Quantitative Live-Cell Kinetic Degradation and ... - ACS Publications

Articles Cite This: ACS Chem. Biol. XXXX, XXX, XXX−XXX

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Quantitative Live-Cell Kinetic Degradation and Mechanistic Profiling of PROTAC Mode of Action Kristin M. Riching,† Sarah Mahan,† Cesear R. Corona,‡ Mark McDougall,‡ James D. Vasta,† Matthew B. Robers,† Marjeta Urh,† and Danette L. Daniels*,† †

Promega Corporation, 2800 Woods Hollow Road, Madison, Wisconsin 53711, United States Promega Biosciences Incorporated, 277 Granada Drive, San Luis Obispo, California 93401, United States

‡

ACS Chem. Biol. Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 09/01/18. For personal use only.

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ABSTRACT: A new generation of heterobifunctional small molecules, termed proteolysis targeting chimeras (PROTACs), targets proteins for degradation through recruitment to E3 ligases and holds significant therapeutic potential. Despite numerous successful examples, PROTAC small molecule development remains laborious and unpredictable, involving testing compounds for endpoint degradation activity at fixed times and concentrations without resolving or optimizing for the important biological steps required for the process. Given the complexity of the ubiquitin proteasomal pathway, technologies that enable real-time characterization of PROTAC efficacy and mechanism of action are critical for accelerating compound development, profiling, and improving guidance of chemical structure−activity relationship. Here, we present an innovative, modular live-cell platform utilizing endogenous tagging technologies and apply it to monitoring PROTAC-mediated degradation of the bromodomain and extraterminal family members. We show comprehensive real-time degradation and recovery profiles for each target, precisely quantifying degradation rates, maximal levels of degradation (Dmax), and time frame at Dmax. These degradation metrics show specific PROTAC and family member-dependent responses that are closely associated with the key cellular protein interactions required for the process. Kinetic studies show cellular ternary complex stability influences potency and degradation efficacy. Meanwhile, the level of ubiquitination is highly correlated to degradation rate, indicating ubiquitination stemming from productive ternary complex formation is the main driver of the degradation rate. The approaches applied here highlight the steps at which the choice of E3 ligase handle can elicit different outcomes and discern individual parameters required for degradation, ultimately enabling chemical design strategies and rank ordering of potential therapeutic compounds.

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which could be used as recruiter molecules for PROTAC design.4,5,12,14,16−20 Of those, currently only two proteins, von Hippel Lindau (VHL) and Cereblon (CRBN), both of which are substrate adaptor components of larger E3 Cullin-RING complexes, have shown significant and broad success against a diverse set of targets.14,17,21−31 Chemical conversion of previously characterized inhibitors to PROTAC degradation compounds have shown therapeutic advantages, including improved potency and prolonged pharmacodynamics,14,21,23,24,28,30,32−35 and in some cases have unexpectedly introduced specificity not observed in the parental pan-selective inhibitor.26,30,36,37 Despite these early successes, multiple challenges still exist in the functional characterization of PROTACs, including assessment of cellular permeability, ternary complex formation (target:PROTAC:E3 ligase component), functional ubiquitination, and degradation. To optimize the likelihood of finding an active compound, an extensive chemical series is typically developed with two main

argeting key drivers of disease for loss or degradation has been a desirable outcome for numerous therapeutic treatments and is often achieved indirectly by modulation of upstream signaling pathways, transcriptional programs, and/or epigenetic events.1−3 The first examples of compounds affecting degradation were demonstrated many years ago using small molecules or peptides to bridge interactions between a target protein with components of the ubiquitin proteasomal system (UPS).4−8 More recently, the approach has been refined for efficacy and efficiency, comprising a heterobifunctional compound termed proteolysis targeting chimera (PROTAC), also known as SNIPER or degronomid.2−6,9−15 Chemically, these compounds consist of a small molecule target binder or inhibitor on one side which is chemically linked to an E3 ligase recruiter compound on the other side1,2 (Figure 1A). PROTACs induce degradation by simultaneously binding the target protein and the E3 ligase complex proteins, bringing the target protein into proximity for ubiquitination and targeting it for degradation through the UPS1,2 (Figure 1A). While there are hundreds of E3 ligases in eukaryotes and hundreds more proteins involved in active E3 ligase complexes, very few have known binding compounds © XXXX American Chemical Society

Received: July 25, 2018 Accepted: August 23, 2018 Published: August 23, 2018 A

DOI: 10.1021/acschembio.8b00692 ACS Chem. Biol. XXXX, XXX, XXX−XXX

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Figure 1. Generation of HiBiT-tagged endogenous BET family members. (A) Schematic showing strategy for monitoring PROTAC-mediated degradation of endogenously tagged HiBiT-BET family in live cells. HiBiT was inserted via CRISPR/Cas9 to the N-terminus of BET family proteins BRD2, BRD3, and BRD4 in a HEK293 cell line stably expressing LgBiT, which complements with HiBiT to form the luminescent NanoBiT luciferase. Following treatment with PROTACs, protein degradation is monitored by the loss of luminescence. Ternary complex formation or ubiquitination is measured with NanoBRET using the HiBiT-BET protein complemented with LgBiT as an energy donor and HaloTag fused to an E3 ligase component or ubiquitin as the respective energy acceptor. (B) HiBiT blot of CRISPR edited clonal cell lines expressing endogenous HiBiT-BRD2, HiBiT-BRD3, or HiBiT-BRD4. Single isoforms of BRD2 and BRD3 were found, while both short and long isoforms of BRD4 were detected. (C) PROTAC-induced degradation of HiBiT-BET family members after 4 h treatment with 1 μM dBET1 or MZ1. Data are represented as mean RLU values (n = 3) of representative experiments. (D) Bioluminescence imaging using an Olympus LV200 microscope of endogenously tagged HiBiT-BRD4 in LgBiT expressing HEK293 cells treated with 1 μM MZ1 for 2 h.

(NanoBRET)42,43 allows for kinetic measurements of intracellular protein interactions along the degradation pathway such as ternary complex formation, ubiquitination, and PROTAC-target engagement (Figure 1A). In these studies we apply this approach to characterize the cellular mechanism of action of two PROTACs, dBET1 and MZ1, which were shown to degrade bromodomain and extra-terminal (BET) family members BRD2, BRD3, and BRD4. 14,30 Both compounds contain the pan-BET inhibitor JQ1,44 serving as the target-recruiting moiety, conjugated via an amide bond on the same attachment point.14,30 Similar to other BET family PROTACs, both MZ1 and dBET1 demonstrate greater potency as degradation compounds relative to parental BET inhibitors.14,17,30,32,34,35,44−46 These two PROTACs were chosen for this study due to availability, the use of the same JQ1 ligand with identical conjugation but to different E3 handles, and because they are reported to have differential degradation patterns.14,30,37 dBET1, which recruits CRBN through a thalidomide handle, shows equivalent degradation between BRD2, BRD3, and BRD4.14 Recent structural studies with related variants of dBET1 show noncooperative binding

variables: linker composition and the E3 ligase complex recruitment handle.2−4,12,16,19 Compounds are then screened for function by assessing target protein levels ± PROTAC treatment at a given time and various concentrations using end-point techniques such as Western blots or mass spectrometry. These approaches, however, cannot be done in live cells, lack adaptability for high-throughput screening, and are difficult to perform in a quantitative fashion. More importantly, if degradation is not observed in these measurements, they provide no information as to the point of failure or which steps in the process could benefit by compound optimization through further structure−activity relationship (SAR) efforts. In this study, we present a modular live-cell platform to monitor the critical steps of PROTAC mode of action (Figure 1A). At the core of the platform, we combine CRISPR/ Cas938,39 endogenous tagging and luminescent technology40,41 to kinetically measure target protein levels with high precision and without disruption of native expression levels or transcriptional regulation (Figure 1A). Coupling this technology with optimized bioluminescence resonance energy transfer B


