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Cell Biology
The BglF sensor recruits the BglG transcription regulator to the membrane and releases it on stimulation
*Department of Molecular Biology, The Hebrew
University–Hadassah Medical School, P.O. Box 12272, Jerusalem 91120,
Israel; andDepartment of Molecular Biology and
Microbiology, Tufts University Health Sciences Campus, Boston, MA 02111
Edited by Charles Yanofsky, Stanford University, Stanford, CA and approved March 17, 2003 (received for review December 15, 2002)
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Abstract |
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-glucoside availability. BglF is a
membrane-bound protein, whereas BglG is a soluble protein, and they are both
present in the cell in minute amounts. How do BglF and BglG find each other to
initiate signal transduction efficiently? Using bacterial two-hybrid systems
and the Far-Western technique, we demonstrated unequivocally that BglG binds
to BglF and to its active site-containing domain in vivo and in
vitro. Measurements by surface plasmon resonance corroborated that the
affinity between these proteins is high enough to enable their stable binding.
To visualize the subcellular localization of BglG, we used fluorescence
microscopy. In cells lacking BglF, the BglG-GFP fusion protein was evenly
distributed throughout the cytoplasm. In contrast, in cells producing BglF,
BglG-GFP was localized to the membrane. On addition of -glucoside,
BglG-GFP was released from the membrane, becoming evenly distributed
throughout the cell. Using mutant proteins and genetic backgrounds that impede
phosphorylation of the Bgl proteins, we demonstrated that BglG-BglF binding
and recruitment of BglG to the membrane sensor requires phosphorylation but
does not depend on the individual phosphorylation sites of the Bgl proteins.
We suggest a mechanism for rapid response to environmental changes by
preassembly of signaling complexes, which contain transcription regulators
recruited by their cognate sensors-kinases, under nonstimulating conditions,
and release of the regulators to the cytoplasm on stimulation. This mechanism
might be applicable to signaling cascades in prokaryotes and eukaryotes.
-glucosides, most transcripts terminate prematurely at one of two
-independent terminators within the operon; in the presence of an
inducer, BglG allows transcription through these sites by binding to the
bgl transcript
(3–5).
BglG also recognizes and interacts with the ' subunit of E.
coli RNA polymerase (6).
BglF regulates BglG activity by reversibly phosphorylating it depending on
-glucoside availability
(7–9),
thus modulating its dimeric state
(10). Interestingly, BglF, an
enzyme II of the phosphoenolpyruvate-dependent phosphotransferase system (PTS)
that catalyzes transport and phosphorylation of -glucosides, uses the
same active site residue, Cys-24, to phosphorylate the sugar and BglG
(11) and to dephosphorylate
BglG (12). Homologues of the
Bgl proteins have been identified in many bacterial species
(13). Activity of these
proteins as antiterminators is regulated by the PTS and for some, negative
regulation by BglF-like sugar phosphotransferases has been demonstrated
(14).
BglF is a membrane-bound sensor, whereas the BglG transcriptional regulator
is soluble, and both are present in small amounts in uninduced cells. How do
BglF and BglG find each other upon inducer-mediated stimulation of BglF? A
possible mechanism, that would ensure an efficient response to environmental
changes, is recruitment of BglG to the membrane by BglF and rapid release of
BglG to the cytoplasm on stimulation. To test this hypothesis, we took several
experimental approaches. We first asked whether BglG binds to BglF. Using the
Far-Western technique and bacterial two-hybrid systems, we demonstrated that
BglG interacts with BglF and with its active site-containing domain both
in vitro and in vivo. Using surface plasmon resonance (SPR),
we showed that the affinity of BglG for the active site containing domain of
BglF is strong (KD 
3 x 10-7) and
supports the idea of stable binding. Using fluorescence microscopy, we showed
that BglG fused to GFP is located at the cell membrane only in the presence
BglF and is released to the cytoplasm after addition of -glucosides.
Neither Cys-24 of BglF, which reversibly phosphorylates BglG, nor the BglG
phosphorylation site His-208 is crucial for BglF–BglG interaction.
However, phosphorylation is required for the interaction, as indicated by the
lack of interaction and recruitment of BglG in a 
pts strain,
or when both proteins lack phosphorylation sites.
