JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M501615200 on May 10, 2005

J. Biol. Chem., Vol. 280, Issue 27, 25665-25673, July 8, 2005
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
280/27/25665    most recent
M501615200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Togashi, M.
Right arrow Articles by Webb, P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Togashi, M.
Right arrow Articles by Webb, P.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Rearrangements in Thyroid Hormone Receptor Charge Clusters That Stabilize Bound 3,5',5-Triiodo-L-thyronine and Inhibit Homodimer Formation*

Marie Togashi{ddagger}, Phuong Nguyen{ddagger}, Robert Fletterick§, John D. Baxter{ddagger}, and Paul Webb{ddagger}||

From the {ddagger}Diabetes Center and the§ Department of Biochemistry and Biophysics, University of California, San Francisco, California 94143-0540

Received for publication, February 11, 2005 , and in revised form, May 9, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In this study, we investigated how thyroid hormone (3,5',5-triiodo-L-thyronine, T3) inhibits binding of thyroid hormone receptor (TR) homodimers, but not TR-retinoid X receptor heterodimers, to thyroid hormone response elements. Specifically we asked why a small subset of TR{beta} mutations that arise in resistance to thyroid hormone syndrome inhibit both T3 binding and formation of TR{beta} homodimers on thyroid hormone response elements. We reasoned that these mutations may affect structural elements involved in the coupling of T3 binding to inhibition of TR DNA binding activity. Analysis of TR x-ray structures revealed that each of these resistance to thyroid hormone syndrome mutations affects a cluster of charged amino acids with potential for ionic bond formation between oppositely charged partners. Two clusters (1 and 2) are adjacent to the dimer surface at the junction of helices 10 and 11. Targeted mutagenesis of residues in Cluster 1 (Arg338, Lys342, Asp351, and Asp355) and Cluster 2 (Arg429, Arg383, and Glu311) confirmed that the clusters are required for stable T3 binding and for optimal TR homodimer formation on DNA but also revealed that different arrangements of charged residues are needed for these effects. We propose that the charge clusters are homodimer-specific extensions of the dimer surface and further that T3 binding promotes specific rearrangements of these surfaces that simultaneously block homodimer formation on DNA and stabilize the bound hormone. Our data yield insight into the way that T3 regulates TR DNA binding activity and also highlight hitherto unsuspected T3-dependent conformational changes in the receptor ligand binding domain.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Thyroid hormone receptors (TR{alpha} and TR{beta})1 are conditional transcription factors that play important roles in development, metabolism, and homeostasis (1–4). TRs regulate gene transcription in the presence of 3,5,3'triiodo-L-thyronine (T3) and in the absence of ligand (5). Current efforts to modulate TR activities have focused on development of selective agonists that mimic the beneficial effects of T3 upon circulating cholesterol and body weight without producing unwanted effects of the hormone on heart rate (6). However, there is also a need for TR antagonists, which could represent improved and faster acting treatments for hyperthyroidism and cardiac arrhythmias (6, 7). Furthermore observations from TR{alpha}/TR{beta} knock-out mice suggest many clinical manifestations of hypothyroidism are due to actions of unliganded TRs (8, 9). Thus, drugs that specifically reverse actions of unliganded TRs could be useful for treating hypothyroidism and would avoid risk of thyroid hormone excess (7). Improved understanding of unliganded TR structure and ways that unliganded TRs rearrange in response to T3 will facilitate development of all of these drugs.

Presently the organization of unliganded TR is only partly understood (10, 11). X-ray structures of liganded TR C-terminal ligand binding domains (LBDs) reveal a canonical {alpha}-helical structure with T3 buried in the core of the protein (12–16), but there are no equivalent structures of unliganded TRs. It has proven possible, however, to use a combination of x-ray structural information and targeted mutagenesis to learn about the organization of unliganded TRs. For example, T3 blocks transactivation and transrepression activities of unliganded TRs by promoting release of corepressors such as N-CoR and SMRT (silencing mediator of retinoid and thyroid receptors) (5) and induces a T3-dependent activation function (AF-2) that binds coactivators such as the p160s (17). Functional analysis of TR mutants reveals that AF-2 is comprised of surface-exposed residues from helices (H) 3, 5, and 12 and that the corepressor binding surface overlaps AF-2 but extends below the position of H12 in the liganded state (18–21). Thus, it is possible to infer that H12 is displaced in the unliganded state and that T3 binding leads to repositioning of H12 over the lower part of the corepressor binding surface, simultaneously promoting corepressor release and completing the coactivator binding site (5).

T3 also regulates TR DNA binding activity (1). TRs utilize their DNA binding domain to recognize specific thyroid hormone response elements (TREs) comprised of AGGTCA repeats and bind these elements either as heterodimers with the closely related retinoid X receptor (RXR) or as homodimers and monomers. T3 does not affect RXR-TR interactions with TREs but does promote release of TR homodimers from some TREs (inverted palindromes (F2/IP-6) and direct repeats (DR-4)) but not from TREs at which TRs bind as monomers or paired monomers (palindromes, TREpal) (22). TR homodimers bind N-CoR more strongly than RXR-TR heterodimers (23, 24), and the extent of TR homodimer binding to different TREs in vitro correlates with the extent of repression from these elements in vivo (25, 26). Thus, it is thought that T3-dependent inhibition of homodimer formation relieves transcriptional repression by unliganded TRs. Nevertheless the mechanisms involved in coupling of T3 binding to inhibition of DNA binding are not clear; TRs utilize the same surface at the junction of H10 and H11 in homodimer and heterodimer formation on DNA (27). The structural elements that render homodimers sensitive to T3 are not known.



View larger version (55K):
[in this window]
[in a new window]
  		FIG. 1.
Location of RTH mutations in charge clusters that are adjacent to the dimer surface. A, charge Clusters 1 and 2 are adjacent to the TR{beta} dimer surface. The figure shows a space-filling model of the TR{beta} LBD. Residues in the dimerization surface (Leu400, Pro419, Leu422, Met423, and Met430) are shown in green. Residues in Cluster 1 (Arg338, Lys342, Asp351, and Asp355) and Cluster 2 (Glu311, Arg383, and Arg429) are shown in blue (positively charged) and red (negatively charged). B and C, closer view interactions between the residues that comprise charge Clusters 1 (B) and 2 (C). Positively charged residues are shown in blue, and negatively charged residues are shown in red. Asterisk represents residues mutated in RTH. D, alignment of Cluster 1 residues in TR and other NRs. E, alignment of Cluster 2 residues in TR and other NRs. hTR, human TR; hRAR, human retinoic acid receptor; hLXR, human liver X receptor; hER, human estrogen receptor; hPXR, human pregnane X receptor; hVDR, human vitamin D receptor; hCAR, human constitutive androstane receptor.

