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Applied and Environmental Microbiology, March 2004, p. 1307-1314, Vol. 70, No. 3
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.3.1307-1314.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Physiology and Nutrition of Treponema primitia, an H₂/ CO₂-Acetogenic Spirochete from Termite Hindguts

Joseph R. Graber and John A. Breznak^*

Department of Microbiology and Molecular Genetics and Center for Microbial Ecology, Michigan State University, East Lansing, Michigan 48824

Received 12 August 2003/ Accepted 27 November 2003

Top Abstract Introduction Materials and Methods Results Discussion References

 
Treponema primitia strains ZAS-1 and ZAS-2, the first spirochetes to be isolated from termite hindguts (J. R. Leadbetter, T. M. Schmidt, J. R. Graber, and J. A. Breznak, Science 283:686-689, 1999), were examined for nutritional, physiological, and biochemical properties relevant to growth and survival in their natural habitat. In addition to using H₂ plus CO₂ as substrates, these strains were capable of homoacetogenic growth on mono- and disaccharides and (in the case of ZAS-2) methoxylated benzenoids. Cells were also capable of mixotrophic growth (i.e., simultaneous utilization of H₂ and organic substrates). Cell extracts of T. primitia possessed enzyme activities of the Wood/Ljungdahl (acetyl coenzyme A) pathway of acetogenesis, including tetrahydrofolate-dependent enzymes of the methyl group-forming branch. However, a folate compound was required in the medium for growth. ZAS-1 and ZAS-2 growing on H₂ plus CO₂ displayed H₂ thresholds of 650 and 490 ppmv, respectively. Anoxic cultures of ZAS-1 and ZAS-2 maintained growth after the addition of as much as 0.5% (vol/vol) O₂ to the headspace atmosphere. Cell extracts exhibited NADH and NADPH peroxidase and NADH oxidase activities but neither catalase nor superoxide dismutase activity. Results indicate that (i) T. primitia is able to exploit a variety of substrates derived from the food of its termite hosts and in so doing contributes to termite nutrition via acetogenesis, (ii) in situ growth of T. primitia is likely dependent on secretion of a folate compound(s) by other members of the gut microbiota, and (iii) cells possess enzymatic adaptations to oxidative stress, which is likely to be encountered in peripheral regions of the termite hindgut.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Spirochetes are among the most abundant microbial groups in termite hindguts, accounting for as much as one-half of the prokaryotic community (42). For more than a century, however, our knowledge of these organisms was largely limited to sporadic reports of their presence in various termite species, their morphological diversity, and their physical association with termite gut protozoa (8). Although elimination of spirochetes from the termite gut led to a decrease in termite survival (20), the specific roles of spirochetes and the factors contributing to their abundance in the hindgut have remained obscure.

Over the past 10 years, our understanding of termite hindgut spirochetes has advanced dramatically. Cultivation-independent molecular approaches revealed that they group within the genus Treponema and that the large majority of 16S rRNA gene clones form a phylogenetically discrete cluster (the "termite cluster") within this genus (34). These studies also revealed a striking degree of phylogenetic diversity among termite gut treponemes, with as many as 21 distinct species occurring within a single termite host species (34). A few years ago, the first pure cultures of these organisms were isolated in our laboratory and were found to possess metabolic capabilities hitherto unknown in the Spirochaetes division of the Bacteria, including acetogenesis from H₂ plus CO₂ (31) and N₂ fixation (33). Both of these processes are unique to prokaryotes and have been demonstrated to be important in the provision of carbon, nitrogen, and energy to termites (6, 46). Acetogenesis plays a particularly prominent role in termite nutrition: 71 to 100% of the insect's energy requirements can be met by oxidation of acetate produced by hindgut microbes, and 10.5 to 33% of this acetate production is attributable to H₂-CO₂ acetogenesis (6, 41, 51).

The availability of pure cultures of termite gut spirochetes has enabled us to explore properties relevant to their growth and survival in situ. In this paper, we report on the nutritional, physiological, and biochemical properties of Treponema primitia strains ZAS-1 and ZAS-2, H₂/CO₂-acetogenic spirochetes isolated from the hindgut of the California dampwood termite, Zootermopsis angusticollis (Hagen) (31). Additional information regarding the taxonomy, nomenclature, and genomic properties of these strains is reported in a companion paper (23).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Media and cultivation methods.
Routine cultivation of T. primitia strains ZAS-1 (DSMZ 12426) and ZAS-2 (DSMZ 12427; ATCC BAA-887) was carried out in butyl rubber-stoppered anaerobe tubes or bottles (30) containing 2YACo medium (31) at no more than one-fifth their volume under an atmosphere of 80% H₂-20% CO₂ (vol/vol). 2YACo medium consisted of inorganic salts including trace elements, a vitamin mixture (10 ng of biotin/ml, 40 ng of 4-aminobenzoic acid/ml, 50 ng [each] of calcium-D-(+)-pantothenate and vitamin B₁₂/ml, 150 ng of pyridoxamine · HCl/ml, and 100 ng [each] of nicotinic acid and thiamine · HCl/ml [final concentrations]) (55), a mixture of 11 cofactors (50 µg [each] of folic acid and thiamine pyrophosphate/ml, 10 µg of riboflavin/ml, 5 µg [each] of pyridoxal · HCl and pyridoxal phosphate/ml, 1.3 µg of hemin/ml, 1 µg [each] of calcium folinate, ß-NAD, flavin adenine dinucleotide, and coenzyme A (CoA)/ml, and 0.5 µg of nicotinamide/ml [final concentrations]) (31), and 2% (vol/vol) laboratory-prepared yeast autolysate (equivalent to ca. 2.2 mg of dry solids/ml) (31). The medium was buffered by inclusion of 70 mM NaHCO₃ and 10 mM 3-N-[morpholino] propanesulfonic acid (MOPS) and was reduced with dithiothreitol (final concentration,1 mM). The pH of the medium prior to inoculation was 7.2. Unless otherwise noted, cultures were grown at 30°C on a reciprocal shaker (50 strokes per min) with vessels held in a horizontal position.