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Materials and Methods |
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pts derivative of SU202
(15) was isolated by
transducing SU202 to TetR by using P1 phage grown on PPA310 that
harbors a deletion of the pts operon, obtained from P. Postma
(University of Amsterdam). A pts derivative of MG1655 was
isolated by transducing MG1655 to KanR using P1 phage grown on
TP2811 (16). Chemicals. N-hydroxysuccinimide, N-ethyl-N-(3-diethylaminopropyl) carbodimide, ethanolamine hydrochloride, and HBS buffer (10 mM Hepes, pH 7.4/150 mM NaCl/3.4 mM EDTA/0.005% P-20) were obtained from BIAcore AB (Uppsala).
Plasmids. All plasmids used in this study and the proteins they encode are listed in Table 3, which is published as supporting information on the PNAS web site, www.pnas.org. Construction of plasmids is provided as supporting information. Plasmids used for the LexA-based two-hybrid system were derived from pMS604 and pDP804 (15). Plasmids used for Far Western and SPR were constructed by cloning the bglF and bglG alleles in pET15b (Novagen) or in pST6#1, a derivative of MBPL–/gp21(338-445) (17), obtained from P. Poumbourios (St. Vincent's Institute of Medical Research, Victoria, Australia). Plasmids used for fluorscence microscopy: pJS185 encodes BglG-GFP; the bglF alleles were cloned in pBAD18 (18).
Two-Hybrid Analysis. SU202 strain and its pts
derivative were cotransformed with the derivatives of pDP804 and pMS604. Cells
were grown until midlogarithmic phase at 37°C in M63 medium
(19) containing succinate
(0.4%) as a carbon source and 0.5 mM isopropyl -D-thiogalactoside.
Interaction between the two hybrid proteins was monitored on
MacConkey–maltose indicator plates and quantitated by
-galactosidase assays
(19).
Affinity Chromatography. MBP-BglG (MBP, maltose-binding protein) was purified as described in ref. 11. His-tagged proteins were expressed in BL21(DE3) and purified as described (20), except that the extracts were incubated for only 1h with the Ni-NTA resin. For His-tagged BglF, disruption of cells was performed by resuspending pelleted cells in lysis buffer (30 mM Tris·HCl, pH 8/20% sucrose/10 mM EDTA), supplemented with 50 µg/ml lysozyme. After 15 min incubation at 37°C, DNase (5 µg/ml) and MgCl2 (15 mM) were added, and the extract was incubated for an additional 15 min. After centrifugation at 20,800 x g in the cold, the membrane fractions were resuspended in 2 ml of PBS buffer (80 mM Na2HPO4/20 mM NaH2PO4/100 mM NaCl) containing 0.5% SDS and 6 M urea.
Far-Western Analysis. Proteins were separated on 10% SDS-polyacrylamide gels. Gels were subjected to Far-Western analysis as described (6) or stained with Coomassie blue.
SPR. SPR measurements were preformed with a BIAcore3000 system (BIAcore, Uppsala, Sweden). All procedures were performed at 25°C. His-tagged WT BglG protein and its mutants (5 µg/ml) in 10 mM sodium acetate, pH 3.5, were immobilized on the dextran surface of CM5 sensorchips by the standard amino coupling method (21), except that 70 µl of the activators N-hydroxysuccinimide, N-ethyl-N-(3-diethylaminopropyl) carbodimide, and the blocker ethanol amine were injected at a flow rate of 10 µl/min. One flow cell with no coupled protein served as reference. Once the His-tagged BglG proteins were immobilized, serial dilutions ranging from 1.25 to 10 µM of IIBbgl or IIBbgl (C24S) were injected at a flow rate of 30 µl/min in HBS buffer containing 1 mg/ml carboxy methyl dextran to minimize nonspecific interactions. A sample injection of 90 µl was followed by buffer flow for 3 min for dissociation. Regeneration was achieved by a 20-µl pulse of 50 mM phosphoric acid. Binding constants were determined by using BIAEVALUATION software. The KD values for the interaction of BglG with IIBbgl were calculated from the apparent kinetic constants by fit to a first-order kinetic model. Values were estimated from at least three independent experiments, using different chips and protein preparations.
