Contacts between membrane proximal regions of the PDGF receptor ectodomain are required for receptor activation but not for receptor dimerization

Abstract

The mechanism of PDGF-receptor β (PDGFRβ) activation was explored by analyzing the properties of mutant receptors designed based on the crystal structure of the extracellular region of the related receptor tyrosine kinase KIT/stem cell factor receptor. Here, we demonstrate that PDGF-induced activation of a PDGFRβ mutated in Arg-385 or Glu-390 in D4 (the fourth Ig-like domain of the extracellular region) was compromised, resulting in impairment of a variety of PDGF-induced cellular responses. These experiments demonstrate that homotypic D4 interactions probably mediated by salt bridges between Arg-385 and Glu-390 play an important role in activation of PDGFRβ and all type III receptor tyrosine kinases. We also used a chemical cross-linking agent to covalently cross-link PDGF-stimulated cells to demonstrate that a Glu390Ala mutant of PDGFRβ undergoes typical PDGF-induced receptor dimerization. However, unlike WT PDGFR that is expressed on the surface of ligand-stimulated cells in an active state, PDGF-induced Glu390Ala dimers are inactive. Although the conserved amino acids that are required for mediating D4 homotypic interactions are crucial for PDGFRβ activation, these interactions are dispensable for PDGFRβ dimerization. Moreover, PDGFRβ dimerization is necessary but not sufficient for tyrosine kinase activation.

The generally accepted mechanism of receptor tyrosine kinase (RTK) activation is that ligand-induced receptor dimerization facilitates transautophosphorylation of critical regulatory tyrosine residues in the activation loop of the catalytic core, a step essential for tyrosine kinase activation. This is followed by autophosphorylation of multiple tyrosine residues in the cytoplasmic domain that serve as binding sites for Src homology 2 (SH2) or phosphotyrosine-binding (PTB) domains of a variety of signaling proteins, which upon recruitment and/or tyrosine phosphorylation transmit signals to a variety of intracellular compartments in a regulated manner (1–3).

Although nearly all RTKs are activated by dimerization, different RTK families have evolved to use different molecular strategies for ligand-induced receptor dimerization and activation. Dimerization and activation of members of the EGF receptor (EGFR) family is mediated by interactions between EGF or TGFα with the extracellular region (ectodomain) of EGFR, which exposes a buried dimerization interface that facilitates receptor-mediated EGFR dimerization and formation of an activated EGF/EGFR 2:2 complex (4). Dimerization of the FGF receptors (FGFRs), however, is mediated by tripartite interactions among monomeric FGF molecules, heparan sulfate proteoglycans (HSPG), and FGFR molecules to stabilize the formation of an active ternary FGF/HSPG/FGFR 2:2:2 complex (5). By contrast to EGF or FGF, all ligands of type III RTKs, including PDGFs, stem cell factor (SCF), colony-stimulating factor (CSF), and Flt3-ligand (Flt3L), are dimeric molecules capable of cross-linking their cognate receptors by bivalent binding to equivalent sites of two neighboring receptor molecules. Each PDGF protomer is composed of a central four-stranded β-sheet with the characteristic cystine knot at one end of the molecule. Two PDGF protomers are arranged in an antiparallel manner and linked to each other by two interchain disulfide bridges (6). By contrast each SCF, CSF, or Flt3L protomer is composed of a short helical fold, and they are connected to each other by noncovalent interactions (7–10). Despite their diverse folds, the two growth factor subtypes bind to and activate their cognate RTKs in a virtually identical manner, resulting in formation of activated ligand/RTK 2:2 complexes (10). All type III RTKs are composed of an extracellular ligand-binding region containing five tandem Ig-like domains followed by a single transmembrane helix and a cytoplasmic tyrosine kinase domain with a large kinase-insert region flanked by regulatory regions that are subject to autophosphorylation and to phosphorylation by heterologous protein kinases (11).

