The In-Cell Western immunofluorescence assay to monitor PROTAC mediated protein degradation

Abstract

The In-Cell Western plate-based immunofluorescence assay is a useful methodology for monitoring protein levels and provides a facile moderate through-put method for PROTAC and degrader optimization. The method is compared to other reported assays used for PROTAC development. The advantages of this method are the greater through-put compared to Western blots due to its plate-based method and the ease to transfer between cells lines. Adherent cell lines are preferred, although suspension cells can be used following recommended modifications and precautions to the protocol. This method requires a high-quality antibody that recognizes the protein epitope in its cellular context, and in general provides data similar to Western blots with higher assay through-put.

1. Introduction

1.1. In-Cell Western assay to monitor PROTAC mediated protein degradation

Targeted protein degraders provide attractive approaches to inhibit challenging drug target proteins that play an important role in cancer and other diseases (Békés, Langley, & Crews, 2022; Chamberlain & Hamann, 2019; Hanzl & Winter, 2020). Targeted protein degraders include heterobifunctional molecules such as proteolysis-targeting chimeras (PROTACs) which contain a high affinity ligand to a target protein tethered to a ligand to recruit an E3 ligase, resulting in poly ubiquitination and proteasomal degradation, or a Molecular glue degrader which is a small molecule that stabilizes the interaction between a protein and an E3 ligase. At present there are at least 6 molecular glues and 12 heterobifunctional degraders in clinical trials (Békés et al., 2022) emphasizing the translational importance of this approach. Assays that can be broadly utilized for lead optimization are of great importance. Analysis of target protein levels after treatment with compounds developed as new PROTACs or degraders is an essential assay used for structure activity relationship (SAR) development.

The In-Cell Western assay is a plate-based quantitative immunofluorescent method that can be used to monitor protein degradation levels following PROTAC treatment to compliment Western blotting and provide increased through-put. This assay uses adherent cells cultured in 96-well plates which are treated with compounds synthesized as potential degraders and then fixed and permeabilized in the microplate for immunostaining. The technique provides a robust and sensitive assay due to the use of secondary antibodies conjugated with bright IRdye^® near-infrared fluorescent dyes resulting in a method to monitor protein levels where cells are normalized to cell number using cell or DNA stains. The fluorescent signal generated provides an accurate readout of protein expression levels and cell population in each well. This technique avoids much of the labor intensive and time-consuming steps involved in a traditional Western blot. In addition, this technique monitors proteins in their cellular context, and allows for the rapid and precise observation after cellular treatments. In general, the results are consistent with Western blotting, and this technique provides a higher throughput, less labor-intensive method for PROTAC optimization compared to optimization solely using Western blotting. Therefore, PROTACs or degraders which show activity in the In-Cell Western assay can be prioritized for evaluation in secondary Western blot confirmatory steps rather than using Western blotting as the primary assay.

1.2. General techniques for PROTAC optimization

PROTAC and degrader optimization is complex and requires assay methods to evaluate the affinity towards the target protein of interest, affinity towards the E3 ligase that is being recruited, followed by evaluation of the bifunctional molecule to degrade the protein of interest in a cellular context. This iterative optimization cycle requires moderate through-put so that rapid feedback is provided to medicinal chemists who need to evaluate changes to all three key regions of the PROTAC, the target ligand, the E3 ligase recruiting ligand, and the linker, then synthesize additional analogs to further optimize potential degrader candidates. There are several reported methods to monitor cellular protein degradation after PROTAC and degrader treatment which are used for optimization, and are briefly reviewed below (Liu et al., 2020; Daniels, Riching, & Urh, 2019). Each technique has its strengths and weaknesses which include complexity of use, expertise, instrumentation, and evaluation in an endogenous versus an ectopically expressed system. In general, the In-Cell Western assay compliments these methods by providing a method to evaluate new analogs in a relevant cellular context using a plate-based approach to rapidly provide data back to medicinal chemistry.

Global Mass Spectrometry provides a robust method to evaluate protein ubiquitination, a post translational modification readily assayed by using mass spectrometry (Zheng & Shabek, 2017) or immunoblotting. This technique provides a robust method to detect linkage type endogenous ubiquitination levels. Global proteomics (Brand et al., 2019; Deracinois, Flahaut, Duban-Deweer, & Karamanos, 2013) is a sophisticated method to evaluate degrader selectivity and provides information such as the number of proteins affected due to PROTAC treatment relative to a non-treated control. These methods are excellent in providing an in-depth study of a few compounds but are difficult to implement in high-throughput and are laborious for studying multiple time points. Monitoring degradation in real-time using ectopic expression of target proteins fused to fluorescent proteins, i.e., GFP, provides robust assays to monitor cellular target engagement and ternary complex formation (Dobrovolsky et al., 2019). This technique provides significant advantages but might not reflect the true degradation and regulation of the endogenous target (Zeng et al., 2020). NanoBRET can be used to measure target engagement (Robers et al., 2019). This technique relies on the bioluminescence resonance energy transfer between a fluorophore ligand (BRET acceptor) and a NanoLuc luciferase (BRET donor) which is activated by NanoLuc substrates. Target engagement results in a loss of BRET signal due to displacement of the fluorophore-labeled ligand. The advantage of this approach is the ease of measuring fluorescent signals, however in most cases this system requires over-expressed proteins which may confound PROTAC activity in the native endogenous system and may be limited to available fluorescent probes. CRISPR/cas9 to generate HiBiT tagged proteins with expression of LgBiT for complementation luminescence assays provides the ability to study cellular target engagement in either an endogenous or ectopically tagged fusion (Daniels et al., 2019). This method uses CRISPR/cas9 to tag proteins of interest with HiBiT, an 11 amino acid peptide that results in bright luminescence following complementation with LgBiT (Riching et al., 2018) and provides a sophisticated cell-based system to quantify degradation rates and maximum levels of degradation. However, although this system offers many advantages, it is not readily transferable to many different cells which may have different expression levels of E3 ligase components and show different degradation profiles. Cellular thermal shift assay, CESTA, is a technique that relies on a principle that upon heating the proteins unfold and aggregate (Martinez Molina et al., 2013). The aggregated protein is removed during processing and the remaining soluble protein is quantified (Savitski et al., 2014). In this technique, PROTAC compound binding to the target protein stabilizes the target protein during heating which is visualized by a shift in the thermal melt curve, where the unbound target protein is typically less stable than the complex. The isothermal study over a range of PROTAC concentrations provides information on target occupancy. This technique requires lysis of cells, heating, and detection by antibodies of the stabilized proteins, thus high-quality antibodies are important for this technique. Note that this technique may be complicated due to competing target degradation during observation of thermal stability. Simple Western (capillary electrophoresis immunoassay (CEI)) Systems (Chen et al., 2013) which simplify Western blots include the Simple Western system of automating the protein separation and immune detection steps. This reduces the overall processing time to about 3h. This technique uses capillary electrophoresis immunoassay (CEI) based technology (Moser & Hage, 2008; Voeten, Ventouri, Haselberg, & Somsen, 2018). The process still relies on preparation of cell lysates from treated cells, and selection of antibodies. Antibodies are detecting denatured protein in this technique, similar to a traditional Western blot. The main advantage of this technique is the reduced processing time and the ability to reliably capture and quantify high molecular weight proteins, as well as the reduced sample requirements. They also eliminate run-to-run variability of Western blots by automating loading, transfer, and incubation and wash times. High-throughput flow cytometry is a widely used laser-based technology that provides a medium throughput methodology for monitoring protein degradation in 96-well formats for increased throughput (Black, Duensing, Trinkle, & Dunlay, 2011). Flow cytometry measures fluorescent intensity of a fluorescent-labeled antibody specifically bound to a target protein of interest expressed intracellular or on the surface of the cell. This technique reduces the total processing time from sample preparation to analysis. An advantage with this technique is the ease in use of suspension cells, although adherent cells can also be evaluated, it is preferred to use suspension cells to avoid extra steps to suspend adherent cells enzymatically or mechanically. In this technique treated cells are fixed, permeabilized, washed, then treated with fluorescently labeled antibody for visualization analysis. Flow cytometry measures a single cell at a time and provides an analysis of cell populations. Whilst all these approaches are available, most laboratories still rely on traditional Western blotting approaches.