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Figure 2. BET family member kinetic degradation profiles and quantitation of degradation parameters. (A) HEK293 cells containing endogenously tagged HiBiT-BET family member and coexpressing LgBiT were treated with an 8-point concentration series of 1 μM dBET1 or MZ1 added at t = 0. Luminescence (RLU) was continuously monitored in 5 min intervals over a 24 h time period in the presence of a stabilized furimazine substrate, Endurazine. Profiles are plotted as mean fractional RLU values by normalizing to DMSO control. Error bars are represented as SEM of n = 3 experiments. (B) Representative schematic and equations to determine degradation parameters: degradation rate, Dmax and time at Dmax. Summary of all degradation parameters tabulated for each BET family member treated with the various concentrations of dBET1 and MZ1 from kinetic degradation profiles in (A). The initial degradation portion of each concentration curve was fit using GraphPad Prism to a one-component exponential decay model (red line), and best fit parameters ƛ (degradation rate) and plateau (minimum remaining fraction) were obtained. The maximum degraded fraction, Dmax, was calculated as 1− plateau. Time at Dmax was defined as the amount of time during which protein levels remained below the threshold plateau + 10% of Dmax. (C) Degradation rate and (D) Dmax values expressed as percent degradation from (B) plotted against PROTAC concentration for each family member and PROTAC. (D) DC50 values were obtained by fitting Dmax values to a variable slope dose−response model in GraphPad Prism. Variability is expressed as SD of the mean from n = 3 experiments. (E) Time at Dmax, as determined by the formula in (B), for HiBiT-BET family members treated with 1 μM dBET1 or MZ1 PROTAC. Data are represented in singlicate taken from the combined (n = 3) kinetic traces.

between BET family members with CRBN.47 In contrast, MZ1, which recruits VHL through a VHL ligand handle,48 shows preferential degradation of BRD4.30 Supporting

structural and biophysical studies indicate cooperative binding between VHL and a bromodomain of BRD4 in the presence of MZ1.37 These structural and biochemical studies yield C


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scope showed highly uniform expression, nuclear localization, and rapid degradation of HiBiT-BRD4, as evidenced by loss of luminescence over 2 h in the presence of 1 μM MZ1 (Figure 1D). Together, these data show expected physiological and PROTAC responses of the endogenously tagged HiBiT-BET proteins, indicating insertion of the HiBiT tag at their respective genomic loci did not impact function. Determination of Degradation Profiles and Quantification of Key Degradation Parameters. To investigate the time-dependent effects of PROTAC-mediated degradation on each BET family member, comprehensive degradation profiles of biological triplicates were generated by monitoring luminescence in 5 min continuous intervals over a 24 h period (Figure 2A). Treatment of HiBiT-BRD2, BRD3, and BRD4 with an 8-point dilution series ranging from 1 nM to1 μM dBET1 or MZ1 showed this concentration window resulted in a broad range of responses from very little degradation to near complete degradation as detected by luminescence (Figure 2A). Strikingly, each family member showed a unique signature, which was both PROTAC- and concentrationdependent (Figure 2A). The degradation profiles in general contained three phases: an initial degradation curve showing loss of the target, a Dmax, and then an eventual upward rise indicating target recovery. Recovery curves were highly variable, often multiphasic, and not quantifiable. Fortunately, other parameters such as initial degradation rate, Dmax, and time at Dmax as depicted in Figure 2B were all amenable for quantitation from these profiles. To determine degradation rate and D max at each concentration, only the starting curve representing the time frame to maximal degradation was considered, as defined in Figure 2B. This initial curve fit a single-component exponential decay model, yielding parameters for cellular degradation rate and Dmax for concentrations of each PROTAC and BET family member summarized in Figure 2B. Trends showed that increasing concentration of PROTAC increased rate of degradation, eventually reaching a plateau at higher concentrations (Figure 2C). This trend indicates that beyond the plateau, concentration of compound is not rate-limiting. Moreover, it is expected that very high concentrations will become inhibitory, consistent with observed hook effects for PROTACs17,30 Degradation rate plots revealed emergence of family member differences, particularly for MZ1 treatment, which showed fast degradation of BRD2 and BRD4, while BRD3 exhibited much slower degradation (Figure 2C). Such family differences were not observed with dBET1, which showed similar rates of degradation for BRD2, BRD3, and BRD4 (Figure 2C). These results demonstrate how not only PROTAC-mediated recruitment to either VHL or CRBN E3 ligase complexes can impact the degradation rate, but also how different family members can show varying rates to the same PROTAC (Figure 2C). Despite having differences in the initial degradation rate, family members were able to achieve similar Dmax if given enough time, and total amount of degradation increased with increasing concentrations of PROTAC (Figure 2A and B). Previous dose response curves (DRCs) to determine cellular DC50 potency of PROTAC compounds have calculated degradation at fixed time points,14,30,37,47 overlooking the time dependency to reach Dmax (Figure 2A). We find that for any given family member at any concentration, the time at which Dmax is reached varies greatly at each concentration (Figure 2A). Therefore, DRCs at fixed times could easily

considerable and meaningful insights into the stability and formation of ternary complexes in vitro but are done with isolated BET bromodomains and limited components of the respective multiprotein E3 ligase complexes.37,47 Full-length BET family members are large proteins, however, each contains two bromodomains capable of separately binding compound.44,45 Therefore, in the cell, positioning of the family members within the E3 ligase complexes could be very different, potentially leading to critical differences in degradation activities as well. Here, we complement the biochemical work and provide additional live-cell context by characterizing the kinetic mechanistic differences mediated via the two possible E3 ligase ternary complexes as well as further understanding of family member specificities. Significant insights into the degradation process and differential modes of action of MZ1 and dBET1 are revealed and also show how these technologies could be applied broadly to enhance understanding of any PROTAC or process which impacts protein homeostasis.