The elucidation of the x-ray crystal structure of the entire ectodomain of KIT/stem cell factor receptor before and after SCF stimulation provided valuable insights concerning the mechanism of SCF-induced KIT dimerization and activation (12). The structure shows that the first three Ig-like domains of KIT, designated D1, D2, and D3, are responsible for SCF binding. The main role of SCF binding is to cross-link two KIT molecules to increase the local concentration of KIT on the cell membrane. This facilitates a large conformational change in the membrane-proximal regions of KIT, resulting in a homotypic interaction between D4 or D5 of neighboring KIT molecules. The lateral interactions between D4 of two neighboring KIT molecules occur via direct contacts through two pairs of salt bridges from EF loops of each D4 protomer. The membrane proximal D5 domain provides additional indirect interactions between neighboring KIT molecules to further stabilize and position the membrane proximal part of the ectodomain at a distance and orientation that enables the activation of cytoplasmic tyrosine kinase.

On the basis of the structure-based sequence alignment of type III RTKs ectodomains and a homology model of the PDGF receptor (PDGFR) D4 structure, we identified amino acids in the PDGFRβ D4 domain that may form salt bridges similar to those shown to mediate homotypic D4 interactions essential for SCF-induced KIT activation. In this report, we demonstrate that PDGF-induced activation of PDGFRβ is compromised when Arg-385 and Glu-390 in D4 were mutated to alanine residues. Furthermore, a variety of cellular responses that depend on PDGFRβ activation either are reduced or their kinetics strongly attenuated. We also apply a chemical cross-linking agent to covalently cross-link intact unstimulated or PDGF-stimulated cells to demonstrate that an E390A point mutation in D4 does not interfere with PDGF-induced receptor dimerization. However, unlike the covalently cross-linked WT PDGFRβ dimers that are displayed on the cell surface in an activated state, the covalently cross-linked dimers of the E390A mutant are inactive. These experiments demonstrate that the conserved amino acids that take part in formation of D4 homotypic interactions play a critical role in PDGFRβ activation. However, the D4 homotypic interactions are dispensable for PDGFRβ dimerization. Although bivalent PDGF binding is the driving force for PDGFRβ dimerization, dimerization itself is necessary but not sufficient for tyrosine kinase activation.

Results

Similar to the D4 domain of KIT, D4 of PDGFRα and PDGFRβ lack a characteristic disulfide bond that bridges cysteine residues located in B5 and F5 in Ig-like domains. The amino acid sequence alignment presented in supporting information (SI) Fig. S1 shows that 13 of 20 fingerprint residues of the I-set IgSF fold are conserved in D4 of PDGFRs, and that the number and length of strands corresponding to the fingerprint residues are highly conserved in D4 of KIT, PDGFRα, PDGFRβ, and CSF1R (12).

D4 of KIT is composed of two β sheets, each containing four strands, with the arrangement ABED/A′GFC, and the homotypic D4 contacts are mediated by the EF loop of D4 projecting from two neighboring KIT molecules (12). The KIT structure shows that Arg-381 and Glu-386 in the EF loop form salt bridges and van der Waals contacts across a twofold axis of KIT dimer. In addition, the side chains of Arg-381 of each protomer form hydrogen bonds with the main chain carbonyl of the corresponding residue of neighboring KIT molecules (12). Structure-based sequence alignment has shown that the size of the EF loop and the critical amino acids comprising the D4–D4 interface are conserved in KIT, PDGFRα, PDGFRβ, and CSF1R. In PDGFRα, Glu-386 is replaced by an aspartic acid, a residue that may also function as a salt-bridge partner. In addition, a pair of basic and acidic (Glu/Asp) residues is strictly conserved in PDGFRα and PDGFRβ of different species ranging from Takifugu rubripes to Homo sapiens (Fig. S1), providing further support for the functional importance of this region.

PDGF-Induced PDGFR Activation Is Compromised by Mutations in D4.

