K.M.R, S.D.M, M.U. and D.L.D are all employees of Promega Corporation

1. Endpoint degradation studies with HiBiT CRISPR target proteins in lytic format with optional cell viability fluorescence analysis
<ol>
	<li>Preparation and plating of mammalian adherent or suspension cell line
	<ol>
		<li>Adjust the cell density to 2.22 x 105/mL by dilution in appropriate cell media used for passaging and cell growth.</li>
		<li>Dispense cells into plates with a minimum of 3 wells per experimental and control condition. Dispense 90 &#181;L (20,000 cells) per well of cell suspension into 96-well white plates. For 384-well format, dispense 36 &#181;L (8,000 cells) per well of cell suspension into 384-well white plates.</li>
	</ol>
	</li>
	<li>Preparation and addition of compounds
	<ol>
		<li>Prepare serially diluted PROTAC or degrader test compound plates at 1,000x final concentration in 100% DMSO. Then dilute it to 10x final concentration in the cell culture medium. Add an equal volume of DMSO to the medium, to be used as a no compound DMSO control.</li>
		<li>For 96-well format add 10 &#181;L of 10x compound and control solutions to 90 &#181;L of cells. For 384-well format add 4 &#181;L of 10x compound and control solutions to 36 &#181;L of cells.</li>
		<li>Incubate the plates in an incubator at 37 &#176;C and 5% CO2 for the desired amount of time or in conditions that are optimal for their growth. 
		NOTE: As this is an endpoint assay, testing of multiple time points will require preparation of separate degradation plates for each time point, as described in above step 1.1.2. Incubation times to detect compound-mediated degradation are highly variable and are also likely dependent on compound concentration. Suggested initial time points would be 6 h and 24 h.</li>
		<li>If measuring endpoint luminescent detection without the optional cell viability measurement, proceed directly to step 1.3 below. If performing multiplexing with cell viability measurement, proceed to the next section 1.4 below.</li>
	</ol>
	</li>
	<li>Lytic measurement of cells
	<ol>
		<li>Immediately prior to HiBiT lytic measurements, prepare 2x lytic detection reagent by adding 20 &#181;L of lytic substrate and 10 &#181;L of LgBiT protein per every 1 mL of the lytic buffer. Prepare enough 2x detection reagent for number of wells to be assayed, including extra volume to account for pipetting error (i.e., number of wells + 10%).</li>
		<li>Add prepared lytic detection reagent to cells. For 96-well format, add 100 &#181;L of 2x lytic detection reagent to each well containing 100 &#181;L of cells. For 384-well format, add 40 &#181;L of 2x lytic detection reagent to each well containing 40 &#181;L of cells. Mix the plate on a microplate vortex mixer for 10-20 min at 350 rpm.</li>
		<li>Measure luminescence on a luminometer capable of reading luminescence in a 96- or 384-well plate.</li>
	</ol>
	</li>
	<li>Optional cell viability multiplexing 
	NOTE: This step is performed using a commercially available CellTiter-Fluor (CTF) kit (see Table of Materials).
	<ol>
		<li>30-40 min prior to the desired endpoint measurement, prepare a 6x cell viability detection reagent solution by adding 10 &#181;L of the substrate to 2 mL of the assay Buffer. Prepare enough 6x reagent for each well to be assayed, including extra volume for pipetting error (i.e., number of wells + 10%).</li>
		<li>Add the prepared reagent to wells. For 96-well format add 20 &#181;L of 6x reagent to each well already containing a 100 &#181;L volume. For 384-well format add 8 &#181;L of 6x reagent to each well containing 40 &#181;L of cells. Mix briefly on a microplate vortex mixer, then incubate the plate for 30 min in a 37 &#176;C incubator.</li>
		<li>At the desired endpoint of measurement (i.e., 6 or 24 h post treatment, step 1.2.3), measure fluorescence on an instrument capable of reading fluorescence (380-400nmEx/505nmEm) in 96- or 384- well format.</li>
		<li>Prepare 2x lytic detection reagent by adding 20 &#181;L of lytic substrate and 10 &#181;L of LgBiT protein per 1 mL of lytic buffer. Prepare enough 2x detection reagent for number of wells to be assayed, including extra volume to account for pipetting error (e.g., number of wells + 10%).</li>
		<li>Add the prepared lytic detection reagent to wells. For 96-well format add 120 &#181;L of 2x lytic detection reagent to each well already containing a 120 &#181;L volume. For 384-well format add 48 &#181;L of 2x lytic detection reagent to each well already containing a 48 &#181;L volume. Mix the plate on a microplate vortex mixer for 10-20 min.</li>
		<li>Measure luminescence on a luminometer capable of reading luminescence in 96- or 384- well plates.</li>
	</ol>
	</li>
	<li>Quantification of degradation and cell viability
	<ol>
		<li>Average the relative light units (RLU) from the DMSO control at the measured time point. Use this value as the baseline protein level of the target to calculate fractional degradation by normalizing all other treatments tested at this same time point to this value. For example, if the average RLU for the DMSO control wells at 6 h was 10,000, and the RLU for a given compound treatment at 6 h was 5,000, the fractional degradation would be calculated as 5,000 &#247; 10,000 = 0.5 (Equation 1). 
		Equation 1: <img alt="Equation " src="/files/ftp_upload/61787/61787equ01.jpg" /></li>
		<li>Determine the percentage degradation from Fractional RLU: 
		Equation 2: <img alt="Equation " src="/files/ftp_upload/61787/61787equ02.jpg" /></li>
		<li>Plot fractional RLU or % degradation at specific time points to rank the activity of compounds.</li>
		<li>Optionally, analyze relative fluoresecence unit (RFU) data&#160;for the cell viability assay measurement by comparing the values from all treatments to the DMSO control. If a significant decline in RFU is observed for any treatment relative to the DMSO control, the degradation data can be additionally normalized to the cell viability assay data to determine changes in protein level relative to losses in cell viability.</li>
	</ol>
	</li>
</ol>
2. Real-time kinetic degradation of HiBiT CRISPR target proteins and optional cell viability luminescence assay
