Department of Microbiology, Molecular Biology
and Biochemistry, University of Idaho, Moscow, Idaho
83844-3052,1 and Department of Biology,
The John Muir College, University of California at San Diego, La
Jolla, California 92093-01162
The glucose analog 2-deoxyglucose (2dGlc) inhibits the growth and
multicellular development of Myxococcus xanthus. Mutants of
M. xanthus resistant to 2dGlc, designated hex
mutants, arise at a low spontaneous frequency. Expression of the
Escherichia coli glk (glucokinase) gene in M. xanthus
hex mutants restores 2dGlc sensitivity, suggesting that these
mutants arise upon the loss of a soluble hexokinase function that
phosphorylates 2dGlc to form the toxic intermediate,
2-deoxyglucose-6-phosphate. Enzyme assays of M. xanthus
extracts reveal a soluble hexokinase
(ATP:D-hexose-6-phosphotransferase; EC 2.7.1.1) activity
but no phosphotransferase system activities. The hex
mutants have lower levels of hexokinase activities than the wild type,
and the levels of hexokinase activity exhibited by the hex
mutants are inversely correlated with the ability of 2dGlc to inhibit
their growth and sporulation. Both 2dGlc and N-acetylglucosamine act as inhibitors of glucose turnover
by the M. xanthus hexokinase in vitro, consistent with the
finding that glucose and N-acetylglucosamine can antagonize
the toxic effects of 2dGlc in vivo.
 |
INTRODUCTION |
The myxobacteria, including their
best-characterized representative, Myxococcus xanthus, are
social prokaryotes that undergo multicellular development. When a large
number (>105) of M. xanthus cells are starved
for essential nutrients, individual cells within a starved population
coordinate their movements to form a fruiting body that supports the
differentiation of a subset of vegetative cells into spores. Spores
represent only a small minority (1 to 10%) of developing cells after
carbon starvation and are more resistant to heat, UV light, and
sonication than vegetative cells. Spores ensure the survival of this
organism under extreme environmental conditions (17).
Development is a rapid process. Spores mature within fruiting bodies in
the short span of 36 h. The cycle leading to spore maturation
involves a cascade of gene expression, in which successive subsets of
developmental stage-specific promoters are activated (22).
Development is accompanied by dramatic changes in the flow of carbon
through metabolic pathways. The early stages of development involve the
oxidative catabolism of vegetative biopolymers to generate intermediary
metabolites. These metabolites fuel a burst of primarily gluconeogenic
anabolism during the later stages of development. This gluconeogenic
burst begins about 4 to 6 h after starvation, and, in turn,
supports the assembly of the spore coat, comprised mainly of
polysaccharides (48).
Although many studies have focused on the cascade of gene expression
that occurs during development, very little is known about the
metabolic changes that accompany this adaptive response. Most of the
genes known to be required for development function early in this
cycle. In contrast, few genes that function late in development have
been identified. Thus, little is known about the genes involved in
spore maturation, the regulation of gene expression during late
development, or the spatial and temporal coordination of polysaccharide
biosynthesis with spore coat assembly (17).
To understand how polysaccharide biosynthesis and spore maturation are
coordinated, we are exploring the regulation of the gluconeogenic
pathway during development. Previous studies of intermediary metabolism
have shown that M. xanthus makes many of the enzymes that
catalyze steps in this pathway. These include phosphoenolpyruvate (PEP)
carboxykinase (EC 4.1.1.32), glyceraldehyde-3-phosphate dehydrogenase (EC 1.2.1.12), aldolase (EC 7.1.2.7),
fructose-1,6-phosphate phosphatase (EC 3.1.3.11), and
phosphoglucoisomerase (EC 5.3.1.9), as well as phosphoglucomutase (EC
2.7.5.1) and UDPG pyrophosphorylase (EC 2.7.7.9) (46).
Expression of the gluconeogenic enzymes is induced during an alternate
developmental pathway, in which cells sporulate in response to high
external concentrations of short-chain polyols (10).
The gluconeogenic pathway in M. xanthus is unusual in two
respects. The M. xanthus PEP carboxykinase, which catalyzes
the first committed step in gluconeogenesis, requires GTP or ITP as a
phosphoryl donor (46), like eukaryotic PEP carboxykinases and unlike many of their ATP-dependent prokaryotic counterparts. In
addition, the M. xanthus glycolytic pathway, which shares
many enzymes with the gluconeogenic pathway, initially appeared
incomplete. When extracts were prepared from vegetative M. xanthus cells and assayed for glycolytic activities, neither
ATP-dependent hexokinase (EC 2.7.1.1) nor pyruvate kinase (EC 2.4.1.40)
activities were detected (46).
Their inability to detect intracellular ATP-dependent hexokinase,
pyruvate kinase, and high-affinity glucose permease activities led
Watson and Dworkin (46) to speculate that M. xanthus may exclude glucose for some selective advantage. These
results also could suggest that the apparently glycolytic activities
produced by M. xanthus may have roles only in the anabolic
pathway of gluconeogenesis. However, this notion is not satisfying,
because M. xanthus makes an ATP-dependent
phosphofructokinase (EC 2.7.1.11), which catalyzes a committed step in
glycolysis that must be bypassed in the gluconeogenic pathway
(46).
During vegetative growth, M. xanthus is predatory and is
thought to derive its energy primarily from the oxidative catabolism of
proteins released by the lysis of prey organisms by a battery of
secreted enzymes. The minimal requirements for the growth of M. xanthus in defined media suggest that proteins comprise the core
of its rich diet. M. xanthus is auxotrophic for leucine, isoleucine, and valine, the three branched-chain amino acids, and for
methionine or cobalamin. M. xanthus is also a phenylalanine bradytroph, and many nonessential amino acids stimulate its growth (2, 7, 18).
Surprisingly, hexose monosaccharides including glucose (Glc), fructose
(Fru), galactose, and glucosamine (2-amino-2-deoxyglucose; GlcN), as
well as a variety of their disaccharides, do not stimulate the growth
of this obligate aerobe on defined medium. In contrast, acetate and
pyruvate, as well as the citric acid cycle intermediates malate and
succinate, do stimulate growth (2, 18). Again, these results
suggest that M. xanthus cannot transport or phosphorylate Glc efficiently or, at the least, utilize Glc as an energy source. Only
a small fraction of label added as [14C]glucose is
incorporated into biopolymers with high molecular mass during growth on
defined media (18, 46), reinforcing the idea that M. xanthus does not require Glc for growth.
On the other hand, it is clear that simple sugars can play critical
roles during M. xanthus development. One subset of simple sugars, including the reduced triose glycerol (8) and the
pentoses ribose and xylose (34), elicit the synchronous and
rapid conversion of rod-shaped vegetative cells in liquid culture into
spherical spores. These short-chain polyols trigger an abbreviated
unicellular developmental pathway leading to the formation of spores
that are more heat resistant than vegetative cells but less heat
resistant than the more structurally complex spores produced during
starvation-induced multicellular development (42).
Furthermore, addition of the GlcN, but not the closely related hexose
Glc or N-acetylglucosamine (2-deoxy-2-N-acetylamino-D-glucose; GlcNAc), to
exponentially growing cells at high concentrations elicits cell lysis
(29). Therefore, it is inviting to speculate that M. xanthus uses these sugars or their phosphorylated derivatives as
signals to trigger pathways that define distinct cell fates during development.
M. xanthus is sensitive to the antibiotic 2-deoxyglucose
(2dGlc). In most organisms, the analog 2dGlc is transported by
mechanisms that also transport glucose and must be phosphorylated to
form 2-deoxyglucose-6-phosphate (2dGlc6P) to exert its toxic effect. Starting with this discovery, we obtained genetic evidence implying that M. xanthus phosphorylates Glc with a soluble hexokinase
and have identified this hexokinase activity in vitro.
| |
MATERIALS AND METHODS |
Media and chemicals.