The amino acid sequence alignment presented in Fig. S1 demonstrates that Arg-385 and Glu-390 in the EF loop of PDGFRβ may mediate homotypic D4 interactions similar to the salt bridges formed between Arg-381 and Glu-386 of KIT that are responsible for mediating homotypic D4 interactions between neighboring KIT receptors (Fig. 1). To investigate whether a similar mechanism is used by PDGFRβ, Arg-385, and Glu-390, each alone (R385A, E390A) or in combination (RE/AA), were substituted by alanine residues. An additional conserved Lys-387 residue in the loop region was also substituted by an alanine (RKE/AAA) residue to examine its potential role in control of PDGF-induced PDGFRβ activation. WT and mutant PDGFRβ s were stably expressed in fibroblasts derived from mouse embryos (MEFs) deficient in both PDGFRα and PDGFRβ (13–15). MEFs expressing WT or mutant PDGFRβ s matched for expression level were used in the experiments described below. The experiment presented in Fig. 2A shows that PDGF-induced tyrosine autophosphorylation of PDGFRβ is strongly compromised in cells expressing the E390A, R385A, RE/AA, and RKE/AAA mutants of PDGFRβ; both the magnitude (Fig. 2A) and kinetics (Fig. S2) of tyrosine autophosphorylation were reduced and attenuated, respectively. These experiments demonstrate that Arg-385 and Glu-390 in the EF loop of D4 play an important role in PDGF-induced stimulation of PDGFRβ, which suggests that a similar pair of salt bridge to those identified in KIT structure may exist in activated PDGFRs. Direct interaction between D4 of neighboring receptor within the ligand–receptor complex may represent a common mechanism used for ligand-induced activation of type III RTKs. We have consistently and reproducibly observed that PDGF-induced receptor autophosphorylation is more strongly compromised in cells expressing the E390A compared with cells expressing the R385A, RE/AA, or the RKE/AAA mutants. Although the precise mechanism responsible for the difference between these mutants is not clear it is possible that the positive local surface charge at the D4 interface may cause electrostatic repulsion to maintain D4 of neighboring receptors apart before ligand stimulation. Whereas substitution of Arg-385 by an alanine residue will prevent salt bridge formation this change may also decrease the net positive charge in the D4-D4 interface resulting in weaker inhibition of PDGFR activation.

Fig. 1.

Homology modeling of membrane proximal region of PDGFRs. The membrane proximal region of PDGFRβ ectodomain is shown as ribbons with transparent molecular surface (D4 colored in gold and D5 in magenta) (Left). A closer view (Right) of the D4–D4 interface of two neighboring PDGFRβ molecules demonstrates that interactions between D4 are mediated by residues Arg-385 and Glu-390 projected from two adjacent EF loop. Key amino acids are labeled and shown as a stick model.

Fig. 2.

PDGF-induced PDGFR activation is compromised by mutations in D4. (A) PDGFRα/β−/− MEFs expressing WT PDGFRβ and various PDGFRβ D4 mutants (R385A, E390A, RE/AA, and RKE/AAA) were serum-starved overnight and stimulated with indicated PDGF concentration for 5 min at 37°C. Cell lysates were immunoprecipitated with anti-PDGFR antibodies, followed by immunoblotting with antiphosphotyrosine antibodies 4G10. Membranes were stripped off and reblotted with anti-flag tag antibodies. (B) PDGFRα/β−/− MEFs expressing WT (■), R385A (▴), E390A (▾), and RE/AA (♦) PDGFRβ were incubated with 5 ng/ml ¹²⁵I-PDGF at 4°C for 90 min in the presence of increasing concentration of native PDGF. Cell-associated ¹²⁵I-PDGF were collected with 0.5 M NaOH solution and quantitated with a scintillation counter. The IC₅₀ values were determined by curve fitting with Prism4 (GraphPad). (C) MEFs expressing WT, R385A, E390A, RE/AA, and RKE/AAA PDGFR were serum-starved overnight and lysed. Cell lysates were immunoprecipitated with anti-PDGFR antibodies, and immunopellets were subjected to in vitro autophosphorylation assay in the absence (−) or presence (+) of 1 mM ATP and 10 mM Mg²⁺ for 10 min at room temperature. Pellets were resolved with SDS/PAGE followed by immunoblotting with antiphosphotyrosine antibodies and antiflag antibodies.