1.3. Comparison of In-Cell Western to Western blot

Western blotting is typically used for analysis of protein levels. In this technique a mixture of proteins from a cell lysate are subjected to gel electrophoresis and separated by molecular weight. These are then transferred to a membrane made of nitrocellulose or PVDF (polyvinylidene fluoride) where application of an electrical current induces the proteins to migrate from the gel to the membrane. The membrane can then be further processed with antibodies specific for the target of interest and visualized using secondary antibodies and detection reagents, producing a band for the protein of interest in the mixture (Mahmood & Yang, 2012). The process is labor intensive requiring heating the cell lysate, gel preparation, accurate sample loading, running the sample through the gel under a current, transfer of the protein from the gel to a membrane under an electrical current, and processing the membrane with a primary antibody for labeling and a secondary antibody for visualization of the protein of interest.

Western blotting as a method of protein quantification is widely adopted as the method of choice since its inception in 1979. The technique is inherently simple and inexpensive resulting in widespread use of this technique. Thus, Western blotting has been used extensively to measure changes in protein expression resulting from pharmacological treatment and is considered the gold standard. However, the Western blotting method has several drawbacks when considered as a primary technique as a rapid assay for structure activity relationship (SAR) development. Mainly the technique is labor intensive and considered low throughput. Due to the extensive sample processing involved in the technique this may limit the number of experimental conditions that are explored such as a limited time course and limited dose concentrations tested for a dose responsive effect of pharmacological treatment.

The In-Cell Western (ICW) assay is an alternative method based on the same principals as Western blotting. ICW is a medium to high throughput technique for quantifying protein levels and signaling events. The technique is plate based, typically using 96-well plates or 96-well half volume plates to save reagents, so that many samples can be evaluated and processed in parallel. This technique is well suited for adherent cells due to the number of washing steps, but suspension cells can be used following modifications to the method (LI-COR protocols http://bit.ly/2IEtwqT). The change in fluorescent signal is linear to cell number, where the change in fluorescent signal coming from the primary antibody signal is relative to a secondary fluorescent control signal from a cell or DNA stain that is constant. This provides an easy to monitor fluorescent signal that can be quantitatively compared for different degraders using various time points and concentrations. The system can readily be transferred across different cell lines. This technique provides moderate throughput for sample processing, for example 20 compounds can readily be evaluated with a 2–3-day turn-around-time with 8–10 concentrations per compound to provide rapid feedback to medicinal chemists optimizing a PROTAC series. This technique does not separate proteins on a gel therefore non-specific primary antibody binding will reduce the signal. Importantly the primary antibodies will detect protein in a more relevant cellular conformation, instead of a denatured protein which is detected by traditional Western blot. Like any technique that relies on antibody recognition, it is possible that ubiquitination may block the epitope recognized by the primary antibody or if the protein is in complex, such as bound to chromatin, an antibody that is suitable for Western blot may not be optimal for ICW. Generally, ICW assays yield comparable data with the added advantage of increased throughput, improved accuracy, and reduction in sample preparation (Aguilar, Zielnik, Tracey, & Mitchell, 2010; Hoffman, Moerke, Hsia, Shamu, & Blenis, 2010; Ma et al., 2017; Paguirigan, Puccinelli, Su, & Beebe, 2010).

1.4. Selection of antibodies

Western blot, Simple Western, Flow Cytometry, and the In-Cell Western assay all are reliant on the availability of high-quality antibodies. For Flow cytometry and ICW, since proteins are not separated on a gel, specific and non-specific signals cannot be differentiated based on association with a reference protein of the correct molecular weight, therefore monoclonal primary antibodies are preferred to reduce nonspecific binding and background noise. However, although monoclonal antibodies are preferable due to batch-to-batch consistency, they may be too specific in some cases, or more susceptible to loss of a single epitope due to processing. Also, consider that Western blot antibodies recognize fully denatured proteins and may not bind to the same protein in its native folded state, therefore antibodies selected for ICW assay development should be compatible with immunohistochemistry staining, providing optimal recognition of the protein epitope in context to the cellular environment. Thus, the ICW assay is expected to be readily adapted to the use of any antibody that is suitable for immunofluorescence-based detection. Another consideration is whether the target signal can be observed in multiple cell compartments, i.e., nuclear or cytosol, or both after membrane permeabilization (Kunze & Berger, 2015; Lu et al., 2021).

1.5. Advantages of In-Cell Western assay

The main advantage of the ICW technique over traditional Western blot is the throughput and processing advantages. Another is that once a suitable antibody is identified to monitor the protein of interest the analysis can be readily transferred to other cell lines providing a facile method to analyze many cell lines using one technique. The ICW assay normalizes to cell number using a cell or DNA stain which increases the accuracy by correcting for well-to-well variations in cell number. Near-IR dyes offer several advantages over fluorescent dyes in the visible spectrum such as reduced light scattering, reduced background noise due to autofluorescence, and less signal loss due to absorbance ion (Kunze & Berger, 2015; Lu et al., 2021). Thus, the near-IR dyes provide robust signals at lower antibody dilutions compared to using dyes in the visible spectrum. Protein degradation is measured in situ, in fixed cells providing a read-out of activity in a relevant cellular context. Experimental variability is reduced due to reduced processing which allows for better uniformity across experiments, and improved reproducibility. This usually results in lower standard deviations compared to Western blots. The ICW assay thus provides results comparable to Western blot and is well suited as a rapid primary screening assay for PROTAC or degrader lead optimization.