CTPM liquid medium (1% Casitone, 10 mM
Tris [pH 7.6], 1 mM potassium phosphate [pH 7.5], 5 mM
MgSO4) was used for growth of M. xanthus. TPM
buffer, used for developmental assays, is CTPM without Casitone. Stocks
of monosaccharides used to supplement media were dissolved in water and
filter sterilized. Antibiotics, sugars, enzymes, substrates, and other
chemicals were from Sigma or Aldrich, with the exception of ATP, which
was from Pharmacia. 14C-labeled hexoses used as substrates
in the assays for glucose-specific phosphotransferase system (PTS)
activities were from ICN or NEN-DuPont. Oligonucleotides used for
plasmid construction and mutagenesis were made by Biosource Inc.
Restriction endonucleases and DNA-modifying enzymes were from New
England Biolabs and were used under recommended conditions.
Bacteria and plasmids.
Escherichia coli JM107
(50) was used for the construction of plasmids by standard
methods and for the preparation of plasmid DNA (37).
Plasmids were introduced into JM107 by electroporation (43).
Derivatives of JM107 with plasmids were grown in LB medium supplemented
with ampicillin (100 µg/ml) and/or kanamycin (40 µg/ml). M. xanthus strains are derivatives of the wild-type strain DK1622
(19) or the nonmotile strain DZ1 (3). Strains
XS200 and XS201, which carry the hex1 and hex3
mutations, respectively, were selected by plating 108
exponentially grown cells from independent cultures of DK1622 on CTPM
medium with 1% 2dGlc. In some experiments, the rates of growth of
M. xanthus cultures were monitored by measuring relative light scattering in Klett-Summerson units (KU), using a Klett-Summerson colorimeter (model 800; 640- to 700-nm filter). Light scattering is
proportional to the numbers of viable wild-type cells grown in CTPM
medium at 32°C over the range of 10 to 120 KU. The assay of viable
M. xanthus cells in culture was made by the soft-agar overlay method, in which aliquots of cultures diluted in CTPM medium
were resuspended in 3 ml of CTPM soft agar (0.7%) and distributed over
the surfaces of CTPM agar (1.5%) plates.
Integrative plasmid pAY703 (26) carries the M. xanthus
mglBA operon subcloned into kanamycin-resistant (Kmr)
vector pBGS18 (41). To express the E. coli glk
(glucokinase) gene as part of the mglBA operon, template DNA
isolated from E. coli JM107 was amplified with primers GLK1
(5' AAAGGTACCATGAGGAGGTGTACATGACAAAGTATGCATTAGTCG) and GLK2
(5' CCCGAATTCTCTAGAGAATGTGACCTAAGGTCTGGCG). The
amplified product was cleaved with Acc65I and
EcoRI and ligated to the same sites of pAY703 to make
pAY1108. Plasmids were introduced into M. xanthus by
electroporation (20).
Assays for the multicellular development of M. xanthus.
To initiate development, derivatives of DK1622 were grown
to a density of 5 × 108/ml in CTPM medium at 32°C,
concentrated by low-speed centrifugation, and resuspended in 0.1× TPM
buffer. For standard assays (23), multiple spots (20 µl)
were made on TPM (1.5%) agar; plates were incubated at 32°C for
120 h. Sets of five 20-µl spots were harvested after incubation
at 50°C for 2 h and scraped together into 1 ml of TPM buffer.
Suspensions were sonicated for 15 s at 5 W on a Fisher model 60 Sonic Dismembranator to disperse spores. Serial dilutions of spore
suspensions were plated on CTPM plates by the soft-agar overlay method
and scored after 72 h at 32°C.
To determine the time(s) after the onset of starvation at which 2dGlc
inhibits development, we modified the standard assay. Aliquots (1 ml)
of exponentially growing cells suspended in 100 µl of TPM buffer were
spotted on the surfaces of nitrocellulose filters (0.025-µm-pore-size
type VS; Millipore) that had been sterilized by autoclaving in
distilled H2O. Spots were allowed to dry for 2 h at
25°C; then the plates with filters were placed at 32°C. At each
12-h interval after the 2-h initial drying period and throughout the
5-day incubation period, duplicate filters were transferred with
sterile forceps from the surfaces of TPM agar plates to TPM agar plates
supplemented with 0.1% 2dGlc. Two filters were retained on TPM plates,
and not transferred to plates with 2dGlc, to ensure that fruiting body
formation and sporulation were not affected by cell contact with the
filters. After transfer, filters on plates with 2dGlc were incubated at
32°C to complete the incubation time of 120 h and then incubated
at 50°C for 2 h to kill vegetative cells. Filters were quartered
by using sterile scissors and placed into a 2-ml microcentrifuge tube
containing 1 ml of TPM buffer, and samples were sonicated for 15 s
at 5 W. Serial dilutions of each sonicate were plated on CTPM agar
plates. Colonies arising from germinated spores were counted after
72 h at 32°C.
To determine the precise time after the initiation of the developmental
cycle at which spores mature when cells are allowed to develop on
filters, we set up a parallel experiment. Cells were grown, harvested,
and spotted onto filters on the surfaces of TPM agar plates with or
without 0.1% 2dGlc. At 12-h intervals after the initiation of
development, filters were removed, and incubated at 50°C for 2 h
to kill vegetative cells prior to assays for heat-resistant spores. The
results of this control experiment showed that spores first acquire
heat resistance at about 36 h after the initiation of development,
as they do when cells are placed directly upon the TPM agar surface.
PTS activity assays.
Wild-type strain DK1622 and its mutant
derivatives were grown to a density of 5 × 108
cells/ml in CTPM medium supplemented with 2% of each of the PTS sugars
Glc, GlcNAc, Fru, and mannitol (Mtl) at 32°C and concentrated by
low-speed centrifugation. Cell pellets were suspended in 1/10 volume 20 mM Tris buffer (pH 8.0)-1 mM EDTA-5 mM 2-mercaptoethanol and
disrupted by sonication with four 10-s bursts at 5 W in a Branson W140D
Sonifier cell disruptor. Assays for partial PTS activities
(35a), for fructose-1-phosphate kinase activity
(1a), and for mannitol-1-phosphate dehydrogenase activity
(35a) were performed as described elsewhere.
Hexokinase activity assays.
Wild-type strain DK1622 and its
mutant derivatives were grown to a density of 5 × 108
cells/ml in CTPM medium at 32°C, concentrated by low-speed
centrifugation, and suspended in 1/10 volume TPM buffer at 4°C. Cell
extracts were prepared by sonication of cultures in TPM for 10 s
at 100 mW in a model 60 Fisher Sonic Dismembranator and then subjected to microcentrifugation at 20,000 × g for 5 min to
remove cell debris. Extracts were maintained at 4°C no longer than
4 h prior to assay.
Hexokinase activities were determined by using the coupled reaction of
Fraenkel and Horecker (12), with minor modifications. Standard reaction mixtures (1 ml) contained 50 mM Tris (pH 9.1)-10 mM
MgCl2 (hexokinase buffer), 20 mM glucose, 10 mM ATP (pH
9.1), 10 to 100 µl of cell extract (0.1 to 2.0 mg of protein), and
500 to 1,000 U of Leuconostoc mesenteroides
glucose-6-phosphate (Glc6P) dehydrogenase. (One unit corresponds to 1 nmol of Glc6P min
1 mg1 Glc6P oxidized to
6-phospho-D-gluconate in the presence of NADP at pH 7.6 at
25°C.) In experiments used to determine apparent values of
Km for ATP and Glc and Ki
for 2dGlc, the concentrations of these compounds were varied in the
standard reaction mixtures. Assays were initiated by the addition of
NADP to 200 µM. The initial velocity of NADP reduction
(
NADPH = 6.22 × 106 cm2)
was measured at 25°C as the change in absorbance at 340 nm at 30-s
intervals. Spectrophotometry was done with a Perkin-Elmer Lambda 12 UV/VIS spectrophotometer supported by a Dell OptiPlex XMT590
workstation and the uv-WinLab software package. Initial velocities were
found to be linear for at least 5 to 10 min and proportional to the
protein concentration of extracts (Fig.
