
==== Front
J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02117-3
10.1016/j.jbc.2024.107616
107616
Research Article
Bivalent target-binding bioPROTACs induce potent degradation of oncogenic SHP2
Hoffman Megan 12
Krum David 13
Wittrup K. Dane wittrup@mit.edu
123∗
1 Koch Institute for Integrative Cancer Research, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
2 Department of Biological Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
3 Department of Chemical Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
∗ For correspondence: K. Dane Wittrup wittrup@mit.edu
31 7 2024
9 2024
31 7 2024
300 9 1076163 4 2024
2 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Targeted protein degradation is an emergent and rapidly evolving therapeutic strategy. In particular, biologics-based targeted degradation modalities (bioPROTACs) are relatively under explored compared to small molecules. Here, we investigate how target affinity, cellular localization, and valency of bioPROTACs impact efficacy of targeted degradation of the oncogenic phosphatase src-homology 2 containing protein tyrosine phosphatase-2 (SHP2). We identify bivalent recruitment of SHP2 by bioPROTACs as a broadly applicable strategy to improve potency. Moreover, we demonstrate that SHP2-targeted bioPROTACs can effectively counteract gain-of-function SHP2 mutants present in cancer, which are otherwise challenging to selectively target with small molecule constructs. Overall, this study demonstrates the utility of bioPROTACs for challenging targets, and further explicates design principles for therapeutic bioPROTACs.

Keywords

protein degradation
protein engineering
proteasome
E3 ubiquitin ligase
cancer
BioPROTAC
PROTAC
SHP2
SPOP
targeted protein degradation
Abbreviations

bioPROTAC biologic PROTAC

BSA bovine serum albumin

DMEM Dulbecco's modified Eagle's medium

ERK extracellular signal-regulated kinase

FBS fetal bovine serum

MAPK mitogen-activated protein kinase

NLS nuclear localization sequence

PROTAC proteolysis-targeting chimera

sfGFP superfolder GFP

SHP2 src-homology 2 containing protein tyrosine phosphatase-2

SPOP speckle-type pox virus and zinc finger protein

TPD targeted protein degradation

UPS ubiquitin proteasome system

xK lysine free

Reviewed by members of the JBC Editorial Board. Edited by George DeMartino
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pmcTargeted protein degradation (TPD) is an emerging therapeutic paradigm that holds immense promise in the field of drug discovery. Unlike conventional approaches centered on stoichiometric activity inhibition, TPD achieves efficacy by reducing target protein abundance. This method offers numerous advantages over traditional inhibition techniques. Firstly, degradation-inducing constructs (degraders) can counteract a broader spectrum of target proteins, including those previously deemed "undruggable," by using ligands that bind to noncatalytic pockets on the target protein's surface. Unlike inhibitors, degraders are reusable, enabling a single degrader to eliminate multiple target proteins in a substoichiometric manner (1). Moreover, the degradation mechanism of action results in more comprehensive and sustained effects on downstream signaling pathways (2).

The most prevalent mechanism to achieve TPD is through chemical compounds known as proteolysis-targeting chimeras or PROTACs (3). PROTACs take advantage of the endogenous proteasomal degradation pathway to selectively eliminate proteins of interest. More specifically, PROTACs function as heterodimeric ligands that bring a target of interest into proximity with components the ubiquitin proteasome system (UPS). If the orientation of the target protein is favorable relative to the UPS, ubiquitin molecules are ligated to lysine residues on the surface of the target protein, which triggers proteasomal degradation of the target protein.

The majority of PROTACs are small molecule-based and are often derived from existing inhibitor compounds, used as target-recruiting ligands, linked to ligands that recruit E3 ligases, and the component of the UPS that is responsible for substrate recruitment prior to ubiquitination. However, small molecule PROTACs have a limited range of E3 ligases they can leverage. Only about a dozen out of the nearly 500 known E3 ligases currently have reported small molecule ligands (4). As an alternative approach, biologic PROTACs (bioPROTACs) are protein-based constructs designed to achieve targeted degradation using the proteasome (5, 6, 7). BioPROTACs operate similarly to small molecule PROTACs but instead use a target-binding protein linked to a full E3 ligase or an E3 ligase subunit. Because bioPROTACs incorporate the E3 itself into the construct, ligand expression, or availability is not a limiting factor and essentially any E3 ligase can be used. Additionally, protein-based ligands can have greater affinity and specificity for the target protein than small molecules and can leverage display platforms for rapid development. The combination of enhanced E3 ligase diversity and improved target ligand recruitment gives bioPROTACs the potential for highly effective degradation.

Although bioPROTACs are potentially efficacious therapies, they remain a relatively under studied modality within the field of targeted protein degradation. To address this, we previously used a cell-based screening platform to identify several novel bioPROTACs, which were derived from preexisting target-recruiting scaffolds and validated E3 ligases (8, 9), that were capable of degrading src-homology 2 containing protein tyrosine phosphatase-2 (SHP2), a relevant target for a multitude of disease states, including cancer (10, 11). From our screening data, we elucidated design principles and identified trends contributing to successful bioPROTACs, but further characterization of degrader design principles is needed. In this study, we aim to deepen our understanding of bioPROTAC properties and evaluate how these properties either enhance or inhibit degradation activity. To achieve this, we developed a novel loss-of-signal cell line engineered to minimize false positives. This assay platform enabled us to investigate how modifications to affinity, localization, and avidity influence the degradation efficiency of SHP2 bioPROTACs, particularly using the E3 ligase speckle-type pox virus and zinc finger protein (SPOP). Our findings highlight multivalent target recruitment as a potentially impactful strategy for enhancing bioPROTAC potency across various E3 ligases. Furthermore, we provide evidence that bioPROTACs show indications of antitumor efficacy against oncogenic SHP2 mutants and data to suggest that bioPROTACs may exhibit enhanced activity compared to small molecule PROTACs. Overall, this study advances our knowledge of designing potent bioPROTACs for therapeutic development and underscores the potential advantages of bioPROTACs over small molecule alternatives.

