ml glass vials containing 70% ethanol (Triplehorn and Johnson 2004, Woolfolk et al. 2006). The BIFA/HIFAs were then identified to species using keys developed by Trager (1991) and Mississippi Entomological Museum (2003a,b). 618 Southeastern Naturalist Vol. 6, No. 4 Enumeration and isolation of microorganisms. Two nutrient media were employed for bacterial isolation and two for fungi during the study. For bacteria, nutrient glucose agar (NGA) medium, consisting of 3.0 g beef extract, 5.0 g peptone, 2.5 g glucose, and 15.0 g of agar added to 1.0 L of distilled water, was used. The second medium used was King’s medium B agar (KB Difco™, Detroit, MI) containing 20 g proteose peptone #3 (Difco™), 1.5 g of K2HP04, 1.5 g of MgS04⋅7H20, 15.0 g of agar, and 10 ml of glycerol. For fungi, potato dextrose agar (PDA, Difco™) was prepared by adding 4.0 g potato infusion from solids, 20.0 g dextrose, and 15.0 g agar to 1.0 L distilled water, and water agar (WA) was prepared by adding 18 g of agar to 1.0 L of distilled water. The NGA was amended with 50 mg/L of nystatin (Sigma, St. Louis, MO) to inhibit fungi, and the PDA had 300 mg/L streptomycin sulfate (Sigma) and 100 mg/L of chlorotetracyc1ine (Sigma) to inhibit bacteria. Isolation from soil. For each 500-g sample per mound, mound plant debris, BIFA/HIFAs bodies, and soil were removed at random and plated onto the four media described above. For soil isolation, serial dilutions were prepared using 1.0 g of soil from each mound and placed into sterile test tubes containing 9.0 ml of sterile distilled water. The samples were vigorously shaken for 1 min and then 1 ml of the solution was added to another tube containing 9 ml of sterile distilled water (10-1) and shaken for 1 min. An additional dilution was done as before to obtain a 10-2 dilution. The dilutions were processed by placing 100 μL of the solution on each of the four media types and replicated four times by dilution factor. The 100 μL was evenly spread across each plate with a sterile glass rod. The Petri plates were placed at room temperature on a laboratory bench to allow growth for up to 1 week. All fungi growing from the soil were subcultured onto PDA or NGA for bacteria. Isolation from mound plant debris. All mound plant debris was removed from the 500 g of soil by visual removal. From those pieces, 1-cm2 pieces were randomly selected so that 16 pieces per isolation medium or 64 per mound were plated. The 1-cm2 pieces were surface-sterilized using 10% sodium hypoc1orite (w/v 0.534) for 30 sec., and then aseptically placed onto the plates using two per plate. The plates were incubated as in the soil isolation, and isolates were subcultured as before. Isolation from ant bodies. Ant bodies refer to external body regions of the BIFA/HIFAs. Carbon dioxide was used to paralyze the BIFA/HIFAs prior to plating. This was accomplished by placing Nalgene® tubing (0.5-cm diameter) connected to the carbon dioxide tank into a Ziploc® bag containing the BIFA/HIFAs, soil, and debris from the mounds for 30–40 min. A total of 16 ants per mound were plated onto each medium, or 64 ants per mound for four media. Each BIFA/HIFA was submerged into 1% sodium hypoc1orite (w/v 0.0534) containing 0.01% Tween-80® (Sigma, St. Louis, MO.) for 1 min. and rinsed twice with sterile distilled water. Two ants per plate were placed on each of the four media. All fungi or bacteria growing from the tissues 2007 R. Baird, S. Woolfolk, and C.E. Watson 619 on each plate were subcultured onto their respective medium for up to two weeks as per soil dilutions or plant debris methods. Characterization of bacteria and fungi. Since total isolations of fungi and bacteria would be impossible to evaluate for each sampling date, a method was developed for estimating diversity and densities