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RESULTS AND DISCUSSION Generation of Endogenously Tagged HiBiT-BET Family Members. Protein overexpression can impart defects on normal protein turnover and homeostasis; therefore, we developed a means to monitor protein levels in real-time without disruption of endogenous expression or regulation. To this end, we used CRISPR/Cas9 genome editing38,39 to append an 11 amino acid peptide, termed HiBiT,41 to the Nterminus of BET family members natively expressed in HEK293 cells, BRD2, BRD3, and BRD4 (Figure 1A and Supporting Information Table 1). The HiBiT peptide, which shows high efficiency for CRISPR insertion given its small size, has pM affinity for and spontaneously complements with the 18 kDa LgBiT protein, forming the luminescent and bright NanoBiT luciferase41 (Figure 1A). LgBiT can be introduced for luminescent detection of HiBiT fusion proteins in a variety of ways, but to enable live cell detection, we performed CRISPR endogenous tagging in HEK293 cells stably expressing LgBiT. CRISPR clonal selection was performed, and genomic analysis by Sanger sequencing revealed homozygous HiBiT allelic insertions for BRD3 and BRD4, and a heterozygous insertion for BRD2 (data not shown). Confirmation of size and expression for each HiBiT-BET family member was confirmed by SDS-PAGE with luminescent blotting as described in the Materials and Methods. Single bands for BRD2 and BRD3 were observed, while both long and short isoforms of BRD4 were detected, as expected given the placement of HiBiT on the N-terminus (Figure 1B). Comparative luminescence expression levels of each family member showed similar expression levels of BRD2 and BRD4, while expression of BRD3 was approximately 60% lower relative to these (Figure 1C and Supporting Information Table 2). End-point analysis of each endogenously tagged HIBiTBET family protein treated with 1 μM dBET1 or MZ1 showed substantial degradation at 4 h (Figure 1C). To show specificity of degradation, HiBiT-BRD4 cells treated with either dBET1 or MZ1 for 3 h were then treated with a 10-fold molar excess of JQ1 (Supporting Information Figure 1). This resulted in increased rates of recovery of BRD4 in the presence of JQ1, the parental competitive binding inhibitor which does not induce degradation (Supporting Information Figure 1). To assess proper localization and response to PROTAC treatment, bioluminescence imaging using an Olympus LV200 microD


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found at both 100 nM and 10 nM treatments (Supporting Information Figures 3A and B). Furthermore, the time at Dmax for each family member followed the trends in family member potency for a particular PROTAC (Figure 2D and E). For example, BRD4 showed greatest MZ1 potency and greatest time at Dmax, while BRD2 had lowest MZ1 potency and shortest time at Dmax, despite having a fast initial degradation rate (Figure 2C, D, and E). These data suggest many factors play a role in time at Dmax and recovery. The finding that BET family members show longer times at Dmax after treatment with MZ1 as compared to dBET1 (Figures 2A, B, and E and Supporting Information Figure 3) support previous reports of reduced cellular compound stability of dBET1.14 The differential family member response however to the same PROTAC (Figure 2A, B, and E and Supporting Information Figure 3) indicate other events are impacting these phases. This is most apparent for BRD2, which shows faster recovery as compared to BRD3 and BRD4 (Figure 2A); therefore, we investigated whether inhibition alone of BET proteins could lead to compensatory changes in BRD2 protein levels. To do this, we performed kinetic analysis of HiBiT-BRD2 after treatment with JQ1 alone. In these studies, we saw a rapid and significant increase in HiBiT-BRD2 levels (Supporting Information Figure 4A), not observed for HiBiT-BRD3 and HiBiT-BRD4 (Supporting Information Figures 4B and C) similarly treated with JQ1, indicating potential feedback on BRD2 synthesis as a consequence of BET inhibition. These results and others that have shown a similar effect on endogenous BRD2 by mass spectrometry49 suggest that the rapid increase in BRD2 following BET family degradation may be due to compensatory feedback mechanisms, and demonstrate how these can outcompete degradation to drive faster recovery of specific family members over others. Kinetic Monitoring of Ternary Complex Formation and Ubiquitination. To relate the degradation profiles to the specific interactions in the degradation pathway, we investigated the kinetics of intracellular ternary complex formation. To do so, we utilized proximity-based NanoBRET technology42 comprising the same endogenously tagged HiBiT-BET family members complemented with LgBiT as luminescent energy donors, and ectopically expressed HaloTag-CRBN or HaloTag-VHL fusions as energy acceptors. (Figure 1A). Due to its ratiometric nature,42 NanoBRET is particularly suited for these studies as loss of the donor (target protein) will not impact the readout of ternary complex formation (Supporting Information Figure 5), and is capable of detecting transient interactions. To first demonstrate ternary complex specificity by NanoBRET, we examined ternary complex formation of ectopically expressed NanoLuc-BRD4 with either HaloTagVHL or HaloTag-CRBN in the presence of either PROTAC. Only the specific pairings of NanoLuc-BRD4 with the respective E3 ligases and PROTACs, showed a dose-dependent increase in ternary complex formation, as measured by NanoBRET (Supporting Information Figure 5A). While detection of ternary complexes within short periods of time (30−60 min) was possible, longer term kinetic analyses were challenging due to the significant loss of the target proteins. To mitigate this loss and study the ternary complex stability over several hours, MG132, a proteasomal inhibitor, was added to prevent degradation, and resulted in a more robust assay window (Supporting Information Figures 5B and C). Because MG132 can be toxic to cells, we tested cell viability by

misrepresent DC50 potency values depending on the time chosen for the measurement. For example, measuring degradation of BRD2 at 2 h would show either PROTAC is potent for degradation, but at 20 h it would appear that neither PROTAC had any effect, because degradation had already occurred and transitioned to recovery (Figure 2A). With these understandings and to remove the impact of time, we sought a more accurate method to calculate DC50 values by determining Dmax at every concentration irrespective of time (Figure 2B and D). This approach reflects and takes into account the actual cellular response and the true capacity for degradation. Comparison of DC50 values calculated in this fashion allowed for rank ordering potency, with MZ1 showing BRD4 > BRD3 > BRD2 and with dBET1, BRD3 > BRD4 > BRD2 (Figure 2D). The window of family member potencies exhibited a broader range with MZ1 (9X) as compared to dBET1 (4X), but significant differences in degradation capabilities only emerged at very low concentrations of PROTAC (Figure 2D), matching previous studies with the endogenous proteins analyzed by Western blotting.30 As other PROTAC cellular studies have utilized ectopic expression of fluorescent proteins to determine compound degradation potencies,47 we wanted to examine whether this approach influenced the kinetic degradation rate and Dmax. To do so, comparative experiments were performed with NanoLuc40 (the parental luciferase of the HiBiT+LgBiT complementation technology) fused to BRD4 and transiently transfected in HEK293 cells. In comparison to the profile for endogenous HiBiT-BRD4 treated with MZ1 in Figure 2A, ectopically expressed NanoLuc-BRD4 treated with MZ1 shows a significantly different degradation signature (Supporting Information Figure 2A). Overall, the degradation window was compressed, showing only a small population of the NanoLucBRD4 was degraded, even at high concentrations of MZ1 (Supporting Information Figure 2A). Determination of the degradation rate showed degradation of the ectopic NanoLucBRD4 was slower (Supporting Information Figure 2B), and the DC50 showed a 30× reduction in apparent potency (Supporting Information Figure 2C). Moreover, the profiles showed more rapid protein recovery, not matching the prolonged time at Dmax observed for endogenously tagged BRD4 (Figure 2A). Because constitutive expression from a non-native promoter lacks the relevant epigenetic and transcriptional control found at the endogenous target loci and ectopic expression will produce excess target protein levels in the cells, degradation profiles cannot be meaningfully interpreted (Supporting Information Figure 2A), and quantitative parameters will be skewed (Supporting Information Figures 2B and C). While the degradation rate and DC50 values are readouts of efficiency and potency, they do not capture the time at Dmax in the cells, which speaks to the time of sustained compound efficacy. To quantify this time frame, we defined the time at Dmax as the period in which protein levels were within a threshold window of 10% of Dmax: (plateau + 0.1(Dmax)) (Figure 2B). Calculation of time at Dmax in this fashion showed this parameter increased with increasing concentration of PROTAC, with a few outliers were observed at the extremely low levels of degradation where the threshold window approached the level of experimental noise (Figure 2B). From this analysis, we found that MZ1 treatment showed longer times at Dmax for all BET family members as compared to dBET1 treatment at 1 μM (Figure 2E), with similar trends E