Fluorescence Microscopy and Photography. MG1655 and its
pts derivative were transformed with the indicated plasmids.
Cells were grown to early log phase at 30°C in either LB or M9 medium
(19) containing 0.4% glycerol.
Expression of the bglF alleles, cloned in pBAD18, was induced by
adding 0.1% arabinose for 1 h. Expression of the BglG-GFP fusion was not
induced, to keep the level of the fusion protein low. When indicated, 0.5%
arbutin was added to the growing culture and BglG-GFP localization was
examined at 5-min intervals, after the addition of arbutin, for 2 h.
Microscopy was performed by using an Olympus (Melville, NY) BX60 microscope with an Olympus PlanApo 100 x 1.4 n.a. oil-immersion objective. An Olympus MWIB photocube transmitting a wavelength of 460–490 nm was used to stimulate GFP fluorescence. Images were obtained by using cells that had been washed and resuspended in saline and placed on a microscope slide under a coverslip. Pictures were obtained by using a Hamamatsu Photonics (Hamamatsu City, Japan) C4742-95 digital charge-coupled device camera and an automatic light shutter (MAC2000, LudI Electronics, Hawthorne, NY). The exposure time was the same in all cases. OPENLAB 3.02 imaging software (Improvision, Lexington, MA) controlled image acquisition and control of the light shutter, as well as colorization of the greyscale images acquired.
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To test whether BglG binds to BglF in vivo, we used the LexA-based bacterial two-hybrid system (15). In this system, the proteins of interest are fused either to a WT LexA repressor DNA-binding domain (LexADBD) or to an altered specificity LexADBD and introduced into a strain that harbors a chromosomal copy of lacZ under the control of a LexA hybrid operator (SU202). Transcriptional repression is achieved on coexpression of both hybrid proteins, provided they bind to each other. The combination of IIBbgl fused to the WT LexADBD and BglG fused to mutant LexADBD resulted in a stable, biologically active heterodimer, as indicated by the 76% transcriptional repression (Table 1). The leucine zipper domains of Fos and Jun fused to the WT and mutant LexADBD, respectively, and chimeras between BglG and the two LexADBD served as positive controls (98% and 97% repression), and the two LexADBD served as a negative control (0% repression) (Table 1). Similar results were obtained with the adenylate cyclase-based two-hybrid system (22) (see Supporting Text, which is published as supporting information on the PNAS web site).
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We anticipated that fusion of the full-length BglF protein to LexADBD would result in a membrane-anchored protein, which might not bind DNA efficiently. Nevertheless, we fused BglF to the mutant LexADBD, expressed from a reduced copy number plasmid, to avoid overproduction. Coexpression of this fusion with BglG fused to WT LexADBD yielded 59% repression (Table 2). Therefore, despite the constraint caused by anchoring one of the repressor monomers to the membrane, repression was evidently achieved, corroborating BglG–BglF interaction.
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Taken together, the results with the two-hybrid system demonstrate that BglG interacts with the active site-containing domain of BglF, as well as with the entire BglF protein, in vivo.
Kinetic Analysis of BglG–IIBbgl Interaction. To study the kinetics of the interaction between BglG and IIBbgl, we used SPR, which enables the measurement of rates of association (ka) and dissociation (kd). BglG was immobilized on a sensorchip, leaving one flow-cell blank, and various concentrations of IIBbgl were passed over the chip. The response from the reference surface was subtracted from the response in the channel with BglG to give a signal (resonance units) that is directly proportional to the amount of bound compound. As can be seen in Fig. 2A, BglG and IIBbgl gave binding signals in a concentration-dependent manner. The binding signal decayed very slowly after completion of the injection, indicating that the complex is highly stable. An equilibrium constant (KD) value of 2.93 x 10-7 was calculated for the BglG–IIBbgl complex. The high affinity between BglG and IIBbgl is characterized by a low dissociation rate constant (kd 2.43 x 10-4·sec-1).
