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Dual Allosteric Inhibition of SHP2 Phosphatase Michelle Fodor, Edmund Price, Ping Wang, Hengyu Lu, Andreea Argintaru, Zhouliang Chen, Meir Glick, Huai-Xiang Hao, Mitsunori Kato, Robert Koenig, Jonathan R LaRochelle, Gang Liu, Eric McNeill, Dyuti Majumdar, Gisele A. Nishiguchi, Lawrence B. Perez, Gregory Paris, Christopher M Quinn, Timothy Ramsey, Martin Sendzik, Michael David Shultz, Sarah L. Williams, Travis Stams, Stephen C. Blacklow, Michael G Acker, and Matthew J. LaMarche ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/acschembio.7b00980 • Publication Date (Web): 05 Jan 2018 Downloaded from http://pubs.acs.org on January 6, 2018

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Allosteric site 1

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Allosteric site 2

SHP099 (1)

SHP244 (2)

SHP2 Phosphatase with Allosteric Inhibitors SHP099 (1), SHP244 (2).


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Dual Allosteric Inhibition of SHP2 Phosphatase Michelle Fodor,† Edmund Price,† Ping Wang,† Hengyu Lu,† Andreea Argintaru,† Zhouliang Chen,† Meir Glick,† Huai–Xiang Hao,† Mitsunori Kato,† Robert Koenig,† Jonathan R. LaRochelle, ‡Gang Liu,† Eric McNeill,† Dyuti Majumdar,† Gisele A. Nishiguchi,† Lawrence B. Perez,† Gregory Paris,† Christopher M. Quinn,† Timothy Ramsey,† Martin Sendzik,† Michael David Shultz,† Sarah L. Williams,† Travis Stams,† Stephen C. Blacklow,‡ Michael G. Acker,†* and Matthew J. LaMarche†* †

Novartis Institutes for Biomedical Research, Cambridge, MA 02139, United States.

‡

Department of Biological Chemistry & Molecular Pharmacology, Harvard Medical School and Department of Cancer Biology, Dana–Farber Cancer Institute, Boston, Massachusetts 02215, United States. KEYWORDS: SHP2, PTPN11, protein tyrosine phosphatase, phosphatase, allosteric inhibitor, structure activity relationship, cancer, immuno–oncology, MAP kinase pathway, SHP099, dual inhibition. ABSTRACT SHP2 is a cytoplasmic protein tyrosine phosphatase encoded by the PTPN11 gene and is involved in cell proliferation, differentiation, and survival. Recently we reported an allosteric mechanism of inhibition that stabilizes the auto–inhibited conformation of SHP2. SHP099 (1) was identified and characterized as a moderately potent, orally bioavailable, allosteric small molecule inhibitor, which binds to a tunnel–like pocket formed by the confluence of three domains of SHP2. In this report, we describe further screening strategies that enabled the identification of a second, distinct small molecule allosteric site. SHP244 (2) was identified as a


1


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weak inhibitor of SHP2 with modest thermal stabilization of the enzyme. X–ray crystallography revealed that 2 binds and stabilizes the inactive, closed conformation of SHP2, at a distinct, previously unexplored binding site– a cleft formed at the interface of the N–terminal SH2 and PTP domains. Derivatization of 2 using structure–based design resulted in an increase in SHP2 thermal stabilization, biochemical inhibition, and subsequent MAPK pathway modulation. Downregulation of DUSP6 mRNA, a downstream MAPK pathway marker, was observed in KYSE–520 cancer cells. Remarkably, simultaneous occupation of both allosteric sites by 1 and 2 was possible, as characterized by cooperative biochemical inhibition experiments and X–ray crystallography. Combining an allosteric site 1 inhibitor with an allosteric site 2 inhibitor led to enhanced pharmacological pathway inhibition in cells. This work illustrates a rare example of dual allosteric targeted protein inhibition, demonstrates screening methodology and tactics to identify allosteric inhibitors, and enables further interrogation of SHP2 in cancer and related pathologies.

Allosteric site 1

Allosteric site 2

Cl Cl

NH2 N

N

N NH2

SHP099 (1)

SHP244 (2)

SHP2 Phosphatase with Allosteric Inhibitors SHP099 (1), SHP244 (2).


