A cell-based receptor/adaptor translocation assay to specifically monitor FGFR3 activity. Plant extracts have long been recognized as a valuable source of bioactive molecules for pharmacotherapy and drug development (16). In this study, our focus was to identify natural compounds from plant extracts that specifically inhibit FGFR3 at early activation stages. To achieve this aim, we established a cell screening system capable of specifically monitoring early FGFR3 activation, while minimizing interference from autofluorescence or fluorescence-quenching compounds present in plant extracts. The activation of FGFR3 initiates a cascade of events, including receptor dimerization, tyrosine phosphorylation in the receptor kinase domain, adaptor protein recruitment, and internalization of the activated FGFR3-adaptor protein complex (3). We designed a system to monitor early FGFR3 activation by tracking the subcellular localization of an adaptor/GFP fusion protein that specifically interacts and internalizes with activated FGFR3. The schematic representation of the whole-cell FGFR3 assay system is depicted in Figure 1A. Notably, cells ectopically expressing either wild-type (WT) or activated mutant FGFR3, together with adaptor/GFP proteins that selectively bind to activated FGFR3, are expected to exhibit intracellular punctate fluorescent signals. These signals indicate the presence of internalized vesicles containing activated FGFR3 and adaptor/GFP complexes. By tracking changes in the subcellular localization of an FGFR3-specific interacting adaptor/GFP, we aimed to identify natural compounds from plant extracts that can inhibit FGFR3 at an early activation stage.

SH2 (SH2Bβ)/GFP specifically interacts with activated FGFR3 to generate an internalized cytoplasmic spot pattern in U2OS cells. (A) Schematic illustration shows the assay design concept by tracking the subcellular localization of an adaptor/GFP fusion protein that specifically interacts and internalizes with activated FGFR3. (B) Schematic representation shows the SH2 domain (underlined) of SH2Bβ and PLCγ proteins fused to the N-terminus of GFP. (C) U2OS cells transiently expressing GFP, SH2(SH2B)/GFP, or SH2(PLCγ)/GFP were observed by confocal live imaging. Scale bars, 10 μm. (D) Schematic representation shows the relative positions of different mutations in FGFR3 known to cause skeletal dysplasias. TDI, thanatophoric dysplasia 1; Ig I, II, and III; immunoglobulin-like domains; TM, transmembrane I domain; TK1 and 2, tyrosine kinase domains. (E) U2OS cells stably coexpress SH2 (SH2B)/GFP and various FGFR3s (WT, ACH, and TDI) or empty vector control (mock). The levels of FGFR3 and SH2 (SH2B)/GFP fusion proteins were detected by immunoblotting. (F) SH2 (SH2B)/GFP fluorescence signals (green) in the indicated cell lines were observed by confocal live imaging. Scale bars, 10 μm. (G) The direct interaction of SH2(SH2B)/GFP and FGFR3 was detected by co-immunoprecipitation (IP) using FGFR3 antibody followed by immunoblotting for FGFR3 and GFP. (H–J) Inhibition of FGFR3 activation by mutated Y724 and Y760 phosphorylation sites in TDI FGFR3 abolished the internalized spot pattern in U2OS cells coexpressing SH2(SH2B)/GFP cells TDI FGFR3. (H) Schematic representation shows the substitution of phenylalanine for tyrosine at Y724F and Y760F in TDI-FGFR3. (I) U2OS-SH2(SH2B)/GFP cells stably expressing empty vector control (mock), TDI-FGFR3, or Y724F/Y760F TDI-FGFR3 were observed by confocal live imaging. Scale bars, 10 μm. (J) The expression of FGFR3 was detected by immunoblotting.