1). Control experiments showed that the
concentrations of inhibitors used in the assays that we report did not
decrease the rate of the coupled, NADP-dependent oxidation of Glc6P by
Glc6P dehydrogenase to make this the rate-limiting step in these
assays. Cell supernatants used in enzyme assays were assayed for total
protein content by the bicinchoninic acid method (40);
reagents for assay were from Pierce. Samples (10 µl) were placed in
the wells of a 96-well microtiter plate, and 200 µl of reagent was
added. Microtiter plates were incubated at 37°C for 30 min and then
absorbance at 540 nm was measured with a Titertech Multiscan MCC340
spectrometer. Protein concentrations were extrapolated from a standard
bovine serum albumin curve.
View larger version (16K):
[in this window]
[in a new window]
|
FIG. 1.
Hexokinase activity in extracts from wild-type M. xanthus cells is a linear function of protein concentration.
Activities of hexokinase are plotted as a function of total protein in
extracts prepared from wild-type M. xanthus cells, are the
averages of three determinations, and had standard deviations of
<±0.032 nmol min1. Extracts were incubated in 50 mM
Tris-10 mM MgCl2 (pH 9.1) with 500 to 1,000 U of Glc6P
dehydrogenase plus Glc (20 mM) and ATP (10 mM) (filled circles), ATP
(10 mM) (open circles), or Glc (20 mM) (squares). The observed rates of
NADP reduction in the absence of added ATP (squares) are similar in
reactions with or without added extract, activity is dependent on the
addition of Glc, and the addition of Glc decreases the rate of
spontaneous NADP reduction in the complete assay without added
extract.
|
|
When hexokinase activity is assayed directly by the method of Darrow
and Colowick (5), which measures the rate at which protons
are generated concomitant with Glc phosphorylation, a competing
activity that consumes protons at a rate faster than their liberation
is observed. This is true even in extracts prepared from M. xanthus cells expressing E. coli glucokinase. We chose to assay hexokinase activity at pH 9.1 in the coupled assay, because we
found that E. coli glucokinase is about twofold more active (and has lower Kms for both ATP and Glc) at pH
9.1 than at pH 7.6, whereas M. xanthus hexokinase has
comparable activities at both pH values. Hexokinase activity in
M. xanthus extracts is labile. Activity is lost within
48 h if extracts are stored at 4°C and does not survive repeated
cycles of freezing at 
20°C and thawing. Therefore, we took care to
assay extracts immediately after the sonication of cells. This
instability may account for the negative results of Watson and Dworkin
(46), as may a relatively high background of hexokinase
activity present in preparations of Saccharomyces cerevisiae
Glc6P dehydrogenase, given that the specific activity of M. xanthus hexokinase is quite low.
| |
RESULTS |
2dGlc inhibits the growth of M. xanthus.
Wild-type
M. xanthus DK1622 does not form colonies efficiently on
medium containing the Glc analog 2dGlc (Table
1). Concentrations of 2dGlc as low as
0.04% (2.4 mM) inhibit the efficiency of plating of wild-type cells by
a factor of greater than 105. This result is surprising in
light of the previous findings that cell extracts from vegetative
M. xanthus cells lack an ATP-dependent glucokinase activity
(46) and that less than 2% of label added to exponentially
growing cells in the form of [14C]glucose is incorporated
into intracellular materials with higher molecular mass (18,
46). In other microbes, 2dGlc is most often transported by
mechanisms that facilitate the transport of glucose.
Wild-type M. xanthus cells plate with high efficiencies in
the presence of higher concentrations of a variety of other hexoses, including Glc, methyl-
-D-glucopyranoside (MeGlc),
GlcNAc, Fru, and mannitol Mtl. However, the growth of M. xanthus is inhibited by at least one other hexose, glucosamine
(GlcN). We find, as did Mueller and Dworkin, that wild-type cells do
not form colonies on rich medium with 2% GlcN (Table 1). This is
consistent with the observation that the addition of GlcN to
exponentially growing cells at high concentrations elicits cell lysis
(29).
2dGlc, unlike glycerol or GlcN, is not a morphogen.
To address
how 2dGlc inhibits growth, we tested whether 2dGlc, like GlcN, might
act as a morphogen to induce cell lysis. We also tested whether 2dGlc
might behave like another subset of sugars that elicit a different
morphogenetic response when added to exponentially growing cells. These
sugars, including the reduced triose glycerol (8) and the
pentoses ribose and xylose (34), trigger the rapid
sporulation of vegetative M. xanthus cells without the
concomitant formation of fruiting bodies. The development of M. xanthus cells in response to these short-chain polyols is thought
to mimic the later steps in starvation-induced multicellular development for two reasons. A subset of mutants resistant to glycerol-induced sporulation do not undergo multicellular development (9), and the accumulation of intracellular glycerol within starving cells coincides with later steps in starvation-induced development (13).
Wild-type cells were grown to exponential density in rich medium, and
each of a variety of sugars was added to the growing cells. The growth
of the cultures and the microscopic appearances of cells within each
culture were monitored at various times after the addition of these
sugars. As shown in Fig. 2, when a
culture of exponential cells is supplemented to 0.5 M glycerol, these glycerol-treated cells tend to aggregate in liquid suspension as they
differentiate into spores. Consequently, the turbidity of a
glycerol-treated culture decreases significantly. The majority of cells
change from long rods to more refractile spheres within 3 h (Fig.
3B). In contrast, when cells are treated
with 2% GlcN, the turbidity of the culture continues to increase,
although not so rapidly as that of an untreated culture (Fig. 2).
Microscopic examination of the GlcN-treated cells reveals that the
majority has lysed after 3 h of treatment, leaving behind empty,
rod-shaped husks (Fig. 3D). Presumably, these husks retain the ability
to scatter visible light and contribute to the increase in turbidity that accompanies the lysis of GlcN-treated cultures. In contrast, the
turbidity of cultures treated with 1% 2dGlc does not change for 6 h after treatment (Fig. 2), and cells within these cultures neither
round to form spores nor lyse (Fig. 3C).
View larger version (18K):
[in this window]
[in a new window]
|
FIG. 2.
2dGlc arrests the growth of M. xanthus. A
culture of wild-type cells was grown to a density of 5 × 108/ml in CTPM medium at 32°C. At the start of the
experiment, equal subcultures were left untreated (open circles) or
supplemented to 0.1 M 2dGlc (filled circles), 0.1 M GlcN (open
squares), or 0.5 M glycerol (filled squares). The percent relative
turbidity (turbidity of each subculture divided by its turbidity at
0 h) is shown; initial values, determined immediately after the
addition of supplements, ranged from 90 to 110 KU.
|
|
View larger version (172K):
[in this window]
[in a new window]
|
FIG. 3.
2dGlc is not a morphogen. At 3 h after treatment, 1 ml of each culture prepared as described in the legend to Fig. 2 was
concentrated by low-speed centrifugation and suspended in 0.1 ml of
CTPM medium. Aliquots were examined by using differential interference
contrast optics and a 40× objective lens with a Nikon Microphot-FXA
microscope (bottom). (A) Untreated cells; B, cells plus 0.5 M glycerol;
C, cells plus 1% 2dGlc; D cells plus 2% GlcNAc. Photomicrographs of
cells are at the same magnification; the average length of untreated
cells is ca. 10 µm.
|
|
When aliquots of treated cells are plated for viable titers at various
times after treatment, we find that glycerol-induced spores can
germinate to yield the initial titer of cells from which they arose,
consistent with previous results (9). Cells treated with
GlcN show a dramatic decrease in titer within 6 h, consistent with
the results published by Mueller and Dworkin (29). In
contrast, the viable titer of cells after treatment with 1% 2dGlc does
not change for 6 h (Fig. 4). After
prolonged treatment with 2dGlc, cells round up and eventually lyse.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 4.
