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. 2026 Mar 20;21(4):852–862. doi: 10.1021/acschembio.6c00094

A Modular Platform for Quantitative Two-Color Bioluminescence Combining NanoLuc and Red-Shifted NanoPrism Luciferases

Karilyn Porter †, Mike Killoran †, Robin Hurst †, Mark Klein †, Debayan De Bakshi †, Rahele Esmatpour †, Lance P Encell †, Felipe Gama-Franceschi ‡, Joseph Bessetti †, Kristin M Riching †, Thomas Machleidt †, Laura Mezzanotte ‡, Rachel Friedman Ohana †,*
PMCID: PMC13097136  PMID: 41858017

Abstract

Bioluminescent reporters are widely used to monitor and image biological processes. Among these, NanoLuc luciferase and its complementation variants (LgBiT/SmBiT and LgBiT/HiBiT) are commonly used due to their brightness, sensitivity, and compatibility with prolonged kinetic measurements. However, the single-channel emission of these NanoLuc-based systems (460 nm peak) limits their use in multiplexed assays. Prior efforts to shift NanoLuc’s emission employed bioluminescence resonance energy transfer (BRET) to a proximal fluorescent protein or organic fluorophore. Building on this concept, we engineered high-efficiency BRET reporters, termed NanoPrism luciferases, by inserting circularly permuted NanoLuc or LgBiT into a surface loop of the self-labeling HaloTag protein. These NanoPrisms achieve a ∼90% BRET efficiency by optimally positioning NanoLuc variants near a fluorophore covalently bound to HaloTag. The binary design further supports high- and low-affinity complementation, allowing applications in HiBiT knock-in cells and tracking protein–protein interactions, respectively. Pairing red-shifted NanoPrisms with unmodified NanoLuc or its complementation variants, we created a two-color bioluminescent reporter platform featuring bright signals of similar intensity and >100 nm spectral separation, allowing quantitative, simultaneous measurement of two molecular readouts within the same sample. Here, we demonstrate the platform’s utility for monitoring a degradation target alongside a control protein and for tracking two distinct events within a biological pathway, using plate-based detection and bioluminescence imaging. By enabling concurrent measurements within the same sample, the system provides insights into cellular dynamics while reducing variability and complexity associated with parallel single-channel assays.

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Introduction

Bioluminescence detection is widely used in functional biology analyses in vitro and small animal models due to its high sensitivity and low background. , Unlike fluorescent reporters, luciferases emit light through the enzymatic oxidation of luciferin substrates, eliminating the need for external excitation. This avoids autofluorescence and minimizes phototoxicity and photobleaching. ,, As a result, bioluminescent systems offer high signal-to-noise ratios, broad dynamic range, and ability to monitor dynamic biological processes over extended periods. −

However, compared to fluorescent reporters, the emission spectra of luciferases are constrained by luciferase–luciferin chemistry, limiting their use in multiplexed assays. Additionally, bioluminescent measurements can fluctuate due to differences in reporter expression, substrate depletion, or changes in cell viability and experimental conditions. ,, To correct for such variability or to monitor two biological events, parallel single-reporter assays are often performed in separate wells, increasing variability, cost, and workflow complexity. Measuring two reporters within the same well would address these limitations.

In practice, signal deconvolution can be challenging when multiplexing existing luciferase reporters because of vast differences in their luminescence output, overlapping emission profiles, or a combination of both. A common multiplexing strategy employs orthogonal substrates. While this approach can accommodate differences in brightness, it often requires sequential measurements in which one reaction must be terminated before adding a second substrate. , Alternatively, multiplexing can rely on spectrally distinct luciferases combined with unmixing approaches, though this strategy is often limited by insufficient spectral separation and the need for complex deconvolution algorithms. ,,

An ideal two-color bioluminescence system would therefore combine bright reporters with well-separated emission spectra. , Shared, single substrate designs further simplify experimental workflows by eliminating sequential substrate addition and enabling real-time correction for signal decay caused by substrate instability or consumption. Despite this potential, most spectrally resolved bioluminescent systems have been applied primarily to multicolor animal imaging, , and their use in quantitative plate-based assays remains limited to niche applications. ,, This likely reflects the limited availability of sufficiently bright, spectrally distinct luciferases and the lack of simple signal deconvolution tools.

One strategy to extend the emission spectra of luciferases relies on bioluminescence resonance energy transfer (BRET). ,,, In BRET, energy generated by a luciferase-catalyzed reaction is transferred nonradiatively to a nearby fluorophore, which then emits light at a longer wavelength. BRET efficiency depends on proximity (≤10 nm), orientation, and spectral overlap between donor emission and acceptor excitation, making covalently linked configurations especially effective.

NanoLuc luciferase has emerged as an attractive donor for intramolecular BRET systems due to its small size, high stability, and bright emission (peak at 460 nm). , Several prior designs fused NanoLuc to one or two fluorescent proteins to extend its emission spectrum (e.g., LumiFluors, enhanced NanoLanterns, and Antares). ,, An alternative design described by Hiblot et al. incorporated a circularly permuted version of NanoLuc into self-labeling proteins (i.e., HaloTag or SNAP-tag), allowing covalent attachment of interchangeable organic fluorophores for tunable emission.

Building on these prior designs, we engineered high-efficiency intramolecular BRET reporters, termed NanoPrism luciferases, which utilize NanoLuc or its complementation variants (LgBiT/SmBiT or LgBiT/HiBiT) as the donor. NanoPrisms incorporate a circularly permuted NanoLuc or LgBiT into a surface loop of HaloTag, achieving a ∼90% BRET efficiency by positioning the donor in close proximity and a favorable, constrained orientation relative to a covalently bound fluorophore. Additionally, the versatile binary design supports high- and low-affinity complementation, enabling applications in HiBiT knock-in cells under endogenous expression levels and tracking reversible protein–protein interactions, respectively.