Results

Engineering a lysine-free GFP assay to detect SHP2 degradation

We first sought to develop an assay platform that would enable unambiguous quantification of target degradation. In particular, we wanted to ensure that the assay results were not convoluted by off-target ubiquitination, the tagging of ubiquitin to lysine residues not on the target protein but on the fluorescent reporter. Prior studies warn that screening loss-of-signal assays where the target protein is directly fused to GFP can result in false positives where the degraders identified only degrade the GFP-tagged target and not the target alone (12). We hypothesized that engineering a lysine-free GFP protein would enable direct labeling of the target protein while preventing unwanted ubiquitin tagging to GFP. This hypothesis is supported by recent findings in which RIPK1 fused to a lysine-free luciferase was shown to have a higher half-maximal degradation constant (DC50) than RIPK1 fused to WT luciferase, indicating that the DC50 of the RIPK1 fused to WT luciferase may overestimate the capabilities of the PROTACs tested due to ubiquitin tagging on the surface of the luciferase (13). In attempt to make a lysine-free fluorescent protein, we and others have previously tried to express enhanced green fluorescent protein with all lysine residues converted to similarly charged arginine residues but observed poor stability and no fluorescent activity (14, 15). However, Lawrence et al. successfully engineered a supercharged GFP using an alternative “superfolder” GFP (sfGFP) scaffold, indicating that this sfGFP framework might be more stable and amenable to alterations (16). To test this, we expressed sfGFP with all lysine residues mutated to arginine residues (notated as xK to indicate lysine-free), fused to SHP2, our target of interest (Fig. 1A). We found that the lysine-free superfolder GFP (sfGFPxK) expressed well and was fluorescent, although at a partially reduced level compared to the WT (Fig. 1B). Encouraged by these results, we stably overexpressed SHP2-sfGFPxK in HEK293T/17 cells and proceeded to use these cells for characterization of bioPROTAC activity against SHP2.Figure 1 Lysine-free GFP assay robustly detects SHP2 degradation by bioPROTACs.A, schematic of a typical SHP2-GFP loss-of-signal degradation assay (top) with potential off-target ubiquitination of GFP in comparison to the lysine-free GFP assay platform (bottom). Target protein fused to GFP is brought into proximity with the ubiquitin proteasome system using a bioPROTAC consisting of a fibronectin type III (FN3)-derived monobody scaffold that binds to the target protein linked to an E3 ligase component. B, fluorescent expression of GFP constructs measured by flow cytometry 48 h after transient transfection with plasmid DNA encoding for sfGFP with or without lysine residues (xK) coexpressed with mCherry. C, Western blots of SHP2-sfGFPxK stable line HEK293T/17 cells 48 h after transfection with FLAG-tagged bioPROTACs. D fluorescent expression of SHP2-sfGFPxK stable line cells versus WT HEK293T/17 cells. E, fluorescent expression of SHP2-sfGFPxK stable line cells 48 h after transfection with bioPROTACs shown as histograms (left) and bar graphs (right) where row numbers of the histograms correspond with column labels in the bar graphs. F, replicate measurements of fluorescence of SHP2-sfGFPxK stable line cells 48 h after transfection with positive and negative control bioPROTACs. Corresponding Z′ values are listed. Gray shading indicates three standard deviations above and below the mean. Replicate samples in panels F and G are technical replicates. Statistics: GFP MFI compared by one-way ANOVA with Sidak’s multiple comparisons test. ∗∗∗∗p < 0.0001. bioPROTAC, biologic PROTAC; MFI, mean fluorescence intensity; sfGFP, superfolder GFP; xK, lysine free.

We first validated that the SHP2-sfGFPxK assay platform responded to known degraders. We used two SHP2 bioPROTACs, CS3-SPOP, and NSA5-SPOP, identified in our previous work as positive controls to test the new assay platform (14). Notably, CS3 and NSA5 are SHP2-binding monobodies, which are derivatives of the fibronectin type III (FN3) domain developed by Sha et al. that have low nanomolar affinity for SHP2 and bind to distinct epitopes on the C-SH2 and N-SH2 domains of the target protein (8). SPOP is an E3 ligase in which the ligand recruiting MATH domain has been replaced by the monobodies. Although nearly a dozen different bioPROTACs were identified from our prior screen, we chose to pursue CS3-SPOP and NSA5-SPOP due to the consensus from previous reports that use of SPOP in bioPROTACs results in robust and promiscuous degradation (7, 9, 17, 18, 19, 20). Furthermore, SPOP is unique to bioPROTACs, as no small molecule PROTACs are yet reported to leverage SPOP for targeted degradation (21, 22). For all experiments, we used size-matched untargeted negative controls in which the monobodies were mutated such that they no longer bound to SHP2, as validated previously (14). Additionally, bioPROTACs are FLAG tagged and coexpressed with an mCherry transfection marker. For all bioPROTAC transfections, cells are gated on mCherry expression to exclude untreated cells from the analysis.

While characterizing the assay platform, we confirmed by Western blot that both endogenous SHP2 and SHP2-sfGFPxK were degraded by CS3-SPOP and NSA5-SPOP but not by their corresponding negative controls (Fig. 1C). Interestingly, we also observed much lower expression of CS3-SPOP compared to NSA5-SPOP yet both bioPROTACs achieved similar levels of SHP2 degradation, indicating a potential difference in the catalytic activity of these two constructs. These differences are likely due to unintended degradation of the bioPROTAC, which is to be expected to some degree given that evidence shows that SPOP is ubiquitinated upon target engagement (23), although why CS3-SPOP is seemingly more heavily impacted than NSA5-SPOP remains to be validated. For additional assay characterization, we measured changes in GFP mean fluorescence intensity (MFI) by flow cytometry to ensure that the assay platform could robustly detect changes in SHP2-sfGFPxK expression levels (Fig. 1, D and E). We also validated that the SHP2-sfGFP platform responded to incubation with SHP2-D26, a SHP2-targeted small molecule PROTAC (24), by both flow cytometry and Western blot (Fig. S1), indicating that the assay platform is suitable for detecting degradation from different types of PROTACs. Finally, we used a Z-factor analysis to validate the robustness of the assay for detecting SHP2-sfGFPxK degradation (Fig. 1F) (25). Notably, due to a lack of any known false positive PROTACS that cause off-target ubiquitination of SHP2-GFP, we were unable to validate with certainty that the SHP2-GFPxK platform negates all false positives. However, we hypothesized that screening with the lysine-free GFP yields results more consistent with endogenous SHP2 degradation than screening with traditional lysine-containing GFP, and proceeded to use the SHP2-GFPxK assay platform for bioPROTAC characterization.

Evaluating impact of target affinity on degradation

We next sought to find ways to improve the degradation activity of CS3-SPOP and NSA5-SPOP. We started by investigating the relationship between the affinity of the bioPROTACs for the target and how much target degradation is achieved. Previous work indicates that affinity is not necessarily a predictor or requisite for degradation of targets by small molecule PROTACs as low affinity target-binding showed similar degrees of degradation compared to higher affinity interactions (26). However, studies of bioPROTACs indicate that higher target affinity may be necessary for elimination of proteins of interest (6, 7). To study the target affinity and target degradation relationship, we performed an alanine scan, systematically mutating residues on CS3 and NSA5 that contact SHP2 (8). We transiently transfected the mutated monobody-SPOP fusions into the SHP2-sfGFPxK cell-based assay platform and measured SHP2 degradation (Fig. 2, A and B). We observed varying degrees of impact on SHP2 degradation ranging from almost no change to complete loss of degradation activity. We next expressed each of the mutant monobodies recombinantly and measured their affinity for SHP2 (Fig. S2). We plotted the relationship between the KD of the monobody for binding SHP2 and the fraction of SHP2-sfGFPxK degraded after 48 h (Fig. 2C). For these monobody-SPOP-fusion bioPROTACs, we found that higher affinity (i.e. lower KD) appeared to monotonically correlate with lower levels of SHP2-sfGFPxK and thus a greater degree of degradation. This result is further supported by findings from KRAS-degrading bioPROTACs. A similar experiment comparing a panel of VHL-mediated bioPROTACs with a range of affinities for KRAS found a correlation between higher affinity and improved degradation of KRAS-GFP (27). Taken together, these findings suggest that in the case of bioPROTACs, improving affinity may lead to optimized elimination of a target, at least down to single-digit nM Kd values, and that these observations are neither target nor E3 ligase specific.Figure 2 BioPROTAC degradation efficiency correlates with target affinity.A and B, fraction of SHP2-sfGFPxK remaining in stable expressing HEK293T/17 cells 48 h after transient transfection with bioPROTACs containing mutant SHP2-binding monobodies CS3 (A) and NSA5 (B) as measured by MFI on flow cytometry normalized to the negative control as 100%. Data shown are assay replicates. C, scatter plot of the relationship between affinity of monobodies for SHP2 (plotted as the KD measured for the interaction on BLI) and fraction of SHP2 remaining after degradation from SPOP-based bioPROTACs with mutant-monobodies. Symbol colors and shapes plotted correspond with data plotted in 2A and 2B. Statistics: Fraction of SHP2-sfGFPxK remaining compared by one-way ANOVA to the corresponding untargeted negative controls (monobody.mut-SPOP) using Dunnett’s multiple comparisons test. ∗∗∗∗p < 0.0001. Replicate samples in panels A and B are technical replicates. bioPROTAC, biologic PROTAC; MFI, mean fluorescence intensity; sfGFP, superfolder GFP; SPOP, speckle-type pox virus and zinc finger protein; xK, lysine free.