Nutritional and growth studies.
The ability of commercial yeast extracts to replace laboratory-prepared yeast autolysate in 2YACo medium was tested by using the following products at a final concentration of 6 mg/ml: Tastone 900, 154, and 310; Amberex 1003 and 695; and Amberferm 5925, 5902, and 5021 (all from Red Star Bioproducts, Juneau, Wis.). Cofactor requirements were evaluated by testing for attenuation of growth in 2YACo medium lacking one of the 11 cofactors (listed above). Candidate individual cofactors were then tested in reciprocal experiments in which they were incorporated alone into 2YACo medium instead of the 11-cofactor mixture.

Substrate utilization studies were performed with cells growing under an N₂-CO₂ (80:20 [vol/vol]) atmosphere. For ZAS-1, 2YACo medium was modified to contain Amberferm 5902 (4 mg/ml) in place of yeast autolysate to minimize background growth in medium lacking a test substrate. Unmodified 2YACo medium was used for ZAS-2, with a small amount of H₂ added to the N₂-CO₂ headspace (final concentration, 16 mM H₂; ca. 95,000 ppmv) to aid in the initiation of growth (see below). H₂ was not added to cultures grown on methoxylated aromatics. An increase in cell yield (>20%) in the presence of a substrate, compared to its absence, was taken to indicate utilization of the substrate as an energy source. Cell growth was determined by measuring the optical densities (OD) of cultures at 600 nm with a Milton Roy Spectronic 20 colorimeter. OD readings were converted to cell numbers by reference to a standard curve relating these quantities. Substrate carbon balances were determined under conditions of substrate-limited growth. Carbon recoveries for methoxylated aromatic compounds were calculated on the basis of the acetate production expected from demethylation of the aromatic (R) substrate according to the following equation: R(-OCH₃)_n + 0.5n CO₂ R(-OH)_n + 0.75n CH₃COOH + 0.5n H₂O. Organic acid production was determined by using a high-performance liquid chromatograph (HPLC) with refractive index detection (4). Aromatic compounds were analyzed by using a Beckman model 127 HPLC equipped with a model 168 photodiode array detector and an Alltech Lichrosorb RP-18 column (250 by 4.6 mm; particle size, 10 µm). The mobile phase was 0.1% phosphoric acid with a methanol gradient, increasing linearly from 48 to 55% in 30 min. The flow rate was 1.5 ml/min.

To test for mixotrophic growth, strain ZAS-2 was grown with shaking as described above in 750-ml bottles containing 100 ml of 2YACo medium with 2 mM maltose and a 650-ml headspace composed of 20% H₂, 20% CO₂, and 60% N₂ (vol/vol). Consumption of H₂ was followed by gas chromatography (5). Maltose consumption and organic acid production were followed by an anthrone assay (1) and HPLC analysis (described above), respectively.

Determination of hydrogen thresholds.
Hydrogen thresholds were determined as described by Lovley (36). In brief, cultures were grown under H₂-CO₂ (80:20 [vol/vol]) in 2YACo medium that was either left unmodified (ZAS-2) or modified to contain 4 mg of Amberferm 5902/ml in place of yeast autolysate (ZAS-1). When cultures reached mid-log phase (at which point further growth of both strains was strictly dependent on the presence of H₂), the headspace was replaced with N₂-CO₂ (80:20 [vol/vol]), followed by the introduction of ca. 6,000 ppmv of H₂ into the gas phase. The basal level to which this H₂ was consumed (i.e., the H₂ threshold) was determined through three cycles of H₂ addition and consumption for each culture. H₂ was measured by using a Trace Analytical RGA2 gas chromatograph equipped with an RGD2 trace gas detection unit.

Enzyme assays.
Cells from mid-log-phase cultures growing on H₂ plus CO₂ (OD, 0.15 to 0.3) were harvested by centrifugation (at 16,000 x g for 10 min) and resuspended at 10 times their original concentration in an appropriate assay buffer (as cited below) containing 10 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, and leupeptin (10 µg/ml) to inhibit proteases. While held at 4°C, cells were disrupted by sonication three times for 30 s each time with a Branson model 450 Sonifier (power setting, 5; 50% duty cycle) equipped with a stepped microtip. The resulting crude cell extracts were assayed for enzyme activities.

Formyltetrahydrofolate (formyl-THF) synthetase, methenyl-THF cyclohydrolase, and methylene-THF dehydrogenase (40) and methylene-THF reductase (38) were assayed as described previously. THF and THF derivatives used in the assays were obtained from Schircks Laboratories (Jona, Switzerland). Catalase was assayed by measuring the rate of decrease in the A₂₄₀ of H₂O₂ (2). Rates of O₂-dependent and H₂O₂-dependent oxidation of reduced pyridine nucleotides (oxidase and peroxidase activities, respectively) were determined as described by Stanton (49). Superoxide dismutase (SOD) was assayed by the xanthine-xanthine oxidase-cytochrome c reduction method (21). The protein content of cell extracts was measured by the Lowry assay (37). Absorbance measurements were made using a Perkin-Elmer Lambda 14 UV/VIS spectrophotometer.