By pairing red-shifted NanoPrisms with unmodified NanoLuc or its complementation variants, we created a two-color bioluminescent reporter platform featuring bright signals with similar intensities and >100 nm spectral separation, enabling quantitative, simultaneous measurement of two molecular readouts within the same well. Here, we demonstrate its utility for monitoring two biological events, including BRD4 degradation alongside a control protein and distinct steps within the EGF-induced epidermal growth factor receptor (EGFR) signaling pathway, using both plate-based detection and bioluminescence imaging. Overall, this two-color approach enables concurrent measurement of two biological events, supporting internal normalization and real-time analysis of pathway dynamics in a single assay format.

Results and Discussion

Engineering Intramolecular BRET-Based Reporters

Envisioning a two-color bioluminescent reporter system for quantitative cell-based assays and in cellulo imaging, we sought to pair NanoLuc (NLuc) known for its bright blue emission with red-shifted intramolecular BRET reporters that utilize the same substrate. We opted to build on the intramolecular BRET design introduced by Hiblot et al., in which a circularly permuted NanoLuc (cpNLuc 67/68), generated by linking the native N- and C-termini and creating new termini at an internal site, was inserted into HaloTag. In our hands, this design achieved ∼64% energy transfer to a TMR ligand covalently bound to HaloTag, compared to ∼20% energy transfer for tandem NLuc-HaloTag fusions, while maintaining comparable overall brightness (Figure S1).

To explore whether BRET efficiency can be further improved, we used the HaloTag 3D structure (PDB # 6U32) to identify alternative insertion sites within the lid domain that are proximal to the bound fluorophore (Figure S1). Three candidate surface loops (residues 164–167, 178–179, and 194–195) were evaluated by inserting cpNLuc 67/68 into each position. Among these constructs, insertion between residues 178–179 was 2.5-fold dimmer than a tandem HaloTag-NLuc fusion but achieved ∼80% BRET efficiency to a covalently bound TMR HaloTag ligand (Figure S1B–E). This configuration was therefore selected for further optimization.

Next, we examined whether alternative circular permutation sites in NanoLuc could further enhance the BRET efficiency (Figure S2). Circularly permuted variants (n = 21) were inserted at the optimized 178–179 position within HaloTag, expressed in E. coli, and evaluated as lysates. Several permutation sites beyond cpNLuc 67/68, specifically cpNLuc 27/28, 64/65, and 103/104, exhibited high apparent BRET efficiencies despite substantially reduced donor and acceptor emission intensities, reflecting the ratiometric nature of BRET. In contrast, cpNLuc 67/68 combined high apparent BRET efficiency with bright acceptor emission and was therefore retained as the optimal donor permutation for this intramolecular BRET configuration.

Flexible linkers connecting domains of the intramolecular BRET reporter can influence protein folding, structural stability, spatial configuration, and ultimately BRET efficiency (Figure S3A). Our lead construct contained a single 12-residue flexible linker (GGTGGSGGTGGS; L 2) connecting the native N and C termini of NanoLuc. To investigate the impact of linker length, L 2 was replaced with a panel of Ser–Gly linkers ranging from 3 to 15 residues. Evaluation of these variants in E. coli lysates revealed that constructs containing 9–15 residue linkers retained high BRET efficiency and bright acceptor emission, whereas those with shorter linkers were up to 5-fold dimmer (Figure S3). Given these results, we retained the 12-residue L 2 linker and explored the effect of additional linkers flanking cpNLuc 67/68 and connecting it to the N and C terminal domains of HaloTag (L 1 and L 3, respectively). Incorporation of a 3-residue L 1 linker modestly enhanced BRET, whereas longer L 1 linkers progressively decreased BRET in a length-dependent manner. Insertion of any L 3 linker caused a marked decrease in BRET (Figure S3D). Biochemical characterization of purified proteins revealed that incorporation of the 3-residue L 1 linker did not significantly affect total bioluminescence or BRET but increased HaloTag ligand binding kinetics by ∼1.4-fold (k = 1.9 × 106 M–1 s–1), possibly reflecting improved protein folding (Figure S4). Addition of two HaloTag mutations (V197E and P206V), previously shown to enhance the stability of certain HaloTag variants (data not shown), further increased ligand binding kinetics by 1.8-fold (k = 3.4 × 106 M–1 s–1). The structure of the resulting optimized construct, termed PrismaLuc, is shown in Figure A, and the sequence is provided in Supporting Information Materials and Methods.

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Designs and biochemical properties of PrismaLuc and PrismaLgBiT. (A) Schematic illustration of PrismaLuc and PrismaLgBiT: HaloTag (red), flexible linkers (gray), cpNLuc or cpLgBiT (blue), and mutation sites (darker shades). The last β-strand of NanoLuc is shown in yellow. Purified NanoPrisms were compared to their corresponding tandem fusions for (B) total luminescence (error bars represent the SD of triplicates), (C) bioluminescence emission spectra to a bound JF549 HaloTag ligand, normalized to the donor peak at 460 nm, and (D) apparent BRET efficiencies calculated as the percent area under the curve AUC(%): [acceptor/(acceptor + donor) × 100]. (E) Boltz-1X-predicted structure of PrismaLuc with bound fluorophore.

