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INTRODUCTION |
D-Ribose is a pentose sugar commonly found as a
component of nucleic acids. It can be utilized by many bacteria as a
sole carbon source. In Escherichia coli, the
sugar is transported via the high affinity RbsACB transporter, a member
of the ABC1-type permeases (1,
2), and phosphorylated by ribokinase (3). Lopilato et al.
(4) showed genetically that at least one low affinity ribose transport
system exists which shares the kinase activity with the high affinity
ribose transporter. Therefore, it has
been believed that a specific low affinity transporter for ribose may
exist, similar to xylE for D-xylose and
araE for L-arabinose. On the other hand, it was
recently reported that D-ribose can be transported through
another high affinity transport system for D-xylose (5) and
D-allose (6), structurally related to D-ribose,
by a mutational derepression allowing an elevated level of transport components.
The bacterial phosphoenolpyruvate:carbohydrate phosphotransferase
system (PTS) couples translocation and phosphorylation for the PTS
sugars, e.g. glucose, mannose, and mannitol. Permeases of
the PTS system differ in their substrate specificity but share the two
general cytoplasmic PTS proteins, the enzymes I encoded by
ptsI and HPr by ptsH, which sequentially transfer
phosphoryl groups from phosphoenolpyruvate to their specific sugars.
The glucose transporter consists of two subunits, IIAGlc
encoded by the crr gene and IIBCGlc encoded by
ptsG. IIAGlc is a hydrophilic protein
phosphorylated in its His-90 residue (7) and is a central regulatory
molecule in catabolite repression. IIBCGlc (PtsG) consists
of the N-terminal domain with eight transmembrane peptides (8) and the
hydrophilic C-terminal domain containing a phosphorylation site at
Cys-421 (9). The sequence of PtsG protein is homologous to that of
N-acetylglucosamine transporter, which has three domains
corresponding to IIBCGlc and IIAGlc in a single
polypeptide, and that of malX gene product, which can
complement ptsG when expressed constitutively (10).
In addition to glucose, PtsG is involved in the transport of other
sugars that are structurally related to glucose, i.e. methyl
-glucoside, 2-deoxyglucose, glucosamine, mannose,
L-sorbose, and 5-thioglucose. Even though transport does
not occur in the PTS system without phosphorylation, there are
mutations in ptsG that allow facilitated diffusion of
glucose without phosphorylation (11) or vice versa (12). Furthermore,
mutation that alters the substrate specificity to allow transport of
mannitol, another PTS sugar, was mapped in one of the transmembrane
helices of PtsG (13). However, nothing is known about the coupling
between phosphorylation and translocation or about the structure
determining substrate specificity. The Mlc protein was at first
reported as a factor affecting glucose transport, causing an enlarged
colony size when overproduced because of a decrease in acetate
production as a result of a reduced glucose utilization (14). It was
recently reported that it is involved in the regulation of several
sugar operons, i.e. manXYZ (14), malT
(15), and ptsG (16, 17). Mlc is a 406-amino acid protein
containing a putative helix-turn-helix motif at its C-terminal residue
that seems to be required for DNA binding involved in a repression.
In searching for secondary D-ribose transporters different
from the high affinity system, we discovered that two mutations were
simultaneously required for the utilization of ribose, one at the
ptsG gene, a glucose-specific PTS permease, and the other at
mlc, a negative regulator of ptsG (16, 17). The
mutations in ptsG alter specificity of substrate to allow
D-ribose transport, whereas the mlc mutation
renders ptsG expression constitutive. As part of
characterizing the mutant phenotype, we first examined the effect of
the mlc mutation on ptsG expression. We further analyzed the ptsG mutations allowing ribose transport in
terms of changes in their substrate specificity to glucose, xylose, and
allose. Patterns of sugar specificities and locations of the mutations
provide insight into an organization of the sugar-binding domains and
their coupling with the phosphorylation domain of PtsG.
