(Received for publication, February 25, 1997, and in revised form, April 23, 1997)
From the Molecular Sciences, Glaxo Welcome, Research
Triangle Park, North Carolina 27709 and the
Department of Molecular Biology and Biochemistry,
University of California, Irvine, California 92697
Transcription from cistrons of the Escherichia coli CytR regulon is activated by E. coli cAMP receptor protein (CRP) and repressed by a multiprotein complex composed of CRP and CytR. De-repression results when CytR binds cytidine. CytR is a homodimer and a LacI family member. A central question for all LacI family proteins concerns the allosteric mechanism that couples ligand binding to the protein-DNA and protein-protein interactions that regulate transcription. To explore this mechanism for CytR, we analyzed nucleoside binding in vitro and its coupling to cooperative CytR binding to operator DNA. Analysis of the thermodynamic linkage between sequential cytidine binding to dimeric CytR and cooperative binding of CytR to deoP2 indicates that de-repression results from just one of the two cytidine binding steps. To test this conclusion in vivo, CytR mutants that have wild-type repressor function but are cytidine induction-deficient (CID) were identified. Each has a substitution for Asp281 or neighboring residue. CID CytR281N was found to bind cytidine with three orders of magnitude lower affinity than wild-type CytR. Other CytR mutants that do not exhibit the CID phenotype were found to bind cytidine with affinity similar to wild-type CytR. The rate of transcription regulated by heterodimeric CytR composed of one CytR281N and one wild-type subunit was compared with that regulated by wild-type CytR under inducing conditions. The data support the conclusion that the first cytidine binding step alone is sufficient to induce.
The transport proteins and enzymes required for nucleoside utilization in Escherichia coli are encoded by genes belonging to the CytR regulon (1). This gene family consists of nine unlinked transcriptional units whose expression is coordinately controlled by the interplay of two gene regulatory proteins. Transcription is activated in response to intracellular cAMP levels by CRP1 and repressed by a three-protein, CRP· CytR·CRP, complex. Transcription is induced when CytR binds cytidine. A central feature of this coordinate regulation is that CytR and CRP bind cooperatively to their respective operators (2). This is so despite the role of CytR as a functional antagonist of CRP. The critical role that cooperativity plays is highlighted by the fact that expression is induced, because this cooperative interaction is lost when CytR binds cytidine. Cytidine binding has no effect on intrinsic CytR binding to DNA.
CytR is a member of the LacI family of bacterial repressors (3). The gene regulatory activity of each of these proteins is modulated by binding a peripheral ligand, which functions as either inducer or co-repressor. The basic DNA binding unit of each of these proteins is a homodimer in which helix-turn-helix domains from both subunits combine to form the DNA binding interface. Since both subunits harbor identical ligand binding sites, the allosteric mechanism that couples inducer or co-repressor binding to changes in the macromolecular interactions that regulate transcription is an important issue to this entire family of proteins.
For both PurR and LacI, conformational transitions that accompany ligand binding have been investigated by x-ray crystallography (4-6). In these two cases, binding of co-repressor or inducer, respectively, causes a change in tertiary structure that alters substantially the dimer interface. In the non-DNA binding conformation, hinge helices that connect the helix-turn-helix motif to the ligand binding globular core domain are destabilized, and the helix-turn-helix motifs from the two subunits are thought to be out of register with successive DNA major grooves. In this manner, cooperative ligand binding (7, 8) to the individual subunits controls a concerted quarternary conformational change of the dimer. These features are consistent with MWC allostery. While the structural mechanisms that couple ligand binding to tertiary conformation differ in the two proteins (4-6), the tertiary and quarternary structural perturbations are remarkably similar.
The structures of the LacI family proteins, including CytR, appear to be highly conserved (5, 6, 9, 10). Given the structural resemblance among family members plus the similarity of allosteric mechanism for LacI and PurR, a similar mechanism might be anticipated for CytR. Yet CytR differs from all LacI family members in that it is cooperativity that is allosterically controlled and not intrinsic DNA binding. Allostery thus appears to have a different structural basis in CytR than in other LacI/PurR proteins.
Understanding the allosteric mechanism is central to understanding coordinate regulation of the CytR regulon genes. Recently, we showed that CytR binds to multiple operators at one CytR regulated promoter, deoP2 (12). CytR binding to the operator responsible for repression interacts cooperatively with CRP binding to flanking CRP sites, CRP1 and CRP2. However, by binding to additional specific sites, CytR competes with CRP for binding to CRP1 and CRP2. The net result of cooperativity and competition is that while CRP recruits CytR to form the repression complex, there is no significant reciprocal recruitment of CRP by CytR. This effect has also been reported for the nupG promoter (13). These interactions presumably function to direct both a multistage activation of transcription, using both Class I and Class II CRP mechanisms (14) and also a similar multistage repression mediated by CytR. We have proposed that this might be a general feature of CytR-mediated gene regulation (12).
The unique mechanism of cytidine mediated induction also suggests a multistage process. The cooperativity to which cytidine binding is linked appears to be complementary pair wise in nature. This follows from the observation that the free energy change characterizing cooperativity in the three protein complex, CRP·CytR·CRP bound to DNA, is equal to the sum of free energy changes characterizing pair wise cooperativity between CytR and CRP bound either to CRP1 or to CRP2 (12). If cooperativity in the three-protein repression complex is pairwise, then it is easy to envision that the two subunits of the dimer might react independently to cytidine binding. This would result in sequential elimination of pairwise, CytR·CRP cooperativity, hence sequential relief from repression, in response to sequential cytidine binding to the subunits.