To broaden utility, we explored whether these design principles could be applied to binary systems capable of both high- and low-affinity complementation, supporting applications in HiBiT knock-in cells and the detection of protein–protein interactions, respectively. To this end, we generated a construct comprising a circularly permuted version of LgBiT (cpLgBiT 67/68) inserted into HaloTag between residues 178 and 179 and incorporating the optimized 3-residue L 1 linker (Figure S5). Upon complementation with a synthetic VS-HiBiT peptide, this construct retained a high BRET efficiency but exhibited a ∼60-fold lower affinity for HiBiT and a ∼66-fold slower ligand binding kinetics, suggesting potential structural instability. To address these impairments, we introduced mutations previously shown to enhance the stability of HaloTag (V197E and P206V) and LgBiT (E4D, Q42M, M106K, R112H, K123E, V127T, and T144D) variants. These mutations significantly improved the construct’s affinity for HiBiT (K D = 14.9 nM) and accelerated HaloTag ligand binding (k = 1.9 × 106 M–1 s–1). Although these values remain ∼5-fold weaker and ∼10-fold slower than those of unmodified LgBiT and HaloTag, respectively, they represent a marked improvement over the unoptimized construct, suggesting enhanced folding and structural stability. The resulting design, termed PrismaLgBiT, is illustrated in Figure A, and the sequence is provided in the Supporting Information Materials and Methods.

To assess the biochemical properties of the two NanoPrisms, we first compared them to tandem NanoLuc- and LgBiT-HaloTag fusions (Figures and S6A–C). PrismaLuc was only ∼3-fold dimmer than NLuc-HaloTag and exhibited ∼5-fold slower binding kinetics of a TMR HaloTag ligand. Upon complementation, PrismaLgBiT was ∼4-fold dimmer than LgBiT-HaloTag and displayed ∼10-fold slower ligand binding kinetics in both complemented and noncomplemented states. These results suggest that NanoPrisms exhibit structural stability, proper folding, and rapid, quantitative labeling with a HaloTag ligand.

Evaluating intramolecular BRET efficiencies and compatibility with multiple energy acceptors, we first compared two HaloTag ligands with identical emission maxima, TMR and Janelia Fluor 549 (JF549). The latter was reported to be a higher quantum yield acceptor in NanoLuc-based BRET systems. NanoPrisms labeled with JF549 displayed brighter acceptor emissions and correspondingly higher BRET efficiencies than those labeled with TMR (Figure S6D–F). Using JF549, PrismaLuc and PrismaLgBiT achieved ∼89% and ∼87% BRET efficiencies, respectively, compared to ∼26% and ∼31% for the corresponding tandem fusions (Figure C,D). Notably, the binary NanoPrism exhibited a BRET efficiency similar to that of the full-length construct, indicating that high energy transfer can be maintained in a complementation-based format. Expanding the evaluation to ten additional Janelia Fluor HaloTag ligands (emission maxima ranging from 535 to 690 nm) confirmed broad fluorophore compatibility and consistently higher BRET efficiencies for NanoPrisms relative to tandem fusions (Figure S7). As expected, the efficiencies of these energy transfers mostly correlated with the extent of donor–acceptor spectral overlaps. ,

Finally, we compared the intramolecular BRET efficiencies of NanoPrisms paired with JF549 (87–89%, depending on the construct) to two representative red-shifted NanoLuc-based reporters under identical experimental conditions (Figure S8). The HaloTag–cpNLuc configuration described by Hiblot et al., which inspired our design, achieved ∼76% energy transfer when paired with JF549. In comparison, Antares, a genetically encoded reporter in which NanoLuc is fused to two CyOFP1 fluorescent proteins (emission maximum ∼589 nm, comparable to that of JF549), exhibited a lower BRET efficiency of ∼68%. Under these conditions, NanoPrisms exhibited a higher intramolecular BRET efficiency and a more complete spectral shift, which is advantageous for applications requiring effective signal separation.

Given that BRET efficiency depends on donor–acceptor proximity and spatial orientation, we used Boltz-1X, an open-source AI model, to predict the 3D structure of PrismaLuc. The model predominantly predicted a “horizontal” orientation of cpNLuc relative to HaloTag’s lid domain, aligning the two in parallel configuration with the bound fluorophore positioned at the interface between them (Figure D). A per-residue confidence analysis suggested that the HaloTag domains were predicted with high confidence, whereas the cpNLuc domains, particularly at the junctions connecting them to HaloTag, were predicted with lower confidence. This may reflect regions with conformational flexibility or model uncertainty (Figure S9). To explore a possible relationship between BRET efficiency and the predicted cpNLuc orientation, we modeled two significantly lower BRET configurations in which cpNLuc was inserted into HaloTag at alternative sites (residues 164–167 and 194–195; Figure S1). Interestingly, for these two variants, the model predicted a more “vertical” orientation of cpNLuc, resulting in reduced parallel alignment with the HaloTag lid domain (Figure S10). Collectively, these structural predictions suggest a correlation between BRET efficiency and the predicted degree of parallel alignment between cpNLuc and the HaloTag lid domain, with the “horizontal” orientation in PrismaLuc appearing to be favorable for efficient BRET.