To establish the system, our first objective was to identify an adaptor protein that specifically interacts with activated FGFR3. We elected to use U2OS cells as a model to identify the desired adaptor protein, as we found this human osteosarcoma cell line endogenously expressed FGFR1, 2, and 4 but had no detectable FGFR3 expression (Supplemental Figure 1; supplemental material available online with this article; https://doi.org/10.1172/jci.insight.171257DS1). To ensure minimal interference from downstream FGFR3 signaling on cell proliferation or viability, we utilized only the binding domain of the adaptor protein rather than the full-length sequence. We investigated the Src homology 2 (SH2) domain of 2 known adaptor proteins that interact with FGFRs or FGFR3 in U2OS cells. The SH2 domain of PLCγ binds to tyrosine-autophosphorylated regions of FGFRs (18), while the SH2 domain of SH2Bβ was identified through a yeast 2-hybrid screen for its interaction with activated kinase domains of FGFR3 (19). To visually examine the intracellular localization and interactions of these SH2 domains with endogenous proteins in U2OS cells, we fused them to GFP (Figure 1B). When U2OS cells expressed SH2(PLCγ)/GFP, we observed punctate signals at the plasma membrane, indicating its interaction with endogenous plasma membrane proteins. In contrast, U2OS cells expressing SH2(SH2B)/GFP exhibited a diffuse fluorescent signal throughout the cytoplasm and nucleus, accompanied by few condensed fluorescent signals in nucleus (Figure 1C). Of note, SH2Bβ is known to undergo nucleocytoplasmic shuttling and has functions in nucleus (20). In the absence of an interaction between SH2(SH2B) and endogenously expressed membrane proteins in U2OS cells, the SH2(SH2B) may interact with proteins located in nucleus, leading to the observed condensed signals within the nucleus. These findings demonstrate that SH2(SH2B)/GFP does not interact with endogenously expressed plasma membrane proteins in U2OS cells, including FGFR1, 2, and 4.

To explore the interaction between FGFR3 and SH2(SH2B)/GFP in U2OS cells, we generated stable cell lines expressing SH2(SH2B)/GFP along with WT or activating mutation forms of FGFR3 (Figure 1D). Western blot analysis verified the expression of SH2(SH2B)/GFP and WT or mutant FGFR3 in U2OS cells (Figure 1E). Notably, U2OS cells coexpressing both SH2(SH2B)/GFP and different forms of FGFR3 (WT, ACH, and TDI) displayed distinct intracellular punctate GFP signals (Figure 1F). These cells also displayed fewer nuclear condensed signals than control cells (mock) that only expressed SH2(SH2B)/GFP (Figure 1F). Additionally, weak plasma membrane signals were observed in U2OS cells coexpressing SH2(SH2B)/GFP and the different forms of FGFR3 (WT, ACH, and TDI) (Figure 1F). These intracellular punctate signals suggest that the recruitment of SH2(SH2B)/GFP with activated FGFR3 (WT, ACH, and TDI) lead to internalization and formation of intracellular puncta. Furthermore, the direct interaction between FGFR3 and SH2(SH2B)/GFP was validated through co-immunoprecipitation of SH2(SH2B)/GFP with FGFR3 variants (Figure 1G).

To directly investigate the relationship between FGFR3 activation and intracellular punctate GFP signal, we examined the impact of inhibiting FGFR3 activity. We inactivated a constitutively active TDI-FGFR3 by substituting 2 critical tyrosine (Y) phosphorylation sites with phenylalanine (F) residues, Y724F and Y760F (Figure 1H). These 2 phosphorylation sites are well established as vital for FGFR3 activation and its interaction with the SH2 domain of SH2B (19). Strikingly, the intracellular punctate GFP signals were completely abolished in U2OS cells coexpressing SH2(SH2B)/GFP and Y724F/Y760F TDI-FGFR3 (Figure 1I), while the expression levels of both FGFR3 isoforms were similar between the groups (Figure 1J).

These findings highlight the successful establishment of a cell-based receptor/adaptor translocation system for specifically monitoring FGFR3 activity. By utilizing SH2(SH2B)/GFP, we demonstrated the direct interaction between SH2(SH2B)/GFP and activated FGFR3, leading to intracellular punctate GFP signals in U2OS cells.

