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Microbial Synergy via an Ethanol-Triggered Pathway

Michael G. Smith, Shelley G. Des Etages,^, and Michael Snyder^*

Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, Connecticut 06520

Received 6 October 2003/ Returned for modification 14 January 2004/ Accepted 30 January 2004

	ABSTRACT

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We have discovered a microbial interaction between yeast, bacteria, and nematodes. Upon coculturing, Saccharomyces cerevisiae stimulated the growth of several species of Acinetobacter, including, A. baumannii, A. haemolyticus, A. johnsonii, and A. radioresistens, as well as several natural isolates of Acinetobacter. This enhanced growth was due to a diffusible factor that was shown to be ethanol by chemical assays and evaluation of strains lacking ADH1, ADH3, and ADH5, as all three genes are involved in ethanol production by yeast. This effect is specific to ethanol: methanol, butanol, and dimethyl sulfoxide were unable to stimulate growth to any appreciable level. Low doses of ethanol not only stimulated growth to a higher cell density but also served as a signaling molecule: in the presence of ethanol, Acinetobacter species were able to withstand the toxic effects of salt, indicating that ethanol alters cell physiology. Furthermore, ethanol-fed A. baumannii displayed increased pathogenicity when confronted with a predator, Caenorhabditis elegans. Our results are consistent with the concept that ethanol can serve as a signaling molecule which can affect bacterial physiology and survival.

	INTRODUCTION

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In the natural environment, microbes exist in a state of constant interaction with other microbes. Most microbial interactions are antagonistic. Microbes compete for nutrients and inhibit the growth of their competitors by producing antimicrobial compounds. The most notable example of antibiosis was the discovery of penicillin by Fleming, in which Penicillium notatumin was found to inhibit the growth of Staphylococcus aureus (9). Positive interactions also occur: a reciprocal transfer of required nutrients has been demonstrated using Lactobacillus arabinosus and Streptococcus faecalis in minimal medium which enabled both to grow (29). While much effort has been devoted to describing bacterial cell-cell interactions, particularly antagonistic ones, surprisingly little attention has been devoted to yeast-microbe interactions.

Saccharomyces cerevisiae is an ideal organism in which to study how eukaryotic cells interact with other microorganisms. Yeasts are ubiquitous in nature, soil dwelling, and are also opportunistic pathogens (13, 26, 35, 37). The wealth of information about this fungus lends itself to the genetic, molecular biological, and microbiological techniques required for dissecting this eukaryote-microbe interaction.

We examined the interactions between naturally occurring strains of the budding yeast, S. cerevisiae, and a wide variety of bacteria. We found that the pathogenic isolates of yeast had the ability to affect the growth of the human pathogens Acinetobacter baumannii and Acinetobacter haemolyticus as well as several natural isolates of Acinetobacter spp. Acinetobacters are commonly found in soil, water, and sewage (15). It has been estimated that acinetobacters comprise as much as 0.001% of the population of heterotrophic aerobic bacteria in soil and water (1), illustrating their prevalence and versatility. Acinetobacters are best known for their ability to transform with DNA readily (18), their ability to utilize and degrade a wide range of carbon sources including petroleum (1, 3, 28), and for their rising incidence of multidrug-resistant, nosocomial-derived strains infecting immunocompromised individuals in hospitals worldwide (5, 10, 41).

Here we describe antagonistic interactions and an unexpected synergistic relationship between S. cerevisiae and Acinetobacter. This synergistic relationship involves the production of a diffusible factor by yeast that allows the bacteria to grow to higher cell density. The diffusible factor responsible for this synergistic relationship was explored further and found to be ethanol. Ethanol-fed acinetobacters can withstand salt stress and are more virulent to the bacterial predator Caenorhabditis elegans. These results suggest that ethanol can act not only as a carbon source but also as a signal which triggers one or more pathways that result in an alteration of a bacterium's physiology and survival.