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BglG Is Recruited to the Cell Membrane Only When BglF Is Expressed in the Cell and Is Released from the Membrane After Addition of the Stimulating Sugar. Does the affinity between the BglG transcriptional regulator and the BglF membrane-bound sensor lead to recruitment of BglG to the cell membrane? Previous results suggested that at least a fraction of the cellular BglG associates with the membrane (O.A.-C., unpublished data; ref. 23). Using fluorescence microscopy, we examined the subcellular localization of a BglG-GFP fusion. This fusion protein is fully functional as a transcriptional antiterminator (results not shown). As shown in Fig. 3A, BglG-GFP was evenly distributed in cells lacking BglF. In contrast, in cells producing BglF, BglG-GFP was observed mainly as a bright ring around the periphery of the cell (Fig. 3B). These results show that BglG-GFP is tethered to the cell membrane only in the presence BglF. When the soluble domain of BglF, IIBbgl, was expressed in the cells, instead of BglF, BglG-GFP was not seen associated with the membrane (Fig. 3C).
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In the experiment presented in Fig. 3, cells were grown in a rich medium. Cells overproducing BglF were longer than normal (Fig. 3, compare B with A and C), a phenomenon frequently observed in cells overproducing membrane proteins. When the experiment was repeated with cells grown in a minimal medium, BglG-GFP was also detected mainly at the periphery of BglF-producing cells (Fig. 4A). However, in this case, cell size was normal, presumably due to lower levels of BglF produced under these conditions. Hence, the subcellular localization of the BglG-GFP is not related to cell length. BglG-GFP in cells lacking BglF or producing IIBbgl that were grown in minimal medium was evenly distributed as in cells grown in rich medium (data not shown).
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Is BglG released to the cytoplasm on stimulation? To test this possibility,
we grew cells in minimal medium containing glycerol as a sole carbon source
and examined the subcellular localization of BglG-GFP at 5-min intervals for a
period of 2 h after the addition of the -glucoside arbutin to the
medium. The fluorescent ring, observed at the cell periphery before arbutin
addition (Fig. 4A),
disappeared 15 min after arbutin addition, and the BglG-GFP became uniformly
distributed within the cells (Fig.
4B) and remained so for the duration of the experiment.
These results imply that BglG is released from the membrane to the cytoplasm
shortly after addition of the stimulating sugar. To see whether BglG can
relocalize to the membrane when the conditions are not in favor of
-glucoside utilization, we grew the cells in minimal medium and added
arbutin for 30 min and then glucose for 30 more minutes. The fluorescent ring
reappeared, indicating that BglG-GFP was retethered to the membrane (data not
shown).
Taken together, the results obtained by fluorescence microscopy demonstrate
that BglG is recruited to the cell membrane, provided that BglF is present in
the membrane. BglG is released to the cytoplasm after stimulation of BglF with
-glucosides and stays there as long as BglF remains stimulated.
Requirements for BglF-BglG Binding and Recruitment of BglG to the Membrane. To study the requirements for the interaction between BglG and BglF, we examined the effects of mutations in BglG and BglF that impair phosphorylation on their interaction. In BglF, a mutation (C24S) in the active site prevents BglG phosphorylation and negative regulation (11). In BglG, a mutation (H208R) in the site that is phosphorylated by BglF prevents BglG regulation and phosphorylation by BglF (24). Two other mutations in BglG, D100N and H160Y, abolish negative regulation of BglG by BglF in vivo, and either severely reduce or completely abolish BglG phosphorylation by BglF in vitro, respectively (7). The three mutant BglG proteins are constitutively active in vivo.
Using the Far-Western technique, we found that the three mutant BglG proteins, each fused to MBP and immobilized on a nitrocellulose filter, gave a signal similar to that seen with WT MBP-BglG when incubated with His-IIBbgl and antibodies against the histidine tag (Fig. 1C). When a filter-immobilized WT MBP-BglG was probed with His-IIBbgl(C24S) and anti-His antibodies, a strong signal was observed (Fig. 1D, lane 1), comparable with the signal observed with His-tagged WT IIBbgl as a probe (Fig. 1A, lane 1). Hence, replacement of the phosphorylated residues on BglG and IIBbgl, as well as other residues in BglG that are required for its phosphorylation by BglF, does not seem to have an effect on BglG–IIBbgl interaction in vitro. However, when we tested the interaction between mutants of BglG and IIBbgl, both lacking their phosphorylation sites, i.e., filter immobilized MBP-BglG(H208R) probed with His-IIBbgl(C24S), a very weak signal was obtained (Fig. 1D, lane 2). It is important to mention that after purification, a fraction of the WT proteins is phosphorylated, as demonstrated by acrylamide-urea gel analysis (data not shown).