M. xanthus cells die slowly in the presence
of 2dGlc. The turbidity (A) and viability (B) of subcultures left
untreated (open circles) or treated with 2dGlc as described in the
legend to Fig. 2 (filled circles) was monitored over a 48-h time
course. Viability was determined by plating serial dilutions of the
subcultures on CTPM agar plates. Note that a twofold loss in viability
is observed after 12 h of treatment with 2dGlc, corresponding to
the period of one doubling in the untreated culture.
|
|
Spontaneous mutants of M. xanthus resistant to 2dGlc
arise at a frequency indicative of the loss of function.
The
sensitivity of M. xanthus to 2dGlc suggests that M. xanthus can phosphorylate this hexose analog. Sensitivity to 2dGlc in many other microbes requires both the transport of this glucose analog and its phosphorylation to form the toxic intermediate, 2dGlc6P.
This can occur by one of two different general mechanisms: group
translocation, in which the transport of hexoses is coupled with their
phosphorylation, or a combination of active transport and
phosphorylation by a soluble hexokinase. For example, E. coli can use the PTS, dependent on the ptsI,
ptsH, and crr genes, to simultaneously transport
2dGlc and phosphorylate this substrate, using PEP as the phosphoryl
donor (31). Many other gram-negative and gram-positive
bacteria have similar PTSs that translocate both Glc and its toxic
analog 2dGlc (33, 35).
When microbes do not couple 2dGlc translocation with its
phosphorylation, sensitivity to 2dGlc depends on the combination of an
independent transport mechanism and a soluble hexokinase activity. Such
is the case for both Streptomyces coelicolor (1) and S. cerevisiae (25). Most mutations that
confer resistance to 2dGlc in these organisms inactivate the genes
encoding hexokinase. Mutations that confer resistance to 2dGlc in yeast
also are found in a gene required for the transport of both Glc and
2dGlc (30). Both known mechanisms by which microbes are
sensitive to 2dGlc involve the transport and phosphorylation of 2dGlc
by proteins that transport and phosphorylate a subset of other hexoses.
This subset often includes Glc.
Because many gram-negative bacteria have one or more PTSs capable of
2dGlc group translocation, initially it was inviting to speculate the
M. xanthus might also have a PTS. This hypothesis would
neatly explain the previous failure to find both ATP-dependent hexokinase and pyruvate kinase activities made by M. xanthus
(46); both PTS activities are coupled and most often PEP
dependent. Therefore, we assayed extracts prepared from M. xanthus cells grown in the presence of each of the four PTS sugars
Glc, GlcNAc, Fru, and Mtl for PTS activities. However, we did not
detect PEP- or ATP-dependent PTS activities with any of these four
hexoses, MeGlc, or 2dGlc as the labeled substrate. In addition, we
could detect neither fructose-1-phosphate kinase activity
characteristic of the Fru PTS or mannitol-1-phosphate dehydrogenase
activity characteristic of the Mtl PTS. These negative results
suggested that, like S. coelicolor (1), M. xanthus has independent mechanisms of 2dGlc (and Glc) transport
and phosphorylation.
To confirm this hypothesis, we began with a genetic approach, selecting
and characterizing mutants of M. xanthus resistant to 2dGlc.
When wild-type strain DK1622 is plated on rich medium supplemented to
1% 2dGlc, it forms colonies with an efficiency of 3 × 106 (Table 1). This is about 10-fold higher than the
frequency observed for spontaneous mutations that inactivate the
uraA gene of M. xanthus (21). This
high frequency of spontaneous mutation suggests that resistance to
2dGlc is due to the loss of function, and mutations that confer
resistance inactivate one or more target genes.
Colonies formed by mutants resistant to 2dGlc have two distinct
morphologies on rich medium with 2dGlc. Under restrictive conditions,
most colonies are large with diffuse, spreading edges, whereas fewer
colonies are smaller (less motile) and have sharp, well-defined edges.
Under permissive conditions (rich medium without 2dGlc), both types of
mutants form large, spreading colonies with the same morphology as the
wild type. We isolated and characterized two independent mutants of the
wild-type strain, carrying hex1 and hex3
mutations, which form larger and smaller colonies, respectively, under
restrictive conditions. These mutants form colonies on rich medium
supplemented with 1% 2dGlc with similar efficiencies as on medium
without 2dGlc (Table 2).
Expression of the E. coli glk (glucokinase) gene in
mutants of M. xanthus resistant to 2dGlc restores
sensitivity to 2dGlc.
If M. xanthus, like E. coli, has a PTS for the coupled transport and phosphorylation of
2dGlc, then we would expect that expression of a soluble hexokinase in
such mutants should not restore sensitivity to 2dGlc. For example,
mutations in E. coli that inactivate the group translocation
system for Glc confer resistance to 2dGlc yet retain the soluble
hexokinase activity made by the glk gene (4).
Alternatively, if M. xanthus, like S. coelicolor
or S. cerevisiae, has independent glucose transport and
hexokinase functions, then mutants of M. xanthus resistant
to 2dGlc should carry mutations that inactivate a gene, hex,
encoding a soluble hexokinase. The expression of a heterologous
glucokinase function in these mutants, such as the product of the
E. coli glk gene, should restore sensitivity to 2dGlc. For
example, when a plasmid with an active glkA gene is
introduced into a mutant strain of S. coelicolor resistant to 2dGlc, it restores sensitivity to 2dGlc (45). Therefore, we constructed derivatives of wild-type M. xanthus DK1622,
and its mutant derivatives resistant to 2dGlc that express the E. coli glk gene, and examined their phenotypes.
The E. coli glk (glucokinase) gene (28) was
amplified by PCR and cloned distal to the mglA gene in the
constitutively expressed mglBA operon (16)
carried by plasmid vector pAY703 (26). Plasmid pAY703 and
its otherwise isogenic derivative with glk, pAY1108, have a
Kmr determinant and an origin that supports autonomous
replication in E. coli but not in M. xanthus.
After electroporation, these plasmids can integrate into the circular
M. xanthus chromosome by homologous recombination between
the plasmid and chromosome within the region of homology shared by the
two elements, the mglBA operon. Integration results in the
formation of Kmr merodiploid strains with two copies of the
mglBA operon that sandwich nonhomologous plasmid DNA (Fig.
5).
View larger version (9K):
[in this window]
[in a new window]
|
FIG. 5.
Plasmids pAY703 and pAY1108. The structures of the
chromosomal mgl locus are shown for wild-type M. xanthus and for recombinant merodiploid strains with integrated
plasmid vectors pAY703 and pAY1108. Recombinant strains with plasmid
pAY1108 express the E. coli glk (glucokinase) gene from the
constitutive mglBA promoter. The level of glucokinase
expression in M. xanthus is the same as its basal level of
expression in E. coli (28).
|
|
A Kmr derivative of DK1622 with integrated plasmid pAY1108
should express the E. coli glk gene as part of the
mglBA operon. We have shown that similar derivatives of
plasmid pAY703 express the myxophage Mx8 mox (26)
and int (36) genes, as well as the M. xanthus sglK gene (47). Kmr electroporants
of host DK1622 carrying plasmid pAY1108 were obtained with the same
efficiency after electroporation as with parental plasmid pAY703. Also,
after purification, we find that strain DK1622::pAY1108 has
the same doubling time as strain DK1622::pAY703 in rich
medium at 32°C.
