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Bactericidal and Fungicidal Activity of Ant Chemicals on Feather Parasites: an Evaluation of Anting Behavior As a Method of Self- Medication in Songbirds

Posted on: Saturday, 13 November 2004, 03:00 CST

ABSTRACT.-

Songbirds apply ants to their feathers during anting behavior, possibly as a method of reducing feather parasites. We tested polar and nonpolar ant secretions and pure formic acid for bactericidal and fungicidal effects on microbial ectoparasites of feathers. Microbial inhibition trials were run with the bacteria Bacillus liceniformis (strains OWU 138B and OWU 1432B) and B. subtilis; and with the fungi Chaetomium globosum, Penicillium chrysogenum, and Trichoderma viride. Ant chemicals were derived from Camponotus pennsylvanicus, Pheidole dentata, Aphaenogaster rudis, Crematogaster lineolata, and Lasius flavus worker-caste ants. Although pure formic acid strongly inhibited all bacteria and fungal hyphae tested, concentrations of formic acid found in the bodies of formicine ants did not. Neither hexane ant-chemical extracts nor ant suspensions in deionized water inhibited the microbial species. Consequently, the hypothesis that birds apply ants to control feather parasites was not supported. Received 2 May 2003, accepted 24 June 2004.

RESUMEN. -Las aves paserinas aplican hormigas a sus plumas en el comportamiento de hormigueo, posiblemente como un mtodo para reducir la carga parasitaria de las plumas. En este estudio probamos la capacidad bactericida y fungicida de secreciones polares y no polares de hormigas y formas puras de cido frmico sobre ectoparsites microbianos de plumas. Las pruebas de inhibicin bacteriana fueron realizadas con Bacillus licheniformis (cepas OWU 138B y OWU 1432B) y B. subtilis, y con los hongos Chaetomium globosum, Penicillium chrysogenum y Trichoderma viride. Los qumicos de hormigas fueron derivados de las castas obreras de las especies Camponotus pennsylvanicus, Pheidole dentata, Aphaenogaster rudis, Crematogaster lineolata y Lasius flavus. A pesar de que las formas puras de cido frmico inhibieron fuertemente a todas las bacterias e hifas micticas probadas, las concentraciones de cido frmico encontradas en los cuerpos de las hormigas de la subfamilia Formicinae no lo hicieron. Los extractos qumicos de hexanos de hormigas y hormigas suspendidas en agua deionizadas tampoco inhibieron a las especies de microbios. Consecuentemente, la hiptesis de que las aves se aplican hormigas para controlar los parsitos de las plumas no fue apoyada.

ONE HYPOTHESIS PROPOSED to explain "anting behavior" in songbirds (i.e. individuals applying ants to their own feathers) is that they are extracting ant chemical secretions to control feather parasites (Ehrlich et al. 1986). Although anting behavior has been shown to have no effect on feather mites or lice (Kelso and Nice 1963, Potter 1989, Judson and Bennett 1992, clayton 1999), it may reduce the abundance of feather-degrading microbes (Ehrlich et al. 1986, Clayton 1999). Beer (1963), who reviewed the frequency and kinds of microorganisms on feathers, found large numbers of both bacterial and fungal species. Fungi and bacteria can harm feathers by breaking down their structural integrity through keratinolytic activities (Pugh 1965, Burtt and Ichida 1999, Muza et al. 2000).

Most songbird anting displays identified in the literature involve use of worker ants of the subfamily Formicinae (Formicidae: Hymenoptera) (McAtee 1938; Staebler 1942; Ivor 1943; Nice 1945; Brackbill 1948; Groskin 1950; Potter 1970, 1985; Whyte 1981). Apparently, songbirds favor formicine ants more than ants of other subfamilies (Simmons 1957, Whitaker 1957). Formicine ants secrete a variety of chemicals, including formic acid, a corrosive and cytotoxic acid capable of causing dermal necrosis in large doses (BIum 1992, Judson and Bennett 1992). Formicine ants have a vestigial sting, and they use a dilute solution of formic acid as a primary defensive mechanism. They produce it as a spray that usually consists of a 60% aqueous solution containing formic acid, free amino acids, and small peptides (Blum 1992). At certain concentrations, formic acid has bactericidal and fungicidal properties.