Oxygen tolerance.
Cells growing under anoxic conditions were tested for the ability to maintain growth after the addition of various concentrations of O₂ to the headspace. Replicate cultures were grown under H₂-CO₂ with reciprocal shaking (above) in 18-mm-diameter anaerobe tubes containing 5 ml of 2YACo medium modified by the inclusion of 10 mM maltose but containing no dithiothreitol reducing agent. When cells reached mid-log phase, the headspace (ca. 21 ml) was balanced to atmospheric pressure with H_2--CO₂, and sterile O₂ was injected to a final headspace concentration of 0.5, 1, 2.5, or 5% (vol/vol). Cultures were then immediately reincubated in a horizontal position with shaking to facilitate equilibration of gaseous O₂ with the liquid phase, and further growth was monitored as described above.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Nutrition and growth of T. primitia strains ZAS-1 and ZAS-2.
Cells of both strains grew in 2YACo medium under H₂ plus CO₂ within an initial pH range of 6.5 to 7.8, with an optimum at pH 7.2. No growth was observed in media with an initial pH of 6.0 or 8.0. Both strains grew well within a temperature range of 23 to 32°C, with an optimum at 30°C. No growth occurred at 4 or 34°C. Under optimum conditions, the shortest doubling time of cells was 22 h for ZAS-1 (with glucose [Table 1]) and 29 h for ZAS-2 (with H₂ plus CO₂ plus maltose [see below]).

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TABLE 1. Substrates utilized by T. primitia

A search for commercial yeast extracts to replace laboratory-prepared yeast autolysate in 2YACo medium was expanded from an unsuccessful earlier attempt (31). Tastone 900, Amberferm 5925, and Amberferm 5902 could replace yeast autolysate for the growth of strain ZAS-1, whereas Tastone 154, Tastone 310, Amberex 1003, and Amberex 695 were suitable replacements for the growth of strain ZAS-2. No single product was effective for both organisms. Growth rates and cell yields of ZAS-1 and ZAS-2 in media containing commercial yeast extracts (6 mg/ml) were similar to those measured in 2YACo medium.

Besides H₂ plus CO₂, a variety of hexoses, pentoses, and disaccharides were also used as energy sources and were fermented homoacetogenically (Table 1). Curiously, however, when grown on organic substrates under N₂-CO₂, strain ZAS-2 displayed prolonged lag phases (72 h) prior to exponential growth. The provision of small amounts of H₂ to the headspace (ca. 95,000 ppmv) or the use of larger inocula (>5% [vol/vol]) eliminated such lags (Fig. 1). Strain ZAS-2 was additionally able to utilize methoxylated aromatic compounds (syringate, ferulate, vanillate, and trimethoxybenzoate) as energy sources when they were supplied at 2.5 mM (higher concentrations inhibited growth). Cell doubling times were typically longer than 100 h on these substrates, and acetate production was consistent with demethylation of the compounds (i.e., the aromatic ring did not appear to be cleaved) (Table 1). This was confirmed for trimethoxybenzoate, which was quantitatively converted to gallic acid and acetate. Neither strain grew on other C₁ compounds or methyl group donors tested (methanol, formate, CO, betaine, or choline). Relatively low concentrations of CO were inhibitory to both strains; addition of 1% CO (vol/vol) to the headspace of actively growing cultures resulted in the immediate cessation of growth.

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FIG. 1. H2 stimulation of chemoorganotrophic growth of T. primitia strain ZAS-2. Either 10 mM maltose (•), 16 mM H2 (), 10 mM maltose plus 16 mM H2 (), or no substrate () was added to 2YACo medium under N2-CO2 (80:20 [vol/vol]).

It was noticed previously that the growth of ZAS-1 and ZAS-2 on H₂ plus CO₂ in the medium used for enrichment (JM4 medium [30]) was markedly improved if the nutrient broth and bovine rumen fluid present in JM4 were replaced by laboratory-prepared yeast autolysate (2 or 4% [vol/vol]) and a mixture of 11 cofactors, resulting in 2YACo or 4YACo medium, respectively (31). However, little or no growth was obtained if either yeast autolysate or the cofactor mixture was omitted from the medium. In order to determine which cofactors were required, ZAS-1 was grown through three successive transfers in 2YACo medium lacking one of the 11 cofactors. Cultures deficient in folinate (formyl-THF) displayed successively decreasing growth yields with each transfer. Further testing demonstrated that calcium folinate alone (final concentration, 500 ng/ml; equivalent to 0.98 µM) could replace the 11-cofactor mixture for both strains. Folate (the unsubstituted, unreduced form of the cofactor, at the same final concentration) could replace folinate for strain ZAS-2 but not for ZAS-1. If yeast autolysate contains folate compounds, they are obviously present at a concentration insufficient to support detectable growth of ZAS-1 and ZAS-2 in 2YACo medium. However, yeast autolysate contains at least one other required factor not present in most commercial yeast extracts or autolysates or in the 11-cofactor or 7-vitamin mixture included in 2YACo medium (described above).

Mixotrophic growth.
Strain ZAS-2 was capable of mixotrophic growth, i.e., the simultaneous utilization of organic substrates and H₂ for energy. Growing in the presence of both maltose and H₂ plus CO₂, strain ZAS-2 displayed a significantly shorter doubling time (29.2 ± 1.2 h) than when growing on maltose (65.9 ± 2.4 h) or H₂ plus CO₂ (50.3 ± 2.1 h) alone. Moreover, maltose and H₂ were consumed simultaneously during growth (Fig. 2), and cell yields and acetate production from maltose plus H₂ plus CO₂ were close to the sum of cell yields and acetate production when strain ZAS-2 was grown on either substrate alone (Table 2). Strain ZAS-2 also appeared to be capable of mixotrophy with H₂ plus CO₂ plus other organic substrates: doubling times during growth on H₂ plus CO₂ plus trimethoxybenzoate (38.1 ± 2.5 h) or xylose (35.2 ± 1.5 h) were significantly shorter than during unitrophic growth on either organic substrate alone (Table 1).