 
In this study, we utilized targeted mutagenesis to explore elements of the TR that are specifically required for homodimer formation on TREs and tested the hypothesis that the same elements are involved in coupling T3 binding to inhibition of DNA binding. Whereas most TR{beta} mutations that arise in resistance to thyroid hormone syndrome (RTH) reduce the affinity of TR{beta} for T3 (3, 28, 29), a small subset also inhibits binding of TR{beta} homodimers, but not heterodimers, to TREs (30, 31). Here we report that these RTH mutations affect clusters of charged amino acids in the LBD with potential for electrostatic stabilization of TR conformation but that distinct arrangements of charged residues are needed for stable T3 binding and DNA binding by unliganded TRs. We propose that the charge clusters rearrange upon T3 binding to block homodimer formation and create new ionic bonds that stabilize bound hormone.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
TR Mutants—The pCMX vector was used for expression of the full-length human TR{beta} (17). Mutations within TR-encoding sequences were created using the QuikChange XL site-directed mutagenesis kit (Stratagene). Mutation of target sequences was verified by automated DNA sequence (Elim Biopharmaceuticals, Inc., Hayward, CA).

Transfections—HeLa cells were maintained in Dulbecco's modified Eagle's H-21 4.5 g/liter glucose medium containing 10% fetal bovine serum, 2 mM glutamine, 50 units/ml penicillin, and 50 mg/ml streptomycin. For transfection, cells were collected and resuspended in Dulbecco's phosphate-buffered saline (0.5 ml/4.5 x 107 cells) containing 0.1% dextrose and typically 4 µg of reporter, 1 µg of TR expression vector or empty vector control, and 2 µg of pCMV-{beta}-galactosidase (17). Cells were electroporated at 240 V and 960 microfarads, transferred to fresh media, and plated into 12-well plates. After incubation for 24 h at 37 °C with T3 or vehicle, cells were collected, and pellets were lysed by addition of 150 µl of 100 mM Tris-HCl, pH 7.8, containing 0.1% Triton X-100. The reporters contained two copies of each TRE (DR-4, F2, and TREpal) upstream of the herpes simplex virus thymidine kinase promoter TATA box linked to luciferase coding sequence. Luciferase and {beta}-galactosidase activities were measured by using a luciferase assay system (Promega) and Galacto-Light Plus {beta}-galactosidase reporter gene assay system (Applied Biosystems).

Glutathione S-Transferase Pull-down Assays—Full-length human TR{beta} was expressed in a coupled transcription translation system (TNT, Promega). N-CoR (amino acids 1944–2453) and GRIP1 (amino acids 563–1121) were expressed in Escherichia coli strain BL21 as a fusion protein with glutathione S-transferase according to the manufacturer's protocol (Amersham Biosciences). Bindings were performed by mixing glutathione-linked Sepharose beads containing 4 µg of glutathione S-transferase fusion proteins (Coomassie Plus protein assay reagent, Pierce) with 1–2 µl of 35S-labeled human TR{beta} in 150 µl of binding buffer (20 mM HEPES, 150 mM KCl, 25 mM MgCl2, 10% glycerol, 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride, and protease inhibitors) containing 20 µg/ml bovine serum albumin for 1.5 h. Beads were washed three times with 200 µl of binding buffer, the bound proteins were resuspended in SDS-PAGE loading buffer, and proteins were separated using 10% SDS-polyacrylamide gel electrophoresis and visualized by autoradiography.

T3 Binding Assay—TRs were expressed using the TNT T7 quick coupled transcription translation system (Promega). The affinities of T3 binding were determined using a saturation binding assay. Briefly 15 fmol of each in vitro translated protein were incubated overnight at 4 °C with varying concentrations of [125I]T3 (PerkinElmer Life Sciences) in 100 µl of E400 buffer (400 mM NaCl, 20 mM KPO4, pH 8, 0.5 mM EDTA, 1.0 mM MgCl2, 10% glycerol), 1 mM monothioglycerol, and 50 µg of calf thymus histones (Calbiochem). The bound [125I]T3 was isolated by gravity flow through a 2-ml Sephadex G-25 (Amersham Biosciences) column and quantified using a {gamma}-counter (COBRA, Packard Instruments). Off rate (koff) was determined by adding a 1000-fold molar excess of unlabeled T3 to a mixture containing TR and 1 nM [125I]T3 incubated previously overnight at 4 °C; aliquots were taken at the indicated time points to determine how rapidly the labeled ligand dissociates from TR. These aliquots were applied to Sephadex G-25 columns, and TR-bound [125I]T3 was quantified using a {gamma}-counter. As each T3-TR complex dissociates at a random time, the amount of specific binding follows an exponential dissociation equation: Y = Span·e–k·x + Plateau where x is time (min), Y is total binding (cpm), Span is the difference between binding at time 0 and plateau (cpm), and k is the dissociation rate constant (koff, expressed in min–1). Binding curves were fit by nonlinear regression, and dissociation constant (Kd) and koff values were calculated using the one-site saturation binding, one-phase exponential decay, and one-phase exponential association models, respectively, contained in the Prism version 3.03 program (GraphPad Software, Inc., San Diego, CA).

Gel Shifts—Binding of TR to DNA was assayed by mixing 20 fmol of TRs produced in a reticulocyte lysate system, TNT T7 (Promega), with 300,000 cpm [{gamma}-32P]ATP-radiolabeled DR-4 and F2 oligonucleotides and 1 µg of poly(dI-dC) (Amersham Biosciences) in a 20-µl reaction (32). In cases in which TR ligand binding activity was severely affected by Cluster 1 mutations, the overall amount of translated TRs in the extracts was also verified independently by Western blot. The binding buffer contained 25 mM HEPES, 50 mM KCl, 1 mM dithiothreitol, 10 µM ZnSO4, 0.1% Nonidet P-40, 5% glycerol. After 30 min at room temperature, the mixture was loaded onto a 5% nondenaturing polyacrylamide gel that was previously run for 30 min at 200 V. To visualize the TR-DNA complexes, the gel was run at 4 °C for 120–180 min at 200 V in a running buffer containing 45 mM Tris borate (pH 8.0) and 1 mM EDTA. The gel was then fixed, dried, and exposed for autoradiography.

