DNA oligonucleotides used for cloning and construction of gene libraries were purchased from Hokkaido System Science. The sequences of all the oligonucleotides used in this study are provided in Supplementary Table 3. KOD-Plus (Toyobo Life Science) was used for non-mutagenesis PCR amplification. Products of PCR and restriction enzyme (RE) digestion were routinely purified using preparative agarose gel electrophoresis followed by DNA isolation using a QIAEX II gel extraction kit (Qiagen). Restriction endonucleases were purchased from New England Biolabs or Takara-bio, and used according to the manufacturer’s recommended protocol. Ligations were performed using T4 ligase in Rapid Ligation Buffer (Promega). Small-scale plasmid DNA preparation was performed by alkaline lysis of the bacterial pellet obtained from 1.5 ml of LB liquid culture followed by ethanol precipitation of the DNA. Large-scale plasmid DNA preparations were performed by alkaline lysis of the bacterial pellet derived from 200 ml of Luria-Bertani (LB) liquid culture followed by isopropanol precipitation, PEG 8000 precipitation and two rounds of phenol/chloroform extraction. cDNA sequences for all constructs were read by dye terminator cycle sequencing using the BigDye Terminator v1.1 Cycle Sequencing kit (Life Technologies). The luminescent substrate coelenterazine-h was purchased from Wako Chemicals, and furimazine was synthesized according to a previous report4. Cultured cells were not tested for the presence of Mycoplasma, as such contamination would not affect the conclusions made on the basis of our results.
The primary amino-acid sequence of Nluc4 was used as input to the I-TASSER structure prediction server30 using default parameters. The generation of figures depicting structures was done using UCSF chimera31.
Construction of five colour eNL variants
cDNA of mNeonGreen was provided by Allele Biotechnology. The deletion mutant libraries of mNeonGreen–Nluc and Venus–Nluc fusion constructs were generated as follows32. The cDNAs of C-terminally deleted mNeonGreen mutants (mNGΔC0-10) and Venus mutants (VenusΔC0-12) were amplified by PCR. They were digested with BamHI and KpnI. The cDNAs of N-terminally deleted Nluc mutants (NlucΔN1-5 for mNG and NlucΔN1-6 for Venus) were amplified by PCR and digested with EcoRI and KpnI. The digested PCR fragments were gel-purified, mixed together and cloned in-frame into the BamHI/EcoRI sites of pRSETB (Invitrogen) for bacterial expression (Supplementary Fig. 1). After screening, the linker amino acids (encoded by KpnI, –Gly–Thr–) of deletion mutants was randomized by inverse PCR techniques (nucleotide sequence NNKNNK, where N=A, G, C and T; and K=G and T) to generate 400 amino-acid combinations (1,024 nucleotide combinations; Supplementary Fig. 3).
Considering the 230th–239th residues of mTQ2 and 464th–476th residues of tdTomato could be removed without disrupting the function and the ΔN4Nluc was selected in the screening of GeNL, we chose mTQ2ΔC10-GT-ΔN4Nluc and tdTomatoΔC8-GT-ΔN4Nluc as templates for CeNL and ReNL development. We constructed linker libraries by systematic truncation of the connecting region between either mTQ2 or tdTomato and Nluc using inverse PCR techniques (Supplementary Fig. 5). For each library, two randomized amino-acid residues (nucleotide sequence NNKNNK, where N=A, G, C and T; and K=G and T) were placed between the C terminus of either mTQ2 or tdTomato and the N terminus of Nluc, to generate 400 amino-acid combinations (1,024 nucleotide combinations).
The cDNAs of mKOκ with various length flexible linkers (no linker to GGSGGSGGS for 5′-end of mKOκ and no linker to TLGMDELYK for 3′-end) were amplified by PCR. They were digested with XhoI and SacI, then gel-purified and mixed together. XhoI and SacI restriction sites were also introduced at the 50th/51th insertion site of the Nluc moiety in pRSETB_Nluc. This fragment was then ligated with the mKOκ fragment mixture to yield the pRSETB_mKOκ@51_Nluc library (Supplementary Fig. 7c).
The nucleotide sequences of GeNL, YeNL, CeNL, ReNL and OeNL are in Supplementary Note 4.
Covalent labelling of Nluc with eosin maleimide dye
Eosin-5-maleimide was purchased from Molecular Probes (no. E-118). To achieve the labelling at specific positions, we substituted the endogenous cysteine of Nluc (166th residue) with alanine, and four glycine residues (50th, 66th, 97th and 136th) were substituted with cysteine, respectively. The mutated Nluc was expressed in Escherchia coli and purified as explained below. The protein (50 μM) was pre-incubated with 5 mM dithiothreitol in 20 mM HEPES buffer (pH=7.4) for 30 min at room temperature. The solutions were then changed to fresh 20 mM HEPES buffer (pH=7.4) using a NAP-5 column (GE-Healthcare). The proteins were incubated with 100 μM eosin-5-maleimide in 20 mM HEPES buffer (pH=7.4) for 1 h at room temperature. The solutions were changed to fresh 20 mM HEPES buffer (pH=7.4) using a NAP-5 column to remove the free eosin-5-maleimide. The luminescent spectral measurement was performed as described in ‘LP characterization’ below.
Screening of mutants with bright and high-FRET efficiency
After transforming E. coli JM109 (DE3) with the mutagenized DNAs, we spread the bacterial cells homogeneously over 95 mm agar plates and incubated them at 37 °C for 12 h, before sitting for 2 days at room temperature to allow the chromophore maturation. Bright mutants with high-FRET efficiency were screened and selected in two steps. Initially, the E. coli colonies were poured with a phosphate-buffered saline (PBS) solution supplemented with 5 μM coelenterazine-h and examined with an LAS-1000 luminescence imaging system (GE Healthcare). Bright colonies were picked, and those mutants were inoculated in the 96-well plate supplemented with 100 μl liquid LB medium. The 96-well plate was incubated at 23 °C for 3 days before luminescence spectra were measured using a micro-plate reader (SH-9000; Corona Electric). A final concentration of 5 μM coelenterazine-h was used as the luminescent substrate for this measurement. Luminescent spectra were normalized at the 450 nm luminescence intensity and mutants with a high-FRET ratio (ratio of peak intensity at 520/450 nm for mNeonGreen, 530/450 nm for Venus, 565/450 nm for mKOκ and 585/450 nm for tdTomato) picked and subjected to DNA sequencing and protein characterization. Because the emission peaks of Nluc (donor, ∼460 nm) and mTQ2 (acceptor, 480 nm) were too close to discern, the cyan variants were directly purified and screened in vitro on the basis of brightness and FRET efficiency.