1.6. Limitations of In-Cell Western assay

The key limitations of this assay are identification of a suitable antibody and the preference for adherent cells for accurate results due to the number of washing steps and signal linearity to cell number. Notably, the development of the assay for a new lab can be cumbersome. The initial setup may require optimization of the blocking buffer, fixative, and permeabilization reagents. The ICW assay is a well-based, rather than a cell-based assay and does not provide high-resolution information that would be better suited for high-content or confocal imaging. This technique may be suitable for the study of phospho-proteins or other proteins from whole tissue, as long as proteins are in high enough levels to be detected from the cellular material in a single well. Therefore, low abundance proteins may be difficult to detect (Aguilar et al., 2010).

2. A case studies using the ICW to evaluate CDK6 degraders

2.1. A linearity study for CDK6 in SK-MEL-2 cells

The first step in the ICW assay development is to identify the linear range of the assay. This is performed prior to evaluating potential PROTACs or degraders. The reason a linearity study is performed first is to determine the correct conditions for the assay in terms of how many cells to use per well and what antibody dilution is required so that the results based on the fluorescent readout are directly proportional to the concentration of the protein being measured. Thus, the linearity study provides the relationship between cell number and signal intensity, and a clear understanding of the dynamic range of the assay. Measurements within the linear range parameters will provide an accurate assay. Measurements outside of the linear range will be poorly reproducible, i.e., above the linear range strong signals will saturate the capacity of the detector and produce a signal less than expected, and measurements below the linear range will be difficult to detect from background.

2.2. Materials and equipment

• ICW assay plate: Greiner Bio-One cell culture microplate 96 well half area black, flat clear bottom: Greiner Bio-one μClearTM 96-well half area microplate, Cat# 675090

• Cell line SK-MEL-2 (RRID:CVCL_0069); Source: ATCC

• Primary antibody: anti-CDK6 rabbit mAb: (Abcam, Cat# ab124821)

• 4% Para-Formaldehyde solution in PBS (Thermo Scientific, Cat #J19943-K2)

• 0.1% Triton X-100 TBS solution (Triton X-100 (Roche, Cat #14327722, 10xTBS solution: Bio-Rad Cat #1706435)

• Washing Solution: 0.1% Tween-20 in 1XTBS (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR Intercept Blocking Buffer (LI-COR Cat #927–60001)

• 0.2% Tween-20 LI-COR Intercept Blocking Buffer (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR IRDye 800CW secondary antibodies (LI-COR Cat #926–32232)

• LI-COR CellTag 700 stain (LI-COR Cat #926–410909)

• Odyssey^® CLx Infrared Imaging System (LI-COR Biosciences)

• LI-COR Image Studio Software^® version 5.2

Note: LI-COR has recently updated their analysis software to Empiria Studio Software^® providing post-processing, Data Integrity Software for quantitative protein expression analysis which now includes the In-Cell Western^™ assay.

2.3. Step-by-step method for a CDK6 linearity study

Step 1. Seed SK-MEL-2 Cells

The first step is to determine the number of cells to seed in each 96-well plate and the primary antibody dilution factor.

1. Seed SK-MEL-2 cells in Greiner Bio-one 96-well half area black, clear bottom plates at 15K, 10K, 8K, 6K, 4K and 2K cells per well with 100μL, in duplicate. Then the cells are incubated at 37°C, in a 5% CO₂ atmosphere overnight. See Fig. 1 which shows a plate map for plating the cells

2. Out-lying wells are filled with PBS alone to reduce edge effects due to evaporation during the incubation; they are not used for analysis

Fig. 1.

The plate map for seeding SK-MEL-2 cells. Out-lying wells are not used for analysis due to edge effects and are filled with PBS to help avoid evaporation.

Note: The outer wells can be used but care must be taken as to how the cells are handled so that edge effects are minimized or eliminated (Lundholt, Scudder, & Pagliaro, 2003).

Step 2. Fix Cells

In this step the cell media will be removed, and the cells will then immediately be treated with fixative reagent to fix the cells.

1. Dump the cell media, then using the multichannel pipette carefully add 50μL of 4% paraformaldehyde to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate at room temperature on the bench top for 20min without shaking or agitation

Step 3. Permeabilize Cells

In this step the fixative solution will be removed and replaced with permeabilization solution.

1. Make a 0.1% Triton X-100 TBS solution: add 1mL of Triton X-100 (Cat #14327722) and 100mL of 10xTBS solution (Cat #1706435), then add H₂O to 1000mL

2. Dump the fixing solution into an appropriate chemical waste container. Caution to avoid skin and eye contact with the solution which contains formaldehyde

3. Using a multichannel pipette, add to each well with 50μL of 0.1% Triton X-100 TBS which has been warmed to room temperature. Carefully add the solution down the sides of the wells to avoid detaching the cells

4. Gently agitate the plate using a flatbed plate shaker at room temperature for 5min

5. Repeat these steps, removing the old solution and adding with 50μL of new 0.1% Triton X-100 TBS four more times, then removing the final solution carefully without disturbing the cells

Step 4. Block Cells

In this step blocking buffer will be added to the cells followed by incubation.

1. Dump the permeabilization solution, then add 50μL of Intercept Blocking Buffer carefully to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate for 1h at room temperature with moderate shaking on a flatbed plate shaker to allow for blocking

Step 5. Incubate with Primary Antibody

In this step the primary antibody for CDK6 will be diluted and added to the plate.

1. Make a 0.2% Tween-20 in Intercept Blocking Buffer solution by adding 100μL of Tween-20 (Bio-Rad, Cat #1706531) to 50mL of LI-COR Intercept Blocking Buffer (Cat #927–60,001).

2. Dilute primary antibody with 0.2% Tween-20 in Intercept Blocking Buffer at 1:1000, 1:3000, 1:5000 and 1:10000

3. Dump the Intercept Blocking Buffer. To the cell seeding testing wells, add 15μL of each dilution of the primary antibody to the testing wells and 0.2% Tween-20 in Intercept Blocking Buffer to background control wells following the Scheme in Fig. 2. i.e., Columns 2 and 3 are duplicate wells for dilution 1:1000, columns 4 and 5 are duplicate wells for dilution 1:3000, columns 6 and 7 are duplicated wells for dilution 1:5000, columns 8 and 9 are duplicate wells for dilution 1:10,000. Columns 10 and 11 will be used to measure background from the secondary antibody alone

4. Incubate with primary antibody for 2.5h at room temperature (also can incubate the plate overnight at 4°C) with gentle shaking

Fig. 2.

The plate map for the primary and secondary antibody dilution. The out-lying wells are not used for the assay.

Step 6. Wash Plate

1. Make a 1 × TBS solution with 0.1% Tween-20 washing solution: add 1mL of Tween-20 and 100mL of 10 × TBS, then add H₂O to 1000mL

2. Dump the primary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

3. Gently agitate the plate on a flatbed plate shaker for 5min

4. Repeat washing steps 4 more times

Step 7. Incubate with Secondary Antibody

In this step the secondary antibody conjugated to the IRDye^® will be incubated with the cells for visualization. The secondary antibody used to visualize the protein is diluted 800 times as recommended in the LI-COR Application Guide “In-Cell Western Assay Development Handbook.” The LI-COR CellTag 700 stain is used to normalize to cell number and is diluted 1000 times as recommended in the LI-COR Application Guide “In-Cell Western Assay Development Handbook.”