The effect of the above mutations on BglG-IIBbgl and BglG-BglF binding in vivo was examined using the LexA-based two-hybrid system. As shown in Tables 1 and 2, all four mutant proteins repressed transcription when expressed in combination with a WT partner protein, although the interaction of BglG(D100N) with IIBbgl or BglF and of BglG(H160Y) with BglF was somewhat reduced. However, the combination of IIBbgl(C24S) with BglG(H208R) reduced the interaction more profoundly (52% repression compared with 76% given by WT IIBbgl with WT BglG; Table 1). The combination of BglF(C24S) and either BglG(H208R) or BglG(H160Y) showed almost no interaction (14% and 13% repression, respectively). Hence, the results concerning the effect of the above mutations on the interaction between the studied proteins, obtained in vitro and in vivo, are in accord.
Can a mutant BglF protein that lacks the C24 phosphorylation site recruit WT BglG to the membrane? As shown in Fig. 3D, BglG-GFP was recruited to the membrane in cells expressing BglF(C24S).
Using SPR, we were able to quantify the effect of the mutations in BglG and BglF on BglG–BglF interaction. The three BglG mutant proteins were immobilized on the sensorchip, and IIBbgl, at various concentrations, was injected over these channels. A single sensorgram curve, obtained at 10 µM of IIBbgl, is presented for each channel (Fig. 2B). The binding profiles, the association and dissociation constants, and the KD values obtained for IIBbgl binding to the three BglG mutants were very similar to those obtained for IIBbgl binding to WT BglG (Fig. 2B).
A different binding profile was observed for the interaction between BglG and IIBbgl(C24S) (Fig. 2C, blue curve). The binding signal decayed rather rapidly, compared with the complex of WT IIBbgl and BglG, indicating that the BglG-IIBbgl(C24S) complex is less stable. Although kd and ka values could not be calculated accurately from the curve obtained with IIBbgl(C24S), curve simulations enabled prediction of values around 10-2·sec-1 for kd, and around 104·M-1·sec-1 for ka, compared with 10-4·sec-1 and 103·M-1·sec-1 for kd and ka of BglG-IIBbgl, respectively. These values suggest that IIBbgl(C24S) associates with and dissociates from BglG more rapidly than WT IIBbgl. Because both association and dissociation are faster, it is possible that at equilibrium the amount of IIBbgl(C24S) bound to BglG is comparable with the amount of IIBbgl. A visual comparison of the binding response obtained for BglG-IIBbgl and BglG-IIBbgl(C24S) shows a higher response level in the first case. Because the height of the plateau of the curves reflects the amount of the analyzed proteins bound to the BglG bearing channel, it is quite evident that the C24S mutation reduced the affinity toward BglG to 54% but did not abolish it (compare Fig. 2 A and B, blue curves). The binding profiles obtained for the interaction of IIBbgl(C24S) with the BglG mutants impaired in phosphorylation are similar to the profile of WT BglG with IIBbgl(C24S), i.e., fast association and dissociation. However, the affinity between IIBbgl(C24S) and the mutant BglG proteins, calculated on the basis of the height of the plateau of the curves, declined in comparison to the affinity between a pair of WT proteins or between a WT protein and a mutant protein: 33% for BglG(H160Y)-IIBbgl(C24S), 29% for BglG(D100N)-IIBbgl(C24S), and as low as 17% for BglG(H208R)-IIBbgl(C24S) (Fig. 2C, green, purple, and orange curves, respectively).
Our results imply that each individual phosphorylation site of the interacting proteins, BglG and BglF, is not crucial, by itself, for their binding. Yet, binding requires the presence of one phosphorylation site, either on BglG or on BglF, raising the possibility that the presence of a phosphoryl group on one of the interacting partners is essential for the binding.