2

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INTRODUCTION SHP2 is a non–receptor protein tyrosine phosphatase encoded by the PTPN11 gene, and contains two N–terminal Src homology 2 (SH2) domains, a protein tyrosine phosphatase (PTP) domain, and a poorly ordered C–terminus. X–ray crystallographic studies have shown that SHP2 inhibits its own phosphatase activity by blocking access to the catalytic site on the PTP domain using the N–terminal SH2 domain.1,2

Others have previously demonstrated3 that bis–

phosphotyrosyl proteins or peptides (e.g., IRS–1) bind to the SH2 domains of SHP2, disrupting the N–terminal SH2–PTP domain interaction. This binding event allows substrate access to the catalytic site and activates the phosphatase. Germline or somatic mutations in PTPN11 that cause hyperactivation4 of SHP2 have been identified in a number of pathophysiologic states: in the developmental disorder Noonan syndrome (50%),5 in hematological malignancies including juvenile myelomonocytic leukemia (35%), myelodysplastic syndrome (10%), B–cell acute lymphoblastic leukemia (7%), acute myeloid leukemia (4%),6 and in solid tumors at low frequency.7,8 Interestingly, these activating mutations frequently occur at the interface of the N–terminal SH2 domain and PTP domain, likely weakening this quaternary protein interaction and activating the phosphatase.9 SHP2 is located in the cytoplasm and transduces cell signaling from a variety of receptor– tyrosine kinases, and thus is involved in numerous oncogenic signaling cascades (e.g., RAS– ERK, PI3K–AKT, JAK–STAT).

However, its role in these pathways is not yet fully

understood.10,11 In 2015, SHP2 was reported to bind and dephosphorylate RAS, increase RAS– RAF association, and activate downstream proliferative RAS/ERK/MAPK signaling.12 SHP2 also has been reported to participate in the T cell programed cell death/checkpoint pathway (PD– L1/PD–1) and contribute to immune evasion.13 Activated PD–1 recruits SHP2 to the membrane where it dephosphorylates costimulatory receptor CD28 and suppresses T–cell function.14


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Investigating the inhibition of SHP2 with small molecule modalities for cancer immunotherapy is of great interest given recent clinical success of anti–PD–1 and PD–L1 therapeutics.15 Since the initial discovery of SHP2 as a signal transducer and potential anticancer target,16 the identification of pharmacologically relevant inhibitors has attracted broad interest in the scientific community.17

Many groups have reported active site inhibitors, but these

compounds typically suffer from low potency, modest selectivity, and poor pharmaceutical properties.

These limitations are largely due to the highly conserved, polar, and charged

environment of the phosphatase active site, which greatly complicates drug discovery and has slowed the advancement of the phosphatase field.18,19,20,21 Alternatively, allosteric modes of phosphatase inhibition have been reported (e.g., targeting WIP1),22,23,24 but these efforts have yet to progress viable chemical matter to the clinic. Recently, we reported a novel allosteric mechanism of phosphatase inhibition, whereby small molecules bind and stabilize the inactive conformation of SHP2.25,26 SHP099 (1) was identified and characterized (Figure 1A) as a selective and moderately potent SHP2 inhibitor in biochemical experiments (IC50 = 0.070 µM) and in an EGFR–amplified esophageal squamous cell carcinoma model, KYSE–520 (phospho–ERK IC50 = 0.250 µM; antiproliferation EC50 = 2.5 µM).

High aqueous solubility, selectivity, and oral bioavailability enabled in vivo

characterization of 1, which proved efficacious in RTK–dependent cancer xenograft models. An engineered, double mutation in the allosteric pocket (SHP2T253M/Q257L) disrupted the binding of 1 (Figure 1B) and rescued pathway inhibition in cancer cells, indicating that the cellular activity of 1 was attributable to SHP2 inhibition.25,26 Further medicinal chemistry optimization of the aminopyrazine chemical series will be reported in due course. Here we report the discovery and


4

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characterization of a second SHP2 allosteric pocket through predictive modeling, and selective screening of the engineered SHP099 binding–deficient mutant, SHP2T253M/Q257L. A.

SHP099 (1) 1–525 SHP2 IC50 = 0.070 µM T253M/Q257L SHP2 IC50 > 10 µM KYSE–520 p–ERK IC50 = 0.250 µM DSF (∆Tm) = 7.5 °C

B.

T253M

Q257 L Figure 1. A. Structure, biochemical and cellular activity, and thermal stabilization of SHP099 (1). B. Model of engineered double mutant (SHP2T253M/Q257L) which disrupts the binding of 1. SHP2WT colored in wheat, SHP2T253/Q257 sidechains colored in magenta.


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RESULTS and DISCUSSION While SHP2T253M/Q257L was successfully used for cellular characterization of 1, we also considered the double mutant as a unique tool to discover further small molecule modalities. Analysis of ligandable pockets of SHP2 using SiteMap27,28 in Maestro29 revealed three potential small molecule binding sites, including the allosteric “tunnel” previously reported for 1 (Figure 2A).28 Additional allosteric binding sites were predicted on both sides of the N–SH2/PTP domain interface: the “latch” binding site, located approximately 20 Å from the tunnel (Figure 2B), and the “groove” site (Figure 2C) on the opposite side of the protein. For both the “latch” and “groove” predicted pockets, small molecules could be envisioned to stabilize the closed form of SHP2 by forming interactions with both N–SH2 and PTP domains.