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EXPERIMENTAL PROCEDURES |
Bacterial Strains, Plasmids, and Phages--
All strains used
are E. coli K12 derivatives that are listed in Table
I. MC4100 or ZSC112
G was used as a
parent for most strains. Cells were grown in Luria-Bertani medium,
tryptone broth modified from the preparation according to Miller (18),
and H1 minimal medium (19) containing 0.05% ribose, 0.2% glucose, or
0.2% xylose. Antibiotics were added to the growth medium when needed
(ampicillin, 100 µg/ml; kanamycin, 25 µg/ml; chloramphenicol, 30 µg/ml; tetracycline, 15 µg/ml), and for minimal medium, they were
supplied at 50%. The transposon hopping phage,
::TnphoA132, was obtained from B. L. Wanner (20).
DNA Manipulation--
The recombinant DNA techniques were
performed as described by Sambrook et al. (21). A
2.1-kilobase DNA fragment of the chromosome containing the
mlc gene was amplified by polymerase chain reaction. The
primer complementary to the 5' end of mlc created an
XbaI restriction site at 50 bp upstream of the
mlc start codon. The 3' end region was generated with
PvuI, which cuts 600 bp downstream of the mlc
stop codon. The resulting 1.9-kilobase DNA fragment was inserted into
the pUC19 vector previously cleaved with XbaI and
SmaI. The nucleotide sequence was confirmed by DNA
sequencing. The clones were named pOH106 (mlc+)
and pOH108 (mlc-101).
Genetic Procedures--
The P1vir phage was used for
transduction experiments (18). To characterize the ptsG
mutation, the TnphoA132 transposon was used to obtain an insertion
linked to the mutation. Random insertion of the transposon was
generated with
::TnphoA132 for the MC4100 strain from
which P1 lysate was prepared to obtain a pool of 20,000 independent
insertions. Using the P1 lysate, transduction was carried out for
CP1046 showing enhanced growth on ribose, and small colonies that
resulted from an exchange of the mutated gene with the wild type were
isolated on 0.05% ribose minimal plate containing tetracycline.
Auxotrophic mutations were excluded by examining growth on 0.2%
glucose minimal medium. The candidate insertions were tested for their
cotransduction frequencies (Tetr) with the ribose growth
phenotype to measure their linkages. For localization of the insertion
sites, the transposons were transferred to the
polAts strain (22). After appropriate
recombination, the flanking regions of the insertions were
characterized by DNA sequencing and through a data base search with
BLAST. All sequencing reactions were carried out with the DNA sequenase
version 2.0 (U. S. Biochemical Corp.) and ABI PRISMTM
377 DNA sequencer (Perkin-Elmer).
Construction of the ptsG-lacZ Transcriptional Fusion--
A
1074-bp fragment with KpnI ends, comprising part of the
ptsG structural gene and the 455-bp upstream sequence, was
obtained as a polymerase chain reaction product whose sequence was
confirmed by DNA sequencing. The fragment containing the promoter
region of ptsG was cloned into pUC19 using KpnI,
obtained by EcoRI and BamHI digestions, and
cloned into EcoRI and BamHI sites of pRS415, a
lacZ operon fusion plasmid (23), yielding pOH115. The
promoter region was transferred to
RS45 and lysogenized into an
appropriate host. A single-copy
prophage was confirmed by
polymerase chain reaction with three primers recognizing the
attB, attP, and int genes (24). Cells were grown
to an optical density of 0.4-0.8 at A600 in H1
minimal medium (19) containing 0.4% glycerol and 0.2% glucose.
-Galactosidase activity was measured according to the method of
Miller (18). All data were averaged from at least three independent experiments.
DNA Mobility Shift Assay--
CP1035/pOH106
(mlc+) and CP1036/pOH108 (mlc-101)
were grown to late log phase in Luria-Bertani medium containing 100 µg/ml ampicillin. Cells were harvested by centrifugation (5,000 × g) for 10 min, resuspended in 0.05 volume of 10 mM Tris-HCl (pH 8.0), 10% (v/v) glycerol, and 1 mM EDTA and sonicated in ice. The mixture was then
centrifuged to remove insoluble fraction. Protein concentration in the
soluble fraction was measured using the bicinchoninic acid protein
assay reagent (Pierce). The supernatant was kept frozen at
20 °C.