To examine the possibility of whether mutation in D4 of PDGFR may have affected cell membrane expression and ligand-binding affinity of mutant PDGFRβs, we next performed quantitative PDGF-binding experiments to cells expressing WT or mutant PDGFRβs. Cells expressing WT, R385A, E390A, or the RE/AA PDGFRβ mutants were incubated with a buffer solution containing ¹²⁵I-PDGF for 90 min at 4°C in the presence of increasing concentration of native PDGF. Cell-bound radioactivity was measured by using a scintillation counter. The EC₅₀ values of the displacement curves of WT and mutant PDGFRβs were analyzed by curve fitting with Prism4 (Fig. 2B). We have also compared the amount of WT and mutant PDGFRβs expressed in the transfected MEFs by immunoblotting of total cell lysates with antibodies against PDGFR or antitag antibodies (Fig. 2 A and C). Taken together, these experiments demonstrate that similar amount of WT or mutant PDGFRβs are expressed on the cell surface of the transfected cells. Moreover, similar IC₅₀ values (PDGF concentration that displaces 50% of ¹²⁵I-PDGF binding) were obtained for cells expressing WT (3.7 nM), R385A (6.0 nM), E390A (2.8 nM), or the RE/AA (3.0 nM) mutants. We also examined the possibility of whether the intrinsic tyrosine kinase activity of mutant PDGFRβs was adversely affected by comparing the in vitro tyrosine kinase activities of WT and mutant receptors. In this experiment, cell lysates from serum-starved cells were subjected to immunoprecipitation with anti-PDGFR antibodies, and the immobilized PDGFRs were subjected to in vitro kinase assays in the presence of 1 mM ATP and 10 mM magnesium chloride. After incubation, the samples were analyzed by immunoblotting with antiphosphotyrosine antibodies. The experiment presented in Fig. 2C demonstrates that the R385A, E390A, or RE/AA mutations do not influence the intrinsic tyrosine kinase activity of PDGFR. Together, these experiments demonstrate that the mutations in D4 that affect PDGF-induced stimulation of PDGFRβ do not alter the expression of PDFGRβ on the cell surface, influence the ligand binding affinity of PDFGRβ, or alter the intrinsic tyrosine kinase activities of mutant PDGFRβ.

PDGFR D4 Point Mutants Are Expressed on the Surface of PDGF-Stimulated Cells in the Form of Inactive Dimers.

Because receptor dimerization has been established as critical mechanism underlying RTK activation, we investigated whether reduced tyrosine autophosphorylation of mutant PDGFRβ in response to PDGF stimulation is caused by deficiency in receptor dimerization. We have previously applied chemical cross-linking agents to monitor and follow ligand-induced dimerization of several cell membrane receptors, including WT, and a variety of EGF receptor mutants on the cell surface of living cells (16). In this experiment, cells expressing WT PDGFRβ or the E390A mutant were serum-starved overnight, followed by PDGF incubation for 90 min at 4°C. Several washes were used to remove unbound PDGF, and the cells were incubated with 0.5 mM disuccinimidyl suberate (DSS) in PBS for 30 min at 25°C. Cell lysates from unstimulated or PDGF-stimulated cells were subjected to immunoprecipitation with anti-PDGFR antibodies followed by SDS/PAGE and immunoblotting with either antiflag antibodies to monitor the status of PDGFR dimerization or with antiphosphotyrosine antibodies to monitor the status of PDGFR activation (Fig. 3).

Fig. 3.

PDGF-stimulated PDGFRβ mutated in D4 are expressed on the cell surface in the form of inactive dimers. PDGFRα/β−/− MEFs expressing WT PDGFRβ or E390A mutant were serum-starved overnight, followed by incubation with the indicated amount of PDGF at 4°C for 90 min. After removing the unbound ligand, cells were incubated with 0.5 mM DSS in PBS for 30 min. Cell lysates were immunoprecipitated with anti-PDGFR antibodies, and immunopellets were analyzed by SDS/PAGE and immunoblotted with anti-flag antibodies (Left) and antiphosphotyrosine antibodies (Right), respectively.

The experiment depicted in Fig. 3 demonstrates that, in lysates of unstimulated cells, a band that migrates in SDS gel with an apparent molecular mass of 180 kDa corresponding to PDGFR monomers was detected in lysates from cells expressing either WT PDGFRβ or the E390A mutant. Upon PDGF stimulation, an additional band that migrates in SDS gel with an apparent molecular mass of 360 kDa corresponding to PDGFR dimers was detected in cells expressing both WT PDGFRβ and the E390A mutant. However, immunoblotting of the samples with antiphosphotyrosine antibodies demonstrates that, whereas the band corresponding to dimers of WT PDGFR is strongly tyrosine-phosphorylated, very weak tyrosine phosphorylation of the band corresponding to the dimers of E390A mutant is detected (Fig. 3). This experiment shows that impaired ligand-induced tyrosine autophosphorylation of the E390A mutant is not caused by deficiency in ligand-induced receptor dimerization. It also demonstrates that the covalently cross-linked WT PDGFRβ are displayed on the cell surface of PDGF-stimulated cells in the form of active dimers, whereas the E390A mutant is displayed on the surface of PDGF-stimulated cells in the form of inactive dimers. We conclude that the D4 homotypic interactions in PDGFR are dispensable for receptor dimerization, and that PDGF-induced receptor dimerization is necessary (17) but not sufficient for tyrosine kinase activation (Fig. 3).