The most general possibilities for coupling between ligand binding and
transcription initiation are presented in Fig. 1. We have combined biophysical chemical and molecular genetic approaches to
investigate these possibilities. First, CytR binding to CRP-saturated deoP2 was analyzed to evaluate the total contribution from
cooperativity. Subsequently, CytR binding titrations were conducted as
a function of cytidine concentration. The shape of the transition
characterizing loss of cooperativity as cytidine binds CytR indicates
that induction is an all or nothing process that occurs concomitant
with only one of the cytidine binding steps. Second, CytR mutants were
isolated and characterized as fully functional repressors, but which do not induce. The only defect in these mutants is inability to bind cytidine. By co-expressing cytidine induction-defective subunits and
wild-type subunits, we evaluated whether the resulting heterodimers would support induction with only one subunit capable of binding cytidine. The combined data from these studies indicate that induction results when cytidine binds to the first subunit of the CytR dimer.
Table
I lists the bacterial strains and plasmids used in this
study. The CID cytR allele, cytRD281N, was
transferred to the bacterial chromosome as described by Winans et
al. (15). First, the cat gene was inserted into plasmid
pCB071-161 at a position 44 bp 3 of the cytR termination
codon, resulting in plasmid, pCB122. Second, E. coli strain
VJS803 was transformed with linearized pCB122, and a recombinant
strain, SS6140, was selected as chloramphenicol-resistant (Cmr) and ampicillin-sensitive (Amps). The
cytRD281N allele was subsequently transferred to other strains by P1 transduction. The presence of the cytRD281N
allele was verified in each Cmr isolate by enzyme assays.
The tsx-lac gene fusions carried by strains GP4, Tsx-lac500,
Tsx-lac501, Tsx-lac502 and Tsx-lac503 (16) were transferred into strain
SS6003 by P1 transduction. The cytR::Tn10dTet
insertion was then moved from SS6018 into each SS6003 derivative,
yielding strains SS6117 through SS6121 (Table I).
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To express CytR, the coding sequences of wild-type and mutant cytR alleles were subcloned as an NdeI/BamHI fragment downstream of the T7 promoter carried by plasmid pSS584. Strain BL21(DE3) (17, 18) was transformed with each construct. The control plasmid for these experiments is pCB135, a pSS584 derivative devoid of cytR coding sequence.
Bacteria were collected from exponentially growing cultures for enzyme assays. The medium contained Vogel and Bonner salts (19) supplemented with vitamin B1 at 5 µg/ml, 0.02% casamino acids, and 0.4% glycerol (20). BL21(DE3) derivatives harboring CytR plasmids were grown in a 1% Bacto-tryptone, 0.4% glycerol medium containing Vogel and Bonner salts (TV medium). Either L-broth or 2 × YT was used for transformations and plasmid preparations (21). The Lac ± phenotypes of the various strains were determined on solid TTC-Lac medium as described previously (20). When used, the final cytidine concentration was 2 mM. Antibiotic concentrations used in the media were: ampicillin, 100 µg/ml; tetracycline, 15 µg/ml; chloramphenicol, 20 µg/ml; and kanamycin, 25 µg/ml in minimal medium or 50 µg/ml in rich medium.
Generation, Identification, and Characterization of CytR MutantsA mixture of mutagenic oligonucleotides complementary to cytR codons 276 through 284 was synthesized using an Applied Biosystems model 381A DNA synthesizer. The spiking protocol of Hutchison et al. (22, 23) was used to create degeneracy in the oligonucleotide sequence. The mutagenic oligonucleotide mixture and a site-directed mutagenesis kit from Amersham Corp. was used to mutate the cytR gene on an M13mp19cytR10 template. Both single and multiple mutations were obtained, the frequency of single mutations being about 30%. Phage pooled from about 5000 mutagenized M13mp19cytR10 plaques was propagated in E. coli strain JM103 by incubation for 4 h in 2 × YT medium. RF-M13 DNA was prepared as described (24). The cytR gene fragment containing the mutagenized sequence bounded by ApaI and BamHI cleavage sites was subcloned into pCB093 by fragment exchange (20).
The recombinant plasmid pool was transferred into SS6018 (cytR), which was grown on TTC-Lac-Kan medium containing 2 mM cytidine, to identify mutant CID repressors. The dominant negative phenotype of CytR mutants was established using CytR+ strain, SS6004, as described previously (20). The steady-state level of wild-type and mutant CytR was measured using a Western immunoblot analysis (20). Each cytR mutation that yielded a stable mutant protein was identified by DNA sequencing of the mutagenized cytR gene segment on a purified, double-stranded template (20).
Enzyme AssaysBacteria used for enzyme assays were grown
and cell extracts prepared as described previously (25). Cytidine
deaminase (CDA) and uridine dephosphorylase (UDP) spectroscopic assays
were performed as described previously (25, 26) except that the CDA
assay mixture contained 50 mM Tris-HCl (pH 7.5) and 0.5 mM cytidine. The -galactosidase activity of
exponentially growing bacteria was measured as described by Miller
(21). To express the enzyme activity as specific activity of the cell
extracts, the total protein concentration of the extracts was measured
by the Bradford assay (27) using bovine serum albumin as a
standard.