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Figure 3. NanoBRET ternary complex formation and ubiquitination of BET family members. (A and B) HEK293 cells containing endogenously tagged HiBiT-BET family members and expressing LgBiT were transiently transfected with either HaloTag-VHL or HaloTag-CRBN (A) or HaloTag-Ubiquitin (B), which encodes for the first ubiquitin repeat, amino acids 1−76, of the polyubiquitin-B precursor protein. Forty-eight hour post-transfection, NanoBRET ratios were calculated to measure intracellular ternary complex formation (A) or target ubiquitination (B) in the presence of 1 μM dBET1 or MZ1 (t = 0). Kinetic monitoring of NanoBRET was performed for 6 (A) or 2 (B) h in the presence of both stabilized furimazine substrate, Vivazine, and NanoBRET618 fluorescent ligand. Experiments were also set up for all assays without NanoBRET618 fluorescent ligand as a control and allowed for subtraction of background within the NanoBRET assay, as detailed in the Materials and Methods. Data are represented as fold increase in NanoBRET by normalizing to DMSO control. Variability expressed as SEM from n = 3 experiments. (C) HEK293 cells containing endogenously tagged HiBiT-BET family members and expressing LgBiT were treated at the indicated times with 1 μM dBET1 or MZ1 PROTACs. Cells were lysed with digitonin and incubated for 10 min with both primary polyclonal anti-Ubiquitin and Alexa-594 fluorescent secondary antibody to determine NanoBRET ratios. Data are represented as fold increase in NanoBRET by normalizing to t = 0 time point. Variability expressed as SEM from n = 3 experiments. (D) Degradation rate at 1 μM PROTAC from Figure 2C is plotted against ubiquitination fold increase at the 1 h time point shown in (C).

CellTiter-Glo after 10 μM MG132 treatment over 6 h and found no impact on cell number (Supporting Information Figure 5D). We then performed kinetic analysis of ternary complex formation in the presence of MG132 for 6 h with endogenously tagged HiBiT-BET family members complemented with LgBiT. Complexes between all BET family members and CRBN in the presence of dBET1 showed very rapid association kinetics, peaking at 1 h and then either holding steady or declining (Figure 3A). In contrast, complexes of BET family members with VHL and MZ1 exhibited slower association kinetics, yet continued to increase over time showing longer complex stability (Figure 3A). The prolonged stability of the ternary complexes mediated by MZ1 compared to dBET1 for all BET family members agreed well with the trend that MZ1 treatment resulted in longer time at Dmax (Figures 2A and E), indicating ternary complex stability directly influenced the duration of degradation efficacy. Because our readout of ternary complex formation is only a measurement of binding and not activity of the complex, we measured downstream target ubiquitination. Intracellular ubiquitination kinetics were measured by NanoBRET using the same HiBiT-BET family members as donors, but paired with HaloTag-Ubiquitin as an acceptor. Robust and rapid ubiquitination of BRD2 and BRD4 was observed with MZ1 treatment, while BRD3 showed both slower kinetics and reduced ubiquitination levels (Figure 3B). For BET family

members treated with dBET1, ubiquitination signals were similar in fold change over time and in general slower as compared with MZ1 treatments (Figure 3B). As an orthogonal approach, NanoBRET was also used to probe endogenous ubiquitination of HiBiT-BET family members, utilizing a polyclonal ubiquitin primary antibody and Alexa594-conjugated secondary antibody as an acceptor. These data show identical trends to those observed in the live-cell kinetic experiments with HaloTag-Ubiquitin (Figure 3C). The different NanoBRET ratios obtained for the same target treated with either MZ1 or dBET1 indicate that ubiquitination, either in pattern or extent, is different dependent on the E3 ligase complex to which it is recruited. The degradation rates determined in Figure 2C show striking similarity to the trends in ubiquitination for respective family members and PROTACs. Plotting degradation rates against the relative increase in ubiquitination at the same concentration of PROTAC revealed a positive and linear correlation, indicating the step of ubiquitination is the predictive step of degradation and controls the rate (Figure 3D). Lytic and Live Cell Target Engagement and Cell Permeability Assessment. Given the large molecular size of heterobifunctional degraders, a significant step in the process of chemical optimization is characterization of cell permeability, target occupancy, and binding affinities to both the target and E3 ligase complex. To facilitate these studies, we F