Developing a Two-Color Bioluminescent Reporter System

Following biochemical characterization, we evaluated the performance of NanoPrisms in mammalian cells using two experimental configurations. In one, HeLa cells were transfected with mRNA encoding PrismaLuc (Figure S11). In the other, genome-edited HeLa cells expressing HiBiT-tagged glyceraldehyde-3-phosphate dehydrogenase (GAPDH-HiBiT) were transfected with mRNA encoding PrismaLgBiT, allowing for HiBiT/PrismaLgBiT complementation (Figure ). Transfected cells were incubated overnight with 100 nM Janelia Fluor HaloTag ligands with emission maxima ranging from 535 to 690 nm. Covalent binding ensured full occupancy of the expressed NanoPrisms, and the specificity of BRET eliminated the need to washout unbound ligand. Upon substrate addition, BRET-driven spectral shifts were evaluated by bioluminescence emission scans and filtered bioluminescence imaging. Emission scans revealed characteristic energy-transfer profiles with decreased donor emission and increased acceptor emission (Figure A), closely matching those obtained in biochemical analyses. Filtered luminescence images were acquired using a GloMax Galaxy imager equipped with a 460/50 nm bandpass (BP) filter for donor detection and long-pass (LP) filters (500, 575, or 600 nm) for acceptor detection (Figure B). Images captured at identical exposure times and displayed on a uniform intensity scale demonstrated BRET-driven spectral shifts with the extent of donor emission loss and acceptor emission gain correlating with the degree of donor–acceptor spectral overlap. These analyses demonstrated that the NanoPrisms support highly efficient BRET-driven spectral shifts in live cells across a range of HaloTag ligands. Ligand selection, however, is application-dependent. For two-color analyses pairing NanoLuc and NanoPrism, optimal ligands should combine favorable cellular permeability, high quantum yield, efficient BRET-mediated spectral shifting, and marked spectral separation from NanoLuc. Considering these criteria, the JF549 HaloTag ligand was selected for subsequent experiments. JF549 provides a favorable spectral overlap with NanoLuc for efficient energy transfer and high quantum yield, resulting in a signal that is > 100 nm red-shifted from NanoLuc’s 460 nm peak while minimizing residual donor emission (Figure A,B). Despite this substantial spectral separation between NanoLuc (or its complementation variants) and NanoPrisms paired with JF549, a small degree of overlap between detection channels remains unavoidable.

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BRET-driven spectral shifts for PrismaLgBiT paired with Janelia Fluor HaloTag ligands. HeLa cells coexpressing PrismaLgBiT and GAPDH-HiBiT were incubated with 100 nM Janelia Fluor HaloTag ligands and treated with Nano-Glo luciferase substrate. (A) Bioluminescence emission spectra acquired on a Tecan Infinite M1000 and normalized to the 460 nm donor peak emission. (B) Filtered luminescence images acquired using a GloMax Galaxy imager equipped with the indicated filters. All images were captured with 2 min exposures and displayed with a common intensity scale. Grayscale images are provided in Figure S12.

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Two-color bioluminescent reporter system: spectral profiles and correction factors for signal deconvolution. (A) Bioluminescence emission spectra for HeLa cells expressing NanoLuc or PrismaLuc. (B) Corresponding emission spectra for genome-edited HeLa cells expressing GAPDH-HiBiT and either LgBiT or PrismaLgBiT. Cells were incubated overnight with 100 nM JF549 HaloTag ligand, and emission scans were acquired following addition of Nano-Glo luciferase substrate using a Tecan Infinite M1000 plate-reader. (C) Correction factors determined for plate-reader and imaging-based detection. Corrections for spectral bleed-through were determined using HEK293 cells expressing NLuc-BRD4 and calculated as the fraction of the red-channel signal relative to the blue-channel signal (red/blue). Corrections for the residual NanoPrism donor emission were determined using HeLa cells expressing PrismaLuc or PrismaLgBiT together with GAPDH-HiBiT and calculated as the fraction of the blue-channel signal relative to the corrected red signal (blue/corrected red). Correction factors represent mean ± SD (n = 3), and differences reflect instrument-specific filter sets and detector sensitivity, which influence the relative amount of light captured in each channel (see Figure S13). These correction factors were used to resolve NanoLuc and NanoPrism signals in two-color analyses as described in the Materials and Methods.

To accurately resolve the two signals within the same sample, we implemented a correction strategy that accounts for two factors: spectral bleed-through from the blue channel into the red channel and residual donor emission from the NanoPrism reporter. Although NanoPrisms exhibit high BRET efficiency, a fraction of donor emission remains detectable in the blue channel and must be corrected (Figure C). Because these corrections are context-dependent, they must be determined empirically for each NanoPrism/fluorophore pairing and instrument/filter configuration.

Spectral bleed-through was quantified using genome-edited HEK293 cells expressing NLuc-BRD4, plated in a 96-well plate or an 8-chamber slide and incubated overnight with or without a 100 nM JF549 HaloTag ligand. Following substrate addition, filtered luminescence was measured using a GloMax Discover plate-reader or imaged using a GloMax Galaxy imager (blue channel: 450/8 nm or 460/50 nm filters and red channel: 600 or 575 nm long-pass filters, respectively). Spectral bleed-through calculated as the fraction of the red-channel signal relative to the blue-channel signal was unaffected by the presence of an unbound JF549 HaloTag ligand. These correction factors for bleed-through (0.01 ± 0.0003 for the plate-reader and 0.025 ± 0.0015 for the imager) were highly reproducible for a given instrument and filter configuration and were used to correct red-channel signals in subsequent analyses.

Residual NanoPrism donor emission was quantified using HeLa cells expressing either PrismaLuc or coexpressing GAPDH-HiBiT and PrismaLgBiT, which were incubated overnight with a 100 nM JF549 HaloTag ligand. Because BRET efficiency is concentration-independent and constant for a given donor–acceptor configuration, the residual donor emission can be expressed as a fixed fraction of the corrected red-channel signal. Residual donor contributions were 0.36 ± 0.02 (PrismaLuc) and 0.52 ± 0.02 (PrismaLgBiT) for the plate-reader and 0.1 ± 0.006 and 0.14 ± 0.009 for the imager, respectively. These differences primarily reflect instrument-specific filter sets and detector sensitivity, which influence the relative amount of light captured in each channel (Figure S13). While the lower residual donor fractions measured with the imager more closely reflect the high BRET efficiency (∼90%), the absolute values are less important than their consistency. For both systems, these correction factors were highly reproducible for a given NanoPrism/fluorophore configuration and instrument setup.

Together, the corrections for spectral bleed-through and residual donor emission form the basis of our two-color signal deconvolution strategy. By correcting the red channel for spectral bleed-through and subtracting the NanoPrism donor contribution from the blue channel, this approach enables accurate resolution of the NanoPrism- and NanoLuc-derived signals within the same sample.