1. Dilute the fluorescently labeled secondary antibody IRDye 800CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800 and LI-COR CellTag 700 stain at 1:1000

2. Dump the last wash solution, add 15μL of diluted secondary antibody solution with CellTag 700 stain to testing wells. To the background wells, add 15μL of secondary antibody solution without CellTag 700 stain. Follow the Scheme shown in Fig. 2

3. Incubate for 1h at room temperature with gentle shaking. Protect the plate from light during incubation

4. The Background (BG) wells (See Fig. 2) will contain only secondary antibody IRDye 800CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800

Step 8. Wash Plate

1. Dump the secondary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min. Protect plate from light during washing

3. Repeat washing steps 4 more times

Step 9. Scan

1. Dump the final wash solution, turn the plate upside down and tap or gently blot with paper towels to remove traces of the wash buffer

2. Clean the bottom plate surface with 70% alcohol and the scanning bed with a moist, lint-free paper towel or cloth

3. Scan the plate on the LI-COR Odyssey^® CLx Infrared Imaging System with the 700 and 800nm channel. Select macro plate and use 3 or 3.5mm for Focus Offset

Step 10. Data Analysis

In this step, the plates are analyzed using the In-Cell Western procedure of the Image Studio Software^®.

1. Analyze data with Image Studio Software^® under the In-Cell Western Software protocol (see Fig. 3).

2. Normalize the data to CellTag 700 stain (see Fig. 4).

3. Select Well Types (Fig. 4) then define the Testing Wells (see Fig. 5).

4. The qualification data was exported in Excel by selecting the Grid sheet tab and selecting the report to define the quantification data to export (see Fig. 6).

5. The linearity of cell number and primary Ab dilution was graphed in Prism GraphPad 8 (see Fig. 7).

6. The LI-COR CellTag 700 stain was used to define the signal linearity between the fluorescent signal vs. the cell number. The CellTag 700 stain data was graphed to show the linearity of the CellTag 700 stain to the cell number (see Fig. 8A).

7. The ratio of primary Ab to background was generated (see Fig. 8B) to show how strong the signal would be over the background

8. The image of the primary antibody signal, the CellTag 700 stain, and the merged image can also be saved at this step. (See Fig. 9).

9. The final conditions chosen for the degradation study of CDK6 after PROTAC treatment will be seeding 8000 cells per well and a 1:5000 primary antibody dilution

Fig. 3.

Screen shot of LI-COR Image Studio Software^® image under the In-Cell Western protocol.

Fig. 4.

In-Cell Western analysis normalized to CellTag 700.

Fig. 5.

In-Cell Western analysis to define the well types.

Fig. 6.

In-Cell Western analysis under the Grid Sheet tab to define the Qualification data to export to an Excel file.

Fig. 7.

Signal Linearity. (A) In-Cell Western Analysis to show the signal linearity between the fluorescent signal from the secondary antibody IRDye 800CW to the cell number normalized to CellTag 700 stain. (B) The partial graph of signal linearity is used to calculate a more accurate R value.

Fig. 8.

(A) Signal linearity for CellTag 700 stain at various cell densities without addition of secondary antibody IRDye 800CW. (B) Signal to Noise ratio calculated for various primary antibody dilutions and cell number.

Fig. 9.

(A) Plate Image for visualization of CDK6 with secondary antibody IRDye 800CW. (B) Plate Image for visualization with CellTag 700 stain. (C) Plate Image A and B merged.

2.4. Treatment study for CDK6 protein levels in SK-MEL-2 cells

Cyclin dependent kinases (CDKs) are important for their role in regulating cell cycle. Cyclin D-dependent kinases CDK4 and CDK6, and the peptide inhibitor CDK4/6 inhibitor p16^INK4 play a key role in regulating mammalian cell proliferation from G1 into S phase of the cell-division cycle (Sherr, Beach, & Shapiro, 2016). FDA approved CDK4/6 inhibitors such as palbociclib, ribociclib, and abemaciclib are important therapeutic agents for hormone-positive breast cancer and are standard or care therapies in combination with endocrine therapy as first- and second-line treatment for metastatic breast cancer (George, Qureshi, Omene, Toppmeyer, & Ganesan, 2021). Importantly CDK4/6 inhibition is being explored as potential treatments for other cancers, including acute myeloid leukemia, AML (Uras, Sexl, & Kollmann, 2020) and Ph⁺ acute lymphoblastic leukemia (Ph⁺ ALL) (De Dominici et al., 2020; Porazzi, De Dominici, Salvino, & Calabretta, 2021). Development of targeted CDK6 degraders is of high therapeutic interest as potential therapies against resistance, i.e., for breast cancer, (Li et al., 2022) and to improve safety and efficacy (De Dominici et al., 2020).

This protocol demonstrates the use of the ICW assay for evaluation of CDK6 protein levels after pharmacological treatment. The assay is important for use in lead identification and optimization due to the ease of throughput compared to a Western blot. Adherent cells versus suspension cells were utilized in this ALL drug discovery effort since this provided a facile assay for prioritizing and optimizing potential PROTAC analogs during lead optimization which were then confirmed in Western blot assays in Ph+ALL suspension cells (De Dominici et al., 2020).

2.5. Materials and equipment

• ICW assay plate: Greiner Bio-One cell culture microplate 96 well half area black, flat clear bottom: Greiner Bio-one μClearTM 96-well half area microplate, Cat# 675090

• Cell line SK-MEL-2 (RRID:CVCL_0069); Source: ATCC

• Primary antibody: anti-CDK6 rabbit mAb: (Abcam, Cat# ab124821)

• 4% Para-Formaldehyde solution in PBS (Thermo Scientific, Cat #J19943-K2)

• 0.1% Triton X-100 TBS solution (Triton X-100 (Roche, Cat #14327722, 10xTBS solution: Bio-Rad Cat #1706435)

• Washing Solution: 0.1% Tween-20 in 1XTBS (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR Intercept Blocking Buffer (LI-COR Cat #927–60001)

• 0.2% Tween-20 LI-COR Intercept Blocking Buffer (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR IRDye 800CW secondary antibodies (LI-COR Cat #926–32232)

• LI-COR CellTag 700 stain (LI-COR Cat #926–410909)

• Odyssey^® CLx Infrared Imaging System (LI-COR Biosciences)

• LI-COR Image Studio Software^® version 5.2

Note: LI-COR has recently updated their analysis software to Empiria Studio Software^® providing post-processing, Data Integrity Software for quantitative protein expression analysis which now includes the In-Cell Western™ assay.

2.6. Step-by-step method to evaluate CDK6 levels after compound treatment

Step 1. Seed Cells and Apply Treatment to Cells

1. Seed 8K cells based on the linearity assay data performed a day before. For a typical plate configuration see plate Map in Fig. 10

2. Dissolve compound in culture medium with 0.1% DMSO for all the compound treatment start from 10μM or 1uM to 0μM with 3-fold dilution factor

3. Apply 100μL of compound solution to cells for 24h

Fig. 10.