Impaired Stimulation of Cells Signaling in Cells Expressing D4 PDGFR Mutants.

We next examined the impact of PDGFR D4 mutations on cell signaling in response to PDGF stimulation. Lysates from unstimulated or PDGF-stimulated cells expressing either WT or PDGFR D4 mutants were subjected to immunoprecipitation with antiphospholipase Cγ (anti- PLCγ) antibodies followed by SDS/PAGE and immunoblotting with either anti-PLCγ or anti-pTyr antibodies. The experiment presented in Fig. S3 shows that tyrosine phosphorylation of PLCγ is severely compromised in cells expressing the R385A, E390A, RE/AA, or the RKE/AAA PDGFR mutants. Impaired stimulation of additional PDGF-induced cellular responses are observed in cells expressing PDGFR D4 mutants. The experiment presented in Fig. 4A shows that MAPK response and Akt stimulation were strongly compromised in cells expressing the R385A, E390A, RE/AA, or RKE/AAA PDGFR mutants, as compared with similar responses induced by PDGF in MEFs expressing WT PDGFRs. Overall, ≈10-fold higher concentrations of PDGF were required for a similar level of MAPK response and Akt stimulation in cells expressing the E390A, RE/AA, or RKE/AAA PDGFR mutants.

Fig. 4.

PDGF-induced cellular responses are compromised by mutations in PDGFRβ D4 mutant. (A) Cells were stimulated with 10 ng/ml PDGF for 5 min, as described above. Total lysates were subjected to SDS/PAGE and analyzed by immunoblotting with antiphospho-MAPK, MAPK, phospho-Akt, and Akt antibodies. (B) Cells seeded on coverslips were serum-starved for 16 h and either left untreated or stimulated with 50 ng/ml PDGF for 2, 5, 10, or 30 min. Coverslips were stained with FITC-phalloidin, and the percentage of cells showing dorsal actin rings were quantitated and presented linearly in C. WT (■), R385A (□), E390A (▴), and RE/AA (Δ).

One of the hallmarks of PDGF stimulation of cultured fibroblasts is a typical formation of membrane ruffles and circular actin ring structures on the dorsal surface of PDGF-stimulated cells. The experiment presented in Fig. 4 B and C shows that PDGF stimulation of actin ring formation is compromised in MEFs expressing PDGFR D4 mutants. Although ≈83% of MEFs expressing WT PDGFR exhibited circular actin ring formation, only 5% of PDGFR D4 mutant cells showed similar circular actin ring formation after 2-min stimulation with 50 ng/ml of PDGF. Furthermore, the transient circular actin ring formation that peaks in MEFs expressing WT PDGFR after 2–5 min of PDGF stimulation was weakly detected in cells expressing the R385A, E390A, or the RE/AA PDGFR mutants.

Reduced Internalization and Degradation of D4 PDGFR Mutants.