NOTE: The ability to perform kinetic screening and degradation requires LgBiT protein co-expression in the cell, which has been described previously18,19,63. This can be achieved via transient transfection of a LgBiT vector, use of BacMam LgBiT, or by performing HiBiT CRISPR insertion into a LgBiT stable cell line.
<ol>
	<li>Plating of adherent cell lines.
	<ol>
		<li>Remove medium from cell flask by aspiration, wash cells with DPBS, dissociate cells with 0.05% trypsin-EDTA, and allow cells to dissociate from the flask bottom. For suspension cell lines, proceed to section 2.2.</li>
		<li>Neutralize trypsin using serum-containing cell culture medium, mix to collect and resuspend cells, and transfer cell suspension to a conical tube.</li>
		<li>Spin down cells at 125 x g for 5 min. Discard the cell culture medium and resuspend in an equal volume of fresh cell culture medium.</li>
		<li>Plate cells into assay plates with a minimum of triplicate wells per experimental and control condition. For 96-well format count to estimate cell density, adjust density to 2 x 105 cells/mL in assay medium and dispense 100 &#181;L (20,000 cells) per well in a 96-well plate. For 384-well format count to estimate cell density, adjust density to 4.44 x 105 cells/mL in the assay medium and dispense 18 &#181;L (8,000 cells) per well.</li>
		<li>Incubate plates at 37 &#176;C, 5% CO2 overnight or in conditions which are optimal for their growth.</li>
	</ol>
	</li>
	<li>Plating of suspension cells
	<ol>
		<li>Adjust cell density to 2.22 x 105 cells/mL in CO2-independent medium supplemented with 10% FBS and 1x Endurazine (1:100 dilution of the stock reagent).</li>
		<li>Plate cells into assay plates with a minimum of 3 wells per experimental and control condition. For 96-well format dispense 90 &#181;L (20,000 cells) per well. For 384-well format dispense 36 &#181;L (8,000 cells) per well. 
		NOTE: For suspension cell lines which have low signal to background (S:B) luminescence, e.g., when working with CRISPR pools rather than clones, it is possible to increase the luminescence by increasing the number of cells plated, up to 100,000 cells/well in 96-well format, or 40,000 cells/well in 384-well format.</li>
	</ol>
	</li>
	<li>Kinetic degradation assays using HiBiT CRISPR cells expressing LgBiT
	<ol>
		<li>For suspension cells already containing Endurazine, which was included at the plating step in 2.2., proceed directly to step 2.3.3. For adherent cell lines prepare Nano-Glo Endurazine solution. For 96-well format, prepare a 1x solution of Endurazine by diluting stock reagent 1:100 into CO2-independent medium supplemented with 10% FBS. For 384-well format, prepare a 2x solution of Endurazine by diluting stock reagent 1:50 into CO2-independent medium supplemented with 10% FBS.</li>
		<li>Add Endurazine solution to each well of adherent cells. For 96-well format aspirate medium and add 90 &#181;L of 1x Endurazine solution. For 384-well format, add 18 &#181;L of 2x Endurazine solution to 18 &#181;L of cells. Do not aspirate medium as the degradation assay is performed in a 50:50 mixture of culture medium and CO2 independent medium in 384-well format.</li>
		<li>Incubate suspension or adherent cell plates containing Endurazine for 2.5 h in an incubator at 37 &#176;C and 5% CO2 to allow luminescence to equilibrate.</li>
		<li>Prepare a 10x concentration of test PROTAC titration in CO2-independent medium and add 10 &#181;L to each well of 96-well plate or 4 &#181;L for 384-well plate. For compounds with unknown efficacy, a final concentration of 1-10 &#181;M at the highest point is recommended as a starting point.</li>
		<li>Collect kinetic measurements of luminescence in luminometer pre-equilibrated to 37 &#176;C for a period between 0-48 h. The time increments of measurement can be customized for each experiment, but a recommended initial experiment would be luminescence measurements every 5-15 min for 24 h or the desired amount of time.</li>
	</ol>
	</li>
	<li>Optional cell viability same-well multiplex analysis after final kinetic measurement 
	NOTE: This assay is performed with a commercially available CellTiter-Glo (CTG) kit (see Table of Materials).
	<ol>
		<li>Equilibrate CTG reagent to room temperature.</li>
		<li>Following degradation measurement at the last time point of the kinetic analysis, add 100 &#181;L (96-well plate) or 40 &#181;L (384-well plate) of the reagent per well of the plate, and mix on a plate shaker at 500-700 rpm (96-well plate) or a microplate vortex mixer (384-well plate) for 5 min.</li>
		<li>Incubate the plate at room temperature for 30 min to allow for cell lysis and quenching of HiBiT signal.</li>
		<li>Measure total luminescence on a luminometer by following manufacturer&#8217;s recommendation.</li>
	</ol>
	</li>
	<li>Quantification of kinetic degradation profiles
	<ol>
		<li>Using the kinetic luminescence measurements collected, normalize the raw RLUs for each PROTAC concentration to the replicate averaged DMSO condition at every time point to account for changes in free furimazine concentration over time. Calculate Fractional RLU using Equation 1. 
		Equation 1: <img alt="Equation " src="/files/ftp_upload/61787/61787equ01.jpg" /></li>
		<li>From the degradation curves, fit a single-component exponential decay model using Equation 2 to the initial degradation portion of each curve to the point where the data reaches a plateau. 
		NOTE: It may be helpful to exclude from the fit the first few data points as there may be a brief lag before degradation is observed. 
		Equation 2: <img alt="Equation " src="/files/ftp_upload/61787/61787equ03.jpg" /></li>
		<li>From Equation 2, determine the parameter &#411;, which represents the degradation rate constant and the Plateau, which represents the lowest amount of protein remaining.</li>
		<li>Calculate Dmax, which is the maximum fractional amount of degraded protein and is calculated as 1-Plateau.</li>
		<li>Plot Dmax for each concentration of PROTAC to determine a time-independent degradation potency curve.</li>
		<li>Determine the Dmax50 value for the plot in 2.3.5 to analyze efficacy of compounds. 
		NOTE: To determine a DC50 at a specific time point, plot the calculated percent degradation for each concentration at the chosen time. This can be specified as DC50 t=4 h or DC50 t=12 h.</li>
	</ol>
	</li>
</ol>