Plate Map for ICW Analysis.

Step 2. Fix Cells

In this step the cell media will be removed, and the cells will then immediately be treated with fixative reagent to fix the cells.

1. Dump the cell media, then using the multichannel pipette carefully add 50μL of 4% paraformaldehyde to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate at room temperature on the bench top for 20min without shaking or agitation

Step 3. Permeabilize Cells

In this step the fixative solution will be removed and replaced with permeabilization solution.

1. Make a 0.1% Triton X-100 TBS solution: add 1mL of Triton X-100 (Cat #14327722) and 100mL of 10xTBS solution (Cat #1706435), then add H₂O to 1000mL

2. Dump the fixing solution into an appropriate chemical waste container. Caution to avoid skin and eye contact with the solution which contains formaldehyde

3. Using a multichannel pipette, carefully add to each well add 50μL of 0.1% Triton X-100 TBS which has been warmed to room temperature. Carefully add the solution down the sides of the wells to avoid detaching the cells

4. Gently agitate the plate using a flatbed plate shaker at room temperature for 5min

5. Repeat these steps, removing the old solution and adding with 50μL of new 0.1% Triton X-100 TBS four more times, then removing the final solution carefully without disturbing the cells

Step 4. Block Cells

In this step blocking buffer will be added to the cells followed by incubation.

1. Dump the permeabilization solution, then add 50μL of Intercept Blocking Buffer carefully to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate for 1h at room temperature with moderate shaking on a flatbed plate shaker to allow for blocking

Step 5. Incubate with Primary Antibody

In this step the primary antibody for CDK6 will be diluted and adding to the plate.

1. Dilute primary antibody with 0.2% Tween-20 in Intercept Blocking Buffer at 1:5000

2. Dump the Intercept Blocking Buffer, then add 15μL of diluted primary antibody solution to the testing wells from column 2 to 11 and 0.2% Tween-20 in Intercept Blocking Buffer to background control wells of column 12 following Fig. 11 scheme

3. Incubate with primary antibody for 2.5h at room temperature or alternatively incubate the plate overnight at 4°C with gentle shaking

Fig. 11.

Plate Map for Compound Treatment, Primary and secondary Antibody.

Step 6. Wash Plate

1. Dump the primary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min 3. Repeat washing steps 4 more times

3. Repeat washing steps 4 more times

Step 7. Incubate with Secondary Antibody

In this step the secondary antibody conjugated to the IRDye^® will be incubated with the cells for visualization. The secondary antibody used to visualize the protein is diluted 800 times as recommended in the LI-COR Application Guide “In-Cell Western Assay Development Handbook.” The LI-COR CellTag 700 stain is used to normalize to cell number and is diluted 1000 times as recommended in the LI-COR Application Guide “In-Cell Western Assay Development Handbook.”

1. Dilute the fluorescently labeled secondary antibody IRDye 800 CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800 and LI-COR CellTag 700 stain at 1:1000

2. Dump the last wash solution, add 15μL of diluted secondary antibody solution with CellTag 700 stain to testing wells from column 2 to 11. To the background wells of column 12, add 15μL of secondary antibody solution without CellTag 700 stain

3. Incubate for 1h at room temperature with gentle shaking. Protect the plate from light during incubation

4. The Background (BG) wells (See Fig. 11 for Plate MAP) will contain only secondary antibody IRDye 800 CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800

Step 8. Wash Plate

1. Dump the secondary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min. Protect plate from light during washing

3. Repeat washing steps 4 more times

Step 9. Scan

1. Dump the final wash solution, turn the plate upside down and tap or gently blot with paper towels to remove traces of the wash buffer

2. Clean the bottom plate surface with 70% alcohol and the scanning bed with a moist, lint-free paper towel or cloth

3. Scan the plate on the LI-COR Odyssey^® CLx Infrared Imaging System with the 700 and 800nm channel. Select macro plate and use 3 or 3.5mm for Focus Offset

Step 10. Data Analysis

In this step, the plates are analyzed used the In-Cell Western procedure of the Image Studio Software^®.

1. Analyze data with Image Studio Software^® under the In-Cell Western Software protocol (see Fig. 3).

2. Normalize the data to CellTag 700 stain (see Fig. 4).

3. Select Well Types under the analysis tab (Fig. 4) to define the testing well types (see Fig. 5).

4. The qualification data is exported in Excel under the Grid sheet tab by selecting report (see Fig. 6).

5. The data for the degradation study was graphed in Prism GraphPad^® (see Fig. 13).

6. The CellTag 700 stain data was also graphed to show the cell viability (see Fig. 14).

Fig. 13.

ICW Analysis of CDK6 levels after pharmacological treatment. Treatment of nine compounds monitoring CDK6 protein levels at 9 different concentrations. (A) Compounds tested starting from 10μM include AC-03-092, a resynthesis of BSJ-03-123 used as a reference compound and Palbociclib. (B) Compounds tested starting from 1μM include YX-2-107 a reported CDK6 degrader.

Fig. 14.

ICW Analysis of Live SK-MEL-2 cells after pharmacological treatment. Treatment of nine compounds monitoring SK-MEL-2 cell number at 9 different concentrations.

2.7. Discussion

A Western blot analysis was performed using the Abcam CDK6 antibody in the SK-MEL-2 cells. The antibody shows a single, strong band at 36kDa for CDK6 confirming that this is a suitable antibody for the study (see Fig. 12).

Fig. 12.

Western Blot for CDK6 in SK-MEL-2 cells.

In this example nine compounds were evaluated for their effect on CDK6 protein levels and cell viability (cell number) after a 24-h treatment. SK-MEL-2 cells were used for the analysis due to their cell growth characteristics, the good CDK6 expression levels, and that they are adherent cells. The compounds were tested in 3 plates at 9 different concentrations either starting at 10μM (Fig. 13A) or 1μM (Fig. 13B) plus a DMSO control using a 3-fold dilution scheme performed in duplicate. The data is normalized to cell number using the CellTag 700 stain. If a treatment time course is desired, multiple treatment times simply require duplicate plates incubated at various time points. The Odyssey^® CLx Infrared Imaging System scans one plate at a time and takes about 5min per plate. The data is analyzed in Prism GraphPad^® and provides 9-point dose response curves which simplify comparison for SAR analysis during lead optimization. In addition, since the analysis method depends on cell number, this parameter is evaluated and provides an additional readout on the cytotoxicity of compounds used under the treatment conditions (Fig. 14).

Visualization of the plates can be saved during the analysis step (Fig. 15). In this example, plates 1–3 contain compounds. The primary antibody is visualized in green depicting the protein levels, the cell stain is in red depicting cell number, and the merged plate is shown. The highest compound concentration is on the left, with a 3-fold decrease in concentration, BG depicts an empty well used for the background signal.

Fig. 15.

ICW analysis visualizing the plates. Plates 1–3 in green are for visualization of the primary antibody, in red for visualization of the cell stain depicting cell number, and the merged plate. Compounds are plated in duplicate with the highest concentration on the left. BG is an empty well for background reading.