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FIG. 2. Mixotrophic growth of T. primitia strain ZAS-2 on H2 (plus CO2) and maltose.

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TABLE 2. Cell yields and acetate production by T. primitia strain ZAS-2 during unitrophic and mixotrophic growth

Hydrogen thresholds.
H₂ thresholds of mid-log-phase cultures growing on H₂ plus CO₂ were 490 ± 40 ppmv for ZAS-1 and 650 ± 50 ppmv for ZAS-2 (means ± standard deviations [SD]; n = 3). The H₂ threshold of ZAS-2 decreased moderately, to 350 ± 30 ppmv, when ZAS-2 was grown mixotrophically on H₂ plus CO₂ plus trimethoxybenzoate (Fig. 3).

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FIG. 3. Determination of H2 thresholds for two (of three) replicate cultures of T. primitia strain ZAS-2 growing mixotrophically on H2 plus CO2 plus trimethoxybenzoate (• and ). H2 additions are indicated by arrows. The H2 threshold is indicated by a dashed line. Symbols stand for two of three replicate cultures.

H₂/CO₂-acetogenic enzyme activities.
Previous work in our laboratory demonstrated that ZAS-1 and ZAS-2 possessed hydrogenase, formate dehydrogenase, and acetyl-CoA synthase (measured as CO dehydrogenase); the latter is a key enzyme of CO₂-reductive acetogenesis via the Wood/Ljungdahl pathway (31). To further confirm the presence of this pathway in T. primitia, enzymes associated with the conversion of formate to the methyl group of acetate were assayed. Cell extracts of ZAS-1 and ZAS-2 displayed activities for formyl-THF synthetase, methenyl-THF cyclohydrolase, methylene-THF dehydrogenase, and methylene-THF reductase, the four characteristic enzymes of the methyl group-forming branch of the Wood/Ljungdahl pathway (Table 3). The formyl-THF synthetase activities of both strains were dependent on the presence of ATP, formate, and THF. The activity of methylene-THF-dehydrogenase was NADP dependent in both strains; no activity was observed with NAD. In all cases, enzyme activity was abolished in cell extracts that had been boiled for 10 min.

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TABLE 3. Enzyme activities relevant to H2/CO2 acetogenesis in T. primitia

Oxygen tolerance and oxidative stress enzymes.
Mid-log-phase cultures of strains ZAS-1 and ZAS-2 were able to maintain growth after the addition of 0.5% O₂ to the headspace atmosphere, albeit with a slight and transient decrease in the growth rate (Fig. 4). Addition of 1% O₂, however, led to the rapid cessation of growth. The tolerance of cells to limited O₂ exposure prompted assays for enzymes associated with oxidative stress protection. Cell extracts of both strains showed low levels of NADH and NADPH peroxidases, whereas neither exhibited catalase or SOD activity. Both strains had low levels of NADPH oxidase, but relatively high levels of NADH oxidase activity were seen only in ZAS-2 (Table 4). Exposure of cultures to 0.5% O₂ 12 h prior to enzyme assays did not significantly alter the levels of any of the enzyme activities tested in either of the strains, and for all enzymes tested, activity was abolished in boiled cell extracts.

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FIG. 4. Effect of O2 on the growth of T. primitia strain ZAS-2. Cultures were grown under H2-CO2 (80:20 [vol/vol]), and O2 was introduced at 125 h (arrow) to yield a concentration (expressed as a percentage of headspace volume) of 0% (control) (), 0.5% (), 1% (), 2.5% (), or 5% ().

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TABLE 4. Oxygen- and peroxide-detoxifying enzyme activities in T. primitia

Top Abstract Introduction Materials and Methods Results Discussion References

 
T. primitia strains ZAS-1 and ZAS-2 possess a variety of properties typical of traditional "homoacetogens," including enzymes of the acetyl-CoA (Wood/Ljungdahl) pathway for synthesis of acetyl-CoA and acetate from H₂ plus CO₂ for energy generation and CO₂ fixation (15). Operation of this pathway, first suggested by the presence of acetyl-CoA synthase/CO dehydrogenase in cells (31), was further supported here by demonstration of the corresponding enzymes catalyzing conversion of CO₂ to the methyl group of acetate (Table 3). To the extent that enzyme activities measured with crude extracts and permeabilized cells can be correlated to the rate of acetate formation by growing cells, we compared the activity of the acetate-forming enzyme system, dA_E/xdt, to the rate of acetate production by cells during growth on H₂ plus CO₂, dA_G/xdt, where dA stands for micromoles of acetate produced by x milligrams of cell protein per unit of time (dt, in minutes), as inferred from enzyme activity (E) or growth measurements (G). This comparison was done for strain ZAS-2, which showed no background growth in 2YACo basal medium in the absence of H₂ (31). Inasmuch as optimum conditions for each enzyme activity reported in Table 3 have not yet been established, for dA_E/xdt we used the mean specific activity of all ZAS-2 enzymes, after dividing hydrogenase activity by 4. This gave 0.33 ± 0.25 µmol of acetate · min^-1 · mg of protein^-1 (mean ± SD). We calculated dA_G/xdt as being equal to Y_A/G · k, where Y_A/G, the acetate yield from growing cells, is calculated as 1/(4 · Y_hydrogen), expressed as micromoles of acetate · mg of protein^-1, and k, the specific growth rate constant, is calculated as 0.693/doubling time in minutes (39) (Table 1). For simplicity, we ignored the small fraction of H₂-derived electrons used for biomass synthesis, which probably accounted for an electron recovery, in acetate produced, of only 96% (Table 1, footnote c). This is reflected by using a rounded-off value of 4 in the term 4 · Y_hydrogen. The calculated value of dA_G/xdt is then 0.55 µmol · min^-1 · mg of protein^-1. Thus, even by a rough approximation, the mean activity of measured enzymes could account for as much as 60% of acetate production by growing cells. Hence, it seems reasonable to assume that the acetyl-CoA pathway is probably the major, if not the sole, route of acetate production from CO₂ in these spirochetes.