Investigating SPOP localization and dimerization

Next, we evaluated contributions of different domains of the SPOP E3 ligase component of the CS3-SPOP bioPROTAC toward SHP2 degradation. Notably, SPOP is a multifaceted protein containing several different interaction domains (Fig. S3). To carry out ubiquitination, SPOP acts as an adapter for Cul3, which is recruited via the 3Box motif (28). Endogenous SPOP recruits antigens via the MATH domain, which we removed and replaced with the SHP2-targeted monobodies to ensure selective bioPROTAC targeting (29). SPOP also contains two different dimerization domains, BTB and BACK, that enable it to form oligomers (Fig. 3A) and a nuclear localization domain (NLS), which together with the dimerization domains results in the localization of SPOP to nuclear speckles (28, 30).Figure 3 SPOP dimerization and localization contribute to degradation efficiency.A, schematic of SPOP dimerization via both BTB and BACK domains. B and C, fraction of SHP2-sfGFPxK remaining in stable line HEK293T/17 cells 48 h after transient transfection with bioPROTACs containing mutations to various dimerization domains (B) or nuclear localization domains (C) in SPOP. Data shown are assay replicates. D, microscopy images of FLAG-tagged bioPROTAC localization in HEK293T/17 cells 24 h after transient transfection. Images are representatives of results observed in single cells from two replicate experiments. Arrowheads indicate nuclear speckles where SHP2 appears to colocalize with SPOP. Statistics: Fraction of SHP2-sfGFPxK remaining compared by one-way ANOVA to the corresponding untargeted negative controls (monobody.mut-SPOP) using Dunnett’s multiple comparisons test. ∗∗∗∗p < 0.0001. Replicate samples in panels B and C are technical replicates. bioPROTAC, biologic PROTAC; sfGFP, superfolder GFP; SPOP, speckle-type pox virus and zinc finger protein; xK, lysine free.

We first investigated the impact of SPOP dimerization on SHP2 degradation. We mutated residues on SPOP required for dimerization of the BTB (L186D, L196D, L193D, and I217K) or BACK (Y353E) domains alone (notated as SPOPxBTB and SPOPxBACK) or in combination (notated as SPOPxBTBxBACK or as SPOPxBB) and expressed these mutants as CS3-SPOP bioPROTACs (Fig. 3B). We observed no significant SHP2-sfGFPxK degradation from SPOPxBTB or SPOPxBTBxBACK whereas SPOPxBACK achieved a 26% reduction in SHP2-sfGFPxK expression compared to the negative control. This is in contrast to nearly 82% reduction of SHP2-sfGFPxK for the fully functional CS3-SPOP construct, indicating that SPOP dimerization is favorable for SHP2 degradation. This is in agreement with previous studies, which found that inhibiting dimerization of SPOP abrogated degradation of H2B-GFP (18), indicating that SPOP dimerization may broadly benefit degradation of different targets. We also expressed SPOP with the 3Box region removed to inhibit interactions with Cul3 and effectively prevent ubiquitination (Fig. 3B). As expected, we did not observe significant degradation from SPOPx3box, which also validates that degradation is SPOP-mediated and is the result of proteasome dependent activity.

Next, we evaluated the impact of nuclear localization of CS3-SPOP on SHP2 degradation. In previous studies, Ju Shin et al. used vhhGFP4-SPOP to target multiple GFP-tagged proteins for degradation and found that removing the nuclear localization sequence (xNLS) from SPOP had minimal impact on degradation of targets localized to the nucleus, presumably because dimerization of vhhGFP4-SPOPxNLS with endogenous SPOP still indirectly promoted nuclear localization (18). In contrast, Wang et al. found that removal of the SPOP nuclear localization sequence enhanced degradation of the microtubule-associated protein tau by enabling better colocalization of the degrader and target in the cytosol (20). Taken together these studies indicate that the impact of SPOP localization on degradation is target specific and likely depends on target localization and the target’s ability to translocate to the nucleus. Notably, SHP2 is widely expressed in the cytoplasm of most cell types but is also reported to translocate to the nucleus (10, 31). To determine how SPOP localization impacts SHP2 degradation, we removed the nuclear localization sequence from SPOP on CS3-SPOP and measured degradation in our SHP2-sfGFPxK assay (Fig. 3C). We found that degradation efficiency was reduced from 82% degraded by CS3-SPOP to only 24% degraded by CS3-SPOPxNLS. This indicates that SPOP nuclear localization is favorable for SHP2 degradation by CS3-SPOP.

To better understand the impact of SPOP dimerization and localization on SHP2 degradation, we used fluorescent microscopy to visualize how SPOP bioPROTAC constructs were distributed throughout the cell (Fig. 3D). To achieve this, we transiently cotransfected bioPROTACs with full length HA-SPOP that could dimerize and colocalize with endogenous SPOP as shown previously for efficient labeling of SPOP localization (30). We validated that the additional overexpression of SPOP had minimal impact if any on the overall degradation occurring in the cells (Fig. S4). We found that CS3-SPOP colocalized with HA-SPOP in the nucleus, quantification of which indicated more than 90% of both proteins were colocalized (Fig. S5). It also appeared that a portion of SHP2 colocalized in nuclear speckles, presumably induced by CS3-SPOP recruitment of SHP2 and translocation to the nucleus. Notably, a considerable amount of SHP2 was still present in the cells observed, likely due to the shorter incubation time (24 h posttransfection compared to 48 h for previous assays). When the BTB and BACK dimerization domains of SPOP were mutated, the bioPROTAC and SHP2 no longer appeared to colocalize in the nucleus with HA-SPOP, and CS3-SPOPxBTBxBACK was expressed diffusely around the cell. Additionally, the nuclear speckles observed for CS3-SPOPxBTBxBACK were visibly smaller, which is in agreement with prior observations that smaller speckles result from less SPOP present in the nucleus (30). The nontargeted CS3.mut-SPOP construct showed colocalization with HA-SPOP in nuclear speckles but did not appear to induce SHP2 translocation to the nucleus. Finally, we evaluated CS3-SPOPxNLS, which had some cytosolic expression but still appeared to largely localize to the nucleus, presumably driven by dimerization with HA-SPOP and endogenous SPOP that still contained localization sequences. The nuclear speckles observed in the CS3-SPOPxNLS treated cells also appeared smaller indicating that less SPOP accumulated in the nucleus of these cells, which may contribute to the decrease in SHP2 degradation observed in the SHP2-sfGFPxK assay. Overall, these images contribute to our understanding of how SPOP-based SHP2 bioPROTACs induce degradation through dimerization and localization to the nucleus.