A more detailed analysis is shown for YX-2-107, a reported CDK6 degrader,³¹ AC-03-092, a re-synthesis of the reported CDK6 degrader BSJ-03-123, (Brand et al., 2019) and palbociclib, a clinically used CDK4/6 inhibitor (Fig. 16). The adherent CDK6 expressing SK-MEL-2 cell line used for PROTAC optimization is very useful for prioritizing potent degraders for confirmation in Western blot assays in more relevant cell lines, patient derived cells, or tissues after pharmacological treatment in vivo (De Dominici et al., 2020). The two reported degraders clearly show potent concentration dependent degradation in the analysis. Interestingly, Palbociclib shows an up regulation of CDK6 expression levels after a 24-h treatment, which is consistent with the compensatory increase in CDK6 expression seen with clinically used CDK4/6 kinase inhibitors (Li et al., 2018; Yang et al., 2017).

Fig. 16.

ICW Analysis highlighting 2 PROTACs and the CDK4/6 inhibitor, palbociclib. (A) YX-02-107 and AC-03-092 (BSJ-03-123) are reported CDK6 degraders and Palbociclib is a clinically approved CDK4/6 inhibitor. In this cell line and under these analysis conditions YX-02-107 (DC50=0.01μM) and AC-03-092 (DC50=0.06μM) show potent CDK6 degradation consistent with their reported values. Palbociclib shows a modest increase in CDK6 levels after a 24-h treatment. (B) Compound treatment did not reduce cell number. (C) Structures of the compounds are shown.

A Western blot was then performed to confirm the results of the ICW assay (Fig. 17). SK-MEL-2 cells are treated with YX-2-107 for 24h at six different concentrations followed by Western blot analysis. This analysis confirms the dose responsive effect of YX-2-107 on CDK6 protein levels which is consistent with the ICW assay data thus confirming the use of this technique to provide a higher through-put assay for lead optimization.

Fig. 17.

Western blot analysis of CDK6 protein levels in SK-MEL-2 cells after 24h treatment. SK-MEL-2 cells were treated with YX-02-107 at six different concentrations and Palbociclib at three different concentrations for 24h prior to Western blot analysis. CDK6 has a molecular weight of 36kDa, β-Actin (molecular weight=42kDa) was chosen as a loading control, expression levels of this protein do not vary drastically due to cellular treatment. Protein loading=15μg.

3. A case studies using the ICW to evaluate PARP1 degraders in SNU719 cells

3.1. A linearity study for PARP1 degraders in SNU719 cells

The linearity study was performed in the SNU719 as the first step in the ICW assay development to identify the linear range of the assay. This is performed prior to evaluating potential PROTACs or degraders.

3.2. Materials and equipment

• ICW assay plate: Greiner Bio-One cell culture microplate 96 well half area black, flat clear bottom: Greiner Bio-one μClear^™ 96-well half area microplate, Cat# 675090

• Cell line SNU-719 (RRID:CVCL_5086) human gastric carcinoma, EBV-associated gastric cancer

• Primary Ab Anti-PARP1 antibody (Abcam cat # 191217)

• 4% Para-Formaldehyde solution in PBS (Thermo Scientific, Cat #J19943-K2)

• 0.1% Triton X-100 TBS solution (Triton X-100 (Roche, Cat #14327722, 10xTBS solution: Bio-Rad Cat #1706435)

• Washing Solution: 0.1% Tween-20 in 1XTBS (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR Intercept Blocking Buffer (LI-COR Cat #927–60001)

• 0.2% Tween-20 LI-COR Intercept Blocking Buffer (Tween-20: Bio-Rad, Cat #1706531)

• LI-COR IRDye 800CW secondary antibodies (LI-COR Cat #926–32232)

• LI-COR CellTag 700 stain (LI-COR Cat #926–410909)

• Odyssey^® CLx Infrared Imaging System (LI-COR Biosciences)

• LI-COR Image Studio Software^® version 5.2

3.3. Step-by-step method for a linearity study for PARP1

Step 1. Seed SNU719 Cells

The first step is to determine the number of cells to seed in each 96-well plate and the primary antibody dilution factor.

1. Seed SNU719 cells in 96-well half area black, clear bottom plates at 15K, 10K, 8K, 6K, 4K and 2K cells per well with 100μL, in duplicate. Then the cells are incubated at 37°C, in a 5% CO₂ atmosphere overnight. See Fig. 18 which shows a plate map for plating the cells

Fig. 18.

The plate map for seeding SK-MEL-2 cells. Columns 1, 2, 11 and 12 are not used for analysis due to edge effects and are filled with PBS to help avoid evaporation.

Step 2. Fix Cells

Remove cell media and immediately treat the cells with fixative reagent to fix the cells.

1. Dump the cell media, then using the multichannel pipette carefully add 50μL of 4% paraformaldehyde to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate at room temperature on the bench top for 20min without shaking or agitation

Step 3. Permeabilize Cells

In this step the fixative solution will be removed and replaced with permeabilization solution.

1. Dump the fixing solution into an appropriate chemical waste container. Caution to avoid skin and eye contact with the solution which contains formaldehyde

2. Using a multichannel pipette, carefully add to each well add 50μL of 0.1% Triton X-100 TBS which has been warmed to room temperature. Carefully add the solution down the sides of the wells to avoid detaching the cells

3. Gently agitate the plate using a flatbed plate shaker at room temperature for 5min

4. Repeat these steps, removing the old solution and adding with 50μL of new 0.1% Triton X-100 TBS four more times, then removing the final solution carefully without disturbing the cells

Step 4. Block Cells

In this step blocking buffer will be added to the cells followed by incubation.

1. Dump the permeabilization solution, then add 50μL of Intercept Blocking Buffer carefully to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate for 1h at room temperature with moderate shaking on a flatbed plate shaker to allow for blocking

Step 5. Incubate with Primary Antibody

In this step the primary antibody for PARP1 will be diluted and added to the plate.

1. Dilute primary antibody with 0.2% Tween-20 in Intercept Blocking Buffer at 1:200, 1:300, 1:500

2. Dump the Intercept Blocking Buffer. To the cell seeding testing wells, add 15μL of each dilution of the primary antibody to the testing wells and 0.2% Tween-20 in Intercept Blocking Buffer to background control wells following the Scheme in Fig. 19, i.e., Columns 3 and 4 are duplicate wells for dilution 1:200, columns 5 and 6 are duplicate wells for dilution 1:300, columns 7 and 8 are duplicated wells for dilution 1:500, Columns 9 and 10 will be used to measure background from the secondary antibody alone

3. Incubate with primary antibody for 2.5h at room temperature (also can incubate the plate overnight at 4°C) with gentle shaking

Fig. 19.

The plate map for the primary and secondary antibody dilution. The out-lying wells are not used for the assay.