Strains ZAS-1 and ZAS-2 are also similar to other homoacetogens in being nutritionally versatile (16) and capable of homoacetogenic growth on a variety of organic substrates likely to be present in the termite gut, including hexoses, pentoses, disaccharides, and (in the case of ZAS-2) the methyl group of methoxylated aromatic compounds. Although the high-molecular-weight core lignin of lignocellulose does not appear to be degraded significantly during passage through the guts of wood-feeding termites (7, 11, 19, 25), there is evidence for the demethylation of aromatic lignin moieties (19). As such, it seems likely that T. primitia contributes to termite nutrition via acetogenesis from fermentation and demethylation of organic substrates in situ, as well as from H₂ and CO₂ produced by other members of the gut microbiota.

Strain ZAS-2 (and apparently also ZAS-1 [31]) was capable of mixotrophy, i.e., simultaneous use of H₂ plus CO₂ and organic substrates as energy sources (Fig. 2; Table 2). This capability has been demonstrated in other acetogens, including those isolated from termite guts (5). If H₂ concentrations in the hindgut lumen of Z. angusticollis are as high as those of other xylophagous termites (ca. 5 kPa or 50,000 ppmv in Reticulitermes flavipes) (17), mixotrophy may be a common mode of growth for T. primitia in situ. Indeed, some spirochetes may have become adapted to the constant presence of H₂ in termite hindguts such that they now require it for optimal growth. This was suggested by the stimulatory effect of H₂ on the utilization of organic substrates by strain ZAS-2 (Fig. 1), which may indicate a requirement for H₂ as a low-potential electron donor for some step(s) in biosynthesis. Although ZAS-2 possesses a hydrogenase (Table 3) and presumably forms some H₂ during the fermentation of organic substrates, at low cell densities such free H₂ may diffuse away from cells rapidly and thereby limit the rate of growth. Only as cell density (and hence H₂-forming capacity per milliliter) increases and H₂ accumulates would the specific growth rate increase incrementally. This interpretation is supported by the broadly concave shape of the growth curve of ZAS-2 on maltose in the absence of added H₂ (Fig. 1).

The mean H₂ thresholds of ZAS-1 and ZAS-2 (490 and 650 ppmv, respectively) were also well within the range reported for other H₂-utilizing acetogens (12). For ZAS-2, the mean H₂ threshold was somewhat lower during mixotrophic growth on H₂ plus trimethoxybenzoate (350 ppmv), but it was not lowered to a level that would make ZAS-2 competitive with H₂-utilizing methanogens under H₂-limited growth conditions. In any case, direct competition with methanogens for H₂ may be largely irrelevant: if high luminal H₂ concentrations exist in Z. angusticollis, spirochetes may never encounter seriously H₂ limited conditions in vivo.

In contrast to those of other homoacetogens, the substrate-specific biomass yields (Y_substrate) of T. primitia ZAS-1 and ZAS-2 were surprisingly low: 0.1 and 0.2, respectively, during growth on H₂ (plus CO₂) and 7.9 and 6.3, respectively, during growth on glucose (Table 1). These biomass yields are substantially lower than those of other acetogens, which range from 0.5 to 4.2 with H₂ and from 32.7 to 53.0 with hexoses (13, 18, 45, 52). However, this seemingly inefficient coupling of acetogenesis to growth may contribute to the effectiveness of T. primitia as a hindgut symbiont. Acetate produced by gut microbes is the primary energy source for termites (6), and low Y_substrate values mean that ZAS-1 and ZAS-2 will carry out homoacetogenesis without a concomitantly large increase in biomass and nutrient demand that might be detrimental to the host.

Previous studies have shown that the peripheral region of the termite hindgut is hypoxic (9, 17). O₂ concentrations near the gut epithelium are 50 to 100 µM and decrease steeply to anoxia within about 200 µm of the gut wall. It seems reasonable to assume that spirochetes inhabiting the hindgut lumen are likely to encounter this zone periodically. Although ZAS-1 and ZAS-2 are "anaerobes" in the traditional sense (i.e., they are incapable of growth in air), their ability to tolerate low levels of oxygen exposure could be of adaptive value in their natural habitat. Both ZAS-1 and ZAS-2 could maintain growth in the presence of 0.5% (vol/vol) O₂ in the gas phase (equivalent to a dissolved O₂ concentration of 6 µM). This finding suggests that the spirochetes could survive transient exposure to substantially higher concentrations of O₂ (a situation that is perhaps more analogous to that which they would encounter in vivo). Both strains possessed NADH and NADPH peroxidase activities that could afford protection against H₂O₂. ZAS-2 also possessed the O₂-consuming enzyme NADH oxidase. The activities of peroxidase and oxidase were similar to those of Brachyspira hyodysenteriae, an aerotolerant anaerobic spirochete that colonizes the gastrointestinal tracts of pigs (49). Studies have shown that free-living acetogens display various degrees of aerotolerance and can grow under atmospheres ranging from 0.3% oxygen (Acetobacterium woodii) to 21% oxygen (Sporomusa strain ST-1) (28; K. Küsel, A. Gößner, C. R. Lovell, and H. L. Drake, Abstr. 103rd Gen. Meet. Am. Soc. Microbiol. 2003, abstr. Q-375, p. 582, 2003). A recent study revealed a similar degree of aerotolerance, as well as active reduction of O₂ by using H₂ or organic compounds as reductants, in a variety of nonspirochetal acetogens isolated from termite hindguts (3). Like the acetogens examined in those studies, the T. primitia strains lack SOD activity. The ability to tolerate low levels of O₂ has also been observed in the termite hindgut-associated methanogens Methanobrevibacter cuticularis and Methanobrevibacter curvatus (30), suggesting that aerotolerance may be a common trait among anaerobes in the termite hindgut.

