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Journal of Bacteriology, January 2007, p. 169-178, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00792-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

ß-D-Allose Inhibits Fruiting Body Formation and Sporulation in Myxococcus xanthus{triangledown}

Marielena Chavira,1 Nga Cao,1 Karen Le,1 Tanveer Riar,1 Navid Moradshahi,1 Melinda McBride,1 Renate Lux,2 and Wenyuan Shi1,2*

Department of Microbiology, Immunology and Molecular Genetics,1 School of Dentistry, University of California, Los Angeles, California2

Received 1 June 2006/ Accepted 3 October 2006


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ABSTRACT
 
Myxococcus xanthus, a gram-negative soil bacterium, responds to amino acid starvation by entering a process of multicellular development which culminates in the assembly of spore-filled fruiting bodies. Previous studies utilizing developmental inhibitors (such as methionine, lysine, or threonine) have revealed important clues about the mechanisms involved in fruiting body formation. We used Biolog phenotype microarrays to screen 384 chemicals for complete inhibition of fruiting body development in M. xanthus. Here, we report the identification of a novel inhibitor of fruiting body formation and sporulation, ß-D-allose. ß-D-Allose, a rare sugar, is a member of the aldohexose family and a C3 epimer of glucose. Our studies show that ß-D-allose does not affect cell growth, viability, agglutination, or motility. However, ß-galactosidase reporters demonstrate that genes activated between 4 and 14 h of development show significantly lower expression levels in the presence of ß-D-allose. Furthermore, inhibition of fruiting body formation occurs only when ß-D-allose is added to submerged cultures before 12 h of development. In competition studies, high concentrations of galactose and xylose antagonize the nonfruiting response to ß-D-allose, while glucose is capable of partial antagonism. Finally, a magellan-4 transposon mutagenesis screen identified glcK, a putative glucokinase gene, required for ß-D-allose-mediated inhibition of fruiting body formation. Subsequent glucokinase activity assays of the glcK mutant further supported the role of this protein in glucose phosphorylation.


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INTRODUCTION
 
Although Myxococcus xanthus exists as individual vegetative rods, the myxobacteria are fundamentally social organisms capable of intercellular communication, coordinated group movements, and the formation of multicellular structures. Upon starvation, a developmental program is initiated wherein bacterial cells aggregate into dome-shaped, macroscopic structures called fruiting bodies. The signaling and genetic systems orchestrating this program have received much attention in M. xanthus research (21, 22, 47). In particular, investigation of exogenous compounds that inhibit fruiting body formation have provided useful starting points for identifying the signaling pathways and molecular components involved in this highly regulated process. Previous studies investigating inhibition of fruiting body formation by various amino acids have shown that methionine-mediated inhibition is associated with a dramatic reduction in the production of S-adenosylmethionine (SAM) (6, 42). SAM, a methyl donor for the M. xanthus frz chemosensory system, affects fruiting through the regulation of motility (5, 45). Lysine, threonine, leucine, and isoleucine were also found to inhibit fruiting body formation through a reduction of intracellular levels of SAM. A separate study found that glycine can block purine-induced fruiting body formation, suggesting that purine-containing compounds, such as cyclic AMP and ADP, induce fruiting through a single mechanism (38).

To identify additional compounds that completely inhibit fruiting body formation, we utilized Biolog phenotype microarrays (PMs) to conduct a high-throughput screen of 384 substrates. The PMs are 96-well microtiter plates that include various carbon-, nitrogen-, sulfur-, and phosphorus-containing compounds. Through this screen, we identified ß-D-allose, a novel inhibitor of M. xanthus morphogenesis. ß-D-Allose, a C3 isomer of glucose, is a rare sugar found predominantly in plant polysaccharides (8, 13). In Aerobacter aerogenes and Escherichia coli, D-allose is incorporated into the glycolytic pathway through conversion to D-fructose-6-phosphate (14, 26). ß-D-Allose has also become the focus of several biomedical studies which show that it has anti-inflammatory and antiproliferative activity in eukaryotic models, such as Lewis rats and human ovarian carcinoma cells (19, 20, 49, 50). While these and other studies demonstrate strong biological activity, very little is currently known about ß-D-allose activity at the molecular level. To explore the possible mechanisms of ß-D-allose-mediated fruiting inhibition, we examined its effects on growth, viability, agglutination, sporulation, motility, and developmental gene expression. In addition, we investigated the ability of various aldohexose sugars to rescue fruiting in the presence of ß-D-allose. Finally, through a transposon-mediated mutagenesis screen, we identified, glcK, a putative glucokinase required for ß-D-allose sensitivity. Although ß-D-allose is not abundant in nature, it is clear that it has strong biological properties which may prove useful for guiding our attention to new pathways or molecular mechanisms central to the regulation of fruiting body formation.


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MATERIALS AND METHODS
 
Bacterial strains and culture conditions. All M. xanthus strains used in this study are listed in Table 1. Cells were grown in CYE (10 g Casitone per liter, 5 g yeast extract per liter, 8 mM MgSO4 in 10 mM morpholinepropanesulfonic acid [MOPS] buffer, pH 7.6) at 32°C on a rotary shaker at 225 rpm. Kanamycin (40 µg/ml) was added for growth of mutant strains where indicated.