The ptsG promoter region (568 bp from the KpnI to
PvuII sites) was inserted into pUC19 with KpnI
and HincII. The target DNA fragment containing the
ptsG promoter was isolated by EcoRI and
HindIII digestions, which were dephosphorylated with calf intestinal phosphatase and labeled with [
-32P]ATP
using T4 polynucleotide kinase. The assay was carried out in a 20-µl
solution of 20 mM Tris-HCl (pH 8.0), 50 mM KCl,
1 mM EDTA, 1 mM dithiothreitol, and 10%
glycerol with crude extract (0.5-5 µg of protein) mixed with about
0.3 ng of the radiolabeled ptsG promoter DNA and 1 µg of
sonicated salmon sperm DNA as a competitor. The reaction mixtures were
incubated for 20 min at 25 °C and subjected to electrophoresis on
5% polyacrylamide gel (60:1) with 0.5× TBE buffer (45 mM
Tris borate, 1 mM EDTA) at room temperature. The gels were
exposed to x-ray film (Eastman Kodak Co.) at
70 °C.
Uptake Assay--
Cultures grown to late log phase were
harvested and washed three times with equal volumes of KEP buffer (10 mM KH2PO4 (pH 7.0), 0.1 mM EDTA) at 4 °C. Cell density was then adjusted to 2.5 at A600 for
D-[14C]ribose (51.1 mCi/mmol, New England
Biolabs) uptake and to 0.25 at A600 for
D-[U-14C]glucose (310 mCi/mmol, Amersham
Pharmacia Biotech) and for D-[U-14C]xylose
(91 mCi/mmol, Amersham Pharmacia Biotech). Each 0.5-ml aliquot of
culture was stored in ice. After preincubation for about 15 min at
30 °C, 20 µM D-[14C]ribose,
0.2 µM D-[U-14C]glucose, and
0.2 µM D-[U-14C]xylose were
added to the cultures. For an inhibition experiment, unlabeled
competitor sugar (2 mM D-glucose,
D-xylose, and D-allose) was added with
D-[14C]ribose. A 120-µl sample was taken at
10, 20, 30, and 40 s and filtered through a 0.45-µm size
nitrocellulose filter (Amicon). After drying, radioactivities were
counted (25).
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RESULTS |
Screening of Mutations Enhancing Ribose Growth--
Even with a
defect in the high affinity ribose transport, the ribokinase-proficient
E. coli K12 strain can use ribose as a sole carbon source,
albeit less efficiently (4). Previous studies demonstrated that this
growth deficiency could be partially recovered by a regulatory mutation
in the allose (6) or the xylose (5) operons, both of which enhance
expression of the low affinity transporter for ribose. When the high
affinity transport system as well as the low affinity transporter genes
(
als(R-K) and
xylG::TnphoA'-2) were inactivated in OW1, an
E. coli K12 strain, a mutation-enhancing ribose growth still
appeared spontaneously at a frequency of
10
8-10
9. This indicates that there are
some other loci involved in ribose transport.
To characterize those loci, transposon insertions with TnphoA132 were
generated to isolate a tag associated with the mutations. About 20,000 independent insertions in MC4100 were pooled and transduced into the
OW1 mutant showing enhanced growth on ribose. Analysis of the
mutational locations linked to insertions revealed that two chromosomal
loci, one at 36 min and the other at 25 min, are involved in the ribose
growth. The high frequency (10
8-10
9) of
the original mutation, which lowers the possibility of a double
mutation, led us to suspect that one of the mutations might already be
present in the parental strain. Indeed, the mutation at 36 min was
resident in the OW1 strain, although it was not found in other E. coli strains such as MC4100 and W3110.
Further characterization of the mutations revealed that the mutation at
25 min was found in ptsG, encoding the glucose transporter of the PTS system. A total of seven independent mutants was sequenced, in which five different mutational changes were observed: one for F37Y,
G176D, and G281D and two for I283T and L289Q. Repeated occurrence of
the mutations indicates an apparent saturation of the changes. F37Y and
G176D were found in the predicted periplasmic loops, whereas G281D,
I283T, and L289Q are in the transmembrane region (Fig.