Studies of the nutritional requirements of T. primitia revealed a strict requirement for folate compounds. This was surprising given that THF plays a critical role as a one-carbon carrier in the methyl-group-forming branch of the Wood/Ljungdahl pathway (35). Most bacteria are capable of de novo folate biosynthesis, and numerous homoacetogens grown in defined media have no apparent folate requirements (22, 32, 44). By contrast, many host-associated bacteria require an exogenous source of folate or a folate derivative, including ruminal (Treponema bryantii [48]) and genital (Treponema phagedenis [50]) spirochetes and numerous nonspirochetal members of the bovine rumen microbial community (24, 47). The source of folate compounds for T. primitia in vivo is currently unknown. Although provision of folates by the termite host is a possibility (e.g., folate synthesis has been demonstrated in the mosquito, Aedes aegypti [27, 53, 54]), most insects require a dietary source of preformed folate. A more likely source of folate compounds is other members of the complex hindgut community. A variety of bacteria have been shown to secrete folate compounds in vitro (14, 26) as well as in gastrointestinal tracts (10, 29, 43), and we have recently isolated several strains of bacteria from the hindguts of Z. angusticollis whose extracellular secretions can support the growth of ZAS-1 and ZAS-2 in folate-free medium (J. R. Graber and J. A. Breznak, presented at the 9th International Symposium on Microbial Ecology, Amsterdam, The Netherlands, 26 to 31 August 2001). The nature of these organisms and their putative folate secretions are the subjects of ongoing study in our laboratory.

In summary, T. primitia strains ZAS-1 and ZAS-2 possess an assortment of nutritional and physiological properties that would appear to make them well adapted to life as termite hindgut symbionts. In light of the phylogenetic diversity of termite gut spirochetes revealed by cultivation-independent molecular analyses (34), it seems certain that the few strains currently in culture (31, 33) offer only an introductory glimpseinto the physiological diversity of termite gut spirochetes and their importance to termite nutrition.

 
This research was funded by grants from the National Science Foundation (IBN97-09000 and IBN01-14505).

HPLC analysis of aromatic compounds was performed with equipment provided by the laboratory of James Tiedje, with the aid of Benjamin Griffin. We are grateful to Tom Schmidt and members of his laboratory for helpful discussions and criticisms.