Statistical Analysis—All data are presented as means ± S.D. One-way ANOVA with Tukey's post-test or t test was performed using GraphPad Prism version 3.03 for Windows. Data analyzed referred to at least three independent experiments. A p value of <0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
RTH Mutations That Inhibit T3 and DNA Binding Reside in Charge Clusters—RTH mutations that inhibit homodimer formation on DNA affect positively charged Arg residues (R338W, R429Q, and R316H) (30, 31, 33–35). In addition, we found that another RTH mutation that affects a positively charged Lys residue (K342I) also inhibits homodimer formation on DNA (not shown). Investigation of TR structural models revealed that each of these amino acids lies within separate clusters of closely juxtaposed charged residues (Fig. 1A and Table I). The TR{beta} LBD contains only one similar charge cluster that is not known to be affected by RTH mutations (Cluster 4, see Table I).


View this table:
[in this window]
[in a new window]
  		TABLE I
Location and conservation of TR charge clusters

RTH mutants known to inhibit DNA binding activity are shown in bold.

 
Clusters 1 and 2 are comprised of residues that are exposed or partially exposed on the surface of the LBD and are both adjacent to the classical dimer surface at the junction of H10 and H11 (Fig. 1A and Table I). Unlike many residues that are affected by RTH mutations, none of the residues in the clusters directly contacts T3 or comprises part of a known coregulator binding surface.



View larger version (27K):
[in this window]
[in a new window]
  		FIG. 2.
Charge Cluster 1 mutations inhibit T3 activation. Maximal (Max.) activation and repression (A) and dose of T3 (B) required for half-maximal activation at a T3-regulated reporter gene with an F2/IP-6 TRE. Activities obtained with TR mutants are compared with those obtained with wild type TR{beta}, which is set to 100%. The EC50 T3 concentration for wild type TR{beta} was 15 nM for the F2 driven reporter. In A, no statistical difference was found among mutants and wild type (p > 0.05). In B, different letters over bars indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test. WT or wt, wild type.

 
The residues in Clusters 1 and 2 have the potential to engage in electrostatic interactions with each other. Cluster 1 includes Arg338 and Lys342 on H7 and two negatively charged residues on H8, Asp351 and Asp355, and is completely surface-exposed. We originally suggested that Arg338 and Lys342 engage in parallel ionic pairings with Asp355 and Asp351, respectively, based on analysis of x-ray crystal structures of the TR{alpha} LBD (12). Reinvestigation of TR{beta}-LBD structures (13) suggested another arrangement: Arg338 and Lys342 both pair with Asp351, and Asp355 is not directly engaged in the cluster (Fig. 1B). Cluster 2 includes Arg429 on H11 and Arg383 on H9, both of which are also mutated in RTH but reported not to affect DNA binding (36), and is partially surface-exposed. Here x-ray structures of TR{alpha} and TR{beta} indicated that both Arg residues pair with Glu311 on H6 in the LBD core (Fig. 1C).



View larger version (33K):
[in this window]
[in a new window]
  		FIG. 3.
Effects of Cluster 1 mutations on activities of liganded TRs in vitro. A, mutations in Cluster 1 do not affect coregulator binding. Shown are autoradiograms of SDS-polyacrylamide gels used to separate labeled TRs bound to bacterially expressed GRIP1 (amino acids 563–1121) and N-CoR (amino acids 1944–2453) in pull-down assays. The result is representative of three experiments. B, Kd, equilibrium dissociation constant. Mutants are compared with values obtained with wild type TR, which was 161.4 x 10–12 M and set to 100%. Values represent the averages of at least three determinations. C, kinetics of ligand dissociation from wild type and mutant TRs, koff. Values represent the averages of at least three determinations. In B and C, different letters over bars indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test. WT or wt, wild type; GST, glutathione S-transferase.

 
Residues in Clusters 1 and 2 show considerable conservation. They are conserved in TRs throughout vertebrate species (not shown). Residues equivalent to those in charge Cluster 1 are conserved on H7 and H8 in other NRs, including retinoic acid receptors and PPARs (Fig. 1D). Residues in Cluster 2 show even better conservation (Fig. 1E). Together all of these considerations indicate that the charge clusters play an important, and unappreciated, role in TR activities. Furthermore the fact that mutations in the clusters affect T3 binding and homodimer formation indicates that the clusters must play a role in activities associated with liganded and unliganded TRs.

Cluster 1 Is Required for Optimal T3 Binding—We first examined effects of mutations in Cluster 1 on activities of liganded TRs. Because residues of this cluster are completely surface-exposed it appeared unlikely that these mutations would exert indirect effects on TR function by disrupting internal folding of the LBD. We introduced 1) Ala substitutions, which swap a residue with a small neutral side chain for a residue with a charged side chain and thereby eliminate the potential for electrostatic interactions, and 2) charge reversal mutations, which should disrupt ionic bonds between oppositely charged residues by juxtaposing residues with like charges.

Fig. 2 shows effects of mutations on activity of transfected TR{beta} in mammalian cells. TR{beta} Cluster 1 mutants did not affect maximal activation of transcription from a TRE-driven reporter (F2) in the presence of saturating T3 or repression of basal transcription in the absence of T3 (Fig. 2A). Nevertheless several TR{beta} Cluster 1 mutants displayed altered T3 concentration dependence (Fig. 2B and Table II) both in HeLa cells (shown here) and in other cells (U2-OS and CV-1, not shown). Mutations in two residues (Arg338 and Asp351) led to reduced T3 sensitivity. The TR{beta}R338W RTH mutant required 17-fold more T3 than wild type TR{beta} for half-maximal activation (EC50). TRs bearing Ala substitutions at Arg338 and Asp351 (TR{beta}R338A and TR{beta}D351A) exhibited more modest reductions in T3 sensitivity, and TRs with charge reversal mutations at Arg338 and Asp351 (TR{beta}R338D and TR{beta}D351R) displayed more marked reductions in T3 sensitivity. In contrast, different mutations at Lys342 exhibited divergent effects. A relatively mild Ala substitution mutation (TR{beta}K342A) had either no effect or slightly enhanced T3 sensitivity (Table II). Nevertheless a more severe charge reversal mutation, TR{beta}K342D, exhibited decreased T3 sensitivity as did the TR{beta}K342I RTH mutant (not shown). Finally mutations at Asp355 did not reduce T3 sensitivity. TR{beta}D355A and TR{beta}D355R either exhibited T3 sensitivity comparable to wild type TR{beta} or enhanced T3 sensitivity at some reporters (Fig. 2B and Table II).