1). The mutation at 36 min was
characterized to have a change in mlc that has been
implicated in the regulation of ptsG (16, 17). This allele
in OW1 was named mlc-101 and was sequenced to have C instead
of T at the first base of Gln-369, causing an introduction of the UAG
stop codon.

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Fig. 1.
Locations of mutations in the PtsG
protein. Mutations transporting D-ribose were found in
residues represented by shaded circles. Residues in
open circles are mutations allowing facilitated diffusion
without phosphorylation (11). Residues in open boxes are
mutations that result in poor translocation with phosphorylation (12).
The residue in the shaded box has a change with altered
substrate specificity toward D-mannitol (13).
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Effect of the mlc-101 Mutation on ptsG Expression--
To test
whether ribose transport through the mutated PtsGs results from an
altered level of the transporter, as reported in the Als and Xyl
systems (5, 6), we examined a transcriptional effect of Mlc-101 on
ptsG expression by using DNA mobility shift assay and the
ptsG-lacZ fusion. The 568-bp
KpnI/PvuII fragment containing the
ptsG promoter region was obtained from pOH116 and mixed with
crude extract prepared from CP1035/pOH106 (mlc+)
and CP1036/pOH108 (mlc-101). A band shift was observed only with the wild-type Mlc but not with the truncated Mlc-101 (Fig. 2). This observation is consistent with
the ptsG-lacZ fusion analysis and uptake assay of
D-[14C]glucose in which the ptsG
transcription was derepressed at about 7-fold, whereas the uptake rates
were increased about 33-fold in mlc-101 background (data not
shown). Because Mlc-101 lacks the C-terminal peptide containing a
putative DNA binding motif, it seems likely that the protein loses its
function as a repressor.

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Fig. 2.
DNA mobility shift assay for Mlc and Mlc-101
proteins. About 0.3 ng of DNA of the end-labeled ptsG
promoter region (568 bp from the KpnI to PvuII
sites) was mixed with 0.5, 1, and 5 µg of crude extract from
CP1035/pOH106 (Mlc+) or CP1036/pOH108
(Mlc-101) as described under "Experimental
Procedures." About 6 ng of DNA of the unlabeled fragment of the same
region was used as a competitor.
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Effect of the ptsG Mutations on Ribose Transport--
Growths of
the various ptsG mutants were examined on minimal plate with
0.05% ribose, rates of which were comparable with that of the
Rbs+ strain and found to be correlated with the uptakes of
[14C]ribose (Table II).
Among them, I283T, located in the transmembrane region, exhibited the
highest growth and uptake rates. Although the uptake rates through the
mutated PtsG were far less than that of the Rbs transport system, they
were considerably better than the transport through the Als or Xyl
systems, low affinity transporters for D-ribose (5, 6).
To investigate the effect of PtsG mutations on glucose transport,
growth and uptake rates for glucose were measured. Growths showed
little differences from that of wild-type PtsG (data not shown),
whereas the uptake rates were reduced (Table II). Because the tested
strain contained the manZ and glk mutations, the
coupling between transport and phosphorylation must occur in the mutant proteins (26). In general, the three transmembrane mutations reduce the
specificity toward glucose more than the two periplasmic mutations,
which is inversely correlated with the affinities to ribose. The I283T
mutation with the highest ribose specificity reduced the uptake of
glucose at about 50% of the wild-type PtsG, suggesting that at least
the I283 residue, and perhaps other mutated residues, plays a key role
in determining the substrate specificity.
Unlike glucose, ribose transport through PtsG occurs without
phosphorylation. Thus, it seems possible that PTS and non-PTS sugars
might use different mechanisms for their transports. However, the
ribose transport through PtsG is inhibited by the presence of glucose,
xylose, and allose as shown in Table II. The glucose effect is fairly
strong, whereas D-xylose exerts a weaker inhibition. Unexpectedly, D-allose shows some inhibitory effects on
ribose transport although the sugar was not transported by the mutated PtsG (data not shown). The apparent diversity of the inhibition pattern
may reflect a multiplicity of substrate binding specificity of the transporter.