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  		TABLE 1. M. xanthus strains used in this study

Biolog phenotype microarray screen. The content and construction of the phenotype microarrays have been described previously (3). Additionally, the specific chemical compositions of PM1 to PM4 can be found at http://www.biolog.com/pmMicrobialCells.html (although information regarding the chemical concentrations in the arrays is proprietary). M. xanthus strain DK1622 was grown to log phase (2 x 108 to 3 x 108 cells/ml), washed in MOPS-calcium-magnesium (MCM) buffer (10 mM MOPS, pH 6.8, 4 mM MgSO4, 1 mM CaCl2), and resuspended to a concentration of 2 x 108 cells/ml. The cell suspension was then added to the PM arrays using a multichannel pipettor (50 µl/well). Next, tetrazolium violet, a colorimetric indicator dye that turns dark purple in response to cellular respiration, was dissolved in MCM buffer and added to each well (50 µl/well), generating a final concentration of 20 µM tetrazolium violet and a final cell concentration of 1 x 108 cells/ml per well. To optimize the tetrazolium violet conditions for the PM screen, a range of dye concentrations from 0 to 100 µM was tested to ensure adequate color change without disturbing the process of fruiting body formation. The low concentration of 20 µM tetrazolium violet was found to be optimal for this application. The plates were sealed in a plastic bag along with a moist cloth to maintain hydration and incubated at 32°C for 3 days. The plates were scored visually with a Leica DMIL inverted light microscope. Images were taken using SPOT digital camera/software (Diagnostic Instruments, Inc.).

Determination of ß-D-allose MICs in submerged culture and on MOPS plates. To determine MIC, submerged cultures were prepared as follows. DK1622 cultures were grown to log phase, washed twice in MCM buffer, resuspended to a concentration of 1.5 x 108 cells/ml in MCM buffer, and aliquoted (120 µl/well) into 96-well, polystyrene, flat-bottom plates. ß-D-Allose, dissolved in MCM buffer, was added (30 µl/well) to final concentrations of 0 to 5 mM ß-D-allose per well. Cultures were then incubated at 32°C for 24 h. For MIC determination on MOPS agar plates (MOPS buffer solidified with 1.5% Bacto agar), DK1622 cultures were grown to log phase, washed twice in MCM buffer, and resuspended in MCM buffer to a concentration of 5 x 109 cells/ml. Cells were then spotted (7 µl) onto MOPS agar plates containing 0 to 5 mM ß-D-allose (ß-D-allose was added to molten agar after autoclaving and cooling to 60°C). MOPS plates were incubated at 32°C for 24 h. Images of fruiting bodies in submerged culture and on MOPS plates were taken with a Leica DMIL inverted light microscope and SPOT camera/software (Diagnostic Instruments, Inc.).

Agglutination, swarm plate, and sporulation assays. A slightly modified version of a previously described agglutination assay was utilized (48). In brief, DK1622 cells were grown to log phase, washed once with MCM buffer, and resuspended in MCM to a density of 3 x 108 cells/ml. Absorbance at 600 nm was measured every 10 min for 1.5 h using a Biomate 3 Thermo Spectronic spectrophotometer. Swarm plates (CYE medium solidified with 0.3% Bacto agar) were inoculated as described previously (46). Plates were incubated at 32°C for 2 days before colony diameters were measured or colony edges were imaged. Starvation-induced sporulation was carried out in submerged culture while glycerol-induced sporulation was carried out according to Licking et al. (36). DK1622 culture was grown to log phase, divided into two cultures, and incubated with 0 or 5 mM ß-D-allose for 1 h. Cultures were then incubated in 0.5 M glycerol for 4 h at 32°C followed by a 2-h incubation at 55°C. Aliquots taken immediately after glycerol addition and after 55°C incubation were serially diluted and plated to obtain CFU values. Aliquots from submerged and glycerol-containing cultures were also assessed for sporulation microscopically at x400 (total magnification) with a Leica DMLS light microscope.

Reversal frequency, speed, and percent motility analysis. To determine reversal frequency, speed, and percent motility, 1 x 105 exponentially growing cells (about 1 µl) were diluted 10-fold with MCM and then spotted into 250 µl of 1% methylcellulose in a 24-well polysterene plate or on 1.5% MOPS plates containing 0 or 5 mM ß-D-allose. Cell movements were viewed with a Nikon Eclipse TE200 inverted microscope at x40 and recorded with a Panasonic AG-6040 time-lapse video cassette recorder at x35 and at 1/60 real-time speed. The recordings were played at real time for data analysis. Reversal frequencies were calculated by measuring the amount of time (min) per reversal by hand. For each condition, 20 different cells were measured 10 times each. Speed was calculated by measuring the amount of time (min) it took for a cell to travel a straight distance from an arbitrarily chosen starting point to an ending point. This value was converted to µm/min and then divided by the total magnification factor (x1,400). Twenty speed measurements were calculated for each condition. Motile percentage was calculated by estimating the number of motile versus nonmotile cells in a given screen shot per sample.

Measurement of ß-galactosidase expression from Tn5lac strains. The measurement of ß-galactosidase specific activity was adapted from protocols described by Kroos et al. (31). Tn5lac strains were grown to log phase in 100-ml cultures of CYE plus kanamycin, washed, and resuspended in 7 ml MCM buffer. A 100-µl aliquot was added to 250 µl sodium phosphate buffer (50 mM, pH 7.2) and stored at –80°C for later determination of ß-galactosidase activity during growth (t = 0 sample). For developmental samples, five 20-µl aliquots were spotted onto 1.5% agar MOPS plates, supplemented with 0 or 5 mM ß-D-allose. Plates were prepared 2 to 3 days beforehand by allowing autoclaved agar to cool to 60°C before the addition of ß-D-allose. Plates were then dried and stored on the bench at room temperature until use. After spotting, plates were incubated at 32°C for 2, 4, 6, 8, 12, 24, 36, and 48 h. Cells were harvested by scraping into microcentrifuge tubes containing 250 µl sodium phosphate buffer. Samples were vortexed and stored at –80°C until all samples were collected.

