Evaluation of a Stochastic Inactivation Model for Heat-Activated Spores of Bacillus spp.

Heat activates the dormant spores of certain Bacillus spp., which is reflected in the “activation shoulder” in their survival curves. At the same time, heat also inactivates the already active and just activated spores, as well as those still dormant. A stochastic model based on progressively changing probabilities of activation and inactivation can describe this phenomenon. The model is presented in a fully probabilistic discrete form for individual and small groups of spores and as a semicontinuous deterministic model for large spore populations. The same underlying algorithm applies to both isothermal and dynamic heat treatments. Its construction does not require the assumption of the activation and inactivation kinetics or knowledge of their biophysical and biochemical mechanisms. A simplified version of the semicontinuous model was used to simulate survival curves with the activation shoulder that are reminiscent of experimental curves reported in the literature. The model is not intended to replace current models to predict dynamic inactivation but only to offer a conceptual alternative to their interpretation. Nevertheless, by linking the survival curve''s shape to probabilities of events at the individual spore level, the model explains, and can be used to simulate, the irregular activation and survival patterns of individual and small groups of spores, which might be involved in food poisoning and spoilage.Heat inactivation kinetics of bacterial spores is a well-researched field. Much of the work on its relation to foods has focused on the heat-resistant spores of Clostridia, particularly those of Clostridium botulinum, which to this date serves as the reference organism in sterility calculations of low-acid foods (8, 32). The thermal resistance of Bacilli spores, although also extensively studied, has received less attention in the literature on food preservation. This is primarily because they are unlikely to germinate and produce cells that will survive and divide under the anaerobic conditions in a sterilized food container. Yet the mere possibility of viable Bacillus spores being present in processed foods has become an issue of food safety and a security concern. For this reason, there is a renewed interest in these spores'' heat resistance (2, 3, 6, 7, 16, 30). One of the peculiarities of certain Bacillus spores, like those of Bacillus sporothermodurans or Bacillus stearothermophilus, is that many of them can remain dormant unless activated by heat. The result is a survival curve that exhibits an “activation shoulder,” as shown schematically in Fig. ​Fig.11 and with published data in Fig. ​Fig.2.2. Thus, modeling this survival pattern, where the number of spores initially grows rather than declines, must account for the heat''s dual role of being a lethal agent and activator at the same time.Open in a separate windowFIG. 1.A schematic view of a survival curve having an activation shoulder. S(t) is the ratio between the number N(t) of viable spores at time t and the initial number N₀. Notice the discrepancy between the two ways to estimate the number of dormant spores, represented by the dashed and dotted gray lines.Open in a separate windowFIG. 2.Demonstration of the fit of equation 1 (solid line) and equation 2 (dashed line) to survival curves of B. stearothermophilus spores at two temperatures. Notice the postpeak concavity of the curves. In such cases, the estimated number of dormant spores reached by the tangent method will depend on the experiment duration. The original experimental data are from Sapru et al. (25).Traditionally, the thermal inactivation of both Clostridia and Bacilli spores has been thought to follow first-order kinetics (9, 12, 31), an assumption that has been frequently challenged in recent years (18, 21, 33, 35). The most publicized model of the simultaneous heat activation and inactivation of Bacillus spores in food is that proposed by Sapru et al. (24, 25), which is an improved version of models proposed earlier by Shull et al. (29) and Rodriguez et al. (23). All of these authors and others (1, 17) assumed that the activation of dormant spores follows first-order kinetics and so does their inactivation before and after activation. The temperature dependence of the corresponding exponential rate constants was assumed to follow the Arrhenius equation.Peleg (18, 20) and van Boekel (33, 35) have shown that none of the above assumptions was necessary and that the same survival data on Bacillus stearothermophilus reported by Sapru et al. (25) and other investigators (5) can be described by different kinds of alternative four-parameter empirical models, which have a slightly better fit. This was evident not only visually (Fig. ​(Fig.2)2) but also as judged by statistical criteria (34). Fig. ​Fig.22 shows the fit of the “double Weibullian” model proposed by van Boekel (33). It has the following form: (1) where S(t) = N(t)/N₀ is the survival/activation ratio, N₀ and N(t) are the initial and momentary number of countable spores, respectively, and b₁, b₂, n₁, and n₂ are adjustable temperature-dependent constants. Figure 2 also shows the fit of an ad hoc empirical model, a hybrid between the double Weibullian model and one previously proposed (20) that can be written in the following form: (2) or (3) where a₁, b₁, t_c2, and m₂ are adjustable temperature-dependent parameters. According to this model, a₁ is the asymptote of the first term on the right, b₁ is a time characteristic of the activation, t_c2 is a characteristic time of the inactivation, and m₂ is a parameter that represents the curve''s postpeak concavity. The structure of equation 2 or 3 dictates that the number of dormant spores must be finite and cannot exceed N₀ × 10^a1, if the logarithm is base 10, or N₀ × exp(a₁), if it is base e. (A demonstration that generates realistic-looking activation/inactivation curves using equation 3 as a model is available from Wolfram Research [http://demonstrations.wolfram.com/SurvivalCurvesOfBacilliSporesWithAnActivationShoulder/].)Corradini and Peleg (5) proposed a way to estimate the initial number of dormant spores from survival curves having an activation shoulder using a similar model, which was originally described in Peleg (20). They suggested that the intersection of a tangent to the survival curve drawn at its postpeak region with the time axis (Fig. ​(Fig.1)1) is not a recommended method to estimate the number of dormant spores and that it can render unrealistically high values if used. Also, where there is no evidence that the survival curve in the postpeak region ever becomes a straight line; the same survival curve will yield different estimates of the dormant spores'' initial number depending on the experiment''s duration. Moreover, if in the postpeak region the survival ratio drop rate progressively increases, as it most probably does (Fig. ​(Fig.2)2) (20, 33), then the number of dormant spores estimated by the tangent extrapolation method will grow indefinitely, despite the fact that it must be finite (1). Also, since the exponential inactivation rate can be a function of time as well as of temperature, the applicability of the Arrhenius equation as a secondary model might come into question. The same can also be said about the log-linearity of the D value''s temperature dependence if used instead of the Arrhenius equation.The question that arises in light of all the above is whether one can construct a conceptual population dynamic model of the activation/inactivation of spores without assuming any fixed kinetic order. The biochemical and biophysical mechanisms that govern bacterial spore germination, activation, and inactivation have been thoroughly investigated (11, 13, 14, 15, 22, 26-28). Still, it is not clear how processes within an individual spore can be translated into activation and survival patterns at the population level and how their manifestation can be expressed in a mathematical model. Whenever a system has inherent variability and knowledge of its working is incomplete or merely insufficient to develop a model from basic principles, one can, and sometimes must, resort to a probabilistic modeling approach. The general objective of this work has been to explore the merits and limitations of this option by developing a stochastic model of Bacilli spores'' heat activation and inactivation and examining its properties. The goal has not been to develop a new method to predict the spores'' survival under dynamic conditions—rate versions of the existing empirical models such as equation 1, 2, or 3 seem to be quite suitable for that—but to offer an alternative interpretation of the patterns reported and discussed in the literature.     

Chenodeoxycholate Is an Inhibitor of Clostridium difficile Spore Germination

Some cholate derivatives that are normal components of bile can act with glycine to induce the germination of Clostridium difficile spores, but at least one bile component, chenodeoxycholate, does not induce germination. Here we show that chenodeoxycholate inhibits the germination of C. difficile spores in response to cholate and taurocholate.The anaerobic human pathogen Clostridium difficile must be in the spore form to survive for extended periods of time outside the colonic environment (6). Spores are also the form of the organism most likely to be ingested by a host. To cause disease, however, C. difficile spores must germinate in the gastrointestinal tract and reach the anaerobic environment of the colon, where they can grow out as vegetative bacteria (2). The vegetative form produces two toxins that damage the colonic epithelium and lead to C. difficile-associated diseases, such as diarrhea, pseudomembranous colitis, and toxic megacolon (4, 15). Extending the work of Wilson and colleagues (17, 18), we have shown that certain bile salts and glycine act as cogerminants for C. difficile spores (13). Primary bile salts produced by the liver are composed mainly of cholate (CA) and chenodeoxycholate (CDCA) derivatives conjugated with either taurine or glycine (11). Since CA derivatives are found in the relatively aerobic proximal ileum (9), we reasoned that C. difficile might benefit if its germination were inhibited until the spores reached the anaerobic environment of the large intestine.Inhibitors of germination are typically structurally similar to the germinant whose activities they inhibit. For example, l-alanine-mediated germination of Bacillus subtilis spores is inhibited by d-alanine (16) and 6-thioguanosine inhibits inosine-mediated germination in Bacillus anthracis (1, 16). Since CA and CDCA are structurally similar but CA induces the germination of C. difficile spores (13) and CDCA does not, we tested whether CDCA could act as an inhibitor of germination. C. difficile strain CD196 (10) spores were produced and their concentration determined as described previously (13). After the vegetative bacteria were killed by incubation at 60°C for 20 min, spores were incubated in water containing various concentrations and combinations of bile salts for 10 min. Here we took advantage of the finding by Wilson et al. that C. difficile spores germinate very inefficiently on rich medium plates lacking bile salts (18) unless they are preincubated with bile salts (13, 17). After incubation, spores were serially diluted and plated on brain heart infusion agar supplemented with 5 g yeast extract per liter-0.1% l-cysteine (BHIS) (Difco) in the absence of any bile salt (BHIS contains enough glycine to act as a cogerminant). After overnight growth at 37°C, colonies were enumerated. As a positive or negative control, spores were plated on BHIS containing 0.1% taurocholate (TA) [BHIS(TA)] or on BHIS agar alone, respectively. Preincubation of spores with 0.1% TA in water resulted in the recovery of approximately 0.5% of the total number of spores as colonies compared to results for spores plated directly on BHIS(TA). These results are similar to our previous findings that spores germinate and grow out as colonies more efficiently on agar medium containing TA (13). As reported previously, 0.1% CDCA poorly stimulated colony formation by C. difficile spores (13), yielding only 0.006% spore recovery (Fig. ​(Fig.1A).1A). When TA and CDCA were combined, both at 0.1%, colony formation by C. difficile spores was reduced 21-fold to 0.024% compared to the effect of TA alone. This result indicates that CDCA blocks TA-stimulated colony formation and suggests that CDCA may be an inhibitor of C. difficile spore germination. Increasing the ratio of TA to CDCA suppressed the inhibitory effect of CDCA, increasing colony formation by spores (Fig. ​(Fig.1A).1A). Thus, CDCA seems to block colony formation by competing with TA.Open in a separate windowFIG. 1.CDCA inhibits colony formation by C. difficile spores in response to TA and CA. (A) Spores were prepared and preincubated with TA or CDCA or both in water for 10 min before serial dilution and plating on BHIS agar in the absence of TA. Spores plated on BHIS(TA) served as a positive control for 100% colony formation (CFU). Based on comparisons of total spore counts obtained by microscopy and by colony formation on BHIS(TA) plates, the efficiency of colony formation on BHIS(TA) was estimated at 83%. (B) Spores were prepared as described for panel A and exposed to CA or CDCA or both. Values shown are the averages for three independent experiments, and error bars represent one standard deviation from the mean.CA and other cholate derivatives (e.g., TA, glycocholate, and deoxycholate [DCA]) are also germinants for C. difficile spores (13, 17). To test if CDCA prevents colony formation induced by CA, spores were preexposed to 0.1% CA with and without CDCA. Exposure to CA alone resulted in approximately 1% spore recovery, whereas exposure to 0.1% CA and 0.1% CDCA together led to a decrease in colony formation to 0.075% (Fig. ​(Fig.1B).1B). The effect of CDCA on CA-mediated colony formation was relieved by increasing the concentration of CA to 1.0%, raising colony formation to 2.6% (Fig. ​(Fig.1B).1B). These results indicate that CDCA blocks colony formation induced by CA, as well as that induced by TA, and may be an inhibitor of germination by C. difficile spores that acts competitively in both cases.Spore germination per se is classically measured as a decrease in the optical density of a spore suspension occurring concomitantly with a release of Ca²⁺-dipicolinate from the spore core, rehydration of the core, and degradation of the cortex (8, 12). As determined by this assay, TA is the most effective bile salt for inducing rapid germination (13). To test if CDCA is an inhibitor of germination as opposed to an inhibitor of some other step between germination and colony formation, spores were purified as described previously (13). Spores did not germinate in BHIS medium alone or when this medium was supplemented with 0.1% CDCA (Fig. ​(Fig.2).2). When C. difficile spores were suspended in BHIS containing 0.1% TA, the optical density of the suspension rapidly decreased, indicating that the spores were germinating. However, the optical density of the spores suspended in BHIS with 0.1% TA plus 0.1% CDCA did not decrease over time, indicating that CDCA inhibited TA-mediated germination (Fig. ​(Fig.2).2). When the concentration of TA was increased from 0.1% to 1.0% in the presence of 0.1% CDCA, spores were able to germinate (Fig. ​(Fig.2).2). After overnight incubation in BHIS with 0.1% TA plus 0.1% CDCA, 84% of the spores remained phase bright, while only 11% of spores remained phase bright in BHIS with 1.0% TA plus 0.1% CDCA, indicating that CDCA blocks germination at a very early step. Thus, CDCA is an inhibitor of germination by C. difficile spores that functions by competing with TA and possibly with CA.Open in a separate windowFIG. 2.CDCA inhibits germination of Clostridium difficile spores. Spores were prepared as described previously (13). C. difficile spores were suspended in BHIS alone (•), BHIS plus 0.1% CDCA (▾), BHIS plus 0.1% TA (⧫), BHIS plus 0.1% TA-0.1% CDCA (▪), or BHIS plus 1.0% TA-0.1% CDCA (▴). The ratio of the OD₆₀₀ at the various time points to the OD₆₀₀ at time zero is plotted versus time. Data points are the averages of three independent experiments, and error bars represent one standard deviation from the mean.We previously suggested a role for bile salts in determining the ability of C. difficile to colonize and cause disease (13). In this model, germination of C. difficile spores depends on interaction with glycine and certain bile salts. We show here that the primary bile salt CDCA inhibits germination of C. difficile spores. As mentioned above, germination inhibitors are commonly structurally related to the germinant they inhibit. The structures of CA derivatives and CDCA derivatives are very similar; they differ only insofar as CDCA lacks the 12α hydroxyl group (11).CDCA and CA derivatives are present in approximately equal concentrations in the cecum (5). Under such conditions, CDCA would compete with CA derivatives for binding to putative germinant receptors on C. difficile spores. Mekhjian and colleagues measured the colonic absorption rates of CDCA, CA, and DCA that were introduced into the cecum and collected at the distal colon (7). They found that CDCA was absorbed by the colon at 10 times the rate for CA (7). Thus, when spores reach the distal large intestine, they encounter a decreased ratio of CDCA to CA. Such a change in ratio might allow CA derivatives to act as effective germinants. Thus, C. difficile spores would not be expected to germinate until they reach the colon, which also provides the anaerobic environment required for C. difficile growth.The colonic microflora, which is known to protect the host against C. difficile infection, plays a significant role in the metabolism of bile salts (3, 11). Many different species express on their cell surfaces bile salt hydrolases that serve to remove the conjugated tauryl or glycyl groups from primary bile salts (11). After deconjugation, CA and CDCA are further metabolized by a small percentage of the bacterial species in the cecum to the secondary bile salts deoxycholate and lithocholate, respectively (11, 14). Deoxycholate is an inhibitor of C. difficile growth (13, 17). CDCA inhibits both germination and growth (13). The use of CDCA either as prophylaxis or as a therapy for C. difficile-associated disease might be helpful for patients who are undergoing antibiotic regimens or who are colonized by this bacterium. For example, when an antibiotic that is known to be associated with an increased risk of inciting C. difficile-associated disease is administered, the coadministration of CDCA might protect that individual from colonization by C. difficile through inhibiting spore germination. Alternatively, administering CDCA to individuals who are already being given vancomycin or metronidazole for C. difficile-associated disease may have the benefit of preventing spore germination and further vegetative growth (13) after antibiotic therapy is stopped. This strategy may reduce the already significant risk of a relapse.     