We next examined the effect of PDGFR D4 mutations on PDGF internalization, PDGFR degradation, and PDGFR ubiquitination. MEFs expressing WT PDGFR or the PDGFR D4 mutants were treated with 5 ng/ml of ¹²⁵I-labeled PDGF for 90 min at 4°C followed by brief washes with PBS (pH 7.4) to remove the excess ligand in the medium. Prelabeled cells were warmed to 37°C to initiate the endocytosis of ligand–receptor complex for various time intervals up to 4 h. Cell surface-bound, intracellular, and degraded ¹²⁵I-PDGF in medium were collected, quantitated by using a scintillation counter, and presented as percentage of total cell-associated ¹²⁵I-PDGF radioactivity after 90-min incubation (t = 0) at 4°C (mean ± SD). The experiment presented in Fig. 5A shows that the kinetics of internalization of ¹²⁵I-labeled PDGF bound to MEFs expressing WT PDGFR is much faster than the kinetics of internalization of ¹²⁵I-labeled PDGF bound to cells expressing the E390A, R385A, or the RE/AA (data not shown) PDGFR mutants. After 30 min, ≈75–80% of ¹²⁵I PDGF was removed from the cell surface and accumulated inside the cells expressing WT receptors compared with <50% in cells expressing mutant receptors. The low molecular mass degradation product of ¹²⁵I-PDGF became detectable after 30 min. The release of degraded ¹²⁵I-PDGF was much slower in E390A mutant cells than in WT cells (Fig. 5A). Reduced PDGF internalization and degradation were reflected in reduced degradation of PDGFR D4 mutants. Cells expressing WT or the R385A, E390A, or RE/AA PDGFR mutants were first incubated 30 min with cycloheximide, to prevent the biosynthesis of new PDGFR molecules during the degradation experiment. The experiment presented in Fig. 5B shows that the kinetics of degradation of R385A, E390A, or the RE/AA PDGFR mutants was strongly attenuated, whereas half of WT PDGFRs were degraded within 1.5 h of PDGF stimulation, the half-life for PDGFR D4 mutants was extended to ≈4 to 6 h. The experiment presented in Fig. S4 shows that PDGF-induced stimulation of ubiquitination of the E390A PDGFR was also strongly reduced as compared with WT PDGFR under similar conditions. Taken together, these experiments demonstrate that PDGFR internalization and ubiquitin-mediated PDGFR degradation are compromised by mutations in D4 of PDGFR.

Fig. 5.

Altered kinetics of the ligand–receptor complex internalization and receptor degradation in cells expressing PDGFR D4 mutants. (A) Cells were incubated with 5 ng/ml ¹²⁵I-PDGF for 90 min, and unbound ligand was removed. Cells were transferred to 37°C for the indicated time intervals. Cell surface receptor-associated (■), internalized (▴), and degradation product (▾) of ¹²⁵I-PDGF were determined and expressed as percentage of total binding at t = 0 min. Each point was performed in triplicates and presented as mean ± standard error. (B) Cells were pretreated with 10 μg/ml cycloheximide for 30 min before PDGF stimulation. PDGF (20 ng/ml) was added for the indicated time. Cell lysates were immunoprecipitated with PDGFR antibodies and immunoblotted with anti-flag antibodies. Total cell lysates were immunoblotted with anti-actin antibodies as control.

Discussion

The extracellular domains of all members of type III RTKs, including PDGFRα, PDGFRβ, CSF1R, Flt3, and KIT, are composed of five Ig-like domains, of which the first three function as binding sites for the dimeric ligand molecule, which upon binding stimulates receptor dimerization and activation. Because the molecular architecture, ligand-binding characteristics, and mechanism of receptor dimerization of type III RTKs are highly conserved, the mechanism of SCF-induced KIT activation revealed by the crystal structures of the complete extracellular domain of KIT before and after SCF stimulation may represent a general mechanism of activation of all type III RTKs. Moreover, phylogenic analysis of RTKs containing Ig-like domains in their extracellular domains suggests a common evolutionary origin for types III and IV RTK, a family including VEGFR1 (Flt1), VEGFR2 (KDR), and VEGFR3 (Flt4) (18). Moreover, both VEGF and PDGF belong to the same cystine-knot family, homodimeric growth factors, sharing similar topology, size, and receptor-binding strategy. The salient features of KIT activation revealed by the x-ray structural analysis of its extracellular domain may, therefore, also apply for ligand-induced activation of type IV RTKs.

The structural analysis of KIT has shown that a pair of salt bridges formed between Glu-386 and Arg-381 of two neighboring D4 domains are responsible for mediating homotypic D4 interactions that are essential for SCF-induced KIT activation. Comparison of the amino acid sequences of type III RTKs demonstrates that an identical sequence motif exists in the EF loop region of D4 of PDGFRα, PDGFRβ, and CSF1R (Fig. 1 and Fig. S1), suggesting that a similar salt bridge may also be formed between D4 of type III RTKs. Indeed, substitution of Arg-385 or Asp-390 in D4 of PDGFRβ by alanines has compromised PDGF stimulation of PDGFRβ activation resulting in impairment of a variety of cellular responses that are stimulated by PDGF in cells expressing WT PDGFRβ. The mechanism of ligand-induced KIT activation revealed by analysis of KIT structure may therefore apply for the activation of all type III RTKs.