Monovalent versus bivalent bioPROTACs

We next sought to modify our bioPROTACs to maximize degradation activity. We took the approach of experimenting with increased valency since previous reports demonstrated improved degradation of BRD2 and BRD4 with trivalent small molecule PROTACs (32) and avid bioPROTACs showed heightened activity against EED (6). We created the bivalent SHP2-recruiting SPOP-based SHP2 degrader, CS3-SPOP-NSA5. From our previous studies, we found that CS3 and NSA5 were similarly effective at achieving degradation when linked to either the N terminus or C terminus of SPOP. We made size-matched controls of CS3-SPOP-NSA5 in which one monobody was functional while the other was mutated to test differences between size-matched monovalent and bivalent degraders. We also produced a fully-nontargeted negative control where both monobodies are mutated. When tested in triplicate in the SHP2-sfGFPxK assay, all constructs resulted in a significant decrease in GFP MFI with a slight improvement from the bivalent degrader over the monovalent controls (Fig. 4A). In an attempt to further parse out differences between the bivalent and monovalent constructs, we titrated the amount of functional bioPROTAC plasmid transfected into each cell (Figs. 4B and S6). We found that the bivalent degrader achieved slightly more SHP2-sfGFPxK degradation at low plasmid concentrations compared to monovalent equivalents, indicating marginally improved potency of the bivalent bioPROTAC at the 48-h time point tested. Interestingly, we found that CS3-SPOP-NSA5 degraded 50% of SHP2 with as little as 0.3% of the total plasmid load transfected expressing functional bioPROTAC. These results also illustrate that bioPROTAC titrations are critical for deconvoluting potency and maximum activity; a given level of degradation can be accomplished by many combinations of these two parameters, and so single-concentration results are at best only qualitative threshold indicators for the presence of bioPROTAC activity.Figure 4 Bivalent bioPROTACs achieve increased potency over monovalent.A, triplicate measurements of SHP2-sfGFPxK MFI 48 h after transient transfection in stable line HEK293T/17 cells with multivalent SPOP bioPROTACs and sized match monovalent and nonbinding controls. Data shown are assay replicates. B, titrations of bivalent bioPROTAC plasmids transfected into SHP2-sfGFPxK-expressing cells where “% bioPROTAC” indicates what fraction of the total plasmid payload transfected expresses the bioPROTAC compared to an mCherry only expressing negative control. Results are the average of triplicate measurements. C, MFI of endogenous SHP2 (left, labeled with AF647 measured in the APC channel) and FLAG-tagged bioPROTAC expression (right, labeled with AF405 measured in the BV421 channel) measured by intracellular flow cytometry 48 h after transient transfection of bioPROTACs into WT HEK293T/17 cells. Data shown are assay replicates. D, response curves of recombinant bivalent bioPROTACs and size-matched monovalent and negative controls binding SHP2 as measured by Bio Layer Interferometry (BLI). Kinetic values fit using GraphPad Prism are listed. E, degradation of SHP2-sfGFPxK 48 h after transfection with monovalent (blue) or bivalent (pink and red) bioPROTACs normalized as a fold-change decrease in SHP2-sfGFPxK MFI relative to non-binding negative controls. Statistics: GFP MFI compared by one-way ANOVA using Dunnett’s multiple comparisons test. ∗∗∗∗p < 0.0001. Replicate samples in panels B–D are technical replicates. bioPROTAC, biologic PROTAC; MFI, mean fluorescence intensity; sfGFP, superfolder GFP; SPOP, speckle-type pox virus and zinc finger protein; xK, lysine free.

We next tested the impact of the CS3-SPOP-NSA5 bivalent bioPROTAC on endogenous SHP2 levels (Figs. 4C and S7A). At 48 h after transfection, we detected reduced levels of endogenous SHP2 remaining in all cells treated with active bioPROTACs, both monovalent and bivalent, but not in the nonfunctional bioPROTAC treated cells. We also measured FLAG labeling, indicative of bioPROTAC expression levels, and found that CS3-SPOP-NSA5 had the lowest expression levels but still achieved the most SHP2 degradation, an unexpected result but consistent with other bioPROTACs examined here. We further validated that SHP2 degradation by CS3-SPOP-NSA5 was abrogated by the presence of the proteasome inhibitor MG132, indicating that CS3-SPOP-NSA5 induces proteasome-dependent degradation (Fig. S7B). Taken together, these data indicate that bivalent bioPROTACs may drive increased potency compared to monovalent equivalents.

We examined several mechanisms of action for bivalency improvements to potency (Fig. S8A). First, bivalent degraders could capture two target proteins for ubiquitination at the same time. Second, bivalent degraders with two distinct binding epitopes like CS3-SPOP-NSA5 could bind in cis to the target protein resulting in an avidity that creates increased target affinity resulting in more efficient degradation. Third, CS3-SPOP-NSA5 could cluster SHP2 together and somehow promoting more rapid degradation. To test this third theory, we coexpressed CS3-E3-CS3 and NSA5-E3-NSA5 constructs together, which could induce clustering, but did not see improvements to degradation at similar levels of CS3-SPOP-NSA5 (Fig. S8B).

To test the first two hypotheses, we recombinantly expressed the bivalent target-binding CS3-SPOP-NSA5 degrader and corresponding size-matched monovalent target-binding controls and characterized their binding interactions with SHP2 using BLI (Figs. 4D and S9). Notably, constructs were expressed with the nondimerizing SPOPxBTBxBACK (SPOPxBB) to eliminate any interactions other than that of the monobodies with SHP2. When biotinylated bivalent bioPROTACs were loaded onto the BLI tip, all samples had similar loading response of approximately 1 nm (data not shown). When the loaded BLI tip was incubated with SHP2, we found that the bivalent degrader showed nearly double the amount of binding response compared to the monovalent controls but had similar overall kinetics and affinities. This suggests that the first theory is most consistent with the data, and that the multivalent constructs bind two SHP2 proteins at once.

We next wanted to determine if the improved activity of the multivalent E3 ligases was SPOP specific or was broadly applicable to other bioPROTACs. We chose to test a panel of E3 ligases reported previously (14) to comparatively test monovalent target-binding (monobody-E3 or E3-monobody) versus bivalent target-binding (monobody-E3-monobody) SHP2 bioPROTACs using our SHP2-sfGFPxK assay (Fig. 4E). Interestingly, for several E3 ligases tested (FBW7, SKP2, VHL, pVHL, SPOP, ASB1, SOCS2, and IPAH9.8) we observed improved degradation from bivalent bioPROTACs over equivalent monovalent constructs. In particular, bivalent constructs with two different monobodies, rather than two of the same monobodies, yielded the greatest fold-improvement in SHP2 degradation in the majority of E3 ligases tested.

Curious to see if higher order valency could continue to improve bioPROTAC potency, we engineered multivalent bioPROTACs with four monobodies attached to a single E3 ligase (Fig. S8C). None of the multivalent constructs showed improvements over the bivalent equivalents, indicating there is a limit to the effectiveness of this valency improvement. Overall, these results indicate that bivalent bioPROTACs show improved potency and degradation, and that valency should be considered as a strategy for future bioPROTAC development.