Streptomyces Development in Colonies and Soils

Streptomyces development was analyzed under conditions resembling those in soil. The mycelial growth rate was much lower than that in standard laboratory cultures, and the life span of the previously named first compartmentalized mycelium was remarkably increased.Streptomycetes are gram-positive, mycelium-forming, soil bacteria that play an important role in mineralization processes in nature and are abundant producers of secondary metabolites. Since the discovery of the ability of these microorganisms to produce clinically useful antibiotics (2, 15), they have received tremendous scientific attention (12). Furthermore, its remarkably complex developmental features make Streptomyces an interesting subject to study. Our research group has extended our knowledge about the developmental cycle of streptomycetes, describing new aspects, such as the existence of young, fully compartmentalized mycelia (5-7). Laboratory culture conditions (dense inocula, rich culture media, and relatively elevated temperatures [28 to 30°C]) result in high growth rates and an orderly-death process affecting these mycelia (first death round), which is observed at early time points (5, 7).In this work, we analyzed Streptomyces development under conditions resembling those found in nature. Single colonies and soil cultures of Streptomyces antibioticus ATCC 11891 and Streptomyces coelicolor M145 were used for this analysis. For single-colony studies, suitable dilutions of spores of these species were prepared before inoculation of plates containing GYM medium (glucose, yeast extract, malt extract) (11) or GAE medium (glucose, asparagine, yeast extract) (10). Approximately 20 colonies per plate were obtained. Soil cultures were grown in petri dishes with autoclaved oak forest soil (11.5 g per plate). Plates were inoculated directly with 5 ml of a spore suspension (1.5 × 10⁷ viable spores ml⁻¹; two independent cultures for each species). Coverslips were inserted into the soil at an angle, and the plates were incubated at 30°C. To maintain a humid environment and facilitate spore germination, the cultures were irrigated with 3 ml of sterile liquid GAE medium each week.The development of S. coelicolor M145 single colonies growing on GYM medium is shown in Fig. ​Fig.1.1. Samples were collected and examined by confocal microscopy after different incubation times, as previously described (5, 6). After spore germination, a viable mycelium develops, forming clumps which progressively extend along the horizontal (Fig. 1a and b) and vertical (Fig. 1c and d) axes of a plate. This mycelium is fully compartmentalized and corresponds to the first compartmentalized hyphae previously described for confluent surface cultures (Fig. 1e, f, and j) (see below) (5); 36 h later, death occurs, affecting the compartmentalized hyphae (Fig. 1e and f) in the center of the colony (Fig. ​(Fig.1g)1g) and in the mycelial layers below the mycelial surface (Fig. 1d and k). This death causes the characteristic appearance of the variegated first mycelium, in which alternating live and dead segments are observed (Fig. 1f and j) (5). The live segments show a decrease in fluorescence, like the decrease in fluorescence that occurs in solid confluent cultures (Fig. ​(Fig.11 h and i) (5, 9). As the cycle proceeds, the intensity of the fluorescence in these segments returns, and the segments begin to enlarge asynchronously to form a new, multinucleated mycelium, consisting of islands or sectors on the colony surfaces (Fig. 1m to o). Finally, death of the deeper layers of the colony (Fig. ​(Fig.1q)1q) and sporulation (Fig. ​(Fig.1r)1r) take place. Interestingly, some of the spores formed germinate (Fig. ​(Fig.1s),1s), giving rise to a new round of mycelial growth, cell death, and sporulation. This process is repeated several times, and typical, morphologically heterogeneous Streptomyces colonies grow (not shown). The same process was observed for S. antibioticus ATCC 11891, with minor differences mainly in the developmental time (not shown).Open in a separate windowFIG. 1.Confocal laser scanning fluorescence microscopy analysis of the development-related cell death of S. coelicolor M145 in surface cultures containing single colonies. Developmental culture times (in hours) are indicated. The images in panels l and n were obtained in differential interference contrast mode and correspond to the same fields as in panels k and m, respectively. The others are culture sections stained with SYTO 9 and propidium iodide. Panels c, d, k, l, p, and q are cross sections; the other images are longitudinal sections (see the methods). Panels h and i are images of the same field taken with different laser intensities, showing low-fluorescence viable hyphae in the center of the colonies that develop into a multinucleated mycelium. The arrows in panels e and s indicate septa (e) and germinated spores (s). See the text for details.Figure ​Figure22 shows the different types of mycelia present in S. coelicolor cultures under the conditions described above, depending on the compartmentalization status. Hyphae were treated with different fluorescent stains (SYTO 9 plus propidium iodide for nucleic acids, CellMask plus FM4-64 for cell membranes, and wheat germ agglutinin [WGA] for cell walls). Samples were processed as previously described (5). The young initial mycelia are fully compartmentalized and have membranous septa (Fig. 2b to c) with little associated cell wall material that is barely visible with WGA (Fig. ​(Fig.2d).2d). In contrast, the second mycelium is a multinucleated structure with fewer membrane-cell wall septa (Fig. 2e to h). At the end of the developmental cycle, multinucleated hyphae begin to undergo the segmentation which precedes the formation of spore chains (Fig. 2i to m). Similar results were obtained for S. antibioticus (not shown), but there were some differences in the numbers of spores formed. Samples of young and late mycelia were freeze-substituted using the methodology described by Porta and Lopez-Iglesias (13) and were examined with a transmission electron microscope (Fig. 2n and o). The septal structure of the first mycelium (Fig. ​(Fig.2n)2n) lacks the complexity of the septal structure in the second mycelium, in which a membrane with a thick cell wall is clearly visible (Fig. ​(Fig.2o).2o). These data coincide with those previously described for solid confluent cultures (4).Open in a separate windowFIG. 2.Analysis of S. coelicolor hyphal compartmentalization with several fluorescent indicators (single colonies). Developmental culture times (in hours) are indicated. (a, e, and i) Mycelium stained with SYTO 9 and propidium iodide (viability). (b, f, and j) Hyphae stained with Cell Mask (a membrane stain). (c, g, and l) Hyphae stained with FM 4-64 (a membrane stain). (d, h, and m) Hyphae stained with WGA (cell wall stain). Septa in all the images in panels a to j, l, and m are indicated by arrows. (k) Image of the same field as panel j obtained in differential interference contrast mode. (n and o) Transmission electron micrographs of S. coelicolor hyphae at different developmental phases. The first-mycelium septa (n) are comprised of two membranes separated by a thin cell wall; in contrast, second-mycelium septa have thick cell walls (o). See the text for details. IP, propidium iodide.The main features of S. coelicolor growing in soils are shown in Fig. ​Fig.3.3. Under these conditions, spore germination is a very slow, nonsynchronous process that commences at about 7 days (Fig. 3c and d) and lasts for at least 21 days (Fig. 3i to l), peaking at around 14 days (Fig. 3e to h). Mycelium does not clump to form dense pellets, as it does in colonies; instead, it remains in the first-compartmentalized-mycelium phase during the time analyzed. Like the membrane septa in single colonies, the membrane septa of the hyphae are stained with FM4-64 (Fig. 3j and k), although only some of them are associated with thick cell walls (WGA staining) (Fig. ​(Fig.3l).3l). Similar results were obtained for S. antibioticus cultures (not shown).Open in a separate windowFIG. 3.Confocal laser scanning fluorescence microscopy analysis of the development-related cell death and hyphal compartmentalization of S. coelicolor M145 growing in soil. Developmental culture times (in days) are indicated. The images in panels b, f, and h were obtained in differential interference contrast mode and correspond to the same fields as the images in panels a, e, and g, respectively. The dark zone in panel h corresponds to a particle of soil containing hyphae. (a, c, d, e, g, i, j, and k) Hyphae stained with SYTO 9, propidium iodide (viability stain), and FM4-64 (membrane stain) simultaneously. (i) SYTO 9 and propidium iodide staining. (j) FM4-64 staining. The image in panel k is an overlay of the images in panels i and j and illustrates that first-mycelium membranous septa are not always apparent when they are stained with nucleic acid stains (SYTO 9 and propidium iodide). (l) Hyphae stained with WGA (cell wall stain), showing the few septa with thick cell walls present in the cells. Septa are indicated by arrows. IP, propidium iodide.In previous work (8), we have shown that the mycelium currently called the substrate mycelium corresponds to the early second multinucleated mycelium, according to our nomenclature, which still lacks the hydrophobic layers characteristic of the aerial mycelium. The aerial mycelium therefore corresponds to the late second mycelium which has acquired hydrophobic covers. This multinucleated mycelium as a whole should be considered the reproductive structure, since it is destined to sporulate (Fig. ​(Fig.4)4) (8). The time course of lysine 6-aminotransferase activity during cephamycin C biosynthesis has been analyzed by other workers using isolated colonies of Streptomyces clavuligerus and confocal microscopy with green fluorescent protein as a reporter (4). A complex medium and a temperature of 29°C were used, conditions which can be considered similar to the conditions used in our work. Interestingly, expression did not occur during the development of the early mycelium and was observed in the mycelium only after 80 h of growth. This suggests that the second mycelium is the antibiotic-producing mycelium, a hypothesis previously confirmed using submerged-growth cultures of S. coelicolor (9).Open in a separate windowFIG. 4.Cell cycle features of Streptomyces growing under natural conditions. Mycelial structures (MI, first mycelium; MII, second mycelium) and cell death are indicated. The postulated vegetative and reproductive phases are also indicated (see text).The significance of the first compartmentalized mycelium has been obscured by its short life span under typical laboratory culture conditions (5, 6, 8). In previous work (3, 7), we postulated that this structure is the vegetative phase of the bacterium, an hypothesis that has been recently corroborated by proteomic analysis (data not shown). Death in confluent cultures begins shortly after germination (4 h) and continues asynchronously for 15 h. The second multinucleated mycelium emerges after this early programmed cell death and is the predominant structure under these conditions. In contrast, as our results here show, the first mycelium lives for a long time in isolated colonies and soil cultures. As suggested in our previous work (5, 6, 8), if we assume that the compartmentalized mycelium is the Streptomyces vegetative growth phase, then this phase is the predominant phase in individual colonies (where it remains for at least 36 h), soils (21 days), and submerged cultures (around 20 h) (9). The differences in the life span of the vegetative phase could be attributable to the extremely high cell densities attained under ordinary laboratory culture conditions, which provoke massive differentiation and sporulation (5-7, 8).But just exactly what are “natural conditions”? Some authors have developed soil cultures of Streptomyces to study survival (16, 17), genetic transfer (14, 17-19), phage-bacterium interactions (3), and antibiotic production (1). Most of these studies were carried out using amended soils (supplemented with chitin and starch), conditions under which growth and sporulation were observed during the first few days (1, 17). These conditions, in fact, might resemble environments that are particularly rich in organic matter where Streptomyces could conceivably develop. However, natural growth conditions imply discontinuous growth and limited colony development (20, 21). To mimic such conditions, we chose relatively poor but more balanced carbon-nitrogen soil cultures (GAE medium-amended soil) and less dense spore inocula, conditions that allow longer mycelium growth times. Other conditions assayed, such as those obtained by irrigating the soil with water alone, did not result in spore germination and mycelial growth (not shown). We were unable to detect death, the second multinucleated mycelium described above, or sporulation, even after 1 month of incubation at 30°C. It is clear that in nature, cell death and sporulation must take place at the end of the long vegetative phase (1, 17) when the imbalance of nutrients results in bacterial differentiation.In summary, the developmental kinetics of Streptomyces under conditions resembling conditions in nature differs substantially from the developmental kinetics observed in ordinary laboratory cultures, a fact that should be born in mind when the significance of development-associated phenomena is analyzed.     