High-throughput imaging assay to quantify FGFR3 activity. Our next goal was to establish a high-throughput imaging method to quantify FGFR3 activity for drug screening. The TDI Y373C substitution in the extracellular region of FGFR3 (TDI-FGFR3) results in the formation of intermolecular disulfide bonds that cause receptor dimerization, constitutive kinase activation, and adaptor protein recruitment (2). Since U2OS cells expressing SH2(SH2B)/GFP and TDI-FGFR3 (U2OS-TDI/SH2GFP) exhibit robust intracellular GFP puncta, we used this cell line to establish high-throughput imaging system (Figure 1). With this system, we are able to screen for hits that inhibit early events of FGFR3 activation, including disrupting receptor dimerization, kinase activation, and adaptor protein recruitment.

To optimize the protocol and parameter settings for the high-throughput imaging system, we used PKC412, a small-molecule multiple tyrosine kinase inhibitor, which has been shown to inhibit FGFR3 tyrosine phosphorylation (21). Indeed, treating U2OS-TDI/SH2GFP cells with PKC412 for 1 hour led to a marked reduction in intracellular punctate GFP signals (Figure 2A). To automatically quantify the punctate GFP signals in cytoplasm, we employed nuclei staining using Hoechst to identify individual cells, and cell images were automatically obtained via the ArrayScan VTI HCS Reader. For each cell, the nucleus (circle) and cytoplasmic area (ring) were defined using the parameter settings in the Compartmental Analysis BioApplication software (Figure 2B). The GFP spot number (ring spot count) and spot intensity (ring spot intensity) in ring area were determined.

High-throughput imaging system (receptor/adaptor translocation assay) to quantify FGFR3 activation and identify hits that inhibit FGFR3 activation. (A and B) U2OS-TDI/SH2BGFP cells were treated with 4 μM PKC412 or vehicle control (DMSO) for 1 hour. Scale bars, 20 μm. (A) Confocal images show SH2(SH2B)/GFP signals (green). (B) Representative raw images acquired on the ArrayScan VTI HCS Reader (left panels) and automated identification of spots (yellow dots) within cytoplasmic area (green ring) (right panels). (C and D) Dose-response curves of PKC412 plotted by ring spot count (C) or ring spot intensity (D) relative to vehicle control. (E) FGFR3 activity was quantified based on ring spot counts in U2OS-SH2(SH2B)/GFP cells expressing TDI FGFR3, Y724F/760F TDI FGFR3, or vector control (Mock). (F) U2OS-TDI/SH2BGFP cells were treated with different plant extracts. Ring spot count per cell was plotted relative to vehicle control. Data from 3 independent treatments are shown. Hits: red circles; PKC412: black circles. (G) Linear regression analysis of experiments 1 and 2 showing PKC412 treatments and hits (lower-left corner) and active responder (upper-right corner). (H–J) Dose-response curves of (H) 4 identified hits (Hits 1–4), (I) 2 different batches of Hit 4 (Hit 4 B1 and Hit 4 B2), and (J) 2 additional plant extracts from species closely related to Hit 4: Hit 4-1 (A. viridis) and Hit 4-2 (A. tricolor). Data in C–E and H–J represent the mean ± SEM of triplicates. (K) KMS-11 cells were treated with vehicle control or different doses of plant extracts for 48 hours. Cell viability was analyzed using WST-1 assay and normalized to the vehicle control. Data represent the mean ± SEM of 3 independent experiments. Student’s 2-tailed t tests were performed. *P < 0.05, ***P < 0.001. (L) KMS-11 cells were treated with indicated concentrations of plant extracts, PKC412, or vehicle control (Vehicle) for 1 hour. Total and phosphorylated protein levels of FGFR3 and its downstream effectors were detected by immunoblotting.

To evaluate the principal quantifiable features reflecting the underlying biology, we conducted a dose-response assay with PKC412 and plotted the ring spot counts and ring spot intensity per cell as percentage of DMSO control (Figure 2, C and D). Both parameters reliably showed a dose-dependent response to the inhibitor, indicating their sensitivity to FGFR3 activation. However, considering the potential influence of autofluorescence or fluorescence-quenching compounds present in plant extracts on the ring spot intensity, we decided to use the ring spot count per cell as the preferred method for routine quantification of FGFR3 activation.