Promega Corporation is the commercial owner by assignment of patents of the HiBiT and NanoLuc technologies and applications.

We present here two methods of screening degradation compound activity in either endpoint lytic format or live-cell kinetic mode. These approaches are based upon the same luminescent measurement principles yet provide different levels of detail and understanding. The choice of either approach will likely be dependent upon screening goals and size of compound library. For large compound screening decks or primary screens to observe any detectable degradation, endpoint lytic screening offers sensitive and efficient high-throughput compatibility where other endpoint approaches, such as western blot or mass spectrometry can be either impractical or difficult to adapt14. A starting point for these screens could be performed with a limited number of concentrations and timepoints. Recommended initial concentrations to test are in the range of 100 nM-10 &#181;M, to account for initial degraders having low potency, poor permeability, or in some cases with highly potent compounds, the presence of a hook effect. It is further recommended that a minimum of two different time points should be tested to establish early onset degradation at 4-6 h and latent or sustained degradation at 18-24 h. Compounds that exhibit high degradation potency and on-target mechanism are readily observed within a 4-6 h timeframe, whereas degradation or apparent protein loss observed only at later time points could be due to a variety of mechanisms. It is highly recommended to monitor cell viability at both early and late time points, such that protein loss can be uncoupled from loss due to cell death. Similar to any type of luminescent or fluorescent assay, there is a potential for compounds within libraries to interfere or inhibit signal, therefore, orthogonal follow-up experiments with lead compounds using unrelated fusions or alternative approaches to monitoring protein level will be important for assessing that loss of RLU in these assays is associated directly with target protein degradation.
The ability to screen in live cell kinetic format over extended periods of time relies heavily on the assay signal to background (S:B). Factors that contribute to S:B include the expression level of the target protein itself, which can span several orders of magnitude, the efficiency of LgBiT expression in the cell line chosen for peptide insertion, and the availability of the tagged target for complementation in its various native complexes. We have established a general cutoff requirement consisting of a S:B of 15 to successfully measure degradation in kinetic mode with either Endurazine or Vivazine. The S:B is determined by measuring the baseline signal of the HiBiT-edited cells co-expressing LgBiT relative to unedited parental cells expressing LgBiT alone in the presence of either Endurazine or Vivazine live cell substrates. Vivazine will produce a higher luminescent signal but will decay faster than Endurazine and may limit signal acquisition to 24 hours or less. Furthermore, S:B can also be highly dependent on whether CRISPR pools or clones are used. For targets in cell lines which are more amenable and have high efficiency for CRISPR/Cas9 engineering, a heterogeneous CRISPR pool population of edited cells may have sufficient S:B for kinetic analysis. For targets in more difficult cell lines where less efficient genomic integration via CRISPR results in pools with low S:B, isolating CRISPR clones might be necessary to enrich edited populations and achieve a sufficiently high S:B for kinetic analysis. For any of these scenarios, if S:B is less than 15 with either Endurazine or Vivazine substrates, endpoint lytic screening is advised.
For better understanding and characterization of compounds, including determination of a degradation profile with quantitative parameters, real-time kinetic analysis in live cells is the recommended screening approach14,18. Like endpoint analysis discussed above, initial kinetic screening can be done with a limited number of concentrations in the range of 100nM-10&#181;M in high-throughput fashion. In 384-well format, over 100 compounds can readily be screened in triplicate at one concentration on a single plate. The resulting degradation profiles will provide guidance not only on the extent of degradation observed, but the rate of degradation, the duration of degradation, and potential recovery of the protein14,18 (Figure 2 and Figure 3). The shapes of the degradation profile also yield valuable information. Specific and potent degraders often show an initial rapid loss of the target protein to a plateau in a matter of hours18,53, whereas other mechanisms such as transcriptional feedback or compound toxicity typically result in more linear loss of the protein over time. These details and nuances are missed with endpoint lytic analysis, and with real-time analysis over 24-48 hours, one does not have to predict the time to capture the true Dmax within sets of new or unknown compounds.
Real-time kinetics also allow&#160;for efficient dose response screening to better understand compound efficacy, how compound concentration impacts initial degradation rate, and offers possibilities to rank compounds based on more than one parameter. Classical measurements of degradation potency involve DC50 calculations at a specific point in time based on apparent degradation maxima. In contrast, our kinetic approach to evaluate potency incorporates the true degradation maximum at each concentration regardless of when it occurs in time18. We call this measurement of kinetic degradation potency, the Dmax5018. Analysis in this fashion accounts for compounds that may initiate degradation more slowly at lower concentrations and therefore take a longer time post-treatment to reach their Dmax. It can be especially informative to rank compounds on both degradation rate and Dmax. For the most potent degraders this will further differentiate slow, but potent degraders from those which are both fast and potent. Together, both lytic and live cell kinetic screening utilizing HiBiT CRISPR cell lines are powerful approaches which yield a more comprehensive picture of targeted protein degradation, compound function, and enable the screening process from initial activity assessment to downstream chemical optimization through enhancement of key degradation parameters.