BL21(DE3) derivatives harboring T7 expression plasmids for
either wild-type or mutant CytR were grown at 37 °C in
nucleoside-free TV medium with 0.4% glycerol to
A600 1.0. CytR expression was induced by
adding 1% lactose and 2 mM
isopropyl-
-D-thiogalactopyranoside and incubating for 60 min before harvesting the cells. Cell pellets were washed, resuspended
in 20 mM MOPS (pH 6.8), 2 mM MgSO4,
1 mM Na4EDTA, 200 mM NaCl, and
treated with lysozyme, added to 10 µg/ml. The cells were frozen at
20 °C, thawed at 23 °C, and broken by sonication. The final
cell extract was the clear supernatant remaining following
centrifugation at 100,000 × g for 1 h at
4 °C.
CytR was purified using a simpler protocol than that reported several years ago (2) but which yielded a higher yield of CytR with similar purity. All purification steps were carried out at 4 °C. Pellets from cells harvested 165 min postinduction were resuspended in 20 mM MOPS (pH 6.80), 2 mM MgSO4, 1 mM Na4EDTA, 1 mM dithiothreitol (R-buffer) supplemented with 0.3 M NaCl. The resuspended cells were lysed using two passes through a French press and centrifuged at 50,000 × g for 3 h. The supernatant was adjusted to 0.2 M NaCl and 10% glycerol in R-buffer. Polyethyleneimine was added to a final concentration of 0.04% to precipitate nucleic acids and some proteins. The supernatant from a low speed centrifugation was adjusted to 0.1 M NaCl and chromatographed on two Bio-Rad EconoPac Q cartridges (5 ml each) connected in series using a Pharmacia FPLC. The pooled CytR containing the flow-through peak was loaded on two Bio-Rad Econo-Pac S cartridges connected in series. After washing the column with 0.2 M NaCl R-buffer until the A280 of the wash returned to the buffer base line, the column was eluted using a 0.2-0.6 M NaCl gradient. CytR elutes in a broad peak between 0.3 and 0.4 M NaCl.
CytR concentration was estimated from an extinction coefficient of
0.30 ± 0.03 cm1 mg
1 ml at 280 nm.
This value was calculated from the average extinction coefficients for
amino acid residues in a protein (28-31). The unusually low extinction
is due to the fact that CytR contains no tryptophan. Based on the
calculated extinction, the yield of CytR using this expression and
purification protocol is 1.5-2 mg/g of cell paste. CytR is stored at a
concentration of 2-3 mg/ml in the S-column elution buffer in a liquid
nitrogen dewer after flash freezing as ~25-µl beads in liquid
nitrogen. The DNA and cytidine binding activities remain stable for at
least several years when stored in this manner.
Sedimentation equilibrium analysis shows this material to be homogeneous dimer, with no evidence for either dissociation to monomer or association to higher order polymers over the concentration range, 0.1-10 µM.2 More recent analysis of gel mobility shift assays of CytR binding to DNA has been interpreted to indicate that CytR remains as stable dimer over the range of concentrations at which it binds DNA operators (11). Based on these data, the binding experiments were analyzed according to the simplest model in which CytR exists only as dimer.
Cytidine Binding AssaysBinding of [3H]cytidine to purified wild-type CytR and to both wild-type and mutant CytR containing cell-free extracts was measured using a filter binding assay. Binding reaction mixtures contained either 18-50 nM purified CytR dimer or 15-30 µg of cell extract protein in a 100-µl volume containing 0.04-11.0 µM [3H]cytidine (NEN Life Science Products). Two different buffers were used: 1) 20 mM MOPS (pH 6.8), 2 mM MgSO4, 1 mM NaEDTA, 200 mM NaCl and 2) 10 mM bis-Tris (pH 7.0), 100 mM NaCl, 0.5 mM MgCl2, 0.5 mM CaCl2. Both buffers contained 100 µg/ml bovine serum albumin and 1 µg/ml calf thymus DNA. Following a 5-min incubation at 23 °C, the CytR-bound [3H]cytidine contained in 80 µl of assay mix was collected on a prewashed nitrocellulose filter (Millipore HAWP 02500; Millipore Corp., Bedford, MA). The filters were washed once with 500 µl of assay buffer, air-dried, and then dissolved in 3.5 ml of Packard Filter-Count LSC mixture (Packard Instrument Co.). Radioactivity was measured using a Packard model 1900TR scintillation counter.
For determination of nucleoside binding constants, binding assays were conducted as titrations by varying the nucleoside concentration at constant CytR concentration. The data were analyzed according to a simple Langmuir binding model as described below (see Equation 1) using a nonlinear least squares curve fitting program (32). The CytR concentration used in some titrations was not negligible. To analyze data under these conditions, the conservation polynomials for total cytidine and total CytR were solved for the concentration of free cytidine for each data point and at each iteration of the nonlinear least squares analysis. The program NONLN (33) was used for this purpose.
In experiments to compare [3H]cytidine binding by CytR mutants in cell-free extracts and wild-type CytR, the CytR content of the cell-free extracts was estimated using Western immunoblots as described above. For each extract, 25-200 ng of extract protein was electrophoresed on SDS-16.5% acrylamide gels. Proteins were electrotransferred to Immobilon-P membranes (Millipore Corp.) as described previously (20). CytR was complexed with anti-CytR antibody and 125I-protein A. 125I-Protein A in the complex was quantitated using a Molecular Dynamics PhosphorImager (Molecular Dynamics, Sunnyvale, CA).