Characterization of the Enzymes Encoded by the Anthrose Biosynthetic Operon of Bacillus anthracis

Bacillus anthracis spores, the etiological agents of anthrax, possess a loosely fitting outer layer called the exosporium that is composed of a basal layer and an external hairlike nap. The filaments of the nap are formed by trimers of the collagenlike glycoprotein BclA. Multiple pentasaccharide and trisaccharide side chains are O linked to BclA. The nonreducing terminal residue of the pentasaccharide side chain is the unusual sugar anthrose. A plausible biosynthetic pathway for anthrose biosynthesis has been proposed, and an antABCD operon encoding four putative anthrose biosynthetic enzymes has been identified. In this study, we genetically and biochemically characterized the activities of these enzymes. We also used mutant B. anthracis strains to determine the effects on BclA glycosylation of individually inactivating the genes of the anthrose operon. The inactivation of antA resulted in the appearance of BclA pentasaccharides containing anthrose analogs possessing shorter side chains linked to the amino group of the sugar. The inactivation of antB resulted in BclA being replaced with only trisaccharides, suggesting that the enzyme encoded by the gene is a dTDP-β-l-rhamnose α-1,3-l-rhamnosyl transferase that attaches the fourth residue of the pentasaccharide side chain. The inactivation of antC and antD resulted in the disappearance of BclA pentasaccharides and the appearance of a tetrasaccharide lacking anthrose. These phenotypes are entirely consistent with the proposed roles for the antABCD-encoded enzymes in anthrose biosynthesis. Purified AntA was then shown to exhibit β-methylcrotonyl-coenzyme A (CoA) hydratase activity, as we predicted. Similarly, we confirmed that purified AntC had aminotransferase activity and that purified AntD displayed N-acyltransferase activity.Bacillus anthracis, the causative agent of anthrax, is a Gram-positive, rod-shaped soil bacterium that forms spores when deprived of essential nutrients (15). Spore formation begins with an asymmetric septation that divides the developing cell into a forespore compartment and a larger mother cell compartment, each of which contains a copy of the genome. The mother cell then engulfs the forespore and surrounds it with three protective layers: a cortex composed of peptidoglycan, a closely apposed proteinaceous coat, and a loosely fitting exosporium (10). Mother cell lysis releases the mature spore, which is dormant and capable of surviving in harsh environments for many years (17). When spores encounter an aqueous environment containing nutrients, they can germinate and grow as vegetative cells (21).Recently, interest in B. anthracis spores has intensified in response to their use as agents of bioterrorism. Of particular interest has been the outermost layer of the spore, the exosporium, which serves as a semipermeable barrier to potentially harmful macromolecules (8, 25) and as the vital first point of contact with the immune system of an infected host (11, 18, 30). The exosporium of B. anthracis and of closely related species, such as Bacillus cereus and Bacillus thuringiensis, is comprised of a paracrystalline basal layer and an external hairlike nap (1). The basal layer contains approximately 20 different proteins (20, 23), while the filaments of the nap are formed by trimers of a single collagenlike glycoprotein called BclA (2, 26). The central region of BclA contains a large number of GXX repeats, and the region varies in length in naturally occurring strains of B. anthracis, resulting in hairlike naps of differing lengths (22, 27). Most of the GXX repeats are GPT, and many of the threonine residues are glycosylated. Two major oligosaccharide side chains are present, a pentasaccharide and a trisaccharide, and both are linked to the protein through reducing terminal N-acetylgalactosamine (GalNAc) residues (3). Several studies have demonstrated that the oligosaccharides are antigenic and are exposed on the surface of Bacillus anthracis spores (14, 29). This makes them prime targets for both detection devices and immunoprophylaxis.We previously reported our use of hydrazinolysis to release BclA oligosaccharides from exosporium preparations (3). The primary product was a tetrasaccharide that formed as a result of the undesirable loss of the reducing terminal GalNAc residue of the pentasaccharide, a process called “peeling.” We determined that the oligosaccharide consisted of a linear chain of three rhamnose residues with a novel deoxyamino sugar at its nonreducing terminus. This unusual sugar, 2-O-methyl-4-(3-hydroxy-3-methylbutamido)-4,6-dideoxy-d-glucose, was given the trivial name anthrose.Rhamnose is the major sugar present in both the trisaccharide and the pentasaccharide, and a four-gene rhamnose biosynthetic operon was previously identified (22). Previously, we proposed a pathway for anthrose biosynthesis (Fig. ​(Fig.1)1) and identified a four-gene operon (Fig. ​(Fig.2)2) that is essential for its biosynthesis (5). An in-frame deletion of the first gene of the operon reduced the amount of anthrose by approximately 50%, whereas the deletion of any one of the other three genes totally abolished anthrose synthesis. Here, we describe the characterization of the altered oligosaccharide side chains of the four deletion mutants. We also cloned several genes that we predicted are involved in anthrose biosynthesis and demonstrated that the gene products possessed the expected biochemical activities.Open in a separate windowFIG. 1.Proposed biosynthetic pathway of anthrose. The pathway utilizes dTDP-4-keto-6-deoxy-α-d-glucose, an intermediate in rhamnose biosynthesis, and methylcrotonyl-CoA, derived from leucine catabolism. (Modified from reference 5.)Open in a separate windowFIG. 2.Anthrose operon and flanking genes. The four genes of the anthrose operon are antA (BAS3322), antB (BAS3321), antC (BAS3320), and antD (BAS3319). The operon is flanked by genes that encode a putative collagenase (BAS3323) and a putative methyltransferase (BAS3318). (Modified from reference 5.)     

Spatial and Developmental Differentiation of Mannitol Dehydrogenase and Mannitol-1-Phosphate Dehydrogenase in Aspergillus niger

The presence of a mannitol cycle in fungi has been subject to discussion for many years. Recent studies have found no evidence for the presence of this cycle and its putative role in regenerating NADPH. However, all enzymes of the cycle could be measured in cultures of Aspergillus niger. In this study we have analyzed the localization of two enzymes from the pathway, mannitol dehydrogenase and mannitol-1-phosphate dehydrogenase, and the expression of their encoding genes in nonsporulating and sporulating cultures of A. niger. Northern analysis demonstrated that mpdA was expressed in both sporulating and nonsporulating mycelia, while expression of mtdA was expressed only in sporulating mycelium. More detailed studies using green fluorescent protein and dTomato fused to the promoters of mtdA and mpdA, respectively, demonstrated that expression of mpdA occurs in vegetative hyphae while mtdA expression occurs in conidiospores. Activity assays for MtdA and MpdA confirmed the expression data, indicating that streaming of these proteins is not likely to occur. These results confirm the absence of the putative mannitol cycle in A. niger as two of the enzymes of the cycle are not present in the same part of A. niger colonies. The results also demonstrate the existence of spore-specific genes and enzymes in A. niger.Mannitol has been described as one of the main compatible solutes in fungi (20) and may play a role as a storage carbon source (3) or a protectant against a variety of stresses (10, 16, 20, 22). Mannitol metabolism in fungi has been the subject of study for decades. It was proposed to exist in the form of a cyclic pathway, the mannitol cycle (9). This cycle consists of four steps enabling the conversion of fructose into mannitol and back to fructose (Fig. 1). The main role proposed for this cycle was regenerating NADPH (9, 10). Subsequently, many studies have questioned the existence of a mannitol cycle (reviewed in reference 20), and it has been shown that a mannitol cycle is not involved in NADPH regeneration in Stagonospora nodorum (19), Aspergillus niger (16), and Alternaria alternata (21). However, all enzymes of the cycle were detected in both sporulating and nonsporulating mycelia in A. niger (16), suggesting that a cycle could operate in this fungus. Fungi are able to use mannitol as a sole carbon source but do so in various ways (7).Open in a separate windowFig. 1.Putative mannitol cycle in fungi as proposed by Hult and Gatenbeck (9). HXK, hexokinase (EC 2.7.1.1); MTD, mannitol dehydrogenase (EC 1.1.1.138); MPD, mannitol-1-phosphate dehydrogenase (EC 1.1.1.17); MPP, mannitol-1-phosphate phosphatase (EC 3.1.3.22).d-Mannitol plays an important role in germination of Aspergillus conidia. In A. niger (23) and Aspergillus oryzae (8), mannitol accumulates in conidiospores and is utilized during the initial stages of germination. Production of mannitol appears to be largely dependent on mannitol-1-phosphate dehydrogenase (MPD) while mannitol dehydrogenase (MTD) contributes to a lesser extent (16, 19, 20).In this study we demonstrate that MTD and MPD as well as the expression of the corresponding genes (mtdA and mpdA) are spatially separated in colonies of A. niger. This demonstrates that a mannitol cycle does not exist in this fungus and shows that spores express specific genes that are involved in germination.     