Targeted protein degradation has emerged as one of the fastest growing areas in small molecule drug discovery, bolstered greatly by the therapeutic success of immunomodulatory molecular glue compounds (e.g., IMiD) for cancer treatment, and promising early clinical trial data of Proteolysis Targeting Chimera compounds1,2,3,4,5,6,7,8,9,10,11,12. Targeted protein degradation compounds function by bringing into proximity a target protein with E3 ligase machinery proteins1,2,3,4,5,6,7,8,9,10,11,12. This compound-induced recruitment of the target protein to the E3 ligase leads to the target protein ubiquitination and degradation via the ubiquitin proteasomal pathway (UPP)1,2,3,4,5,6,7,8,9,10,11,12. Historically, small molecule drug discovery screening programs have relied upon initial biochemical assays to assess activity and rank order compounds. This, however, has presented a significant challenge for targeted protein degraders whose ultimate activity, degradation via the proteasome, is dependent upon a cascade of cellular events1,2,4,5,6,11,12,13,14,15,16,17,18. The multiple pathways and complexity of protein complexes required for the successful target degradation necessitate cellular assay approaches for early screening and triaging of initial compounds. Currently, the availability of technologies to monitor target protein degradation in a high-throughput fashion in the context of the cellular environment is severely lacking14. Here we will present protocols for real-time kinetic live cell or endpoint lytic degradation activity assessment using CRISPR/Cas9 endogenously tagged HiBiT target cell lines18,19,20 to monitor the loss of the target protein via luminescent measurement after treatment with degrader compounds10,11,18,19.
To achieve successful degradation of therapeutic targets and to expand the druggable proteome, numerous approaches and types of degraders have emerged which can target a broad range of proteins for destruction, including those localized at or in the plasma membrane, lysosomes, mitochondrial membranes, cytoplasm, and the nucleus21&#8211;57. The two primary classes of compounds most extensively studied are molecular glues and protein targeting cimeras2,4,5,6,7,12,26. Molecular glues are monovalent, thus typically smaller in size, and facilitate a novel protein:protein interaction interface with a target protein upon binding to an E3 ligase component2,12,26. They are most commonly degraders that bind to the Cereblon (CRBN) E3 ligase component2,12,26,55,56,57. Recently though exciting new examples utilizing other E3 ligase machinery such as DCAF1558,59,60 and CDK/Cyclin recruitment to DDB145 show the potential for expansion of this class of compounds. In contrast, PROTACs are larger, bivalent molecules, consisting of a target binding ligand, most often an inhibitor, bridged via a chemical linker to a E3 ligase handle1,3,4,5,7,13. As such, these compounds are capable of direct binding to both the E3 ligase and the target protein1,3,4,5,7,13. Numerous proteins have been shown to be degraded via these bivalent molecules, and the most used E3 ligase handles recruit either CRBN or Von Hippel Lindau (VHL)1,3,4,5,7,13. However, the number of available handles for E3 ligase recruitment in chimeras targeting proteolysis design is rapidly growing, expanding the capabilities of this class of compounds with the potential to degrade diverse target classes as well as enhance cell- or tissue-type specificity24,48,61,62. Combined with the minimal requirement to engage a target protein, even with marginal affinity, degradation compounds hold promise for expanding the druggable proteome.
Characterizing the cellular dynamics of protein loss, as well as potential protein recovery post-treatment, is critical for understanding degradation compound function and efficacy. While it is possible to study endogenous protein level changes in relevant cellular systems with western blot antibody assays or mass spectrometry, these approaches are difficult to adapt to high-throughput screening formats, have limited quantification capability, or ability to measure kinetic changes at many timepoints14. To address these challenges, we have developed a plate-based cellular luminescent system for monitoring changes in endogenous protein levels, which utilizes genomic insertion via CRISPR/Cas9 of the 11 amino acid tag, HiBiT, to the loci of any key degradation targets18,19,20. This peptide complements with high affinity to its binding partner, LgBiT, to produce bright luminescence in the presence of its substrate18,19,20,63, thereby making these tagged endogenous proteins luminescent in cells or lysates18,19,20,63. The relative light units (RLUs) measured with a luminometer instrument are directly proportional to the tagged target protein levels18,19,20,63. With the development of stabilized luciferase substrates, real-time kinetic protein level measurements over 24-48 h time frames are possible18,53,64. This allows for the determination of a complete degradation profile for any given target at any given compound concentration, including quantitative analysis of initial degradation rate, degradation maximum (Dmax), and recovery after compound treatment18,53. If screening large libraries of degradation compounds, however, endpoint analysis can also readily be performed in 384-well format at various drug concentrations and designated times.
The protocols presented in this manuscript represent cellular screening strategies for targeted protein degradation compounds, applicable for all types of degraders. The use of HiBiT CRISPR cell lines along with these protocols, however, are not limited to protein degradation, rather they are general tools for monitoring any endogenous target protein level which could be modulated post-treatment to study impact of compounds or even resistance mechanisms20,65,66. A pre-requisite for these luminescent-based detection methods is a CRISPR endogenously tagged HiBiT target cell line, which is critical as it enables sensitive luminescent detection, while still maintaining endogenous target expression and native promoter regulation18,19,20. Significant advances have been made in utilizing CRISRP/Cas9 for insertion of genomic tags, particularly in scalability20 and with the high sensitivity of detection, in various formats including CRISPR pools or clones with either heterozygous or homozygous allelic insertions18,19,20. Use of exogenous expression of HiBiT or other reporter fusions in cells in lieu of endogenous tagging is possible, but significant caution should be taken using systems with protein overexpression14,18. These can lead to artefacts in understanding true compound potency and protein recovery dynamics14,18, including potential transcriptional feedback loops activated post target degradation. In addition, early stage compounds with low potency could be missed, and present themselves as false negatives in screening. As protein loss could result from compound-induced toxicity and cell death, the protocols described here contain highly recommended, but optional cell viability luminescent or fluorescent assays paired with the degradation protocol. There are two major sections to the protocol, lytic endpoint, and live cell kinetic screening. Within each of those sections, options are included for multiplexed cell viability measurements in endpoint or kinetic formats. Monitoring the changes of the tagged endogenous protein requires complementation with LgBiT in cells. Therefore, the kinetic screening section references important protocols for the introduction of this, which can be achieved via transient or stable expression and is essential for performing the live cell luminescent measurements. All approaches presented here allow rapid rank ordering and activity assessment of compounds, enabling early stage compound screening efforts and more rapid identification of lead degraders.
This protocol is designed for the study of degradation compounds in conjunction with a HiBiT CRISPR cell line. Protocols for generation of HiBiT CRISPR insertions for numerous targets have been outlined in several recent publications18,19,20.