Competition by other nucleosides for [3H]cytidine binding to wild-type CytR in extracts was measured in the MOPS assay buffer. [3H]Cytidine and competitor concentrations were 1 µM and 1 mM, respectively. Cytidine deaminase and adenosine deaminase present in the cell extracts were inhibited by addition of 1 µM tetrahydrouridine and 10 µM deoxycoformycin, respectively. These nucleoside deaminase inhibitors did not affect cytidine binding by purified, wild-type repressor.
CRP Protein Used in DNA Binding StudiesCRP was the kind gift of James C. Lee (University of Texas Medical Branch, Galveston, TX). This protein preparation shows no evidence of any contaminating material by Coomassie staining of overloaded SDS- polyacrylamide gels, from which we conservatively estimate at least 98% purity.
Individual-Site Binding ExperimentsA series of DNase I
footprint titrations of CytR binding to deoP2 was conducted
to assess the effect of cytidine binding to CytR on heterologous
cooperativity between CRP·cAMP and CytR (Fig. 3). An 879-bp
NotI/HincII restriction fragment containing the deoP2 regulatory sequence was isolated from a derivative of
pSS1332 (Table I) in which an 8-bp NotI linker was inserted
into the SmaI site of the polylinker. The fragment contains
the E. coli deo sequence from +151 to 801 from the P2
start site for transcription. The fragment was labeled with
32P at the NotI site by using the Klenow fill-in
reaction and purified as described (34).
Each separate titration of CytR binding to this fragment (Fig. 4) was
conducted at a different, fixed concentration of cytidine ranging from
1 nM to 10 mM and at a saturating concentration
of CRP·cAMP. CytR was the titrating ligand. DNase I footprint
titrations were conducted as described (12) in the bis-Tris (pH 7)
cytidine binding buffer described above. CRP and cAMP were added to
final concentrations of 0.10 µM (as dimer) and 100 µM, respectively. Each titration was analyzed using NONLN
(33) according to Equation 1 to obtain the apparent free energy change
for cooperative binding of CytR at one cytidine concentration
(Gapp =
RTlnkapp),
![]() |
(Eq. 1) |
We previously described two CytR mutants, D281N and D281Y, that are repression-competent but nonresponsive to induction by cytidine (20). We denote such cytidine induction-defective mutants as CID. Degenerate oligonucleotide-directed mutagenesis of cytR codons 276 through 284 was used to identify other residues that are critical for cytidine induction. This mutagenesis yielded 21 mutant cytR genes that produced stable CytR. Twelve of these mutant genes encoded inactive repressors. Eleven of these inactive repressors have amino acid substitutions for either Ile277 or Asp274 and are recessive. The other inactive repressor (G279R) has an inactive, dominant negative phenotype. The remaining nine of these mutant genes produced CID phenotypes. Three of these are previously identified mutants, and six are newly identified mutants. Four of the newly identified mutants have single amino acid substitutions, three for Asp281 (Table II) and one for Gly279. The others carried double substitutions (F280I/D281A and F280S/N282I).
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The CID phenotype of the four new single substitutions was verified by measuring enzyme synthesis from two CytR controlled genes, cdd and udp, in the presence and absence of cytidine. As was found with the original CID mutants, expression of these CytR-controlled genes is repressed by the mutant CytR proteins and is unaffected by cytidine (Table II). Only D281A was a significantly less active repressor than the wild-type protein. However, like all CID mutants, D281A did not respond to cytidine. All CID cytR alleles were expressed identically and all produced steady-state cellular CytR concentrations equal to that of the wild-type protein based on Western blot analysis of soluble extracts from the mutant strains. As found for other CytR mutants (20), the CytR controlled enzyme levels and the phenotype observed for bacteria expressing these CID mutants reflect directly the change in repressor function.
Functional repressor might show the CID phenotype if repression was no
longer dependent on heterologous cooperative interactions with
CRP·cAMP. To determine whether CID mutants require such interactions for repression, we compared the ability of each repressor to regulate transcription from wild-type and mutant tsx-lac reporter
gene fusions. The tsx-lac gene fusions were constructed and
characterized by Gerlach et al. (16). The tsx DNA
of each mutant reporter gene fusion contains a single bp substitution
in CRP2, which greatly reduces its affinity for CRP·cAMP binding.
These point mutations have no direct affect on intrinsic CytR binding;
CytR binding is affected only indirectly via loss of CytR·CRP
cooperativity. This loss of cooperativity prevents repression of mutant
tsxP2 promoters by wild-type CytR in vivo, even
when expressed at high levels from a multicopy plasmid (37). Thus,
expression from tsx-lac gene fusions provides a specific,
direct assessment of CytR·CRP cooperativity. It avoids the use of
bacteria having either cya or crp
mutations
and their pleiotropic effects.
Both wild-type and CID CytR repress -galactosidase synthesis from
wild-type tsxP2. Regulation of mutant tsx-lac
fusions by wild-type and CID repressors was also identical (Table
III). Neither protein represses expression from the
mutant promoters. Therefore, repression by mutant CID CytR is dependent
on cooperative interactions with operator-bound CRP·cAMP, the same as
repression by wild-type CytR. Thus, the only identifiable defect in
function of the CID CytR proteins is their failure to respond to
cytidine induction.