Next, cells were thawed, lysed with 4 µl toluene, vortexed for 30 s, and incubated at 37°C for 5 min. Samples were then placed in the hood with open caps for 10 min to allow toluene evaporation and centrifuged for 15 min at maximum speed (Eppendorf centrifuge 5415D). The supernatant layer was transferred to a fresh tube and stored on ice. To measure ß-galactosidase activity, 10 µl of 4 mg/ml o-nitrophenyl-ß-D-galactoside (ONPG) was added to 100 µl supernatant and incubated at room temperature until development of yellow color (or for 2 h). The reactions were stopped with 140 µl of 1 M NaCO3. A blank sample, containing 100 µl sodium phosphate buffer, was processed along with the other samples. From each sample, a 200-µl aliquot was transferred to a 96-well microtiter plate and the absorbance was measured at 420 nm using the Benchmark Plus microplate reader (Bio-Rad).

Protein concentrations were determined using the Coomassie Plus protein assay (Pierce), by diluting 2 to 15 µl supernatant with double-distilled H2O (ddH2O) to 150 µl and then adding 150 µl prewarmed Coomassie Plus reagent. Absorbance readings were measured at 595 nm, and protein concentrations were extrapolated from a bovine serum albumin standard curve. ß-Galactosidase specific activities were calculated as nanomoles ONP produced per minute per milligram protein as previously described (31).

Development on MOPS plates containing ß-D-allose and various hexose sugars. Fruiting experiments were performed on MOPS agar plates containing 50 mM arabinose, ribose, glucose, galactose, or xylose in combination with 0 or 3 mM ß-D-allose. Plates were prepared 1 day prior to use by autoclaving MOPS agar and allowing agar to cool to 60°C before adding the sugars. DK1622 cultures were prepared, spotted onto MOPS plates, and imaged as described for MIC determination experiments.

magellan-4 mutagenesis screen. To prepare electrocompetent cells, a 10-ml culture of DK1622 was grown to log phase, collected by centrifugation, washed three times in ddH2O, and resuspended in 1 ml ddH2O. An 80-µl aliquot of washed cells was combined with the plasmid, pminiHimar1, which was isolated using the Wizard SV gel and PCR cleanup system (Promega). The plasmid, pminiHimar1, donor of the magellan-4 element, has been described previously (43). Electroporations were performed in the Gene Pulser II (Bio-Rad) with settings at 0.65 kV, 400 {Omega}, and 25 µF, resulting in typical time constants of 9.4 ms. Samples were immediately flushed and recovered in 1 ml CYE and then incubated at 32°C for 6 h on a rotary shaker at 225 rpm. Cells were plated onto 1.5% CF agar plates supplemented with kanamycin and 5 mM ß-D-allose (17). After 14 to 18 days, plates were screened visually to identify colonies with fruiting body development, which indicated a loss of sensitivity to ß-D-allose. A total of 50 mutants were isolated from approximately 26,400 colonies screened. The mutants were purified and retested for resistance to ß-D-allose. Of these, 26/50 mutants demonstrated stable phenotypes. The magellan-4 insertions in these strains were designated mas1 (mutation in allose sensitive) through mas26. To confirm that the mutant phenotypes resulted from single insertions, strains containing mas insertions were backcrossed by electroporation of the mutant genomic DNA into the parent strain, DK1622 (53). After backcrossing, 4/26 mutants demonstrated stable phenotypes and were chosen for sequence analysis.

Cloning and sequencing the magellan-4 insertion sites. To subclone genomic regions neighboring the magellan-4 transposon, genomic DNA was isolated from vegetative cultures of DK1622 mas::magellan-4 strains. Genomic DNA was digested with SacI for 12 to 16 h in a total volume of 20 µl, heat inactivated for 15 min at 65°C, and cleaned with the Wizard SV gel and PCR cleanup system (Promega). Digested DNA samples (12.5 µl) were then ligated into a cloning vector (1 µl) via QuickLigase (New England BioLabs) in a total volume of 30 µl before transformation into the E. coli host DH5{alpha}-{gamma}pir. Approximately 1 to 20 Kmr transformants were obtained per transformation and were recovered on LB plates supplemented with kanamycin after incubation at 37°C for 24 h. Plasmids were isolated (QIAGEN Miniprep kit) and sequenced with primers corresponding to the region immediately downstream of the left inverted repeat (pminiHimar1 5' end, 5' CATTTAATACTAGCGACGCCATCT 3') and immediately upstream of the right inverted repeat (pminiHimar1-lacZ 3' end, 5'GAACTATGTTGAATAATAAAAACGA 3'). Sequencing reactions were analyzed with an ABI 37300 DNA analyzer. Mutants with mas insertions in known genes were designated MC0001 to MC0004 (Table 1).

Glucokinase activity assay. Lysates were prepared as previously described (58), with minor modifications. Strains DK1622 and MC0001 were grown to log phase in 50-ml cultures, collected by centrifugation, washed in 1/2 volume MCM buffer, then resuspended in 1/50 volume cold MCM. Cultures were placed on ice and sonicated for 45 s (in 5-s bursts) at 5 W in a model 60 Fisher sonic dismembranator. Following microcentrifugation at 4°C to remove cellular debris (20,000 x g for 10 min), lysates were kept on ice and assayed within 4 h. Glucokinase activities were determined using a colorimetric reaction, which couples the production of glucose-6-phosphate to the reduction of NADP (11, 16). Our standard reaction mixture (1-ml total volume) contained 50 mM Tris (pH 9.1), 20 mM MgCl2, 20 mM glucose, 10 mM ATP (pH 9.1), 10 to 100 µl cell lysate, and 1 U of Leuconostoc mesenteroides glucose-6-phosphate dehydrogenase (Sigma). Reactions were initiated by the addition of NADP (Sigma) to 1 mM, and the rate of NADP reduction was measured as the change in absorbance at 340 nm, in 30-s intervals, over 15 min at 25°C ({varepsilon}NADPH = 6.22 x 106 cm–2). Absorbance readings were measured with a Spectronic Genesys 5 spectrophotometer. Lysate protein concentrations were determined by the Coomassie Plus protein assay (as described above). Specific activities were calculated according to the equation reported by Goward et al. (16).