 
* Corresponding author. Mailing address: Department of Microbiology and Molecular Genetics, 6190 Biomedical and Physical Sciences Building, Michigan State University, East Lansing, MI 48824-4320. Phone: (517) 355-6463, ext. 1607. Fax: (517) 353-8957. E-mail: breznak{at}msu.edu.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Ashwell, G. 1963. Colorimetric analysis of sugars. Methods Enzymol. 3:73-105.
2
2. Beers, R. F., and I. W. Sizer. 1952. Spectrophotometric method for measuring the breakdown of hydrogen peroxide by catalase. J. Biol. Chem. 195:133-140.[Free Full Text]
3
3. Boga, H. I., and A. Brune. 2003. Hydrogen-dependent oxygen reduction by homoacetogenic bacteria isolated from termite guts. Appl. Environ. Microbiol. 69:779-786.[Abstract/Free Full Text]
4
4. Breznak, J. A., and J. M. Switzer. 1986. Acetate synthesis from H₂ plus CO₂ by termite gut microbes. Appl. Environ. Microbiol. 52:623-630.[Abstract/Free Full Text]
5
5. Breznak, J. A., and J. S. Blum. 1991. Mixotrophy in the termite gut acetogen, Sporomusa termitida. Arch. Microbiol. 156:105-110.[CrossRef]
6
6. Breznak, J. A. 1994. Acetogenesis from carbon dioxide in termite guts, p. 303-330. In H. L. Drake (ed.), Acetogenesis. Chapman & Hall, New York, N.Y.
7
7. Breznak, J. A., and A. Brune. 1994. Role of microorganisms in the digestion of lignocellulose by termites. Annu. Rev. Entomol. 39:453-487.[CrossRef]
8
8. Breznak, J. A., and J. R. Leadbetter. 2002. Termite gut spirochetes. In M. Dworkin (ed.), The Prokaryotes: an evolving electronic resource for the microbiological community. [Online.] Springer-Verlag, New York, N.Y.
9
9. Brune, A., D. Emerson, and J. A. Breznak. 1995. The termite gut microflora as an oxygen sink: microelectrode determination of oxygen and pH gradients in guts of lower and higher termites. Appl. Environ. Microbiol. 61:2681-2687.[Abstract]
10
10. Camilo, E., J. Zimmerman, J. B. Mason, B. Golner, R. Russell, J. Selhub, and I. H. Rosenberg. 1996. Folate synthesized by bacteria in the human upper small intestine is assimilated by the host. Gastroenterology 110:991-998.[CrossRef][Medline]
11
11. Cookson, L. J. 1987. [¹⁴C]lignin degradation by three Australian termite species. Wood Sci. Technol. 21:11-25.
12
12. Cord-Ruwisch, R., H. Seitz, and R. Conrad. 1988. The capacity of hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen depends on the redox potential of the terminal electron acceptor. Arch. Microbiol. 149:350-357.[CrossRef]
13
13. Daniel, S. L., T. Hsu, S. I. Dean, and H. L. Drake. 1990. Characterization of the H₂- and CO-dependent chemolithotrophic potentials of the acetogens Clostridium thermoaceticum and Acetogenium kivui. J. Bacteriol. 172:4464-4471.[Abstract/Free Full Text]
14
14. Deguchi, Y., T. Morishita, and M. Mutai. 1985. Comparative studies on synthesis of water-soluble vitamins among human species of Bifidobacteria. Agric. Biol. Chem. 49:13-19.
15
15. Drake, H. L. 1992. Acetogenesis and acetogenic bacteria, p. 1-15. In J. Lederberg (ed.), Encyclopedia of microbiology, vol. 1. Academic Press, New York, N.Y.
16
16. Drake, H. L., S. L. Daniel, K. Küsel, C. Matthies, C. Kuhner, and S. Braus-Stromeyer. 1997. Acetogenic bacteria: what are the in situ consequences of their diverse metabolic versatilities? Biofactors 6:13-24.[Medline]
17
17. Ebert, A., and A. Brune. 1997. Hydrogen concentration profiles at the oxic-anoxic interface: a microsensor study of the hindgut of the wood-feeding lower termite Reticulitermes flavipes. Appl. Environ. Microbiol. 63:4039-4046.[Abstract]
18
18. Eichler, B., and B. Schink. 1984. Oxidation of primary aliphatic alcohols by Acetobacterium carbinolicum sp. nov., a homoacetogenic anaerobe. Arch. Microbiol. 140:147-152.[CrossRef]
19
19. Esenther, G. R., and T. K. Kirk. 1974. Catabolism of aspen sapwood in Reticulitermes flavipes. Ann. Entomol. Soc. Am. 67:989-991.
20
20. Eutick, M. L., P. Veivers, R. W. O'Brien, and M. Slaytor. 1978. Dependence of the higher termite, Nasutitermes exitiosus and the lower termite, Coptotermes lacteus on their gut flora. J. Insect Physiol. 24:363-368.[CrossRef]
21
21. Flohe, L., R. Becker, R. Brigelius, E. Lengfelder, and F. Otting. 1988. Convenient assays for superoxide dismutase, p. 287-293. In J. Miquel (ed.), CRC handbook of free radicals and antioxidants in biomedicine, vol. 3. CRC Press, Boca Raton, Fla.
22
22. Genthner, B. R. S., C. L. Davis, and M. P. Bryant. 1981. Features of rumen and sewage sludge strains of Eubacterium limosum, a methanol-utilizing and H₂-CO₂-utilizing species. Appl. Environ. Microbiol. 42:12-19.[Abstract/Free Full Text]
23
23. Graber, J. R., J. R. Leadbetter, and J. A. Breznak. 2004. Description of Treponema azotonutricium sp. nov. and Treponema primitia sp. nov., the first spirochetes isolated from termite guts. Appl. Environ. Microbiol. 70:1315-1320.[Abstract/Free Full Text]
24
24. Herbeck, J. L., and M. P. Bryant. 1974. Nutritional requirements of the intestinal anaerobe Ruminococcus bromii. Appl. Microbiol. 28:1018-1022.[Medline]
25
25. Hyodo, F., J. Azuma, and T. Abe. 1999. Estimation of the effect of passage through the gut of a lower termite, Coptotermes formosanus Shiraki, on lignin by solid-state CP/MAS ¹³C NMR. Holzforschung 53:244-246.
26
26. Iwai, K., M. Kobashi, and H. Fujisawa. 1970. Occurrence of Crithidia factors and folic acid in various bacteria. J. Bacteriol. 104:197-201.[Abstract/Free Full Text]
27
27. Jaffe, J. J., J. J. McCormack, E. Meymarian, and H. M. Doremus. 1977. Comparative activity and properties of lactate dehydrogenase, xanthine dehydrogenase, and dihydrofolate reductase in normal and Brugia pahangi infected Aedes aegypti. J. Parasitol. 63:547-553.[CrossRef][Medline]