<table>
	<tbody>
		<tr>
			<td>CellTiter-Glo 2.0 reagent</td>
			<td>Promega</td>
			<td>G9241</td>
			<td>Cell Viability luminescent assay</td>
		</tr>
		<tr>
			<td>CellTiter-Fluor Cell Viability Assay</td>
			<td>Promega</td>
			<td>G6080</td>
			<td>Cell Viability fluorescent assay</td>
		</tr>
		<tr>
			<td>CO2-independent medium</td>
			<td>ThermoFisher</td>
			<td>18045-088</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma Aldrich</td>
			<td>D2650</td>
			<td>For compound dilution and control</td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Gibco</td>
			<td>14190</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Seradigm</td>
			<td>89510-194</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>HEK293 LgBiT stable cell line</td>
			<td>Promega</td>
			<td>N2672</td>
			<td>For complementation with HiBiT to generate luminescence</td>
		</tr>
		<tr>
			<td>HiBiT CRISPR mammalian cell line</td>
			<td>Promega</td>
			<td></td>
			<td>https://www.promega.com/crispr-tpd</td>
		</tr>
		<tr>
			<td>Hygromycin B solution</td>
			<td>Gibco</td>
			<td>10-687-010</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>LgBiT BacMam</td>
			<td>Promega</td>
			<td>CS1956C01</td>
			<td>For complementation with HiBiT to generate luminescence</td>
		</tr>
		<tr>
			<td>LgBiT Expression Vector</td>
			<td>Promega</td>
			<td>N2681</td>
			<td>For complementation with HiBiT to generate luminescence</td>
		</tr>
		<tr>
			<td>Luminometer Plate Reader</td>
			<td></td>
			<td></td>
			<td>Luminomenter capable of measuring luminescence and fluorescence (e.g. GloMax Discover System, Promega GM3000)</td>
		</tr>
		<tr>
			<td>NanoGlo Endurazine live cell substrate</td>
			<td>Promega</td>
			<td>N2570</td>
			<td>Kinetic HiBiT reagent</td>
		</tr>
		<tr>
			<td>NanoGlo Vivazine live cell substrate</td>
			<td>Promega</td>
			<td>N2580</td>
			<td>Kinetic HiBiT reagent</td>
		</tr>
		<tr>
			<td>NanoGlo HiBiT Lytic Detection system</td>
			<td>Promega</td>
			<td>N3030</td>
			<td>Enpoint lytic HiBiT reagent</td>
		</tr>
		<tr>
			<td>Opti-MEM Reduced Serum Medium, no phenol red (ThermoFisher)</td>
			<td>ThermoFisher</td>
			<td>11058-021</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Tissue culture plates, white, 96 well plate</td>
			<td>Costar</td>
			<td>3917</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Tissue culture plates, white, 384 well plate</td>
			<td>Corning</td>
			<td>3570</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Trypsin/EDTA</td>
			<td>Gibco</td>
			<td>25300</td>
			<td>Cell culture</td>
		</tr>
	</tbody>
</table>

high-throughput cellular profiling of targeted protein degradation compounds using hibit crispr cell lines

Targeted protein degradation compounds, including molecular glues or proteolysis targeting chimeras, are an exciting new therapeutic modality in small molecule drug discovery. This class of compounds induces protein degradation by bringing into proximity the target protein and the E3 ligase machinery proteins required to ubiquitinate and ultimately degrade the target protein through the ubiquitin-proteasomal pathway (UPP). Profiling of target protein degradation in a high-throughput fashion, however, remains highly challenging given the complexity of cellular pathways required to achieve degradation. Here we present a protocol and screening strategy based on the use of CRISPR/Cas9 endogenous tagging of target proteins with the 11 amino acid HiBiT tag which complements with high affinity to the LgBiT protein, to produce a luminescent protein. These CRISPR targeted cell lines with endogenous tags can be used to measure compound induced degradation in either real-time, kinetic live cell or endpoint lytic modes by monitoring luminescent signal using a luminescent plate-based reader. Here we outline the recommended screening protocols for the different formats, and&#160;also describe the calculation of key degradation parameters of rate, Dmax, DC50, Dmax50, as well as multiplexing with cell viability assays. These approaches enable rapid discovery and triaging of early stage compounds while maintaining endogenous expression and regulation of target proteins in relevant cellular backgrounds, allowing for efficient optimization of lead therapeutic compounds.

To demonstrate single concentration endpoint lytic degradation analysis, several CDK target proteins; CDK2, CDK4, CDK7, and CDK10 were endogenously tagged with HiBiT at their C-terminus in HEK293 cells and treated with a 1 &#181;M concentration of the pan-kinase Cereblon-based PROTAC, TL12-18654 (Figure 1A). The level of CDK protein was measured at different time points and the fractional RLU relative to the DMSO control was determined (Figure 1A). Each CDK protein showed different extents of degradation in response to the compound treatment and the various time points (Figure 1A). To understand how CDK proteins compared to each other directly in terms of protein loss, the fractional RLUs in Figure 1A were calculated as total % degradation and plotted for each time point in Figure 1B. This shows that even at early time points, 2 or 4 h, some of the CDK family members show high levels of degradation which continue to trend upwards over time (Figure 1B).
To demonstrate kinetic degradation analysis, each of the BET family member proteins; BRD2, BRD3, and BRD4 were endogenously tagged with HiBiT at their N-terminus in HEK293 cells stably expressing the LgBiT protein18. These were then treated with three different concentrations of the pan-BET PROTACs; the Cereblon-based dBET650 (Figure 2A) and the VHL-based ARV-77141 (Figure 2B). Kinetic measurements were collected over a 24 h period, and from the profiles at each concentration, the differences in BET family member response are readily apparent. The ability of BRD2 to initiate a more rapid recovery response post-degradation compound treatment (Figure 2A,B) has been previously observed with other pan-BET PROTACs and is likely due to a transcriptional feedback response competitive to the degradation process18.
Both endpoint and kinetic analysis can be done with full compound dose response treatments. Shown in Figure 3 are kinetic dose response degradation profiles of treatment of Ikaros/IKZF1-HiBiT CRISPR Jurkat cells stably expressing LgBiT protein with four different molecular glue compounds2,26,55,57; lenalidomide (Figure 3A), iberdomide (CC-220) (Figure 3B), thalidomide (Figure 3C), and pomalidomide (Figure 3D). These degraders show significant differences in degradation response amongst the compounds as well as across the concentration series (Figure 3).
To quantitatively assess degradation and rank order the compounds in Figure 3 the dose response profiles were used to calculate key degradation parameters including the degradation rate (Figure 4A), Dmax (Figure 4B), and Dmax50 values (Figure 4B). These analyses show that iberdomide (CC-220) and pomalidomide have very similar rapid initial degradation rates (Figure 4A), yet iberdomide (CC-220) has the highest potency as has been previously seen in orthogonal studies55,57 (Figure 4B). Since Iberdomide exhibits such high potency, and all concentrations tested show greater than 50% degradation, the Dmax50 value obtained for Iberdomide represents an estimate based on the limitation in accurately fitting the data. From the graphs in Figure 3C,D and Figure 4B, neither lenalidomide nor thalidomide degrade the Ikaros/IKZF1 target to completion at the highest concentrations tested. Due to very little degradation observed with thalidomide, the degradation traces could not be accurately fit to an exponential decay model, therefore, degradation rate was not quantified for this treatment. For the most potent degrader, iberdomide (CC-220)55,57 (Figure 4B). Cell viability multiplex assays showed no loss in cell viability for the concentrations tested (Figure 4C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61787/61787fig01.jpg" /> 
Figure 1: CDK endpoint degradation and toxicity with pan-kinase PROTAC, TL12-18654.&#160;(A) Select panel of endogenous CDK target proteins fused with HiBiT on the C-terminus via CRISPR/Cas9 and assessed for degradation with 1 &#181;M TL12-186 PROTAC54 at 2 h, 4 h, 8 h, and 24 h treatment. Values are represented as Fractional RLU relative to a DMSO control measured at each time point. Error bars represent SD of the mean of 3 technical replicates. (B) Percent degradation of panel of CDK target proteins calculated from (A) representing the amount of degradation of each family member observed at 2, 4, 8, and 24 h time points. Error bars represent SD of the mean of 3 technical replicates. <a href="https://www.jove.com/files/ftp_upload/61787/61787fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61787/61787fig02.jpg" /> 
Figure 2: Profiling kinetic degradation selectivity of BET family members with BET degraders, dBET650 and ARV-77141.&#160;Kinetic degradation profiles of endogenous BET family members, BRD2, BRD3, and BRD4, tagged with HiBiT on the N-terminus via CRISPR/Cas9 with treatment of single concentrations of 1 nM (left), 10 nM (middle), or 100 nM (right) dBET650 (A) or ARV-77141 (B) PROTACs. Values are represented as Fractional RLU calculated from a DMSO control at each kinetic time point. Error bars represent SD of the mean of 4 technical replicates. <a href="https://www.jove.com/files/ftp_upload/61787/61787fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61787/61787fig03.jpg" /> 
Figure 3: Live cell kinetic degradation dose response profiles of Ikaros/IKZF1-HiBiT with a molecular glue panel2,26,55,57.&#160;Jurkat cells stably expressing the LgBiT protein were engineered using CRISPR/Cas9 to tag the C-terminus of Ikaros/IKZF1 with the HiBiT peptide. Cells were treated with an 8 point dose response concentration series including DMSO of four different molecular glue compounds2,26,55,57: (A) lenalidomide, (B) iberdomide (CC-220), (C) thalidomide, or (D) pomalidomide. Luminescence was measured every 5 min for a total of 19.5 h. Relative light unit (RLU) data from (A-D) were converted to fractional RLU as described in Step 2.4.1 and graphed as a function of time. Error bars represent SD of 3 technical replicates. <a href="https://www.jove.com/files/ftp_upload/61787/61787fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61787/61787fig04.jpg" /> 
Figure 4: Calculation of degradation rate and Dmax50 for Ikaros/IKZF1-HiBiT, and multiplexing cell health assays.&#160;Kinetic degradation data from Figure 3 were used to calculate quantitative degradation parameters. (A) Degradation rates and (B) degradation maximum values (Dmax) are graphed at each drug concentration for the indicated molecular glue compounds2,26,55,57. (B) Dmax50 values for each compound were calculated using a dose-response model with constrained Hill slope of 1, which can be used to rank order degradation compounds for a target. (C) Cell viability assays with the iberdomide (CC-220)55,57 degradation dose response from Figure 3B was performed as an endpoint measurement upon completion of the kinetic degradation measurements. Error bars represent SD of 3 technical replicates. <a href="https://www.jove.com/files/ftp_upload/61787/61787fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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High-Throughput Cellular Profiling of Targeted Protein Degradation Compounds Using HiBiT CRISPR Cell Lines