Ants produce numerous chemicals in addition to formic acid. For example, growth of both Escherichia coli bacteria and Aspergillus parasiticus fungi is repressed by the 3-methylindole produced by army ants (Neivamynnex nigrescens; Brown et al. 1979). Although pheromone secretions are usually species-specific, unrelated species -including parasitoids-have evolved the ability to identify ants' locations through their chemical secretions (Brown and Feener 1991). Watkins et al. (1969) found that 3-methylindole produced by N. nigrescens repels insectivorous snakes. Apocephalus paraponerae, a phorid fly parasitoid, is attracted to whole-body extracts of Paraponera clavata, a large tropical ant (Feener et al. 1996). Additionally, some colony parasites use ant chemicals to identify appropriate prey. The myrmecophilic beetle Atemeles piibicolis identifies suitable ant colonies through their host odor (Hlldobler 1969). Such semio-chemical communication between ant and nonant species may be involved when birds rub ants on their feathers.

We tested the hypothesis that when birds engage in anting behavior, they are reducing microbial loads on feathers through fungicidal or bactericidal actions of ant-derived chemicals (Ehrlich et al. 1986, Burke et al. 1993, Clayton and Wolfe 1993, Clayton 1999, Furlow 2000). We tested for bactericidal and fungicidal properties of polar (i.e. molecules containing covalent bonds in which asymmetric distribution of electron density creates potential for hydrogen bonding with other polar molecules) and non-polar (i.e. molecules containing symmetric distributions of electron densities) ant secretions, and of pure formic acid, on microbial ectoparasites of bird feathers.

METHODS

The effect of ant chemicals on feather microbes was examined using agar-plate inhibition assays with bacterial and fungal species identified by Pugh (1965), Burtt and Ichida (1999), and Muza et al. (2000) as likely to occur on feathers. Those microbes included the bacteria Bacillus licheniformis (strains OWU138B [ATCC] and OWU 1432B) and B. subtilis; and the fungi Chactomium globosum, Penicillium chrysogenum, and Trichoderma viride. Ant chemicals were derived from worker-caste ants collected in southeastern Virginia, including the Formicinae ants Camponotus pennsilvanicus and Lasius flavus and the Myrmicinae ants Pheidole dentala, Aphaenogaster rudis, and Crematogaster lineolata. We ordered the fungi and B. subtilis from Fisher Scientific (Pittsburgh, Pennsylvania). Bacillus licheniformis strains were provided by E. H. Burtt, Jr. We first subcultured each bacterial and fungal species to produce individual colonies and to confirm identity through visual examination.

Bacillus colonies and mature fungal colonies and spores were suspended in sterile 0.85% NaCl solution. The Bacillus suspension was prepared by using one full loup of a mature colony (>24 h incubation) mixed with 1 mL of saline in a test tube on a Vortex mixer (Labnet, Woodbridge, New Jersey). The suspension of mature fungal hyphae (>72 h incubation) was produced from a 1-mL piece of agar cut from a mature fungal plate and combined with 1 mL saline solution in a tissue homogenizer. The fungal spore suspension was prepared from three plates of mature fungal colonies for each species. Spores were collected on round filter paper (6.5 cm diameter) and then mixed with 1 mL of a 0.85% NaCl solution in a glass test tube using a Vortex mixer. Presence of spores was confirmed through visual inspection of the resulting agar plates under a 200x Nikon dissection microscope.

Agar plates used in inhibition tests contained a 10-L, suspension of each Rncilliis strain, a 100-L mature fungal suspension, or a 400- L fungal spore suspension. All microbial work was conducted under a laminar flow hood (class II, type A) with autoclaved equipment. Agar plates were inoculated with bacteria using a spread-plate methodology to create lawns of bacterial growth. Bactericidal and fungicidal activity of ant secretions and of pure formic acid were tested on filter-paper disks (6 mm diameter) placed into each inoculated agar spread-plate. Each agar plate contained four equidistant filter-paper disks; two disks contained 10 L each of the experimental treatment solution, and two contained 10 L each of the control solution (Fig. 1). Preliminary trials identified no flooding or leakage from 6-mm filter-paper disks when absorbing 10 L of deionized water. Inhibition assays testing fungal spore suspensions were conducted by mixing the 400-micro;L individual spore suspensions with 20 L of one treatment type on an agar plate. Fungal- spore inhibition assays were conducted on each microbial species with both polar and nonpolar extracts from each ant species.