A comparison of the abilities of strains DK1622,
DK1622::pAY703, and DK1622::pAY1108 to form
colonies in the presence of glucose and several glucose analogs is
shown in Table 1. All three strains form colonies with similar
efficiencies on rich medium supplemented with each of the hexoses, with
one exception. Whereas wild-type strain DK1622 and strain
DK1622::pAY703 plate with low but detectable efficiencies
(3 × 106) on medium with 1% 2dGlc, strain
DK1622::pAY1108 does not form colonies with a
measurable efficiency (<108). Expression of the E. coli glk gene in wild-type M. xanthus results in a gain
of function and confers an increased level of sensitivity to 2dGlc upon
the wild-type strain.
The addition of the glk gene to the wild-type M. xanthus genetic background prevents mutation to 2dGlc resistance
in a single step. The simplest hypothesis to account for this finding
is that glk is expressed in M. xanthus, and the
functions of glk and of the target (hex) for
2dGlc resistance mutations in the wild-type M. xanthus
genetic background are similar. Consistent with this hypothesis,
derivatives of the mutant hex1 and hex3 strains
carrying pAY1108 (mglBA-glk) regain sensitivity to 2dGlc,
whereas their otherwise isogenic controls with plasmid pAY703
(mglBA) remain resistant to 2dGlc (Table 2). These two lines
of genetic evidence suggest that the hex mutations confer
resistance to 2dGlc due to the loss of function and that this function
is a soluble hexokinase. A derivative of the wild-type strain
expressing both hex and glk cannot acquire
resistance in a single mutational step, whereas both the wild-type
strain (expressing hex alone) and the complemented mutant
strains (expressing glk alone) can (Table 2).
Three additional results support this explanation. First, the mutant
hex1 and hex3 strains carrying pAY1108
(mglBA-glk) were plated on rich (CTPM) medium with 1%
2dGlc, and 104 independent 2dGlc-resistant mutants arising from each
strain were tested for a Kmr phenotype. Among the
2dGlc-resistant colonies arising from the hex1::pAY1108 and
hex3::pAY1108 strains, 104 of 104 (100%) and 52 of 104 (50%), respectively, were found to have acquired a
Kms phenotype. These Kms segregants most likely
arise due to the loss of integrated plasmid pAY1108 DNA, after
homologous recombination within the merodiploid mgl locus
promotes excision of the integrated plasmid, and a subset of daughter
cells do not inherit the excised plasmid. Second, we isolated a
spontaneous Kms segregant from strain
DK1622::pAY1108 that had lost its integrated plasmid. This
segregant, like its wild-type grandparent, forms colonies on medium
with 2dGlc with an efficiency of 3 × 106. Third,
the multiple mutant, nonmotile M. xanthus strain DZ1 (3), with a different genetic background, carries a
uncharacterized mutation (hex2) that confers resistance to
2dGlc. Again, the introduction of plasmid pAY1108
(mglBA-glk), but not plasmid pAY703 (mglBA), into
this strain restores sensitivity to 2dGlc.
The sensitivity of M. xanthus to 2dGlc can be
antagonized by Glc and GlcNAc but not by Fru or Mtl.
If M. xanthus can transport 2dGlc and, like other microbes,
phosphorylates 2dGlc to make the toxic metabolite 2dGlc6P, then a
subset of hexoses may antagonize this sensitivity to 2dGlc, by
competing with 2dGlc as alternate substrates for phosphorylation or by
inhibiting hexokinase by another mechanism. Therefore, we measured the
efficiency of plating of wild-type strain DK1622 on medium containing
0.1% (6 mM) 2dGlc and various concentrations of potential hexose
antagonists. Wild-type strain DK1622 plates with the low efficiency of
about 3 × 106 on medium containing 0.1% (ca. 6 mM)
2dGlc alone. However, the addition of Glc or GlcNAc to 2% (ca. 100 mM)
permits this strain to form colonies with high efficiencies in the
presence of 2dGlc (Table 3). In contrast,
the addition of either Fru or Mtl to 2% does not antagonize 2dGlc.
Because production of E. coli glucokinase in hex
mutant strains of M. xanthus restores sensitivity to 2dGlc, the hex mutant strains retain the ability to transport
2dGlc. In addition, both Glc and GlcNAc are transported by M. xanthus. From these data, however, we cannot assess whether this
transport mechanism is active or passive.
On the other hand, it is not surprising that M. xanthus
transports these hexoses. M. xanthus is known to transport a
variety of saccharides, including galactose, 
-galactosides, and
sucrose. When the E. coli galK gene is expressed in M. xanthus, growth becomes sensitive to both 2-deoxygalactose and
galactose (44), indicating that galactose is transported but
not phosphorylated efficiently by M. xanthus. The E. coli lacZ gene produces active -galactosidase activity in
M. xanthus, and cells transport the -galactoside
5-chloro-4-bromo-3-indolyl--D-galactopyranoside even in
the absence of a functional lacY permease (6).
M. xanthus strains expressing E. coli lacZ also
are sensitive to o-nitrophenyl--D-galactoside (unpublished results). M. xanthus cells that express the
foreign Bacillus subtilis sacBR genes also acquire
sensitivity to sucrose (49).
Both the fruiting body morphogenesis and sporogenesis of M. xanthus in response to starvation are inhibited by 2dGlc.
The later stages of the M. xanthus developmental cycle
involve dramatic changes in carbohydrate metabolism. Late in
development, a burst of gluconeogenic anabolism fuels the assembly of
the spore coat, comprised primarily of polysaccharides. Because the
toxic metabolite 2dGlc6P may inhibit enzymes common to the glycolytic and gluconeogenic pathways, and the pathway(s) that this metabolite inhibits may be required for development, we also examined whether 2dGlc inhibits the multicellular development of M. xanthus.
Cultures of wild-type and hex mutant strains were grown to
exponential density in rich medium, concentrated by centrifugation, and
spotted on starvation (TPM) medium with various added concentrations of
2dGlc. After 120 h, all three strains were found to form fruiting bodies on TPM medium without 2dGlc (Fig.
6) (4). On TPM medium with as
little as 0.05% 2dGlc, only the hex1 mutant strain forms fruiting bodies. Under these conditions, the wild-type strain forms no
aggregates, and the hex3 mutant strain forms poorly defined, translucent aggregates. Higher concentrations of 2dGlc (>0.5%) inhibit the fruiting body morphogenesis of even the hex1
mutant (data not shown).
View larger version (97K):
[in this window]
[in a new window]
|
FIG. 6.
2dGlc impairs fruiting body morphogenesis during the
development of M. xanthus. The morphologies of developing
wild-type and hex3 and hex1 mutant cells were
photographed with a Nikon SMZ-U stereomicroscope at a magnification of
×15 on TPM medium and TPM medium supplemented with 0.05% 2dGlc
120 h after the onset of starvation. (A) Wild type on TPM; (B)
hex3 on TPM; (C) hex1 on TPM; (D) wild type on
TPM plus 2dGlc; (E) hex3 on TPM plus 2dGlc; (F)
hex1 on TPM plus 2dGlc.
|
|
Cells that had undergone starvation for 120 h were recovered from
plates, resuspended in TPM buffer, incubated at 50°C to kill
vegetative cells, and assayed for the presence of heat-resistant spores. All three strains form a wild-type complement of heat-resistant spores on starvation agar without 2dGlc (Table
4). Because the hex mutations
result in a loss of function, it is likely that the hex gene
is not essential for development, because hex mutants both
fruit and sporulate. Table 4 also shows that at concentrations of both
0.5 and 2%, 2dGlc inhibits the formation of heat-resistant spores by
wild-type and hex3 mutant cells but not by hex1
mutant cells. Thus, hex1 mutant cells can sporulate in the
presence of 2% 2dGlc, even though fruiting is impaired under these
conditions.