View this table:
[in this window]
[in a new window]
  		TABLE II
Average EC50 for T3 response obtained with TRs bearing Cluster 1 mutants at different TREs

Values are compared to wild type TR set at 100%. Mean values ± S.D. are the average of at least three experiments. Different letters in the same horizontal row indicate statistical difference (p < 0.05), and different numbers in the same vertical column indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test.

 



View larger version (56K):
[in this window]
[in a new window]
  		FIG. 4.
Mutations in charge Cluster 1 inhibit or enhance homodimerization on DNA. A–C, autoradiograph of gel shift assays of labeled F2 (A–C) and DR-4 (A and B) element oligonucleotides with wild type TR (TR{beta}wt) and various TR mutants in the presence or absence of both T3 and RXR. A, comparison of TR{beta} with TR{beta}R338W and a dimer surface mutant (L422R). B, comparison of TR{beta} with Ala substitution mutants as indicated. C, comparison of TR{beta} with charge reversal mutants. WT, wild type; Retic., reticulocyte lysate.

 
None of the Cluster 1 mutations impaired binding to a coactivator (GRIP1, Fig. 3A) or to a corepressor (N-CoR, Fig. 3A) in pull-down assays in vitro. This is consistent with the results that show no impairment in the maximal effect of the hormone in transfection assays. By contrast, the same mutations that reduced T3 sensitivity in vivo also reduced the affinity of the TR for T3 (Fig. 3B) and increased T3 dissociation rates (Fig. 3C).

Together our results indicate that Arg338, Asp351, and, to a lesser extent, Lys342 are required for optimal T3 binding and response and that Asp355 is not. This is consistent with the apparent organization of Cluster 1 in TR{beta} crystal structures where Arg338, Lys342, and Asp351 side chains engage in electrostatic interactions with each other, and Asp355 does not (Fig. 1B).

Mutations in Cluster 1 Can Either Impair or Enhance Homodimer Binding to DNA, and Effects Do Not Correlate with T3 Binding—Next we examined effects of mutations in Cluster 1 on TR homodimer formation on DNA. The data in Fig. 4A confirmed that the TR{beta}R338W RTH mutant exhibits defective homodimer formation at F2 and DR-4 elements along with normal levels of heterodimer formation (30, 31). By contrast, an artificial mutation (TR{beta}L422R) in the classical dimer interface abolished both homodimer and heterodimer formation. In parallel, TRs bearing Ala substitution mutants at Arg338 and Asp351 (both required for optimal T3 binding) exhibited reduced homodimer but not heterodimer formation at F2 and DR-4 elements (Fig. 4B) just like the TR{beta}R338W RTH mutant. However, TR{beta}K342A displayed reduced homodimer formation even though it did not inhibit T3 binding (compare Figs. 2B and 4B). More surprisingly, the charge reversal mutants exhibited enhanced DNA binding (Fig. 4C) even though most of these mutations inhibit T3 binding (Fig. 2B). The precise effect of the charge reversal mutants varied; TR{beta}R338D showed enhanced DNA binding in the absence of T3, whereas TR{beta}D351R and TR{beta}K342D showed enhanced DNA binding in the presence or absence of T3. TR{beta}D355R exhibited enhanced DNA binding in the presence of T3, reversing the usual effects of T3 on TR DNA binding activity (Fig. 4C). Again these effects were largely homodimer-specific, although TR{beta}K342D did exhibit somewhat enhanced heterodimer formation. Together our results confirm that Cluster 1 is required for TR homodimer but not heterodimer formation on DNA. Nevertheless the same data also revealed that different arrangements of charge are needed for optimal DNA and T3 binding.



View larger version (31K):
[in this window]
[in a new window]
  		FIG. 5.
Mutations in Cluster 2 differentially affect T3 response and DNA binding. A, Glu311 is required for optimal T3 response. Shown is a summary of relative EC50 values response obtained in for T3 transfections assays performed in HeLa cells with an F2-driven reporter gene as in Fig. 3. B, Arg429 is required for optimal homodimer formation. Shown are electrophoretic mobility shift assays to determine binding of Cluster 2 mutants to an F2 oligonucleotide as in Fig. 4. In A, different letters over bars indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test. WT or wt, wild type.

 
Charge Cluster 2 Residues Differentially Affect T3 Activation and DNA Binding—Mutations in Cluster 2 (Arg383, Arg429, and Glu311) also exhibited differential effects on activity of liganded TRs and DNA binding. Fig. 5A shows that TR{beta}E311A exhibited a much larger reduction in T3 sensitivity than TRs bearing mutations at Arg429 and Arg383 (TR{beta}R429A and TR{beta}R383H) (Fig. 5A). This finding is consistent with previous observations that RTH mutations in these Arg residues only affect T3 sensitivity weakly (30) and is also consistent with the organization of Cluster 2 in TR{beta} structures (Fig. 1C); a mutation in Glu311 that breaks electrostatic interactions with both Arg residues exhibited a more severe defect than mutations at Arg429 and Arg383, which only break one bond. By contrast, TR{beta}E311A (and TR{beta}R383H) bound to DNA as efficiently as wild type TRs in the absence of T3, whereas TR{beta}R429A exhibited decreased homodimer formation on DNA (Fig. 5B). Thus, Cluster 2 requires different charged residues for optimal T3 response and DNA binding just like Cluster 1.

An Arg338-Asp351 Pair Stabilizes Bound T3—To learn how individual residues within Cluster 1 interact, we examined effects of multiple mutations in the cluster upon TR function. First we reversed the positions of two residues that are most important for optimal T3 binding, Arg338 and Asp351. Fig. 6A shows that TR{beta}R338D,D351R displayed markedly reduced T3 sensitivity in transfections like the R338D and D351R single mutants. TR{beta}R338D,D351R also displayed decreased affinity for T3 and increased dissociation rates of bound T3 with normal levels of coregulator binding and TR homodimer formation on DNA (not shown).