Monitoring Degradation of BRD4 Alongside Internal Control

Having established a two-color bioluminescent reporter system and a strategy for resolving the two signals, we next evaluated its utility for quantitative analysis of two proteins within the same well. Quantitative analyses were performed using both plate-based measurements, which report population-averaged luminescence per well, and bioluminescence imaging, which enables visualization of the two reporters in individual cells. First, we applied this approach to targeted protein degradation assays, simultaneously monitoring a degradation target and a nondegrading control within the same well. Inclusion of a nondegrading control allows correction for signal decay unrelated to degradation such as substrate depletion, reporter inhibition, or reduced cell viability, thereby improving degradation assessment.

We chose a well-characterized model involving dBET6, a thalidomide-based PROTAC (Proteolysis Targeting Chimera) that induces ubiquitin-mediated degradation of BRD4, a member of the BET bromodomain family. In this two-color configuration, endogenously tagged NLuc-BRD4 served as the degradation target while the red-shifted NanoPrism reporter monitored a nondegrading control protein. GAPDH was selected as a generic control, whereas GSPT1, a known Cereblon-dependent neosubstrate, was included as a potential off-target control, given that thalidomide-based PROTACs can induce degradation of certain neosubstrates. Accordingly, we first assessed whether dBET6 induces degradation of GSPT1 and confirmed that it does not, supporting its use as a control in this context (Figure S14).

The two reporters were initially expressed in separate cell populations, one expressing NLuc-BRD4 and the other either expressing GAPDH-HiBiT or HiBiT-GSPT1. The populations were coplated at a 1:1 ratio in a 96-well plate or an 8-chamber slide. The following day, cells were transfected with mRNA encoding PrismaLgBiT and incubated overnight with 100 nM JF549 HaloTag ligand. To support extended kinetic measurements, cells were preincubated for 1 h with Vivazine (a slow-release NanoLuc substrate) prior to treatment with 100 nM dBET6. Filtered luminescence readings and images were collected continuously over a 3 h period using the GloMax Discover plate-reader and GloMax Galaxy imager, respectively (Figure A, Supporting Information Videos V1 and V2). Images were processed using Fiji software. For each channel, threshold-guided segmentation was applied to identify cells and extract the total signal intensities over time. Red-channel intensities were corrected for spectral bleed-through, and residual NanoPrism donor contributions were subtracted from blue-channel intensities using the correction factors defined in Figure (Figures B and S15A,B). Filtered luminescence readouts from the plate-reader were processed in the same manner (Figures C and S15C,D).

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BRD4 degradation tracked alongside an internal control in cocultured cell populations. HEK293 cells expressing NLuc-BRD4 were cocultured with cells expressing nondegrading control (GAPDH-HiBiT or HiBiT-GSPT1 complemented with PrismaLgBiT bound to JF549). Following preincubation with Vivazine, cells were treated with 100 nM dBET6. (A) Representative merged 2-channel images acquired over 3 h using a GloMax Galaxy imager (3 min exposures). Entire time courses are provided in Supporting Information Videos V1 and V2. Deconvolved signals for (B) imager and (C) plate-reader (provided in Figure S15), normalized to their initial values (T 0 = 100%), then baseline-corrected to the NanoPrism control. Error bars (plater-reader) represent the SD of 6 replicates.

Despite differences in control expression levels (GAPDH vs GSPT1), normalized analyses from both plate-reader and imager yielded consistent dBET6-induced BRD4 degradation profiles, with maximal degradation observed within 60 min, in agreement with prior reports. The close agreement between population-averaged plate-based measurements and imaging-based analyses demonstrates the robustness of the two-color system and the associated signal deconvolution strategy, supporting its use for quantitative analyses in standard plate-based formats. Notably, in the absence of HiBiT complementation, the presence of PrismaLgBiT/JF549 in the NLuc-BRD4 cells generated no measurable signal in either channel, suggesting minimal crosstalk between reporters in coplated cell populations.

Motivated by these results, we evaluated the system’s ability to resolve two reporters expressed in the same cell. To this end, we generated a double knock-in HEK293 cell line expressing nuclear NLuc-BRD4 and cytosolic HiBiT-GSPT1. Cells were transfected with mRNA encoding PrismaLgBiT and incubated overnight with 100 nM JF549 HaloTag ligand. Although variable transfection efficiency likely limited dual-reporter expression in all cells, both imaging and plate-based detection revealed degradation kinetics similar to those observed in coplated populations with maximal BRD4 degradation occurring within 60 min (Figures ,S16 and Supporting Information Video V3). In this configuration, localization of the two reporters to distinct subcellular compartments minimized unintended signal interference and enabled robust signal deconvolution. Extension of this approach to two reporters localized to the same cellular compartment may require optimization of expression levels to limit nonspecific energy transfer.

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BRD4 degradation tracked alongside an internal control coexpressed in the same cells. HEK293 cells coexpressing NLuc-BRD4 and HiBiT-GSPT1, transfected with PrismaLgBiT’s mRNA and incubated with JF549 HaloTag ligand. Following preincubation with Vivazine, cells were treated with 100 nM dBET6. (A) Representative merged 2-channel images acquired over 3 h using a GloMax Galaxy imager (3 min exposures). The entire time course is provided in Supporting Information Video V3. Deconvolved signals for (B) imager and (C) plate-reader (provided in Figure S16), normalized to their initial values (T 0 = 100%), then baseline-corrected to the NanoPrism control. Error bars (plater-reader) represent the SD of 6 replicates.

Monitoring EGFR Signaling, Internalization and Degradation

We further assessed the ability of our two-color system to resolve distinct but mechanistically linked cellular events within the same sample using epidermal growth factor receptor (EGFR) signaling and trafficking as a model. EGFR undergoes EGF-induced activation and recruitment of adaptor proteins such as Grb2, followed by rapid endocytosis, and subsequent ubiquitin-dependent lysosomal degradation.