RbsD and Ribokinase Are Indispensable for D-Ribose
Transport through PtsG--
To test whether D-ribose
transport through PtsG is coupled to phosphorylation, the requirement
of ribokinase was assessed for F37Y. When the whole rbs
operon was deleted (
rbsD-R), the mutant no longer
maintained its enhanced growth phenotype on ribose (Table
III), indicating that some rbs
component(s) is required. An addition of rbsK on the pYP60
plasmid, however, did not entirely restore the enhanced growth
phenotype. After testing other components of the rbs operon,
we discovered that the maximal level of enhanced growth and transport
was observed when both rbsD and rbsK were expressed. The rbsD gene is a part of the rbs
operon, which codes for a protein with unknown function. The results
indicate that the transported ribose is phosphorylated not by the PTS
system but by ribokinase and that RbsD somehow plays a critical role in
the PtsG-mediated ribose transport. The requirement of rbsD was also demonstrated in other ptsG mutants and in other low
affinity transporters, such as the Xyl and Als systems (data not
shown).
D-Xylose Can Be Transported through PtsG in the
mlc
Background--
Because it was
demonstrated that ribose transport can be mediated by the Als and Xyl
systems, we tested whether D-xylose is recognized by the
mutated PtsG. The ribose uptake through these transporters appears to
be based on the structural similarities of the sugars to
D-ribose as shown in Fig. 3.
Interestingly, it was found that xylose can be transported even through
the wild-type PtsG only when there is a mutation in mlc,
indicating that the xylose transport through PtsG is
dose-dependent in terms of the level of transporter (Table
IV). The experiment was done in the strain with xylF::TnphoA'-1, an insertion in the
gene coding for the xylose-binding protein that inactivates the high
affinity transporter for xylose. The OW1 strain that has
xylA (xylose isomerase) and mlc mutations did not
grow on xylose minimal plate in the absence of xylFGH (data
not shown), indicating that as in ribose transport, xylose is
phosphorylated not by the PTS system but by xylulose kinase encoded by
xylB. The presence of the ribose-specific mutations tends to
reduce the xylose uptake rate in both the periplasmic and transmembrane
mutants. The impairments of glucose uptake (Table II) are more notable
in the membrane mutants than in the periplasmic ones. The degree of
reduction, I283T > G281D > L289Q for both xylose and
glucose, appears to be correlated with the specificity changes to
ribose.

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Fig. 3.
Structure of
D-glucose. The differences of
configurations found in other sugars that are transported
through PtsG are also represented. D-Ribose and
D-xylose are shown in their pyranose forms.
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DISCUSSION |
We report here that in addition to Rbs, Als, and Xyl transporters,
D-ribose can be transported through PtsG when mutated
specifically and derepressed by an inactivation of mlc,
encoding a negative regulator of ptsG (16, 17). Moreover, we
found that D-xylose, structurally analogous to
D-glucose and D-ribose, is transported through
PtsG in the absence of Mlc. D-Ribose exists primarily as a
pyranose form in solution as shown in Fig. 3. It is the one interacting
with the periplasmic binding protein (27). This form shows similarity
to D-allose, D-glucose, and
D-xylose, explaining the cross-specificity between these
sugars in sharing the transporters.
Besides glucose, PtsG is involved in the transport of other sugars that
are structurally related to glucose such as mannose, 2-deoxyglucose,
glucosamine,
-methylglucoside, L-sorbose, and 5-thioglucose (Fig. 3). Among these, the first three sugars differ from
glucose at their C-2 positions, indicating flexibility in recognizing
the configuration around C-2. The same is true for
-methylglucoside
at the C-1 position and 5-thioglucose at C-5. D-Mannitol
(1-deoxymannose) was reported to be taken up through PtsG when Gly-320
is mutated to valine and thus modified to recognize the change of
D-mannose at C-1 (13). Despite the differences in
structures, the glucose analogs have some common features; they can be
transported and phosphorylated by PtsG, which is not the case in
D-ribose and D-xylose, which require their own
kinases for recognizing a furanose form of sugar. The lack of
phosphorylation coupling in transport of these sugars might lie in the
fact that the hydroxyl group at C-5, normally exposed in the furanose
form, is hidden in the transported sugar. The necessity of
mlc mutation for the xylose transport, not found in the
transport of other phosphorylated glucose analogs, may indicate that an
uptake of nonphosphorylated sugar through PtsG requires a higher level
of gene expression.