using previously defined protocols (Inglis and Cohen 2004, Woolfolk and Inglis 2003). Up to ten randomly selected fungal colonies and ten bacterial colonies from the appropriate dilution per treatment (plate) were transferred onto PDA slants for fungi and NGA slants for bacteria. In addition, morphologically different or distinct colonies observed on the plates were also isolated for identification. Bacterial isolates were grown for seven days and stored at 24–27 °C or at 4 °C until identifications were determined. A minimum of two representatives of each taxon were stored in glycerol-Brain Heart Infusion broth (BHI; Difco™) mixture (ratio 1:4) at -80 °C for the permanent collection. Primary identification of the bacteria was obtained from their fatty acid methyl ester (FAME) profiles using gas-liquid chromatography (Microbial Identification System Inc. [MIDI], Newark, DE) with previously developed protocol (Sasser 1990). All fungal isolates were grown on PDA for 14 days and stored in -80 °C until further identifications could be conducted. Four representative isolates of each unknown species were placed into a 1.2-ml sterile cryogenic vial (Coming, Acton, MA) containing a glycerol solution (15% glycerol + 85% sterile distilled water) and stored at -80 °C. To prepare the isolates for identification using cultural morphologies, two mycelium plugs were removed from -80 °C storage, placed on PDA plates (Villarroel et al. 2004), and grown for 14 days at room temperature. Fungi were identified using methods developed by Barnett and Hunter (1998), Domsch et al. (1980), Ellis (1971), Roy et al. (2001), and Sutton (1980). Single spore cultures from colonies initially identified as Fusarium spp. were transferred to carnation-leaf agar and further identified using Toussoun and Nelson (1976) methods. Carnation-leaf agar medium was prepared using four discs of prepared sterile, irradiated carnation leaves 7 mm in diameter (Buckout Laboratory, University Park, PA) placed in sterile Petri dishes with 10 ml of 2% water agar added to cover the pieces. Preparations of fatty acid methyl esters (FAMEs) samples. Previously reported methods for preparation and analysis of FAMEs from the bacteria cellular fatty acids were used (Gitaitas and Beaver 1990). Bacteria were grown on tryticase soy-broth agar (TSBA) for aerobes which consisted of 30 g of tryticase soy broth (Difco™), 15 g of agar (Difco™), and 1 L of distilled water. All methods for preparation and analysis of fatty acids were those developed by MIDI, Inc. specific for their library system (Anonymous 2002, 2003). After removal from -80 °C, the isolates were initially grown on Petri plates containing NGA at 30 °C. To prepare the bacterial isolates for FAMEs characterization, each bacterial isolate was streaked onto TSBA and incubated for 48 hrs as previously described (Anonymous 2002). Approximately 0.1 g of bacteria were removed from the TBSA plates and placed 620 Southeastern Naturalist Vol. 6, No. 4 into a screw-top test tube (13 x 100 mm) for fatty acid extractions. FAMEs compositions were determined by gas chromatography procedures using a Hewlett-Packard-6890 (Hewlett Packard, Pittsburgh, PA). The statistical methods used included species richness values (SR), species diversity indices (H') calculated for different data sets using Shannon-Weaver, coefficient of community (CC), evenness (E), and percentage similarity (PS) indices (Stephenson 1989, Stephenson et al. 2004). Stephenson et al. (1989) provides a thorough description of all formulas for the above analyses. Where warranted, data were further analyzed using oneway analysis of variance (ANOVA). Results A total of 5742 isolates consisting of 58 bacterial and 35 fungal taxa were obtained during the study (Tables 1 and 2). The highest isolation