Step 6. Wash Plate

1. Dump the primary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Gently agitate the plate on a flatbed plate shaker for 5min

3. Repeat washing steps 4 more times

Step 7. Incubate with Secondary Antibody

1. Dilute the fluorescently labeled secondary antibody IRDye 800CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800 and LI-COR CellTag 700 stain at 1:1000

2. Dump the last wash solution, add 15μL of diluted secondary antibody solution with CellTag 700 stain to testing wells. To the background wells, add 15μL of secondary antibody solution without CellTag 700 stain. Follow the Scheme shown in Fig. 19

3. Incubate for 1h at room temperature with gentle shaking. Protect the plate from light during incubation

Step 8. Wash Plate

1. Dump the secondary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min. Protect plate from light during washing

3. Repeat washing steps 4 more times

Step 9. Scan

1. Dump the final wash solution, turn the plate upside down and tap or gently blot with paper towels to remove traces of the wash buffer

2. Clean the bottom plate surface with 70% alcohol and the scanning bed with a moist, lint-free paper towel or cloth

3. Scan the plate on the LI-COR Odyssey^® CLx Infrared Imaging System with the 700 and 800nm channel. Select macro plate and use 3 or 3.5mm for Focus Offset

Step 10. Data Analysis

In this step, the plates are analyzed using the In-Cell Western procedure of the Image Studio Software^®.

1. Analyze data with Image Studio Software^® under the In-Cell Western Software protocol (see Fig. 3).

2. Normalize the data to CellTag 700 stain (see Fig. 4).

3. Select Well Types (Fig. 4) then define the Testing Wells (see Fig. 5).

4. The qualification data was exported in Excel by selecting the Grid sheet tab and selecting the report to define the quantification data to export (see Fig. 6).

5. The linearity of cell number and primary Ab dilution was graphed in Prism GraphPad 8^® (see Fig. 20A).

6. The LI-COR CellTag 700 stain was used to define the signal linearity between the fluorescent signal vs. the cell number. The CellTag 700 stain data was graphed to show the linearity of the CellTag 700 stain to the cell number (see Fig. 20B).

7. The ratio of primary Ab to background was generated (see Fig. 20C) to show how strong the signal would be over the background

8. The image of the primary antibody signal, the CellTag 700 stain, and the merged image can also be saved at this step. (See Fig. 21).

9. The final conditions chosen for the degradation study of PARP1 after PROTAC treatment will be seeding 8000 cells per well and a 1:300 primary antibody dilution

Fig. 20.

Signal Linearity. (A) Signal linearity normalized to CellTag 700. (B) Signal linearity for CellTag700 at various cell densities. (C) Signal to Noise ratio calculated for various primary antibody dilutions and cell number.

Fig. 21.

(A) Plate Image for visualization of PARP1 with secondary antibody IRDye 800CW. (B) Plate Image for visualization with CellTag 700. (C) Plate Image A and B merged.

3.4. Treatment study for PARP1 in SNU719 cells

Poly (ADP-ribose) polymerases (PARPs) are a family of related enzymes that catalyze the transfer of ADP-ribose to target proteins and play important roles in modulation of chromatin structure, transcription, and DNA repair. Tumors defective in homologous recombination mechanisms that rely on PARP-mediated DNA repair for survival are sensitive to PARP inhibition (Bryant et al., 2005; Farmer et al., 2005; Morales et al., 2014). PARP1 was the first member of the PARP family to be characterized (Amé et al., 1999; Amé, Spenlehauer, & de Murcia, 2004). The enzymatic function of PARP1 is to catalyze poly-ADP-ribosylation or PARylation (Chambon, Weill, & Mandel, 1963). When DNA damage is detected PARP1 binds to DNA and activates PARylation using NAD⁺ as a substrate (McCabe et al., 2006). PARP inhibition has been well studied and has led to at least four PARP inhibitors which have received FDA approval including olaparib, niraparib, rucaparib, and talazoparib. Several others are in clinical development, i.e., veliparib and pamiparib. PARP inhibitors have two key mechanisms of action, firstly since PARP inhibitors can block enzymatic activity preventing PARylation and secondly, they can also “trap” PARP enzymes on DNA resulting in increased genomic instability and cytotoxicity (Pommier, O’Connor, & de Bono, 2016). The extent of PARP trapping has been associated with various levels of cytotoxicity but may also contribute to both cancer cell as well as healthy bone marrow toxicity (Hopkins et al., 2019). PARP inhibitors have different “trapping” potencies; however, they exhibit comparable tumor growth inhibition suggesting that “trapping” may not solely be required for highly efficacious therapeutic agents.

PARP1 is also hyperactivated in other non-oncology related pathological conditions (Berger et al., 2018; Kam et al., 2018) and over-stimulation of the catalytic activity of PARP1 may result in NAD⁺ depletion leading to a cascade of events resulting in cell death. PARP1 plays an important role in multiple neurologic diseases through mediation of caspase-independent cell death, designated as parthanatos, which may be related to signaling through PAR polymers (Wang, Dawson, & Dawson, 2009). PARP1 PROTACs are being studied to address mechanistic questions to uncouple trapping from enzymatic inhibition, (Wang et al., 2019) and to study the effects due to enzymatic inhibition and scaffolding effects of PARP1 during PARP1 hyperactivation.

Epstein-Barr Virus (EBV) establishes lifelong latency in human B-cells (Price & Luftig, 2015) and is associated with several malignancies including nasopharyngeal carcinoma and Burkett’s Lymphoma. EBV triggers the activity of the chromatin modifying enzyme PARP1, through the action of the viral protein LMP1. PARP1 and PARylation play a key role in EBV latency maintenance and lytic reactivation. Inhibition of PARylation and PARP1 inhibition in EBV-positive B-cells holds significant therapeutic potential and PARP1 PROTACs are valuable tool compounds to study PARP’s essential role in regulating EBV chromatin structure and latent gene expression (Morgan et al., 2022).

This protocol demonstrates how the ICW assay can be used to facilitate the lead optimization of PARP1 PROTACs and is exemplified using experimental PARP1 degraders.

Step 1. Seed Cells and Apply Treatment to Cells

1. Seed 8K cells based on the linearity assay data performed a day before. Follow the Plate Map for treatment shown in Fig. 10

2. Dissolve compound in culture medium with 0.1% DMSO for all the compound treatment start from 10μM with 3-fold dilution factor, plus the DMSO control

3. Apply 100μL of compound solution to cells for 24h

Step 2. Fix Cells

Remove cell media and immediately treat the cells with fixative reagent to fix the cells.

1. Dump the cell media, then using the multichannel pipette carefully add 50μL of 4% paraformaldehyde to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate at room temperature on the bench top for 20min without shaking or agitation

Step 3. Permeabilize Cells

In this step the fixative solution will be removed and replaced with permeabilization solution.