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 6.
Reversibility of the putative Arg338-Asp351 salt bridge. A, Arg338 and Asp351 cannot be reversed without severe disruption of T3 response. Shown is a comparison of activities of wild type TR {beta} and charge reversal mutants as a function of T3 concentration. Transfections were performed in HeLa cells with an F2-driven reporter. B, Arg338 and Asp351 can be reversed in the absence of interfering charge at Lys342 and Asp355. Ala substitution mutations at Lys342 and Asp355 restore activity of the TR{beta}R338D,D351R charge reversal mutant (D/R). Data are presented as a comparison of EC50 values for different TR{beta} mutants as in Fig. 3C. In B, different letters over bars indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test. wt, wild type.

 
The phenotype of the TR{beta}R338D,D351R double mutant was surprising; it is often possible to reverse the positions of residues in ionic pairs and regenerate wild type protein function. Nevertheless Fig. 6B shows that introduction of Ala substitutions at Lys342 and Asp355 restored the activity of the TR{beta}R338D,D351R double mutant to near wild type levels but not that of a TR{beta} mutant with like charges at both positions (TR{beta}R338D,K342A,D355A). Thus, Arg338 and Asp351 can be reversed without severe loss of TR{beta} function, suggesting that they can form a reversible ionic bond that stabilizes liganded TR{beta}. This effect can only be observed, however, when other charges are removed from the cluster.

Cluster 1 Is Dispensable for Ligand TR Activity—Because Lys342 and Asp355 interfere with TR{beta} activity and T3 binding when the putative Arg338-Asp351 ionic bond is reversed (Fig. 6), we asked whether Lys342 and Asp355 might also interfere with TR{beta} activity and T3 binding in the context of wild type TR{beta}.To do this, we examined effects of multiple Ala substitutions in Cluster 1.

Mutations at Lys342 and Asp355 rescued effects of mutations at Arg338 and Asp351. Fig. 7A shows that a TR{beta} double mutant bearing Ala substitutions at residues that are required for optimal T3 binding (TR{beta}R338A,D351A) displayed reduced T3 sensitivity and T3 binding and increased dissociation rates of bound T3. Furthermore a TR{beta} double mutant bearing Ala substitutions at residues that are not required for optimal T3 binding (TR{beta}K342A,D355A) did not affect TR{beta} activity. These results confirm that Arg338 and Asp351 are needed for optimal hormone binding, and Lys342 and Asp355 are not. More surprisingly, a double mutant that eliminated both positive charges in Cluster 1 (TR{beta}R338A,K342A) exhibited a phenotype that was similar to wild type TR{beta}. Furthermore a double mutant that removed both negative charges (TR{beta}D351A,D355A) exhibited a phenotype that was intermediate between TR{beta}D351A, reduced affinity for T3, and TR{beta}D355A, similar to wild type TR{beta}. Thus, Ala substitution mutations at Lys342 and Asp355 rescue effects of similar mutations at Arg338 and Asp351.

The fact that some mutations in Cluster 1 rescue effects of others was underscored by the observation that elimination of all charge within Cluster 1 with a quadruple Ala substitution (TR{beta}4A) failed to inhibit T3 binding or liganded TR{beta} function. TR{beta}4A displayed enhanced T3 sensitivity in transfections (Fig. 7B), slightly increased affinity for T3 (Table III), and normal levels of coactivator and corepressor binding (not shown). Nevertheless TR{beta}4A exhibited strongly reduced homodimer formation on DNA (Fig. 7C). This reduction in homodimer formation, the largest obtained with any Cluster 1 mutation in this study (not shown), was paralleled by impaired repression at a TR-regulated reporter without T3 (Fig. 7B, inset).


View this table:
[in this window]
[in a new window]
  		TABLE III
Charge Cluster 1 is dispensable for T3 binding

Means ± S.D. are the average of at least three experiments. The same letters in the same column indicate no statistical difference (p > 0.05) according to t test.

 
Together our results show that, whereas two individual residues in the cluster (Arg338 and Asp351) are required for optimal T3 response and T3 binding, Cluster 1 itself is dispensable for the function of liganded TR. Nevertheless Cluster 1 is required for activities associated with unliganded TRs: homodimer formation on DNA and transcriptional repression (see "Discussion").


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we examined how TR DNA binding activity is regulated by its LBD and by ligand. To begin to understand this issue, we asked why some RTH mutations (R316H, R338W, K342I, and R429Q) that reduce the affinity of TR{beta} for T3 also inhibit binding of TR homodimers, but not heterodimers, to TREs (30, 31). We reasoned that these mutations might affect structural elements that are involved in coupling T3 binding to inhibition of DNA binding activity. We report here that each of these RTH mutations affected amino acids that lie within clusters of charged residues with potential for electrostatic interactions between individual residues in the cluster. Two of these clusters (1 and 2) are adjacent to the TR dimer/heterodimer surface (Table I and Fig. 1). The importance of the clusters is underscored by their conservation both in TRs across evolution (not shown) and in other NRs (Fig. 1, D and E) and by our studies, which revealed that mutations in Clusters 1 and 2 lead, variously, to increases and decreases in T3 binding and/or DNA binding.

The existence of functionally important clusters of charged residues on the TR LBD surface was surprising because proteins are largely stabilized by hydrophobic effects in which hydrophobic residues form the interior of the protein and charged side chains are surface-exposed, freely solvated with water (37). Nevertheless electrostatic interactions between oppositely charged side chains have been shown to provide additional stability to proteins in several contexts, including particular conformers of allosteric proteins, protein-protein interaction surfaces, and proteins in thermophilic organisms (37–40). For TRs, two RTH mutations that disrupt ionic bonds, one in Cluster 3 (TR{beta}R316H) and a single surface-exposed ionic bond between Arg243 in H3 and Glu322 at the base of H6 (TR{beta}R243Q), lead to broadening of experimental electron density in the lower part of the LBD in x-ray structures (14, 15). This confirms that electrostatic interactions between oppositely charged TR{beta} residues can stabilize the liganded TR{beta}-LBD.