In addition to the C-6 hydroxyl group accepting phosphorylation,
D-ribose differs from D-glucose in the hydroxyl
group at C-3. Unlike D-ribose, the pyranose form of
D-xylose has exactly the same configuration as
D-glucose at all carbon positions except for C-6, a site
for phosphorylation. Therefore, it seems likely that the configuration
at C-3 is a major specificity determinant, more so than the ones at
C-1, C-2, or C-6, which is consistent with the fact that an additional
mutation is required for the transport of D-ribose and not
for D-xylose. The tendency of the ribose-specific mutations
toward losing specificities to xylose and glucose is exemplified in the
extreme case of I283T. A slight deviation from this general trend is
illustrated in the mutations localized in the periplasmic domain in
which the glucose specificities were not substantially reduced. This
may reflect the differences between the mutations in two locations,
which might form two different pores with altered specificities. It is
conceivable that a unique feature in the periplasmic pore lies in its
discrimination of the C-6 residue. In fact, even though both glucose
and xylose are transported, the xylose uptake rate of
ptsG+ was about half that of glucose, even under
a derepressed condition (Table II), which may indicate that the
phosphorylation of sugar affects transport, perhaps through an
intramolecular signaling. In other words, there might be a coupling
between transport and phosphorylation which is likely to be manifested
by a structural change in PtsG during sugar phosphorylation.
The possibility of two pores in PtsG could also be discussed in terms
of its structural organization. Unlike the ABC transporter, the PTS
system has no periplasmic sugar-binding protein, which may explain the
fact that PTS has broader substrate specificity than the ABC
transporters. It seems likely that in PtsG, the periplasmic region as
well as the transmembrane region confers substrate specificities, and
the absence of a binding protein may be functionally compensated by the
presence of the periplasmic specificity domain, which might be
explained by a model proposed here as the "two-sieve mechanism." A
similar mechanism was reported to occur in the potassium channel, in
which both the narrow selectivity filter recognizing K+
located on the outer surface and the wider, hydrophobic inner pore
structure stabilizing a cation exist (28).
It is of interest to note that the D-mannitol-specific
mutation of G320V was found in helix VIII (13), and three other
mutations involved in facilitated diffusion, including ours, were found in helices VI and VII (11). These observations suggest that these
helices form a channel for sugar specificity and transport. It was
thought that the hydrophobic pocket plays an important role in sliding
of sugar during translocation through the channel of PtsG. Also in the
potassium channel, the lining of hydrophobic amino acids in the channel
was suggested to facilitate a final release of ligand by loosening an
association with the substrate (28). In this regard, a change into
hydrophilic amino acid as found in our mutations, especially ones
located in the transmembrane region, may elicit a negative effect on
phosphorylation-mediated glucose transport but a positive effect on
ribose transport, perhaps occurring as facilitated diffusion.
During the study of the Rbs component involved in ribose
transport though PtsG, we found that RbsD is required in addition to
ribokinase. RbsD was originally proposed as a member of the membrane
permease associating with RbsC (29). However, it apparently lacks a
region predicted to be in the membrane. Furthermore, a mutation in RbsD
does not abolish ribose
transport,2 suggesting that
it is not directly involved in the permease function. There is no
functional homolog found for RbsD based on its sequence similarity. The
only clue about its function is that it enhances the utilization of
ribose when the sugar is transported through a low affinity
transporter. In other words, the role of RbsD is independent of
specific types of transporters, implying that RbsD is involved in the
step after membrane transport, such as acceleration of the ribose
metabolism. The function of RbsD is currently under investigation.