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VHL protein (Kd = 66 nM for MZ137 and 90 nM for VH29848). These results, however, were not recapitulated in live cells, with MZ1 showing a >40-fold reduction in binding affinity relative to that of VH298 (Figures 4C and F). Similar live cell target engagement studies were done with NanoLuc-CRBN and a CRBN tracer and showed a 10-fold reduction in binding of dBET1 as compared to that of thalidomide (Supporting Information Figures 6A−C). To interrogate the live cell binding affinities and kinetics to BRD4 by these PROTACs, competitive displacement studies were done using a BET family tracer with NanoLuc-BRD4. Results showed dBET1 and MZ1 had an 8- and 15-fold right-shifted reduction in binding affinity, respectively, compared to that for JQ1 in live cells (Figures 4D and F). Intracellular kinetics were performed and found that both PROTACs had significantly slower target engagement compared to JQ1 in live cells and apparent IC50 values which decreased over time (Figure 4E). Together, these results indicate reduced and slow cellular permeability of MZ1 and dBET1, yet they remain highly efficacious for degradation at substoichiometric binding affinity concentrations, supporting a catalytic mechanism of action by these PROTACs. Response of cMyc in MV4;11 Cell Lines to Inhibitors and PROTAC Treatments. One of the primary consequences and desired outcomes of BET family inhibition is downregulation of key oncogenic targets, including cMyc.44,45 To quantitatively study cMyc levels post-treatment with BET inhibitors as compared to PROTACs, we tagged endogenous cMyc with HiBiT on the C-terminus using CRISPR/Cas9 in a relevant leukemia line, MV4;11 (Supporting Information Table 1). Treatment of these cells with 10 nM JQ1, dBET1, or MZ1 over a 24 h time period showed differential levels of cMyc expression knockdown, from minimal loss with JQ1, to 50 and 80% loss with dBET1 and MZ1, respectively (Figure 5A). Correlation to cell viability assays at each time point revealed that only treatment with MZ1 resulted in measurable cellular death at this concentration (Figure 5B). At a higher concentration, both compounds showed significant reduction in cMyc expression and concurrent cellular death (Figures 5C and D). While cMyc is one of many transcriptional targets impacted by BET inhibition44,45 and PROTAC-mediated degradation,14 the use of HiBiT endogenous tagging as a downstream reporter of function allowed for understanding of the amount of loss needed to elicit the desired phenotypic outcome.

wanted to transition from monitoring protein:protein interactions to protein:small molecule interactions. To this end, we applied NanoBRET technology using ectopic NanoLuc fusions as donors and fluorescently labeled small molecule tracers as acceptors.43 This approach interrogates the separate events of PROTAC binding to either target or E3 ligase component (Figure 4A), with a key advantage in that it can be performed in both live cell or lysate formats to assess permeability.43 Competitive displacement of the VHL tracer molecule revealed identical binding affinities between MZ1 and a related potent monovalent VHL binder, VH298,48 in lytic format (Figure 4B), in agreement with biophysical measurements in vitro which showed identical binding affinity with recombinant

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Figure 4. NanoBRET target engagement with PROTAC compounds. (A) Schematic showing NanoBRET target engagement strategy with either E3 ligase component or BRD4 using NanoLuc fusions as energy donors and fluorescent small molecule tracers as energy acceptors, which can be competitively displaced by unlabeled compounds. (B and C) Competitive displacement profiles of HEK293 cells transiently transfected with NanoLuc-VHL and incubated with the VHL fluorescent tracer in the presence of serial titrations of either MZ1 or VH298 in digitonin-permeabilized lysate (B) or live cells (C). (D) Live cell competitive displacement profiles of HEK293 cells transiently transfected with NanoLuc-BRD4 and incubated with the BRD4 fluorescent tracer in the presence of serial titrations of either MZ1, dBET1, or JQ1 in live cells. Data are represented as NanoBRET ratios normalized to zero compound. Error bars are expressed as SD of the mean (n = 3) of a representative experiment. (E) NanoBRET kinetic monitoring of IC50 values for JQ1, dBET1, and MZ1 binding to BRD4 in live cells. Data are expressed as singlicate IC50 values from RLU (n = 3) of a representative experiment. (F) Tabulated IC50 values for indicated target, PROTAC, and assay format.

CONCLUSION Generation of heterobifunctional PROTAC compounds, either de novo or from known inhibitors, presents an exciting new direction for drug discovery.1,2,10,12 BET family PROTACs, including those studied here, have shown advantages over BET inhibitors and hold great promise for potential treatment of a wide variety of diseases, including cancer and inflammation.14,17,30,32,34,35,44,45 In this work, we further understanding of the cellular mechanism of action of dBET1 and MZ1 using a suite of cell-based assays combined with CRISPR/Cas938,39 endogenous tagging, allowing for precise and quantitative kinetic analysis of key degradation events. Importantly, we also assess which stages of the degradation process are impacted by recruitment to different E3 ligase complexes, a critical consideration and variable in the development of PROTAC compounds. Currently the primary E3 ligase components utilized are VHL and CRBN, but active research efforts are G


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Figure 5. PROTAC modulation of cMyc-HiBiT expression and viability in MV4;11 cells (A and B) MV4;11 cells containing endogenously tagged cMyc-HiBiT were treated 10 (A) or 100 (B) nM JQ1, dBET1, or MZ1. (C and D) Cell viability was measured by CellTiter-Glo following treatment with 10 (C) or 100 (D) nM for each respective compound. All data are represented as RLU normalized to DMSO control. Error bars are expressed as SD of the mean (n = 3) of a representative experiment.

Figure 6. Model of degradation phases and contributing mechanisms Schematic showing the three phases of degradation: initiation of degradation, degradation maximum (Dmax), and recovery correlated to the key mechanistic processes listed in black for each phase. The target protein response to PROTACs at each phase is represented pictorially, showing first introduction of PROTAC and target protein, followed by loss of target protein, and then recovery. Listed in red are the parameters identified to regulate each phase with arrows depicting the processes which to monitor or optimize.

complex with MZ137 or related dBET compounds47 and their respective E3 ligase partners. In the case of MZ1/VHL, positive cooperativity in binding was observed,37 in agreement with our finding that these complexes were more stable over time in cells. In contrast, for related dBET compounds with CRBN, a lack of cooperativity as well as multiple conformations have been reported,47 suggestive of more dynamic and transient ternary complex formation for this system. Indeed, we see this behavior mirrored inside the cell with complexes forming quickly but not showing increasing stabilization over time with any of the BET family members. Our studies were done in HEK293 cells, and it is worth noting that degradation profiles could vary in other cell lines that may have different expression levels of E3 ligase components or BET family members. However, we would not expect overall trends to be different because ternary complex stability and activity of specific complexes would likely be the same. Because thorough studies comparing functional outcomes imparted by use of different E3 ligase handles have not been previously performed, it is difficult to predict whether changes in the E3 recruitment system will be favorable or

underway to identify other efficacious E3 ligase components to serve as recruiters.2−4,12,16,19 This will result in an expansion of options in the future, directly translating to increased chemical development and furthering the need for robust and relevant technologies to efficiently profile and triage PROTAC cellular activity. To fully characterize and compare PROTAC-mediated degradation via the VHL or CRBN E3 ligase complexes, quantifiable parameters of the cellular degradation profile were determined in these studies. Across all BET family targets studied here, we found that the MZ1/VHL recruitment system elicits faster rates of degradation, slower, but more stable ternary complex formation, enhanced ubiquitination, greater compound potency, and longer-lasting degradation efficacy as compared to the dBET1/CRBN system. Because both compounds contain JQ1 as the target specific handle, we conclude these differences are directly attributed to ternary complex stability and relative positioning within the different E3 ligase systems of each BET target protein for ubiquitination. Our ternary complex kinetic data correlate well with recent structural studies of bromodomains of BRD4 proteins in H