	MATERIALS AND METHODS

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Strains and media. Yeast and bacterial strains used in this study are listed in Table 1. Yeast growth conditions were as described in Guthrie and Fink (13). YPAD contains 1% yeast extract, 2% Bacto-Peptone, 2% glucose, and 300 µM adenine. LB contains 1% Bacto-Tryptone, 0.5% yeast extract, and 1% NaCl. Lactic acid mineral medium (LAMM) contains 0.5% lactic acid, 10 mM KH₂PO₄, 100 mM Na₂HPO₄, 0.2% NH₄Cl, 1 mM MgSO₄, 0.001% CaCl₂, 0.0005% FeSO₄ (pH 6.65) (17).

Microbial interaction assay. Bacteria and yeast were grown separately overnight in YPAD at room temperature with shaking. The bacteria were then diluted and added to molten YPAD with agar (60°C) and overlaid onto cooled YPAD plates. Yeast were diluted and spotted onto the lawns of bacteria, and the plates were incubated overnight at room temperature. For liquid-based assays, yeast were grown to an optical density at 600 nm (OD₆₀₀) of 5 to 7 in YPAD, the cells were pelleted by centrifugation, and the conditioned medium (CY) was filter sterilized using a 0.22-µm-pore-size filter. Conditioned media were added to fresh media to a final volume of 10%. Bacterial precultures were grown overnight at room temperature in YPAD and diluted to an OD₆₀₀ of 0.01 in fresh YPAD, YPAD plus 10% conditioned medium, or YPAD plus 0.1% ethanol. Bacteria were incubated overnight at room temperature with shaking, and the OD₆₀₀ was measured to determine cell density. Aliquots were also removed, serially diluted, and plated onto YPAD plates to determine the number of CFU present in the samples. Pronase E (Sigma Chemical Co.) was added directly into molten agar (or to CY) to a final concentration of 50 µg/ml.

Ethanol determination. Ethanol concentrations were determined using a kit developed by Boehringer Mannheim (Darmstadt, Germany). Briefly, samples were diluted 1:1,000 in water and 100 µl was added to 3 ml of potassium diphosphate buffer. An absorbance reading was taken at 340 nm. An enzyme suspension containing alcohol dehydrogenase and aldehyde dehydrogenase was added, and a second OD₃₄₀ measurement was taken. The difference in the absorbances (A) was then factored into the following equation: concentration (in grams per liter) of ethanol = V x MW(g)/ x d x v x 2,000 x A, where V is the final volume (in milliliters), v is the sample volume (in milliliters), MW is the molecular mass of the substance to be assayed (in grams), d is the light path (in centimeters), and is the extinction coefficient.

C. elegans killing assay. Escherichia coli (strain OP50) cells grown on NGM medium (38) were fed to C. elegans (strain N2) worms. L3/L4-stage worms were placed onto lawns of Acinetobacter spp. grown on NGM, NGM plus 1% ethanol, PGS (1% Bacto-Peptone-1% NaCl-1% glucose-0.15 M sorbitol-1.7% Bacto-Agar), or PGS plus 1% ethanol. Plates were incubated at 20°C. Viability was tested every 24 h by visual examination. Worms were considered dead if they no longer moved or responded to touch.

	RESULTS

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Antagonistic and synergistic interactions between yeast and bacteria. To explore the interactions between yeast and other microbes, we tested natural isolates of S. cerevisiae for their ability to affect bacterial growth on solid media. Thirty yeast strains were tested: 20 strains were from immunosuppressed patients (12), 4 were from cactus plants (36), and 6 were standard laboratory strains (1 produced killer toxin, another was an isogenic killer toxin-defective strain, and the remaining 4 were unrelated killer toxin-defective, well-characterized laboratory strains) (Table 1). These different yeast strains were spotted onto lawns of bacteria, incubated overnight at room temperature, and examined for changes in the growth of the bacterial lawns (Fig. 1A). A total of 61 microbial strains from 15 genera were tested, including strains from Acinetobacter (33 strains), Pseudomonas (6 strains), Staphylococcus (4 strains), Agrobacterium (3 strains), Escherichia (2 strains), Salmonella (2 strains), Enterobacter (2 strains), and others (Table 1).