This protocol describes the quantitative luminescent detection of protein degradation kinetics in living cells that have been engineered using CRISPR/Cas9 to express antibody free endogenous protein detection tag fused to a target protein. Detailed instructions for calculating and obtaining quantitative degradation parameters, rate, Dmax, DC50, and Dmax50 are included.

Targeted protein degradation compounds, including molecular glues or proteolysis targeting chimeras, are an exciting new therapeutic modality in small ...

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9.2K Views.  Promega Corporation. This protocol describes the quantitative luminescent detection of protein degradation kinetics in living cells that have been engineered using CRISPR/Cas9 to express antibody free endogenous protein detection tag fused to a target protein. Detailed instructions for calculating and obtaining quantitative degradation parameters, rate, Dmax, DC50, and Dmax50 are included.

Video: High-Throughput Cellular Profiling of Targeted Protein Degradation Compounds Using HiBiT CRISPR Cell Lines

Using the Dot Assay to Analyze Migration of Cell Sheets

Although complex organisms appear static, their tissues are under a continuous turnover. As cells age, die, and are replaced by new cells, cells move within tissues in a tightly orchestrated manner. During tumor development, this equilibrium is disturbed, and tumor cells leave the epithelium of origin to invade the local microenvironment, to travel to distant sites, and to ultimately form metastatic tumors at distant sites. The dot assay is a simple, two-dimensional unconstrained migration assay, to assess the net movement of cell sheets into a cell-free area, and to analyze parameters of cell migration using time-lapse imaging. Here, the dot assay is demonstrated using a human invasive, lung colony forming breast cancer cell line, MCF10CA1a, to analyze the cells' migratory response to epidermal growth factor (EGF), which is known to increase malignant potential of breast cancer cells and to alter the migratory phenotype of cells.

Cell migration is essential for development, tissue maintenance and repair, and tumorigenesis, and is regulated by growth factors, chemokines, and cytokines. This protocol describes the dot assay, a two-dimensional, unconstrained migration assay to assess the migratory phenotype of attached, cohesive cell sheets in response to microenvironmental cues.

Although complex organisms appear static, their tissues are under a continuous turnover. As cells age, die, and are replaced by new cells, cells move ...

Characterize Disease-related Mutants of RAF Family Kinases by Using a Set of Practical and Feasible Methods

The rapidly accelerated fibrosarcoma (RAF) family kinases play a central role in cell biology and their dysfunction leads to cancers and developmental disorders. A characterization of disease-related RAF mutants will help us select appropriate therapeutic strategies for treating these diseases. Recent studies have shown that RAF family kinases have both catalytic and allosteric activities, which are tightly regulated by dimerization. Here, we constructed a set of practical and feasible methods to determine the catalytic and allosteric activities and the relative dimer affinity/stability of RAF family kinases and their mutants. Firstly, we amended the classical in vitro kinase assay by reducing the detergent concentration in buffers, utilizing a gentle quick wash procedure, and employing a glutathione S-transferase (GST) fusion to prevent RAF dimers from dissociating during purification. This enables us to measure the catalytic activity of constitutively active RAF mutants appropriately. Secondly, we developed a novel RAF co-activation assay to evaluate the allosteric activity of kinase-dead RAF mutants by using N-terminal truncated RAF proteins, eliminating the requirement of active Ras in current protocols and thereby achieving a higher sensitivity. Lastly, we generated a unique complementary split luciferase assay to quantitatively measure the relative dimer affinity/stability of various RAF mutants, which is more reliable and sensitive compared to the traditional co-immunoprecipitation assay. In summary, these methods have the following advantages: (1) user-friendly; (2) able to carry out effectively without advanced equipment; (3) cost-effective; (4) highly sensitive and reproducible.