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The
induction-defective phenotype of CID CytR could result either if the
protein fails to bind inducer or if it binds but fails to respond to
inducer. To assess these options, cytidine binding was studied
directly. Titration of purified wild-type CytR yielded a single binding
transition (Fig. 2). When analyzed according to a simple
binding model (Equation 1), an apparent Kd value of
0.17 (0.11,0.33)3 µM was obtained. The
limiting ratio of cytidine retained to CytR dimer is 0.82. While this
is similar to a stoichiometry of one per dimer, it doesn't account for
CytR protein that isn't retained by the nitrocellulose filters and so
only represents a lower limit to the stoichiometry. Cytidine binding is
unaffected by addition of cell free extracts from an E. coli
strain that is deleted of the cytR gene
[Kd = 0.16 (0.11,0.30) µM]. Cytidine
binding was also the same whether using purified CytR or cell free
extracts from an E. coli strain containing the wild-type
cytR gene (data not shown). Ligand binding is also specific.
In competition assays conducted in the presence of a 1000-fold excess
of nucleoside competitor (1 mM), only adenosine competed
with cytidine (KI = 22.5 ± 1.5 µM). Uridine, 2-deoxycytidine, 2
,3
-dideoxycytidine, cytidine arabanoside, and both 2
- and 3
-O-methylcytidine
had no effect on cytidine binding in vitro, consistent with
the inducer specificity in vivo (38-40).
Since addition of CytR-free E. coli extracts has no effect on cytidine binding to purified CytR, cytidine binding to wild-type and mutant CytRs in cell free extracts can be compared directly. Extracts were prepared from isogenic strains expressing either wild-type CytR, CytR from one of the different classes of mutants, or no CytR. There was no detectable cytidine binding in extracts lacking CytR. Of the mutant CytR containing extracts, only that containing the CID repressor (CytRD281N) showed a significant decrease in cytidine binding (Table IV) as compared with the extract containing wild-type CytR. At a cytidine concentration (11 µM) that essentially saturates wild-type CytR (0.98 saturation), this CID CytR bound less than 1% as much cytidine as wild-type. This indicates a Kd for the mutant CID CytR of 1 mM or greater, an affinity at least 2000-fold reduced from that of wild-type CytR. An effect of such magnitude can only be readily explained by differences between CytR in the different cell-free extracts. By contrast to this result, CytR with amino acid substitutions in domains proposed previously (20) to function in DNA binding (CytRV15A) and in signal transduction (CytRM151I and CytRM151V) bound roughly the same amount of cytidine as wild-type CytR, thus indicating no effect on cytidine binding affinity.
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DNase I footprint titration was used to
determine the apparent affinity of CytR for deoP2 at
different, fixed concentrations of inducer cytidine. The concentrations
of cAMP and of CRP used yield 97 and >99% saturation of CRP1 and
CRP2, respectively (12). Fig. 3 presents the apparent
Gibbs free energy changes (Gapp) for binding
of CytR dimer to the deoP2·(CRP·cAMP)2
complex as a function of cytidine concentration. Each
Gapp was determined from a separate CytR
titration experiment such as that shown in Fig. 4. Under
the conditions of these experiments, heterologous cooperativity
increases the apparent affinity of CytR for deoP2 by
100-fold (Fig. 3 and Ref. 12). Thus, the
Gapp
values plotted reflect a sum of two contributions. One contribution
arises from intrinsic CytR binding (
10.4 ± 0.4 kcal/mol; Fig.
5 and Ref. 12); the other is from cooperative
interactions between CytR and CRP·cAMP. In the absence of cytidine,
this sum of contributions yields
Gapp =
13.1 ± 0.2 kcal/mol (Fig. 3 and Ref. 12). The contribution from
cooperativity is reduced when CytR binds cytidine, as indicated by the
smooth transition in Fig. 3. The data in Fig. 5 confirm no effect on
intrinsic binding. At cytidine concentrations above 1 mM,
the apparent free energy change reaches an asymptotic limit of about
10.7 kcal/mol. The cooperative effect is thus reduced from 100-fold
to less than 2-fold.
The equilibria between CytR, cytidine, and the deoP2· (CRP·cAMP)2 complex are shown in Scheme I. An implicit simplifying assumption is that CytR binds to only a single site near deoP2. The effect of CytR binding to additional sites (12) is negligible under the conditions employed in these assays. Scheme I accounts for one cytidine binding site on each subunit of homodimeric CytR. K1 and K2 are macroscopic stepwise association constants, which describe the binding of the first and second cytidine ligands to free CytR dimer. K3, K4, and K5 are association constants for binding of CytR, CytR· cytidine, and CytR·(cytidine)2 to the deoP2·(CRP·cAMP)2 complex. K6 and K7 are macroscopic stepwise association constants for binding of the first and second cytidine ligands to deoP2·(CRP·cAMP)2-bound CytR. The apparent equilibrium constant for CytR binding to deoP2·(CRP·cAMP)2 at any cytidine concentration ([cyt]) is as follows.
![]() |
(Eq. 2) |
Equation 2 was used to analyze the values in Fig. 3
(Gapp =
RTlnKapp) to estimate the Gibbs
free energy changes corresponding to
K1-K5. Results of this
analysis are indicated by the broken curve in Fig. 3. The
maximum likelihood parameter estimates indicate an affinity for binding
of the first cytidine, K1 = 2.7 × 107 M
1. In this model, this
binding step is accompanied by only a negligible effect on
Kapp (1.5-fold decrease). The second subunit
binds cytidine with significantly lower affinity
(K2 = 1.0 × 106
M
1) and is accompanied by the major
reduction in Kapp (40-fold decrease). (Note: K2 is a stepwise macroscopic equilibrium
constant; it corresponds to a microscopic constant for cytidine binding
to the remaining cytidine binding site equal to 2.0 × 106 M
1.) This result suggests
that cytidine binding is negatively cooperative and that the major
allosteric switch occurs at only one of the two cytidine binding
steps.