To co-monitor EGFR–Grb2 interaction and receptor internalization, we generated genome-edited HeLa cells expressing HiBiT–EGFR and transfected them with mRNAs encoding EGFR–PrismaLgBiT and Grb2–SmBiT. In this configuration, EGF stimulation induces EGFR activation and Grb2 recruitment, driving a facilitated SmBiT/PrismaLgBiT complementation and reconstitution of a red-shifted NanoPrism reporter. In parallel, cell-surface complementation of HiBiT–EGFR with purified LgBiT enables tracking of EGFR internalization, as the pH-sensitive HiBiT/LgBiT luminescence decreases when the receptor translocates from the neutral plasma membrane to acidic endosomes. Cells were plated in either 96-well plates or 8-chamber slides, transfected, and incubated overnight with 100 nM JF549 HaloTag ligand. After serum starvation, purified LgBiT and Nano-Glo luciferase substrate were added to initiate surface complementation, followed by treatment with 300 ng/mL EGF or vehicle control. Filtered luminescence readings and images were collected continuously for 30 min (Figure ). Deconvolved plate-based measurements (Figure B), together with complementary time-lapse imaging, revealed rapid EGF-induced EGFR–Grb2 interaction, followed closely by receptor internalization, consistent with reported dynamics of EGFR activation and trafficking.

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Monitoring EGF-induced EGFR signaling and internalization. HeLa cells expressing HiBiT-EGFR were transfected with mRNA for EGFR-PrismaLgBiT and Grb2-SmBiT at a 1:2 molar ratio and incubated with JF549 HaloTag ligand. After 4 h of starvation, Nano-Glo luciferase substrate and purified LgBiT were added followed by treatment with 300 ng/mL EGF or vehicle control. (A) Representative merged 2-channel images acquired over 30 min (3 min exposures). Entire time course provided in Figure S17. (B) Deconvolved signals for plate-reader. Error bars represent the SD of 3 replicates.

We then assessed whether the system could resolve sequential trafficking events within the same samples. For this purpose, we generated an additional genome-edited HeLa cell line expressing EGFR–HiBiT, which upon complementation with PrismaLgBiT forms a red-shifted reporter that remains exposed to the neutral cytosolic environment, thereby the signal decrease signifies EGFR degradation. HeLa cells expressing HiBiT–EGFR and EGFR–HiBiT were coplated at a 1:1 ratio in a 96-well plate or an 8-chamber slide. The following day, cells were transfected with mRNA encoding PrismaLgBiT and incubated overnight with 100 nM JF549 HaloTag ligand. Following serum starvation, Vivazine and purified LgBiT were added to initiate substrate release and surface complementation, followed by treatment with 300 ng/mL EGF or vehicle control. Filtered luminescence images and reading were collected continuously for 2 h using the GloMax Galaxy imager and GloMax Discover plate-reader, respectively (Figure ). Deconvolved plate-based readings (Figure B) and complementary time-lapse images revealed rapid EGF-induced receptor internalization followed by slower degradation kinetics. These temporal dynamics were consistent with reported EGFR trafficking dynamics. Together, these results demonstrate that our two-color system is suitable for simultaneous quantitative tracking of two distinct events within the same sample.

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Monitoring EGF-induced EGFR internalization and degradation. HeLa cells expressing either HiBiT-EGFR or EGFR-HiBiT were cocultured, transfected with PrismaLgBiT’s mRNA, and incubated with JF549 HaloTag ligand. After 4 h of starvation with Vivazine and purified LgBiT added at the final hour, cells were treated with 300 ng/mL EGF or vehicle control. (A) Representative merged 2-channel images acquired over 120 min (3 min exposures). Entire time course provided in Figure S18. (B) Deconvolved plate-reader readouts. The EGF-treated signals were further normalized to untreated controls. Error bars represent the SD of 3 replicates.

Conclusions

By pairing NanoLuc-based reporters with engineered red-shifted NanoPrism reporters, we developed a modular two-color bioluminescent platform that operates with a single substrate. The system generates two bright, spectrally distinct emission signals (>100 nm separation), which can be detected and quantitatively deconvolved using standard plate-readers and bioluminescence imagers equipped with appropriate filters. Notably, the use of a shared substrate enables real-time kinetic measurements while simplifying experimental workflows. Together, these features support simultaneous tracking of two molecular readouts within the same sample, either in cocultured cell populations or coexpressed in a single cell.

A key feature of the NanoPrism design is its binary format, which supports both high- and low-affinity complementation. Here, we applied the high-affinity configuration in HiBiT knock-in cells for the sensitive and quantitative detection of endogenously expressed proteins. In targeted protein degradation assays, HiBiT-tagged GAPDH and GSPT1 were used as reference controls alongside NLuc–BRD4 degradation, whereas EGFR–HiBiT was used to monitor ligand-induced receptor internalization and degradation. We also applied the low-affinity, facilitated complementation to monitor the EGF-induced EGFR–Grb2 interaction in real time. These examples illustrate how the binary NanoPrism can be tuned to support distinct biological readouts, ranging from the detection of protein abundance to dynamic tracking of reversible interactions. Beyond these examples, this two-color framework can be extended to additional combinations of complementation-based reporters, including two HiBiT-tagged targets, two interaction-based readouts, or mixed interaction and abundance-based measurements in cocultured cell populations.

Functionally, this two-color platform enables two-parameter analyses in real time. Using protein degradation and receptor trafficking models, we showed that this system can resolve parallel and sequential biological events within the same sample including changes in protein abundance, pathway interactions, and downstream signaling responses. Across these applications, the overall concordance between population-averaged plate-based measurements and bioluminescence imaging analyses supports the robustness of our signal deconvolution strategy.