View larger version (42K): [in this window] [in a new window]   FIG.1. Antagonism and synergism by a diffusible factor. Standard laboratory yeast strains W303 and S288c and clinical isolates YJM145, YJM835, YJM939, and YJM 940 were spotted onto lawns of Acinetobacter strains ADP7594, AD321, and ADP1. (A) Halos of ADP7594 nongrowth surrounding yeast spots, especially around the clinical strains. AD321 growth was enhanced around spots of YJM145 and YJM835. Strain ADP1 showed neither inhibited nor enhanced growth. (B) AD321 cells were mixed with YJM835 yeast cells, serially diluted, and spread onto plates of YPAD. After 24 h of incubation at room temperature, the bacterial colonies closest to the yeast colony (arrow) were observed to be larger in diameter than those bacterial colonies further away from the yeast spot. (C) Cells were incubated in YPAD at room temperature with shaking. Open icons indicate growth in YPAD. Closed icons indicate growth in YPAD supplemented with 10% conditioned YPAD. Medium was conditioned as described in Materials and Methods. Squares, strain AD321; circles, strain ADP1.

Enhanced bacterial growth is due to a diffusible factor. The enhanced growth of Acinetobacter in the presence of yeast was not anticipated and therefore was studied further. Coculturing experiments were performed to determine if the yeast-derived factor was diffusible. While several isolates of Acinetobacter displayed the enhanced growth phenotype, AD321 was used exclusively for the determination of the yeast-derived factor, since it displayed the highest level of growth enhancement of all the strains tested. A low number of YJM835 yeast cells were mixed with Acinetobacter strain AD321, serially diluted, and spread onto YPAD plates. In the experiment illustrated in Fig. 1B, a single yeast colony was surrounded by many bacterial colonies. The AD321 colonies closest to the yeast colony grew to a much larger diameter (about 10 times larger) than those that resided farther away. Thus, the yeast-derived factor is diffusible, and direct contact of the two species is not required to enhance bacterial growth.

To test directly whether the growth enhancement of bacteria requires the presence of yeast cells, strains of Acinetobacter were incubated in fresh medium (YPAD) supplemented with filter-sterilized medium prepared from overnight cultures of YJM835 yeast. The conditioned yeast medium (CY) was added to fresh medium to a final concentration of 10%. A time course of Acinetobacter growth was then plotted using the OD₆₀₀ as a measure of cell growth. We found that Acinetobacter strain AD321 grew to 2.0 ± 0.1 times the cell density (OD₆₀₀, 6.1 ± 0.7 versus 3.1 ± 0.5) in 10% CY compared to cells grown in YPAD alone (Fig. 1C). Analysis of the growth curves indicated that bacterial cells in CY medium grew at the same rates as controls but to higher cell densities. This increase in optical density of the bacterial cultures after 24 h was always accompanied by a corresponding increase in CFU from these cultures ([4.61 ± 1.3] x 10¹¹ CFU/ml for cells grown in 10% CY, compared to [1.67 ± 0.8] x 10¹¹ CFU/ml for cells grown in YPAD alone). Control strain ADP1 was neither enhanced nor inhibited by CY in either the plate assay or the liquid assay (Fig. 1C). These data suggest that yeast enhance bacterial cell growth not by altering growth rates but instead by affecting the final cell densities to which these cells can grow.

One obvious explanation for this effect on bacterial growth is that YPAD is a suboptimal medium for bacterial growth and the yeast are providing nutrients that are normally present in bacterial growth media. To that end, the liquid assay described above was repeated using the bacterial media LB or LAMM with or without the CY supplement. We found that strain AD321 was enhanced by CY to the same extent when added to either LB or LAMM (40.5% ± 7.0% increase) as it is when grown in YPAD. Moreover, the enhanced growth was not due to limited amino acids or nitrogen; doubling the amount of amino acids or primary nitrogen source in the LB medium compared to LAMM did not enhance the growth of bacteria. Thus, the bacterial growth-enhancing factor is not a nutrient that yeast contribute to the media, but a different yeast-derived compound.