frequencies for bacteria included Chryseobacterium indologenes (7.7%), Stenotrophomonas maltophilia (7.3%), Actinomadura yumaensis (2.3%), and Arcanobacterium haemolyticum (1.1%). Bacteria species taxonomy were based on the groupings defined by Holt et al. (1994) and Euzeby (2006). Approximately 31.0% of the fatty acid profiles of the other bacterial isolates could not be identified, and all other species of bacteria were <1%. Approximately 66% of all the fungi cultured belonged to the artificial assemblage Fungi Imperfecti. The common species identified were Curvularia geniculata (12.7%), Penicillium spp. (11.5%), Nigrospora sphaerica (7.5%), and Monoacrosporium leptosporium (7%). Also, the insect pathogen, Beauveria bassiana (6.7%) was isolated from ant bodies, mound soil, and mound plant debris, with the highest percentage from ant bodies. Remaining filamentous organisms included Ascomycota and Zygomycota (≈1%), and sterile or unknown isolates were 37.4%. Species richness values for bacteria and fungi isolated on selective media had a similar trend with species diversity following the same pattern (Table 3). Values for total bacterial taxa was highest from mound soil (n = 53), and bacterial taxa were isolated at a greater rate from KB (n = 42) than NGA (n = 36) media. The mound plant debris, however, had the lowest values on KB (n = 6) and NGA (n = 5) and were also lowest for total taxa (n = 8). Species diversity values for bacteria were higher from mound soil (H' = 0.89) than ant bodies (H' = 0.71) and mound plant debris (H' = 0.53). Species richness values for fungi were highest from total mound soil (n = 30), and fungi were isolated more commonly on PDA (n = 29), but species diversity was slightly lower for mound soil (H' = 1.0) than ant bodies (H' = 1.07) (Table 3). Total fungi values were lowest from ant bodies (n = 22), but fungal species diversity from mound plant debris (H' = 0.94) had the lowest value. Species richness and diversity values were compared by sampling date and isolation condition for bacteria and fungi. Mound soil values for bacteria and fungi were highest in January for all sampling dates, except 2007 R. Baird, S. Woolfolk, and C.E. Watson 621 for bacteria in October and fungi in January (Fig. 1). Mound plant debris diversity values were consistently lower for fungi than for the other two isolation conditions except for December, and were lower for bacteria in Table 1. Percent isolation frequencies of fungal taxa identified from black/hybrid imported fire ant mounds collected in northeast Mississippi. Total % by condition Over- Mound all Ant Mound plant total TaxaA bodies soil debris % Fungi Imperfecti Acremonium strictum W. Gams 0.2 0.4 0.0 <1.0 Aspergillus terreus Tham. 0.0 0.0 1.0 <1.0 Beauveria bassiana (Bals.) Vuill. 8.0 6.1 4.7 6.7 Dipdans nodulosa (Berk. & M.A. Curtis) Shoemaker 0.0 0.0 1.2 <1.0 Candida guillermondii (Castellani) Langeron & Guerra 0.2 1.2 0.0 <1.0 Cochliobolus sativus (Ito & Kuribayashi) Drechs. Ex Dastur 1.4 7.4 5.1 4.3 Curvularia geniculata (Tracey & Earle) Boedijn 4.7 25.0 8.2 12.7 Cytosporella sp. 0.0 0.4 2.0 <1.0 Fusarium merimoides Corda 0.0 0.2 0.0 <1.0 Fusarium oxysporum Fr. 0.0 0.2 0.0 <1.0 Fusarium solani (Marti) Sacc. 1.3 0.4 2.0 1.0 Fusarium spp. (10 spp.) 0.3 2.9 7.0 2.5 Gloeosporium spp. (2 spp.) 0.0 1.2 0.0 <1.0 Monoacrosporium leptosporium (Drechs.) A. Rubner 2.2 12.5 8.2 7.0 Macrophoma spp. (2 spp.) 1.0 0.0 0.0 <1.0 Nigrospora sphaerica (Sacc.) F. Mason 2.2 12.5 8.2 7.5 Nodulosporium spp. (3 spp.) 0.5 3.9 1.2 1.8 Paecilomyces lilacinus (Thom) R. A. Samson 0.2 1.2 0.0 <1.0 Papulospora byssina Hotson 0.0 0.0 1.2 <1.0 Penicillium spp. (4 spp.) 2.7 20.5 14.8 11.5 Phaeoseptaria airae (Grove) R. Sprague 0.0 2.2 0.0 <1.0 Phoma herbarum Westend. 0.3 1.0 5.1 1.4 Rhinocladiella atrovirens Nannf. in Melin & Nannf. 