In this article, we presented a set of practical and feasible methods for characterizing disease-related mutants of RAF family kinases, which include in vitro kinase assay, RAF co-activation assay, and complementary split luciferase assay.

The rapidly accelerated fibrosarcoma (RAF) family kinases play a central role in cell biology and their dysfunction leads to cancers and developmental ...

Establishment of Gastric Cancer Patient-derived Xenograft Models and Primary Cell Lines

The use of preclinical models to advance our understanding of tumor biology and investigate the efficacy of therapeutic agents is key to cancer research. Although there are many established gastric cancer cell lines and many conventional transgenic mouse models for preclinical research, the disadvantages of these in vitro and in vivo models limit their applications. Because the characteristics of these models have changed in culture, they no longer model tumor heterogeneity, and their responses have not been able to predict responses in humans. Thus, alternative models that better represent tumor heterogeneity are being developed. Patient-derived xenograft (PDX) models preserve the histologic appearance of cancer cells, retain intratumoral heterogeneity, and better reflect the relevant human components of the tumor microenvironment. However, it usually takes 4-8 months to develop a PDX model, which is longer than the expected survival of many gastric patients. For this reason, establishing primary cancer cell lines may be an effective complementary method for drug response studies. The current protocol describes methods to establish PDX models and primary cancer cell lines from surgical gastric cancer samples. These methods provide a useful tool for drug development and cancer biology research.

The current protocol describes methods to establish patient-derived xenograft (PDX) models and primary cancer cell lines from surgical gastric cancer samples. The methods provide a useful tool for drug development and cancer biology research.

The use of preclinical models to advance our understanding of tumor biology and investigate the efficacy of therapeutic agents is key to cancer research. ...

Radiosensitivity of Cancer Stem Cells in Lung Cancer Cell Lines

The presence of cancer stem cells (CSCs) has been associated with relapse or poor outcomes after radiotherapy. Studying radioresistant CSCs may provide clues to overcoming radioresistance. Voltage-gated calcium channel &#945;2&#948;1 subunit isoform 5 has been reported as a marker for radioresistant CSCs in non-small cell lung cancer (NSCLC) cell lines. Using calcium channel &#945;2&#948;1 subunit as an example of a CSC marker, methods to study the radiosensitivity of CSCs in NSCLC cell lines are presented. CSCs are sorted with putative markers by flow cytometry, and the self-renewal capacity of sorted cells is evaluated by sphere formation assay. Colony formation assay, which determines how many cells lose the ability to generate descendants forming the colony after a certain dose of radiation, is then performed to assess the radiosensitivity of sorted cells. This manuscript provides initial steps for studying the radiosensitivity of CSCs, which establishes the basis for further understanding of the underlying mechanisms.

The presence of cancer stem cells have been associated with relapse or poor outcomes after radiotherapy. This manuscript describes the methods to study the radiosensitivity of cancer stem cells in lung cancer cell lines.

The presence of cancer stem cells (CSCs) has been associated with relapse or poor outcomes after radiotherapy. Studying radioresistant CSCs may provide ...

Establishment and Characterization of Small Bowel Neuroendocrine Tumor Spheroids

Small bowel neuroendocrine tumors (SBNETs) are rare cancers originating from enterochromaffin cells of the gut. Research in this field has been limited because very few patient derived SBNET cell lines have been generated. Well-differentiated SBNET cells are slow growing and are hard to propagate. The few cell lines that have been established are not readily available, and after time in culture may not continue to express characteristics of NET cells. Generating new cell lines could take many years since SBNET cells have a long doubling time and many enrichment steps are needed in order to eliminate the rapidly dividing cancer-associated fibroblasts. To overcome these limitations, we have developed a protocol to culture SBNET cells from surgically removed tumors as spheroids in extracellular matrix (ECM). The ECM forms a 3-dimensional matrix that encapsulates SBNET cells and mimics the tumor micro-environment for allowing SBNET cells to grow. Here, we characterized the growth rate of SBNET spheroids and described methods to identify SBNET markers using immunofluorescence microscopy and immunohistochemistry to confirm that the spheroids are neuroendocrine tumor cells. In addition, we used SBNET spheroids for testing the cytotoxicity of rapamycin.

Neuroendocrine tumors (NETs) originate from neuroendocrine cells of the neural crest. They are slow growing and challenging to culture. We present an alternative strategy to grow NETs from the small bowel by culturing them as spheroids. These spheroids have small bowel NET markers and can be used for drug testing.

Small bowel neuroendocrine tumors (SBNETs) are rare cancers originating from enterochromaffin cells of the gut. Research in this field has been limited ...

Analysis of Side Population in Solid Tumor Cell Lines

Cancer stem cells (CSCs) are an important cause of tumor growth, metastasis, and recurrence. Isolation and identification of CSCs are of great significance for tumor research. Currently, several techniques are used for the identification and purification of CSCs from tumor tissues and tumor cell lines. Separation and analysis of side population (SP) cells are two of the commonly used methods. The methods rely on the ability of CSCs to rapidly expel fluorescent dyes, such as Hoechst 33342. The efflux of the dye is associated with the ATP-binding cassette (ABC) transporters and can be inhibited by ABC transporter inhibitors. Methods for staining cultured tumor cells with Hoechst 33342 and analyzing the proportion of their SP cells by flow cytometry are described. This assay is convenient, fast, and cost-effective. Data generated in this assay can contribute to a better understanding of the effect of genes or other extracellular and intracellular signals on the stemness properties of tumor cells.

A convenient, fast, and cost-effective method to measure the proportion of side population cells in solid tumor cell lines is presented.

Cancer stem cells (CSCs) are an important cause of tumor growth, metastasis, and recurrence. Isolation and identification of CSCs are of great ...