To verify the capability of the system to quantify FGFR3 inactivation, we used it to compare U2OS-TDI/SH2GFP cells with U2OS-Y724F/760F TDI/SH2GFP cells. The dramatic reduction in the number of intracellular spot counts in U2OS-Y724F/Y760F TDI/SH2GFP cells (Figure 2E) supported the notion that the counts of intracellular spots directly reflect the degree of FGFR3 activation. Therefore, we utilized this high-throughput FGFR3-TDI/SH2GFP imaging system, which we termed R/ATA (receptor/adaptor translocation assay), for subsequent experiments.

Identification of Amaranthus viridis extract as an inhibitor of FGFR3 activation. The R/ATA system was used to screen a library of plant extracts from various species collected in Taiwan, aiming to identify hits that could effectively reduce FGFR3 activation. To identify hits targeting the early events of FGFR3 activation, we treated cells with the plant extracts for 1 hour prior to analysis. A hit was defined as a treatment that exhibited more than 40% inhibition of FGFR3 activity in 3 independent experiments. This criterion was established based on the observation that the control inhibitor (PKC412) suppressed at least 40% of FGFR3 activity in our experiments. Using the 40% inhibition criterion, we identified 4 plant extracts as hits from the initial screen (Figure 2F). The inhibitory activities of these extracts were further verified by applying linear regression analysis to 2 independent experiments, referred to as experiments 1 and 2. The hits were clearly distinguished from nonresponders and active responders, aligning with PKC412-positive controls in the lower left corner of the resulting plots (Figure 2G). Subsequently, the 4 hits were found to exhibit dose-dependent reductions in FGFR3 activities (Figure 2H). Hit 4 is the ethanol extract from Amaranthus spinosus, a widely available food plant in Taiwan and tropical countries. Furthermore, the ethanol extract of Amaranthus mangostanus has been demonstrated to prevent ovariectomy-induced bone loss in mice (22). Thus, it was selected for further validation and analysis. To assess batch-to-batch consistency of hit 4, we prepared 2 different batches of ethanol extract (hit 4 B1 and hit 4 B2) from plant collected from different locations in Taiwan during different seasons. Both batches displayed inhibitory properties against FGFR3 (Figure 2I). Furthermore, we tested ethanol extracts from 2 plant species closely related to hit 4, A. viridis (hit 4-1) and A. tricolor (hit 4-2), which also showed dose-dependent reductions in FGFR3 activation (Figure 2J). These results demonstrate that the R/ATA system could successfully identify hits, and the methods employed for plant extracts preparation were reliable. The consistent inhibitory activity of hit 4 across different batches and related plant species further supports its potential as a reliable and reproducible candidate for further investigation.

The functional effects of hit 4–mediated inhibition of FGFR3 activity were further assessed in a clinically relevant FGFR3-overactivated human MM cell line, KMS-11 cells, which ectopically expressed FGFR3Y373C. The viability of KMS-11 cells is dependent on FGFR3 activation (23). Consistent with this, the viability of KMS-11 cells was reduced following treatment with hit 4 and extracts of a closely related species (hit 4-1 and hit 4-2), whereas the extract from a nonresponder did not affect cell viability (Figure 2K). Of note, the reduction in KMS-11 cell viability correlated with decreased FGFR3 accumulation and downregulation of FGFR3 downstream signaling pathway components, including phosphorylated PLCγ (24), PI3K (25), and ERK1/2 (26), observed at 1 hour posttreatment (Figure 2L). These results validate the bioactivity of hit 4, which includes both hit 4-1 and hit 4-2, as it effectively inhibits cell viability and FGFR3 early activation in FGFR3-activated MM cells. A. viridis (hit 4-1), which is widely cultivated in Taiwan, was used for further bioactive compound discovery.