Studies exploring a variety of receptor mutants or using monoclonal antibodies that bind specifically to individual Ig-like domains of KIT (19), PDGFRs (20), and other type III RTKs have proposed that D4 plays a role in mediating receptor dimerization even when KIT is stimulated by monovalent SCF ligands (21). However, quantitative analysis using microcalorimetry of SCF binding and SCF stoichiometry toward a purified extracellular domain of KIT composed of either the first three Ig-like domains (D1–D3) or all five Ig-like domains (D1–D5) have shown that D4 and D5 are dispensable for SCF stimulation of KIT dimerization. In other words, KIT dimerization is primarily driven by the dimeric nature of SCF binding to KIT (22). Rather than playing a role in receptor dimerization, the homotypic D4 and presumably also homotypic D5 interactions between neighboring receptors are required for precise positioning of the membrane proximal regions of two receptors at a distance and orientation that enable interactions between their cytoplasmic domains resulting in tyrosine kinase activation. Therefore, rather than interfering with receptor dimerization, monoclonal antibodies that bind to D4 of PDGFRs, KIT, or other type III RTK most likely exert their inhibitory effect on receptor activation by preventing critical homotypic interactions between membrane proximal regions of type III RTK that are essential for positioning the cytoplasmic domain at a distance and orientation essential for tyrosine kinase activation.

The experiments presented in this report, together with earlier biophysical (22) and structural (12) studies, demonstrate that dimerization of PDGFRβ, KIT, and other type III RTKs is entirely driven by ligand binding, and that the sole role of ligand binding is to cross-link two receptor molecules to increase their local concentration in the cell membrane. The two salt bridges (with interface of a buried surface area of 360 Ų) responsible for mediating homotypic D4 interactions are too weak to support receptor interactions without the support of ligand-mediated receptor dimerization, which in the case of KIT is mediated by a variety of strong interactions with a total buried surface area of 2,060 Ų for each SCF protomer. We have used an approach based on “average distance to nearest neighbor calculation” (23, 24) to estimate that the apparent concentration of a receptor in the cell membrane of an unstimulated cell expressing 20,000 receptors per cell is ≈1–3 μM. Upon binding a dimeric ligand such as SCF, two occupied receptors are held together at a distance of 75 Å (12). Under this condition, the apparent receptor concentration in the cell membrane calculated by using the average distance to nearest-neighbor approach is increased by >2 orders of magnitude to 4–6 × 10⁻⁴ M. This calculation shows that even weak interactions with a dissociation constant in the range of 10⁻⁴-10⁻⁵ M, such as those mediated by the two salt bridges, could mediate association and direct contacts between membrane proximal regions of two neighboring receptors. The high local concentration in the cell membrane together with the flexibility of the joints connecting D4 and D5 to the rest of the receptor molecule enable movement and formation of homotypic D4 and homotypic D5 contacts that position the membrane proximal region of the receptor at a precise orientation and distance (15 Å in the case of KIT) that enable interactions between neighboring cytoplasmic domains, tyrosine autophosphorylation, and stimulation of tyrosine kinase activity.

Finally, applying a chemical cross-linking agent to covalently cross-link WT or mutant receptors on unstimulated or PDGF-stimulated cells, we demonstrate that an E390A PDGFRβ mutant undergoes PDGF-induced dimerization similar to PDGF-induced dimerization of WT receptors. However, by contrast to WT PDGFRβ that is expressed on the cell surface of PDGF-stimulated cells in the form of activated dimers, the E390A mutant is expressed on the surface of PDGF-stimulated cells in the form of inactive dimers. This experiment demonstrates that homotypic D4–D4 interactions are dispensable for PDGFRβ dimerization, and that PDGFRβ dimerization is necessary but not sufficient for receptor activation.

Experimental Procedures

Cell Lines and Retroviral Infection.

MEFs deficient in both PDGFRα and PDGFRβ (PDGFRα/β) were provided by Philippe Soriano (Fred Hutchinson Cancer Research Center, Seattle) and Andrius Kazlauskas (Harvard Medical School, Boston). PDGFRβ cDNA was provided by Daniel DeMaio (Yale University, New Haven, CT). PDGFRβ cDNA was subcloned into pLXSHD retroviral vector, and a flag-tag was added to the C terminus of the receptor. All mutants in D4 were generated by site-directed mutagenesis according to manufacturer's instruction (Stratagene). Retrovirus encoding WT and mutant PDGFRβ was produced in 293GPG cells (25). After infection, cells were selected with l-histidinol, and pools of selected cells were used in the experiments.