A.

Allosteric site 1: tunnel

B.

Allosteric site 2: latch

C.

Allosteric site 3: groove

Figure 2. Allosteric pocket predictions of the autoinhibited conformation of SHP2. PTP domain is colored in brown, N–SH2 in green, C–SH2 in blue. A. predicted allosteric site 1 (tunnel), and binding site of 1. B. predicted allosteric site 2 (latch) at the N–SH2/PTP domain interface. C. predicted allosteric site 3 (groove) at the N–SH2/PTP domain interface.


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Considering the potential existence of multiple allosteric pockets, a new screening paradigm was conceived using the engineered mutant protein (Figure 3A).

A library of

approximately 1.5 M compounds was screened as previously reported28 using DIFMUP as the substrate and diphosphotyrosyl–IRS–1 to activate SHP2. Near full–length protein (SHP21–525) was employed to identify compounds with all modes of phosphatase inhibition, and active site inhibitors were removed on the basis of their activity in the phosphatase activity assay utilizing the SHP2PTP protein. In order to remove inhibitors that bound to the previously characterized allosteric site (e.g. 1, and recently reported scaffolds30,31), we selected compounds which maintained biochemical activity on the double mutant (SHP2T253M/Q257L). Finally, we employed X–ray crystallography to determine binding mode and differential scanning fluorimetry (DSF) to investigate thermal stabilization. This screening paradigm led to a collection of compounds with unknown modalities of inhibition, including the triazolo–quinazolinone, SHP244 (2, Figure 3B).32 A.


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B. O

Cl N

N

HO

5

N N

O 5

SHP244 (2) SHP2 IC50 = 60 µM T253M/Q257L SHP2 IC50 = 68 µM PTP SHP2 IC50 > 100 µM DSF (∆Tm) = 0.31 °C 1–525

Figure 3. A. Screening paradigm for novel allosteric pocket discovery. B. Structure, biochemical, and biophysical profile of SHP244 (2). Compound 2 proved to be a weak SHP2 inhibitor (SHP21–525 IC50 = 60 µM), was approximately equipotent on the double mutant (SHP2T253M/Q257L IC50 = 68 ± 10 µM), and selective over the phosphatase domain (SHP2PTP IC50 > 100 µM). Compound 2 was synthesized as illustrated in Scheme S1 (see supporting information).

The highly conjugated tricyclic

structure possessed poor aqueous solubility (0.047 mM in pH 6.8 buffer) and high lipophilicity (cLogP = 3.9). Despite weak biochemical inhibition and poor physicochemical properties, we prioritized 2 for further biophysical characterization using X–ray crystallography, and for optimization for improved biochemical inhibition and physicochemical properties. The crystal structure of 2 in complex with SHP2 was successfully determined at 2.2 Å and bound to the predicted binding pocket (vide supra) located at the interface between the N– terminal SH2 and the PTP domains (Figure 4A). The binding site of 2 is located approximately 20 Å away from the binding site of 1. Overall, the shape and conformation of the SHP2–2


8

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protein structure (PDB code 6BMR) was similar to the SHP2–1 protein structure (PDB code 5EHR),25 as well as the previously reported apo–protein structure (PDB code 2SHP).1 However, several protein residues reposition (e.g. E83, R265) in order to accommodate 2, and another 8 residues (H85–E90 and N92–V95) appear more ordered as compared to the SHP2–1 structure (overlay, Figure 4B). The structure revealed several protein ligand interactions, including the following: hydrogen bonding between the phenol of 2 and residues L262 and R265; interaction of the methoxy group of 2 with Q79 via water–mediated binding; triazole nitrogens of 2 make interactions with the backbone NH and side chains of R265 and Q269, respectively; the chlorophenyl and quinazolinone phenyl rings of 2 interact with Q269 and Q79, respectively, via pi stacking. Interestingly, the ketone of 2 makes no polar interactions and is located in a hydrophobic area formed by H84 and Y80. Furthermore, we recognized that the chlorophenyl ring of 2 incompletely fills a hydrophobic groove (formed by Q269, N281, I282, L283, H84, Y80) which terminates in hydrophilic residues K280 and K274 (Figure 4C). We thus exploited this area of the binding pocket via structure–based design. The 5–position of the chlorophenyl ring of 2 appeared as an optimal vector for this purpose. In addition, K266 was oriented 4.4 Å away from the phenolic ring of 2. We thus hypothesized that an acidic moiety positioned para to the phenol of 2 would capture K266 in an intermolecular hydrogen bond.