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RESULTS
 
Identification of ß-D-allose—a novel inhibitor of fruiting body formation. Biolog phenotype microarrays were used to screen for compounds that inhibit fruiting body formation. A diverse set of 384 compounds were included in the Biolog PM screen. PM1 and PM2 contained carbon-based substrates, PM3 contained nitrogen-based compounds and PM4 contained phosphorous and sulfur-based compounds. In addition, to distinguish between metabolically active and inactive cells, tetrazolium violet, a colorimetric indicator dye which functions by shifting from colorless to an irreversible dark purple upon reduction by NADH (Fig. 1A), was included in every well (2). We therefore screened for wells showing a dark purple coloration in addition to a lack of fruiting body development (Fig. 1B).


Figure 1
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  		FIG. 1. Identification of ß-D-allose using Biolog phenotype microarrays. PM microtiter plates containing 20 µM tetrazolium violet were inoculated with DK1622 cell suspensions, and incubated at 32°C for 24 h. (A) Control well with normal fruiting body formation. (B) Test well containing ß-D-allose with complete nonfruiting. Images were captured after 24 h and are representative of phenotypes observed in two, independent PM screen experiments. Total magnification = x40.

The majority of wells (96%) displayed purple coloration and some form of altered fruiting body morphology. For example, wells containing D-galactonic acid-{gamma}-lactone produced clustered mounds, phospho-L-arginine produced small fruiting bodies, and Gly-Gln produced enlarged fruiting bodies. In the remaining wells, which contained chemicals such as butyric acid, Tween 40, and acetone, the cells appeared clear or light yellow, with no visible fruiting body formation. Only eight compounds produced complete inhibition of fruiting along with a dark purple coloration: adenine, ß-D-allose, 2'-deoxyadenosine, DL-ethionine, L-homoserine, L-methionine, L-phenylalanine, and L-tyrosine. These results agree with previous reports: DL-ethionine, L-homoserine, L-methionine, L-phenylalanine, and L-tyrosine have been shown to inhibit fruiting body formation (45). In contrast, ß-D-allose had never been studied in the context of fruiting body formation. Furthermore, ß-D-allose is unusual in that it is the only monosaccharide in the set of chemicals identified; the other inhibitors are amino acids or modified nucleotides. ß-D-Allose, therefore, emerged as a novel inhibitor and was selected for further study.

Following identification of ß-D-allose in the chemical screen, we determined the MICs of ß-D-allose in submerged culture and on MOPS plates. In submerged culture conditions, complete inhibition of fruiting occurs at 3 mM ß-D-allose, while on MOPS plates inhibition occurs at 2 mM ß-D-allose (Fig. 2). It is interesting to note that on MOPS plates as well as in submerged culture, there is a concentration-sensitive change from a fruiting to a nonfruiting phenotype over a 1 mM increase in ß-D-allose: on MOPS plates, fruiting occurs at 1 mM but is blocked at 2 mM ß-D-allose. A similarly abrupt change in phenotype is seen from 2 mM to 3 mM ß-D-allose in submerged culture. This suggests that ß-D-allose must reach a threshold concentration in order for inhibition of development to take place.


Figure 2
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  		FIG. 2. ß-D-Allose MICs on MOPS plates and in submerged culture. DK1622 cells were washed and spotted onto 1.5% agar MOPS plates or added to submerged cultures containing final concentrations of 0 to 5 mM ß-D-allose. Plates were incubated at 32°C and photographed 24 h later. The total magnifications are x20 for MOPS plates and x40 for submerged cultures. Images are representative of three independent experiments.

Effects of ß-D-allose on M. xanthus physiology. In our initial attempts to explore the mechanism of ß-D-allose fruiting inhibition, we examined the effects of ß-D-allose on basic M. xanthus physiology including growth rate, viability, and sporulation. To assess effects on growth rate, turbidity measurements were taken over a period of 30 h from DK1622 liquid cultures prepared with 0, 1, 3, 4, and 5 mM ß-D-allose. For all the concentrations tested, ß-D-allose did not significantly affect the vegetative growth rate (data not shown). Next, we assessed the effects of ß-D-allose on cell viability. Although toxicity had been assessed by the inclusion of tetrazolium violet in the original PM screen, demonstrating nontoxicity under more rigorous conditions would be particularly important for establishing the reliability of specific data sets (such as the Tn5lac data below). Therefore, wild-type cells were washed and resuspended in MCM buffer in the presence of 0, 2, and 5 mM ß-D-allose and then incubated for 32 h. Aliquots of the three cultures were serially diluted and plated in duplicate, and CFU were determined. Again, our results demonstrated that ß-D-allose does not significantly change viability under these conditions (data not shown).

Interestingly, we found that 5 mM ß-D-allose inhibits both starvation-induced and glycerol-induced sporulation. To further examine ß-D-allose effects on starvation-induced sporulation, submerged cultures were prepared with 0 or 5 mM ß-D-allose. Following incubation for 24 to 72 h, samples were viewed microscopically. Cultures prepared with ß-D-allose were found to have fewer spores overall, and these spores had a nonrefractile, irregular-shaped appearance (data not shown). To examine ß-D-allose effects on glycerol-induced sporulation, cultures incubated with 0 or 5 mM ß-D-allose were treated with glycerol and then incubated at 55°C for 2 h. Colony counts revealed that ß-D-allose caused a 98 to 100% reduction in glycerol induced sporulation (data not shown).