The stimulatory growth factor accumulates in mid-log-phase yeast cultures and is a small molecule. To determine the nature of the production of the yeast-derived bacterial growth-enhancing compound, strains of Acinetobacter were incubated in media (YPAD) supplemented with filter-sterilized medium prepared from pre-log-, early-log-, mid-log-, or late-log-phase yeast cultures. We found that the mid-log- and late-log-phase yeast cultures were best able to enhance the growth of bacteria (Fig. 2). This suggests that the factor responsible for enhancing bacterial growth accumulates preferentially in late-phase yeast cultures.

View larger version (15K): [in this window] [in a new window]   FIG. 2. AD321 is further enhanced by older yeast cultures. YJM835 yeast were grown in YPAD at 30°C. The cell densities were determined, and the culture medium was filter sterilized. These media were then added to fresh YPAD at a final concentration of 10% and used as a culture medium for Acinetobacter strain AD321 cells. After overnight growth, the OD600 of the Acinetobacter cultures was determined. Bacterial cell densities are shown as the percent enhancement compared to that of bacterial cells grown in YPAD alone.

The enhanced growth factor is ethanol. One compound that possesses all of the properties of the growth-enhancing factor (produced by late-log cultures of yeast, small, heat and protease resistant, volatile) is ethanol. Thus, we tested the ability of ethanol to affect the growth of AD321 (Fig. 3). A 95% ethanol solution was diluted in YPAD to various concentrations ranging from 0 to 9.5%. Acinetobacter strain AD321 growth was enhanced by 50% in medium containing low levels of ethanol (0.1%), and the bacterial cell density more than doubled in medium containing between 1 and 4% ethanol; at concentrations of 4.5 and 5% ethanol, these cells grew only as well as in YPAD with no ethanol supplement. We then retested several of the isolates that we recovered in our original screen, and all of the isolates were enhanced by low concentrations of ethanol (up to 2 to 3%). We did observe strain-specific differences in the level of enhancement and tolerance to ethanol. For example, A. baumannii was enhanced maximally by 1% ethanol, while A. johnsonii and A. radioresistens grew best in 2 to 3% ethanol. A. haemolyticus was the most sensitive to ethanol concentration, since it could be enhanced by low levels (0.1 to 1%) but was inhibited at higher concentrations (>3%).

View larger version (15K): [in this window] [in a new window]   FIG. 3. Effect of ethanol on AD321 growth. Ethanol concentration was titrated in YPAD from 0 to 9.5% and tested for its ability to enhance the growth of AD321 cells. Bacteria were grown overnight at room temperature, and optical densities were determined. Cell densities are shown as the percent enhancement or inhibition compared to that of cells grown in YPAD alone.

View larger version (10K): [in this window] [in a new window]   FIG. 4. Ethanol, but not other organics, enhances Acinetobacter growth. Bacteria were grown in YPAD or YPAD supplemented with ethanol, methanol, butanol, or dimethyl sulfoxide. The concentration of each organic used was 0.1, 0.5, 1.0, 5.0, or 9.5%. Bacteria were grown overnight at room temperature, and cell densities were measured by the OD600. Cell densities are shown as the percent enhancement or inhibition compared to that of cells grown in YPAD alone.

To determine if ethanol production is required for production of bacterial growth enhancement, we grew YJM835 on a variety of carbon sources and tested its ability to affect bacterial growth (Fig. 5A). Glucose is metabolized via glycolysis into pyruvate, which is subsequently converted into either ethanol (fermentation) or carbon dioxide (respiration). Yeasts ferment sugars even when grown aerobically (22, 23). Some carbon sources, such as glycerol, which is nonfermentable, require respiration and, thus, result in no ethanol production. YJM835 grown in the fermentable sugars glucose, fructose, or sucrose produced 0.93% ± 0.2% ethanol and enhanced bacterial growth by 53.6% ± 6.3%. Cells grown in glycerol neither produced significant amounts of ethanol (0.03% ± 0.1% ethanol) nor enhanced bacterial growth (–2.54% ± 2.97% growth enhancement), suggesting that yeast-derived ethanol may be the stimulatory component.