View larger version (42K):
[in this window]
[in a new window]
  		FIG. 7.
Cluster 1 is dispensable for liganded TR action. A, mutations at Lys342 and Asp355 rescue effects of mutations at Arg338 and Asp351. Shown is a comparison of EC50, Kd, and koff values obtained TR with mutants with single and double Ala substitutions in Cluster 1; data are presented as in Figs. 2 and 3. B, Cluster 1 can be eliminated without loss of function for liganded TR. Shown are hormone activation profiles for human TR{beta} wild type and a quadruple Ala mutant (4xA) at an F2-TRE-regulated reporter. The data represent a single transfection assay in which standard errors are derived from multiple wells, representative of several experiments. The inset shows maximal (Max.) activation and repression obtained with TR{beta} and TR{beta}4A (4xA). C, elimination of Cluster 1 inhibits DNA binding. Shown is a gel shift comparing binding of TR{beta} and TR{beta}4A (4xAla) on an F2-TRE. In A, different letters over bars indicate statistical difference (p < 0.05) according to ANOVA and Tukey's test. WT or wt, wild type.

 
Our mutational analysis supports the notion that Clusters 1 and 2 are stabilizing elements for liganded TR. Mutations that disrupted the predicted ionic bond arrangements in Clusters 1 and 2 led to reduced T3 sensitivity, reduced affinity for T3, and increased T3 dissociation rates. These phenotypes resemble those of aforementioned TR{beta} RTH mutations that destabilize the TR LBD by breaking electrostatic interactions, R316H and R243Q (14, 15). In addition, three lines of evidence indicate that Arg338 and Asp351 form an ionic bond required for stable T3 binding. 1) Placement of like, repelling charges at Arg338 and Asp351 severely inhibited T3 binding (Fig. 2). 2) Arg338 and Asp351 could be reversed without significant disruption of T3 binding, albeit only in the absence of charge at Lys342 and Asp355 (Fig. 6). 3) TRs with double mutations at Arg338 and Asp351 exhibited phenotypes similar to single mutants, suggesting that both residues are parts of the same structural element (Figs. 6 and 7).

Nevertheless our results also suggest that the clusters adopt a different organization in unliganded TRs. Distinct arrangements of charge are required for optimal T3 binding and for DNA binding by unliganded TR homodimers (Figs. 2, 3, 4, 5). Thus, the TR{beta}K342A mutation inhibited DNA but not T3 binding. Furthermore and more strikingly, charge reversal mutations at Arg338 (R338D), Asp351 (D351R), and Lys342 (K342D) all inhibited T3 binding but not DNA binding, and a charge reversal mutation at Asp355 (D355R) did not affect T3 binding yet enhanced TR homodimer formation on DNA in the presence of T3 (Fig. 4).

Other results are hard to reconcile with the simple notion that Clusters 1 and 2 act as static stabilizing elements for liganded and unliganded TRs. Cluster 1 was dispensable for optimal T3 response and T3 binding (Fig. 7) even though Arg338 and Asp351 were required for T3 binding (Figs. 2, 3, and 7). Furthermore two Cluster 1 residues (Lys342 and Asp355) must inhibit T3 binding to some extent as judged by the fact that TR{beta}K342A and TR{beta}D355A mutants exhibited enhanced sensitivity to T3 in transfections and increased affinity for T3 in vitro and that Ala substitutions at both positions rescued effects of similar mutations at Arg338 and Asp351 (Fig. 7).

Our hypothesis to explain these observations is outlined in Fig. 8. We propose that Clusters 1 and 2 are hormone-dependent stabilizing elements for the TR LBD. We suggest that, in the unliganded state, the clusters adopt an unspecified organization that is distinct from that observed in x-ray structures of liganded TR{beta} LBDs but required for optimal homodimer formation on DNA. Given the placement of these residues, we favor the notion that the clusters comprise homodimer-specific extensions of the dimer surface that engage in flexible contacts with oppositely charged residues in partner LBDs or possibly influence homodimer formation via distant stabilizing effects on the dimer surface. We further suggest that T3 binding promotes structural rearrangements in the TR LBD that reposition the charged residues, simultaneously breaking the interactions that are needed for optimal homodimer formation on DNA and creating new ionic bonds that hold the walls of the hormone binding pocket in an appropriate configuration for stable T3 interactions.



View larger version (16K):
[in this window]
[in a new window]
  		FIG. 8.
Model to explain coupling of T3 binding to inhibition of TR{beta} DNA binding activity. The blue spheres represent positively charged residues in Cluster 1, whereas red spheres represent negatively charged residues. The charged residues are in an unspecified organization required for homodimer formation in the absence of hormone and rearrange to form the ionic bond organization detected in our x-ray structures in the presence of hormone.

 



View larger version (44K):
[in this window]
[in a new window]
  		FIG. 9.
Ligand-dependent rearrangements in RXR H7 (equivalent to TR{beta} H8). A schematic shows apoRXR H7 in orange and holo-RXR in gray with ligand (9-cis-retinoic acid (RA)) in gray and red. H7 changes backbone conformation: Lys356 and Glu352, paired in apoRXR, move from inside to outside on ligand binding.

 
Our model suggests explanations for several apparently paradoxical results. 1) Different charge arrangements are required for T3 binding and homodimer formation on TREs because the clusters adopt different organizations in the presence and absence of T3. 2) Cluster 1 could be eliminated without obvious effect on TR{beta} even though mutations in Arg338 and Asp351 inhibit T3 binding because the Arg338-Asp351 ionic bond counteracts the tendency of the Cluster 1 to revert toward the organization in the unliganded state. If Cluster 1 is eliminated, the requirement for the stabilizing element is eliminated. 3) Lys342 and Asp355 inhibited T3 binding because they stabilize the unliganded TR{beta} conformer that binds to DNA as a homodimer but only provide limited stability (Lys342) or no additional stability (Asp355) to the liganded TR{beta} conformer. Here mutation of Lys342 and Asp355 enhanced T3 response and T3 binding by counteracting the tendency of the cluster to revert to its organization in unliganded state.