Identification of bioactive fractions and compounds from A. viridis extracts. Standard bioassay-guided sequential fractionation (27) was used to identify the most potent fractions from the ethanol extract of A. viridis (hit 4-1). During the process of discovering bioactive constituents, the most potent active fractions without cellular toxicity (not affecting the cell/nucleus morphology or cell number) were validated as exerting dose-dependent effects and subjected to further fractionation and purification (Supplemental Figures 2–4). The initial fractionation of A. viridis is presented in Supplemental Figure 2A. Among the 5 initial fractionations, F3, F4, and F5 exhibited inhibitory activities toward FGFR3 activation, with F4 showing the highest potency (Supplemental Figure 2B). We also demonstrated the batch-to-batch preparation consistency of F4 from 2 different batches of A. viridis (hit 4-1 B1-F4 and hit 4-1 B2-F4) in inhibiting FGFR3 activity (Supplemental Figure 2C). Overall, the R/ATA system enabled the consistent identification of active fractions from this complex plant extract and validated their bioactivities.

We then sequentially identified the most potent active fraction (Supplemental Figures 2–4) and finally identified 2 active molecules, pheophorbide a (Pa) and pyropheophorbide a (PyroPa) (Figure 3, A and B), present at high purity in the 2 final potent fractions (as demonstrated using HPLC and MS analysis) (Supplemental Figure 4, A–E). In summary, our approach allowed us to identify and characterize 2 active compounds, Pa and PyroPa, from A. viridis.

Identification of Pa from the active fraction of A. viridis as a specific inhibitor of FGFR3 activity. (A) The structures of Pa and PyroPa are shown. (B) The dose-response curves indicate that Pa and PyroPa inhibit FGFR3 activity using R/ATA system. Data are shown as mean ± SEM of triplicates. (C) The effects of Pa on cell viability in various cancer cell lines. Cells were treated with indicated concentrations of Pa or PyroPa for 48 hours. Cell viability was determined by WST-1 assay. Data are presented as mean ± SEM of 3 to 4 independent experiments. (D) The effects of Pa on protein levels of the FGFR family members were examined in U2OS-SH2BGFP cells or U2OS-TDI-FGFR3/SH2BGFP cells. Cells were treated with indicated concentration of Pa or vehicle control (V) for 1 hour, and protein levels were analyzed by immunoblotting. (E) Relative protein levels for each FGFR family member were quantified and compared with vehicle-treated controls. Data are presented as mean ± SEM of 3 independent experiments. *P < 0.05.

We then validated the effects of Pa and PyroPa on inhibition of FGFR3 activity in KMS-11 cells. Both Pa and PyroPa inhibited the viability of KMS-11 cells (Figure 3C). Since Pa and PyroPa are both porphyrins, we further tested a few other porphyrins (Supplemental Figure 5A) and demonstrated that temoporfin and phyrophenophobide a methyl ester (PyroPa methyl ester) could also inhibit FGFR3 activation in the R/ATA system and reduce viability of KMS-11 cells (Supplemental Figure 5, B and C). However, chlorophyllin could only reduce FGFR3 activation and viability of KMS-11 cells at high concentration (Supplemental Figure 5, B and C). Note that the precursor of porphyrins, 5-aminolevulinic acid, did not inhibit FGFR3 activity, and it did not affect viability of KMS-11 cells (Supplemental Figure 5, B and C). We concluded that the shared porphyrin structure in identified active compounds Pa and PyroPa is important for inhibition of FGFR3 activation in the screening system and reduction of cell viability of KMS-11 cells.

To further investigate the specific inhibitory effect of Pa on FGFR3-dependent cancer cell viability, we evaluated Pa impacts in cancer cell lines for which growth is independent of FGFR3 activity. The cell lines included U2OS, NCI-H441, and NCI-H1975 cells; no endogenous FGFR3 expression was detected in any of these lines (RNA HPA cell line gene data, The Human Protein Atlas, https://www.proteinatlas.org/about/download). NCI-H441is a KRAS-dependent lung adenocarcinoma cancer cell line (28), and NCI-H1975 is a non–small cell lung cancer cell line harboring EGFR L858R/T790M activating mutations (29). In contrast with the dose-dependent inhibitory effect of Pa on cell viability in KMS-11 cells, Pa did not have a dose-dependent inhibitory effect on cell viability in U2OS, NCI-H441, or NCI-H1975 cells (Figure 3C). Only at high doses of Pa did we observe a partial reduction in cell viability in NCI-H441 and NCI-H1975 cells (Figure 3C). These results suggest that Pa exhibits greater potency in inhibiting viability of cancer cells dependent on ectopic expression of FGFR3 compared with cells harboring mutations in RAS family or other receptor tyrosine kinases, such as EGFR.