9


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unoccupied tunnel binding site 1 occupied latch binding site of 2

B.


10

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C.

Figure 4. A. X–ray of SHP2 and SHP244 (2) at 2.2 Å. B. Binding site of 2 (latch) with interaction residues and targeted residues displayed in ball and stick. PTP domain is colored brown, N– SH2 in green, C–SH2 is not shown. Interacting residues and ordered loop residues colored in magenta. 2 is colored in blue. C. Hydrophobic groove shown with surface representation, K280, and K274. Colored by electrostatic charge.

Accordingly, proline–acid 8 was designed (SHP844, Figure 5A) to fill the hydrophobic channel and capture K280 in a hydrogen bond. We hypothesized that the carboxylate of 8 would also lower the lipophilicity (e.g., cLogP of 2 = 3.9; cLogP of 8 = 3.0) and improve the aqueous solubility of 2 (0.047 mM in pH 6.8 buffer). Additionally, SHP504 (9, Figure 5A) was designed to capture K266 and prepared in a similar fashion to 2. Compounds 8 and 9 were next evaluated for SHP2 biochemical inhibition. A modest increase in biochemical inhibition of 8 (SHP21–525 IC50 = 18.9 µM) and 9 (SHP21–525 IC50 = 21 µM) was observed as compared to 2 (SHP21–525 IC50 = 60 µM). Both compounds were inactive against the PTP domain (8, 9: SHP2PTP IC50 >100 µM). A modest thermal stabilization of SHP2 via DSF occurred (8: ∆Tm = 0.58 °C; 9 ∆Tm = 0.39 °C; 2: ∆Tm = 0.31 °C) that was consistent with the increase in biochemical potency.

The carboxylic acid functionality improved the


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aqueous solubility of 8 (0.895 mM in pH 6.8 buffer) and 9 (0.535 mM in pH 6.8 buffer) as compared to 2 (0.047 mM in pH 6.8 buffer). To determine the binding mode and characterize interactions, the crystal structures of 8 and 9 were determined with SHP2 at 2.4 Å (Figure 5B), and 2.1 Å, respectively (PDB codes 6BMX, 6BMV, respectively). The structures revealed that both inhibitors bound in a similar manner to 2, and the overall protein conformation was also similar. As hypothesized, the proline–acid side chain of 8 oriented towards the lipophilic tunnel, and approached within hydrogen bonding distance of the side chains of K280 and Y80 (2.9 Å and 3.1 Å, respectively). Additionally, the carboxylate of 9 formed an intermolecular hydrogen bond with K266 (2.8 Å, Figure 5C). A. SHP844 (8) SHP21–525 IC50 = 18.9 µM SHP2PTP IC50 > 100 µM DSF (∆Tm) = 0.58 °C Aq solubilty: 0.895 mM SHP504 (9) SHP21–525 IC50 = 21 µM SHP2PTP IC50 > 100 µM DSF (∆Tm) = 0.39 °C Aq solubilty: 0.535 mM

SHP357 (10) SHP21–525 IC50 > 100 µM DSF (∆Tm) = no shift

SHP836 (11) SHP21–525 IC50 = 12 µM DSF (∆Tm) = 3.0 °C


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B.

C.

Figure 5. A. Structure, biochemical and biophysical profile of compounds 811. B. X–ray structure of SHP2 bound to SHP844 (8) at 2.4 Å, showing H–bonds to K280 and Y80. C. X–ray of SHP2 and SHP504 (9) at 2.1 Å, showing hydrogen bond to K266. PTP domain is colored brown, N–SH2 in green, C–SH2 is not shown. Critical interacting residues colored in magenta. 8, 9 are colored in blue. From our analysis of X–ray structures of 2, 8, and 9 individually bound to SHP2, we observed that the original allosteric binding pocket of 1 was unperturbed, and that dual occupation of both allosteric sites might be possible. Multiple ligands binding simultaneously to


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the same protein is well precedented and reported in a number of different contexts. Typically, multiple native ligands can bind to the same protein (e.g. hemoglobin), or a native ligand and a small molecule targeted drug can bind simultaneously (e.g., allosteric and orthosteric modulation of GPCRs).33,34 In other contexts, multiple small molecule drugs are reported to bind to non– targeted proteins (e.g. albumin,35 p–glycoprotein,36 etc.). In most cases, the drugs occupy the same binding site.37 Less described in the literature, however, are examples of multiple, small molecule drugs that target a therapeutic protein simultaneously. A recent example of dual, targeted inhibition stems from drug discovery efforts towards the oncogenic fusion protein BCR– ABL.38