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capabilities of high sensitivity over a broad dynamic range.41 In our studies, we were are able to accurately and continuously monitor degradation over a 3-log range with high reproducibility, indicating this approach can enable detection of proteins that have low levels of native expression and/or for characterization of early development PROTAC compounds, which might not show fast or robust degradation. Other approaches are currently also used to study protein:protein and protein:small molecule interactions but do not have the same advantages for degradation research as compared to NanoBRET.42 Given its proximity-based and ratiometric nature, NanoBRET is suited for studying interactions that are transient, and measurement of the interaction is not influenced by changes in the level of donor if this occurs.42 This is why we have specifically chosen the target protein as the energy donor in the configuration of the assay, allowing for determination of complex formation even with active loss of the target. In contrast, other protein interaction technologies are reliant on a single readout, such as fluorescence or luminescent complementation and lack the ability to discern whether changes in signal are due to the modulation of the interaction or changes in levels of one or both partners. A further advantage of NanoBRET is the dual readout capability, which allows simultaneous monitoring of both the interaction (protein:protein or protein:small molecule) by calculation of NanoBRET ratios as well as target loss by direct luminescent monitoring, provided the target protein is the donor. The platform of technologies presented here to characterize the cellular mechanism of action of two BET bromodomain PROTACs, dBET1 and MZ1, not only advance our understanding of these chemical probes, but more broadly provide a comprehensive analysis to advance PROTAC drug discovery and cellular activity assessment. Our cellular analysis shows that both ternary complex stability and subsequent ubiquitination are key steps, but they influence and regulate different phases of degradation (Figure 6), correlations not previously described due to studies done primarily with in vitro assays. These understandings combined with our technology platform to study each parameter in live cells will help further compound optimization, including fine-tuning family member specificity if desired. While the focus of this work was on PROTAC characterization, the approach presented here can be applied to study any process that impacts protein homeostasis, as demonstrated with cMyc (Figure 5) and in our other studies of therapeutic targets HIF1α and β-catenin (data not shown). The diverse capabilities and applications of these technologies coupled with endogenous tagging via CRISPR/Cas9 present a highly enabling and physiologically relevant means to study protein degradation, providing unique insight into the individual and highly complex mechanisms regulating the degradation process.

unfavorable for any given target. Additional BET family PROTACs,17 including more potent versions of dBET1, have been reported.47,50 It would be intriguing to determine which cellular steps and parameters contribute to the improvement in potency for this class of compounds, and then translate this understanding to help guide further chemical design as well as to enhance SAR for BET degraders in general. More broadly, we seek to use the functional relationships determined from analysis of dBET1- and MZ1-mediated degradation to build a mechanistic map of primary contributing events for each phase of the PROTAC cellular degradation profile; degradation initiation, Dmax, and recovery (Figure 6). This model can be used as a guide for compound rank-order profiling, design, and optimization, showing which measurements will be most predictive in improving any given phase in degradation. The important contributors to the initial degradation phase are PROTAC compound permeability, binding affinities to targets and E3 components, and recruitment not simply into a ternary complex, but a productive complex to achieve ubiquitination. Ultimately the level of target ubiquitination, and not necessarily the efficiency of ternary complex formation dictates the rate of degradation. For Dmax, we find, perhaps not unexpectedly, that this phase is related to the long-term stability of the ternary complex and is also highly dependent on any downstream cellular feedback to loss of the target which would promote recovery. This interplay is most evident in our data for BRD2, which shows fast rate of degradation, stabilized ternary complex formation, but by far the shortest time at Dmax. We found this to be due to competing upregulation of protein levels, a response stimulated by JQ1 treatment alone that appeared to be amplified in response to PROTAC treatment from the added layer of BET family degradation compared to inhibition. The protein recovery phase, which is challenging to study given its multiphasic and more complex profiles, will likely vary greatly between targets, as we have found for BET family members, and will also depend on the cellular chemical stability of the PROTAC compound. Together, these correlations help break down some of the barriers in properly assessing PROTAC activity within the highly complex degradation process, providing useful entry points and important parameters to monitor for optimization of key steps required for degradation. Our understanding of the degradation process, particularly the quantitative and kinetic determination of the three phases shown in Figure 6, would not have been possible without the use of the highly sensitive endogenous tagging system, HiBiT,41 and complementary intracellular protein interaction studies using NanoBRET.42,43 Prior to our studies presented here, understanding of PROTAC degradation profiles have been highly granular with limited quantitation, using primarily Western blot analysis to observe target loss and recovery at fixed time points and concentrations. Moreover, other approaches have focused on monitoring ectopically expressed fusion tagged proteins or domains in the cells.47 While this allows for qualitative and visual determination of degradation as well as study of protein interactions as we have shown here, its application for quantitative degradation analysis resulted in alteration of degradation rate, reduced Dmax, right-shifted DC50 values, and non-native recovery profiles. Therefore, to maintain endogenous expression level and regulation, the pairing of HiBiT luminescent technology and CRISPR/Cas9 endogenous tagging was critical. Compared to other tags that could also be used for endogenous insertion, HiBiT provides additional

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MATERIALS AND METHODS

Expression Plasmids. Clones expressing N-terminal HaloTag fusions of human VHL (NM̅ 000551) and Cereblon (AAH17419.1) were obtained from Kazusa DNA Research Institute (Kisarazu, Japan) as pFN21A HaloTag CMV Flexi Vectors (Promega). The N-terminal HaloTag-Ubiquitin constructed consisted of the first ubiquitin repeat, amino acids 1−76, of the human polyubiquitin-B precursor (NM̅ 018955) and was cloned into the pFN28K (Promega) without appending any additional residues to the native C-terminal sequence. N-terminal NanoLuc fusions of human VHL (NM̅ 000551), Cereblon