We tested the significance of phosphorylation for BglG–BglF
interaction by two experimental approaches. First, we tested the interaction
between the Bgl proteins, WTs or mutants, in the LexA-based two-hybrid system
in a pts background, in which the Bgl proteins are not
phosphorylated. No interaction was observed in all cases, as indicated by the
negligible or no repression (Table
2, compare values obtained in pts+ and
pts isogenic strains).
Next, we tested the importance of phosphorylation for tethering BglG to the
membrane. In a pts background, BglG-GFP was poorly, if at all,
recruited to the membrane when coexpressed with WT BglF or with BglF(C24S),
looking similar to its appearance in pts cells expressing
IIBbgl or not expressing any BglF derivative
(Fig. 3 E–H).
The lack of binding between BglF and BglG, when both are nonphosphorylated, is
in accord with the idea that when the sugar dephosphorylates BglF, the latter
dephosphorylates BglG, releasing it to the cytoplasm to act as a
transcriptional antiterminator.
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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The release of BglG from the membrane on stimulation is not absolutely
required for its activity, although it presumably increases transcription
efficiency. Bacterial DNA-binding proteins, such as ToxR in Vibrio
species and CadC in E. coli, function as transcription activators
despite their anchoring to the membrane
(31,
32). Indeed, when BglG was
artificially anchored to the membrane, it could function as a transcriptional
antiterminator (23).
Nevertheless, our results demonstrate that this it not the case with the
native BglG. We show here that, after BglF stimulation by -glucosides,
BglG is released to the cytoplasm and stays there as long as BglF remains
stimulated. Because interaction of BglF with the stimulating sugar changes the
conformation of BglF (33),
this change may induce not only BglG dephosphorylation but also BglF-BglG
separation. The release of factors that are involved in transcription from the
membrane, followed by their translocation to the transcription machinery, is
achieved by various strategies, including activation-induced phosphorylation,
e.g., STATs (29) and SMADs
(34), and regulated
transmembrane proteolysis, e.g., Notch-1, SREBP-1, APP, ErbB-4
(35) and the Bacillus
subtilis 
E protein
(27). The stimulus-triggered
release of BglG from the membrane suggests that prokaryotes and eukaryotes use
similar strategies for signal transmission.
Our results shed light on the requirements for BglG–BglF interaction.
Because the BglF active site, C24, phosphorylates both BglG and
-glucosides (11), we
anticipated that recognition of BglG would not be mediated by this site.
Indeed, as shown here, a mutation in C24 does not abolish BglF interaction
with BglG, although it changes the kinetics of this interaction. This change
in kinetics seems to account for the activity of BglG when expressed in the
cell with BglF(C24S) (11). On
BglG, the three residues that were shown to be essential for its
phosphorylation by BglF (7,
24) are shown here not to be
absolutely essential for the interaction with BglF, although mutating D100 and
H160 somewhat reduces the interaction. However, mutating both BglF and BglG
phosphorylation sites reduces (for IIBbgl) or almost abolishes (for
BglF) the interaction. This suggests that the presence of a phosphate, or the
negative charge that it confers, stabilizes BglF–BglG interaction.
Phosphorylation of BglG in cells containing plasmid-encoded BglF(C24S) or
IIBbgl(C24S) might be accomplished by HPr
(14,
36) or by the
chromosome-encoded BglF, shown to phosphorylate a small portion of BglG in WT
E. coli strains that are bgl0
(8). The role of
phosphorylation in BglF–BglG interaction was substantiated by the
reduced or lack of interaction between BglF and BglG in a pts
background, where neither protein is phosphorylated. Hence, drainage of the
phosphate, required for a stable BglG–BglF complex, by the sugar is
likely to contribute to the release of BglG from the membrane on BglF
stimulation. The relative contribution of this drainage and of the
sugar-induced conformational change in BglF
(33) to the release of BglG to
the cytoplasm is hard to estimate. It is important to mention that recruitment
to the membrane of the BglG mutants, which cannot be phosphorylated by BglF,
has not been demonstrated. We have shown only that they have a certain
capacity to interact with the active site-containing domain of BglF when
overproduced, indicating that each of these residues alone is not crucial for
BglF–BglG interaction.