ß-D-Allose does not affect motility. Finding no significant physiological changes in response to ß-D-allose, we proceeded to investigate whether the sugar might affect M. xanthus motility. We first examined this question by testing whether ß-D-allose alters pilus-exopolysaccharide (EPS) interactions. Agglutination requires effective pilus-EPS binding; therefore, agglutination studies were conducted in the presence of 0 or 5 mM ß-D-allose (8, 35). No differences were detected in the rate of agglutination, indicating that pilus-EPS interactions were unaffected by ß-D-allose (data not shown). Next we focused on the two M. xanthus motility systems: adventurous motility (A-motility) and social motility (S-motility). The ability of M. xanthus to utilize and coordinate its A- and S-motility systems is central to its ability to carry out morphogenesis (34, 51). We assessed the effects of ß-D-allose on A- and S-motility by observing DK122 swarming behavior on agar plates: 0.3% agar plates favor S-motility, and 1.5% agar plates favor A-motility (46). On both 1.5% and 0.3% plates, the pattern of cell movement across the agar as seen at the colony edges did not change in the presence of 5 mM ß-D-allose (Fig. 3). Likewise, a comparison of colony diameters on 0.3% CYE plates containing 0 or 5 mM ß-D-allose revealed no significant differences (Table 2). Although A- or S- motility can be encouraged by the use of 0.3% or 1.5% plates, the two systems ultimately work in a coordinated fashion, which makes it difficult to assess effects on each motility system independently (34). In order to uncouple A- and S-motility and view the effects of ß-D-allose on each system individually, we employed two S-motility mutants, DK1253 and DK1300 and two A-motility mutants, DK1217 and DK1218, in the swarm plate experiment. Predictably, the colony diameters of the mutant strains grown with and without ß-D-allose were not significantly different from each other (Table 2). It would seem that neither A- nor S-motility is affected by ß-D-allose.


Figure 3
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  		FIG. 3. Effect of ß-D-allose on A- and S-motility. DK1622 cells were grown to log phase, washed, and spotted onto 1.5% and 0.3% CYE plates with or without 5 mM ß-D-allose. Colony edge images were taken after 2 days of incubation at 32°C. Total magnification, x200. Images are representative of three data sets.


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  		TABLE 2. Effect of D-allose on colony diameter

We continued our studies by examining motility at the cellular level via time-lapse video microscopy. As with A- and S-motility, ß-D-allose does not appear to affect reversal frequency. We found no significant difference between reversal frequencies in the presence of 0 or 5 mM ß-D-allose on 1.5% agar (10.6 ± 3.3 and 10.3 ± 3.7 reversals/h) and in 1% methylcellulose (10.1 ± 2.7 and 13.7 ± 5.3 reversals/h) (Table 3). Similarly, this hexose had no significant effect on speed. Wild-type cells traversed 1.5% agar containing 0 or 5 mM ß-D-allose at 2.2 ± 0.7 and 1.5 ± 1.0 µm/min and in 1% methylcellulose at speeds of 6.6 ± 2.2 and 4.3 ± 2.0 µm/min (Table 3). The final measurement, percent motility, was calculated by estimating motile versus nonmotile cells in a given screen shot of a video recording. On 1.5% MOPS, the percent motilities were 95% and 94% for 0 and 5 mM ß-D-allose, respectively, while motilities in 1% methylcellulose were 89% and 93%. Taken together, these data demonstrate that ß-D-allose does not affect pilus-exopolysaccharide interactions, A- or S-motility, reversal frequency, or speed of individual cell movements.


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  		TABLE 3. Effect of ß-D-allose on individual cell movements

Mid-developmental genes show reduced expression levels in the presence of ß-D-allose. We proceeded to test whether ß-D-allose might be affecting developmental gene expression by utilizing Tn5lac reporter gene fusions (31). Strains containing these gene fusions are known to express ß-galactosidase at specific stages in development and have been utilized to explore how developmental gene expression is coordinated by extracellular signaling events (29-32). An orderly progression of specific, extracellular signals (designated A to E) is required for the construction of a spore-filled fruiting body (9, 22). In order to assess developmentally regulated gene expression over a broad range of times, we selected six Tn5lac reporter strains: DK4491, DK4300, DK4521, DK4531, DK4414, and DK4500 (Table 1). Strains DK4491, DK4300, and DK4521 contain insertions in early developmental genes and have been shown to initiate ß-galactosidase expression between 0 and 3 h of development (9, 25, 31, 47). Strain DK4300 carries an insertion in the sdeK locus which is induced in an A-signal-independent manner, whereas induction of 4521 requires the A-signal (25, 52). As development continues, strains DK4531, DK4414, and DK4500 express ß-galactosidase at approximately 4 to 7, 6 to 13, and 13 to 14 h, respectively (9, 31, 52). The DK4414 strain carries an insertion in the devRS locus, which requires C signaling and is induced during the aggregation phase of fruiting body formation (52).

To assess the effect of ß-D-allose on developmental gene expression, cells from the Tn5lac strains were spotted on MOPS agar plates supplemented with 0 or 5 mM ß-D-allose, incubated at 32°C, and then collected and analyzed at 0, 2, 4, 6, 8, 12, 24, 36, and 48 h of development (see Materials and Methods). A comparison of ß-galactosidase-specific activities revealed that expression of all genes induced in early development (0 to2 h) was not significantly affected by ß-D-allose (Fig. 4A to B). The Tn5lac strain DK4521, which expresses ß-galactosidase between 1.5 and 3 h of development, demonstrated an intermediate response to ß-D-allose (Fig. 4C). Intriguingly, we found that all genes induced later in development (4 to 14 h) showed significantly lower levels of gene expression in the presence of ß-D-allose (Fig. 4D to F).