Role of GerKB in Germination and Outgrowth of Clostridium perfringens Spores

Previous work indicated that Clostridium perfringens gerKA gerKC spores germinate significantly, suggesting that gerKB also has a role in C. perfringens spore germination. We now find that (i) gerKB was expressed only during sporulation, likely in the forespore; (ii) gerKB spores germinated like wild-type spores with nonnutrient germinants and with high concentrations of nutrients but more slowly with low nutrient concentrations; and (iii) gerKB spores had lower colony-forming efficiency and slower outgrowth than wild-type spores. These results suggest that GerKB plays an auxiliary role in spore germination under some conditions and is required for normal spore viability and outgrowth.Spores of Bacillus and Clostridium species can break dormancy upon sensing a variety of compounds (termed germinants), including amino acids, nutrient mixtures, a 1:1 chelate of Ca²⁺ and pyridine-2,6-dicarboxylic acid (dipicolinic acid [DPA]), and cationic surfactants such as dodecylamine (20). Nutrient germinants are sensed by their cognate receptors, located in the spore''s inner membrane (6), which are composed of proteins belonging to the GerA family (10, 11). In Bacillus subtilis, three tricistronic operons (gerA, gerB, and gerK) expressed uniquely during sporulation in the developing forespore each encode the three major germinant receptors, with different receptors responding to a different spectrum of nutrient germinants (5, 9, 20). Null mutations in any cistron in a gerA family operon inactivate the function of the respective receptor (9, 11). In contrast, Clostridium perfringens, a gram-positive, spore-forming, anaerobic pathogenic bacterium, has no tricistronic gerA-like operon but only a monocistronic gerAA that is far from a gerK locus. This locus contains a bicistronic gerKA-gerKC operon and a monocistronic gerKB upstream of and in the opposite orientation to gerKA-gerKC (Fig. ​(Fig.1A)1A) (16). GerAA has an auxiliary role in the germination of C. perfringens spores at low germinant concentrations, while GerKA and/or GerKC are required for l-asparagine germination and have partial roles in germination with KCl and a mixture of KCl and l-asparagine (AK) (16). In contrast to the situation with B. subtilis, where germinant receptors play no role in Ca-DPA germination (12, 13), GerKA and/or GerKC is required for Ca-DPA germination (16). The partial requirement for GerKA and/or GerKC in C. perfringens spore germination by KCl and AK suggests that the upstream gene product, GerKB, might also have some role in KCl and AK germination of C. perfringens spores. Therefore, in this study we have investigated the role of GerKB in the germination and outgrowth of C. perfringens spores.Open in a separate windowFIG. 1.Arrangement and expression of gerKB in C. perfringens SM101. (A) The arrangement of the gerK locus in C. perfringens SM101 is shown, and the locations of the primers used to amplify the upstream regions of the gerKB gene and the putative promoters of gerKB and gerKA are indicated. The gerKB promoter was predicted to be within the intergenic regions between gerKB and the gerK operon. (B) GUS specific activities from the gerKB-gusA fusion in strain SM101(pDP84) grown in TGY vegetative (filled squares) and DS sporulation (open squares) media were determined as described in the text. Data represent averages from three independent experiments with the error bars representing standard deviations, and time zero denotes the time of inoculation of cells into either TGY or DS medium.To determine if gerKB is expressed during sporulation, 485 bp upstream of the gerKB coding sequence, including DNA between gerKB and gerKA, was PCR amplified with primer pair CPP389/CPP391, which had SalI and PstI cleavage sites, respectively (see Table S2 in the supplemental material). The PCR fragment was cloned between SalI and PstI cleavage sites in plasmid pMRS127 (17) to create a gerKB-gusA fusion in plasmid pDP84 (see Table S1 in the supplemental material). This plasmid was introduced into C. perfringens SM101 by electroporation (3), and Em^r transformants were selected. The SM101 transformant carrying plasmid pDP84 was grown in TGY vegetative growth medium (3% Trypticase soy, 2% glucose, 1% yeast extract, 0.1% l-cysteine) (7) and in Duncan-Strong (DS) (4) sporulation medium and assayed for β-glucuronidase (GUS) activity as described previously (23). Vegetative cultures of strain SM101 carrying plasmid pMRS127 (empty vector) or pDP84 (gerKB-gusA) exhibited no significant GUS activity, and strain SM101 grown in DS medium also exhibited no significant GUS activity (Fig. ​(Fig.1B1B and data not shown). However, GUS activity was observed in sporulating cultures of SM101(pDP84) (Fig. ​(Fig.1B),1B), indicating that a sporulation-specific promoter is located upstream of gerKB. The expression of the gerKB-gusA fusion began ∼3 h after induction of sporulation and reached a maximum after ∼6 h of sporulation (Fig. ​(Fig.1B).1B). The decrease in GUS activity observed after ∼6 h is consistent with the GerKB-GusA protein being packaged into the dormant spore where it cannot be easily assayed and thus with gerKB being expressed in the forespore compartment of the sporulating cell (8). These results confirm that, as with the gerKA-gerKC operon (16), gerKB is also expressed only during sporulation.To investigate the role of GerKB in C. perfringens spore germination, we constructed a gerKB mutant strain (DPS108) as described previously (14-16). A 2,203-bp DNA fragment carrying 2,080 bp upstream of and 123 bp from the N-terminal coding region of gerKB was PCR amplified using primers CPP369 and CPP367, which had XhoI and BamHI cleavage sites at the 5′ ends of the forward and reverse primers, respectively (see Table S2 in the supplemental material). A 1,329-bp fragment carrying 134 bp from the C-terminal and 1,195 bp downstream of the coding region of gerKB was PCR amplified using primers CPP371 and CPP370, which had BamHI and KpnI cleavage sites at the 5′ ends of the forward and reverse primers, respectively (see Table S2 in the supplemental material). These PCR fragments were cloned into plasmid pCR-XL-TOPO, giving plasmids pDP67 and pDP69, respectively (see Table S1 in the supplemental material). An ∼2.2-kb BamHI-XhoI fragment from pDP67 was cloned into pDP1 (pCR-XL-TOPO carrying an internal fragment of gerAA), giving plasmid pDP68, and an ∼1.4-kb KpnI-BamHI fragment from pDP69 was cloned in pDP68, giving pDP73 (see Table S1 in the supplemental material). The latter plasmid was digested with BamHI, the ends were filled, and an ∼1.3-kb NaeI-SmaI fragment carrying catP from pJIR418 (1) was inserted, giving plasmid pDP74. Finally, an ∼4.8-kb KpnI-XhoI fragment from pDP74 (see Table S1 in the supplemental material) was cloned between the KpnI and SalI sites of pMRS104, giving pDP75, which cannot replicate in C. perfringens. Plasmid pDP75 was introduced into C. perfringens SM101 by electroporation (3), and the gerKB deletion strain DPS108 was isolated as described previously (18). The presence of the gerKB deletion in strain DPS108 was confirmed by PCR and Southern blot analyses (data not shown). Strain DPS108 gave ∼70% sporulating cells in DS sporulation medium, similar to results with the wild-type strain, SM101 (data not shown).Having obtained evidence for successful construction of the gerKB mutant, we compared the germinations of heat-activated (80°C; 10 min) gerKB and wild-type spores as previously described (16). Both the gerKB and wild-type spores germinated identically and nearly completely in 60 min at 40°C in brain heart infusion (BHI) broth as determined by the fall in optical density at 600 nm (OD₆₀₀) of germinating cultures and phase-contrast microscopy (data not shown). This result suggests that GerKB plays no essential role in spore germination in rich medium. The role of GerKB in C. perfringens spore germination was also assessed with individual germinants identified previously (16). Heat-activated wild-type and gerKB spores germinated similarly with high (100 mM) concentrations of KCl, l-asparagine, and AK, all in 25 mM sodium phosphate (pH 7.0), and in 50 mM Ca-DPA adjusted to pH 8.0 with Tris base (Fig. 2A to D). These results were also confirmed by phase-contrast microscopy (data not shown). However, with lower (10 to 20 mM) concentrations of KCl, l-asparagine, and AK, gerKB spore germination was very slightly (Fig. ​(Fig.2A)2A) to significantly (Fig. 2B and C) slower than that of wild-type spores. These results suggest that while GerKB is not essential for germination with high concentrations of KCl, l-asparagine, or AK, it plays a significant role in germination with low l-asparagine and AK concentrations and, further, that GerKB is not required for Ca-DPA germination. This latter finding is similar to the situation with B. subtilis spores where germinant receptors play no role in Ca-DPA germination (19, 20). However, in C. perfringens spores, GerKA and/or GerKC do play a significant role in Ca-DPA germination (16).Open in a separate windowFIG. 2.Germination of spores of C. perfringens strains with various germinants. Heat-activated spores of strains SM101 (wild type) (filled symbols) and DPS108 (gerKB) (open symbols) were incubated at an OD₆₀₀ of 1 at 40°C with high (squares) and low (triangles) germinant concentrations of 100 and 10 mM KCl (A), 100 and 20 mM l-asparagine (B), 100 and 10 mM AK (C), and 50 mM Ca-DPA (D) as described in the text, and at various times the OD₆₀₀ was measured. No significant germination was observed when heat-activated spores of SM101 and DPS108 were incubated for 60 min at 40°C in 25 mM sodium phosphate buffer (pH 7.0) (data not shown). The data shown are averages from duplicate determinations with two different spore preparations, and error bars represent standard deviations.Bacterial spores can also germinate with dodecylamine, a cationic surfactant (19). In B. subtilis spores, dodecylamine induces germination most likely by opening channels composed, at least in part, of SpoVA proteins (22), allowing release of the spores'' Ca-DPA (19). Spores of B. subtilis lacking all three functional germinant receptors release DPA, as do wild-type spores, upon incubation with dodecylamine (19), while C. perfringens spores lacking GerKA-GerKC incubated with dodecylamine release DPA slower than wild-type spores (16). However, when C. perfringens gerKB spores at an OD₆₀₀ of 1.5 were incubated with 1 mM dodecylamine in Tris-HCl (pH 7.4) at 60°C (2, 16), gerKB spores released their DPA slightly faster than wild-type spores (Fig. ​(Fig.3)3) when DPA release was measured as described previously (16). These results suggest that GerKB has no role in dodecylamine germination.Open in a separate windowFIG. 3.Germination of spores of C. perfringens strains with dodecylamine. Spores of strains SM101 (wild type) (filled squares) and DPS108 (gerKB) (open squares) were germinated with dodecylamine, and germination was monitored by measuring DPA release as described in the text. There was no significant DPA release in 60 min by spores incubated similarly but without dodecylamine (data not shown). Error bars represent standard deviations.Previous work (16) found that C. perfringens spores lacking GerKA-GerKC had lower viability than wild-type spores on rich medium plates, and it was thus of interest to determine gerKB spore viability, which was measured as previously described (14, 16). Strikingly, the colony-forming ability of gerKB spores was ∼7-fold lower (P < 0.01) than that of wild-type spores after 24 h on BHI plates (Table ​(Table1),1), and no additional colonies appeared when plates were incubated for up to 3 days (data not shown). The colony-forming ability of spores lacking GerKA and GerKC determined in parallel was ∼12-fold lower than that of wild-type spores (Table ​(Table1).1). Phase-contrast microscopy of C. perfringens spores incubated in BHI broth for 24 h under aerobic conditions to prevent vegetative cell growth indicated that >90% of wild-type spores not only had germinated but had also released the nascent vegetative cell, while >85% of gerKA gerKC and gerKB spores remained as only phase-dark germinated spores with no evidence of nascent cell release (data not shown), as found previously with gerKA gerKC spores (16). The fact that >85% of gerKB spores germinated in BHI medium in 24 h but most of these germinated spores did not progress further in development strongly suggests that GerKB is needed for normal spore outgrowth (and see below) as well as for normal spore germination.