Testing bioPROTAC activity against mutant SHP2

Having validated that CS3-SPOP-NSA5 is a highly potent SHP2 degrader, we assessed its potential as a cancer therapeutic. SHP2 has a multifaceted role as a target. Firstly, SHP2 signals upstream of the oncogenic RAS/mitogen-activated protein kinase (MAPK) pathway, a well-established contributor to tumor cell growth and proliferation (33). Small molecule inhibitors of SHP2, such as SHP099, bind to an allosteric pocket on SHP2 that lock the phosphatase in an autoinhibited confirmation (34). Using allosteric inhibitors as SHP2-recruiting ligands, several small molecule PROTACs, such as SHP2-D26 (24), have also been developed (35, 36, 37). In some cancers, however, SHP2 acquires gain-of-function mutations that contribute to overactivation of the MAPK signaling (38). Moreover, SHP2 gain-of-function mutants typically undergo a conformational change that significantly reduces the effectiveness of allosteric ligand binding (39, 40, 41). This means that small molecules, both allosteric inhibitors and PROTACs, are relatively ineffective against SHP2 mutants (Fig. 5A). Conversely, our bioPROTACs utilize monobodies to recruit SHP2 for degradation by binding to SHP2 in areas unaffected by the gain-of-function mutations (Fig. 5A). Thus, we hypothesized that bioPROTACs would be more effective against SHP2 mutants than existing small molecule therapies.Figure 5 BioPROTACs outperform small molecule PROTACs for degrading SHP2 mutants.A, schematic of allosteric small molecule PROTACs binding WT SHP2 but not effectively binding SHP2 with gain-of-function mutations vs bioPROTACs which can bind both. B, activity assay of recombinant SHP2.WT and SHP2.E76K measured by fluorescence activation of DiFMUP substrate. Results are the averages of triplicate measurements. C, schematic of components included in an in vitro ubiquitination assay. D and E, anti-SHP2 western blots of samples from in vitro ubiquitination assays of SHP2.WT and SHP2.E76K coincubated for 1 h with small molecule PROTAC SHP2-D26 with 1 μM of VHL (D) or recombinant bioPROTACs (E). F, schematic of MC38 cells which contain SHP2 with a G503V activating mutation that acts upstream of pERK in the MAPK pathway. G and H, fraction of SHP2 remaining (G) and fraction of cells pERK+ (H) in MC38 cells 48 h after treatment with small molecule inhibitors and PROTACs or transient transfection with bioPROTACs and nondegrading controls as measured by intracellular flow cytometry. Fraction of SHP2 is normalized such that 100% corresponds to MFI from untreated cells and 0% is defined as MFI from cells stained with a concentration matched isotype control. Data shown are assay replicates. I, representative histograms of pERK staining in MC38s from H. Statistics: fraction of cells compared by one-way ANOVA using Dunnett’s multiple comparisons test. ∗p < 0.05, ∗∗∗∗p < 0.0001. Replicate samples in G-H are technical replicates. bioPROTAC, biologic PROTAC; MAPK, mitogen-activated protein kinase; MFI, mean fluorescence intensity; PROTAC, proteolysis-targeting chimera; p-ERK, phospho-extracellular signal-regulated kinase; SHP2, src-homology 2 containing protein tyrosine phosphatase-2.

To test our hypothesis, we evaluated the ability of each type of PROTAC, both small molecule and biologic, to induce SHP2 ubiquitination, a prerequisite for proteasomal degradation. We started by recombinantly expressing the well-characterized gain-of-function mutant SHP2.E76K (40, 42). To validate that the gain-of-function mutant had the expected activating conformational change, we tested SHP2.E76K in comparison with SHP2.WT in a phosphatase activity assay using a probe, DiFMUP, that fluoresces upon dephosphorylation (Fig. 5B). We found that SHP2.E76K rapidly converted DiFMUP to DiFMU consistent with a gain-of-function mutation, while SHP2.WT was much less active, indicating it occupies the expected autoinhibited conformation. Having validated our target antigens, we next assessed how efficiently each PROTAC induced ubiquitin chain formation on SHP2.WT and SHP2.E76K using an in vitro ubiquitination assay, which combines all of the necessary components for ubiquitination as a mock-cellular environment (Fig. 5C). Notably, SHP2-D26 recruits VHL, a different E3 ligase than SPOP, which could contribute to differences between assays. We found that the small molecule PROTAC SHP2-D26 resulted in substantial ubiquitin chain formation on SHP2.WT but less so on SHP2.E76K, as predicted (Fig. 5D). For bioPROTACs, we observed that CS3-SPOP-NSA5 resulted in a very high degree of ubiquitin chain formation on both SHP2.WT and SHP2.E76K (Fig. 5E). Notably, the SPOP used for the expression of recombinant bioPROTACs was the non-dimerizing SPOPxBB, thus the bioPROTACs are not benefiting from SPOP oligomerization effects. Taken together, these results suggest that SHP2 bioPROTACs may be more effective than small molecule PROTACs against SHP2 gain-of-function mutants. Additionally, we found that CS3-SPOP-NSA5 resulted in more ubiquitin chain formation on SHP2.WT compared to incubation with size-matched monovalent bioPROTACs. This agrees with our findings that bivalent target-binding bioPROTACs are more potent than monovalent, seemingly by recruiting two target proteins and forming ubiquitin chains more processively.

Characterizing antitumor effects of SHP2 bioPROTACs

We next sought to validate our hypothesis that bioPROTACs induce potent effects in the context of cancer. We chose to use the MC38 murine adenocarcinoma cell line as proof of concept because the MC38 tumor cell line expresses SHP2.G503V (43), an alternative gain-of-function SHP2 mutant with a similar conformational change as SHP2.E76K (Fig. 5F). Notably, we did not predict any issues translating from human to murine cells given that SHP2 shares 99% sequence homology between these species, and none of the differing amino acids were predicted to interfere with ligand-binding. Additionally, murine and human SPOP are 100% identical and the same VHL-recruiting ligand used in SHP2-D26 achieved targeted degradation of BCL-XL in MC38s (44).

To compare biologics and small molecules, we utilized SHP099 as an allosteric inhibitor, SHP2-D26 to represent small molecule PROTACs, and our highly potent CS3-SPOP-NSA5 bioPROTAC. Notably, NSA5 and CS3 monobodies were developed as inhibitors of the C-SH2 and N-SH2 domains of SHP2, respectively, and were shown to reduce pERK signaling (8). Thus, we wanted to evaluate differences between inhibition by NSA5 and CS3 and degradation by CS3-SPOP-NSA5. To accomplish this, we used CS3-SPOPxBB3-NSA5 as a size-matched construct with inhibition activity from CS3 and NSA5 but no activity from SPOPxBB3, which lacked both BTB and BACK dimerization domains and the 3Box domain. As negative controls, dimethylsulfoxide (DMSO) was used for small molecules and nonfunctional CS3.mut-SPOPxBB3-NSA5.mut was used for biologics.

We first evaluated SHP2 expression in MC38s after treatment with either small molecule or biologic degraders or inhibitors. We found that 1 μM SHP2-D26 significantly induced degradation of SHP2.G503V after 48 h while 1 μM of SHP099 inhibitor did not result in significant changes compared to DMSO controls (Fig. 5G, left). Similarly, the bivalent bioPROTAC CS3-SPOP-NSA5 significantly reduced SHP2 expression while the size-matched inhibitor CS3-SPOPxBB3-NSA5 did not result in any changes compared to the fully nonfunctional bioPROTAC CS3.mut-SPOPxBB3-NSA5.mut (Fig. 5G, right). Notably, although degradation of SHP2.G503V was significant for both the small molecule and bioPROTAC constructs, approximately 46% of SHP2.G503V remained after incubation with SHP2-D26 whereas 12% remained for CS3-SPOP-NSA5. These data suggest that bioPROTACs may be more effective at degrading SHP2.G503V. However, due to differences in the expression levels of the different E3 ligases and the unknown concentrations of bioPROTACs expressed in the cell, a direct comparison between bioPROTACs and small molecule PROTACs cannot be drawn. Furthermore, we validated SHP2-D26 degradation of murine SHP2 by Western blot in multiple other murine cell lines, including the 4T1 murine breast cancer cell line and RAW 264.7 macrophages, indicating that SHP2-D26 is more active in other murine cells and that the reduced effects observed in the MC38 cell line are likely due to the SHP2.G503V mutation (Fig. S10, A–C). We also observed robust SHP2 degradation from bioPROTACs in the 4T1 and RAW 264.7 cell lines (Fig. S10, D–F).