An alternative possibility that is also consistent with this conclusion
is that the allosteric switch occurs at the first cytidine binding
step. If so, the transition in Fig. 3 represents cytidine binding to
only the first of the two CytR subunits. To test this
possibility, the data were analyzed using a simplified version of
Scheme I, in which K2 and
K5 were set equal to zero. Results of this
analysis, indicated by the solid curve in Fig. 3, yielded
K1 = 2.0 × 106
M1. Thus, both models estimate the same
cytidine affinity as being allosterically coupled to cooperative
binding of CytR to the deoP2·(CRP·cAMP)2 complex. Goodness of fit criteria such as the variances and
distributions of residuals provided no basis for discriminating between
the two models. By contrast, all other models tested, such
as the model in which the two subunits have independent
(noninteracting) cytidine binding sites, which are either separately or
together coupled to cooperative operator binding, failed to provide an adequate fit to the data.
The existence of well characterized CID alleles provides the means to test, in vivo, the conclusion that cooperativity is coupled primarily to only one cytidine binding step and also to determine whether the first or second step. To do so, steady-state expression from CytR-regulated promoters was examined in bacteria that co-express both wild-type, inducer-responsive CytR subunits and CID, nonresponsive CytR subunits. Wild-type and CID subunits were expressed by cistrons that had identical promoter and operator regions. Since both wild-type and CID CytR are fully functional repressors, their co-expression should result in heterodimeric CytR with one wild-type subunit and one CID subunit.
To promote heterodimer formation, CID allele cytRD281N was
first recombined in single copy into the chromosome. As shown in Table
V, CytRD281N that is expressed from the bacterial
chromosome is indistinguishable from wild-type CytR in its ability to
repress CDA and UDP synthesis, but retains its CID phenotype. Second, isogenic strains that differ only in their chromosomal cytR
allele (either wild-type or CID) were transformed by a plasmid that
expresses the wild-type cytR gene. A low copy number plasmid
(41, 42) containing a P15A origin was used. These constructions yielded bacteria with the same cytoplasmic CytR level (see "Materials and
Methods") as one another. This is as expected, since expression of
all cytR alleles is identically regulated and both wild-type and CID proteins are functional, stable repressors. The response to
induction was compared using these two strains. To ensure that steady-state transcriptional activity was being compared as opposed to
steady-state enzyme levels governed by protein turnover, the differential rate of CytR-controlled -galactosidase synthesis from a
udp-lac fusion was measured in exponentially growing
cells.
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The differential rate of CytR regulated -galactosidase synthesis in
bacteria that co-express wild-type and CID CytR subunits should depend
on the response of CID/wild-type heterodimers to cytidine. There are
three possibilities. First, if the heterodimer is induction-defective,
then both CID/CID homodimers and CID/wild-type heterodimers would
repress udp-lac expression. Because the wild-type allele is
plasmid-encoded, while the CID allele is chromosomal, these bacteria
have excess wild-type subunits, hence excess wild-type homodimers.
However, under inducing conditions, these homodimers no longer bind
cooperatively to the promoter; their affinity is reduced by 2 orders of
magnitude (Fig. 3). Consequently, even a severalfold excess of
wild-type homodimers would be insufficient to compete effectively with
induction refractive heterodimers and the small proportion of CID
homodimers.4 The differential rate of
-galactosidase synthesis would approximate that of CID homozygous
bacteria, even under inducing conditions. Second, if CID and wild-type
subunits do not associate to form heterodimers, then all CID subunits
expressed must be present as CID homodimers. Again, the level of
noninducible CytR, now comprised of the CID/CID homodimers, would be
sufficient to yield a noninducible phenotype. Third, if (and only if)
cytidine binding by the wild-type subunit of CID/wild-type heterodimers
is sufficient for induction, then the differential rate of
-galactosidase synthesis under inducing conditions would be much
greater than that observed for CID homozygous bacteria, even
approaching that observed for wild-type CytR homozygous bacteria. Full
wild-type levels of induction would not be expected because of
competition by the nonresponsive CID/CID homodimeric CytR, even when
present as a small fraction of the total repressor.
The rates of bacterial growth and -galactosidase synthesis for the
individual strains are shown in Fig. 6. These rates were constant throughout the entire experiment, demonstrating that cytidine
induction is at equilibrium and that the enzyme levels represent
steady-state production in all cultures. The differential rates of
-galactosidase synthesis were calculated from these curves. Under
inducing conditions, the rate for bacteria expressing both wild-type
and CID CytR subunits was found to be 5-fold greater than that measured
for CID homozygous bacteria (2543 units versus 499 units).
It was slightly greater than half of that found for wild-type
homozygous bacteria (4827 units). These results can be explained if
wild-type and CID subunits do associate to form heterodimers, which in
turn do respond to induction by cytidine.
CRP and CytR mediate coordinate regulation of the unlinked genes that encode the proteins necessary for nucleoside utilization in E. coli. The interplay between these regulatory proteins, comprised of both cooperative and competitive interactions, appears to direct both multistage activation and multistage repression of transcription of individual cistrons (12). The critical role of CytR·CRP cooperativity is highlighted by the mechanism of cytidine-mediated induction. This gene regulatory mechanism relies on loss of cooperativity, not on reduction in DNA binding affinity as found with other LacI family members. Understanding the allosteric coupling between cytidine binding and CytR·CRP cooperativity is necessary to understanding the coordinate regulation of this gene family.