0.0 1.0 0.0 <1.0 Rhizoctonia solani Kühn (AG-3) 0.3 0.2 0.0 <1.0 Speggazina sp. 0.0 0.0 0.5 <1.0 Trichoderma spp. (2 spp.) 0.3 0.0 0.4 <1.0 Tubercularia vulgaris Tode: Fr. 0.6 4.5 2.3 2.3 Verticillium dahliae Kleb. 0.8 0.0 0.0 <1.0 Ascomycota Eupenicillium cinnamopurpureum D.E. Scott & A. C. Stolk 0.0 0.2 0.0 <1.0 Unknown 0.5 0.4 0.4 <1.0 Zygomycota Mucor hiemalis Wehmer 0.0 0.2 0.4 <1.0 Zygorhynchus moelleri Vuill. 0.0 1.7 0.0 <1.0 Straminipila Pythium spp. (3 spp.) 1.7 19.7 14.1 10.5 Unknowns 8.0 53.5 78.5 37.4 AMean percent of isolations from ant bodies ÷ 640 (= 20 ant bodies per mound x 8 mounds x 4 sampling dates) x 100; mean percent from mound soil ÷ 512 (= 16 soil dilution plates per mound x 8 mounds x 4 sampling dates) x 100; mean percent of isolations from mound plant debris ÷ 256 (= 8 plant tissue pieces/ mound x 8 mounds x 4 sampling dates) x 100; and overall mean total percentages of total isolates ÷ 1408 (= 640 + 512 + 256) x 100. 622 Southeastern Naturalist Vol. 6, No. 4 Table 2. Percent isolation frequencies of bacterial taxa identified from black/hybrid imported fire ant mounds collected in Northeast Mississippi. Total % by conditionB Over- Mound all Ant Mound plant total TaxaA bodies soil debris % Acidovorax avenae subsp. cattleyae (4) (Pavarino) Willems, 0.0 0.2 0.0 <1.0 Goor, Thielemans, Gillis, Kersters and De Ley Actinomadura yumaensis (28) Labeda, Testa, Lechevalier, 0.8 6.3 0.4 2.7 and Lechevalier Agrobacterium radiobacter (4) (Beijerinck and van Delden) 0.0 0.2 0.0 <1.0 Conn Alcaligenes faecalis (4) Castellani and Chalmers 0.0 0.2 0.0 <1.0 Arcanobacterium haemolyticum (20.0) (McLean, Liebow & 0.2 2.9 0.0 1.1 Rosenberg) Collins, M.D. Jones, D. Schofield Bacillus sp. (18) 0.0 0.0 0.0 <1.0 Bacillus spaericus (18) Meyer and Neide 0.0 0.4 0.0 <1.0 Burkholderia cepacia (5)** (Palleroni and Holmes) Yabuuchi, 0.2 0.2 0.0 <1.0 Kosako, Oyaizu, Yano, Hotta, Hashimoto, Ezaki & Arakawa Burkholderia gladioli (5)** (Severini) Yabuuchi, Kosako, 0.0 0.6 0.0 <1.0 Oyaizu, Yano, Hotta, Hashimoto, Ezaki & Arakawa Burkholderia pyrrocinia (5)** (Imanaka, Kousaka, Tamura, 0.0 0.2 0.0 <1.0 & Arima) Vandamme, Holmes, Vancanneyt, Coenye, Hoste, Coopman, Revels, Lauwers, Gillis, Kersters, & Govan Cedecea davisae (5)** Grimont, Grimont, Farmer, & Asbury 0.0 0.2 0.0 <1.0 Chromobacterium violaceum (4)** Bergonzini 0.0 0.2 0.0 <1.0 Chryseobacterium indologenes (4)** (Yabuuchi, Kaneko, Yano, 1.3 19.5 0.0 <1.0 Moss, & Miyoshi) Vandamme, Bernardet, Segers, Kersters, & Holmes Chryseobacterium indolthetic (4)** (Campbell and Williams) 0.2 0.0 0.0 <1.0 Vandamme, Bernardet, Segers, Kersters, & Holmes Chryseobacterium meningosepticum (4)** (Li) Vandamme, 0.0 0.6 0.0 <1.0 Bernardet, Segers, Kersters, & Holmes Comamonas acidovorans (4) (den Dooren de Jong) Tamaoka, 0.0 0.2 0.0 <1.0 Ha, & Komagata Corynebacterium aquaticum (20.0) (ex Leifson 1962) Evtushenko, 0.0 0.2 0.0 <1.0 Dorofeeva, Subbotin, Cole, and Tiedje Flavobacterium johnsoniae (18) (Stainer) Bernardet, Segers, 0.3 1.4 0.0 <1.0 Vancanneyt, Berthe, Kersters, & Vandamme Flavobacterium resinivorum (18) Delaporte and Daste 0.0 0.6 0.0 <1.0 Klebsiella pneumoniae subsp. pneumoniae (5) (Schroeter) Trevisan 0.0 0.2 0.0 <1.0 Kluyvera ascorbata (5) Farmer, J. J., Fanning, Huntley-Carter, 0.0 0.2 0.0 <1.0 Holmes, Hickman, Richard, & Brenner Kocuria varians (20.0)** (Migula) Stackebrandt, Koch, Gvozdiak, 0.0 0.2 0.0 <1.0 & Schumann Lactobacillus delbrueckii subsp. lactis (19) (Orla-Jensen) Weiss, 0.0 0.4 0.0 <1.0 Schillinger, & Kandler Methylobacterium organophilum (4) Patt, Cole, & Hanson 0.0 0.2 0.0 <1.0 Micrococcus lylae (17) Kloos, Tornabene, & Schleifer 0.0 0.2 0.0 <1.0 emend. Wieser, Denner, Kämpfer, Schumann, Tindall,Steiner, Vybiral, Lubitz, Maszenan, Patel,Seviour, Radax, & Busse