ATP–competitive inhibitors of the kinase active site (e.g. imatinib, nilotinib) and

myristoyl binding site inhibitors (e.g. GNF–2, ABL001) simultaneously bind and occupy the catalytic site and the myristoyl sites, respectively, as characterized by X–ray crystallography and synergism studies. One particularly significant finding was that dual inhibition prevented the emergence of resistance to each individual drug in preclinical animal models.39 To determine whether dual, targeted inhibition of SHP2 was possible utilizing both allosteric binding sites, we conducted biochemical inhibition studies with combinations of inhibitors of both allosteric sites. We observed a dose–dependent decrease in SHP2 activity and a modest enhancement of IC50 for 1 (Figure 6A) with increasing concentrations of 9, indicating possible cooperativity between the two binding modes. This finding contrasted with the results from combinations of 1 and 10, an inactive analog of 9, which showed no changes to either SHP2 activity or IC50 for 1 with increasing concentrations of 10 (see supporting information). As expected, the combination of SHP836 (11)25,26 with 1, both of which bind to the same allosteric (tunnel) pocket, weakened the inhibitory effects of 1 due to competition with a weaker inhibitor for the same binding site.


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Recently, Copeland and colleagues reported a local curve–fitting method for the analysis of four–parameter inhibition curves using multiple inhibitors simultaneously.40 The resulting plots of IC50 of one compound vs. the concentration of a second compound provide mechanistic insight into the relationship of the two inhibitors on the enzyme. We applied these fitting methods to our data and compared the results to those predicted by Copeland et al. Plotting the IC50 of 1 against increasing concentration of 9 resulted in a descending hyperbola (Figure 6B) which indicates synergism between the two inhibitors on the enzyme. Fitting to equation 1 (see methods section) yields the apparent binding constants KI = 0.066 ± 0.003 µM, KJ = 17.6 ± 15 µM, and the interaction constant α = 0.67. Despite the relatively low potency of inhibitors at the latch site, this analysis yields values for KI and KJ that are extremely similar to the IC50 values for 1 and 9 in isolation (0.07 µM and 21 µM, respectively). Combination data using 1 and 11 (SHP836, Figure 6A) and 1 and 10 (see supporting information) were replotted using the above method and as predicted, the resulting fits clearly demonstrated varied interactions between different pairs of molecules. Compound 11 has been demonstrated to bind in the same pocket as 1, albeit at a much lower affinity, and thus binding of these compounds should be mutually exclusive. As such, plotting the IC50 of 1 vs [11] resulted in a linear dependence between these values and fitting to equation 2 (see methods section) yielded apparent binding constants of 0.069 ± 0.004 µM and 3.3 ± 0.23 µM, respectively for 1 and 11 (Figure 6B). Taken together, these data suggest that simultaneous occupation of both the hinge and latch sites of SHP2 by small molecules can enhance the inhibition of SHP2 enzymatic activity.


15


A. Dual inhibition of SHP2 by SHP099 (1) and SHP504 (9)

Dual inhibition of SHP2 by SHP099 ( 1) and SHP836 (11)

[SHP504] (9)

8

[SHP836 (11)]

8

100 µM 6

100 µM 6

50 µM 25 µM

4

Rate

Rate

12.5 µM 6.25 µM

2

50 µM 25 µM

4

12.5 µM 6.25 µM

2

3.13 µM 0 0.0001 0.001

3.13 µM

1.56 µM 0.01

0.1

1

10

100

0 0.0001 0.001

0 µM

1.56 µM 0.01

0.1

1

10

100

0 µM

[SHP099 (1)] (µM)

[SHP099 (1)] (µM)

B. IC50 for SHP099 (1) in combination with SHP504 (9) IC50 for SHP099 (1) in combination with SHP836 (11) 0.8

SHP099 (1) IC50 (µM)

0.08

SHP099 (1) IC50 (µM)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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0.06 0.04 0.02

0.6 0.4 0.2 0.0

0.00 0

20

40

60

80

100

120

0

10

20

30

[SHP836 (11)] (µM)

[SHP504 (9)] (µM)

Figure 6. A. Dual inhibition of 1 and 9, of 1 and 11. (b) Synergistic relationship of 1 and 9; mutually exclusive relationship of 1 and 11. Seeking more biophysical evidence of simultaneous binding, we attempted to determine the X–ray structure of SHP2 with 1 and the site 2 (latch) binders (e.g., 2, 8, 9). Gratifyingly, each of the three second site analogs (i.e., 2, 8, and 9) bound to SHP2 simultaneously with 1 (Figure 7A: SHP2–1–2, 2.1 Å, PDB code 6BMU; see supporting information for SHP2–1–8, PDB code 6BMY, and SHP2–1–9, PDB code 6MBW). The conformation of SHP2 while bound by both allosteric site inhibitors (e.g. 1 and 2) was very similar to the individual, bound conformations, and to the apo–structure conformation (PDB code 2SHP). As characterized previously in the X– ray structure of the SHP2–1 complex,28 movement in R111 was observed in the tunnel allosteric site of the X–ray structure of the SHP2–1–2 complex (Figure 7B). Consistent with the X–ray