We next examined the selectivity of Pa for members of the FGFR family. U2OS-SH2B-GFP cells endogenously express FGFR1, 2, and 4. Therefore, we treated U2OS-SH2B-GFP cells without or with stable expression of an FGFR3 activation mutant (U2OS-FGFR3-TDI) with vehicle or Pa and determined protein levels of FGFRs by Western blot analysis. In Pa-treated cells, the FGFR3 protein level was dramatically reduced; however, Pa had no significant inhibitory effect on other FGFR family members, as shown in the quantitative results (Figure 3, D and E). Thus, we concluded that Pa has the highest potency to reduce FGFR3 accumulation.

Up to this point in the study, we had identified active compounds, Pa and PyroPa, from A. viridis and showed that the shared porphyrin structure is important for inhibition of FGFR3 activity. Furthermore, we found that Pa has the highest potency toward FGFR3 among FGFR family members.

Pa-mediated inhibition of FGFR3 signaling and its downstream effects in FGFR3-overactivated MM cells. In our next experiments, we evaluated Pa-induced inhibition of FGFR3 downstream signaling, which regulates diverse biological activities. It is known that ectopic FGFR3 expression in MM cells, such as KMS-11, induces cell proliferation and blocks apoptosis through ERK signaling and the PI3K/AKT signaling pathway (25, 30–33). Accordingly, we showed that treatment of KMS-11 cells with Pa dramatically reduced FGFR3 protein accumulation and inhibited FGFR3 activation–induced phosphorylation of downstream effectors, including PLCγ1, AKT, and ERK (Figure 4, A–F). Most importantly, Pa inhibited proliferation and antiapoptosis processes that are promoted by FGFR3 activation, as demonstrated by a BrdU incorporation assay and FITC-annexin V/propidium iodide (PI) apoptosis assay (Figure 4, G–I). Taken together, these findings revealed that Pa can effectively reduce FGFR3 accumulation and FGFR3 downstream signaling pathways, which leads to suppression of cell proliferation and induction of apoptosis in MM cells.

Pa reduces FGFR3 signaling, decreases cell proliferation, and induces cell apoptosis processes in MM cells. (A) Pa treatment reduced FGFR3 accumulation and its downstream signaling in KMS-11 cells. KMS-11 cells were treated with vehicle or indicated concentration of Pa for 1 hour. The levels of total and phosphorylated FGFR3 and its downstream effectors were analyzed by immunoblotting. (B–F) The relative levels of phosphorylated FGFR3 and downstream effectors were normalized to the corresponding total protein levels and compared with vehicle-treated controls. Data are shown as mean ± SEM of 3 independent experiments. (G) Pa treatment reduced KMS-11 cell proliferation. KMS-11 cells were treated with vehicle or Pa for the indicated times, and cell proliferation was analyzed by 5′-bromo-2′-deoxyuridine (BrdU) incorporation assay. Data are shown as mean ± SEM of 4 independent experiments. (H and I) KMS-11 cells were treated with vehicle or Pa for 8 hours and stained by annexin V and PI; the proportion of apoptotic cells was analyzed by flow cytometry. (H) Represented data are shown. (I) The percentage of apoptotic cells from 3 independent experiments. Data are shown as mean ± SEM. (B–F) Statistical significance was determined by 1-way ANOVA with Tukey’s multiple-comparison test or (G and I) 2-tailed Student’s t test. *P < 0.05, **P < 0.01.

Pa-induced reduction of FGFR3 half-life in FGFR3-activated MM cells and chondrocytes. We then elucidated the mechanism by which Pa inhibits FGFR3 activity. We observed a reduction in FGFR3 protein levels when KMS-11 cells were treated with Pa for 1 hour (Figure 4, A and C). Activating mutations of FGFR3 have been shown to increase its stability and prolong its half-life (34). Thus, we hypothesized that Pa might reduce FGFR3 accumulation by regulating protein stability. To test this possibility, we determined the half-life of FGFR3 in KMS-11 cells treated with vehicle or Pa in the presence of protein synthesis inhibitor cycloheximide (CHX). The half-life of FGFR3 in Pa-treated KMS-11 cells was shorter than that in vehicle-treated controls (Figure 5, A and B), suggesting that Pa compromises FGFR3 stability in FGFR3-activated MM cells.