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ACS Chemical Biology (AAH17419.1), and BRD4, long isoform (NM̅ 058243) were cloned into pFN31K vectors (Promega). Reagents and Cell Culture. dBET1 was purchased from Cayman Chemical, JQ1 was purchased from Xcessbio Biosciences, MZ1 and VH298 were generously donated by A. Ciulli (University of Dundee), and MG132 was purchased from InvivoGen. HEK293 and MV4;11 cells were obtained from the American Type Culture Collection and cultured at 37 °C, 5% CO2 in DMEM (Gibco) or IMDM (American Type Culture Collection), respectively, containing 10% fetal bovine serum (Seradigm). Generation of HEK293 LgBiT Stable Cell Line. HEK293 cells (8 ×106) plated in a 10 cm2 Petri dish were transfected with 20 μg LgBiT CMV plasmid for 24 h and grown in 2 mg mL−1 G418 selection antibiotic (Promega) until large colonies were present. Clones were isolated by limiting dilution into 96-well plates and after expansion were screened for LgBiT expression by triplicate plating with one plate receiving HiBiT Lytic Detection Reagent (Promega) supplemented with purified HiBiT-HaloTag and another plate receiving CellTiter-Glo (Promega). A clone with high luminescence relative to CellTiterGlo was expanded from the third plate and maintained in 800 μg/mL G418. CRISPR/Cas9-Editing. One nanomole Alt-R CRISPR RNA (crRNA) and 1 nmol Alt-R trans-activating crRNA (tracrRNA) were assembled in 50 μL of Nuclease-Free Duplex Buffer (Integrated DNA Technologies) by incubation at 95 °C for 5 min and cooling to RT to generate gRNA for BRD2, BRD4, and cMyc. CRISPRevolution sgRNA EZ kit (Synthego) was used as the gRNA for BRD3. Singlestranded ultramer DNA oligonucleotides (IDT) were used as the ssODN donor templates. Ribonucleoprotein (RNP) complexes with recombinant Streptococcus pyogenes Cas9 with an N-terminal tag containing a histidine-nuclear localization signal-myc sequence (Aldevron) were assembled as previously described41 by incubating 100 pmol of Cas9 and 120 pmol of gRNA for 10 min at ambient temperature. For HiBiT knock-in to BRD2, BRD3, and BRD4, 2 × 105 HEK293 cells were resuspended in 20 μL of 4D Nucleofector solution SF, and RNP complexes along with 100 pmol ssODN template were electroporated into cells with the 4D Nucleofector System (Lonza) using program CM-130. For HiBiT knock-in to cMyc, MV4;11 cells were resuspended in 20 μL of 4D Nucleofector solution SE, and RNP complexes along with 100 pmol ssODN template were electroporated into cells with the 4D Nucleofector System (Lonza) using program DJ-100. Immediately following electroporation, cells were incubated at ambient temperature for 10 min before transferring to a 12-well plate for culturing. Edited pools were analyzed for HiBiT insertion by assaying for luminescence on a GloMax Discover (Promega) 48−72 h postelectroporation. Generation and Validation of Clonal HiBiT Insertions. Clonal populations of edited cells were obtained by sorting live singlets using a FACSMelody cell sorter (Becton Dickinson) into 96- or 384-well plates. Clones were expanded and screened by luminescence, and genomic DNA was isolated using the Wizard Genomic DNA Purification kit (Promega) from 8 to 10 clones per edited pool. Genomic DNA was amplified using primers designed to anneal ∼200 bases either upstream or downstream from the inserted ssODN template sequence, subcloned into a pF5K vector backbone (Promega), transformed into JM109 cells (Promega), and 24 individual colonies per clonal population were sequenced by Sanger sequencing. Clones were selected based on 100% sequence conformity with no insertions or deletions in the HiBiT-containing open reading frame. HiBiT Blotting. HEK293 clonal cells (2 ×105) expressing HiBiTBET family members were plated in 24-well dishes overnight and then lysed in 100 μL of mammalian lysis buffer (Promega) supplemented with Protease Inhibitor Cocktail (Promega) and RQ1 RNase-Free DNase (Promega) on ice. One hundred microliters of 2× SDS-PAGE gel loading buffer (120 mM Tris-HCl, pH 6.8, 1.5 mM bromophenol blue, 25% glycerol, 200 mM dithiothreitol, and 1% SDS) was added to each sample and incubated for 5 min at 95 °C. Lysates were separated by SDS-PAGE and transferred to a nitrocellulose membrane. Following transfer, the membrane was briefly incubated in TBST

(20 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 0.1% Tween 20) with 1 mM dithiothreitol (DTT) before replacing buffer with NanoGlo Blotting Buffer containing 100 nM purified LgBiT protein (Promega) and incubating at 4 °C overnight with shaking. NanoGlo Luciferase Assay Substrate (Promega) was added, and luminescence was captured on an ImageQuant LAS 4000 digital imaging system (GE Healthcare Life Sciences). Bioluminescence Imaging. Clonal HiBiT-BRD4 edited cells were plated in 8-well glass bottom chamber slides (Nunc Lab-Tek II) at a density of 8 × 104 per well in 200 μL of growth medium and incubated at 37 °C, 5% CO2 overnight. Following incubation, medium was replaced with Opti-MEM (Gibco) containing 20 μM Vivazine extended release substrate (Promega) and incubated for an additional 1 h at 37 °C, 5% CO2 to allow equilibration of substrate and stabilization of luminescent signal. Real-time imaging was performed using the LV200 bioluminescence imaging system (Olympus) equipped with an ImagEM X2 EM-CCD camera (Hamamatsu) and a temperature-controlled stage. Images were collected for a period of 2 h with cellSens software (Olympus) using a 60× oil-immersion objective, electron multiplying EM gain of 1200, and 10 s exposure times. Average projections of 10 sequential images were generated, and dynamic range adjustments and pseudocolor rendering was performed using FIJI. Kinetic Live Cell HiBiT Detection. Clonal cell lines edited for HiBiT insertion to BET family members were plated in white 96-well tissue culture plates at a density of 2 × 104 cells per well in 100 μL of growth medium and incubated overnight at 37 °C, 5% CO2. For experiments with transient expression of NanoLuc-BRD4, 8 × 105 HEK293 cells were first transfected with 0.02 μg of NanoLuc-BRD4 diluted in 2 μg of promoterless carrier DNA using FuGENE HD (Promega), and the following day, were plated in white 96-well tissue culture plates at a density of 2 × 104 cells per well in 100 μL of growth medium and incubated overnight at 37 °C, 5% CO2. For end point live cell detection, wells were treated with dBET1 or MZ1 compounds for the indicated time frames before addition of NanoGlo Live Cell Substrate (Promega) and reading luminescence on a GloMax Discover. For continuous live cell detection out to 24 h, medium was replaced with CO2-independent medium (Gibco) containing 20 μM Endurazine, an extended time-released substrate (Promega), and plates were incubated at 37 °C, 5% CO2, for 2.5 h before addition of a 3-fold serial dilution of 1 μM final concentration dBET1, MZ1, or JQ1 compounds. Plates retaining the plate lids were then read every 5 min for a period of 24 h on a GloMax Discover (Promega) set to 37 °C. Quantitation of Degradation Kinetics. Degradation rate and degradation plateau were calculated by fitting only the initial degradation portion of each kinetic concentration curve to the equation:

y = (y0 − plateau)e−ƛt + plateau where ƛ = degradation rate in units of hr−1. The degraded fraction, Dmax, was calculated as 1 − plateau. For each curve, the first few data points before onset of degradation were excluded from the fits. Time at Dmax was determined by calculating the length of time during which the fractional population remained below a threshold defined as plateau + 0.1(Dmax). Lytic HiBiT Detection and Viability of MV4;11 Cells. Clonal cMyc-HiBiT edited cells were replicate plated at a density of 5 × 104 cells per well in solid, white 96-well tissue culture plates (Corning Costar #3917), and cultured overnight. The following day, cells were treated with either 10 nM or 100 nM JQ1, dBET1, or MZ1 for the indicated time frames. Following each treatment time point, one replicate plate was assayed for luminescence of cMyc-HiBiT by addition of an equal volume of NanoGlo HiBiT Lytic Reagent (Promega N3030) to each well, and another replicate plate was assayed for cell viability by addition of an equal volume of CellTiterGlo 2.0 to each well (Promega). Plates were shaken on an orbital shaker for 10−20 min before reading luminescence on a GloMax Discover (Promega). J