FIG. 1. Experimental agar plate design. (C) Control filter-paper disk. (T) Treatment filter-paper disk.

Ant-chemical treatments were performed by two methods. The first treatment (hereafter "hexane extract") was performed to isolate nonpolar ant-secretions. Fifty ants of each species were frozen and then placed in a beaker with 5 mL of hexane for 30 min. For P dentata, which has both soldier and worker-ant castes, 16 soldiers and 34 workers were soaked in th\e hexane. Next, 5 mL of hexane was added to the beaker, and the ants soaked for an additional 30 min. Ant carcasses were removed from the solution after soaking for 1 h. Thus, 10 mL of ant extract (per 50 ants) was prodviced for each ant species. Next, each 10 L of ant extract was pipetted onto a filter- paper disk (6 mm diameter). Controls for the ant extract consisted of 10 L pure hexane. Paper disks with an ant-extract treatment and a hexane control were allowed to dry before placement onto agar plates containing the microbial cultures. The second ant-chemical treatment was produced as an ant suspension in deionized water (hereafter "ant suspension") to test polar ant-secretions. Three frozen ants of each species were ground in separate tissue-homogenizers with 700 L deionized water. For P. dentata, one soldier and two worker ants were selected. Ten milliliters of each ant suspension and 10-pL deionized-water controls were pipetted onto 6-mm paper disks; two treatment disks and two control disks were placed on each inoculated agar plate. A third treatment consisted of 10 L of formic acid (Fisher Scientific) added to two 6-mm disks and compared with two untreated filter-paper disks.

Three ants to 700 L deionized water (ant suspension) and 50 ants to 10 L hexane (hexane extract) both resulted in approximately the same proportion of solute to solvent: 4.3 x 10^sup -3^ and 5 x 10^sup -3^, respectively. We used those concentrations to produce the amounts of solutions needed to cover one feather with an approximate area of 1.5 x 4 cm.

Each of the three treatments was tested on all bacteria and on hyphae and spores of each fungal species. Five replicate plates were prepared for each treatment and each ant-microbe species combination, except for those with the fungal spores, for which three replicate plates were prepared for each ant-fungus combination.

Bacterial growth was examined 1, 4, 12, and 24 h after inoculation. Fungal growth cultured from hyphal strands was examined 4, 5, and 6 days after incubation. For both bacteria and fungi, zones of inhibition were visually measured in millimeters as the diameter of no growth around each filter-paper disk. Fungal-spore inhibition trials were examined 1, 24, and 36 h after inoculation. Spore germination was determined using a Nikon phase-contrast microscope set at 200x magnification. Counts were made 1 and 36 h after incubation for germinating and nongerminated spores in three fields of each agar plate.

Cultural conditions for the bacteria were as follows. (1) Bacillus subtilis: motile, gram-positive rods cultured aerobically at 30C on tryptic soy agar (TSA) growth medium (Burtt and Ichida 1999). (2) Bacillus licheniformis strains OWU 138B and OWU 1432B: keratinolytic (i.e. known to degrade feathers) grampositive rods reared from wild birds on TSA and cultured at 30C on TSA growth medium (Burtt and Ichida 1999). Cultural conditions for the fungi were as follows. (1) Chaetomium globosum (Ascomycetes), a cellulose- decomposer, cultured at 25C on potato dextrose agar (PDA) growth medium (Hubalek 1978). (2) Penicillium chrysogenum (Deuteromycetes) and T. viride (Deuteromycetes) cultured at 25C on PDA growth medium (Hubalek 1978).

Data collected from all experimental trials were assessed through visual inspection. Paired t-tests showing effects of formic acid on C. globosum, Pe. chrysogenum, and T. viride spore germination were calculated using the SPSS statistical package (SPSS, Chicago, Illinois). Percentages, standard error, and histograms were produced using Microsoft EXCEL software.

RESULTS

Neither formicine (Ca. pennsylvanicus and L. flavus) nor myrmicine ants (A. ruais, P. dentata, and Cr. lineolata) produced chemicals that inhibited common feather microbes. Neither the hexane extracts nor the ant suspensions in deionized water produced inhibition in any of the microbial species. Formic acid strongly inhibited all bacteria and fungal hyphae tested. Average inhibition zones (n = 10) produced by formic acid after 12 h were 30.7 mm for B. subtilis, 44.5 mm for B. licheniformis strain OWU 1432B, and 47.1 mm for B. licheniformis strain OWU 138B. For fungal hyphae cultures, average inhibition zones after 4 days were 32.0 mm for C. globosum, 22.0 mm for Pe. chrysogenum, and 49.0 mm for T. viride (Fig. 2). There were no inhibition zones around any of the controls or untreated disks.