As a control for this experiment, we also tested whether the
sporulation of wild-type and hex mutant cells is sensitive
to GlcN. As shown in Table 4, the development of all three strains is
inhibited by GlcN. Unlike their response to 2dGlc, however, the
development of all three strains suffers to similar extents in the
presence of different concentrations of this hexose.
Cells in two of the colonies formed by the rare, heat-resistant spores
made by wild-type cells in the presence of 2% 2dGlc were purified and
found to plate with high efficiencies on rich medium with 2dGlc. Thus,
the selection for the ability to produce heat-resistant spores in the
presence of 2dGlc also yields mutants resistant to 2dGlc.
Both early and late development are inhibited by 2dGlc.
The
sporulation of M. xanthus in response to starvation involves
the catabolism of vegetative biopolymers and the anabolism of resting
biopolymers, including the spore coat polysaccharides which protect the
resting cell against harsh environmental conditions. From a metabolic
point of view, many of the mutations that confer defects in the early
stages of M. xanthus development appear to be blocked in
pathways of amino acid catabolism and act during the first 6 h
after the onset of starvation. Presumably, these mutations result in a
decrease of the flow of carbon into and through catabolic pathways that
can feed gluconeogenesis. In contrast, the later stages of development
appear to be involved primarily in gluconeogenic anabolism or in spore
coat assembly, which is fueled by gluconeogenesis (17).
Because we have found that 2dGlc blocks the development of wild-type
cells, we determined at which times during development at which 2dGlc
exerts its inhibitory activity. We might expect, for example, that if
2dGlc6P inhibits glycolysis and glycolysis is required only early in
development, that developing cells would be refractory to 2dGlc after
early developmental stages have been completed. We initiated the
development of wild-type cells on starvation medium without 2dGlc and,
at various times after the initiation of development, transferred
developing cells to starvation medium with 0.1% 2dGlc. Cells were
allowed to develop for a total time of 120 h under a combination
of permissive and restrictive conditions, and spore production was
assayed. We find that if cells are transferred to starvation medium
with 2dGlc within the first 26 h of development, even after
fruiting bodies have formed, 2dGlc inhibits the production of
heat-resistant spores by a factor of at least 1,000-fold (Fig.
7). In contrast, cells transferred to
medium with 2dGlc at 38 h or more after the initiation of
development produce a full complement of heat-resistant spores. This
time coincides with the first appearance of mature, heat-resistant spores on medium without 2dGlc. Thus, 2dGlc blocks the development of
wild-type cells throughout the developmental cycle.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 7.
Sporulation is inhibited by 2dGlc throughout the entire
developmental cycle. Wild-type cells were grown to exponential density
in rich medium, concentrated, spotted on nitrocellulose filters placed
on the surfaces of plates with starvation agar, and incubated at
32°C. At the indicated times after the initiation of development,
filters were removed from TPM plates and placed onto TPM plates
supplemented with 0.1% 2dGlc, and incubation was continued at 32°C.
Development on TPM buffer and then on TPM buffer with 2dGlc was allowed
to proceed for a total duration of 120 h; then filters were heat
treated and assayed for the presence of spores as described in
Materials and Methods. Closed circles, spore titers of developing cells
transferred from TPM agar to TPM agar with 2dGlc; open circles, spore
titers of developing cells maintained on TPM agar.
|
|
The hex mutations decrease the specific activity of a
soluble hexokinase from M. xanthus.
To show that the
hex mutations decrease the specific activity of a soluble
hexokinase, we prepared extracts from wild-type cells and assayed these
extracts for glucose- and ATP-dependent hexokinase activity, using the
coupled assay of Fraenkel and Horecker (12). In this assay,
hexokinase activity is measured as the rate of reduction of NADP
concomitant with the oxidation of Glc6P, the product of hexokinase, by
Glc6P dehydrogenase. As shown in Table 5,
extracts prepared from wild-type M. xanthus cells have considerable hexokinase activity. A similar level of activity is
present in extracts made from the derivative of wild type with integrated plasmid vector pAY703 (mglBA). Extracts prepared
from the derivative of the wild type with plasmid pAY1108
(mglBA-glk) have an approximately 40-fold-higher level of
activity. This result confirms that the E. coli glk gene is
expressed in M. xanthus. In support of this claim, we find
that the apparent Km for ATP of this activity at
pH 9.1 in our assays is about 2 mM (data not shown). This value is
comparable to that of the remarkably high Km
(3.8 mM) that E. coli glucokinase has for this substrate in assays at pH 7.65 (28). In contrast, we find that the
M. xanthus hexokinase from wild-type cells has an apparent
Km of 200 ± 100 µM for ATP.
Extracts made from the 2dGlc-resistant mutants, with the
hex1 and hex3 mutations, have lower levels of
hexokinase activity than the wild type (Table 5). The hex1
mutant, as well as its derivative with plasmid pAY703, expresses about
15% of the level of wild-type activity, whereas the hex3
mutant and its derivative with pAY703 express about 30% of the
wild-type level. Both mutants with plasmid pAY1108 express much higher
levels of activity, presumably because they express the E. coli
glk gene from the constitutive mgl promoter. No
hexokinase activity is detectable in these extracts if glucose is
omitted from the assay (Fig. 1).
There is a simple correlation between (i) the specific activities of
hexokinase activity present in the wild-type and mutant hex1
and hex3 extracts and (ii) the behavior of these strains during growth and sporulation in the presence of 2dGlc (Tables 2 and
4). The wild type, with the highest hexokinase activity, is sensitive
to 2dGlc during both growth and sporulation. The hex3
mutant, with intermediate hexokinase activity, forms small nonmotile
colonies upon growth and cannot sporulate with 2dGlc present. The
hex1 mutant, with the lowest activity, can grow and sporulate well in the presence of 2dGlc. This correspondence between phenotype in vivo and hexokinase activity in vitro shows that resistance to 2dGlc in M. xanthus is due to a decreased
level of hexokinase activity.
Both 2dGlc and GlcNAc, but neither Fru nor Mtl, inhibit the
turnover of Glc by M. xanthus hexokinase.
Although
2dGlc inhibits the growth of wild-type M. xanthus cells, the
hexoses Glc and GlcNAc can antagonize this toxic effect of 2dGlc in
vivo (Table 3). If Glc and GlcNAc act as antagonists of 2dGlc by
inhibiting its phosphorylation by hexokinase in vivo, then both
2dGlc and GlcNAc should be inhibitors of Glc turnover in the reaction
catalyzed by hexokinase in vitro. Therefore, we tested whether
2dGlc and GlcNAc, as well as other hexoses, could inhibit the
phosphorylation of Glc in extracts prepared from wild-type cells. In
these assays, Glc was added to 240 µM (ca. 1.5 × Km), and inhibitors were added to 0.2% (1 to 1.5 mM).
Each hexose tested as a potential inhibitor could decrease measured
activity by acting either as an inhibitor of hexokinase that is not
turned over or as an alternative substrate, because the phosphorylated
products of these potential antagonists are not substrates for Glc6P
dehydrogenase in our coupled assay. As shown in Table
6, both 2dGlc and GlcNAc, but neither Fru
nor Mtl, antagonize the rate of glucose turnover by M. xanthus hexokinase. Curiously, GlcNAc is a more potent antagonist of activity, suggesting that GlcNAc or a derivative metabolite may be
an alternative substrate for M. xanthus hexokinase. In addition, the results in Table 6 show that GlcN also inhibits the
phosphorylation of Glc by hexokinase; it too may be an alternate substrate for this enzyme. Like most known hexokinase activities, the
M. xanthus enzyme is refractile to
-methyl-D-glucopyranoside.