Figure 4
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  		FIG. 4. Effect of ß-D-allose on developmental gene expression. Six strains containing Tn5lac reporter insertions were used to compare expression of time-dependent developmental genes, as measured by ß-galactosidase specific activity, in the absence (squares) or presence (circles) of 5 mM ß-D-allose. (A and B) ß-Galactosidase is induced in strains DK4300 and DK4491 between 0 and 2 h of development, (C) in strain DK4521 between 1.5 and 3 h, (D) in DK4531 between 4 and 7 h, (E) in DK4414 between 6 and 13 h, (F) and in strain DK4500 between 13 and 14 h. These data represent the average of three independent trials.

The effect of ß-D-allose on specific stages of fruiting body development was further examined by studies in which the hexose sugar was added to wild-type cells in submerged culture (5 mM final concentration) at 0, 5, 8, 12, 24, and 48 h of development (Fig. 5). These studies revealed that inhibition of fruiting body formation occurred only when ß-D-allose was added before 8 h of development. When ß-D-allose was added at 12 h of development or later, fruiting body formation proceeded to completion. These results, along with the Tn5lac reporter studies, demonstrate that the time period before 12 h of development—in particular the 4- to 12-h time window—is critical for ß-D-allose-mediated fruiting inhibition.


Figure 5
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  		FIG. 5. Addition of ß-D-allose at various times in development. Strain DK1622 was grown to log phase, washed in MCM buffer and aliquotted into 96-well plates for submerged culture. At 0, 5, 8, 12, 24, and 48 h, ß-D-allose was added to a final concentration of 5 mM. (A) Images were taken immediately after addition of ß-D-allose to 5 mM and (B) again after 72 h of development. Total magnification, x40.

Galactose and xylose rescue development in the presence of ß-D-allose. In addition to revealing the inhibitory activity of ß-D-allose, the PM arrays demonstrated that this inhibitory activity is unusual; other sugars, such as arabinose, ribose, glucose, galactose, and xylose did not inhibit fruiting body formation in the arrays (data not shown). However, we were interested in determining whether or not these sugars were capable of antagonizing ß-D-allose-mediated inhibition. First, to confirm our observations from the PM arrays, we added these sugars to submerged cultures and found that they were incapable of developmental inhibition—even up to 50 mM concentrations (Fig. 6). The same set of sugars, at 50 mM, was tested for the ability to restore fruiting in the presence of 3 mM ß-D-allose. Ribose and arabinose were unable to restore development, while glucose had a partial rescue effect. Interestingly, galactose and xylose demonstrated the ability to fully rescue fruiting in the presence of ß-D-allose (Fig. 6).


Figure 6
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  		FIG. 6. Effect of hexose sugars on ß-D-allose-mediated fruiting inhibition. MOPS plates were prepared with 50 mM arabinose, ribose, glucose, galactose, or xylose in combination with 0 or 3 mM ß-D-allose (BDA). DK1622 cells were grown to log phase, washed, spotted onto plates, and incubated at 32°C for 24 h. Total magnification, x20. Images are representative of four independent trials.

glcK is required for BDA-mediated inhibition of fruiting body formation. The Himar1 transposon, isolated from the horn fly Haematobia irritans, is part of the mariner superfamily of mobile genetic elements (43). Because mariner elements do not require host cofactors, they have been successfully transposed in prokaryotes such as E. coli, mycobacteria, and M. xanthus via transposase-containing suicide vectors (43, 56, 57). In order to generate and identify ß-D-allose-resistant mutants in M. xanthus, we utilized a Himar1-derived mini-element, magellan-4, to conduct a genomewide mutagenesis screen (56). Following transposon-mediated mutagenesis, mutants capable of fruiting body development in the presence of 5 mM ß-D-allose were selected for further study (Fig. 7).


Figure 7
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  		FIG. 7. Fruiting body formation in the presence of ß-D-allose in a glcK mutant. Strains DK1622 (wild type) and MC0001 ({Delta}glcK) were grown to log phase, washed, spotted onto MOPS plates with 0 or 5 mM ß-D-allose, and incubated at 32°C. Images were captured after 48 h. Total magnification, x20. Images are representative of four trials.

Among the 26,400 colonies screened, 26 mutants demonstrated the ability to form fruiting bodies in the presence of 5 mM ß-D-allose. These mutants were backcrossed to the parental strain, DK1622, via electroporation of mutant gDNA, to ensure that the ß-D-allose-resistant phenotype resulted from single, specific mutations. Of these, four mutants demonstrated stable phenotypes. We then determined the DNA sequences at the insertion sites by cloning and sequencing the transposon edge and flanking DNA regions. The four sequences were analyzed using BLAST (1, 15) and the nearly completed TIGR genome sequence of M. xanthus (http://www.tigr.org/tdb/mdb/mdbinprogress.html). All four sequences showed 100% sequence identity to the open reading frame MXAN6497. The predicted protein sequence of MXAN6497 was then aligned against the NCBI database, revealing 53% sequence identity and 65% similarity to the Anaeromyxobacter dehalogenans glucokinase (glcK).

Glucokinase specific activity is decreased in a glcK mutant. To test glucokinase specific activities, we prepared cell lysates from DK1622 (wild type) and MC0001 ({Delta}glcK) cells and then added the lysates to a colorimetric reaction mixture in which glucokinase activity is measured as the rate of reduction of NADP coupled with the oxidation of glucose-6-phosphate by glucose-6-phosphate dehydrogenase (11, 16). Our results show that DK1622 lysates contain a soluble glucokinase activity which increases as a linear function of protein concentration (Fig. 8A). Control reactions show that this activity is both ATP and glucose dependent (Fig. 8A). Lysates from DK1622 had an average specific activity of 12.4 (nmol x min–1 x mg–1), while MC0001 lysates showed average activities of 0.2 (nmol x min–1 x mg–1) with standard deviations of ± 1.1 and ± 0.02 (nmol x min–1 x mg–1), respectively (Fig. 8B). These results agree with findings from a previous study which reported glucokinase activities of 9.2 nmol x min–1 x mg–1 for the wild-type strain and 1.3 (nmol x min–1 x mg–1) in a mutant with a disruption in a putative hexokinase gene (58). These results further support the role of GlcK in glucose phosphorylation and provide additional evidence that ß-D-allose-mediated inhibition of fruiting body formation is dependent upon glucokinase activity.