1. Dump the fixing solution into an appropriate chemical waste container. Caution to avoid skin and eye contact with the solution which contains formaldehyde

2. Using a multichannel pipette, carefully add to each well add 50μL of 0.1% Triton X-100 TBS which has been warmed to room temperature. Carefully add the solution down the sides of the wells to avoid detaching the cells

3. Gently agitate the plate using a flatbed plate shaker at room temperature for 5min

4. Repeat these steps, removing the old solution and adding with 50μL of new 0.1% Triton X-100 TBS four more times, then removing the final solution carefully without disturbing the cells

Step 4. Block Cells

In this step blocking buffer will be added to the cells followed by incubation.

1. Dump the permeabilization solution, then add 50μL of Intercept Blocking Buffer carefully to each well by pipetting the solution down the sides of the wells to avoid detaching the cells

2. Incubate for 1h at room temperature with moderate shaking on a flatbed plate shaker to allow for blocking

Step 5. Incubate with Primary Antibody

In this step the primary antibody for PARP will be diluted and adding to the plate.

1. Dilute primary antibody with 0.2% Tween-20 in Intercept Blocking Buffer at 1:300

2. Dump the Intercept Blocking Buffer. Add 15μL of each dilution of the primary antibody to the testing wells and 0.2% Tween-20 in Intercept Blocking Buffer to background control wells following the Scheme in Fig. 19

3. Incubate with primary antibody for 2.5h at room temperature or alternatively incubate the plate overnight at 4°C with gentle shaking

Step 6. Wash Plate

1. Dump the primary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min. Protect plate from light during washing

3. Repeat washing steps 4 more times

Step 7. Incubate with Secondary Antibody

In this step the secondary antibody conjugated to the IRDye^® will be incubated with the cells for visualization.

1. Dilute the fluorescently labeled secondary antibody IRDye 800CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800 and LI-COR CellTag 700 stain at 1:1000

2. Dump the last wash solution, add 15μL of diluted secondary antibody solution with CellTag 700 stain to testing wells. To the background wells, add 15μL of secondary antibody solution without CellTag 700 stain

3. Incubate for 1h at room temperature with gentle shaking. Protect the plate from light during incubation

4. The Background (BG) wells (See Fig. 19 for Plate MAP) will contain only secondary antibody IRDye 800CW in 0.2% Tween-20 Intercept Blocking Buffer at 1:800

Step 8. Wash Plate

1. Dump the secondary antibody solution, add 50μL of 1 × TBS with 0.1% Tween-20 washing solution. Carefully add the solution down the sides of the wells to avoid detaching cells

2. Allow plate to shake on a plate shaker for 5min. Protect plate from light during washing

3. Repeat washing steps 4 more times

Step 9. Scan

1. After the final wash, remove wash solution completely from wells. Turn the plate upside down and tap or blot gently on paper towels to remove traces of wash buffer

2. Clean the bottom plate surface and the scanning bed with moist, lint-free paper

3. Scan the plate in LI-COR Odyssey^® CLx Infrared Imaging System with the 700 and 800nm channel. Select macro plate and use 3 or 3.5mm for Focus Offset

Step 10. Data Analysis

In this step, the plates are analyzed used the In-Cell Western procedure of the Image Studio Software^®.

1. Analyze data with Image Studio Software^® under the In-Cell Western Software protocol (see Fig. 3).

2. Normalize the data to CellTag 700 stain (see Fig. 4).

3. Select Well Types under the analysis tab (Fig. 4) to define the testing well types (see Fig. 5).

4. The qualification data is exported in Excel under the Grid sheet tab by selecting report (see Fig. 6).

5. The data for the degradation study was graphed in Prism GraphPad^® (see Fig. 23A).

6. The CellTag 700 stain data was also graphed to show the cell viability (see Fig. 23B).

Fig. 23.

ICW Analysis for 3 compounds in SNU719 cells. (A) AC-030-086 and AC-03-088 are PARP1 degraders and JG-01-070 is a control compound. In this cell line under these analysis conditions AC-03-086 (DC50=42nM) and AC-03-088 (DC50=65nM) show potent PARP1 degradation. The control compound JG-01-070 did not decrease PARP1 levels. (B) AC-03-088 treatment reduced cell number at concentrations around 1–3μM.

3.5. Discussion

A Western blot analysis was performed using the Abcam PARP1 antibody in the SNU719 cells. The antibody shows a strong band at 113kDa for PARP1, however there are additional non-specific bands observed in this Western blot which may reduce the signal when evaluated in the ICW (Fig. 22). If the signal for the positive control is greater than the negative control, and the assay has a Z’ factor score > 0.5, the nonspecific bands are typically not a concern.

Fig. 22.

Western Blot with PARP1 in SNU-719.

For this example, three compounds were evaluated for their effect on PARP1 protein levels and cell viability (cell number) after a 24-h treatment in human EBV-associated gastric cancer SNU719 cells. The compounds were tested in a single 96 well plate starting at 10μM (Fig. 23A) using a 3-fold dilution scheme performed in duplicate. The data is analyzed in Prism GraphPad^® and provides 9-point dose response curves to compare compound treatment. Pharmacological effect on cell number is evaluated and provides a readout on the cytotoxicity of compounds (Fig. 23B). The results from this study show that compounds AC-03-086 and AC-03-088 degrade PARP1 levels close to 100% compared to JG-01-070 which shows minimal or no-degradation in the SNU719 cells. Interestingly, AC-03-088 affects cell viability at concentrations around 1–3μM.

Visualization of the plate can be saved during the analysis step (Fig. 24). In this example, the primary antibody is visualized in green depicting the protein levels after compound treatment, the cell stain is in red depicting cell number, and the merged plate is shown. The highest compound concentration is on the left, with a 3-fold decrease in concentration, BG depicts an empty well used for the background signal.

Fig. 24.

ICW analysis visualizing the plates. Plate in green is for visualization of the primary antibody, in red for visualization of the cell stain depicting cell number, and the merged plate. Compounds are plated in duplicate with the highest concentration on the left. BG is an empty well for background reading.

Western blot assays are being completed to further confirm PARP1 degradation in the same cell lines used for the ICW assay. In summary, this study provides a method to quickly identify PARP1 PROTACs and prioritize analogs for Western blot and additional characterization studies.

4. Expected outcomes

The ICW assay is shown to be a facile method to analyze protein levels after PROTAC or degrader treatment. The results are generally similar to Western blot analysis. The quality of the antibody used in the analysis is important and it should be optimized for binding to the cellular epitope conformation, not the de-natured protein.

5. Quantification and statistical analysis

The data is generated in duplicate, and replicates can be performed in different experiments to provide biological replicates. The data is analyzed in Prism GraphPad^® to facilitate calculation of standard deviation or standard error of the mean.

5.1. Advantages

The major advantage of this technique is the ability to process compounds in 96 well plates to increase throughput compared to Western blot. The method is less time-consuming facilitating analysis of 20–30 compounds in several days’ time, meeting the needs of medicinal chemistry lead optimization. The method is also readily transferable to different cell lines or cell types without the need of extensive optimization or cell engineering.

5.2. Limitations

The major limitation is the need for a good antibody. Antibodies which are non-specific reduce the signal to noise in the assay and reduce accuracy and reliability.

Abbreviation

PROTAC

proteolysis targeting chimera