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ACS Chemical Biology NanoBRET Live Cell Ternary Complex and Ubiquitination. HEK293 clonal cells (8 ×105) expressing the HiBiT-BET family members were transfected with FuGENE HD (Promega) and 2 μg of HaloTag-CRBN, HaloTag-VHL, or HaloTag-UBB in 6-well plates. For transient NanoBRET experiments with NanoLuc-BRD4, 8 × 105 HEK293 cells were transfected with FuGENE HD (Promega) and 0.02 μg of NanoLuc-BRD4 with either 2 μg of HaloTag-CRBN, or HaloTag-VHL. The following day, 2 × 104 transfected cells were replated into white 96-well tissue culture plates in the presence or absence of HaloTag NanoBRET 618 Ligand (Promega) and incubated overnight at 37 °C, 5% CO2. The following day, medium was replaced with Opti-MEM (Gibco) containing 20 μM Vivazine, an extended time-released substrate (Promega), and plates were incubated at 37 °C, 5% CO2, for 1 h before addition of DMSO or 1 μM final concentration dBET1 or MZ1 compounds. Plates were then read every 3 min for a period of 6 h on a CLARIOstar equipped with an atmospheric control unit (BMG Labtech) set to 37 °C and 5% CO2. Dual filtered luminescence was collected with a 460/80 nm bandpass filter (donor, NanoBiT-BET protein) and a 610 nm long pass filter (acceptor, HaloTag NanoBRET ligand) using an integration time of 0.5 s. Background subtracted NanoBRET ratios expressed in milliBRET units were calculated from the equation:

NanoGlo Substrate and Extracellular NanoLuc Inhibitor (Promega) were added according to the manufacturer’s recommended protocol, and filtered luminescence was measured on a GloMax Discover luminometer equipped with 450 nm BP filter (donor) and 600 nm LP filter (acceptor) using 0.3 s integration time. For real-time target engagement analysis, NanoBRET NanoGlo Substrate and Extracellular NanoLuc Inhibitor (Promega) and 1× NanoBRET tracer were added to the cells 30 min prior test compound addition. To measure NanoBRET in permeabilized cells, digitonin was added to the cells to a final concentration of 50 ug/mL and Extracellular Nluc inhibitor was omitted during the detection step. Milli-BRET units (mBU) are calculated by multiplying the raw NanoBRET values by 1000.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acschembio.8b00692. Tables of CRISPR guide and template sequences and comparative HiBiT-BET family expression; figures of kinetic competition of dBET1 and MZ1 with JQ1, impact of ectopic expression on degradation, BET family member time at Dmax at 10 and 100 nM PROTAC, BET family protein level profiles after JQ1 inhibition, ternary complex specificity and impact of proteasomal inhibition, and NanoBRET target engagement of dBET1 with NanoLuc-CRBN, (PDF)

acceptor channel (no ligand) zyz ji acceptor channel zz1000 mBRET ratio = jjjj − donor channel donor channel (no ligand) z{ k

Fold increase in BRET was calculated by normalizing mBRET ratios to the average mBRET ratios for DMSO controls. NanoBRET Ubiquitination ImmunoAssay. Clonal cell lines edited for HiBiT insertion to BET family members were plated in white 96-well tissue culture plates at a density of 2 × 104 cells per well in 100 μL of growth medium and incubated overnight at 37 °C, 5% CO2. The following day, 1 μM dBET1 or MZ1 was added to the plate for the indicated time frames before replacing medium with OptiMEM (Gibco) containing 200 μg/mL digitonin, 1:200 dilution of primary anti-Ub antibody (Enzo Life Sciences, BML-PW8810), 1:500 dilution of secondary antimouse Alexa 594 antibody (Cell Signaling Technologies, 8890), and 20 μM NanoGlo substrate. Additional control wells received no antibodies (control for background NanoBRET) or no primary antibody (control for specificity). Plates were placed on an orbital shaker for 10 min and NanoBRET measurements were collected on a CLARIOstar (BMG Labtech). NanoBRET calculations were made according to the equation in the section: NanoBRET Live Cell Ternary Complex and Ubiquitination. Fold increase in NanoBRET was calculated by normalizing mBRET ratios to the untreated, t = 0 wells. NanoBRET Target Engagement. NanoLuc-CRBN, VHL-NanoLuc, and NanoLuc-BRD4 were transfected in HEK-293 cells using FuGENE HD (Promega) according to the manufacturer’s protocol. Briefly, NanoLuc/target fusion constructs were diluted into Transfection Carrier DNA (Promega) at a mass ratio of 1:10 (mass/mass), after which FuGENE HD was added at a ratio of 1:3 (μg DNA: μL FuGENE HD). One part (vol) of FuGENE HD complexes thus formed were combined with 20 parts (vol) of HEK-293 cells suspended at a density of 2 × 105, followed by incubation in a humidified, 37 °C/5% CO2 incubator for 20 h. Following transfection, cells were washed and resuspended in Opti-MEM. NanoBRET assays were performed in white, 96-well plates (Corning 3600) at a density of 2 × 104 cells/well. All chemical inhibitors were prepared as concentrated stock solutions in DMSO (Sigma-Aldrich) and diluted in Opti-MEM (unless otherwise noted) to prepare working stocks. For end point analysis of target engagement, cells were equilibrated for 2 h with energy transfer probes and test compound prior to NanoBRET measurements. NanoBRET tracers were prepared at a working concentration of 20× in tracer dilution buffer (12.5 mM HEPES, 31.25% PEG-400, pH 7.5). VHL NanoBRET Tracer was added to the cells at a final concentration of 1 μM, CRBN NanoBRET tracer was added to cells at a final concentration of 0.5 μM, and NanoBRET BRD Tracer-02 was added to cells at a final concentration of 0.25 μM. To measure NanoBRET in live cells, NanoBRET

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AUTHOR INFORMATION

Corresponding Author

*Tel: +1-608-274-4330; Fax: +1-608-277-2601; E-mail: [email protected]. ORCID

Danette L. Daniels: 0000-0002-7659-3020 Notes

The authors declare the following competing financial interest(s): All authors are employees of Promega Corporation and Promega Corporation is the commercial owner by assignment of patents of the HaloTag, NanoLuc, NanoBRET target engagement, NanoBiT, and HiBiT technologies and their applications.

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ACKNOWLEDGMENTS We wish to thank A. Ciulli for providing MZ1 and VH298 compounds, T. Machleidt and M. Schwinn for guidance on HiBiT CRISPR/Cas9 editing and development of LgBiT stable cell line, M. Slater for advice on CRISPR clonal selection, C. Zimprich for help with target engagement, T. Worzella for Glo-Max Discover instrumentation support, and G. Tarpley for thoughtful advice and continued support.

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REFERENCES

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