View larger version (26K): [in this window] [in a new window]   FIG. 5. Ethanol is required for bacterial growth enhancement. (A) YJM835 cells were grown in YPAD or YP plus each of the carbon sources listed to a cell density of 5 x 107 to 7 x 107 cells/ml. Media were sterilized, and ethanol concentration was determined as described in Materialsand Methods. Bacteria were grown overnight in CY medium at room temperature, and cell densities were measured by the OD600. Cell densities are shown as the percent enhancement or inhibition compared to that of cells grown in YPAD alone. (B) Gene deletions were performed as described in Materials and Methods. Cells were grown in YPAD, conditioned media were sterilized, ethanol concentrations were determined, and bacteria were cultured as described above. (C) YJM835 cells were grown in YPAD or YP plus glucose at the concentrations listed to a cell density of 5x 107 to 7 x 107 cells/ml. Conditioned media were sterilized, ethanol concentrations were determined, and bacteria were cultured as described above. For all panels, columns represent bacterial growth enhancement and lines indicate ethanol concentration.

We reasoned that if ethanol concentration were the primary determinant for enhancement of bacterial growth, then increasing the amount of ethanol produced by yeast should also result in an increase in bacterial growth enhancement up to a certain percentage, as shown in Fig. 3. For these experiments, we grew yeast strain YJM835 overnight in YPAD containing 4, 8, or 10% glucose (standard YPD contains 2% glucose). In all cases, we measured the amount of ethanol produced by the yeast and the level of bacterial growth enhancement. YJM835 yeast produced 0.9% ± 0.19% ethanol from 2% glucose, whereas 2.0% ± 0.18% ethanol was produced from 4, 8, or 10% glucose (Fig. 5C). Clearly, saturation was reached, since these yeast could not produce more than 2% ethanol despite the increasing amounts of glucose provided. Nevertheless, bacterial growth enhancement was ethanol dependent. For example, additional ethanol was produced with between 2 and 4% glucose, and under these conditions bacterial growth was enhanced by 17 to 18% (Fig. 5C). Yeast incubated without glucose failed to grow or produce ethanol and were also ineffective at stimulating bacterial growth. These studies indicate that ethanol is necessary and sufficient to stimulate acinetobacter growth.

Ethanol induces a specific cell tolerance response. Along with salt and heat, ethanol is a commonly used stimulus to induce the general stress response in many bacteria (14, 30, 32). While the typical ethanol concentrations used for general stress stimulation are considerably higher than the concentrations produced by yeast in YPAD (4 versus 1%), it is possible that low concentrations of ethanol can also elicit a stress response. In low doses, ethanol might serve a signaling role by specifically altering the physiology of the bacterial cells. It has long been known that ethanol can induce thermotolerance in yeast and mammals (24, 31), and we reason that ethanol may similarly induce tolerance of other stresses in Acinetobacter.

We incubated Acinetobacter cells in high salt, H₂O₂, or high temperatures in the presence or absence of yeast-conditioned medium or 0.1% ethanol. Addition of NaCl to the culture medium restricted the growth of all strains tested: 1% NaCl inhibited growth 16.8 to 21.1%, 2.5% NaCl inhibited growth by up to 65.9%, and 5% NaCl inhibited growth by more than 94.6% (Fig. 6A). However, addition of either yeast-conditioned medium or exogenously added ethanol protected all bacterial strains from the negative effect of the high salt, as depicted by the optical density and relative increase in growth (Fig. 6A and B). Similar results were obtained when the cells were challenged with KCl or CaCl₂ (data not shown). Unlike the protection from high salt, AD321 cells challenged with high temperature or oxidative damage were not protected by conditioned yeast medium or ethanol. Combined, these data suggest that ethanol is not stimulating the general stress response but is instead inducing a specific response, in this case, enabling growth on high salt.