Concentrations of mature-spore suspensions varied among fungal types. Forty-milliliter suspensions cultured on agar plates and analyzed through three 200x fields of view produced an average of 5 spores of T. viride, 88 spores of Pe. chrysogenum, and 54 spores of C. globosum.

FIG. 2. Diameter (mm) of inhibition (mean SE) in microbial inhibition trials with formic acid (n = 10 for formic acid; n = 50 for each microbial species). Abbreviations: B.s. = Bacillus subtilis, B.I. OWU 1432B = B. licheniformis strain OWU 1432B, B.I. 138B = B. licheniformis strain 138B, C.g. = Chaetomium globosum, T.v. = Trichoderma viride, and P.c. = Penicillium chrysogenum.

Fungal spore germination was not inhibited by the ant-suspension or hexane-extract treatments. However, fungal spore germination was moderately inhibited by the formic acid treatment; all the C. globosum spores were inhibited, an average of 3 of the total 5 spores of T. viride were inhibited, and 87 of the 88 spores of Pe. chrysogenum were inhibited (Table 1). Germination of C. globosum fungal spores was significantly inhibited by formic acid (paired t- test = -2.936, df = 8, P = 0.019). However, the formic acid treatment did not have a significant effect on spore germination in Pc. chrysogenum (paired f-test= -2.137, df = 5, P = 0.086) or T. viride (paired t-test = -1.437, df = 8, P = 0.189).

DISCUSSION

Feather ectoparasites have the ability to reduce host fitness (Burtt 1999). Potential effects are present in every aspect of avian life, including mate selection (Clayton and Tompkins 1995), fecundity and successful rearing of hatchlings (Burtt and Ichida 1999), and host survival (Booth et al. 1993). Bird feathers serve important functions in both flight and thermal protection (Proctor and Lynch 1993). Microorganisms can degrade feathers and may act as a selective force in the evolution of molting (Burtt and Ichida 1999). Ability to suppress ectoparasites through self-medication has potential to increase fitness levels among individuals and within populations (Clayton and Wolfe 1993).

TABLE 1. Paired t-test results showing effects of formic acid on Chactomium globosum, Penicillium chrysogenum, and Trichoderma viride spore germination. "Mean germination" represents mean number of spores contained in three 200x microscopic fields of view.

Although microfibrils of twisted keratin in feathers may resist biological degradation, keratinolytic bacteria and fungi have been identified as potential destructive agents (Burtt and Ichida 1999). Microorganisms occurring in high frequencies in cultures from birds were identified by Brittingham et al. (1988). Most of those feather- degrading bacteria also occur commonly in soils (Hubalek 1978, Brittingham et al. 1988, Burtt and Ichida 1999). Most songbirds come in contact with soil particles and may thus acquire multiple microorganisms harmful to their feathers.

Preening behaviors in birds-including bathing in water or dust and coating feathers with oils from preen glands -are commonly thought to improve feather condition. Those behaviors may not substantially reduce feather ectoparasites. Clayton and Wolfe (1993) documented that surgical removal of the preen gland does not lead to increased populations of feather lice. It is possible that anting behavior combined with preening activities can decrease harmful feather parasites (Goodwill 1955, 1956; Simmons 1957; Ehrlich et al. 1986; Clayton and Vernon 1993; Clayton 1999). Songbirds in the genus Pitohui from New Guinea have evolved a chemical defense against feather ectoparasites in the steroidal alkaloid homobatrachotoxin (Dumbacher et al. 1992). The same chemical found in the dendrobatid poison-dart frogs (Phyllobates spp.), homobatrachotoxin is lethal to chewing lice. Pitohui species deter ectoparasitic feather lice with homobatrachotoxin sequestered in feathers and muscle tissue (Dumbacher 1999).