We recognize that our model cannot yet be verified directly because apoTR dimer structures are not available. Nevertheless analysis of liganded and unliganded RXR crystal structures revealed evidence that is consistent with the basic predictions of our model. First charged residues in the region of RXR that is equivalent to H8 rearrange in response to ligand binding (Fig. 9). RXR Glu352 and Lys356 form an ionic bond within the interior of the unliganded LBD. Binding of 9-cis-retinoic acid twists the helix, exposing the charged side chains on the protein surface where they can pair with PPARs in RXR-PPAR heterodimers (41–45). We suggest that TR{beta} charge clusters (on H7, H8, and H11) must undergo similar ligand-dependent rearrangements. This model implies that functionally important conformational rearrangements that accompany T3 binding are not restricted to H12 and that T3 induces reorganization of the opposite face of the TR near the dimer surface.

Finally our results also lend support to the notion that TR homodimers are highly active in mediating transcriptional repression in vivo (25, 26). We observed that a TR{beta} mutant that strongly inhibited homodimer formation on TREs (TR{beta}4A) impaired the ability of unliganded TRs to suppress transcription in the absence of hormone (Fig. 7B). We predict that mutations such as those described here that either specifically inhibit or stabilize particular oligomeric forms of TR will help us to further dissect the relative roles of RXR-TR heterodimers and TR homodimers in vivo.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants DK41482 and DK51281 (to J. D. B.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Deputy director and consultant to Karo Bio AB, a biotechnology company with commercial interests in nuclear receptors. Back

|| To whom correspondence should be addressed: Diabetes Center, University of California School of Medicine, HSW1210, 513 Parnassus Ave., San Francisco, CA 94143-0540. Tel.: 415-476-6789; Fax: 415-564-5813; E-mail: webbp{at}itsa.ucsf.edu.