TABLE 1.

Colony formation by spores of C. perfringens strains^a

Cellulase Production from Spent Lignocellulose Hydrolysates by Recombinant Aspergillus niger

A recombinant Aspergillus niger strain expressing the Hypocrea jecorina endoglucanase Cel7B was grown on spent hydrolysates (stillage) from sugarcane bagasse and spruce wood. The spent hydrolysates served as excellent growth media for the Cel7B-producing strain, A. niger D15[egI], which displayed higher endoglucanase activities in the spent hydrolysates than in standard medium with a comparable monosaccharide content (e.g., 2,100 nkat/ml in spent bagasse hydrolysate compared to 480 nkat/ml in standard glucose-based medium). In addition, A. niger D15[egI] was also able to consume or convert other lignocellulose-derived compounds, such as acetic acid, furan aldehydes, and phenolic compounds, which are recognized as inhibitors of yeast during ethanolic fermentation. The results indicate that enzymes can be produced from the stillage stream as a high-value coproduct in second-generation bioethanol plants in a way that also facilitates recirculation of process water.Energy security, petroleum depletion, and global warming have been the main driving forces for the development of renewable fuels that can replace petroleum-derived fuels, such as gasoline and diesel. Bioethanol is currently the most commonly used renewable automobile fuel. It is largely produced by fermentation of sugar- or starch-containing feedstocks, such as cane sugar, corn, and wheat. However, the supply of these crops is relatively limited and many of them can be considered human food resources. Lignocellulose is a more-abundant and less-expensive raw material with the potential to give a high net energy gain (11, 17).In the production of bioethanol from lignocellulosic materials, hydrolytic enzymes, such as cellulases and cellobiases, can be used to convert the lignocellulosic polysaccharides to monosaccharides. Microorganisms can be used to ferment the monosaccharides to ethanol. The yeast Saccharomyces cerevisiae is one of the most suitable microorganisms for ethanol production and is favored in industrial processes. However, S. cerevisiae only metabolizes hexose sugars. Many lignocellulosic materials consist of a significant proportion of xylan and arabinan, which give rise to pentose sugars. The cost of enzymes for the hydrolysis of polysaccharides and the inability of S. cerevisiae to utilize pentose sugars have been pointed out as two bottlenecks for commercialization of cellulosic ethanol production (9, 25). Considerable research efforts have therefore been focused on reducing the enzyme cost by producing more-efficient enzymes from cheaper growth media (25). Other efforts have been focused on different approaches to convert pentose sugars to ethanol by using recombinant microorganisms (3, 10).A novel approach to reduce the enzyme cost and to optimally utilize all sugars generated from lignocellulose would be to produce hydrolytic enzymes, such as cellulases, from the pentose fraction remaining after consumption of hexoses by S. cerevisiae (Fig. ​(Fig.1).1). The cellulases produced can then be used on site in the next round of hydrolysis of the lignocellulosic feedstock and thereby reduce the dependence on externally produced enzymes.Open in a separate windowFIG. 1.Schematic representation of the experimental approach and on-site enzyme production in a cellulose-to-ethanol process.Furthermore, it is desirable to recycle the process water in an ethanol production plant to minimize the production costs. However, lignocellulose hydrolysates are very complex and contain a wide range of different compounds. Some of these compounds, such as furan aldehydes, aliphatic acids, and phenolic compounds, inhibit the yeast S. cerevisiae, which results in lower ethanol yield and productivity. Recycling of the process water can lead to a buildup in the concentration of inhibitors, which is a phenomenon that has been pointed out as an obstacle to reusing the stillage stream (1, 35). There are several methods to avoid inhibitor-related problems, but they are often associated with additional process cost (40). However, A. niger is an organism that can utilize a broad range of compounds as nutrients, possibly including compounds that inhibit S. cerevisiae. It would be convenient if the A. niger cells could metabolize such compounds and thereby, due to the removal of inhibitors, make it more feasible to reuse the process water.In this study, we explored the possibility of utilizing sugarcane bagasse and spruce wood for ethanol production and using the spent hydrolysates (stillage) for production of the Hypocrea jecorina cellulase Cel7B (formerly called endoglucanase I) by a recombinant strain of Aspergillus niger. Simultaneously, the Cel7B-producing recombinant A. niger strain also removed inhibitory lignocellulose-derived products, thus facilitating recycling of process water.     

Cloning of a Novel Pyrethroid-Hydrolyzing Carboxylesterase Gene from Sphingobium sp. Strain JZ-1 and Characterization of the Gene Product

A novel esterase gene, pytH, encoding a pyrethroid-hydrolyzing carboxylesterase was cloned from Sphingobium sp. strain JZ-1. The gene contained an open reading frame of 840 bp. Sequence identity searches revealed that the deduced enzyme shared the highest similarity with many α/β-hydrolase fold proteins (20 to 24% identities). PytH was expressed in Escherichia coli BL21 and purified using Ni-nitrilotriacetic acid affinity chromatography. It was a monomeric structure with a molecular mass of approximately 31 kDa and a pI of 4.85. PytH was able to transform p-nitrophenyl esters of short-chain fatty acids and a wide range of pyrethroid pesticides, and isomer selectivity was not observed. No cofactors were required for enzyme activity.Pyrethroid pesticides are now the major class of insecticides used for insect control in agriculture and households as a replacement for more toxic organophosphorus pesticides, and their usage is continuing to grow (10). Although pyrethroid pesticides generally have lower acute oral mammalian toxicity than organophosphate insecticides, exposure to very high levels of pyrethroid pesticides might cause endocrine disruption, lymph node and spleen damage, and carcinogenesis (6, 12). In addition, most pyrethroid pesticides possess acute toxicity to some nontarget organisms, such as bees, fish, and aquatic invertebrates, often at concentrations of less than 0.5 μg/kg (19, 22). Great concerns have been raised about the persistence and degradation of pyrethroid pesticides in the environment.In general, pyrethroid pesticides are degraded by both abiotic and biotic pathways, including photooxidation, chemical oxidation, and biodegradation. Microorganisms play the most important role in degradation of pyrethroids in soils and sediments. Many pyrethroid-degrading microorganisms have been isolated from soils (13, 16, 24, 27).The major routes of pyrethroid metabolism in pyrethroid-resistant insects and pyrethroid-degrading microorganisms include oxidation by cytochrome P450s and ester hydrolysis by carboxylesterases (9). Carboxylesterases are a family of enzymes that are important in the hydrolysis of a large number of endogenous and xenobiotic ester-containing compounds, such as carbamates, organophosphorus pesticides, and pyrethroids. Carboxylesterases from Bacillus cereus SM3 (17), Aspergillus niger ZD11 (13), Nephotettix cincticeps (2), and mouse liver microsomes (23) hydrolyzing the carboxyl ester linkage of the pyrethroids were purified to homogeneity and characterized. Genes encoding the pyrethroid-hydrolyzing carboxylesterases from mouse liver microsomes and Klebsiella sp. strain ZD112 were cloned and functionally expressed (23, 27).Pyrethroids differ from many other pesticides in that they contain one to three chiral centers; the chirality may arise from the acid moiety, the alcohol moiety, or both (Fig. ​(Fig.1).1). A pyrethroid compound therefore consists of two to eight isomers. Isomers of a chiral compound often differ from each other in biological properties. Isomer selectivity has been widely observed in insecticidal activity for the isomers of a pyrethroid compound. Recently, studies have shown that biodegradation of pyrethroids also exhibits significant isomer selectivity (15, 23).Open in a separate windowFIG. 1.Molecular structures of pyrethroids tested. Chiral centers are indicated by black dots.In this study, we described the isolation and identification of the pyrethroid-degrading Sphingobium sp. strain JZ-1, the cloning and expression of the gene pytH encoding a novel pyrethroid-hydrolyzing carboxylesterase, and the characterization of the purified enzyme.     

Physical Characteristics of Spores of Food-Associated Isolates of the Bacillus cereus Group

All 47 food-borne isolates of Bacillus cereus sensu stricto, as well as 10 of 12 food-borne, enterotoxigenic isolates of Bacillus thuringiensis, possessed appendages. Spores were moderately to highly hydrophobic, and each had a net negative charge. These characteristics indicate that spores of food-associated B. thuringiensis and not only B. cereus sensu stricto have high potential to adhere to inert surfaces.Bacillus cereus is a worldwide food-borne pathogen causing diarrheal or emetic-type illnesses. The presumptive toxins have been identified in each case (2, 9, 11). We recently reported that the diarrheal type was the more common toxigenic type in U.S. retail rice and seafood (5, 24). Spores of B. thuringiensis and B. mycoides were also isolated from rice. Best known for the insecticidal activity of its parasporal crystals, Bacillus thuringiensis has been associated with gastroenteritis and isolated in rare cases from outbreaks of food-borne illness (17). Isolates of B. thuringiensis, including those isolated from commercial insecticides, have been shown to produce one or both of the enterotoxins HBL and NHE (6, 14, 18, 22, 23, 28).Bacterial spores can adhere to inert surfaces of food processing equipment due to their surface properties and structures (8, 29). Spores of certain Clostridium and Bacillus species possess appendage-like structures (1, 12, 15, 19) which may contribute to biofilm formation (3, 30). Previous studies of the physical properties of spores of the B. cereus group have focused primarily on environmental isolates. Here the spore morphology, hydrophobic characteristics, and net negative charge of food-borne and food poisoning-associated isolates of the B. cereus group were investigated including potentially enterotoxigenic B. thuringiensis.The diarrheal-type B. cereus strains 85 and 133, B. thuringiensis strains 105 and 129, and B. mycoides strain 157 isolated from rice were examined in detail in this study. These B. cereus and B. thuringiensis strains were selected on the basis of the presence of the nhe or hbl gene along with the ability to produce the corresponding gene product at elevated titers as previously described (5). Bacillus subtilis (ATCC 6633) was used as a comparative reference in the surface charge and hydrophobicity studies. In addition to the above isolates, the presence of appendages on spores of the following was also examined by negative staining: 32 isolates of diarrheal B. cereus isolated from seafood (24), an NHE-positive control strain (ATCC 1230/88), 12 emetic-type B. cereus isolates from food poisoning outbreaks (20), and 10 food-borne isolates of B. thuringiensis (in addition to the above two) (5). Spores were produced as previously described (4).The spores of B. thuringiensis were separated from the inclusion bodies (IB) by centrifuging in step gradients of 0.6, 1.0, 1.4, and 1.8 g/ml sucrose. Cleaned spore suspensions in sodium-potassium phosphate buffer were diluted to an A₆₀₀ of 0.4. Two to three ml of the diluted suspension was layered on top of the gradient. The gradients were centrifuged for 2.5 h in a swinging bucket rotor (Dupont-Sorvall) at 450 × g at 10°C. A visible white layer of spores was collected from the bottom with a Pasteur pipette. The layer was washed with 0.85% saline (at least twice) and stored in the same at 4°C. Following these procedures, spore suspensions had <20% inclusions (relative to spores) as observed by phase-contrast microscopy.Spore hydrophobicity was measured using the bacterial adhesion to hydrocarbon (BATH) assay (26) modified from the observations reported elsewhere (16). The mean and the standard error were calculated from a minimum of seven measurements. ζ potentials of the spores were measured in a Malvern Ζetasizer model nano 2S (Malvern Instruments, Westborough, MA) using the Smoluchowski equation (26). The spores were suspended in saline (0.15 M) at a pH of 6 to 7. The ζ potentials were determined from a mean value of five measurements.Spores of two diarrheal B. cereus strains, two B. thuringiensis strains, and one B. mycoides strain were examined by transmission electron microscopy (TEM). Appendages were observed on B. cereus (Fig. 1a, b, and c) and B. thuringiensis (Fig. 1d and e) but not B. mycoides (Fig. ​(Fig.1f).1f). In contrast, exosporia were seen in all the isolates examined (Fig. ​(Fig.1).1). Similar observations for environmental and clinical samples of these species have been reported by others (13, 16, 21, 29). The number of appendages observed here varied among strains. In the case of B. cereus these ranged from three to four (isolate 133) to four to nine (isolate 85). On the other hand, B. thuringiensis 129 had a higher number (12 to 18) of appendages per spore. The appendage length for B. cereus varied from 0.45 to 3.8 μm. Appendages appeared tube-like in appearance, with an average diameter of 13.6 nm (Fig. ​(Fig.1b)1b) as determined by Image J software (http://rsb.info.nih.gov/ij). For each species, examined appendages were often lophotrichous (Fig. ​(Fig.1e)1e) though peritrichous appendages were more common. All 35 food-borne B. cereus isolates examined in this study and one B. cereus NHE control strain possessed appendages, as did 12 of 12 food poisoning-associated, emetic-type B. cereus isolates. Appendages are a common but not universal feature of the B. cereus group. Whether the number and length of spore appendages of the B. cereus group are species associated or due to their fragility and loss during the preparation procedures (16, 30) remains a possibility. That all 47 B. cereus sensu stricto isolates examined here possessed spore appendages suggests that these structures are characteristic of this species. There is some controversy as to their role in adhesion (27).Open in a separate windowFIG. 1.Electron micrographs of Bacillus cereus group spores. (a) Shadowed image of B. cereus 85 showing appendages and exosporium; (b) appendage alone; (c) B. cereus 133 showing appendage and exosporium; (d) B. thuringiensis 129 showing appendage, exosporium, and inclusion; (e) negative stain of B. thuringiensis 129 showing lophotrichous appendages; (f) shadowed image of B. mycoides 157 showing exosporium and lack of appendages.Large differences in the relative hydrophobicity of five food-borne isolates representing three Bacillus species of the B. cereus group were not observed among the species examined (Table ​(Table1).1). The hydrophobicity values for the isolates tested were in a narrow range of 42.4 to 55.6%. Similar values for spores of B. cereus sensu stricto have been reported by others (16). B. subtilis ATCC 6633 was included for comparative purposes. Its relative hydrophobicity (Table ​(Table1)1) was the lowest among spores examined and similar to that reported by Husmark and Ronner (16) for this strain. Using the same procedures, Doyle et al. (7) reported adherence values of 38.3% and 61.4% for two isolates of B. thuringiensis, compared to 42.4% and 55.6% observed here for B. thuringiensis (Table ​(Table1).1). From the values obtained here, our isolates can be classified to be moderately to highly hydrophobic. Exosporia have been proposed to be responsible for surface hydrophobicity of spores (16, 25). As mentioned above, exosporia were observed here in all isolates examined.