The specific question addressed herein is whether induction is a concerted process coupled to the quarternary state of the CytR dimer or a sequential process coupled to the ligation state of the individual subunits. The former holds for LacI and PurR, which undergo an MWC-type allosteric transition between quarternary T and R states (4-6). CytR presumably does not experience the same global conformational change upon ligand binding as LacI and PurR, since inducer binding is not coupled to DNA binding. Moreover, CytR·CRP cooperativity appears to be pairwise and complementary, in nature (12). A multistage induction such as would result from a sequential coupling between binding of cytidine to an individual subunit, and a tertiary conformational change affecting only that subunit's cooperative interaction, could play a significant role in differential expression of the unlinked genes.
Three allosteric mechanisms can be considered: first, a classic MWC mechanism featuring an equilibrium between two symmetric quarternary states, one that interacts cooperatively with CRP and one that does not (Fig. 1A); second, a strictly sequential KNF mechanism in which the tertiary conformation of each subunit switches to a noncooperative state concomitant with cytidine binding to that subunit (Fig. 1B); third, a sequential but concerted mechanism in which distinct quarternary states are formed as each cytidine site is filled (Fig. 1C). Elimination of CytR·CRP cooperativity and induction of transcription might occur when the first cytidine binds, when the second binds, or in part when both bind in proportion to the overall fractional saturation.
We investigated the cytidine-mediated transition from cooperative to noncooperative CytR binding to CRP-saturated deoP2 to distinguish among these possibilities. The data were analyzed according to a general formulation (Scheme I; Equation 2) that encompasses all three allosteric mechanisms. Only two numerical solutions of Equation 2 were found to be consistent with the data. Two common features of these solutions both point to the third allosteric mechanism (Fig. 1C) and are inconsistent with MWC and KNF mechanisms (Fig. 1, A and B). First, the analysis suggests that the transition from cooperative to noncooperative CytR binding is coupled to a single cytidine binding event. Second, cytidine binding is characterized by negative cooperativity.
That loss of cooperativity is coupled to only a single cytidine binding step is reflected by the characteristic shape of the transition curve. Other mechanisms, such as a cooperative transition between pre-existing states (MWC; Fig. 1A) or sequential coupling to each cytidine binding step (e.g. KNF; Fig. 1B) yield either sharper or shallower transitions. These are inconsistent with the data. The finding of negative cooperativity in cytidine binding to CytR is also inconsistent with an MWC allosteric mechanism. A concerted transition between pre-existing states necessarily yields positive cooperativity in ligand binding.
The two numerical solutions to Equation 2 do differ in detail regarding the negative cooperativity. When both cytidine binding steps are considered in the analysis, the cytidine binding constants estimated indicate approximately a 7-fold effect. When Equation 2 is truncated to consider only one cytidine binding step, this is equivalent to the assumption that linkage reflects only the first cytidine binding step. This would mean that binding of the second cytidine is insignificant over the concentration range investigated, suggesting a much higher degree of negative cooperativity.
We cannot distinguish between these possibilities, even based on the titration data for cytidine binding to free CytR (Fig. 2). These data were reanalyzed by considering cytidine binding as comprising two steps. This analysis (Fig. 1) estimated an intrinsic cytidine binding affinity equal to 0.4 µM, nearly identical to that identified as being coupled to loss of cooperativity in DNA binding (0.5 µM; Fig. 3) and negative cooperativity accounting for greater than an 80-fold affect. It also yielded a limiting ratio of cytidine retained to CytR dimer equal to 1.60, consistent with a stoichiometry of two per dimer. According to this model, the transition in Fig. 1 corresponds primarily to the first cytidine binding step; the plateau continues sloping upward, reflecting the second binding step that occurs at higher cytidine concentration. These features mirror the result obtained from the DNA binding data in Fig. 3, when the latter are analyzed simply by assuming linkage to only the first cytidine binding step.
However, the cytidine titration data in Fig. 2 are also reasonably described by the alternative analysis of the DNA binding data. The data in Fig. 2 were analyzed using the stepwise cytidine binding constants, K1 and K2, obtained from analysis of the DNA binding data in Fig. 3 as fixed input parameters. According to this interpretation, cytidine binding in Fig. 2 looks like a single transition, because the negative cooperativity is too moderate to produce separate binding transitions. This analysis estimates a limiting ratio of cytidine retained to CytR dimer equal to 0.9. While this is quite low compared with the model's stoichiometry of two per dimer, the discrepancy could reflect poor CytR retention efficiency in the filter assay.
Despite uncertainty in details, these analyses support three conclusions: first, that cytidine binding to CytR is negatively cooperative; second, that cooperative binding of CytR to deoP2·(CRP·cAMP)2 is primarily coupled to only one of the two cytidine binding steps; and third, the intrinsic cytidine binding affinity in free CytR that is coupled to this transition is 0.2-0.5 µM. Thus, cytidine binding must switch CytR between three conformational states, one corresponding to each cytidine ligation state, as represented by Fig. 1C. However, these data do not identify which conformational change eliminates CytR·CRP cooperativity. To address this issue, it is necessary to evaluate the behavior of the intermediate state with only one subunit liganded. For this, wild-type and CID CytR alleles were co-expressed, thus allowing assembly of hybrid CytR dimers, which have only one subunit capable of binding cytidine. The behavior of these hybrids in vivo was used to assess the induction competency of the intermediate ligation state.