Ochrobactrum anthropi (4) Holmes, Popoff, Kiredjian, & Kersters 0.0 0.2 0.0 <1.0 Paenibacillus peoriae (18)** (Montefusco, Nakamura, and Labeda) 0.0 0.2 0.0 <1.0 Heyndrickx, Vandermeulebroecke, Kersters, DeVos, Logan, Aziz, Ali, & Berkeley 2007 R. Baird, S. Woolfolk, and C.E. Watson 623 Table 2, continued. Total % by conditionB Over- Mound all Ant Mound plant total TaxaA bodies soil debris % Paenibacillus polymyxa (18)** (Prazmowski) Ash, Priest, & Collins 0.0 0.8 0.0 <1.0 Paenibacillus apiarus (18)** (ex Katznelson) Nakamura 0.0 0.2 0.0 <1.0 Pseudomonas aeruginosa (4) (Schroeter) Migula 0.0 2 0.4 <1.0 Pseudomonas chlororaphis (4) (Guignard and Sauvageau) Bergey 0.2 2.2 0.4 <1.0 Pseudomonas fluorescens (4) Migula 0.0 0.2 0.0 <1.0 Pseudomonas putida (4) (Trevisan) Migula 0.3 3.9 0.4 1.6 Salmonella typhimurium (5) (Loeffler) Castellani and Chalmers 0.0 0.4 0.0 <1.0 Sphingobacterium multivorum (4) (Holmes, Owen, & Weaver) 0.2 1.8 0.0 <1.0 Yabuuchi, Kaneko, Yano, Moss, & Miyoshi Sphingobacterium spiritivorum (4) (Holmes, Owen, & Hollis) 0.0 1.2 0.0 <1.0 Yabuuchi, Kaneko, Yano,Moss, & Miyoshi Staphylococcus aureus (17) Rosenbach 0.2 0.0 0.0 <1.0 Staphylococcus hominis (17) Kloos and Schleifer 0.2 0.2 0.0 <1.0 Stenotrophomonas maltophilia (4)** (Hugh) Palleroni and Bradbury 2.8 15.6 2 7.3 Tsukamurella paurometabola (22) (Steinhaus) Collins, Smida, 0.2 0.0 0.0 <1.0 Dorsch, & Stackebrandt Unknown 8.4 70.9 8.2 31.1 Variovorax paradoxus f. A (4) (Davis) Davis, Doudoroff, Stanier, 0.3 0.2 0.0 <1.0 & Mandel Variovorax paradoxus f. B (4) (Davis) Davis, Doudoroff, Stanier, 0.0 1.2 0.0 <1.0 & Mandel Vibrio furnisii (5) Brenner, Hickman-Brenner, Lee, Steigerwalt, 0.0 0.2 0.0 <1.0 Fanning, Hollis, Farmer, Weaver, Joseph, & Seidler Xanthobacter agilis (4) Jenni & Aragno 0.0 0.6 0.0 <1.0 Xanthomonas campestris subsp. campestris (4) (Pammel) Dowson 0.0 0.4 0.0 <1.0 Xanthomonas translucens subsp. translucens (4) (ex Jones, 0.0 0.2 0.0 <1.0 Johnson, and Reddy) Vauterin, Hoste, Kersters, and Swings Xenorhabdus luminescens (5)** Thomas and Poinar 0.0 1 0.0 <1.0 Xenorhabdus nematophilus (5)** (Poinar and Thomas) Thomas 0.0 1.8 0.4 <1.0 and Poinar Yersinia pseudotuberculosis (5) (Pfeiffer) Smith and Thal. 0.0 0.6 0.0 <1.0 Bacillus cereus (18) Frankland & Frankland 0.0 0.2 0.0 <1.0 Chryseobacterium balustinum (4)** (Harrison) Vandamme 0.0 0.2 0.0 <1.0 Gordona bronchialis (22) (Tsukamura) Stackebrandt, Smida, 0 0.2 0 <1.0 & Collins Neisseria flavescens (4) Branham 0.0 0.2 0.0 <1.0 Nocardia nova (22) Tsukamura 0.0 0.2 0.0 <1.0 Paenibacillus pabuli (18)** (Nakamura) Ash, Priest, & Collins 0.0 0.0 0.4 <1.0 Pseudomonas flectens (4) Johnson 0.0 0.6 0.0 <1.0 Pseudomonas huttiensis (4) Leifson 0.0 0.2 0.0 <1.0 Rhodococcus luteus (22) Nesterenko, Nogina, Kasumova, 0.2 0.0 0.0 <1.0 Kvasnikov, & Batrakov ANumbers in parentheses represent groups of bacteria as listed by Holt et al. (1994) and Euzeby (2006). The latter is indicated by “**.” BMean percent of isolations from ant bodies ÷ 640 (= 20 ant bodies per mound x 8 mounds x 4 sampling dates) x 100; mean percent from mound soil ÷ 512 (= 16 soil dilution plates per mound x 8 mounds x 4 sampling dates) x 100; mean percent of isolations from mound plant debris ÷ 256 (= 8 plant tissue pieces/ mound x 8 mounds x 4 sampling dates) x 100; and overall mean total percentages of total isolates ÷ 1408 (= 640 + 512 + 256) x 100. 624 Southeastern Naturalist Vol. 6, No. 4 October and November. No consistent trends in species diversity values were noted for any of the isolation conditions or by sampling date (Fig. 2). Species richness, diversity, and evenness values were also calculated across sampling dates for total bacteria and fungi (Table 4). Richness for bacteria was highest in December (n = 29) and lowest in October (n = 11). However, the diversity was greatest in November (H' = 0.96) and lowest in October (H' = 0.64) for bacteria. Fungi had the highest richness values in October (n = 26) and the lowest in November (n = 18) and December (n = 18). Diversity for fungi was greatest in October (H' = 1.08) and lowest in December (H' = 0.94) and January (H' = 0.97). Evenness values for bacteria and fungi during each sampling date had a moderate to high relative abundance within the two domains at J = ≈0.75 indicating similarity of taxa for bacteria and between fungi. Figure 1. Species richness for bacteria and fungi identified by sampling date and isolation conditions. Table 3. Species richness and diversity (H’) of bacteria and fungi isolated from black/hybrid imported fire ant mounds, using two selective media. Species richness MediaA Isolation Bacteria Fungi Total # of taxa Species diversity Condition KB NGA PDA WA Bacteria Fungi Bacteria Fungi Ant bodies 16 12 19 18 16 22 0.71 1.07 Mound soil 42 36 29 18 53 30 0.89 1.00 Mound plant debris 6 5 22 14 8 25 0.53 0.94 AKB = King’s B medium, NGA = nutrient glucose agar, PDA = potato dextrose agar, WA = water agar. 2007 R. Baird, S. Woolfolk, and C.E. Watson 625 The coefficient of community (CC) values for pooled bacteria and fungal species composition compared between mound plant debris and soil-isolate data was highest at 0.89. The CC values were highest for ant bodies and soil data at 0.81, and lowest for ant bodies and mound plant debris data at 0.77. Isolation data from mound soil, ant bodies, and mound plant debris were compared using coefficient of community values for bacteria and fungi (data not shown). The average value for all possible combinations of bacteria using coefficient of community was 0.30, and ant bodies-mound plant debris were higher than mound soil-mound plant debris at 0.23. Overall, similarities of bacterial species were lower than the fungi from the three isolation conditions. The average coefficient of community value for all fungi was 0.82. Mound soil and mound plant debris comparison had the highest similarity value at 0.89, and the similarity value was lowest for ant bodies-mound plant debris at 0.77. Figure 2. Species diversity (H') of bacteria and fungi identified by sampling date and isolation conditions. Table 4. Species richness (n), diversity (H'), and evenness (J) for bacteria and fungi identified from black/hybrid imported fire ant mounds over four sampling dates in northeast Mississippi. Richness (n) Diversity (H') Evenness (J) Sampling date Bacteria Fungi Bacteria Fungi Bacteria Fungi October (2002) 11 26 0.64 1.08 0.72 0.77 November (2002) 25 18 0.96 1.01 0.80 0.76 December (2002) 29 18 0.83 0.94 0.72 0.76 January (2003) 26 20 0.67 0.97 0.78 0.74 626 Southeastern Naturalist Vol. 6, No. 4 Coefficients of community values were obtained comparing data between the four sampling dates for bacteria and fungi (Table 5). For bacteria and fungi, the October–November and December–January comparisons had the lowest CC value reflecting the differences in common taxa between those sampling dates. With the exception of October and December (0.45) for bacteria, and November and January (0.79) for fungi, the coefficient of community values for both domains were greatest for adjacent samplingdate comparisons of October and November, November and December, and December and January. The CC values for October and January comparison was lower than for any other sampling-date comparisons (0.57) and had the longest period of time between samples. Discussion This investigation included the first major survey of bacterial species associated with fire ant mounds. A higher population of bacterial species was identified over the four sampling dates than fungi (Tables 1 and 2). In a previous study, mounds of RIFA