TABLE 1.

Relative hydrophobicity and charge of spores of food-borne isolates of the B. cereus group

Identification of the Biosynthetic Gene Cluster for the Pseudomonas aeruginosa Antimetabolite l-2-Amino-4-Methoxy-trans-3-Butenoic Acid

l-2-Amino-4-methoxy-trans-3-butenoic acid (AMB) is a potent antibiotic and toxin produced by Pseudomonas aeruginosa. Using a novel biochemical assay combined with site-directed mutagenesis in strain PAO1, we have identified a five-gene cluster specifying AMB biosynthesis, probably involving a thiotemplate mechanism. Overexpression of this cluster in strain PA7, a natural AMB-negative isolate, led to AMB overproduction.The Gram-negative bacterium Pseudomonas aeruginosa is an opportunistic pathogen that causes a wide range of human infections and is considered the main pathogen responsible for chronic pneumonia in cystic fibrosis patients (7, 23). P. aeruginosa also infects other organisms, such as insects (4), nematodes (6), plants (18), and amoebae (20). Its ability to thrive as a pathogen and to compete in aquatic and soil environments can be partly attributed to the production and interplay of secreted virulence factors and secondary metabolites. While the importance of many of these exoproducts has been studied, the antimetabolite l-2-amino-4-methoxy-trans-3-butenoic acid (AMB; methoxyvinylglycine) (Fig. ​(Fig.1)1) has received only limited attention. Identified during a search for new antibiotics, AMB was found to reversibly inhibit the growth of Bacillus spp. (26) and Escherichia coli (25) and was later shown to inhibit the growth and metabolism of cultured Walker carcinosarcoma cells (28). AMB is a γ-substituted vinylglycine, a naturally occurring amino acid with a β,γ-C=C double bond. Other members of this family are aminoethoxyvinylglycine from Streptomyces spp. (19) and rhizobitoxine, made by Bradyrhizobium japonicum (16) and Pseudomonas andropogonis (15) (Fig. ​(Fig.1).1). As inhibitors of pyridoxal phosphate-dependent enzymes (13, 17, 21, 22), γ-substituted vinylglycines have multiple targets in bacteria, animals, and plants (3, 5, 10, 21, 22, 29). However, the importance of AMB as a toxin in biological interactions with P. aeruginosa has not been addressed, as AMB biosynthesis and the genes involved have not been elucidated.Open in a separate windowFIG. 1.Chemical structures of the γ-substituted vinylglycines AMB, aminoethoxyvinylglycine, and rhizobitoxine.     

Novel Technique for Quantifying Adhesion of Metarhizium anisopliae Conidia to the Tick Cuticle

The present study describes an accurate quantitative method for quantifying the adherence of conidia to the arthropod cuticle and the dynamics of conidial germination on the host. The method was developed using conidia of Metarhizium anisopliae var. anisopliae (Metschn.) Sorokin (Hypocreales: Clavicipitaceae) and engorged Rhipicephalus annulatus (Say) (Arachnida: Ixodidae) females and was also verified for M. anisopliae var. acridum Driver et Milner (Hypocreales: Clavicipitaceae) and Alphitobius diaperinus (Panzer) (Coleoptera: Tenebrionidae) larvae. This novel method is based on using an organic solvent (dichloromethane [DCM]) to remove the adhered conidia from the tick cuticle, suspending the conidia in a detergent solution, and then counting them using a hemocytometer. To confirm the efficacy of the method, scanning electron microscopy (SEM) was used to observe the conidial adherence to and removal from the tick cuticle. As the concentration of conidia in the suspension increased, there were correlating increases in both the number of conidia adhering to engorged female R. annulatus and tick mortality. However, no correlation was observed between a tick''s susceptibility to fungal infection and the amount of adhered conidia. These findings support the commonly accepted understanding of the nature of the adhesion process. The mechanism enabling the removal of the adhered conidia from the host cuticle is discussed.The entomopathogenic fungus Metarhizium anisopliae var. anisopliae (Metschn.) Sorokîn (1883) infects a broad range of arthropod hosts and can be used as a biopesticide against different insect and tick species (8, 22, 35, 36). The adhesion of the conidia of entomopathogenic fungi to the host cuticle is the initial stage of the pathogenic process and includes both passive and active events (5, 10). The hydrophobic epicuticular lipid layer plays an important role during both the attachment process and the germination of the conidia on the surface of the host (15, 19). According to Boucias et al. (7), the attachment of conidia to the host cuticle is based on nonspecific hydrophobic and electrostatic forces. The conidia of most entomopathogenic fungi, including M. anisopliae, have an outer cell layer made up of rodlets (6). The hydrophobins, specific proteins present in the rodlet layer, mediate the passive adhesion of conidia to hydrophobic surfaces, such as the cuticles of arthropods (16, 45, 46). However, as germination commences, the hydrophobins are replaced by an adhesion-like protein, Mad1, which promotes tighter and more-specific adhesion between the conidia and the host (44). Many factors may affect the adhesion and persistence of conidia on the host cuticle (i.e., characteristics of the pathogen, including its virulence [2, 18, 48], conditions under which the pathogen is cultured [17], type of spores [7, 16], topographical and chemical properties of the host cuticle [9, 38, 42], host surface hydrophobicity [15, 23], host behavior [31, 33], and environmental conditions [33]). Conidia of M. anisopliae have shown an affinity to cuticular regions containing setae or spines (7, 38) and to highly hydrophobic cuticle regions, such as mosquitoes'' siphon tubes (23) and intersegmental folds (43). Sites with higher numbers of adhered conidia varied among host species. However, in general, the membranous intersegmental regions were often particularly attractive sites for conidial attachment (26). Variation in the distribution of conidia across different anatomical regions has also been noted in studies of several tick species inoculated with entomopathogenic fungi (3, 21, 22). An evaluation of the attachment of Beauveria bassiana conidia to three tick species, Dermacentor variabilis, Rhipicephalus sanguineus, and Ixodes scapularis, demonstrated that the distribution patterns of the different conidia on the ticks'' bodies were not uniform (22). The density of the conidia and their germination varied dramatically across different anatomical regions of Amblyomma maculatum and A. americanum that had been inoculated with B. bassiana (21). Arruda et al. (3) demonstrated that mass adhesion of M. anisopliae conidia to engorged Boophilus microplus females occurs predominantly on ticks'' legs, suggesting its association with the presence of setae.There are a few approaches for assessing the adhesion of conidia to the host cuticle that are based on direct observation of the conidia on the arthropod cuticle. They involve examining a few areas on the surface of an arthropod by means of scanning electron microscopy (SEM) (11, 15, 30), transmission electron microscopy (TEM) (4), or fluorescence microscopy following vital staining of the conidia (2, 28, 29, 37). These methods are expensive, time-consuming, and relatively inaccurate due to the uneven distribution of conidia on the host surface.In this work, we describe a quantitative method for determining the total amount of conidia that have adhered to a whole host cuticle. This method is based on removing adhered conidia from the tick cuticle using an organic solvent, separating the conidia from the extract by centrifugation, resuspending the conidia in a detergent solution, and then counting the conidia in a hemocytometer. The efficacy of the method was evaluated by comparing the results of this procedure with those of a supplementary examination of conidial removal using SEM.The term “adhered” is often used to define conidia in different states: washed or unwashed after inoculation, present on the host cuticle immediately after inoculation, or kept for several hours (1, 2, 38). In this paper, the term “adhered conidia” refers to conidia that remained on the cuticle after washing by vortexing the inoculated and dried host in an aqueous solution of Triton X-100 and rinsing of the material under tap water.     