Functional Assessment of BRCA1 variants using CRISPR-Mediated Base Editors

Recent studies have investigated the risks associated with BRCA1 gene mutations using various functional assessment methods such as fluorescent reporter assays, embryonic stem cell viability assays, and therapeutic drug-based sensitivity assays. Although they have clarified a lot of BRCA1 variants, these assays involving the use of exogenously expressed BRCA1 variants are associated with overexpression issues and cannot be applied to post-transcriptional regulation. To resolve these limitations, we previously reported a method for functional analysis of BRCA1 variants via CRISPR-mediated cytosine base editor that induce targeted nucleotide substitution in living cells. Using this method, we identified variants whose functions remain ambiguous, including c.-97C&#62;T, c.154C&#62;T, c.3847C&#62;T, c.5056C&#62;T, and c.4986+5G&#62;A, and confirmed that CRISPR-mediated base editors are useful tools for reclassifying the variants of uncertain significance in BRCA1. Here, we describe a protocol for functional analysis of BRCA1 variants using CRISPR-based cytosine base editor. This protocol provides guidelines for the selection of target sites, functional analysis and evaluation of BRCA1 variants.

Functional Assessment of BRCA1 variants using CRISPR-Mediated Base Editors

People with BRCA1 mutations have a higher risk of developing cancer, which warrants accurate evaluation of the function of BRCA1 variants. Herein, we described a protocol for functional assessment of BRCA1 variants using CRISPR-mediated cytosine base editors that enable targeted C:G to T:A conversion in living cells.

Recent studies have investigated the risks associated with BRCA1 gene mutations using various functional assessment methods such as fluorescent reporter ...

High-Throughput In Vitro Assay using Patient-Derived Tumor Organoids

Patient-derived tumor organoids (PDOs) are expected to be a preclinical cancer model with better reproducibility of disease than traditional cell culture models. PDOs have been successfully generated from a variety of human tumors to recapitulate the architecture and function of tumor tissue accurately and efficiently. However, PDOs are unsuitable for an in vitro high-throughput assay system (HTS) or cell analysis using 96-well or 384-well plates when evaluating anticancer drugs because they are heterogeneous in size and form large clusters in culture. These cultures and assays use extracellular matrices, such as Matrigel, to create tumor tissue scaffolds. Therefore, PDOs have a low throughput and high cost, and it has been difficult to develop a suitable assay system. To address this issue, a simpler and more accurate HTS was established using PDOs to evaluate the potency of anticancer drugs and immunotherapy. An in vitro HTS was created that uses PDOs established from solid tumors cultured in 384-well plates. An HTS was also developed for assessment of antibody-dependent cellular cytotoxicity activity to represent the immune response using PDOs cultured in 96-well plates.

High-Throughput In Vitro Assay using Patient-Derived Tumor Organoids

A highly accurate in vitro high-throughput assay system was developed to evaluate anticancer drugs using patient-derived tumor organoids (PDOs), similar to cancer tissues but are unsuitable for in vitro high-throughput assay systems with 96-well and 384-well plates.

Patient-derived tumor organoids (PDOs) are expected to be a preclinical cancer model with better reproducibility of disease than traditional cell culture ...

Profiling Sensitivity to Targeted Therapies in EGFR-Mutant NSCLC Patient-Derived Organoids

Novel 3D cancer organoid cultures derived from clinical patient specimens represent an important model system to evaluate intratumor heterogeneity and treatment response to targeted inhibitors in cancer. Pioneering work in gastrointestinal and pancreatic cancers has highlighted the promise of patient-derived organoids (PDOs) as a patient-proximate culture system, with an increasing number of models emerging. Similarly, work in other cancer types has focused on establishing organoid models and optimizing culture protocols. Notably, 3D cancer organoid models maintain the genetic complexity of original tumor specimens and thus translate tumor-derived sequencing data into treatment with genetically informed targeted therapies in an experimental setting. Further, PDOs might foster the evaluation of rational combination treatments to overcome resistance-associated adaptation of tumors in the future. The latter focuses on intense research efforts in non-small-cell lung cancer (NSCLC), as resistance development ultimately limits the treatment success of targeted inhibitors. An early assessment of therapeutically targetable mechanisms using NSCLC PDOs could help inform rational combination treatments. This manuscript describes a standardized protocol for the cell culture plate-based assessment of drug sensitivities to targeted inhibitors in NSCLC-derived 3D PDOs, with potential adaptability to combinational treatments and other treatment modalities.

This protocol describes a standardized evaluation of drug sensitivities to targeted signaling inhibitors in NSCLC patient-derived organoid models.

Novel 3D cancer organoid cultures derived from clinical patient specimens represent an important model system to evaluate intratumor heterogeneity and ...

Exploring Mitochondrial Energy Metabolism of Single 3D Microtissue Spheroids Using Extracellular Flux Analysis

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing cells as two-dimensional, single-cell monolayers (2D culture), spheroid cell culture promotes, regulates, and supports physiological cellular architecture and characteristics that exist in vivo, including the expression of extracellular matrix proteins, cell signaling, gene expression, protein production, differentiation, and proliferation. The importance of 3D culture has been recognized in many research fields, including oncology, diabetes, stem cell biology, and tissue engineering. Over the last decade, improved methods have been developed to produce spheroids and assess their metabolic function and fate.
Extracellular flux (XF) analyzers have been used to explore mitochondrial function in 3D microtissues such as spheroids using either an XF24 islet capture plate or an XFe96 spheroid microplate. However, distinct protocols and the optimization of probing mitochondrial energy metabolism in spheroids using XF technology have not been described in detail. This paper provides detailed protocols for probing mitochondrial energy metabolism in single 3D spheroids using spheroid microplates with the XFe96 XF analyzer. Using different cancer cell lines, XF technology is demonstrated to be capable of distinguishing between cellular respiration in 3D spheroids of not only different sizes but also different volumes, cell numbers,&#160;DNA content and type.
The optimal mitochondrial effector compound concentrations of oligomycin, BAM15, rotenone, and antimycin A are used to probe specific parameters of mitochondrial energy metabolism in 3D spheroids. This paper also discusses methods to normalize data obtained from spheroids and addresses many considerations that should be considered when exploring spheroid metabolism using XF technology. This protocol will help drive research in advanced in vitro spheroid models.

These protocols will help users probe mitochondrial energy metabolism in 3D cancer cell-line-derived spheroids using Seahorse extracellular flux analysis.

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing ...