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.
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
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).
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
Ant Mound plant total
TaxaA bodies soil debris %
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
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
Mucor hiemalis Wehmer 0.0 0.2 0.4 <1.0
Zygorhynchus moelleri Vuill. 0.0 1.7 0.0 <1.0
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
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
Agrobacterium radiobacter (4) (Beijerinck and van Delden) 0.0 0.2 0.0 <1.0
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,
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
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
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
Variovorax paradoxus f. B (4) (Davis) Davis, Doudoroff, Stanier, 0.0 1.2 0.0 <1.0
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
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
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
Table 3. Species richness and diversity (H’) of bacteria and fungi isolated from black/hybrid
imported fire ant mounds, using two selective media.
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 =
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
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.
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.
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
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.
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. Also, thanks go to Nicole Lee, Daisy Goodman, Chelsey Wilson,
and Becky Baker for laboratory support during the investigation. Approved for
publication as Journal Series No. J-11045 of the Mississippi Agricultural and Forestry
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