The Dixon plot shown in Fig. 8 reveals
that 2dGlc, as expected, acts as an inhibitor of Glc in our assay. Its
kinetics of inhibition are simply consistent with its predicted role as
an alternate substrate whose phosphorylated product cannot be oxidized by Glc6P dehydrogenase. The initial velocity measured in the hexokinase assay is plotted in Fig. 8 as a function of inhibitor (2dGlc) concentration at several different concentrations of substrate (Glc).
Because the plots at all values of substrate concentration converge at
a single value of -Ki corresponding to
1/Vmax for the enzyme, inhibition is either
competitive or of the linear mixed type (39). Although
competitive inhibition is the simpler model, we cannot as yet exclude
the possibility that both 2dGlc and Glc can bind hexokinase
simultaneously to form an inactive ternary complex.
View larger version (18K):
[in this window]
[in a new window]
|
FIG. 8.
Competitive inhibition of M. xanthus
hexokinase by 2dGlc. Extracts (100 µl) made from M. xanthus wild-type strain DK1622 were incubated in 50 mM Tris-10
mM MgCl2 (pH 9.1) with Glc (160 µM [squares], 200 µM
[open circles], or 300 µM [filled circles]), ATP (10 mM), 500 to
1,000 U of Glc6P dehydrogenase, and Glc6P at concentrations of 0, 1.2, 2.0, 3.0, and 4.6 mM. Results from a representative experiment are
plotted as 1/V (absorbance at 340 nm × 105
min1)1 versus [I], the
concentration (millimolar) of the inhibitor 2dGlc. Plots are linear
least-square fits to the data.
|
|
In extracts made from wild-type M. xanthus cells, we find
hexokinase to have an apparent Km of 170 ± 50 µM for glucose and an apparent Ki of 2 ± 1 mM for 2dGlc. Remarkably, this apparent Ki
that we measure for 2dGlc in vitro is in close agreement with the
concentration of 2dGlc (2.5 mM) required to inhibit the formation of
colonies by M. xanthus in vivo. However, we must be cautious about our interpretation of these kinetic constants, because they were
determined with whole-cell lysates that may include enzymes or
metabolites that adversely affect their accuracy. More accurate measurements of these kinetic constants and of the substrate
specificity of M. xanthus hexokinase await its purification.
| |
DISCUSSION |
We have provided both indirect genetic and corroborating, direct
biochemical evidence that M. xanthus makes a hexokinase that can catalyze the first step in its glycolytic pathway. Currently, we
refer to this enzyme as a hexokinase, not a glucokinase, because it may
phosphorylate substrates in addition to Glc, including GlcN. Studies
are in progress to purify this enzyme by classical methods to test this
directly and to clone the hex structural gene by using
reverse genetics. The isolation and sequence of the hex gene
will set the stage for the high-level expression of its product and for
the purification of sufficient amounts of hexokinase for detailed
kinetic and mechanistic studies. Because hex mutants, which
make lower levels of hexokinase activity, are resistant to 2dGlc, we
are also attempting to clone the hex gene by
complementation. That is, integrative plasmids with M. xanthus inserts that restore 2dGlc sensitivity to hex
mutants may carry the hex structural gene, if the
hex mutations reduce the activity of the hexokinase
structural gene and not a positive regulatory gene.
The main role of hexokinase in M. xanthus metabolism remains
unknown. Clearly, it is not the utilization of polysaccharides as
carbon and energy sources, as for the cellulose-degrading myxobacterial species Polyangium (Sorangium)
cellulosum (38). Hexokinase may play a key role
in the utilization of spore coat polysaccharides during spore
germination. McBride and Zusman (27) have shown that a large
fraction of the carbon in mature spores is present as trehalose, which
is catabolized during germination. Although the hexokinase-deficient
hex mutants make viable spores (which germinate), these
mutants retain partial, residual hexokinase activities (Table 5) that
may suffice for trehalose degradation. These residual activities in
part may be due to the presence of a second hexokinase, a hypothesis
that we are now testing. The closely related species Myxococcus
coralloides has been shown to make two distinct hexokinase
activities (15).
The target and mechanism of 2dGlc toxicity in M. xanthus and
many other organisms also remains unknown. Lee and Cerami
(24) have shown that the accumulation of Glc6P in Hfr
strains of E. coli that carry a plasmid target for mutation
results in a higher mutation rate. They speculated that the
accumulation of adducts formed by Glc6P and other reducing sugars with
DNA may account for this increased mutation rate. On the basis of their
results, one might imagine that 2dGlc6P accumulation may be toxic to
M. xanthus because it leads to DNA damage. However, this
explanation is not satisfying for two reasons. First, unlike the
effects of other DNA-damaging agents, the response of M. xanthus to toxic levels of 2dGlc is reversible for several
generations (Fig. 2) and does not result in an increased frequency of
mutation (unpublished results). Indeed, we now know that the conditions
Lee and Cerami (24) used to elicit the accumulation of Glc6P
in stationary-phase E. coli cells are likely to induce a
hypermutable state, in which events dependent on an F factor lead to an
increased rate of mutation (11, 14, 32). Consistent with
this interpretation, the majority of mutations that they observed were
plasmid rearrangements presumably catalyzed by mobile elements, such as
those residing on the integrated F episome.
Our data indicate that 2dGlc must be phosphorylated to form 2dGlc6P in
order to act as an antibiotic in M. xanthus, as in other
microbes. Preliminary results suggest that the accumulation of 2dGlc6P
leads to the depletion of intracellular ATP in M. xanthus, a
consequence that may account simply for the profound inhibitory effects
of 2dGlc on both the growth and development of wild-type cells.
Therefore, we will focus our future studies on potential enzyme targets
of 2dGlc6P to understand the mechanism of its toxicity at a
physiological level. Our evidence that M. xanthus makes a hexokinase calls into question the prior claim that this organism has
an incomplete glycolytic pathway (46). Currently, we are searching for its putatively missing pyruvate kinase activity. If we
find that M. xanthus has a complete glycolytic pathway, however, we still must contend with a regulatory puzzle. Glc does not
stimulate the growth of this obligate aerobe on defined medium whereas
acetate does, leading us to suspect that the regulation of carbon flow
through the M. xanthus glycolytic pathway during vegetative
growth is governed by a mechanism that is novel among gram-negative
bacteria. On the other hand, this is not surprising, given that
M. xanthus has relegated several pentoses and hexoses to the
roles of signals that can trigger pathways of cellular differentiation
during development.
This work was supported initially by National Institutes of Health
grants GM53392 to P.Y. and AI14176 to M.H.S. and subsequently by
National Science Foundation grant MCB 9808848 to P.Y.
| 1.
|
Angell, S.,
E. Schwarz, and M. J. Bibb.
1992.
The glucose kinase gene of Streptomyces coelicolor A3(2): its nucleotide sequence, transcriptional analysis and role in glucose repression.
Mol. Microbiol.
6:2833-2844[Medline].
|
| 1a.
|
Baumann, P., and L. Baumann.
1975.
Catabolism of D-fructose and D-ribose by Pseudomonas doudoroffii. I. Physiological studies and mutant analysis.
Arch. Microbiol.
105:225-240[Medline].
|
| 2.
|
Bretscher, A. P., and A. D. Kaiser.
1978.
Nutrition of Myxococcus xanthus, a fruiting myxobacterium.
J. Bacteriol.
133:763-768[Abstract/Free Full Text].
|
| 3.
|
Campos, J. M.,
J. Geisselsoder, and D. R. Zusman.
1978.
Isolation of bacteriophage MX4, a generalized transducing phage for Myxococcus xanthus.
J. Mol. Biol.
119:167-178[Medline].
|
| 4.
|
Curtis, S. J., and W. Epstein.
1975.