View larger version (32K): [in this window] [in a new window]   FIG. 6. Ethanol increases salt tolerance of acinetobacters. Bacteria were grown in YPAD or YPAD supplemented with 0.1% ethanol. (A) Cells were challenged with various concentrations of NaCl as follows: no salt (A and E); 1% (B and F); 2.5% (C and G); and 5% NaCl (D and H). Bacteria were grown overnight at room temperature, and cell densities were measured by the OD600. Cell densities are shown as absolute values. (B) Resulting cell densities, shown as the percent enhancement or inhibition compared to that of cells growth without ethanol. The blue bars represent media containing no salt, red bars represent media containing 1% NaCl, and the yellow bars represent media containing 5% NaCl.

Ethanol increases the pathogenicity of A. baumannii. Chemical stresses are not the only stresses that confront organisms in nature. To survive predation, prey species have evolved a myriad assortment of defense mechanisms. In some cases, prey species have even become parasitic to their predator-host. Several Acinetobacter species are parasitic; in particular, A. baumannii has become a problematic human pathogen worldwide. It is often the case that the extent to which a parasite disables its host is directly correlated with its own reproductive capacity. It is reasonable to suggest, then, that conditions which lead to an increased reproductive capacity of A. baumannii could also lead to an increase in pathogenicity.

To test this hypothesis, we utilized the free-living nematode C. elegans as predator-host of A. baumannii. C. elegans has been used in the past to study the pathogenicity of several bacteria, including Pseudomonas aeruginosa and Serratia marcescens (20, 21, 39, 40). These studies illustrated not only that bacteria can infect and kill worms, but also that composition of the medium can influence pathogenicity. Rich medium induced the expression of virulence genes which resulted in a "fast killing" phenotype (39, 40). Thus, we tested the effect of ethanol on the ability of A. baumannii to kill worms. Incubation of L4-stage worms on lawns of A. baumannii grown on NGM (minimal medium; see Materials and Methods) resulted in the proliferation of worms at a rate comparable to the growth observed when the worms were fed E. coli control strain OP50. We conclude that C. elegans can consume Acinetobacter spp. cells and use them as a food source. Incubation of L4 worms on lawns of A. baumannii grown on the rich medium PGS resulted in worm lethality. The LT₅₀ (time for half of the worms to die) on A. baumannii grown on PGS was 180 ± 36 h (7.5 days; n = 77) (Fig. 7A). The LT₅₀ for worms that were fed A. baumannii grown on PGS plus 1% ethanol was 126 ± 12 h (5.5 days; n = 86). This represents a 30.0% increase in the rate of death of the worms supplied with ethanol-fed bacteria. In contrast, worms fed E. coli OP50 grown on PGS displayed an LT₅₀ of 264 ± 32 h (11 days; n = 52) in the absence of ethanol and 256 ± 32 h (10.7 days; n = 50) (Fig. 7B) in the presence of ethanol in the media under our experimental conditions. These data indicate that C. elegans can be used as a model system for A. baumannii pathogenesis and that ethanol stimulates an increase in A. baumannii virulence.

View larger version (17K): [in this window] [in a new window]   FIG. 7. Ethanol promotes virulence of A. baumannii. Bacteria were grown overnight in LB and spread onto PGS agar plates with (open squares) or without (solid squares) the addition of 1% ethanol. L3/L4-stage worms were placed onto each lawn of bacteria and incubated at 20°C. The plates were then scored for live worms every 24 h. Worms were considered dead when they no longer responded to touch. (A) Worms fed E. coli OP50; (B) worms fed A. baumannii.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Ethanol stimulates acinetobacters to grow to higher cell density, as measured both by optical density and by counting the number of CFU. Since ethanol is consumed by these bacteria (unpublished observation), it is likely utilized as a carbon source (albeit a poor one, since it is present at low levels). However, the benefit of ethanol to acinetobacters extends beyond increasing cell number. Ethanol stimulates salt tolerance but not thermotolerance or resistance to oxidative damage in acinetobacters. Ethanol-fed acinetobacters can also kill a natural predator more efficiently than bacteria fed other carbon sources. These data indicate that ethanol also induces signaling pathways required for specific stress tolerance and virulence.