Construction of mammalian expression vectors
To ensure the robust expression of GeNL in mammalian cells, we replaced the wild-type codon with synthesized cDNA encoding the mNeonGreen with mammalian favourable codons obtained from Life Technologies (GeneArt Strings DNA Fragments). PCR-amplified eNLs were inserted into a pcDNA3 mammalian expression vector using BamHI and EcoRI RE sites. We localized eNLs to mitochondria, plasma membrane and nucleus, respectively, by replacing the Nano-lantern sequence with the eNL sequence in pcDNA3-CoxVIIIx2-Nano-lantern (a duplicated mitochondrial localization sequence derived from the subunit-VIII precursor of human cytochrome c oxidase (Cox-VIII) at the N terminus); pcDNA3-Nano-lantern-H2B (a DNA-binding protein histone 2B (H2B) at C terminus); and pcDNA3-lyn-Nano-lantern (a myristoylation and palmitoylation sequence from lyn kinase at N terminus)9. For peroxisome localization, PCR-amplified eNLs with SKL (a peroxisome localization sequence) were inserted into a pcDNA3 mammalian expression vector using BamHI and EcoRI RE sites. For nucleolus and ER localization, we replaced Phamret with eNLs in pcDNA3-Phamret-fibrillarin or pcDNA3-Phamret-ER (signal peptide from calreticulin at the N terminus and an ER retention signal at the C terminus) using BamHI and EcoRI or BamHI and KpnI RE sites33. A 5-amino-acid linker (GGSGGT) was inserted between the eNLs and ER retention signal sequences. For paxillin, zyxin and vimentin localization, we replaced the Kohinoor sequence with the GeNL sequence in pcDNA3-paxillin-Kohinoor, pcDNA3-zyxin-Kohinoor or pcDNA3-vimentin-Kohinoor using BamHI and EcoRI RE sites34. For β-actin localization, we replaced the Kohinoor sequence with the eNLs sequence in the pcDNA3-Kohinoor-β-actin vector using HindIII and KpnI RE sites. The open reading frames (ORFs) of vimentin, paxillin and zyxin included a 17-amino-acid linker peptide (GTGSGGGGSGGGGSGGS), and β-actin included a 20-amino-acid linker peptide (GGSGGSGGSGGSGGEFQIST). For clathrin localization, we replaced the Kohinoor sequence with the eNLs sequence in pEGFP-N1-Kohinoor-clathrin using NheI and BglII RE sites. Clathrin had a 12-amino-acid linker peptide (RSRAQASNSAVD). For β-tubulin localization of GeNL, we replaced the Kohinoor sequence with the GeNL sequence in pEGFP-N1-β-tubulin-Kohinoor using SalI and NotI RE sites. A 21-amino-acid linker (QSTGSGGGGSGGSTVPRARDP) was inserted between the GeNL and β-tubulin sequences. For β-tubulin localization of CeNL, YeNL, OeNL and ReNL, a β-tubulin sequence was inserted upstream of CeNL, YeNL, OeNL and ReNL in pcDNA3-CeNL, pcDNA3-YeNL, pcDNA3-OeNL and pcDNA3-ReNL using HindIII and EcoRI RE sites. The ORFs of β-tubulin included a 23-amino-acid linker peptide (QSTVPRARDPGSGGGSGGGSGEF). For vinculin lozalization, a vinculin sequence was inserted downstream of GeNL in pcDNA3-GeNL using KpnI and EcoRI RE sites. For lysosome localization, a LAMP sequence was inserted upstream of eNLs in pcDNA3-eNLs using HindIII and BamHI RE sites. The ORFs of vinculin and LAMP included a 16-amino-acid linker peptide (GTGGGGSGGGGSGGSG) and a 17-amino-acid linker peptide (GTGSGGGGSGGGGSGGS). The cDNAs of LAMP and vinculin were amplified from the plasmids, mRuby2-Lysosomes-20 and pEGFP Vinculin, which were gifts from Michael Davidson and Kenneth Yamada (Addgene plasmid # 55902 and 50513, respectively). See Supplementary Table 3 for the polynucleotides used in this study.
Protein expression and purification
LP with an N-terminal polyhistidine tag was expressed in E. coli (JM109 (DE3)) at 23 °C for 65 h in LB bacterial growth medium supplemented with 0.1 mg ml−1 carbenicillin. Cells were collected and ruptured with a French press (Thermo Fisher Scientific), and recombinant proteins were purified from the supernatant using Ni-NTA agarose affinity columns (Qiagen) followed by buffer-exchange (20 mM HEPES, pH 7.4) gel filtration (PD-10 column, GE Healthcare). The whole purification process after rupture was conducted on ice to avoid protein degradation. The protein concentration was determined by Bradford method (Protein assay kit, Bio-Rad).
Recombinant proteins and furimazine were diluted with 20 mM HEPES buffer (pH 7.4), and emission spectra were measured with a photonic multichannel analyser PMA-12 (Hamamatsu Photonics) at room temperature using 500 ms exposures. The final concentrations of proteins and substrate were 100 nM and 25 μM, respectively. Furimazine was used for eNLs and Nluc as a substrate; coelenterazine-h was used for the other proteins. Experiments were performed at least in triplicate, and the averaged data were used for further analysis.
Luminescent quantum yield and kinetic parameters
The luminescent quantum yields were estimated from the total light output by the complete consumption of 0.05 pm of furimazine. The luminescent quantum yields were measured in triplicate with a micro-plate reader (SH-9000, Corona Electric). The photon sensitivity of the detector was calibrated with luminol chemiluminescence under following reaction mixture: 200 nM horseradish peroxidase; 200 nM luminol; and 2 mM hydrogen peroxide in K2CO3 aqueous solution35. The concentration of luminol (Wako, Osaka, Japan) was characterized by the absorbance at 347 nm with using the extinction coefficient of 7,640 M−1 cm−1 as reported previously35. The wavelength characteristics of the detector were adjusted in relation to the photonic multichannel analyzer PMA-12 (Hamamatsu Photonics). The final concentrations of LP and furimazine were 1 nM and 500 pM, respectively. The protein and substrate solutions were diluted with HEPES buffer (50 mM HEPES, pH 7.5) supplemented with <0.1% casein.
Kinetic parameters were measured from the reactions of final 10 pM LP with final furimazine concentrations of 0.025, 0.051, 0.10, 0.20, 0.41, 0.81, 1.6, 3.3, 6.5 and 13 μM, respectively. The initial reaction velocities were measured as the integrated luminescence intensities for the initial 12 s. Michaelis–Menten constants (Km) and maximum reaction velocities (Vmax) were estimated from the nonlinear fitting to the Michaelis–Menten equation using Origin7 software (OriginLab).
Detection of luminescence from single-molecule GeNL
To immobilize the GeNL protein on the glass surface, we prepared the Ni-NTA agarose glass coverslips following a previous study36. The fluorescent beads were used for focusing on the glass surface. First, fluorescent beads (FluoSpheres sulfate microspheres, 0.2 μm diameter, yellow-green fluorescent, no. F8848, Invitrogen) in MOPS/KCl buffer (10 mM MOPS and 100 mM KCl, pH=7.2) were placed on the Ni-NTA agarose glass and incubated for 5 min and removed. Subsequently, 10 pmol of GeNL purified protein in MOPS/KCl buffer was placed on it and incubated for 5 min and washed with MOPS/KCl buffer three times to remove the unbound proteins. Just before observation, 50 μM furimazine was added. To detect the luminescence from a single molecule, an inverted microscope (LV-200, Olympus) equipped with a × 100 objective (Olympus, UPlanSApo, numerical aperture 1.4) and × 0.5 relay lens was used. Emission signals were detected by a cooled EM-CCD (electron-multiplying, charge-coupled device) camera (ImagEM, Hamamatsu Photonics) with 1 × 1 binning settings and 180 s exposure time.
A series of images was processed by Fiji. The background (defined as mean intensity at a region where no luminescence spot was detected) was subtracted using ImageJ’s built-in function. Images were transformed to a 32-bit float and the count of each pixel was converted to photon number (designated as raw images; see also Supplementary Note 2). A median filter (radius size=1) was used to smooth the images. Then luminescence spots were segmented by means of the Particle Track Analysis (PTA ver1.2) plug-in of ImageJ (https://github.com/arayoshipta/projectPTAj) to define square ROIs (filtered by minimum intensity 0.9, size >5 pixel2). The total photon number at each ROI was calculated from the raw images and plotted in a histogram (Fig. 2b) using Origin7 software.
Preparation and luminescence imaging of HeLa cell
HeLa (RIKEN BRC) cells were cultured on collagen-coated 35 mm glass-bottom dishes in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS). The next day, HeLa cells (∼70% confluency) were transfected with 4.0 μg plasmid DNA using Lipofectamine 2000 Transfection Reagent (Life Technologies) according to the manufacture’s recommended protocol. Medium was replaced after 8 h, and the cells grown for an additional 16 h in a CO2 incubator (Sanyo) at 37 °C in 5% CO2. HeLa cells were washed with phenol red-free DMEM/F12 and imaged in phenol red-free DMEM/F12. Just before observation, 20 μM furimazine was added to the imaging medium. To observe eNL signals in living HeLa cells, an inverted microscope (LV-200, Olympus) equipped with a × 100 objective (Olympus, UPlanSApo, numerical aperture 1.4) and × 0.5 relay lens was used. Emission signals were detected by an EM-CCD camera (ImagEM, Hamamatsu Photonics) with 1 × 1 (for eNL) or 2 × 2 (for GeNL(Ca2+)) binning settings. To observe the localization of clathrin light chain labelled with CeNL in living HeLa cells, an inverted microscope (IX83, Olympus, Japan) equipped with a × 100 objective (Olympus, UPlanSApo, numerical aperture 1.4) was used. Emission signals were detected by an EM-CCD camera (Evolve 512, Photometrics) with 1 × 1 binning.
Preparation and Ca2+ imaging of GH3 cells
GH3 (rat pituitary tumour, ATCC CCL-82.1) cells were cultured on poly-D-lysine-coated 35 mm glass-bottom dishes in DMEM/F12 supplemented with 15% horse serum and 2.5% FBS. For Ca2+ imaging using either GCaMP3 or GeNL(Ca2+), GH3 cells (∼70% confluency) were transfected with 4.0 μg plasmid DNA using Lipofectamine 2000 Transfection Reagent (Life Technologies) according to the manufacture’s recommended protocol. The medium was replaced after 8 h, and the cells were grown for an additional 16 h in a CO2 incubator (Sanyo) at 37 °C in 5% CO2. For Ca2+ imaging using Fura-2, 5.0 μM Fura-2-AM (Dojindo Laboratories, F016) was loaded onto GH3 cells in Hanks’ Balanced Salt solution (Sigma) supplemented with 1 × PowerLoad (Invitrogen) for 1 h at 37 °C. The GH3 cells were washed with phenol red-free DMEM/F12 and imaged in phenol red-free DMEM/F12. Just before observation of GeNL(Ca2+), 20 μM furimazine was added to the imaging medium.
Images were captured using an inverted microscope (IX83, Olympus, Japan) equipped with a × 40 objective (Olympus, UApo/340, numerical aperture 1.35) or a × 60 objective (Olympus, PlanApo, numerical aperture 1.4), EM-CCD camera (Evolve 512, Photometrics). The dual-excitation ratio imaging of Fura-2 was conducted with Semrock FF01-340/26 for shorter excitation wavelengths, FF01-387/11 for longer excitation wavelengths, an FF409-DiO2 dichroic mirror and an FF01-510/84 emission filter. The excitation filters for dual excitation images of Fura-2 were alternated using a filter exchanger with 100 ms exposure times for each channels, while sample were illuminated only during exposure time. Images of Fura-2 or GeNL(Ca2+) were acquired at 1.3 Hz frame rate with 2 × 2 binning. The Ca2+ imaging of GCaMP3 was conducted with Omega 475AF20 excitation filter, Semrock FF495-Di03 dichroic mirror and an FF01-525/45 emission filter. The images of GCaMP or GeNL(Ca2+) were acquired at 33 Hz frame rate with 2 × 2 binning and 30 ms exposure time. Cells were maintained on the microscope at 37 °C by using a stage-top incubator (Tokai Hit, Fujinomiya, Japan).
Multicolour imaging with linear unmixing
For multicolour luminescence imaging, five images were acquired with Semrock FF01-447/60 filter (for Nluc), Olympus BA460-510CFP (for CeNL), Semrock FF01-525/35 filter (for GeNL), Semrock FF01-562/40 (for OeNL) and Semrock FF01-593/40 (for ReNL). Five HeLa cells (expressing each fusion construct) were imaged with identical conditions to determine the coefficients for linear unmixing. The signals from NLuc, CeNL, GeNL, OeNL and ReNL were then separated by linear unmixing using these coefficients by PrizMage software (Molecular Devices).
Searching for insertion site of NanoLuc
The Mutation Generation System Kit (Thermo Fisher Scientific) was used to construct the insertion mutation library of Nluc variants. Following the manufacturer’s protocols, an in vitro transposition reaction was performed with 280 ng of pRSETB-Nluc and the M1-KanR Entranceposon that contains the kanamycin resistance gene. The reaction mixture was used to transform E. coli XL10-Gold. The transformation reaction was plated on two LB/agar/amp plates supplemented with kanamycin (25 μg ml−1) and incubated for 12 h at 37 °C. Bacterial colonies were scraped from all the plates and then plasmid DNA was purified. The plasmid DNA was digested with BamHI and EcoRI and the resulting fragments were separated by agarose gel electrophoresis. The 1.6 kb (0.5 kb Nluc gene+Entranceposon) band was excised, isolated and ligated with appropriately digested pRSETB. The ligation reaction was used to transform E. coli as described above. The transformation reaction was diluted into LB/amp and incubated overnight at 37 °C with shaking at 225 r.p.m. Plasmid DNA was isolated from this culture, digested with NotI to remove the KanR gene, and the resulting fragments were separated by agarose gel electrophoresis. The 3.4 kb fragment (modified Nluc gene+pRSETB) was excised and the DNA isolated as described above. The isolated DNA fragment was self-ligated to circularize the plasmid and thus provide the library of Nluc variants with random 15 bp (five codon) insertions.
The plasmid library of Nluc with random insertions was used to transform the chemically competent E. coli strain JM109(DE3). The transformed bacteria were plated on LB agar. Plates were incubated for 12 h at 37 °C. Following a 20 h incubation at 23 °C, a PBS solution containing 5 μM coelenterazine-h was added to the E. coli colonies, and were examined using an LAS-1000 luminescence imaging system (GE Healthcare). The 30 brightest colonies were picked up and cultured in liquid LB medium for 12 h. They were subjected to plasmid DNA isolation and subsequent DNA sequencing as described above.
To determine the insertion sites of Nluc that yield a large signal increase with Ca2+, we used the TorA protein export plasmid (pTorPE) in which a Ca2+ indicator localized to the E. coli periplasm can be placed on Ca2+-bound states and easily extracted by cold osmotic shock37. We subcloned Nluc into pTorPE using SalI and HindIII RE sites. To obtain cDNA fragment encoding the CaM–M13 peptide of Nano-lantern(Ca2+), we digested a pRSETB_Nano-lantern(Ca2+)_600 with NcoI and SacI. These restriction sites were also introduced at the identified insertion sites of pTorPE_Nluc (Supplementary Fig. 15). These fragments were then ligated to yield pTorPE_CaM-M13_Nluc and then mixed together. DH10B bacterial cells were transformed and spread on LB/amp agar plates supplemented with L-arabinose (0.02%). Following a 12 h incubation at 37 °C, a PBS solution containing 5 μM coelenterazine-h was added to the E. coli colonies, before examination using an LAS-1000 luminescence imaging system (GE Healthcare). The brightest colonies were picked and cultured in liquid LB medium for 12 h. The brightest colonies were subjected to further DNA purification and sequencing. An aliquot of E. coli suspensions was also used for secondary screening, in which Ca2+-dependent change in luminescence was assessed using the protein extracted from periplasmic fraction of E. coli. Extraction of periplasmic protein from E. coli was performed by cold osmotic shock procedure as described before37. Briefly, bacterial cells were collected by centrifugation at 13,000g for 2 min at 4 °C and gently resuspended in 500 μl of ice-cold pH 8.0 buffer containing 30 mM Tris-HCl, 1 mM EDTA and 20% sucrose. After 5 min of gentle agitation on ice, the bacteria were again pelleted by centrifugation (9,000g for 5 min at 4 °C), resuspended in 400 μl of ice-cold 5 mM MgSO4 and gently agitated for 10 min on ice. Following centrifugation to pellet the intact bacteria (9,000g for 5 min at 4 °C), the supernatant (the osmotic shock fluid containing the periplasmic protein fraction) was collected. The Ca2+-dependent change in luminescence was measured by a micro-plate reader as described in ‘In vitro characterization of GeNL(Ca2+)’.
Construction of GeNL(Ca2+)
NcoI and SacI restriction sites were introduced at the 66/67 and 69/70 insertion sites of the Nluc moiety in pRSETB_GeNL. This fragment was then ligated with a CaM–M13 fragment to yield pRSETB_CaM-M13@67_GeNL and pRSETB_CaM-M13@70_GeNL. To restore the brightness of pRSETB_CaM-M13@67_GeNL, we performed error-prone PCR for CaM–M13 and a C-terminal fragment of Nluc under conditions to achieve a mutation rate of 5 bp kb−1. Primers containing NcoI/EcoRI were used for re-insertion of CaM–M13_Nluc (67th–171th) mutant cDNA into pRSETB_CaM-M13@67_GeNL. JM109(DE3) bacterial cells were transformed and then spread on LB/amp agar plates. Following a 12 h incubation at 37 °C, a PBS solution containing 5 μM coelenterazine-h was added to the E. coli colonies before examination using a LAS-1000 luminescence imaging system (GE Healthcare). The brightest colonies were picked and cultured in liquid LB medium for 12 h. They were subjected to further DNA purification and sequencing. The mutant with the largest brightness and signal increase was designated GeNL(Ca2+)_480 (latter number indicates Kd value to Ca2+).
To generate affinity variants of GeNL(Ca2+), we introduced various length linkers between CaM and M13 by Quik Change site-directed random mutagenesis38. For mammalian expression experiments, GeNL(Ca2+) cDNA digested with BamHI and EcoRI was cloned into the BamHI/EcoRI site of the pcDNA3 vector. The nucleotide sequences of GeNL(Ca2+)_60, GeNL(Ca2+)_250, GeNL(Ca2+)_480 and GeNL(Ca2+)_520 are in Supplementary Note 4.
For the AAV expression system, pHelper and pAAV-DJ were obtained from Cell Biolabs, Inc39. The cDNA of GeNL(Ca2+)_520 was amplified by PCR using a sense primer containing a BamHI site followed by a Kozak sequence and a reverse primer containing an EcoRI site followed by a stop codon. The cDNA for GeNL(Ca2+)_520 replaced the ArchT-GFP sequence in pAAV-CAG-ArchT-GFP. pAAV-CAG-ArchT-GFP was a gift from Edward Boyden (Addgene plasmid #29777).
In vitro characterization of GeNL(Ca2+)
Emission intensity of the purified proteins was measured using a micro-plate reader (SH-9000, Corona Electric). A final concentration of 5 μM coelenterazine-h was used as the luminescent substrate for these measurements. Experiments were performed at least in triplicate, and the averaged data were used for further analysis. Ca2+ titrations were performed by the reciprocal dilution of Ca2+-saturated and Ca2+-free buffers containing 10 mM MOPS, 100 mM KCl and 10 mM EGTA with or without 10 mM Ca2+ added as CaCO3, at pH 7.2, 25 °C. The free Ca2+ concentrations were calculated using 0.15 μM for the apparent Kd value of EGTA for Ca2+. The Ca2+ titration curve was used to calculate the apparent Kd value by nonlinear regression analysis. The averaged data were fitted to a single Hill equation using Origin7 software (OriginLab).
AAV production and infection
HEK293T (RIKEN BRC Cell Bank RCB2202) cells were grown in DMEM (Sigma) containing heat-inactivated 10% FBS at 37 °C in 5% CO2. Equal amounts of pHelper, pAAV-DJ and pAAV_CAG_GeNL(Ca2+)_520 were transfected by FuGENEHD transfection reagent (Roche) following the manufacturer’s protocol. Three days after transfection, the viruses were collected. The solution containing AAV was distributed into small aliquots and stored at −80 °C.
hiPSC-CM culture and imaging
Human iPS cells (hiPS, 201B7, RIKEN BRC) were cultured with a primate ES medium (Reprocell Inc.) and a 4 ng ml−1 human basic fibroblast growth factor (Wako Pure Chemical Industries, Ltd.) in an incubator with 5% CO2 at 37 °C on a mouse feeder cell layer (SNL, CBA-316, Cell Biolabs, Inc.). Embryoid bodies of hiPSC-CMs were prepared with the following procedure40. After carefully removing feeder cells with CTK solution (Reprocell Inc.), the one 60 mm dish of 80–90% confluent hiPS colonies were collected to the 15 ml centrifuge tube and centrifuge at 1,000 r.p.m. for 30 s, aspirated ES medium and change medium to 1 ml differentiation medium (RPMI+PVA), pipetting with 1 ml tip (WATSON, BIO LAB) for 20 times to obtain cell cluster size around 150 μm. The RPMI+PVA medium consist of RPMI Media 1640 (with L-glutamine), 4 mg ml−1 polyvinyl alcohol (P8136 Sigma-Aldrich, St. Louis, MO), 400 μM 1-thioglycerol, 1% chemically defined lipid concentrate (Thermo Fisher Scientific, 11905-031), 10 μg ml−1 recombinant human insulin (I9278, Sigma-Aldrich), 25 ng ml−1 human BMP4, 5 ng ml−1 human FGF2 (both from R&D systems) and 10 μM Y-27632 (Stemgent, Cambridge, MA). The suspended hiPS cells were transferred to 35 mm dish (Ultra Low Attachment Culture Dish, Corning) and place in incubator with 5% O2 and CO2 at 37 °C. Two days after incubation, the medium was replaced with the RPMI+FBS medium consisting of RPMI Media 1640, 20% FBS and 400 μM 1-thioglycerol, and the dishes were transferred to an incubator with 5% CO2 at 37 °C. Four days after incubation, 8–10 embryoid bodies were transferred to gelatinized 0.1% coverslips in six-well plates. The medium was replaced with fresh RPMI+FBS medium every 3 days afterwards. The embryoid bodies were visually assessed for contraction on 9 days after incubation. Then they were treated with AAV for 4 days before observation.
The hiPSC-CM was washed with Tyrode solution (Sigma) and imaged in Tyrode solution. Just before observation, 40 μM furimazine was added to the Tyrode solution. An inverted microscope (LV-200, Olympus) equipped with a × 100 objective (Olympus, UPlanSApo, numerical aperture 1.4) and × 0.5 relay lens was used. Emission signals were detected by an EM-CCD camera (ImagEM, Hamamatsu Photonics) with 4 × 4 (for GeNL(Ca2+)) binning settings. During the entire imaging period, the temperature was kept at 28 °C by a stage-top incubator. The background drift was manually subtracted using Origin7 software (OriginLab).
The data that support the findings of this study are available from the corresponding author on request. The nucleotide sequences of CeNL, GeNL, YeNL, OeNL, ReNL, GeNL(Ca2+)_60, GeNL(Ca2+)_250, GeNL(Ca2+)_480 and GeNL(Ca2+)_520 have been deposited to GenBank/EMBL/DDBJ database under the following entry IDs: LC128714; LC128715; LC128716; LC128717; LC128718; LC128719; LC128720; LC128721; and LC128722, respectively. The nucleotide sequences of all constructs are also in Supplementary Note 4.
Bioluminescence methodologies have been extraordinarily useful due to their high sensitivity, broad dynamic range, and operational simplicity. These capabilities have been realized largely through incremental adaptations of native enzymes and substrates, originating from luminous organisms of diverse evolutionary lineages. We engineered both an enzyme and substrate in combination to create a novel bioluminescence system capable of more efficient light emission with superior biochemical and physical characteristics. Using a small luciferase subunit (19 kDa) from the deep sea shrimp Oplophorus gracilirostris, we have improved luminescence expression in mammalian cells ∼2.5 million-fold by merging optimization of protein structure with development of a novel imidazopyrazinone substrate (furimazine). The new luciferase, NanoLuc, produces glow-type luminescence (signal half-life >2 h) with a specific activity ∼150-fold greater than that of either firefly (Photinus pyralis) or Renilla luciferases similarly configured for glow-type assays. In mammalian cells, NanoLuc shows no evidence of post-translational modifications or subcellular partitioning. The enzyme exhibits high physical stability, retaining activity with incubation up to 55 °C or in culture medium for >15 h at 37 °C. As a genetic reporter, NanoLuc may be configured for high sensitivity or for response dynamics by appending a degradation sequence to reduce intracellular accumulation. Appending a signal sequence allows NanoLuc to be exported to the culture medium, where reporter expression can be measured without cell lysis. Fusion onto other proteins allows luminescent assays of their metabolism or localization within cells. Reporter quantitation is achievable even at very low expression levels to facilitate more reliable coupling with endogenous cellular processes.
Bioluminescence is found across a diversity of life that includes bacteria, insects, fungi, and an abundance of marine organisms.1 It occurs when a photon-emitting substrate (luciferin) is oxidized by a generic class of enzymes called luciferases. These enzymes have been popular as reporters of cellular physiology because of their ability to provide highly sensitive quantitation with broad linearity. Firefly (Fluc, 61 kDa) and Renilla (Rluc, 36 kDa) luciferases have accounted for the majority of such applications, particularly for elucidating molecular processes coupled to gene expression. More recently, bioluminescence has been applied to other aspects of cellular analysis. Fluc has been configured into assay reagents for quantitation of cell viability, apoptosis, and various processes linked to cellular metabolism.2,3 Luciferases have been fused to other proteins to monitor their metabolism4 and interactions,5 circularly permuted to create intracellular biosensors,6 and split into fragments to monitor protein interactions in living cells.7
The widely recognized utility of bioluminescence has spurred investigation of alternative luciferases, predominantly from marine organisms. Luciferase genes have been derived from the copepods Gaussia princeps (20 kDa)8 and Metridia longa (24 kDa),9 the ostracod Cypridina noctiluca (61 kDa),10 the dinoflagellate Pyrocystis lunula (40 kDa),11 and the deep sea shrimp, Oplophorus gracilirostris (106 kDa).12Gaussia luciferase in particular has been used as a secreted reporter in mammalian cells,13 reportedly providing increased assay sensitivity owing to its bright luminescence and accumulation in the cell culture medium.8 However, the light intensity decays rapidly under most conditions, thus requiring luminometers equipped with injectors to measure the transient peak luminescence. Furthermore, the coelenterazine substrate is prone to chemical instability and high autoluminescence background,14 properties that make sample handling difficult and decrease assay sensitivity.
Although bright luminescence is generally desirable, a sustained signal with low background is necessary to enable efficient assay methods with high sensitivity. Preferably the luciferase should be small, monomeric, and structurally stable to environmental conditions. The luciferase from the deep sea shrimp, Oplophorus, suggested a route for achieving these capabilities.15 The native luciferase is secreted by the shrimp in brilliant luminous clouds as a defense mechanism against predation. Like many marine luciferases, it utilizes coelenterazine in an ATP-independent reaction to produce blue light (spectral maximum 454 nm). The native enzyme was found to be structurally stable with a high specific activity and quantum yield.15 Although it has a heteromeric structure consisting of two 35 kDa subunits and two 19 kDa subunits, cDNA clones revealed that bioluminescence activity was associated only with the smaller subunit (Oluc-19).12 Unfortunately, this smaller recombinant subunit does not retain many of the desirable features evident in the native enzyme, as it is unstable and poorly expressed in the absence of the 35 kDa partner.12
Our examination of the Oluc-19 amino acid sequence revealed an association with the family of intracellular lipid binding proteins (iLBPs), indicating that the underlying protein architecture should support development of a stable structure. A program to achieve a more optimal enzyme structure was undertaken, which also afforded an opportunity to explore variants of the luminogenic substrate. This resulted in a novel bioluminescent system consisting of an engineered luciferase called NanoLuc (Nluc) coupled with a novel coelenterazine analogue called furimazine. The combination of these generates much brighter luminescence than Fluc or Rluc, provides improved physical and chemical characteristics, and is generally compatible with mammalian cells. Overall, we found that Nluc performed exceptionally well as a reporter and anticipate that it will further advance the use of bioluminescence for cellular analysis.
Results and Discussion
Prediction of Stabilizing Amino Acids
There is no experimentally determined structure for Oluc-19, and standard sequence search methods failed to uncover significant similarities with known proteins.12 Using fold-recognition,16 we identified remote similarities between Oluc-19 and the well-characterized family of iLBPs. This protein family exhibits a strongly conserved structural motif, where an Arg or Lys of strand 10 hydrogen bonds with strand 1 and packs against a conserved Trp residue.17 This stabilizing interaction is only partially retained in Oluc-19, where the conserved Arg/Lys is replaced by Asn (Supplementary Figure s1). We hypothesized that a more stable variant of Oluc-19 could be created by mutating Asn to either Arg or Lys. Both substitutions produced higher enzyme stability and improved luminescence output in bacterial lysates, further substantiating the structural similarity to iLBPs. The Arg variant (Oluc-N166R) was most improved (∼50% increased stability at 37 °C and ∼3-fold higher luminescence intensity) and was used as the template for directed evolution.
Structural Optimization and Screening of Novel Coelenterazine Substrates
The luminescence expression of Oluc-N166R was optimized in E. coli in three phases. The first phase utilized a single round of random mutagenesis and screening in bacterial lysates with coelenterazine for brighter luminescence. Eight beneficial mutations (A4E, Q11R, A33K, V44I, A54F, P115E, Q124K, and Y138I) were combined to produce the variant C1A4E. When analyzed in HEK293 cell lysates, C1A4E was approximately 29,000-fold brighter than Oluc-N166R (Table 1). Western blot analysis indicated C1A4E was produced more efficiently than Oluc-N166R in cells (Supplementary Figure s2), accounting for much of the increased luminescence. The increased expression is consistent with improved enzyme stability of C1A4E at 37 °C, where the half-life of activity retention was increased 65-fold over that of Oluc-N166R (Table 2). Despite the increased stability, gel permeation chromatography revealed that the purified protein was largely aggregated (Supplementary Figure s3).
Luminescence in HEK293 Lysates
Signal and Enzyme Stability in HEK293 Lysates
The second phase of optimization focused on identification of a superior substrate and screening for further increases in luminescence. Twenty-four novel coelenterazine analogues were synthesized (Supplementary Figure s4) containing different motifs at positions 2, 6, and 8 of the imidazopyrazinone core (Figure 1). A preferred substrate would yield brighter luminescence while also having greater chemical stability and lower background autoluminescence. We anticipated that efficient catalytic utilization of a novel substrate may require corresponding modifications to the enzyme structure. Thus, together with native coelenterazine, 11 representative analogues were used to screen a random library of C1A4E mutants for brighter luminescence. Variants exhibiting brighter luminescence were then screened again with the entire panel of compounds.
Chemical structures. (a) Coelenterazine. (b) Coelenterazine imidazopyrazinone core (with numbering scheme). (c) Furimazine and presumed reaction products.
Although some of the library mutants revealed improvements specific to particular substrate analogues, many exhibited increased luminescence across multiple substrates. These mutations (Q18L, F54I, F68Y, L72Q, M75K, and I90V) were apparently generally stabilizing to the enzyme structure, and their combination further enhanced enzyme stability (∼10-fold over C1A4E at 37 °C) and luminescence expression. The substrate producing the brightest luminescence signal with this enzyme variant (∼25-fold over coelenterazine) was 2-furanylmethyl-deoxy-coelenterazine (furimazine; Figure 1). Furimazine was found to be more stable in cell culture media and produce lower autoluminescence (Supplementary Figure s5a,b) than coelenterazine or coelenterazine h (R2 = benzyl; Figure 1b).
The final phase of optimization focused on maximizing luminescence with the furimazine substrate. Furimazine was used to screen another random library for brighter luminescence. Beneficial mutations were identified (L27V, K33N, K43R, and Y68D) and combined to produce NanoLuc (Nluc). In total, 16 amino acid substitutions were identified over the wild-type Oluc-19, constituting alteration to about 10% of the amino acid sequence (Supplementary Figure s6). An amended gene sequence encoding Nluc was designed by optimizing codon usage for expression in mammalian cells (www.kazusa.or.jp/codon), removing potentially strong mRNA secondary structure (http://mfold.rna.albany.edu), and removing consensus promoter sequences, other transcription factor binding sites, and potential eukaryotic mRNA splice sites.
Enzymological and physical attributes resulting from this development process are summarized in Tables 1 and 2 relative to Fluc and Rluc. Although several variants of Fluc and Rluc have been described,18−20 we chose forms that have been routinely used as benchmarks in comparative studies. Comparisons to these reporters were done using glow-type assay formats, which are generally preferred for achieving reproducible measurements across multiple samples.
Nluc paired with furimazine produced 2.5 million-fold brighter luminescence in mammalian cells relative to Oluc-19 with coelenterazine (Table 1). Because light intensity is typically correlated inversely with signal duration, these assay characteristics generally should be considered together. The luminescence produced by Nluc decayed with a half-life >2 h, significantly longer than for C1A4E. This is also longer than in the glow-type assays used for Fluc and Rluc, yet Nluc produced 75- and 89-fold more luminescence in mammalian cells, respectively.
Light intensity produced by Rluc was affected only modestly by increasing the substrate concentrations to 50 μM (Km = 15 μM for both coelenterazine and coelenterazine h). Increased coelenterazine raised the light intensity by 1.6-fold; however, the signal decayed twice as fast (Supplementary Figure s7a,b). Coelenterazine h was less efficient, raising light intensity by 1.3-fold but with similarly rapid signal decay (Supplementary Figure s7a,b).
The dramatic improvement in light output for Nluc was due in part to the change of substrate, as Nluc, C1A4E, and Oluc-N166R all produced more luminescence with furimazine compared to coelenterazine. Nluc in particular was ∼30-fold brighter with furimazine, although for Rluc it produced >1,000-fold less luminescence relative to that of coelenterazine. Nonetheless, the increased luminescence was gained mostly through improvements in protein stability, where Nluc showed markedly greater retention of activity in lysates following incubation at 37 °C (Table 2). Moreover, in contrast to C1A4E, purified Nluc appeared only as a monomer by gel permeation chromatography (Supplementary Figure s8).
NanoLuc Characterization and Comparison to Fluc and Rluc
The assay conditions for Nluc (see Methods for buffer composition) were optimized for high luminescence intensity, sustained signal duration, and good working stability under typical laboratory conditions. The reagent is added in equal volume directly to cells in culture medium to elicit steady-state luminescence with a half-life routinely >2 h. Upon combining furimazine with the assay buffer, the working solution at ambient temperature loses potency with a half-life of ∼2 days.
The apparent Km for purified Nluc using either furimazine or coelenterazine was ∼10 μM (Figure 2a), while the maximum luminescence (i.e., apparent Vmax) was ∼30-fold higher for furimazine than for native coelenterazine. This difference in brightness is consistent with the data from lysates of cells expressing Nluc (Table 1). Purified Nluc produced ∼150-fold more luminescence than either Fluc or Rluc on a per mole basis under these glow-type assay conditions (Figure 2b). Quantum yield alone cannot account for this increased luminescence, as the values for Rluc (0.05–0.1)21,18 and Fluc (0.4)22 are already relatively high. Accordingly, the higher light intensity of Nluc is more likely caused by increased catalytic turnover. Strong linearity was evident for all of the luciferases, although non-linearity can occur for Nluc at higher concentrations because the higher turnover rate can deplete available substrate. However, this is a minor practical impediment since it occurs near the saturation limit of common laboratory luminometers.
(a) Furimazine and coelenterazine titrations using Nluc for determining relative signal intensities and Km (n = 3). Note the left and right axes have different scales. (b) Comparison of luminescence intensity (at 10 min) for purified Nluc, Fluc, and Rluc....
The spectral profile of Nluc revealed an emission maximum of 460 nm, consistent with the reported spectrum of the wild-type luciferase.15 The spectrum is 20 nm blue-shifted relative to that of Rluc and about 20% narrower (Figure 2c). As a consequence, Nluc should be well suited for applications involving multiplexing with longer wavelength luminescence reporters,23 providing dual luciferase assays with well separated spectra to support greater composite dynamic range and sensitivity. It may be particularly beneficial to pair Nluc with a beetle luciferase having an emission maximum greater than 600 nm.24,20 Furthermore, given the widespread use of Rluc in bioluminescence resonance energy transfer (BRET), Nluc may be preferable due to its brighter luminescence and narrow spectrum. In contrast, the blue-shifted emission may hinder usage of Nluc in animal models, where shorter wavelengths do not readily penetrate mammalian tissues.
The enzymatic activity of Nluc was found to be more robust than that of Fluc when compared under a variety of environmental conditions. Nluc showed greater thermal stability, retaining activity following incubation at 55 °C for 30 min, whereas Fluc began losing activity below 30 °C (Figure 3a). Using buffers ranging from pH 5 to 9 (Figure 3b), Nluc displayed a broader optimal range between pH 7 and 9 and retained significant activity under more acidic conditions. In contrast, Fluc showed a narrower pH profile with activity dropping sharply below pH 8. Urea sensitivity (Figure 3c) was compared by treatment for 30 min followed by 20,000-fold dilution into assay buffer containing 50 μM furimazine for Nluc or into ONE-Glo for Fluc. Nluc maintained its activity following exposure up to 8 M, whereas Fluc lost activity above 2 M. Both Nluc and Fluc tolerated high concentrations of NaCl, but Nluc showed less inhibition at 10 M (Figure 3d).
Comparison between purified Nluc and Fluc for sensitivity to (a) elevated temperature (n = 4), (b) pH (n = 3), (c) urea (n = 3), and (d) NaCl (n = 3).
Performance as a Genetic Reporter
In addition to producing bright and robust luminescence, it is important that reporters be expressed without bias in their experimental hosts. By originating from a marine invertebrate and having modifications in both the enzyme and gene, Nluc is unlikely to exhibit biases unique to mammalian cells. Nonetheless, the possibility of spurious interactions was examined. Immunodetection (Figure 4a) and luminescence imaging (Figure 4b,c) both suggested that Nluc was expressed uniformly in mammalian cells, including the nuclei. Moreover, no morphological differences were evident between cells expressing Nluc and control cells (not shown). Western blots prepared from cells expressing Nluc revealed only a single band of the expected molecular weight (Supplementary Figures s2, s11). Total mass analysis by mass spectrometry (LC–MS), using purified enzyme from E. coli and mammalian cells, indicated the absence of post-translational modifications. The proteins from both sources revealed identical molecular masses, matching the calculated mass for the expected unmodified protein.
Intracellular distribution of Nluc determined by (a) confocal imaging/ICC of transient expression in U2OS cells fixed and processed with anti-Nluc IgG/Alexa488-conjugated secondary IgG (left panel = fluorescence; right panel = DIC); scale bar = 20 μm....
The intracellular lifetime of a genetic reporter can substantially shape its expression characteristics. A stable reporter can persist longer in cells and thus accumulate to greater levels, allowing greater assay sensitivity and reduced variability from aberrant fluctuations in gene expression. However, by this same consideration, a stable reporter has diminished ability to detect changes in transcriptional rate. Reduced lifetime will yield less signal but may provide better response dynamics.25 Which scenario is appropriate depends on the experimental objectives.
To achieve better response dynamics, Fluc having an appended 41-amino-acid PEST sequence is often used to shorten its intracellular lifetime.25 We evaluated this approach by making a similar fusion to Nluc (NlucP; Supplementary Figure s6) and inserted both Nluc and NlucP into expression plasmids containing tandem cAMP-response elements (CRE). These plasmids, along with analogous plasmids containing Fluc or FlucP, were introduced into mammalian cells, and gene expression was induced by titrated addition of forskolin (FSK). FSK activates adenylate cyclase, causing an increase of intracellular cAMP to stimulate the CRE sites. All four reporters yielded similar response profiles and showed identical EC50 values (Figure 5a). As expected, NlucP was dimmer than Nluc but showed a faster rate of signal increase and greater overall response (2,000-fold versus 750-fold) following pathway activation. When compared to their Fluc counterparts over a range of similar experiments, Nluc was generally brighter than Fluc (on average 80-fold), and NlucP was brighter than FlucP (on average 10-fold).
(a) Reporter induction by tandem cAMP response elements (CRE). Nluc, Fluc, NlucP, and FlucP were transiently expressed in HEK293 cells under multiple CRE linked to a minimal promoter; luminescence measured 5 h after adding varying concentrations of FSK...
It was surprising to find that the PEST sequence had a larger effect on brightness and response dynamics when combined with Nluc rather than Fluc. We estimated intracellular lifetime by adding cycloheximide to block protein synthesis and measuring the decline of residual activity in the cells (Figure 5b). Both Nluc and Fluc were stable for at least 6 h. From multiple experiments, Nluc appears to have a longer lifetime than Fluc, which is consistent with the increased physical stability of Nluc. However, quantitative assessments by this method are difficult for half-lives beyond 6 h due to the toxicity of cycloheximide. As expected, FlucP had a shorter half-life (2 h) due to degradation induced by the PEST sequence. Yet, despite the high physical stability of Nluc, the intracellular half-life of NlucP (20 min) was 6-fold shorter than for FlucP.
It can be expected, therefore, that NlucP may provide better coupling to transcriptional dynamics while still providing good assay sensitivity. The influence of the intracellular lifetime is evident by the relative response of different reporters coupled to an NFκB-response element (Figure 5c). Upon stimulation with TNFα, the least relative response was achieved by Nluc, although it produced the most luminescence. The greatest relative response was achieved by NlucP, where the signal was produced faster and with higher signal over background. Both Fluc and FlucP produced intermediate responses. The greater responsiveness of NlucP as a reporter may be particularly important when the underlying genetic response of interest is subtle (Supplementary Figure s9).
Transcriptional reporters are commonly used for the high-throughput screening (HTS) of diverse compound libraries, where Fluc in particular has gained widespread use. However, the influence of chemical compounds on reporter activity, such as by inhibition or stabilization, can lead to false hits in the screening results.26 The performance of Nluc in the context of a diverse chemical collection was assessed by screening purified enzyme against the LOPAC1280 library (Supplementary Figure s10a). Relative to a parallel screen using Fluc (Supplementary Figure s10b), the data for Nluc showed a tighter distribution with lower inhibitor potency. As a percentage of total activity, 1.2% of the library compounds inhibited Nluc by >10%, 0.5% of these inhibited by >20%, and no compounds inhibited by >30%. For Fluc, 1.9% of compounds inhibited by >10%, where 0.8% inhibited by >20% and 0.5% inhibited by >30%. We speculate that greater structural rigidity associated with the increased thermal stability of Nluc may reduce its potential to bind nonspecifically to small molecules.