contained almost 25% more fungal taxa, and the species composition was different (Zettler et al. 2002). In that investigation, the most frequently isolated fungus was Papulospora byssina Hotson. Other common fungi included 11 species each of Penicillium and Fusarium, 5 of Trichoderma, and 10 species of Zygomycotina. The differences in mycobiota composition between the current and previous study may be due to geographical or ant species variability. The four most common bacteria isolated (C. indologens, S. maltophilia, A. yumaensis, and A. haemolyticum) were isolated more commonly from mound soils and ant bodies than from mound plant debris. Previous studies identifying bacteria from ants were from internal tissues (midguts) rather than exterior sources (Bouwma et al. 2006, Peloquin and Greenberg 2003, Van Borm et al. 2001), but none of these studies observed the four common species identified in the current investigation. Other taxa identified common to the current and previous studies were species of Burkholderia, Flavobacterium, Kluyvera, Methylobacterium, Pseudomonas, and Staphylococcus (Li et al. 2005; Peloquin and Greenberg 2003; Van Borm et al. 2002a, 2002b). However, similarity was only at the generic level. Wolbachia sp. infections are common in ant midgut regions (Van Borm et al. 2001), but no isolations were obtained since this bacterium cannot be cultured. The four most common taxa of fungi identified in this study are common saprophytes (Barnett and Hunter 1998, Ellis 1971). All occur in Table 5. Coefficient of community values for microbiota from black/hybrid imported fire ant mounds collected in northeast Mississippi, 2002–2003. Oct.–Nov. Oct.–Dec. Oct.–Jan. Nov.–Dec. Nov.–Jan. Dec.–Jan Bacteria 0.39 0.45 0.32 0.41 0.41 0.40 Fungi 0.68 0.64 0.57 0.83 0.79 0.84 2007 R. Baird, S. Woolfolk, and C.E. Watson 627 soil, although some infrequently, and survive on plant debris or are plant pathogens. These fungi occur in many different habitats and their presence in fire ant mounds may be secondary. The insect pathogen B. bassiana was isolated more commonly from ant bodies (8%) than mound soil (6.1%) or debris (4.7%). However, no insect pathogens were observed from ant macerates in 1007 colonies of RIFA and 83 of BIFA in the southeastern United States (Jouvenaz et al. 1977) or from S. invicta mounds surveyed at Clemson, SC (Zettler et al. 2002). Briano et al. (1995) conducted a survey of S. richteri and S. invicta pathogens in Argentina, but B. bassiana was not isolated in that study. The current study confirmed that fire ant mounds could serve as a potential reservoir for the fungus. Paecilomyces lilacinus, which was isolated from ant bodies and mound soil in the study, was found to significantly lower colony populations under controlled conditions (R. Baird, unpubl. data). However, the isolation frequencies from mounds were less than 1% for this fungus. Some Paecilomyces spp. are known to be entomopathogenic, and a few were reported to be anamorphs of Cordyceps spp. (Humber 1997). Total species richness values for bacteria and fungi were always highest from mound soils compared to ant bodies and mound plant debris. When those data were compared by sampling date, species richness values were also highest from mound soils with few exceptions. Soils are known to contain diverse microbial communities (Barron 1972) compared to ant bodies (midguts) (Li et al. 2005, Peloquin and Greenberg 2003, Van Borm et al. 2002a), but insect gut regions, including midguts, have limited bacterial and fungal communities that are adapted to and form mutalistic relationships with the insect. The soil provides a widely diverse nutrient base enabling microorganism populations to have greater diversity, but the majority of the microbial community is saprophytic