Contributions of Four Cortex Lytic Enzymes to Germination of Bacillus anthracis Spores

Bacterial spores remain dormant and highly resistant to environmental stress until they germinate. Completion of germination requires the degradation of spore cortex peptidoglycan by germination-specific lytic enzymes (GSLEs). Bacillus anthracis has four GSLEs: CwlJ1, CwlJ2, SleB, and SleL. In this study, the cooperative action of all four GSLEs in vivo was investigated by combining in-frame deletion mutations to generate all possible double, triple, and quadruple GSLE mutant strains. Analyses of mutant strains during spore germination and outgrowth combined observations of optical density loss, colony-producing ability, and quantitative identification of spore cortex fragments. The lytic transglycosylase SleB alone can facilitate enough digestion to allow full spore viability and generates a variety of small and large cortex fragments. CwlJ1 is also sufficient to allow completion of nutrient-triggered germination independently and is a major factor in Ca²⁺-dipicolinic acid (DPA)-triggered germination, but its enzymatic activity remains unidentified because its products are large and not readily released from the spore''s integuments. CwlJ2 contributes the least to overall cortex digestion but plays a subsidiary role in Ca²⁺-DPA-induced germination. SleL is an N-acetylglucosaminidase that plays the major role in hydrolyzing the large products of other GSLEs into small, rapidly released muropeptides. As the roles of these enzymes in cortex degradation become clearer, they will be targets for methods to stimulate premature germination of B. anthracis spores, greatly simplifying decontamination measures.The Gram-positive bacterium Bacillus anthracis is the etiologic agent of cutaneous, gastrointestinal, and inhalational anthrax (24). An anthrax infection begins when the host is infected with highly resistant, quiescent B. anthracis spores (1, 24). Within the host, the spore''s sensory mechanism recognizes chemical signals, known as germinants, and triggers germination, which leads to the resumption of metabolism (36). Spores that have differentiated into vegetative cells produce a protective capsule and deadly toxins. These virulence factors allow the bacteria to evade the host''s immune system and establish an infection resulting in septicemia, toxemia, and frequently death (24). Although vegetative cells produce virulence factors that are potentially fatal, these cells cannot initiate infections and are much more susceptible to antimicrobial treatments than spores (24). Therefore, efficient triggering of spore germination may enhance current decontamination methods.Spores are highly resistant to many environmental insults because the spore core (cytoplasm) is dehydrated, dormant, and surrounded by multiple protective layers, including a modified layer of peptidoglycan (PG) known as the cortex (36). The cortex functions to maintain dormancy and heat resistance by preventing core rehydration (9). It is composed of alternating N-acetylglucosamine (NAG) and N-acetylmuramic acid (NAM) sugars (Fig. ​(Fig.1).1). Peptide side chains on the NAM residues are either involved in interstrand cross-linking, cleaved to single l-alanine side chains, or fully removed with accompanying formation of muramic-δ-lactam (2, 31, 38). After germination is initiated by either nutrient or nonnutrient germinants, the cortex is depolymerized, resulting in complete core rehydration, resumption of metabolic activity, and outgrowth (33, 36).Open in a separate windowFIG. 1.Spore PG structure and hydrolysis. The central structure shows a representative spore PG strand with alternating NAG and NAM or muramic-δ-lactam (MδL) residues and with tetrapeptide or l-Ala side chains on the NAM residues. Forked arrows originate at sites of hydrolysis by the indicated enzymes and point to muropeptide products. The indicated “aG” muropeptide names are as previously published (7, 11). SleB lytic transglycosylase activity produces muropeptides terminating in anhydro-NAM. Cleavage at adjacent NAM residues produces the tetrasaccharide aG7a or aG7b, while cleavage further apart can produce octasaccharides or larger fragments. These can be further cleaved by muramidase treatment, resulting in the production of tetrasaccharide N, which terminates in NAM. The N-acetylglucosaminidase activity of SleL produces tetrasaccharides terminating in NAG, which can be further cleaved by muramidase to trisaccharides terminating in NAM.Cortex hydrolysis is driven by autolysins called germination-specific cortex lytic enzymes (GSLEs) that recognize the cortex-specific muramic-δ-lactam residues (2, 4, 21, 32). GSLEs fall into two classes: spore cortex lytic enzymes (SCLEs), which are thought to depolymerize intact cortical PG, and cortical fragment lytic enzymes (CFLEs), which further degrade partially hydrolyzed cortex (21). Both SCLEs and CFLEs have been identified in a variety of spore-forming species, including B. anthracis (11, 18, 19), Bacillus cereus (4, 20, 26), Bacillus megaterium (8, 34), Bacillus subtilis (13, 16, 25), Bacillus thuringiensis (12), and Clostridium perfringens (5, 23). Of the four GSLEs identified in B. anthracis, CwlJ1, CwlJ2, and SleB are predicted to be SCLEs (11), whereas SleL is thought to be a CFLE (18).Recently, independent studies showed that CwlJ1 and the lytic transglycosylase SleB (Fig. ​(Fig.1)1) play partially redundant roles and that either is sufficient for spore germination and outgrowth (10, 11). However, these same studies report conflicting results concerning the role of CwlJ2 during germination. Heffron et al. found no effect of CwlJ2 on the biochemistry of cortex hydrolysis or on colony-forming efficiency of spores (11). Giebel et al. reported that loss of CwlJ2 caused a minor defect in germination kinetics and that in the absence of SleB and CwlJ1, further loss of CwlJ2 had a major effect on colony forming efficiency (10). SleL in Bacillus anthracis is proposed to be an N-acetylglucosaminidase (Fig. ​(Fig.1)1) whose role is to further degrade cortex fragments resulting from SCLE hydrolysis (18). SleL is not essential for the completion of germination but does promote the release of small muropeptides to the spore''s surrounding environment (18).This study reports the effects of multiple deletion mutations affecting GSLEs on spore germination efficiency and kinetics of cortex hydrolysis. The data confirm the dominant roles played by CwlJ1 and SleB in the initiation of cortex hydrolysis and the major role of SleL in release of small cortex fragments. A minor role of CwlJ2 in nutrient-triggered germination and the contributions of CwlJ1 and CwlJ2 to Ca²⁺-dipicolinic acid (DPA)-triggered germination were revealed.     

Inhibition of the Activity of Both Domains of Lysostaphin through Peptidoglycan Modification by the Lysostaphin Immunity Protein

Resistance to lysostaphin, a staphylolytic glycylglycine endopeptidase, is due to a FemABX-like immunity protein that inserts serines in place of some glycines in peptidoglycan cross bridges. These modifications inhibit both binding of the recombinant cell wall targeting domain and catalysis by the recombinant catalytic domain of lysostaphin.Lysostaphin is a glycylglycine endopeptidase produced by Staphylococcus simulans biovar staphylolyticus (18) that lyses susceptible staphylococci by hydrolyzing the polyglycine cross bridges in their cell wall peptidoglycans (3). The lysostaphin gene sequence was independently determined in 1987 by two groups (8, 13). BLAST analysis (1) of mature lysostaphin revealed two domains: an N-terminal catalytic domain (CAT), which is a member of the M23 family of zinc metalloendopeptidases, and a C-terminal cell wall targeting domain (CWT), which is a member of the SH3b domain family (Fig. ​(Fig.11 A).Open in a separate windowFIG. 1.(A) Schematic diagram of mature lysostaphin, the recombinant catalytic domain (rCAT) (lysostaphin residues 1 to 148), and the recombinant cell wall targeting domain (rCWT) (lysostaphin residues 149 to 246). The numbers represent the beginning and end of the domains, and the solid boxes indicate the N-terminal His₆ tag of the recombinant proteins. (B) SDS-PAGE analysis of rCAT and rCWT purified by a nickel affinity column. Mobilities of molecular mass standards are given on the left side of the gel.The lysostaphin endopeptidase resistance gene (epr or lif) encodes a FemABX-like immunity protein that is located adjacent to the lysostaphin gene on the plasmid pACK1 in S. simulans bv. staphylolyticus (4, 7, 20). Members of the FemABX family of proteins are nonribosomal peptidyl transferases that are involved in the addition of cross bridge amino acids during peptidoglycan subunit synthesis in the cytoplasm (15). In S. simulans bv. staphylolyticus, the lysostaphin immunity protein inserts serines in place of some glycines during peptidoglycan synthesis, which provides resistance to lysostaphin (4, 20).Originally it was suggested that the incorporation of serines in these peptidoglycan cross bridges gave increased resistance to lysostaphin because of the inability of the enzyme to hydrolyze glycyl-serine or seryl-glycine bonds (4, 14, 16). Others later reported that the CWT specifically binds to the polyglycine cross bridges in staphylococci (6) and the binding of CWT to producer-strain cells was less than that to susceptible cells (2). However, the ability of the enzyme or its targeting domain to bind to purified peptidoglycans from staphylococci containing the lysostaphin resistance gene has not been determined. Therefore, we determined if the modification to staphylococcal peptidoglycan cross bridges made by the lysostaphin immunity protein affected the activity of the binding domain, the catalytic domain, or both.     

Metabolic Engineering of the Tricarboxylic Acid Cycle for Improved Lysine Production by Corynebacterium glutamicum

In the present work, lysine production by Corynebacterium glutamicum was improved by metabolic engineering of the tricarboxylic acid (TCA) cycle. The 70% decreased activity of isocitrate dehydrogenase, achieved by start codon exchange, resulted in a >40% improved lysine production. By flux analysis, this could be correlated to a flux shift from the TCA cycle toward anaplerotic carboxylation.With an annual market volume of more than 1,000,000 tons, lysine is one of the dominating products in biotechnology. In recent years, rational metabolic engineering has emerged as a powerful tool for lysine production (16, 18, 22). Hereby, different target enzymes and pathways in the central metabolism could be identified and successfully modified to create superior production strains (1, 2, 5, 8, 10, 17-20). The tricarboxylic acid (TCA) cycle has not been rationally engineered so far, despite its major role in Corynebacterium glutamicum (6). From metabolic flux studies, however, it seems that the TCA cycle might offer a great potential for optimization (Fig. ​(Fig.1),1), which is also predicted from in silico pathway analysis (13, 22). Experimental evidence comes from studies with Brevibacterium flavum exhibiting increased lysine production due to an induced bottleneck toward the TCA cycle (21). In the present work, we performed TCA cycle engineering by downregulation of isocitrate dehydrogenase (ICD). ICD is the highest expressed TCA cycle enzyme in C. glutamicum (7). Downregulation was achieved by start codon exchange, controlling ICD expression on the level of translation.Open in a separate windowFIG. 1.Stoichiometric correlation of lysine yield (%), biomass yield (g/mol) and TCA cycle flux (%; entry flux through citrate synthase) determined by ¹³C metabolic flux analysis achieved by paraboloid fitting of the data set (parameters were determined with Y0 = 105.1, a = −1.27, b = 0.35, c = −9.35 × 10⁻³, d = −11.16 × 10⁻³). The data displayed represent values from 18 independent experiments with different C. glutamicum strains taken from previous studies (1-3, 11, 12, 15, 23).     

Antibody Recognition Force Microscopy Shows that Outer Membrane Cytochromes OmcA and MtrC Are Expressed on the Exterior Surface of Shewanella oneidensis MR-1