Pa compromises FGFR3 protein stability in FGFR3-activated MM cells and chondrocytes. (A and B) KMS-11 cells were treated with either vehicle or 20 μM Pa in the presence of 80 μg/mL cycloheximide (CHX) for the indicated times. (A) Protein levels were detected by immunoblotting. (B) The relative levels of FGFR3 protein were quantified, normalized to β-actin, and compared with time 0 of CHX treatment. (C–F) WT and ACH-FGFR3–expressing ATDC5 cells were treated with vehicle or 10 μM Pa in the presence of 100 μg/mL CHX for indicated times. (C and D) Protein levels were detected by immunoblotting. (E and F) The relative protein levels were quantified, normalized to GAPDH, and compared with time 0 of CHX treatment. (B, E, and F) Data are shown as mean ± SEM of 3 to 4 independent experiments.

To begin addressing the possibility that Pa might be able to functionally alter ACH signs and symptoms, we first tested whether Pa can also compromise FGFR3 stability in chondrocytes. For this purpose, we established ATDC5 cells, a chondrocyte cell line, that stably expresses WT FGFR3 or the G380R mutant (ACH) of FGFR3. Similar to our results in MM cells, Pa reduced the protein stability of both WT and ACH FGFR3 in ATDC5 cells (Figure 5, C–F). To further investigate the impact of Pa on the stability of FGFR3 in a clinically relevant system, we conducted experiments in chondrocytes isolated from FGFR3ACH mice. Pa significantly reduced the half-life of FGFR3 in chondrocytes from FGFR3ACH mice, as compared with vehicle-treated controls (Supplemental Figure 6). These results demonstrated that Pa reduces FGFR3 stability in chondrocytes. Thus, our data reveal that Pa can reduce FGFR3 stability in both MM cells and chondrocytes.

To explore the molecular mechanisms by which Pa reduces FGFR3 stability, we examined a known mechanism regulating FGFR3 stability. Activating mutations have been shown to disrupt FGFR3 protein ubiquitination, which acts as a signal for targeted protein degradation (34, 35). We therefore investigated whether Pa could increase FGFR3 ubiquitination to induce protein degradation. Unexpectedly, less ubiquitination on FGFR3 was observed in KMS-11 cells treated with Pa (Supplemental Figure 7A). We then tested whether Pa could affect ubiquitin-independent macroautophagy protein degradation by examining the conversion of an autophagosomal marker, the cytosolic form of microtubule-associated protein 1A/1B-light chain 3 (LC3-I), to its lipidated form, LC3-II (36). No difference in LC3 conversion was observed in KMS-11 cells treated with or without Pa, according to Western blotting (Supplemental Figure 7B). Thus, it is unlikely that Pa reduces FGFR3 stability by increasing FGFR3 ubiquitination or acting on the macroautophagy protein degradation pathway.

We demonstrated that Pa reduced FGFR3 protein abundance by shortening its half-life, which subsequently attenuated FGFR3 downstream signaling in both cancer cells and chondrocytes. In some cases, drugs that perturb specific signaling targets can induce positive or negative feedback loops that may alter the expression of the target and compromise long-term drug efficacy (37). As such, we probed the effects of Pa on FGFR3 gene expression. We found that after treatment with Pa, the mRNA levels of FGFR3 remained relatively low in both FGFR3-activated MM cells and chondrocytes (Supplemental Figure 8, A–C). These results suggest that Pa is unlikely to induce positive feedback leading to an increase in gene transcription.