In summary, this two-color platform provides a simple and modular approach for quantitative, multiplexed analyses of cellular events using standard plate-readers. Its compatibility with plate-based formats supports scalability to higher-throughput two-color assays, particularly in settings in which sample availability is limited.

Methods

All methods are included in the Supporting Information.

Supplementary Material

cb6c00094_si_001.pdf (2.8MB, pdf)
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The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acschembio.6c00094.

  • Supporting Information Materials and Methods (experimental procedures, materials, and construct sequences); Supporting Information Figures S1–S18 (design of intramolecular BRET reporters, predicted structural models, two-color bioluminescent reporter system, simultaneous tracking of target degradation with an internal control and distinct steps in a ligand-induced receptor trafficking pathway) (PDF)

  • Time courses of target degradation with internal control (Videos V1–V3). Video V1 (AVI)

  • Video V2 (AVI)

  • Video V3 (AVI)

The authors declare the following competing financial interest(s): Eleven authors are Promega employees.

References

  1. Love A. C., Prescher J. A.. Seeing (and Using) the Light: Recent Developments in Bioluminescence Technology. Cell Chem. Biol. 2020;27(8):904–920. doi: 10.1016/j.chembiol.2020.07.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Roda A., Guardigli M., Michelini E., Mirasoli M.. Bioluminescence in analytical chemistry and in vivo imaging. Trends Anal. Chem. 2009;28:307–322. doi: 10.1016/j.trac.2008.11.015. [DOI] [Google Scholar]
  3. Sakama A., Orioka M., Hiruta Y.. Current advances in the development of bioluminescent probes toward spatiotemporal trans-scale imaging. Biophys. Physicobiol. 2024;21(Supplemental):e211004. doi: 10.2142/biophysico.bppb-v21.s004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Zambito G., Chawda C., Mezzanotte L.. Emerging tools for bioluminescence imaging. Curr. Opin. Chem. Biol. 2021;63:86–94. doi: 10.1016/j.cbpa.2021.02.005. [DOI] [PubMed] [Google Scholar]
  5. Yao Z., Zhang B. S., Prescher J. A.. Advances in bioluminescence imaging: new probes from old recipes. Curr. Opin. Chem. Biol. 2018;45:148–156. doi: 10.1016/j.cbpa.2018.05.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Branchini B. R., Southworth T. L., Khattak N. F., Michelini E., Roda A.. Red- and green-emitting firefly luciferase mutants for bioluminescent reporter applications. Anal. Biochem. 2005;345(1):140–148. doi: 10.1016/j.ab.2005.07.015. [DOI] [PubMed] [Google Scholar]
  7. Ni Y., Rosier B., van Aalen E. A., Hanckmann E. T. L., Biewenga L., Pistikou A. M., Timmermans B., Vu C., Roos S., Arts R.. et al. A plug-and-play platform of ratiometric bioluminescent sensors for homogeneous immunoassays. Nat. Commun. 2021;12(1):4586. doi: 10.1038/s41467-021-24874-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Michelini E., Cevenini L., Mezzanotte L., Ablamsky D., Southworth T., Branchini B., Roda A.. Spectral-resolved gene technology for multiplexed bioluminescence and high-content screening. Anal. Chem. 2008;80(1):260–267. doi: 10.1021/ac7016579. [DOI] [PubMed] [Google Scholar]
  9. Liu S., Su Y., Lin M. Z., Ronald J. A.. Brightening up Biology: Advances in Luciferase Systems for in Vivo Imaging. ACS Chem. Biol. 2021;16(12):2707–2718. doi: 10.1021/acschembio.1c00549. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Hattori M., Wazawa T., Orioka M., Hiruta Y., Nagai T.. Creating coveted bioluminescence colors for simultaneous multi-color bioimaging. Sci. Adv. 2025;11(4):eadp4750. doi: 10.1126/sciadv.adp4750. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Yao Z., Brennan C. K., Scipioni L., Chen H., Ng K. K., Tedeschi G., Parag-Sharma K., Amelio A. L., Gratton E., Digman M. A.. et al. Multiplexed bioluminescence microscopy via phasor analysis. Nat. Methods. 2022;19(7):893–898. doi: 10.1038/s41592-022-01529-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Breton B., Sauvageau E., Zhou J., Bonin H., Le Gouill C., Bouvier M.. Multiplexing of multicolor bioluminescence resonance energy transfer. Biophys. J. 2010;99(12):4037–4046. doi: 10.1016/j.bpj.2010.10.025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Ruigrok H. J., Shahid G., Goudeau B., Poulletier de Gannes F., Poque-Haro E., Hurtier A., Lagroye I., Vacher P., Arbault S., Sojic N.. et al. Full-Spectral Multiplexing of Bioluminescence Resonance Energy Transfer in Three TRPV Channels. Biophys. J. 2017;112(1):87–98. doi: 10.1016/j.bpj.2016.11.3197. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Xu Y., Piston D. W., Johnson C. H.. A bioluminescence resonance energy transfer (BRET) system: application to interacting circadian clock proteins. Proc. Natl. Acad. Sci. U. S. A. 1999;96(1):151–156. doi: 10.1073/pnas.96.1.151. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Hall M. P., Unch J., Binkowski B. F., Valley M. P., Butler B. L., Wood M. G., Otto P., Zimmerman K., Vidugiris G., Machleidt T.. et al. Engineered luciferase reporter from a deep sea shrimp utilizing a novel imidazopyrazinone substrate. ACS Chem. Biol. 2012;7(11):1848–1857. doi: 10.1021/cb3002478. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Machleidt T., Woodroofe C. C., Schwinn M. K., Mendez J., Robers M. B., Zimmerman K., Otto P., Daniels D. L., Kirkland T. A., Wood K. V.. NanoBRET--A Novel BRET Platform for the Analysis of Protein-Protein Interactions. ACS Chem. Biol. 2015;10(8):1797–1804. doi: 10.1021/acschembio.5b00143. [DOI] [PubMed] [Google Scholar]