On the basis of theoretical models for molecular crowding and in vitro studies that measured phosphate flux through the glucose PTS enzymes under conditions that presumably mimic intramolecular conditions, it has been speculated that PTS enzymes form multiprotein complexes that transport PTS carbohydrates into the cell and phosphorylate them (37). Enzyme I of PTS was shown to localize to the inner surface of the cytoplasmic membrane (38). It remains to be seen whether BglG and its homologues are part of these PTS complexes.
Binding of a transcription repressor, Mlc, to the membrane-bound Enzyme II of glucose, PstG, has been reported (39, 40). However, unlike BglG and BglF, Mlc and PstG do not preassemble in the absence of stimulus. On the contrary, Mlc does not bind to the phosphorylated nonstimulated PstG. It binds to PstG only after dephosphorylation of PstG by glucose. The purpose suggested for Mlc-PstG binding is membrane sequestration of Mlc to prevent it from binding to its operator. Hence, binding of Mlc to PstG occurs in response to an environmental change and cannot play a role in increasing the efficiency or the speed of initiation of signal transduction.
Most of the information gathered on signaling pathways was obtained from biochemical studies that focused mainly on the chemical changes that the signaling proteins undergo. However, to gain better understanding of signaling processes, more information on the subcellular localization of the participating proteins, their diffusion rates, and the kinetics of their interaction is needed. Preassembly of kinases and their cognate substrates seems suitable for sensory systems that control a single process and produce one type of response, as opposed to systems that produce multiple outputs, and thus require divergent circuitry that relies on recruitment of various combinations of proteins to transient complexes. Many of the bacterial two-component sensory systems, composed of histidine kinase-response regulator pairs (41), fall into this category. Indeed, interaction of the phosphoryl transfer domain of the histidine kinase UhpB with the response regulator UhpA was suggested on the basis of indirect evidence (42). Probable recognition surfaces on several response regulators for their cognate histidine kinases have also been identified (e.g., refs. 43–45). The stability and the kinetics of these interactions remain to be studied. The preassembly of the Bgl proteins may prove to be a useful paradigm for other sensory systems that control a single process and need to respond rapidly to extracellular stimuli.
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Acknowledgements |
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Footnotes |
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Abbreviations: PTS, phosphotransferase system; SPR, surface plasmon resonance; MBP, maltose-binding protein; LexADBD, LexA repressor DNA-binding domain.
To whom correspondence should be addressed. E-mail:
amster{at}cc.huji.ac.il.
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N. Joly, A. Bohm, W. Boos, and E. Richet MalK, the ATP-binding Cassette Component of the Escherichia coli Maltodextrin Transporter, Inhibits the Transcriptional Activator MalT by Antagonizing Inducer Binding J. Biol. Chem., August 6, 2004; 279(32): 33123 - 33130. [Abstract] [Full Text] [PDF]
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 S. Deng, R. A. Stein, and N. P. Higgins Transcription-induced barriers to supercoil diffusion in the Salmonella typhimurium chromosome PNAS, March 9, 2004; 101(10): 3398 - 3403. [Abstract] [Full Text] [PDF]
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L. Fux, A. Nussbaum-Shochat, and O. Amster-Choder A Fraction of the BglG Transcriptional Antiterminator from Escherichia coli Exists as a Compact Monomer J. Biol. Chem., December 19, 2003; 278(51): 50978 - 50984. [Abstract] [Full Text] [PDF]
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L. Fux, A. Nussbaum-Shochat, and O. Amster-Choder Interactions between the PTS Regulation domains of the BglG Transcriptional Antiterminator from Escherichia coli J. Biol. Chem., November 21, 2003; 278(47): 46203 - 46209. [Abstract] [Full Text] [PDF]
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B. Gorke Regulation of the Escherichia coli Antiterminator Protein BglG by Phosphorylation at Multiple Sites and Evidence for Transfer of Phosphoryl Groups between Monomers J. Biol. Chem., November 21, 2003; 278(47): 46219 - 46229. [Abstract] [Full Text] [PDF]
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