Phosphorylation of D-glucose in Escherichia coli mutants defective in glucosephosphotransferase, mannosephosphotransferase, and glucokinase.
J. Bacteriol.
122:1189-1199[Abstract/Free Full Text].
|
| 5.
|
Darrow, R. A., and S. P. Colowick.
1962.
Hexokinase from baker's yeast.
Methods Enzymol.
5:226-235.
|
| 6.
|
Downard, J. S., and D. R. Zusman.
1985.
Differential expression of protein S genes during Myxococcus xanthus development.
J. Bacteriol.
161:1146-1155[Abstract/Free Full Text].
|
| 7.
|
Dworkin, M.
1962.
Nutritional requirements for vegetative growth of Myxococcus xanthus.
J. Bacteriol.
84:250-257[Abstract/Free Full Text].
|
| 8.
|
Dworkin, M., and S. M. Gibson.
1964.
A system for studying microbial morphogenesis: rapid formation of microcysts in Myxococcus xanthus.
Science
146:243-244[Abstract/Free Full Text].
|
| 9.
|
Dworkin, M., and W. Sadler.
1966.
Induction of cellular morphogenesis in Myxococcus xanthus. I. General description.
J. Bacteriol.
91:1516-1519[Abstract/Free Full Text].
|
| 10.
|
Filer, D.,
S. H. Kindler, and E. Rosenberg.
1977.
Myxospore coat synthesis in Myxococcus xanthus: enzymes associated with uridine 5'-diphosphate-N-acetylgalactosamine formation during myxospore development.
J. Bacteriol.
131:745-750[Abstract/Free Full Text].
|
| 11.
|
Foster, P. L., and J. M. Trimarchi.
1995.
Conjugation is not required for adaptive reversion of an episomal frameshift mutation in Escherichia coli.
J. Bacteriol.
177:6670-6671[Abstract/Free Full Text].
|
| 12.
|
Fraenkel, D. G., and B. L. Horecker.
1964.
Pathways of D-glucose metabolism in Salmonella typhimurium. A study of a mutant lacking phosphoglucose isomerase.
J. Biol. Chem.
239:2765-2771[Free Full Text].
|
| 13.
|
Frasch, S. C., and M. Dworkin.
1994.
Increases in the intracellular concentration of glycerol during development in Myxococcus xanthus.
FEMS Microbiol. Lett.
122:321-325[Medline].
|
| 14.
|
Galitski, T., and J. R. Roth.
1995.
Evidence that F plasmid transfer replication underlies apparent adaptive mutation.
Science
268:421-423[Abstract/Free Full Text].
|
| 15.
|
Gonzalez, F.,
A. Fernandez-Vivas,
J. M. Arias, and E. Montoya.
1990.
Polyphosphate glucokinase and ATP glucokinase activities in Myxococcus coralloides D.
Arch. Microbiol.
154:438-442.
|
| 16.
|
Hartzell, P., and D. Kaiser.
1991.
Upstream gene of the mgl operon controls the level of MglA protein in Myxococcus xanthus.
J. Bacteriol.
173:7625-7635[Abstract/Free Full Text].
|
| 17.
|
Hartzell, P. L., and P. Youderian.
1995.
Genetics of gliding motility and development in Myxococcus xanthus.
Arch. Microbiol.
164:309-323[Medline].
|
| 18.
|
Hemphill, H. E., and S. A. Zahler.
1968.
Nutrition of Myxococcus xanthus FBa and some of its auxotrophic mutants.
J. Bacteriol.
95:1011-1017[Abstract/Free Full Text].
|
| 19.
|
Kaiser, D.
1979.
Social gliding is correlated with the presence of pili in Myxococcus xanthus.
Proc. Natl. Acad. Sci. USA
76:5952-5956[Abstract/Free Full Text].
|
| 20.
|
Kashefi, K., and P. L. Hartzell.
1995.
Genetic suppression and phenotypic masking of a Myxococcus xanthus frzF-defect.
Mol. Microbiol.
15:483-494[Medline].
|
| 21.
|
Kimsey, H. H., and D. Kaiser.
1991.
Targeted disruption of the Myxococcus xanthus orotidine 5'-monophosphate decarboxylase gene: effects on growth and fruiting-body development.
J. Bacteriol.
173:6790-6797[Abstract/Free Full Text].
|
| 22.
|
Kroos, L.,
A. Kuspa, and D. Kaiser.
1986.
A global analysis of developmentally regulated genes in Myxococcus xanthus.
Dev. Biol.
117:252-266[Medline].
|
| 23.
|
Kroos, L.,
A. Kuspa, and D. Kaiser.
1990.
Defects in fruiting body development caused by Tn5 lac insertions in Myxococcus xanthus.
J. Bacteriol.
172:484-487[Abstract/Free Full Text].
|
| 24.
|
Lee, A. T., and A. Cerami.
1987.
Elevated glucose 6-phosphate levels are associated with plasmid mutations in vivo.
Proc. Natl. Acad. Sci. USA
84:8311-8314[Abstract/Free Full Text].
|
| 25.
|
Lobo, Z., and P. K. Maitra.
1977.
Resistance to 2-deoxyglucose in yeast: a direct selection of mutants lacking glucose-phosphorylating enzymes.
Mol. Gen. Genet.
157:297-300[Medline].
|
| 26.
|
Magrini, V.,
D. Salmi,
D. Thomas,
S. K. Herbert,
P. L. Hartzell, and P. Youderian.
1997.
Temperate Myxococcus xanthus phage Mx8 encodes a DNA adenine methylase, Mox.
J. Bacteriol.
179:4254-4263[Abstract/Free Full Text].
|
| 27.
|
McBride, M. J., and D. R. Zusman.
1989.
Trehalose accumulation in vegetative cells and spores of Myxococcus xanthus.
J. Bacteriol.
171:6383-6386[Abstract/Free Full Text].
|
| 28.
|
Meyer, D.,
C. Schneider-Fresenius,
R. Horlacher,
R. Peist, and W. Boos.
1997.
Molecular characterization of glucokinase from Escherichia coli K-12.
J. Bacteriol.
179:1298-1306[Abstract/Free Full Text].
|
| 29.
|
Mueller, C., and M. Dworkin.
1991.
Effects of glucosamine on lysis, glycerol formation, and sporulation in Myxococcus xanthus.
J. Bacteriol.
173:7164-7175[Abstract/Free Full Text].
|
| 30.
|
Novak, S.,
T. D'Amore, and G. G. Stewart.
1990.
2-Deoxy-D-glucose resistant yeast with altered sugar transport activity.
FEBS Lett.
269:202-204[Medline].
|
| 31.
|
Postma, P. W.,
J. W. Lengeler, and G. R. Jacobson.
1996.
Phosphoenolpyruvate: carbohydrate phosphotransferase systems, p. 1149-1174.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: molecular and cellular biology. ASM Press, Washington, D.C.
|
| 32.
|
Radicella, J. P.,
P. U. Park, and M. S. Fox.
1995.
Adaptive mutation in Escherichia coli: a role for conjugation.
Science
268:418-420[Abstract/Free Full Text].
|
| 33.
|
Reizer, J.,
M. H. J. Saier,
J. Deutscher,
F. Grenier,
J. Thompson, and W. Hengstenberg.
1988.
The phosphoenolpyruvate: sugar phosphotransferase system in gram-positive bacteria: properties, mechanism, and regulation.
Crit. Rev. Microbiol.
15:297-338[Medline].
|
| 34.
|
Sadler, W., and M. Dworkin.
1966.
Induction of cellular morphogenesis in Myxococcus xanthus. II. Macromolecular synthesis and mechanism of inducer action.
J. Bacteriol.
91:1520-1525[Abstract/Free Full Text].
|
| 35. |
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
Abstract
Introduction