1 The abbreviations used are: TR, thyroid hormone receptor; T3, 3,5',5-triiodo-L-thyronine; RXR, retinoid X receptor; TRE, thyroid hormone response element; RTH, resistance to thyroid hormone syndrome; H, helix; LBD, ligand binding domain; N-CoR, nuclear receptor corepressor; AF, activation function; DR, direct repeat; IP, inverted palindrome; GRIP1, glucocorticoid receptor-interacting protein 1; ANOVA, analysis of variance; PPAR, peroxisome proliferator-activated receptor; NR, nuclear hormone receptor. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Apriletti, J. W., Ribeiro, R. C. J., Wagner, R. L., Feng, W., Webb, P., Kushner, P. J., West, B. L., Nilsson, S., Scanlan, T. S., Fletterick, R. J., and Baxter, J. D. (1998) Clin. Exp. Pharmacol. Physiol. Suppl. 25,S2 –S11[Medline] [Order article via Infotrieve]
  2. Ribeiro, R. C. J., Apriletti, J. W., Wagner, R. L., West, B. L., Feng, W., Huber, R., Kushner, P. J., Nilsson, S., Scanlan, T. S., Fletterick, R. J., Schaufele, F., and Baxter, J. D. (1998) Recent Prog. Horm. Res. 53,351 –394[Medline] [Order article via Infotrieve]
  3. Yen, P. M. (2001) Physiol. Rev. 81,1097 –1142[Abstract/Free Full Text]
  4. Zhang, J., and Lazar, M. A. (2000) Annu. Rev. Physiol. 62,439 –466[CrossRef][Medline] [Order article via Infotrieve]
  5. Glass, C. K., and Rosenfeld, M. G. (2000) Genes Dev. 14,121 –141[Free Full Text]
  6. Baxter, J. D., Dillmann, W. H., West, B. L., Huber, R., Furlow, J. D., Fletterick, R. J., Webb, P., Apriletti, J. W., and Scanlan, T. S. (2001) J. Steroid Biochem. Mol. Biol. 76,31 –42[CrossRef][Medline] [Order article via Infotrieve]
  7. Webb, P., Nguyen, N. H., Chiellini, G., Yoshihara, H. A., Cunha Lima, S. T., Apriletti, J. W., Ribeiro, R. C., Marimuthu, A., West, B. L., Goede, P., Mellstrom, K., Nilsson, S., Kushner, P. J., Fletterick, R. J., Scanlan, T. S., and Baxter, J. D. (2002) J. Steroid Biochem. Mol. Biol. 83,59 –73[CrossRef][Medline] [Order article via Infotrieve]
  8. Gothe, S., Wang, Z., Ng, L., Kindblom, J. M., Barros, A. C., Ohlsson, C., Vennstrom, B., and Forrest, D. (1999) Genes Dev. 13,1329 –1341[Abstract/Free Full Text]
  9. Wondisford, F. E. (2003) J. Investig. Med. 51,215 –220[Medline] [Order article via Infotrieve]
  10. Ribeiro, R. C., Apriletti, J. W., Wagner, R. L., Feng, W., Kushner, P. J., Nilsson, S., Scanlan, T. S., West, B. L., Fletterick, R. J., and Baxter, J. D. (1998) J. Steroid Biochem. Mol. Biol. 65,133 –141[CrossRef][Medline] [Order article via Infotrieve]
  11. Weatherman, R. V., Fletterick, R. J., and Scanlan, T. S. (1999) Annu. Rev. Biochem. 68,559 –581[CrossRef][Medline] [Order article via Infotrieve]
  12. Wagner, R. L., Apriletti, J. W., McGrath, M. E., West, B. L., Baxter, J. D., and Fletterick, R. J. (1995) Nature 378,690 –697[CrossRef][Medline] [Order article via Infotrieve]
  13. Wagner, R. L., Huber, B. R., Shiau, A. K., Kelly, A., Cunha Lima, S. T., Scanlan, T. S., Apriletti, J. W., Baxter, J. D., West, B. L., and Fletterick, R. J. (2001) Mol. Endocrinol. 15,398 –410[Abstract/Free Full Text]
  14. Huber, B. R., Desclozeaux, M., West, B. L., Cunha-Lima, S. T., Nguyen, H. T., Baxter, J. D., Ingraham, H. A., and Fletterick, R. J. (2003) Mol. Endocrinol. 17,107 –116[Abstract/Free Full Text]
  15. Huber, B. R., Sandler, B., West, B. L., Cunha Lima, S. T., Nguyen, H. T., Apriletti, J. W., Baxter, J. D., and Fletterick, R. J. (2003) Mol. Endocrinol. 17,643 –652[Abstract/Free Full Text]
  16. Borngraeber, S., Budny, M. J., Chiellini, G., Cunha-Lima, S. T., Togashi, M., Webb, P., Baxter, J. D., Scanlan, T. S., and Fletterick, R. J. (2003) Proc. Natl. Acad. Sci. U. S. A. 100,15358 –15363[Abstract/Free Full Text]
  17. Feng, W., Ribeiro, R. C. J., Wagner, R. L., Nguyen, H., Apriletti, J. W., Fletterick, R. J., Baxter, J. D., Kushner, P. J., and West, B. L. (1998) Science 280,1747 –1749[Abstract/Free Full Text]
  18. Marimuthu, A., Feng, W., Tagami, T., Nguyen, H., Jameson, J. L., Fletterick, R. J., Baxter, J. D., and West, B. L. (2002) Mol. Endocrinol. 16,271 –286[Abstract/Free Full Text]
  19. Hu, X., and Lazar, M. A. (1999) Nature 402,93 –96[CrossRef][Medline] [Order article via Infotrieve]
  20. Nagy, L., Kao, H. Y., Love, J. D., Li, C., Banayo, E., Gooch, J. T., Krishna, V., Chatterjee, K., Evans, R. M., and Schwabe, J. W. (1999) Genes Dev. 13,3209 –3216[Abstract/Free Full Text]
  21. Perissi, V., Staszewski, L. M., McInerney, E. M., Kurokawa, R., Krones, A., Rose, D. W., Lambert, M. H., Milburn, M. V., Glass, C. K., and Rosenfeld, M. G. (1999) Genes Dev. 13,3198 –3208[Abstract/Free Full Text]
  22. Ribeiro, R. C., Kushner, P. J., Apriletti, J. W., West, B. L., and Baxter, J. D. (1992) Mol. Endocrinol. 6,1142 –1152[Abstract]
  23. Cohen, R. N., Brzostek, S., Kim, B., Chorev, M., Wondisford, F. E., and Hollenberg, A. N. (2001) Mol. Endocrinol. 15,1049 –1061[Abstract/Free Full Text]
  24. Liang, F., Webb, P., Marimuthu, A., Zhang, S., and Gardner, D. G. (2003) J. Biol. Chem. 278,15073 –15083[Abstract/Free Full Text]
  25. Williams, G. R., Zavacki, A. M., Harney, J. W., and Brent, G. A. (1994) Endocrinology 134,1888 –1896[Abstract]
  26. Yoh, S. M., and Privalsky, M. L. (2001) J. Biol. Chem. 276,16857 –16867[Abstract/Free Full Text]
  27. Ribeiro, R. C., Feng, W., Wagner, R. L., Costa, C. H., Pereira, A. C., Apriletti, J. W., Fletterick, R. J., and Baxter, J. D. (2001) J. Biol. Chem. 276,14987 –14995[Abstract/Free Full Text]
  28. Chatterjee, V. K., and Beck-Peccoz, P. (1994) Bailliere's Clin. Endocrinol. Metab. 8,267 –283[CrossRef][Medline] [Order article via Infotrieve]
  29. Kopp, P., Kitajima, K., and Jameson, J. L. (1996) Proc. Soc. Exp. Biol. Med. 211,49 –61[Abstract]
  30. Collingwood, T. N., Adams, M., Tone, Y., and Chatterjee, V. K. (1994) Mol. Endocrinol. 8,1262 –1277[Abstract]
  31. Kitajima, K., Nagaya, T., and Jameson, J. L. (1995) Thyroid 5,343 –353[Medline] [Order article via Infotrieve]
  32. Ribeiro, R. C., Apriletti, J. W., Yen, P. M., Chin, W. W., and Baxter, J. D. (1994) Endocrinology 135,2076 –2085[Abstract]
  33. Ando, S., Nakamura, H., Sasaki, S., Nishiyama, K., Kitahara, A., Nagasawa, S., Mikami, T., Natsume, H., Genma, R., and Yoshimi, T. (1996) J. Endocrinol. 151,293 –300[Abstract]
  34. Takeda, T., Suzuki, S., Nagasawa, T., Liu, R. T., and DeGroot, L. J. (1999) Biochimie (Paris) 81,297 –308
  35. Takeda, T., Nagasawa, T., Miyamoto, T., Minemura, K., Hashizume, K., and Degroot, L. J. (2000) Thyroid 10,11 –18[Medline] [Order article via Infotrieve]
  36. Clifton-Bligh, R. J., de Zegher, F., Wagner, R. L., Collingwood, T. N., Francois, I., Van Helvoirt, M., Fletterick, R. J., and Chatterjee, V. K. (1998) Mol. Endocrinol. 12,609 –621[Abstract/Free Full Text]
  37. Perutz, M. F. (1978) Science 201,1187 –1191[Abstract/Free Full Text]
  38. Perutz, M. F. (1989) Trends Biochem. Sci. 14,42 –44[CrossRef][Medline] [Order article via Infotrieve]
  39. Serrano, L., Horovitz, A., Avron, B., Bycroft, M., and Fersht, A. R. (1990) Biochemistry 29,9343 –9352[CrossRef][Medline] [Order article via Infotrieve]
  40. Strop, P., and Mayo, S. L. (2000) Biochemistry 39,1251 –1255[CrossRef][Medline] [Order article via Infotrieve]
  41. Bourguet, W., Ruff, M., Chambon, P., Gronemeyer, H., and Moras, D. (1995) Nature 375,377 –382[CrossRef][Medline] [Order article via Infotrieve]
  42. Renaud, J. P., Rochel, N., Ruff, M., Vivat, V., Chambon, P., Gronemeyer, H., and Moras, D. (1995) Nature 378,681 –689[CrossRef][Medline] [Order article via Infotrieve]
  43. Gampe, R. T., Jr., Montana, V. G., Lambert, M. H., Miller, A. B., Bledsoe, R. K., Milburn, M. V., Kliewer, S. A., Willson, T. M., and Xu, H. E. (2000) Mol. Cell 5,545 –555[CrossRef][Medline] [Order article via Infotrieve]
  44. Bourguet, W., Vivat, V., Wurtz, J. M., Chambon, P., Gronemeyer, H., and Moras, D. (2000) Mol. Cell 5,289 –298[CrossRef][Medline] [Order article via Infotrieve]
  45. Gampe, R. T., Jr., Montana, V. G., Lambert, M. H., Wisely, G. B., Milburn, M. V., and Xu, H. E. (2000) Genes Dev. 14,2229 –2241[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?
</