28
28. Karnholz, A., K. Küsel, A. Gößner, A. Schramm, and H. L. Drake. 2002. Tolerance and metabolic response of acetogenic bacteria toward oxygen. Appl. Environ. Microbiol. 68:1005-1009.[Abstract/Free Full Text]
29
29. Krause, L. J., C. W. Forsberg, and D. L. O'Connor. 1996. Feeding human milk to rats increases Bifidobacterium in the cecum and colon which correlates with enhanced folate status. J. Nutr. 126:1505-1511.
30
30. Leadbetter, J. R., and J. A. Breznak. 1996. Physiological ecology of Methanobrevibacter cuticularis sp. nov. and Methanobrevibacter curvatus sp. nov., isolated from the hindgut of the termite Reticulitermes flavipes. Appl. Environ. Microbiol. 62:3620-3631.[Abstract]
31
31. Leadbetter, J. R., T. M. Schmidt, J. R. Graber, and J. A. Breznak. 1999. Acetogenesis from H₂ plus CO₂ by spirochetes from termite guts. Science 283:686-689.[Abstract/Free Full Text]
32
32. Leigh, J. A., F. Mayer, and R. S. Wolfe. 1981. Acetogenium kivui, a new thermophilic hydrogen-oxidizing, acetogenic bacterium. Arch. Microbiol. 129:275-280.[CrossRef]
33
33. Lilburn, T. G., K. S. Kim, N. E. Ostrom, K. R. Byzek, J. R. Leadbetter, and J. A. Breznak. 2001. Nitrogen fixation by symbiotic and free-living spirochetes. Science 292:2495-2498.[Abstract/Free Full Text]
34
34. Lilburn, T. G., T. M. Schmidt, and J. A. Breznak. 1999. Phylogenetic diversity of termite gut spirochaetes. Environ. Microbiol. 1:331-345.[CrossRef][Medline]
35
35. Ljungdahl, L. G. 1984. Other functions of folates, p. 555-579. In R. L. Blakley and S. J. Benkovic (ed.), Folates and pterins, vol. 1. John Wiley and Sons, New York, N.Y.
36
36. Lovley, D. R. 1985. Minimum threshold for hydrogen metabolism in methanogenic bacteria. Appl. Environ. Microbiol. 49:1530-1531.[Abstract/Free Full Text]
37
37. Lowry, O. H., N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275.[Free Full Text]
38
38. Meßmer, M., G. Wohlfarth, and G. Diekert. 1993. Methyl chloride metabolism of the strictly anaerobic, methyl chloride-utilizing homoacetogen strain MC. Arch. Microbiol. 160:383-387.
39
39. Neidhardt, F. C., J. L. Ingraham, and M. Schaechter. 1990. Physiology of the bacterial cell. Sinauer Associates, Sunderland, Mass.
40
40. O'Brien, W. E., and L. G. Ljungdahl. 1972. Fermentation of fructose and synthesis of acetate from carbon dioxide by Clostridium formicoaceticum. J. Bacteriol. 109:626-632.[Abstract/Free Full Text]
41
41. Odelson, D. A., and J. A. Breznak. 1983. Volatile fatty acid production by the hindgut microbiota of xylophagous termites. Appl. Environ. Microbiol. 45:1602-1613.[Abstract/Free Full Text]
42
42. Paster, B. J., F. E. Dewhirst, S. M. Cooke, V. Fussing, L. K. Poulsen, and J. A. Breznak. 1996. Phylogeny of not-yet-cultured spirochetes from termite guts. Appl. Environ. Microbiol. 62:347-352.[Abstract]
43
43. Rong, N., J. Selhub, B. R. Goldin, and I. H. Rosenberg. 1991. Bacterially synthesized folate in rat large intestine is incorporated into host tissue folyl polyglutamates. J. Nutr. 121:1955-1959.
44
44. Schink, B. 1984. Clostridium magnum sp. nov., a non-autotrophic homoacetogenic bacterium. Arch. Microbiol. 137:250-255.[CrossRef]
45
45. Sharak-Genthner, B. R., and M. P. Bryant. 1987. Additional characteristics of one-carbon-compound utilization by Eubacterium limosum and Acetobacterium woodii. Appl. Environ. Microbiol. 53:471-476.[Abstract/Free Full Text]
46
46. Slaytor, M., and D. J. Chappell. 1994. Nitrogen metabolism in termites. Comp. Biochem. Physiol. 107:1-10.[CrossRef]
47
47. Slyter, L. L., and J. M. Weaver. 1977. Tetrahydrofolate and other growth requirements of certain strains of Ruminococcus flavefaciens. Appl. Environ. Microbiol. 33:363-369.[Abstract/Free Full Text]
48
48. Stanton, T. B., and E. Canale-Parola. 1980. Treponema bryantii sp. nov., a rumen spirochete that interacts with cellulolytic bacteria. Arch. Microbiol. 127:145-156.[CrossRef][Medline]
49
49. Stanton, T. B. 1989. Glucose metabolism and NADH recycling by Treponema hyodysenteriae, the agent of swine dysentery. Appl. Environ. Microbiol. 55:2365-2371.[Abstract/Free Full Text]
50
50. Steinman, H. G., V. I. Oyama, and H. O. Schulze. 1954. Carbon dioxide, cocarboxylase, citrovorum factor, and coenzyme A as essential growth factors for a saccharolytic treponeme (S-69). J. Biol. Chem. 211:327-335.[Free Full Text]
51
51. Tholen, A., and A. Brune. 2000. Impact of oxygen on metabolic fluxes and in situ rates of reductive acetogenesis in the hindgut of the wood-feeding termite Reticulitermes flavipes. Environ. Microbiol. 2:436-449.[CrossRef][Medline]
52
52. Tschech, A., and N. Pfennig. 1984. Growth yield increase linked to caffeate reduction in Acetobacterium woodii. Arch. Microbiol. 137:163-167.[CrossRef]
53
53. Venters, D. 1971. The mode of systemic insecticidal action of sulphaquinoxaline against mosquitoes. Trans. R. Soc. Trop. Med. Hyg. 65:24-25.[Medline]
54
54. Venters, D. 1972. Folate metabolism in the muscles of Aedes aegypti. Insect Biochem. 2:153-160.
55
55. Widdel, F., and N. Pfennig. 1981. Studies on dissimilatory sulfate-reducing bacteria that decompose fatty acids. I. Isolation of new sulfate-reducing bacteria enriched with acetate from saline environments. Description of Desulfobacter postgatei gen. nov., sp. nov. Arch. Microbiol. 129:395-400.[CrossRef][Medline]

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Applied and Environmental Microbiology, March 2004, p. 1307-1314, Vol. 70, No. 3
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