Antibody recognition force microscopy showed that OmcA and MtrC are expressed on the exterior surface of living Shewanella oneidensis MR-1 cells when Fe(III), including solid-phase hematite (Fe₂O₃), was the terminal electron acceptor. OmcA was localized to the interface between the cell and mineral. MtrC displayed a more uniform distribution across the cell surface. Both cytochromes were associated with an extracellular polymeric substance.Shewanella oneidensis MR-1 is a dissimilatory metal-reducing bacterium that is well known for its ability to use a variety of anaerobic terminal electron acceptors (TEAs), including solid-phase iron oxide minerals, such as goethite and hematite (8, 10). Previous studies suggest that S. oneidensis MR-1 uses outer membrane cytochromes OmcA and MtrC to catalyze the terminal reduction of Fe(III) through direct contact with the extracellular iron oxide mineral (2, 8, 10, 15, 16, 20, 21, 23). However, it has yet to be shown whether OmcA or MtrC is actually targeted to the external surface of live S. oneidensis MR-1 cells when Fe(III) serves as the TEA.In the present study, we used atomic force microscopy (AFM) to probe the surface of live S. oneidensis MR-1 cells, using AFM tips that were functionalized with cytochrome-specific polyclonal antibodies (i.e., anti-OmcA or anti-MtrC). This technique, termed antibody recognition force microscopy (Ig-RFM), detects binding events that occur between antibodies (e.g., anti-OmcA) on an AFM tip and antigens (e.g., OmcA) that are exposed on a cell surface. While this is a relatively new technique, Ig-RFM has been used to map the nanoscale spatial location of single molecules in complex biological structures under physiological conditions (5, 9, 11, 13).Anti-MtrC or anti-OmcA molecules were covalently coupled to silicon nitride (Si₃N₄) cantilevers (Veeco or Olympus) via a flexible, heterofunctional polyethylene glycol (PEG) linker molecule. The PEG linker consists of an NHS (N-hydroxysuccinimide) group at one end and an aldehyde group at the other end (i.e., NHS-PEG-aldehyde). AFM tips were functionalized with amine groups, using ethanolamine (6, 7). The active NHS ester of the NHS-PEG-aldehyde linker molecule was then used to form a covalent linkage between PEG-aldehyde and the amine groups on the AFM tips (6, 7). Next, anti-MtrC or anti-OmcA molecules were covalently tethered to these tips via the linker molecule''s aldehyde group. This was accomplished by incubating the tips with antibody (0.2 mg/ml) and NaCNBH₃ as described previously (7). The cantilevers were purchased from Veeco and had spring constant values between 0.06 and 0.07 N/m, as determined by the thermal method of Hutter and Bechhoefer (12).Prior to conducting the Ig-RFM experiments, the specificity of each polyclonal antibody (i.e., anti-OmcA and anti-MtrC) for OmcA or MtrC was verified by Western blot analysis as described previously (24, 28). Proteins were resolved by both denaturing and nondenaturing polyacrylamide gel electrophoresis (PAGE). Briefly, 2.5 μg of purified OmcA or MtrC (23) was resolved by sodium dodecyl sulfate-PAGE or native PAGE, transferred to a polyvinylidene difluoride membrane, incubated with either anti-OmcA or anti-MtrC, and then visualized using the Amersham ECL Plus Western blotting detection kit. Anti-OmcA bound exclusively to OmcA, anti-MtrC bound exclusively to MtrC, and neither antibody showed cross-reactivity with the other cytochrome. Antibody specificities of anti-OmcA and anti-MtrC were also validated by immunoblot analysis of S. oneidensis whole-cell lysate (28).To determine if MtrC or OmcA was expressed on the external surface of live bacteria when Fe(III) served as the TEA, Ig-RFM was conducted on wild-type versus ΔomcA ΔmtrC double mutant cells. For these experiments, bacteria were cultivated anaerobically with Fe(III), in the form of Fe(III) chelated to nitrilotriacetic acid (NTA), serving as the TEA (19, 23). Growth conditions have been described elsewhere (3, 15) and were based on previous studies (3, 15, 16, 18) that suggest that S. oneidensis MR-1 targets OmcA and MtrC to the cell surface when Fe(III) serves as the TEA.An Asylum Research MFP-3D-BIO AFM or a Digital Instruments Bioscope AFM (16, 17) was used for these experiments. The z-piezoelectric scanners were calibrated as described previously (17). Cells were deposited on a hydrophobic glass coverslip and immersed in imaging buffer (i.e., phosphate-buffered saline [pH 7.4]). The hydrophobic glass coverslips were made as described previously (17) using a self-assembling silane compound called octadecyltrichlorosilane (OTS; Sigma-Aldrich). S. oneidensis MR-1 cells readily adsorbed onto OTS glass coverslips and remained attached to the coverslips during the entire experiment. No lateral cell movement was observed during the experiment, consistent with previous studies that used OTS glass to immobilize bacteria (15, 17, 18, 27).The AFM tip was brought into contact with the surface of a bacterium, and the antibody-functionalized tip was repeatedly brought into and out of contact with the sample, “fishing” for a binding reaction with cytochrome molecules that were exposed on the external cell surface. Binding events were observed upon separating anti-OmcA- or anti-MtrC-functionalized tips from wild-type S. oneidensis MR-1 cells (Fig. ​(Fig.1).1). For the wild-type cells, we observed both nonspecific and specific interactions (Fig. ​(Fig.11).Open in a separate windowFIG. 1.Retraction force curves for anti-MtrC-functionalized tips (A) and anti-OmcA-functionalized tips (B) that are being pulled away from the surface of living ΔomcA ΔmtrC double mutant (gray dotted line) or wild-type (solid black line) S. oneidensis MR-1. These bacteria were adsorbed onto OTS glass coverslips. (C) Retraction curves exhibiting nonspecific binding, specific binding, or no binding between the AFM tip and the cell surface.The distinction between “specific” and “nonspecific” adhesion is made by observing the change in slope of the force curve during the retraction process (26). During specific binding (Fig. ​(Fig.1C),1C), the cantilever is initially relaxed as it is pulled away from the sample. Upon further retraction, the ligand-receptor complex becomes stretched and unravels, resulting in a nonlinear force profile as noted in references 26 and 16. On the other hand, nonspecific adhesion (Fig. ​(Fig.1C)1C) maintains the same slope during the retraction process because only the cantilever flexes (26).Figure ​Figure22 summarizes the frequency or probability of observing a binding event for both anti-OmcA and anti-MtrC tips. Each bar in Fig. ​Fig.22 represents one experiment in which 500 to 1,000 force curves were collected between one AFM tip and two to four live bacterial cells. This figure does not make a distinction between specific and nonspecific binding. It simply shows the frequency of observing an attractive interaction as the antibody-functionalized tip was pulled away from the surface of S. oneidensis MR-1. Binding events occurred with roughly the same frequency when wild-type S. oneidensis MR-1 cells were probed with anti-MtrC-functionalized tips as when they were probed with anti-OmcA-functionalized tips (Fig. ​(Fig.22).Open in a separate windowFIG. 2.Histograms showing the frequency of observing a binding event for anti-MtrC-functionalized (blue) or anti-OmcA-functionalized (red) AFM tips on live wild-type S. oneidensis MR-1 (solid bars) or ΔomcA ΔmtrC double mutant (diagonally hatched bars) cells. The downward arrows designate injection of free antibody into the imaging buffer. The solid gray bars correspond to results obtained with unbaited AFM tips.A number of control experiments were performed to verify the detection of OmcA and MtrC on the surface of wild-type S. oneidensis MR-1. First, 0.1 μM of free anti-OmcA (or anti-MtrC) was added to the imaging fluid to block binding between the antibody-functionalized AFM tip and surface-exposed cytochromes (11, 16). This decreased the adhesion that was observed between the antibody-functionalized tip and the cell surface (Fig. ​(Fig.22).Second, we performed force measurements on ΔomcA ΔmtrC double mutant S. oneidensis MR-1 cells. This mutant is deficient in both OmcA and MtrC (19, 23, 24) but produces other proteins native to the outer surface of S. oneidensis MR-1. The resulting force spectra showed a noticeable reduction in binding events for the ΔomcA ΔmtrC double mutant cells (Fig. ​(Fig.2).2). The binding events that were observed for the double mutant were only nonspecific in nature (Fig. ​(Fig.1).1). This indicates that the antibodies on the tip do not participate in specific interactions with other proteins on the surface of S. oneidensis MR-1 cells.As a final control experiment, force measurements were conducted on wild-type S. oneidensis MR-1 cells, using Si₃N₄ tips conjugated with the PEG linker but not functionalized with polyclonal antibody (unbaited tips). Like the results with the double mutant, the unbaited tips were largely unreactive with the surface of the bacteria (Fig. ​(Fig.2).2). Those binding events that were observed were nonspecific in nature. Taken together, these results demonstrate that the antibody-coated tips have a specific reactivity with OmcA and MtrC molecules. Furthermore, these force measurements show that MtrC and OmcA are present on the external cell surface when Fe(III) serves as the TEA.To map the distribution of cytochromes on living cells, Ig-RFM was conducted on living S. oneidensis MR-1 cells that were growing on a hematite (α-Fe₂O₃) thin film. The conditions for these experiments were as follows. A hematite film was grown on a 10-mm by 10-mm by 1-mm oxide substrate via oxygen plasma-assisted molecular beam epitaxy (14, 16). The cells were grown anaerobically to mid-log phase with Fe(III)-NTA serving as the TEA. Cells were deposited onto the hematite thin film along with anaerobic growth medium that lacked Fe(III)-NTA. The cells were allowed to attach to the hematite surface (without drying) overnight in an anaerobic chamber. The following day, the liquid was carefully removed and immediately replaced with fresh anaerobic solution (pH 7.4). Ig-RFM was performed on the cells by raster scanning an antibody-functionalized AFM tip across the sample surface, thereby creating an affinity map (1). Force curves were collected for a 32-by-32 array. The raw pixilated force-volume data were deconvoluted using a regularized filter algorithm. The total time to acquire a complete image was approximately 20 min.As noted above, attractive interactions between an antibody tip and cell resulted in relatively short-range, nonspecific and longer-range, specific adhesive forces (Fig. ​(Fig.1C).1C). To distinguish between these two interactions, we integrated each force curve beginning at >20 nm and ending at the full retraction of the piezoelectric motor (∼1,800 nm). This integration procedure quantifies the work of binding, measured in joules, between the antibody tip and a particular position on the sample. While this integration procedure does not totally exclude nonspecific binding, it does select for those events associated primarily with specific antibody-antigen binding. Figure ​Figure33 is the antibody-cytochrome recognition images for MtrC and OmcA. The corresponding height (or topography) images of the bacterial cells are also shown in Fig. ​Fig.33.Open in a separate windowFIG. 3.Ig-RFM of live S. oneidensis MR-1 cells deposited on a hematite (α-Fe₂O₃) thin film. Height image (A) and corresponding Ig-RFM image (B) for a bare unfunctionalized Si₃N₄ tip. Height and corresponding Ig-RFM image for a tip functionalized with anti-MtrC (C and D) or anti-OmcA (E and F). Each panel contains a thin white oval showing the approximate location of the bacterium on the hematite surface. A color-coded scale bar is shown on the right (height in micrometers [μm], and the work required to separate the tip from the surface in attojoules [aJ]).OmcA molecules were concentrated at the boundary between the bacterial cell and hematite surface (Fig. 3E and F). MtrC molecules were also detected at the edge of a cell (Fig. 3C and D). Some MtrC, unlike OmcA, was observed on the cell surface distal from the point of contact with the mineral (Fig. 3C and D). Both OmcA and MtrC were also present in an extracellular polymeric substance (EPS) on the hematite surface (Fig. 3D and F), which is consistent with previous results showing MtrC and OmcA in an EPS produced by cells under anaerobic conditions (19, 24). This discovery is interesting in light of the research by Rosso et al. (22) and Bose et al. (4), who found that Shewanella can implement a nonlocal electron transfer strategy to reduce the surface of hematite at locations distant from the point of cell attachment. Rosso et al. (22) proposed that the bacteria utilize unknown extracellular factors to access the most energetically favorable regions of the Fe(III) oxide surface. The Ig-AFM results (Fig. ​(Fig.3)3) suggest the possibility that MtrC and/or OmcA are the “unknown extracellular factors” that are synthesized by Shewanella to reduce crystalline Fe(III) oxides at points distal from the cell. Additional experiments showing reductive dissolution features coinciding with the extracellular location of MtrC and/or OmcA would need to be performed to test this hypothesis.It is important to note that these affinity maps were collected on only a few cells because it so challenging to produce large numbers of quality images. Future work should be conducted on a population of cells. Until this time, these affinity maps can be used to provide a crude, lowest-order estimate of the number of cytochromes on the outer surface of living S. oneidensis MR-1. For example, there were 236 force curves collected on the bacterium shown in Fig. ​Fig.3D.3D. Thirty-eight of these curves exhibited a distinct, sawtooth-shaped, antibody-antigen binding event. In other words, MtrC molecules were detected in one out of every six force curves (16%) that were collected on the cell surface.This probability can be compared to other independent studies that estimated the density and size of MtrC and OmcA molecules from S. oneidensis MR-1. Lower et al. (16) estimated that S. oneidensis has 4 × 10¹⁵ to 7 × 10¹⁵ cytochromes per square meter by comparing AFM measurements for whole cells to force curves on purified MtrC and OmcA molecules. Wigginton et al. (25) used scanning tunneling microscopy to determine that the diameter of an individual cytochrome is 5 to 8 nm. These values can be used to create a simple, geometric, close-packing arrangement of MtrC or OmcA molecules on a surface. Using this approach, cytochromes could occupy 8 to 34% of the cell surface.This estimate is consistent with the observed number of putative MtrC molecules shown in Fig. ​Fig.3D.3D. Therefore, it appears that these affinity maps can be used as a lowest-order estimate for the number of cytochromes on S. oneidensis MR-1 even though we do not know a priori the exact configuration of the antibody tip (e.g., the concentration of antibody on the tip, the exact shape of the tip, the binding epitopes within the antibody).In summary, the data presented here show that S. oneidensis MR-1 localizes OmcA and MtrC molecules to the exterior cell surface, including an EPS, when Fe(III) is the TEA. Here, the cytochromes presumably serve as terminal reductases that catalyze the reduction of Fe(III) through direct contact with the extracellular iron-oxide mineral.