Pa-induced rescue of defective growth in cultured femurs from FGFR3ACH mice. To further assess the therapeutic potential of Pa for ACH, we determined how Pa treatment affects FGFR3 downstream signaling in chondrocytes using ATDC5 cells expressing WT or ACH FGFR3. In ACH pathogenesis, FGFR3 activation suppresses chondrocyte proliferation and differentiation in the growth plate, thereby limiting long bone growth (38). It is known that FGFR3 activation inhibits chondrocyte proliferation via activation of STAT1 (39), and it inhibits chondrocyte differentiation through activation of MAPK/ERK signaling (40, 41). Meanwhile, FGFR3 activation negatively modulates chondrocyte apoptosis through activation of PI3K/AKT signaling (42). Correspondingly, ACH FGFR3 expression in ATDC5 cells increased FGFR3 accumulation and activation, which induced the phosphorylation of STAT1, ERK1/2, PI3K, and AKT (Figure 6, A–G). Treatment of the cells with Pa suppressed FGFR3 accumulation and phosphorylation, along with FGFR3 activation–induced phosphorylation of STAT1, PI3K, and AKT in both WT and ACH FGFR3–expressing ATDC5 cells (Figure 6, A–F). However, Pa induced ERK1/2 phosphorylation in WT FGFR3–expressing cells (Figure 6, A and G). Overall, these results suggest that Pa can partially reverse FGFR3 activation–induced signaling events that are known to impact chondrocyte proliferation, differentiation, and apoptosis.

Pa counteracts overactive FGFR3 signaling in chondrocytes and rescues defective growth of cultured femurs from FGFR3ACH mice. (A) ATDC5 cells stably expressing either WT or ACH FGFR3 were treated with vehicle or the indicated concentrations of Pa in the presence of FGF2 (20 ng/mL) for 2 hours. Levels of total and phosphorylated FGFR3 and downstream effectors were examined by immunoblotting. (B–G) The relative phosphorylated protein levels were normalized to the corresponding total protein levels and compared with vehicle-treated controls. Data are shown as mean ± SEM of 3 to 4 independent experiments and were analyzed by 1-way ANOVA with Tukey’s multiple comparison; *P < 0.05, **P < 0.01, ***P < 0.001. (H–K) Femurs from WT, FGFR3ACH/+, and FGFR3ACH/ACH mice at E16.5 were isolated and cultured for 6 days in the presence of vehicle or indicated concentrations of Pa. (H) Representative images of E16.5 femurs before and after 6 days of culture are shown. Scale bar: 1 mm. (I) The increased femur lengths after treatment were calculated. Data are expressed as mean ± SD (n = 9–35) and analyzed by 1-way ANOVA. *P < 0.05, ***P < 0.001; ****P < 0.0001. (J) Representative images of H&E-stained and collagen X–stained femurs cultured for 6 days. Scale bar: 200 μm. (K) Quantification of the collagen X–stained area in the femurs. Data are shown as mean ± SEM; n = 3–8. Statistical significance was determined by Student’s 2-tailed t test. *P < 0.05 and **P < 0.01.

We therefore directly evaluated the therapeutic potential of Pa on defective long bone growth using ex vivo femur cultures of WT and FGFR3ACH mice (17). Mouse femurs were isolated at embryonic day 16.5 (E16.5) and cultured in the presence or absence of Pa for 6 days. Pa markedly increased the femur length gains in WT, FGFR3ACH/+, and FGFR3ACH/ACH mice. Importantly, the Pa-increased femur length gains in FGFR3ACH/ACH tissues occurred in a dose-dependent manner (Figure 6, H and I). Most significantly, we noticed that Pa increased the area of hypertrophic chondrocytes in the femurs, according to histological analysis of H&E-stained tissues. Hypertrophic chondrocytes are considered to be in a terminal state of chondrocyte differentiation and are essential for long bone growth; the cells exhibit characteristic high levels of type X collagen expression (14, 40). We therefore immunostained hypertrophic chondrocytes in the femurs for type X collagen. Pa treatment significantly increased the area of the hypertrophic zone in femurs across all genotypes, as compared with vehicle-treated controls (Figure 6, J and K). These results suggest that Pa enhances chondrocyte differentiation and that Pa can reverse FGFR3 activation signaling in chondrocytes to promote long bone growth in femur cultures.