  17. Hiblot J., Yu Q., Sabbadini M. D. B., Reymond L., Xue L., Schena A., Sallin O., Hill N., Griss R., Johnsson K.. Luciferases with Tunable Emission Wavelengths. Angew. Chem. Int. Ed. 2017;56(46):14556–14560. doi: 10.1002/anie.201708277. [DOI] [PubMed] [Google Scholar]
  18. Dixon A. S., Schwinn M. K., Hall M. P., Zimmerman K., Otto P., Lubben T. H., Butler B. L., Binkowski B. F., Machleidt T., Kirkland T. A.. et al. NanoLuc Complementation Reporter Optimized for Accurate Measurement of Protein Interactions in Cells. ACS Chem. Biol. 2016;11(2):400–408. doi: 10.1021/acschembio.5b00753. [DOI] [PubMed] [Google Scholar]
  19. Schwinn M. K., Machleidt T., Zimmerman K., Eggers C. T., Dixon A. S., Hurst R., Hall M. P., Encell L. P., Binkowski B. F., Wood K. V.. CRISPR-Mediated Tagging of Endogenous Proteins with a Luminescent Peptide. ACS Chem. Biol. 2018;13(2):467–474. doi: 10.1021/acschembio.7b00549. [DOI] [PubMed] [Google Scholar]
  20. Encell L. P., Friedman Ohana R., Zimmerman K., Otto P., Vidugiris G., Wood M. G., Los G. V., McDougall M. G., Zimprich C., Karassina N.. et al. Development of a dehalogenase-based protein fusion tag capable of rapid, selective and covalent attachment to customizable ligands. Curr. Chem. Genomics. 2012;6:55–71. doi: 10.2174/1875397301206010055. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kincaid V. A., Wang H., Sondgeroth C. A., Torio E. A., Ressler V. T., Fitzgerald C., Hall M. P., Hurst R., Wood M. G., Gilden J. K.. et al. Simple, Rapid Chemical Labeling and Screening of Antibodies with Luminescent Peptides. ACS Chem. Biol. 2022;17(8):2179–2187. doi: 10.1021/acschembio.2c00306. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Thirukkumaran O. M., Wang C., Asouzu N. J., Fron E., Rocha S., Hofkens J., Lavis L. D., Mizuno H.. Improved HaloTag Ligand Enables BRET Imaging With NanoLuc. Front. Chem. 2020;7:938. doi: 10.3389/fchem.2019.00938. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Chu J., Oh Y., Sens A., Ataie N., Dana H., Macklin J. J., Laviv T., Welf E. S., Dean K. M., Zhang F.. et al. A bright cyan-excitable orange fluorescent protein facilitates dual-emission microscopy and enhances bioluminescence imaging in vivo. Nat. Biotechnol. 2016;34(7):760–767. doi: 10.1038/nbt.3550. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Wohlwend J., Corso G., Passaro S., Reveiz M., Leidal K., Swiderski W., Portnoi T., Chinn I., Silterra J., Jaakkola T.. et al. Boltz-1 Democratizing Biomolecular Interaction Modeling. bioRxiv. 2024:624167. doi: 10.1101/2024.11.19.624167. [DOI] [Google Scholar]
  25. Stryer L.. Fluorescence energy transfer as a spectroscopic ruler. Annu. Rev. Biochem. 1978;47:819–846. doi: 10.1146/annurev.bi.47.070178.004131. [DOI] [PubMed] [Google Scholar]
  26. Winter G. E., Mayer A., Buckley D. L., Erb M. A., Roderick J. E., Vittori S., Reyes J. M., di Iulio J., Souza A., Ott C. J.. et al. BET Bromodomain Proteins Function as Master Transcription Elongation Factors Independent of CDK9 Recruitment. Mol. Cell. 2017;67(1):5–18 e19. doi: 10.1016/j.molcel.2017.06.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Nishiguchi G., Keramatnia F., Min J., Chang Y., Jonchere B., Das S., Actis M., Price J., Chepyala D., Young B.. et al. Identification of Potent, Selective, and Orally Bioavailable Small-Molecule GSPT1/2 Degraders from a Focused Library of Cereblon Modulators. J. Med. Chem. 2021;64(11):7296–7311. doi: 10.1021/acs.jmedchem.0c01313. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Schindelin J., Arganda-Carreras I., Frise E., Kaynig V., Longair M., Pietzsch T., Preibisch S., Rueden C., Saalfeld S., Schmid B.. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods. 2012;9(7):676–682. doi: 10.1038/nmeth.2019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Riching K. M., Mahan S. D., Urh M., Daniels D. L.. High-Throughput Cellular Profiling of Targeted Protein Degradation Compounds using HiBiT CRISPR Cell Lines. J. Vis. Exp. 2020;165:e61787. doi: 10.3791/61787. [DOI] [PubMed] [Google Scholar]
  30. Schultz D. F., Billadeau D. D., Jois S. D.. EGFR trafficking: effect of dimerization, dynamics, and mutation. Front. Oncol. 2023;13:1258371. doi: 10.3389/fonc.2023.1258371. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Boursier M. E., Levin S., Zimmerman K., Machleidt T., Hurst R., Butler B. L., Eggers C. T., Kirkland T. A., Wood K. V., Friedman Ohana R.. The luminescent HiBiT peptide enables selective quantitation of G protein-coupled receptor ligand engagement and internalization in living cells. J. Biol. Chem. 2020;295(15):5124–5135. doi: 10.1074/jbc.RA119.011952. [DOI] [PMC free article] [PubMed] [Google Scholar]

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Supplementary Materials

cb6c00094_si_001.pdf (2.8MB, pdf)
Download video file (4.4MB, avi)
Download video file (4.2MB, avi)
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