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structure of the SHP2–2 complex, E83 and R265 move to accommodate 2 in the X–ray structure of the SHP2–1–2 complex. Residues H85–E90 and N92–V95 are also more ordered in SHP2–1– 2 as compared to SHP2–1 (overlay, Figure 7C). Furthermore, thermal stabilization of SHP2 in the presence of 1 and 2 (∆Tm = 8.46 °C), 1 and 8 (∆Tm = 8.77 °C), and 1 and 9 (∆Tm = 8.51 °C) was greater than with either compound alone (1: ∆Tm = 7.59 °C; 2: ∆Tm = 0.31; 8: ∆Tm = 0.58 °C; 9 ∆Tm = 0.39 °C), which is consistent with the biochemical results, X–ray analysis, and the simultaneous binding hypothesis. A.

B.

C.

occupied tunnel binding site of 1 occupied latch binding site of 2

Figure 7. Dual allosteric binding of SHP2. A. X–ray structure of SHP2–1–2 at 2.1 Å showing simultaneous occupation of tunnel and latch allosteric sites. B. Binding site 1 (tunnel) overlay from SHP2–1–2 and SHP2–1. C. Binding site 2 (latch) overlay from SHP2–1–2 and SHP2–1. PTP domain is colored in brown, N–SH2 in green, C–SH2 in blue. Critical interacting residues and ordered loop residues colored in magenta. 1 is colored in yellow, 2 is colored in blue. We next examined whether the identified allosteric latch site inhibitors (e.g. 2, 8, 9) could inhibit SHP2 in cells. From our previous cellular experiments with 1 and related analogs, we understood that modulation of downstream MAPK pathway markers in EGFR amplified KYSE– 520 cells, using DUSP6 mRNA levels, would likely require sub–micromolar biochemical inhibition concentrations. Given the limited biochemical inhibition of 2, 8, and 9 and the


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predicted poor permeability of those compounds due to the presence of ionizable functional groups, we chose 10 and 30 µM assay concentrations. DUSP6 mRNA levels, a commonly used pharmacodynamic marker of the MAPK pathway downstream of SHP2, was measured after 2 h of compound treatment and 1 was included as the positive control. A significant and dose– dependent downregulation of DUSP6 transcription in KYSE–520 cells with 2, 8 and 9 was observed (Figure 8A). Cellular treatment with 2, 8 and 9 at 30 µM reduced DUSP6 levels by 65%, 43%, and 55%, respectively. As expected, cellular treatment with 1 at 0.2 µM reduced DUSP6 levels by 50% and near complete reduction was achieved at 3 µM. These results were consistent with observed potency differences in biochemical assays for 1, 2, 8, and 9. In contrast, the inactive derivative SHP357 (10, Figure 5A), showed only a modest decrease of DUSP6 levels (26% at 30 µM), suggesting the activity of 2, 8, and 9 was likely mediated by inhibiting SHP2. More importantly, 1 (3 µM), 2 (30 µM), and 9 (30 µM) did not downregulate DUSP6 levels in the A375 melanoma cell line bearing the BRAFV600E mutation (Figure 8B). This finding further suggests that the effects were upstream of BRAF within the MAPK pathway and via SHP2. Taken together, these data support that the site 2 allosteric inhibitors described here exhibit activity against endogenous SHP2 in its native environment in cells. Since we had observed biochemical enhancement of IC50 via dual inhibition and observed dual occupancy of both allosteric sites via X–ray structures, we next characterized combinations of site 1 and site 2 inhibitors in KYSE–520 cells. Indeed, combination treatment of KYSE–520 cells with 9 (30 µM) and 1 (0.2 µM) improved DUSP6 downregulation compared to either of the single agents (30 µM of 9, p= 0.0064; 0.2 µM of 1, p= 0.0007), although the combination effect was not observed with 10 µM of 9 (Figure 8C). As expected, the inactive compound 10 (10 µM or 30 µM) provided no combination benefit with 0.2 µM of 1. Combination benefits with 1 were


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not observed with 2 and 8 (see supporting information). This may be related to the weak cellular activity of these site 2 inhibitors, which was expected given their biochemical inhibition, or the small assay window of DUSP6 levels.

In summary, the combination data supports the

simultaneous binding of site 1 and site 2 allosteric inhibitors in cells. A.

B.

C.

Figure 8. Cellular activity of site 2 inhibitors and combinations with site 1 inhibitor SHP099. A. Downregulation of MAPK pathway pharmacodynamics marker, DUSP6 transcript levels, by individual site 1 and site 2 SHP2 inhibitors in KYSE–520 cells after 2 h treatment with indicated concentrations. B. Absence of DUSP6 modulation by SHP2 inhibitors in BRAFV600E melanoma cell line A375. C. Combination effects of 1 with latch site inhibitors SHP504 and inactive derivative SHP357 on DUSP6 transcript level in KYSE–520 cells. DUSP6 levels were normalized to β–actin transcript levels. Data represent average from four replicates and error bars denote standard deviation. **, P< 0.01; ***, P< 0.001.


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Historically, proteins with engineered binding site mutations are powerful tools to characterize the cellular mechanisms of ligand binding. In this article, we describe an alternative methodology that utilizes such modified proteins to identify an additional small molecule modality of SHP2 inhibition. Inhibitor 2 was identified via a screening paradigm which first removed undesired active site inhibitors, and later, allosteric site 1 inhibitors. Thermal protein stabilization was employed to prioritize hits for further biophysical characterization. Characterization of 2 in biochemical inhibition experiments and structure determination of SHP2 with 2 revealed a second, distinct allosteric pocket at the interface of the N–terminal SH2 and PTP domains. This binding site discovery correlated with in silico pocket predictions made a priori.

Protein–ligand interactions were characterized and structure–based drug design was

employed to improve the biochemical inhibition and physicochemical properties of 2, resulting in 8 and 9. Compound 9 enhanced the inhibition of 1 in biochemical phosphatase experiments and 9 enhanced downregulation of the MAPK pharmacodynamic marker DUSP6 by 1 in cellular experiments. Mutual exclusivity biochemical experiments supported dual occupation of both allosteric binding sites, and X–ray crystallography confirmed unambiguously that inhibitors of both allosteric sites can bind to SHP2 simultaneously. This phenomenon represents a rare example of dual, simultaneous binding to a therapeutic target protein, and the only fully allosteric example that we are aware of. Small molecule modulation of SHP2 is of increasing therapeutic interest given its importance in known oncogenic pathways and emerging role in immuno–oncology.

This

additional, novel allosteric inhibition modality of 2 and its derivatives, also stabilizes SHP2 in the auto–inhibited and inactive conformation and complements our previous reports describing the inhibition modality of 1. Combining two distinct but compatible SHP2 inhibitors may offer advantages in enhancing SHP2 inhibition and overcoming resistance. We speculate that the


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screening approaches described in this article could be further applied to SHP2 to uncover additional inhibition modalities beyond that of 1 and 2, perhaps targeting the groove site (Figure 1C). Upon further optimization, the triazolo–quinazolinone compound class (e.g. 2, 8, and 9) should enable further interrogation of the second allosteric binding site, and the multifaceted roles of SHP2 in cancer and related molecular pathologies.

The screening methodology

described in this communication, if employed upon other protein–ligand pairs of interest, may enable the identification of additional dual–targeted modalities for therapeutic proteins. METHODS. SHP2 pocket prediction. Potential small molecule SHP2 binding sites were assessed using the Maestro program from the Schrödinger suite.27,28 Briefly, the SHP2 crystal structure (PDB entry 2SHP) was prepared using the standard parameters available in the protein preparation workflow, with hydrogens added at pH 7.0 and water and other co–crystallized small molecules removed.41 A single polypeptide chain (A) was analyzed for potential small molecule binding pockets using the SiteMap algorithm, applying the default requirement of at least 15 site points per site. The druggability of the predicted sites was assessed through calculation of the following physicochemical properties: hydrophilicity, volume, solvent exposure, contacts, hydrophobicity, and hydrogen–donor/acceptor sites (see supporting information, table S1). Dual inhibition and mutual exclusivity.

Dual inhibition biochemical experiments were

conducted as described in the supporting information, with slight modifications: compounds were dispensed pairwise in matrix format to enable combinations of various inhibitors and concentrations; final reaction volume was 10 µL; reaction was measured in kinetic mode over 15 min on a Spectramax M2 (Molecular Devices) using excitation and emission wavelengths of 340 nm and 450 nm, respectively.

Inhibitory data was analyzed using GraphPad Prism 7.0c


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(GraphPad Software, Inc.) and resulting curves were fit using the dual inhibition equation, or reductions thereof, as described by Copeland and colleagues.40 (1)

In the circumstance of mutually exclusive binding where both molecules occupy the same binding site, α = infinity and the dual inhibition equation simplifies to:

(2) DUSP6 qPCR assay. KYSE–520 and A375 cells were seeded in 6–well plates (500,000 cells/well) and after overnight culturing cells were treated with indicated SHP2 site 2 inhibitors at 10 µM or 30 µM alone or in combination with 0.2 µM site 1 inhibitor 1 for 2 h. Total RNA was extracted by QIAcube (QIAGEN) using RNeasy Mini columns. cDNA was synthesized using qScript™ cDNA SuperMix (Quanta Biosciences). DUSP6 transcript level was measured using TaqMan™ Real Time PCR assay (Applied Biosystems; assay ID: Hs00169257_m1).