Finally, to test the effects of treatments on MC38 cells, we evaluated how SHP2.G503V degradation or inhibition impacted pERK expression as a proxy for MAPK pathway activation in response. We found that SHP2-D26 and SHP099 reduced expression to 33% and 31%, respectively, as a fraction of all cells positive for pERK compared to DMSO treated cells which were 44% pERK+ (Fig. 5, H and I). When treated with biologics, pERK expression was significantly reduced to only 9% of cells (Fig. 5, H and I). Interestingly, the functional bioPROTAC, CS3-SPOP-NSA5, and the size-matched inhibitor, CS3-SPOPxBB3-NSA5, both showed similar degrees of pERK attenuation at this time point; however a control in which CS3 and NSA5 were coexpressed but not linked was somewhat less effective at reducing pERK in MC38 cells, indicating there is benefit to degradation and bivalent binding (Fig. S11). Overall, these data suggest that bioPROTACs are effective at downregulating MAPK signaling in tumor cells, even in cases where oncogenic gain-of-signal SHP2 mutants are present, and show potential advantages compared to complementary small molecules.

Discussion

Targeted protein degradation is a valuable tool with broad applications for mechanistic investigations and therapeutic development. Strategies to identify degraders and characterization of degrader potency is an active area of research. In this work, we reported a novel lysine-free sfGFP, which we used to establish a loss-of-signal assay for identifying proteasomal targeted degradation constructs with a presumed reduced risk of false positives from off-target ubiquitination of lysine residues on GFP. We validated our SHP2-sfGFPxK assay to be robust and suitable for quantifying targeted degradation; however, future work is needed to fully characterize the sfGFPxK assay platform’s ability to prevent off-target ubiquitination and false positives. None of the constructs identified in our work were selective for SHP2-GFP over endogenous SHP2, likely because SHP2 is more than twice the size of GFP, making it a relatively easy target to ubiquitinate. This is unlike KRAS, which is smaller than GFP and has previously reported to suffer from of off-target ubiquitination (9). Nonetheless, we theorize that using the lysine-free GFP system generates results that better predict endogenous degradation.

In addition to developing new tools to aid in degrader discovery, we evaluated how different characteristics of bioPROTACs contributed to degradation efficiency of SHP2. Our findings suggest that bioPROTACs benefit from higher affinity interactions with their target, typically in the low nanomolar range, in agreement with prior reports for other targets using bioPROTACs (6, 7, 27) but in contrast to findings for small molecules that low affinity can be sufficient (26). Notably, the intracellular concentrations of the constructs tested were not known and thus were not considered in this analysis, the impact of which could be assessed in future work. It remains unclear if these observations are target specific, E3 ligase specific, or if there is a KD below which degradation is no longer improved or an optimal kinetic profile rather than binding affinity, all of which could be explored in future studies.

Interestingly, the posttransfection expression levels of the bioPROTAC constructs tested here appeared to range considerably yet similar levels of SHP2 degradation were achieved. It is reasonable that some of the bioPROTACs may get ubiquitinated and degraded along with the target protein, meaning that they are not leveraging maximal catalytic turnover and reuse, which is considered to be a primary advantage of PROTACs. Further studies are needed to better characterize how to optimize this property of bioPROTACs, perhaps through removal of accessible lysine residues on the surface of the bioPROTAC using similar strategies as those used for engineering lysine-free GFP.

We contribute here to the growing understanding of SPOP-based bioPROTACs by characterizing how SPOP’s multifaceted interactions affect degradation of SHP2. We find that SPOP dimerization promotes efficient degradation of SHP2, in agreement with reports from others (18). Given the sequestered and phase-separated nature of SPOP oligomers, we theorize that SPOP is successful at degrading diverse target proteins because, when a target is colocalized with an SPOP oligomer, said target may be surrounded by an abundance of UPS components capable of inducing ubiquitination from multiple angles, negating the need for maximized cooperativity as proposed for most other PROTACs.

Unlike dimerization of SPOP, which appears to be widely beneficial for different targets, a requirement for nuclear localization of SPOP seems to be target specific. For SHP2, despite being primarily a cytosolic protein, maintaining nuclear localization of SPOP in the SHP2-bioPROTAC was favorable. We observed that SHP2 appeared to translocate to the nucleus and colocalize in SPOP-rich nuclear speckles. However, other studies indicate that removal of the nuclear localization sequence from SPOP may be a viable strategy to improve degradation of targets incapable of translocating to the nucleus, although it is unclear if targets relying on cytosolic SPOP degradation still benefit from any SPOP oligomerization outside of the nuclear speckles.

As a seemingly broad strategy not specific to any particular E3 ligase, we found that increasing the valency of bioPROTACs, such that each E3 ligase can recruit two target proteins, one on either polypeptide terminus, promoted a greater degree of degradation. At longer time points, 48 h post transfection in multiple cell types, we observed only small differences in SHP2 degradation between the bivalent CS3-SPOP-NSA5 construct and sized-matched monovalent controls. However, from in vitro ubiquitination assays, we observed that considerably longer ubiquitin chains were formed on SHP2 by bivalent CS3-SPOP-NSA5 within a one-hour time period, indicating an advantage for bivalent bioPROTAC activity at shorter time points and suggesting a more rapidly induced degradation. Interestingly, we observed a greater degree of degradation from bivalent constructs with two different target-recruiting ligands compared to those with the same ligands across E3 ligases, and there were also differences in degradation observed between E3 ligases. Our data did not suggest cis-binding or avidity promoting these interactions, thus why heterobivalent target recruitment appeared to be superior to homobivalent target recruitment remains unclear. We did not observe any indications that multivalency or avidity from constructs with four SHP2-recruting domains was more beneficial than bivalent constructs with two SHP2-recruiting domains. Notably, for all multivalent constructs, steric interactions that differently orient the target protein or sterically hinder it from binding altogether may contribute to the differences in degradation that we observed. Future studies are needed to assess this. In this work, we focused our efforts on characterizing bivalent SPOP-based SHP2 bioPROTACs but future research into bivalent degraders using different E3 ligases against different targets would be beneficial. Considering these findings, we suggest that valency be considered as a strategy for future bioPROTAC development.

We demonstrated that bioPROTACs are able to robustly target gain-of-function SHP2 mutants for degradation and result in reduced MAPK signaling in MC38 tumor cells, more so than their small molecule counterparts. These results highlight the utility of bioPROTACs for cases where specificity against a given target is not sufficiently achieved by small molecules. However, we observed similar pERK signaling reduction for degraders as well as inhibitors; thus, it is unclear if using biologics for SHP2 inhibition alone is sufficient for therapeutic effect or if degradation of SHP2 with bioPROTACs is superior. Future studies are needed to parse out if there are advantages to bioPROTACs from either more sustained impacts on pERK signaling at later time points or from improved effectiveness at lower expression levels. Future studies are also needed to further characterize if SHP2 degradation by bioPROTACs translates to antitumor efficacy in an in vivo setting, as has been shown for small molecule SHP2 PROTACs (45). SHP2 KO or inhibition is also reported to have therapeutic benefit in other cell types beyond tumor cells, such as macrophages and T-cells (43, 46). Future efforts to characterize bioPROTACs in these other cell times would broadly add to our understanding of the potential of targeted SHP2 degradation therapies.

In summary, this research highlights a specific target for which bioPROTACs excel, and informs on design strategies for effective bioPROTACs for improved therapeutic outcomes.

Experimental procedures

Stable line generation

GeneBlocks of double-stranded DNA encoding for murine SHP2 and superfolder GFP were purchased from Integrated DNA Technologies (sequences in SI). SHP2 and GFP inserts were cloned into the lentiviral transfer plasmid pLJM1 (Addgene #19319, a gift from David Sabatini) using In-Fusion ligation (Takara Bio). Stable lines were achieved by lentivirus transduction into HEK293T/17 cells (American Type Culture Collection # CRL-11268), as previously described (14). Briefly, HEK293T/17 cells were transduced with lentivirus particles to generate cells with stable SHP2-sfGFPxK expression. Transduced cells were isolated using a combination of selective media containing 2 μg/ml puromycin and single cell sorting to isolate and expand clonal populations of cells expressing SHP2-sfGFPxK. All cell lines used were annually tested for mycoplasma and confirmed negative.

BioPROTAC plasmid preparation

BioPROTACs were cloned into pIRES2 backbone (Clontech) coexpressed with mCherry, as previously described (14). Bivalent bioPROTACs were cloned using NEBridge Golden Gate Assembly Kit BsmBI-v2 (NEB) to ligate monobody DNA fragments, amplified by CloneAmp HiFi PCR Premix (Takara Bio) and isolated using the NucleoSpin Gel and PCR Clean-Up Kit (Takara Bio), into existing monovalent bioPROTAC backbones. Amino acid sequences for protein components available in the supporting information.

Western blots

Cells were prepared for Western blot using Cellstripper (Corning) to remove cells from the plate. To evaluate bioPROTACs by Western blot in HEK293T/17 cells, 10 cm dishes of cells seeded overnight to approximately 70% confluency were transfected with 1.5 ml of Opti-MEM (Gibco) with plasmid DNA diluted to 10 μg/ml with 3.5 μl of TransIT-293 transfection reagent (Mirus Bio) per 1 μg of DNA. For cells transiently transfected with bioPROTACs, approximately one million mCherry + cells were collected using a Sony MA 900-1 sorter. Cells treated by small molecules (SHP2-D26, Med Chem Express) were all collected for lysis. Cells were washed once with PBS (corning) prior to lysis with 1× radioimmunoprecipitation assay buffer (Abcam) plus 1× Halt Protease Inhibitor (Thermo Fisher Scientific). Lysate was isolated according to the manufacturer’s instructions and the concentration of the lysate was measured by Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Subsequently, 20 μg with 1× Protein Sample Loading Buffer (LI-COR) was loaded into each well of a NuPAGE 4 to 12% Bis-Tris Protein Gel (Invitrogen). Gels were then transferred to a nitrocellulose membrane using an iBlot Gel Transfer Device (Invitrogen) and blocked Intercept TBS Blocking Buffer (LI-COR). Blots were probed for SHP2 (Abcam #Ab187040), β-Actin (Licor #926-42210), and FLAG tag (Sigma-Aldrich #F1804). Proteins were visualized using IRDye680 donkey anti-mouse (LI-COR) and IRDye800 goat anti-rabbit (LI-COR) secondary antibodies. Blots were imaged on a LI-COR Odyssey CLx scanner. Antibodies used were validated by manufacturer.

SHP2-GFPxK assay

SHP2-GFPxK stable HEK293T/17 cells were seeded in a 96-well plate overnight at a density of ∼12,000 cells per well in a volume of 100 μl Dulbecco's modified Eagle's medium (DMEM) + 10% fetal bovine serum (FBS). The following day, cells were transiently transfected with 10 μg L of plasmid DNA diluted to 10 μg/ml in Opti-MEM (Gibco) with 3.5 μl of TransIT-293 transfection reagent (Mirus Bio) per 1 μg of DNA, according to the manufacturer’s instructions. Transfected cells were incubated for approximately 48 h at 37 °C and 5% CO2. Cells were detached from the plate using Cellstripper (Corning) and stained for viability in 3 μM 4′,6-diamidino-2-phenylindole (BioLegend) for 10 min on ice. Cells were washed twice and resuspended in PBS + 1% bovine serum albumin (BSA) for flow cytometry on a BD LSRFortessa Cell Analyzer. For plasmid titrations, the same amount of total plasmid was transfected into each well but the ratio of functional bioPROTAC plasmid and empty plasmid (expressing mCherry only) was modified. All transfections were performed with a minimum of two independent replicate experiments.

Z-score testing

Stable line cells were transfected with replicate wells of bioPROTACs encoding for positive and negative controls. A Z′ score was calculated according to the following equation (25).Z′=1−3σp+3σn|μp−μn|

Recombinant protein expression and purification

Recombinant proteins were prepared as described previously (14). Briefly, all constructs (murine SHP2, monobodies, and bioPROTACs) were cloned into the pE-H6x-SUMO backbone. Proteins were expressed as SUMO-fusions in Rosetta 2 (DE3) Escherichia coli cells (Novagen) and his tag purified from cell lysate using a gravity column with Talon resin (Takara Bio). Proteins were buffer exchanged into PBS (monobodies & bioPROTACs) or TBS (SHP2) using Amicon filters (EMD Millipore). Proteins were incubated with his tagged sumo protease overnight at 4 °C to remove the HIS-SUMO tag. Cleaved proteins were isolated after passing over fresh Talon resin. Protein products were verified to be pure and of the correct molecular weight by running samples on SDS PAGE gels. If necessary, proteins were further purified by size exclusion chromatography on an AKTA fast protein liquid chromatography system (GE HealthCare).

Fluorescent microscopy

Approximately 25,000 WT HEK293T/17 cells were seeded overnight in 300 μl of media in each well of an 8-well Nunc Lab-Tek II Chamber Slide (Thermo Fisher Scientific). Glass chambers were pretreated with poly-L-lysine (Sigma-Aldrich) according to manufacturer’s instructions prior to seeding to improve cell adherence. Cells were cotransfected with plasmids encoding for bioPROTACs and HA-SPOP (full-length) which were each diluted in OptiMEM (Gibco) to a final concentration of 10 μg/ml and combined at a 2:1 ratio. In addition, 3.5 μl of TransIT-293 transfection reagent (Mirus Bio) was added per 1 μg of DNA and 30 μl was transfected into each well of cells. Transfected cells were incubated for 24 h at 37 °C and 5% CO2. Following incubation, media were removed, and cells were washed twice with PBS before fixing with 4% formaldehyde (Electron Microscopy Sciences) for 30 min at room temperature. Cells were then washed twice with PBS and permeabilized for 20 min in perm buffer (PBS + 0.1% TritonX-100 + 5% goat serum). Primary antibodies were diluted into perm buffer and incubated for 2 h at room temperature. Primary antibodies and dilutions: 1:100 rabbit anti-SHP2 (Abcam #300579), 1:250 rat anti-HA (Roche #11867423001), 1:500 mouse anti-FLAG (Sigma-Aldrich, F1804). Wells were washed three times with PBS followed by a 1 h incubation at room temperature protected from light with secondary antibodies diluted in perm buffer. Secondary antibodies and dilutions: 1:1000 goat anti-rabbit AF647 (Invitrogen #A32733), 1:1000 goat anti-rat AF555 (Invitrogen #A48263), 1:1000 goat anti-mouse AF488 (Invitrogen #A32723). Cells were washed three times with PBS. Cells were stained with 300 nM 4′,6-diamidino-2-phenylindole (BioLegend) for 5 min at room temperature protected from light followed by three more washes with PBS. Chambers were removed and cover slips were applied with SlowFade Diamond (noncuring) mounting reagent (Invitrogen). Cells were kept at 4 °C protected from light prior to imaging with a Leica SP8 scanning confocal microscope. Images were processed with Fiji (https://imagej.net/software/fiji). Transfections and imaging were performed in two independent replicate experiments, and representative high-quality images were chosen for publication. Antibodies used were validated by the manufacturer.

In vitro ubiquitination assays

All proteins necessary to induce ubiquitination were purchased from R&D Systems and combined in a PCR tube to a final volume of 10 μl. All assays received 1× ligase buffer (#B-71), 10 mM MgATP (#B-20), 100 nM of Ube1 (#E-305), 100 μM of ubiquitin (#U-100H) and 2 μM of SHP2 substrate (either SHP2.WT or SHP2.E76K). For assays with SHP2-D26 (Med Chem Express), 1 μM of the small molecule PROTAC was added along with 1 μM of Ubch5a (#E2-616) and 1 μM of VHL (#E3-655). For assays with bioPROTACs, 1 μM of bioPROTAC protein was added along with 1 μM of Ubch5b (#E2-622) and 1 μM of CUL3 (#E3-436). Mixtures were incubated at 37 °C for 1 h followed by addition of 20 μl of water plus 10 μl of Protein Sample Loading Buffer (LI-COR) and a 10 min incubation at 100 °C. Subsequently, 10 μl of diluted assay mixture was run on Western blot as described above. A minimum of two replicate experiments were performed for each assay.

SHP2 activity assays

Recombinant SHP2.WT and SHP2.E76K were diluted to a working concentration of 6.66 nM in assay buffer (60 mM HepespH 7.2, 75 mM NaCl, 75 mM KCl, 1 mM EDTA, and 0.05% Tween-20). DiFMUP (Invitrogen) was titrated from 375 μM down to 0 μM. A total of 15 μl of diluted protein and 10 μl of diluted DiFMUP were added to each well of a black 384 well plate for a final SHP2 concentration of 4 nM with DiFMUP concentration ranging from 0 to 150 μM. A standard curve of DiFMUP substrate and DiFMU product were added to the plate. Immediately after combining reagents in the wells, the florescence (excitation 360 nm & emission 460 nm, optimal gain) of the plate was measured every 2 minutes for 15 min using a Tecan M200 plate reader. Two replicate experiments were performed for this assay.

SHP2 and FLAG detection by intracellular flow cytometry

For intracellular staining of SHP2 and FLAG, cells were detached from wells using Cell Stripper (Corning) and washed once with PBS (Corning) before viability staining with LIVE/DEAD Fixable Near IR Stain (Invitrogen) according to the manufacturer’s protocol. Cells were then washed twice in PBSA (PBS + 1% BSA) and fixed with BD Cytofix for 30 min at 4 °C. Cells were then washed into permeabilized with eBioscience Permeabilization Buffer (Invitrogen) and allowed to incubate for 20 min at room temp. Primary antibodies, rabbit anti-SHP2 (Abcam #300579, 1:250) and mouse anti-FLAG (Sigma-Aldrich #F1804, 1:400), were diluted into permeabilization buffer and incubated with the cells for 1 h at room temperature. A rabbit isotype control (Cell Signaling Technology #3900S) was used at a matched molar concentration to the rabbit anti-SHP2 antibody for approximate quantification of background labeling. Cells were washed twice with permeabilization buffer prior to a 30 min room temperature incubation with secondary antibodies, goat anti-rabbit AF647 (Invitrogen #A32733) and goat anti-mouse AF405 (Invitrogen #A48255), at a 1:200 dilution. After two final washes in permeabilization buffer, cells were resuspended in PBSA and analyzed by flow cytometry on a BD FACSymphony A3 Cell Analyzer. A minimum of two replicate experiments were performed for each assay. Antibodies used were validated by manufacturer.

Phospho-ERK detection by intracellular flow cytometry

Approximately 50,000 MC38 cells were seeded in a 48-well plate in 300 μl DMEM + 10% FBS culture media. MC38s were transiently transfected with 30 μl of transfection mixture containing plasmid DNA encoding for bioPROTACs diluted to 10 μg/ml in Opti-MEM (Gibco) with 3.5 μl of Lipofectamine 2000 transfection reagent (Invitrogen) per 1 μg of DNA, according to manufacturer’s instructions. Alternatively, cells were treated with small molecules (SHP2-D26 and SHP099, both from Med Chem Express) at a final concentration of 1 μM of the compound and 0.1% DMSO. Cells were incubated at 37 °C for 48 h. Prior to fixation, treatment media was removed and fresh DMEM + 10% FBS supplemented with 10 ng/ml murine epidermal growth factor (BioLegend) was added to the cells and allowed to incubate at 37 °C for 10 min. Working as quickly as possible, media were removed, cells were detached from the plate with Cellstripper (Corning), and cells were fixed with prewarmed Cytofix buffer (BD Biosciences) for 10 min at 37 °C. Cells were washed once with PBS and then permeabilized with ice cold Phosflow Perm Buffer III (BD Biosciences) for 30 min on ice. Cells were washed twice with staining buffer (1× PBS, 1% FBS, and 0.09% sodium azide). Cells were then stained with Alexa Flour anti-ERK1/2 Phospho (Tyr202/Tyr204) antibody (BioLegened #369507) at a concentration of 5 μl per reaction for 1 h at room temperature. Cells were washed twice with staining buffer and evaluated by flow cytometry on a BD FACSymphony A3 Cell Analyzer. Two replicate experiments were performed for this assay. Antibodies used were validated by manufacturer.

Bio-layer interferometry

After purification, aliquots of bivalent bioPROTACs and SHP2 were biotinylated with Ez-Link Sulfo-NHS-Biotin (Thermo Fisher Scientific) according to the manufacturer’s protocols. Biotinylated proteins were diluted into running buffer (1X PBS, 0.1% BSA, 0.02% tween-20, and 0.05% sodium azide). For the alanine scan of mutant monobodies, biotinylated SHP2 was loaded onto streptavidin tips and incubated with varying concentrations of monobodies. Run parameters were baseline 60 s, loading 360 s, baseline2 120 s, association 180 s, and dissociation 360 s. For the binding assay to detect interactions of bivalent bioPROTACs with SHP2, the bioPROTACs were loaded onto streptavidin tips and incubated with various concentrations of SHP2. Run parameters were baseline 300 s, loading 150 s, baseline2 120 s, association 300 s, and dissociation 600 s. No ligand and no analyte reference well controls were used to normalize the data. Data were aligned to baseline and the Octet Analysis Studio software (Sartorious, www.sartorius.com) and Prism (GraphPad, www.graphpad.com) were used to asses global curve fits.

Statistics

GraphPad Prism was used to perform all statistical tests. Statistical significance and specific tests are indicated in figure captions.

Data availability

All data and methods used to support the conclusions of this study are available in the main text, figures, and supplementary materials.

Supporting information

This article contains supporting information. Additional methods, figures, protein sequences, and raw Western blot images can be found in the supporting information.

Conflict of interest

K. D. W. is a founding shareholder and consultant for 76Bio, Inc, which develops bioPROTACs for therapeutic applications. The other authors do not have a conflict of interest with the contents of the article.

Supporting information

Supporting information

Acknowledgments

We thank the Swanson Biotechnology Center Koch Institute Core facilities, especially the Flow Cytometry and Microscopy cores, for use of their instrumentation and technical support. We thank members of the Wittrup lab and MIT community for productive discussions and feedback related to this work. Thank you to the Ludwig Center at MIT's Koch Institute for support.

Author contributions

M. H., D. K., and K. D. W. conceptualization; M. H. and K. D. W. methodology; M. H. validation; M. H. investigation; M. H. formal analysis; M. H. writing–original draft; D. K. and K. D. W. writing–review and editing; K. D. W. supervision; K. D. W. funding acquisition.

Funding and additional information

Funding for this work was provided by the Ludwig Center at MIT's Koch Institute and NIH EB031082. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
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