In Vitro Phosphorylation Assay for PDGFR.

Cells were serum-starved for 16 h and solubilized in lysis buffer containing 150 mM NaCl, 50 mM Hepes (pH 7.4), 1 mM EDTA, 25 mM NaF, 0.1 mM sodium orthovanadate, 5 μg/ml leupeptin and aprotinin, 1 mM PMSF, and 1% Nonidet P-40. Lysates were immunoprecipitated with anti-PDGFRβ antibodies, and immunopellets were incubated in reaction buffer containing 50 mM Hepes (pH 7.4), 1 mM ATP, and 10 mM MgCl₂ at room temperature for 5 min. After incubation, pellets were analysis by SDS/PAGE followed by immunoblotting with antiphosphotyrosine antibodies. Membrane was stripped off and reblotted with anti-Flag tag antibodies for determination of total PDGFRβ level.

Chemical Cross-Linking of Receptor Dimers.

Cells were serum-starved for 16 h before incubation with the indicated concentration of PDGF in DMEM containing 50 mM Hepes (pH 7) at 4°C. After 90 min, the cells were extensively washed with PBS (pH 7.4). Plates were transferred to room temperature, and disuccinimidyl suberate (DSS) was added to a final concentration of 0.5 mM. The cross-linking reaction was quenched after 30 min by incubation with 10 mM Tris buffer. Cell lysates were immunoprecipitated with anti-PDGFR antibodies and resolved by SDS/PAGE. Nitrocellulose membrane was immunoblotted with antibodies against flag-tag or antiphosphotyrosine (4G10) antibodies to detect the total receptor and phosphorylated receptor level, respectively.

PDGF-Induced Actin Cytoskeletal Reorganization.

Followed by overnight serum starvation, cells were either treated with 50 ng/ml PDGF for 2, 5, 10, or 30 min or left untreated. Cells were fixed in 4% paraformaldehyde in PBS, permeabilized with 0.1% Triton and stained with FITC-phalloidin (Sigma). Coverslips were mounted with Prolong Antifade mounting medium (Invitrogen), and images were acquired with Nikon fluorescence microscope. Approximately 400 cells on each coverslip were analyzed, and the percentage of cells showing actin ring formation was calculated and presented linearly.

PDGF Binding and Internalization Experiments.

PDGF was labeled by using Bolton–Hunter reagent (Pierce) before iodination by using Iodo-gen iodination tubes (Pierce), according to the manufacturer's instructions. Cells were washed twice in cold DMEM containing 20 mM Hepes (pH 7.4) and 0.1% BSA. Triplicate wells were incubated with 5 ng/ml of ¹²⁵I-PDGF in the presence of increasing amounts of native PDGF. Binding was allowed to proceed at 25°C for 1 h. Cells were then washed in cold PBS and solubilized in 0.5 M NaOH. The radioactive content of the samples was determined by using a LS6500 scintillation counter (Beckman Coulter), and the data were analyzed by using PRISM software (GraphPad).

For the internalization experiment, cells were incubated with 5 ng/ml ¹²⁵I-PDGF in DMEM/0.1% BSA/50 mM Hepes, pH 7.4, for 90 min at 4°C. Unbound ligand was removed by washing with ice-cold PBS (pH 7.4). Prewarmed DMEM/0.1% BSA/50 mM Hepes was added to the cells and incubated at 37°C for the time indicated. Cell surface-associated ligand was collected with ice-cold acidic buffer containing PBS (pH 3) and 0.1% BSA for 10 min. Internalized ligands were collected by solubilization with 0.5 M NaOH. The amount of degraded ¹²⁵I-PDGF was determined by precipitation of the incubation medium with 10% trichloroacetic acid (TCA) and counting the supernatant for the TCA soluble fraction. Radioactive content of the samples was determined by using a LS6500 scintillation counter (Beckman Coulter). The amount of surface-bound, intracellular, and degraded PDGF was expressed as a percentage of total cell-associated radioactivity after 90-min incubation on ice (t = 0 min). Each time point was performed in triplicate, and the results were expressed as mean ± SE.

For additional experimental procedures, see SI Experimental Procedures.