Figure 8
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  		FIG. 8. Glucokinase activities in M. xanthus wild-type and glcK mutant lysates. (A) Glucokinase activities are plotted as a function of total protein in cell lysates prepared from strain DK1622 (wild type). Lysates were added to reaction mixtures containing 50 mM Tris (pH 9.1), 20 mM MgCl2, 20 mM glucose, 10 mM ATP (pH 9.1), 1 glucose-6-phosphate dehydrogenase, and 1 mM NADP (black circles). Reactions were also performed without glucose (gray squares) and without ATP (white triangles). These data represent an average of three independent trials. (B) A comparison of glucokinase activities in lysates prepared from strains DK1622 (wild type) and MC0001 ({Delta}glcK). These data are an average of four independent determinations.


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DISCUSSION
 
Biolog phenotype microarrays were designed to test for metabolic activity in the presence of various substrates distributed in a set of 96-well plates. In a slight twist on their original purpose, we used the PMs to screen for chemicals capable of inhibiting fruiting body formation. The identification of five known inhibitors, validated this approach while the identification of ß-D-allose, a novel inhibitor of fruiting body formation, led to more detailed experiments (6, 38, 45). In the first phase of this study, we found that ß-D-allose had no significant effect on vegetative growth, viability, or agglutination. Likewise, our motility studies revealed that A- and S-motility (Fig. 3) as well as speed and reversal frequencies (Table 3) were unaffected by ß-D-allose.

Although ß-D-allose does not appear to modulate growth, cytotoxicity, agglutination, or motility, our data suggests that the hexose sugar affects developmental gene expression. Our experiments utilizing Tn5lac reporter strains revealed that developmental genes induced between 4 and 14 h of development show significantly lower levels of gene expression in the presence of ß-D-allose (Fig. 4D to F). These data agreed with the results obtained when ß-D-allose was added to submerged cultures at various time points. The time point study demonstrated that ß-D-allose only blocks development if added before 12 h, thus confirming that the 4- to 12-h time window is crucial for ß-D-allose inhibitory activity (Fig. 5). Considering the fact that aggregation is known to begin approximately 6 to 8 h after starvation (9), our results from the reporter fusions and time-point studies correlate well with the observation that cells exposed to ß-D-allose are blocked from even simple aggregate formation.

If gene expression levels are reduced in response to ß-D-allose between 4 and 14 h of development, then what specific, extracellular signals might be affected? The earliest signal brought about by nutrient limitation is the transient increase of intracellular guanosine penta- and tetraphosphate ([p]ppGpp) (37). Another early signal, the A-signal, is a group of small peptides and amino acids which serve as a cell density detector (33). Expression of ß-galactosidase in strain DK4521 is dependent upon both increasing [p]ppGpp levels and the quorum-sensing A-signal (25, 32). There is also evidence of an early developmental, A-signal-independent pathway required for normal fruiting body formation. The sdeK locus, in strain DK4300, is activated by high [p]ppGpp levels, but does not require the A-signal (52). Our results show that DK4521 reporter expression is moderately decreased in response to ß-D-allose. In light of the fact that ß-D-allose had virtually no effect on DK4300 reporter expression (Fig. 4A), it seems unlikely that [p]ppGpp signaling is affected by ß-D-allose. Rather, the change in DK4521 reporter expression may be explained by modulation of the quorum-sensing A-signal - although this connection would need further study. The strain DK4491, contains a reporter induced during early development and showed no change in response to ß-D-allose (Fig. 4B). Overall, the Tn5lac reporter data suggests that, while ß-D-allose may have a modest effect on the A-signal, its affect on early developmental signaling is low in comparison to genes expressed during later stages of development.

The subsequent stages of fruiting body development, aggregation and sporulation, are controlled by the C-signal (27, 28). The C-signal, a 17-kDa membrane associated protein, is known to induce the expression of many developmental genes after 6 h of starvation, including devRS (27, 30). According to our results, developmental gene expression levels are greatly reduced between 4 and 14 h—a time period which coincides with the initiation of C-signaling (6 to 8 h). In addition, one of the reporter genes suppressed during this time period contains a reporter fusion in the devRS locus. While these results as well as the nonaggregating phenotype hint at a possible defect in C signaling, we examined ß-D-allose effects on C-signal production via Western blot analysis and found no change in CsgA protein expression levels (data not shown). Hence, the connection between C-signaling and ß-D-allose-mediated fruiting inhibition remains unclear. Here, we have shown that ß-D-allose inhibits aggregation and that this morphological phenotype is correlated with a significant reduction in developmental gene expression between 4 and 14 h of development.

While ß-D-allose appears to suppress developmental gene expression, the mechanisms involved are not known. We wondered whether ß-D-allose was involved in a specific interaction that could be antagonized by other sugars. This question was explored in competition studies, where we found that 50 mM galactose and xylose completely restore fruiting despite the presence of 5 mM ß-D-allose (Fig. 6). Although we cannot make a definitive conclusion regarding the molecular mechanism of fruiting rescue, we have observed that all sugars capable of full or partial rescue carry the hydroxyl groups on carbons 2 and 3 in a trans conformation. Previous studies with aldohexose sugars have shown that the position of the C3 hydroxyl group in relation to its neighboring groups can determine whether or not specific interactions occur (41, 55). For example, a study examining the interaction of ß-D-allose with the insulin-stimulated adipocyte transporter found that the critical hydrogen bonding positions involved the cis conformation of the C1 and C3 ring oxygens (41). Though further study is required to determine the exact mechanism of ß-D-allose inhibitory activity, loss of function in the presence of excess sugar suggests that ß-D-allose is involved in a specific molecular interaction that can be antagonized by alternate substrates.