Although our studies were confined to the laboratory, we expect them to be pertinent to nature, as they involve organisms that we predict to interact in nature. Standard laboratory strains of yeast were shown to be unable to produce the quantity of ethanol required to enhance bacterial growth. In contrast, natural isolates and, in particular, pathogenic isolates of yeast were able to generate sufficient amounts of ethanol to affect the growth of acinetobacters. S. cerevisiae and Acinetobacter are ubiquitously found in soil, water, and vegetation, they both prefer acidic pH (5.5 to 6.0) environments (2, 3, 13, 37), and they can both be opportunistic human pathogens (5, 10, 26, 41). Thus, we expect them to occupy the same ecological niches and have the potential for direct interactions in the wild.

It is possible that the relationship between the two microbes is proto-commensalistic. That is, in certain environments the bacteria benefit, while in others the yeast benefit. The preferred carbon source for the yeast Saccharomyces is sugar, as its name indicates. Yeast typically ferment sugar into ethanol and carbon dioxide, even in the presence of oxygen, which is curious given the unfavorable energetics of the reaction (7, 22, 23). Acinetobacters are obligate aerobes and are known to consume a wide range of carbon sources, although only a few strains can utilize glucose (3, 16). Thus, many acinetobacters would be dependent on yeast to convert a plentiful sugar source into a more readily catabolized one, namely ethanol. The reciprocal situation may also exist, in which acinetobacters metabolize compounds that yeast cannot and provide yeast with a more suitable metabolite.

Alternatively, but not exclusively, the yeast may benefit indirectly from the bacteria via a form of mutualistic antipredation. Recently, C. elegans has been shown to be able to consume yeast as a food source (27). Since ethanol-fed acinetobacters can kill worms more rapidly than those without ethanol, both microbes stand to gain from the elimination of their common predator.

The deleterious effect of ethanol-fed A. baumannii on worms may be due to expression of bacterial virulence genes that cause physical harm to the worms. Alternatively, the bacteria may be responding to the presence of ethanol by altering their physiology such that they provide the worms with a lesser-quality food source and lead, ultimately, to the decreased worm life span observed. A third possibility is that the presence of ethanol in the medium may affect worms independently of its effect on bacteria. While ethanol in the medium, per se, does not adversely affect worm life span as evidenced by the normal life spans observed with ethanol-fed OP50, ethanol may negatively affect the immune system of the worms. These immunocompromised worms would then be more susceptible to A. baumannii infection. Separating these possibilities will be the subject of future experiments.

	ACKNOWLEDGMENTS

 
We thank John McCusker for the pathogenic yeast strains and Andre Lachance for the natural yeast isolates. We also thank Nick Ornston, Arnold Barton, Thomas Burr, and Stefan Pukatzki for bacterial strains. The work involving C. elegans could not have been performed without the assistance of Frank Slack and his lab, in particular, Monica Vella and Stephen Johnson. We also thank William Firshein, Anthony Infante, Dan Nowakowski, Dan Gelperin, Antonio Casamoyer, and Anthony Borneman for their critical reviews of the manuscript. M.G.S. especially thanks Dave Young for strains, suggestions, and invaluable support.

	FOOTNOTES

 
* Corresponding author. Mailing address: Department of Molecular, Cellular and Developmental Biology, 266 Whitney Ave., New Haven, CT 06520. Phone and fax: (203) 432-6161. E-mail: michael.snyder{at}yale.edu.

M.G.S. and S.G.D.E. contributed equally to this work.

Present address: Genomic and Proteomic Sciences, Pfizer Global Research and Development, Pfizer, Inc., Groton, Conn.

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