Human β–actin (Applied

Biosystems; catalog no.: 4310881E) was used as an internal control.

Differential Scanning Fluorimetry. Differential Scanning Fluorimetry (DSF) was used as a method to identify compounds that stabilize SHP2 from thermal denaturation. The following assay conditions were used: 100 µg mL-1 SHP2, 5× SYPRO Orange dye (5000× concentrate in DMSO; Life Technologies), 100 mM Bis–Tris (pH 6.5), 100 mM NaCl, 0.25 mM TCEP, and 5 % DMSO. The final concentration of each compound evaluated was 100 µM. To carry out the experiment, 9.5 µL DSF assay solution was dispensed into an assay plate (LightCycler; 480 Multiwell Plate 384 White) containing 500 nL of compound dissolved in DMSO then mixed. The final assay volume was 10 µL per well in a 384–well format. Plates were then sealed after reagent addition, centrifuged at 1000 rpm for 1 minute, and read on a Bio–Rad C1000 Thermal


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Cycler with a CFX384 Real Time System using an excitation of 465 nm and an emission at 580 nm. The temperature was ramped from 25 °C to 75 °C and measurements were taken at 0.5 °C increments. The melting temperature (Tm) of the raw fluorescence data was identified as the midpoint of the transitions via a semi–parametric fit. The reported Tm values are the average of 5 individual experiments. The ∆Tm was determined by comparing the individual Tm values for each compound with the mean Tm of the apo–SHP2 protein controls containing DMSO only. Crystallization and Structure Determination. Sitting drop vapor diffusion method was used for crystallization, with the crystallization well containing 17% PEG 3350 and 200 mM ammonium phosphate and a drop with a 1:1 volume of SHP2 protein and crystallization solution. Crystals were formed within five days, and subsequently soaked in the crystallization solution with 2.5 mM of 2, 8, or 9 alone or in addition to 1 mM of 1.

This was followed by

cryoprotection using the crystallization solution with the addition of 20% glycerol and 1 mM 2, 8, or 9 alone or in addition to 0.5 mM 1, followed by flash freezing directly into liquid nitrogen. Diffraction data for the SHP2–1 complex is reported elswhere25, 26 and SHP2/compound 2, 8, 9 complex were collected on a Dectris Pilatus 6M Detector at beamline 17ID (IMCA–CAT) at the Advanced Photon Source at Argonne National Laboratories. The data were measured from a single crystal maintained at 100 K at a wavelength of 1 Å, and the reflections were indexed, integrated, and scaled using XDS.42 The spacegroup of the complexes was P21 with 2 molecules in the asymmetric unit. The structure was determined with Fourier methods, using the SHP2 apo–structure1 (PDB accession 2SHP) with all waters removed. Structure determination was achieved through iterative rounds of positional and simulated annealing refinement using BUSTER,43 with model building using COOT.44 Individual B–factors were refined using an overall anisotropic B–factor refinement along with bulk solvent correction.


The solvent,

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phosphate ions, and inhibitors were built into the density in later rounds of the refinement. Data collection and refinement statistics are shown in Tables S2–S7 found in the supporting information. ANCILLARY INFORMATION PDB ID codes: 6BMR for SHP2 in complex with compound 2. 6BMX for SHP2 in complex with compound 8. 6BMV for SHP2 in complex with compound 9. 6BMU for SHP2 in compex with compounds 1 and 2. 6BMY for SHP2 in complex with compounds 1 and 8. 6MBW for SHP2 in complex with compounds 1 and 9. Authors will release the atomic coordinates upon article acceptance. Corresponding author information. Email: [email protected]; telephone: 617– 871–7729. Email: [email protected], telephone: 617–871–4005. Novartis Institutes for Biomedical Research, Inc. 250 Massachusetts Avenue, Cambridge, MA 02139. Acknowledgements: Use of the IMCA–CAT beamline 17–ID at the Advanced Photon Source was supported by the companies of the Industrial Macromolecular Crystallography Association through a contract with Hauptman–Woodward Medical Research Institute. Use of the Advanced Photon Source was supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE–AC02–06CH11357.

JRL gratefully

acknowledges postdoctoral fellowship funding from the American Cancer Society® (Award 128126). The authors thank the entire SHP2 team and our collaborators. Supporting information available: includes chemistry experimentals, biochemistry procedures, cellular pharmacology, X–ray figures, and X–ray data tables.


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