Given additional evidence of a specific ß-D-allose target, we wondered whether the sugar might be affecting M. xanthus development through interactions with the transport or metabolic systems of other sugars. Prior work by Kim et al. led to the discovery of a 6-gene operon devoted to the acquisition and metabolism of D-allose in E. coli. The alsRBACEK operon includes a complete ATP dependent ABC transport system as well as D-allose-regulated enzymes (26). Although the M. xanthus genome does not encode alsRBACEK homologues, it does encode three complete ABC transporters with high sequence similarity to ribose/maltose ABC transporters. Ribose transporters are known to transport ß-D-allose at low levels due to the close structural similarity of the two sugars (7, 26). However, we found that inactivation of the ribose and the two maltose ABC transporter operons through insertional mutagenesis produced no change in sensitivity to ß-D-allose (data not shown).

In a second attempt to find additional members of the ß-D-allose-mediated pathway of inhibition, we conducted a genome-wide mutagenesis screen via magellan-4 transposon insertion (43). Four distinct mutants were found to have transposon insertions in MXAN6497, an ORF with predicted sequence homology to A. dehalogenans glcK, a glucokinase gene. Subsequent glucokinase activity assays demonstrated that cell lysates from a glcK mutant contain 10-fold lower glucokinase activity levels (Fig. 8B). Glucokinases and hexokinases catalyze the phosphorylation of glucose to glucose-6-phosphate in the first step of glycolysis. Whether or not M. xanthus is capable of glycolysis remains unclear. Following an analysis of intermediary metabolism, Watson and Dworkin concluded that M. xanthus contains an incomplete glycolytic pathway due to a lack of glucokinase and pyruvate kinase activity (54). In addition, nutritional studies have repeatedly demonstrated that amino acids and peptides, rather than sugars, are essential for growth (4, 18). On the other hand, our identification of an active glucokinase agrees with an earlier study where hexokinase activity (EC 2.7.1.1) was detected in cell extracts of M. xanthus (58). Furthermore, the M. xanthus genome contains two loci, MXAN3514 and MXAN6299, with sequence similarity to pyruvate kinases (66% to T. tengcongensis pyruvate kinase and 94% to S. aurantiaca pyruvate kinase, respectively [EC 2.7.1.40]).

While evidence for a functional glycolytic pathway continues to mount, the role of glucokinase in M. xanthus carbohydrate metabolism remains far from certain. Whether or not M. xanthus utilizes glcK in its glycolytic pathway, ß-D-allose is not a growth inhibitor and is therefore unlikely to be engaged in disruption of energy-producing pathways. It is also evident that the glucokinase itself is not directly involved in the construction of fruiting bodies; we found that mutations in glcK do not affect development (Fig. 7). Rather, the significance of glcK in ß-D-allose-mediated fruiting inhibition lies in the implication that the hexose likely requires phosphorylation to block development. The inhibition of development by a phosphorylated hexose has been observed before. Youderian et al. found that hexokinase activity is required for the inhibition of development by the glucose analog 2-deoxyglucose (58). Additionally, ß-D-allose (this study) and 2-deoxyglucose were found to inhibit glycerol-induced sporulation in DK1622 liquid cultures (58). The fact that both ß-D-allose and 2-deoxyglucose can block glycerol-induced sporulation suggests that a second process-independent of fruiting body formation-may be targeted. A recent study by Nariya and Inouye has shown that glycogen accumulation during stationary phase and early development is essential for efficient sporulation during fruiting body formation (40). Since glcK is a component of the glycogen synthesis pathway, it is tempting to speculate that phosphorylated ß-D-allose disrupts this pathway through competitive inhibition. Furthermore, if polysaccharide biosynthesis is critical for aggregation, a block in glycogen synthesis could explain both ß-D-allose-sensitive developmental and sporulation phenotypes. These hypotheses await further study.

While many questions remain, it appears that certain hexose sugars are capable of inhibiting development through a common pathway requiring hexose phosphorylation. Our results and previous observations highlight the multitude of biological processes affected by monosaccharides in M. xanthus: glycerol and ribose induce sporulation (10, 44), glucosamine activates cell lysis (39), and ß-D-allose along with 2-deoxyglucose inhibit development (58). As we continue to examine the ß-D-allose-mediated pathway of inhibition, this and future studies promise to expand our understanding of the varied and interchangeable roles simple sugars play in prokaryotic biology.


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ACKNOWLEDGMENTS
 
This work was supported by National Institutes of Health grants GM54666 (W. Shi) and 5F31GM070396 (M. Chavira).

We thank Barry Bochner for patient assistance with the Biolog PM arrays, Daniel Yarbrough for review of the manuscript, Heidi Kaplan for providing the DH5{alpha}-{gamma}pir::pminiHimar1 strain, and Lotte Sogaard-Andersen for supplying the CsgA antibody.


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FOOTNOTES
 
* Corresponding author. Mailing address: UCLA School of Dentistry, CHS 20-114, 650 Charles E. Young Dr. South, Box 951668, Los Angeles, CA 90095-1668. Phone: (310) 825-8356. Fax: (310) 794-7109. E-mail: wenyuan{at}ucla.edu. Back

{triangledown} Published ahead of print on 20 October 2006. Back


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Journal of Bacteriology, January 2007, p. 169-178, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00792-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




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