Ants are known to produce numerous structurally diverse chemicals, and many have been identified in common ants of the U.S. east coast. Hlldobler and Wilson's (1990) definitive work identified the many glandular and pheromone-producing organs in ants. In the experiments described here, we used ants from two subfamilies and five genera. Glandular chemicals have been identified in four of those genera: Aphaenogaster (Wilson 1971, Hlldobler 1995, Hlldobler et al. 1995), Crematogaster (Hlldobler and Wilson 1990), Camponotus (Wilson 1971, Hlldobler and Wilson 1990, Brand and Morgan 1999, Pempo et al. 2000), and Lasius (Wilson 1971, Hlldobler and Wilson 1990).

We extracted ant-chemical compounds from the entire ant body to simulate possible application of chemicals during anting behavior. Our experiments tested the hypothesis that ant chemical secretions inhibit microbial growth. Anting behavior might be considered adaptive if growth of bacteria or fungal hyphae, or germination of fungal spores, were reduced by application of ant chemicals. However, neither the hexane extracts nor the ant suspensions produced such an inhibitory effect in our trials. In contrast, formic acid inhibited growth of all microbial species tested and significantly inhibited spore germination in C. globosum (though not in Pe. chrysogenum or T. viride). Thus, although formic acid clearly has an inhibitory effect on feather-degrading bacteria and fungi, our results suggest that ants are not selected for anting behavior on the basis of an ability to inhibit microbial growth.

The 60% aqueous chemical mixture produced by some ants contains dilute conce\ntrations of formic acid (Blum 1992). In the secretions of the ant species we studied, formic acid was present in concentrations lower than those at which it has been shown to be effectively antimicrobial. Our results show that, as a component within secretions of Co. pennsylvanicus and L. fluvus worker-caste ants, formic acid does not inhibit microbial growth. Similar results have been found for arthropods: formic acid was shown to kill feather mites, chewing lice, and ticks in vitro (Eichler 1936 in Judson and Bermett 1992); but applications of formic acid as a component of ant secretions failed to produce mortality in an aviary setting (Judson and Bennett 1992).

The hypothesis that deleterious microbial colonies commonly found on feathers can be reduced by application of ant-chemical extracts is not supported by the present study, though we tested ants from both Formicinae and Myrmicinae, the two most commonly identified ant subfamilies used in anting behavior (Simmons 1957, Whitaker 1957). Six other hypotheses have been suggested to explain why birds perform anting behavior: (1) to remove stale lipids from skin and feathers (Kelso 1946, Kelso and Nice 1963, Simmons 1966); (2) to provide autoerotic stimulation (Whitaker 1957); (3) to store ants in feathers as a reservoir food supply (Groskin 1943); (4) to facilitate molting and soothe feathers (Potter 1970); (5) to prepare food by removal of chemical secretions of ants, including formic acid (Judson and Bennett 1992); and (6) to reduce feather mites, ticks, and lice (Goodwin 1955, 1956; Simmons 1966; clayton and Vernon 1993).

Reduction of arthropod ectoparasites tends to attract widespread attention and appears to be the most accepted hypothesis among birders (Cape Hennery Audubon Society pers. comm., Outer Banks Audubon Society pers. comm., Zuni Hunt club pers. comm.). It is also one of the few hypotheses that has been quantitatively examined; however, experimental results do not support it (Judson and Bennett 1994, Clayton 1999). Whitaker (1957) found no evidence of ectoparasite mortality after anting episodes. Additionally, experiments testing anting substitutes similar to acids found in many ant secretions, such as lime (Citrus sp.) juice containing citric acid, produced no detrimental effects on feather lice (CIayton and Vernon 1993). Our results, combined with those earlier accounts, support the importance of examining alternative hypotheses to explain anting behavior.

ACKNOWLEDGMENTS

We thank E. H. Burtt, Jr. (Department of Zoology, Ohio Wesleyan University) for supplying B. licheniformis cultures. We are grateful for the support and encouragement of E. H. Burtt, Jr., D. Sonenshine, R. Rose, D. Naik, K. Nesius, and the support facility in the Department of Biological Sciences, Old Dominion University.

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Associate Editor: M. Brittingham

HANNAH C. REVIS1 AND DEBORAH A. WALLER

Department of Biological Sciences, Old Dominion University, Norfolk, Virginia 23529, USA

1 Present address: U.S. Pacific Basin Agricultural Research Center, P.O. Box 4459, Hilo, Hawaii 96720, USA. E-mail: hrevis@pbarc.ars.usda.gov

Copyright American Ornithologists' Union Oct 2004

Source: Auk, The

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