For this approach, it was necessary to find a CytR mutant whose only defect is inability to bind cytidine. We focused the search on a region of the CytR sequence in which CID mutants had been identified previously (20). The newly identified CID mutants expressed wild-type levels of protein, and most had repressor activity equal to wild-type (Table II). One allele, cytRN281N, was found independently both in the previous screen following random mutagenesis of the entire CytR gene (20) and in the present screen following targeted mutagenesis. CytRD281N was shown to require cooperative interaction with CRP for repression as does wild-type CytR (Table III). From this we infer that repression remains coupled to the mechanism that underlies induction. Cytidine binding assays demonstrated the only defect found, a 2000-fold or greater reduction in cytidine binding affinity (Table IV).
The finding that cytidine binding affinity is reduced as a result of
amino acid substitutions for Asp281 is consistent with
observations on other LacI proteins. This is a conserved aspartate in
the sequences of many of the LacI repressors as well as the E. coli periplasmic binding proteins (3). The equivalent aspartate is
essential for ligand binding in LacI, PurR, and the periplasmic
sugar-binding proteins for glucose, ribose, and arabinose (43-45). The
substitutions for CytR Asp281 that yield the CID phenotype
(Asn, Ala, Glu, and Ile) are understandable if Asp281
participates directly in cytidine binding as a hydrogen bond partner to
sugar hydroxyls. Perhaps, the decreased affinity of CID mutants for
cytidine is functionally equivalent to the inability of 2- or
3
-deoxycytidine to compete with cytidine for binding to wild-type
CytR. The substitutions that yield the CID phenotype suggest that
charge (D281N) and side chain size (D281E) as well as hydrogen bonding
potential (D281A) are important to this interaction. Whether this
interaction provides only affinity and specificity, or is also
important to the coupling between cytidine binding and CytR·CRP
cooperativity, cannot be determined from our data. However, we note
that in LacI, only affinity is affected (46).
Evaluation of the capacity of the hybrid dimers to support induction in vivo assumes that the steady-state levels of wild-type and CID subunits are proportional to the gene copy number. Since the wild-type and CID alleles are identically regulated, this assumption is supported by the finding of comparable CytR levels in extracts made from bacteria expressing either wild-type or mutant CytR. Thus, for bacteria expressing both alleles and assuming a plasmid copy number of ~6-8 per cell (41, 42) and random assortment of subunits, then CytR dimer is proportioned about 75% as wild-type homodimer, 23% as heterodimer, and the remainder as CID homodimer. Uncertainty in plasmid copy number has a negligible effect on this distribution. If heterodimers respond to cytidine, this distribution yields a 50-fold excess of inducible over noninducible dimers. When offset by the higher affinity of non-induced dimers, some reduction of the extent of de-repression is expected, perhaps in line with what was observed (Fig. 6). If heterodimers do not respond to cytidine binding, then only a 3-fold excess of inducible over noninducible dimers results. This is insufficient to compete effectively for DNA binding under inducing conditions and de-repression should not be observed. Similarly, if CID and wild-type subunits do not assemble as heterodimers, then the CID subunits must be assembled as CID homodimers to account for the fact that CID CytR is functional repressor. Again, the excess of inducible over noninducible dimer would not be sufficient to support significant de-repression. Therefore, the in vivo data support the conclusion that hybrid dimers respond to cytidine. Presumably, half-saturated wild-type dimers behave in the same manner.
These results, obtained by monitoring the effect of cytidine on CytR-regulated gene expression in vivo, are remarkably consistent with those obtained by in vitro investigation of allosteric coupling between cytidine binding to CytR and cooperative interaction of CytR and deoP2·(CRP·cAMP)2. This consistency supports a molecular model in which cytidine binding switches CytR between three states and binding of cytidine to the first subunit of CytR dimer yields complete induction. Because cytidine binding to CytR is negatively cooperative, the third CytR state, that generated by ligation of both subunits by cytidine, forms only at very high cytidine concentrations. Such concentrations are not normally attained in E. coli nor were they attained in our experiments. We note the similarity to CRP, which also has three distinct conformational and functional states corresponding to its cAMP ligation states (47).
Why does CytR differ from other LacI family members for which ligand binding affects the poise of an equilibrium between DNA binding and non-DNA binding conformations? We envision two possibilities. First, perhaps the CytR inducer binding core domain does undergo a similar conformational change when cytidine binds, but this is uncoupled from the structure of the DNA binding domain. Comparing the sequences of the LacI repressors, we note that where other family members have a pair of conserved alanines in the hinge helix sequence, CytR has proline (Pro57) and glycine (Gly59) (3). Perhaps, instead of hinge helices, CytR has an extended coil that only loosely tethers the Core and DNA binding domains, such that DNA binding is unaffected by the conformation of the Core dimer. Second, perhaps cytidine binding is uncoupled from the T-state to R-state transition, but is instead coupled to transitions between subconformations belonging to the quarternary R-state. With this scenario, it is interesting to speculate whether saturation by cytidine might induce a T-state transition and whether this would have an effect on DNA binding.
We thank Jim Lee and Thomas Heyduk for the gift of the purified CRP used in this study. We thank Richard Brennen for sharing the PurR coordinates with us prior to their publication and for his insights into structure-function relationships in LacI family repressors. We thank Michael Brenowitz and Howard Nash for their critical review of earlier versions of the manuscript. We thank prior members of our laboratories, particularly Laura Perini, for their technical assistance.