and probably has little or no direct involvement with the ants in their mounds (Barnett and Hunter 1998). However, select bacteria and fungi form mutualistic or parasitic associations with ants and other insects such as termites (Aanen et al. 2002, Degnan et al. 2004, Hyodo et al. 2000). These microorganisms can occur in nests and hosts. They serve as a direct food source, as antagonists to invading pests, or as decomposers of degrade plant material later utilized by the insects. None of these microbes were identified in this study. Another possibility why species richness from ant bodies was lower than from soils is that ants are reported to have antimicrobial defenses that significantly lower populations and species of fungi that are tolerant to living in ant bodies (Zettler et al. 2002). Other studies observed that secretions from metapleural glands can significantly decrease fungal growth of several species including B. bassiana and P. lilacinus (Beattie et al. 1985). Species diversity showed a similar trend as species richness for both bacteria and fungi. Bacteria growing in mound soil had the highest species diversity levels in comparison to ant bodies or mound plant debris. Fungi mound soil and ant bodies had similar diversity levels even though species 628 Southeastern Naturalist Vol. 6, No. 4 richness was highest for fungi from the mound soils. Abundance or total isolations of fungi from ant bodies were similar to mound soil even though species richness was greater in the mound soil. Species richness and diversity values showed no apparent trends by sampling date. One exception was that in October species richness and diversity was lowest for bacteria but highest for fungi. Possibly as a result of the reduced antagonism by bacteria during the first sampling date, fungal populations were able to increase due to the reduced competition. No other trends were noted based on isolation data. The coefficient of community values comparing sampling dates for bacteria and fungi were greatest between the first and last sampling date (October and January). However, the values compared between corresponding sampling dates were almost always similar with few exceptions previously noted. These latter results indicate a trend towards a gradual change in microbial community composition over time. Environmental factors such as temperature and rainfall can influence microorganism diversity and densities. In this study, temperature variability during the four sampling dates may have been a contributing factor on coefficient-of-community values between October and January. Even though temperature differences did not vary greatly between consecutive months, average temperatures were warmer in October (14.4 ºC to 28.9 ºC) than January (-2.8 ºC to 10.0 ºC). The temperature differences between those two months may have directly affected diversities and densities (coefficient-of-community values) even though the gradual temperatures changes between corresponding months may not have had significant effects. Also, average monthly precipitation data were almost identical throughout the study period. In conclusion, a diverse microbial community of bacteria and fungi occur within BIFA/HIFA mounds as shown in this current study. The fungi varied from those found previously, and the purpose of their occurrences in the mounds is unknown. The bacteria and fungi isolated during the investigation will be used to identify the potential interactions of the microbes with the ants and possible biological controls for BIFA/HIFA and RIFA. Acknowledgments I would like to acknowledge USDA-ARS Specific Cooperative Agreement for providing support for this research both years of the study under Project Numbers 6402-22320-00300D. 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