The secretion of bioluminescence by the Oplophorus shrimp suggested that Nluc might retain the capacity for efficient secretion from mammalian cells. Like other secreted proteins, the Oplophorus luciferase contains multiple cysteine residues, which are thought generally to provide stability in extracellular environments through disulfide bonds.27 For instance, Gaussia luciferase is a secreted enzyme that contains 11 cysteines. These cysteines all contribute to enzymatic activity but can hinder expression of functional enzyme within the reducing environment of the cell interior.28 Although the Oplophorus luciferase contains 12 cysteines, all but one are located in the larger 35 kDa subunit.12 Nluc, derived from the smaller subunit, contains only one non-essential29 cysteine and thus has no disulfide bonds. Hence, it is suited for both intracellular and secreted reporter configurations.
Initial reports of the recombinant Oluc-19 suggested that the signal sequence may not work effectively in mammalian cells.12 Thus we evaluated the capacity for secretion by appending both the native Oluc-19 secretion peptide and the secretion signal from human IL630 to the N-terminus of Nluc. Although both peptides caused Nluc to accumulate in the media, the IL6 construct (secNluc; Supplementary Figure s6) produced a brighter signal when transiently expressed in HEK293 and Hela cells, with ∼99% of the total luminescence localized to the cell culture medium. The half-life of secreted Nluc in the culture medium at 37 °C was 4.2 ± 0.2 days for HEK293 cells (DMEM + 10% FBS) and 7.2 ± 0.3 days for CHO cells (F12 + 10% FBS). Thus, secreted reporter activity can be retained in culture conditions with negligible loss (<10%) for over 15 h. These values are comparable to what has been observed for secreted Gaussia luciferase.13
Western blot analysis indicated the presence of processed secNluc in the medium, while Nluc without a secretion sequence was detected only in cell lysates (Supplementary Figure s11). The blot also indicates a similar size for secNluc and Nluc in cell lysates, consistent with simultaneous translation and signal peptide processing of secNluc. Images of U2OS cells by immunocytochemistry show a punctate staining pattern consistent with vesicular transport through the secretory pathway31 (Supplementary Figure s12). The performance of secNluc as a transcriptional reporter was evaluated using a plasmid as before containing multiple CRE sites. Following a medium change to remove secNluc activity resulting from basal expression, cells were treated with FSK to observe both the kinetics of signal accumulation (Figure 5d) and a dose response (Supplementary Figure s13). As expected, FSK caused a steady accumulation of secNluc in the medium at levels significantly higher than cells treated with vehicle alone.
Performance as a Fusion Reporter
The enhanced luminescence and small size of Nluc make it an attractive candidate as a protein fusion tag for a variety of applications.32 Luciferase fusions have become increasingly popular for measuring regulated changes in intracellular protein lifetime via the ubiquitin/proteasome system (UPS).33,4 For instance, such fusions have been used to monitor stress response pathways, where the activity of a transcription factor is typically regulated by altering intracellular abundance.34 We applied this approach by fusing Nluc to the C-terminus of p53, which is known to degrade rapidly in unstressed cells. However, DNA damaging agents (e.g., etoposide) cause the transcription factor to accumulate within cells by decoupling this degradation. Upon treatment with etoposide we observed a dose-dependent increase in signal (15-fold response) for cells expressing the p53 fusion, while no effect was evident for cells expressing Nluc alone (Figure 6). This result indicates the potential for Nluc as a fusion tag for monitoring intracellular protein lifetimes as indicators of cellular stress.
Use of Nluc for monitoring regulated changes in p53 stability. HEK293 cells transiently expressing p53-Nluc or Nluc were treated with etoposide for 6 h (n = 5). Response was calculated by comparing treated samples to untreated controls.
We also examined the ability to monitor subcellular localization by bioluminescence imaging (BLI) using Nluc fusions to proteins with distinct static and dynamic localization patterns. We observed that a Histone H3-Nluc fusion was appropriately localized to the nucleus (Supplementary Figure s14a–c). When fused (with a secretion signal) to the N-terminus of the β2-adrenergic receptor (β2-AR), luminescence was localized to the plasma membrane, indicating proper trafficking through the secretory pathway (Supplementary Figure s14d). BLI has the potential for revealing protein dynamics in living cells without the need for repeated sample excitation, a cytotoxic artifact associated with use of fluorescent proteins. However, the limited brightness of existing luciferase enzymes has hindered the common use of BLI for real-time measurements. Using a Nluc-glucocorticoid receptor (GR) fusion, we monitored the expected translocation from the cytosol to the nucleus upon treatment with dexamethasone (Figure 7a,b). Finally, a fusion of Nluc to protein kinase C alpha (PKCα) was properly recruited to the plasma membrane following treatment with phorbol-12-myristate-13-acetate (PMA) (Figure 7c,d).
Monitoring translocation of Nluc fusion proteins using BLI. Hela cells transiently expressing Nluc-GR fusions show (a) cytosolic localization and (b) nuclear accumulation after 15 min of dexamethasone (500 nM) treatment. U2OS cells transiently expressing...
The behavior of the PKCα and GR fusion proteins was consistent with the subcellular localization dynamics of endogenous PKCα35 and GR,36 suggesting that Nluc did not significantly perturb the functionality of the fusion partner. Of significant note, the required sample exposure times in these experiments ranged from 1 to 5 s, in contrast to the 3–10 min previously reported when using an enhanced beetle luciferase adapted specifically for BLI.37 Exposure times exceeding 1 min would make video-rate analysis of rapid (<20 min) translocation events prohibitively difficult, whereas the short exposure times used for the Nluc fusions enabled continuous monitoring of both Nluc-GR (Supporting Video 1) and Nluc-PKCα (Supporting Video 2). Gaussia luciferase has been used for high frame-rate BLI of insulin secretion;38 however, its reduced intracellular activity may limit its general use for this type of application.
Bioluminescent images were acquired immediately following addition of 20 μM furimazine to the culture medium. No significant morphological changes were evident under these conditions (Figure 4b,c; Supplementary Figure s15a–f). Quantitation of cell viability following 2 h of exposure showed that furimazine was tolerated up to about 20 μM (Supplementary Figure s16a). Similar viability profiles were observed for both native coelenterazine and coelenterazine h. Measurement of luminescence intensity from cells expressing Nluc revealed that maximal signal was attained at about 10–20 μM furimazine (Supplementary Figure s16b). These results indicate that high light intensity can be achieved from Nluc in living cells with minimal perturbance to their normal physiology.
Bioluminescence has been associated primarily with quantifying genetic processes, although increasingly it is proving valuable to other aspects of cellular analysis. The potential for achieving new capabilities has motivated the search for new luminescent chemistries able to deliver greater sensitivity and adaptability to experimental programs. We recognized that the Oplophorus luciferase offered such an opportunity by the inherent luminescent efficiency of the native enzyme combined with the discovery that the small catalytic subunit was structurally related to iLBPs. Because these proteins exhibit well-behaved structural properties and are ubiquitously expressed in vertebrates and invertebrates, this protein scaffold provided a good candidate for development of a robust luminescent reporter. The reportedly broad substrate specificity of Oplophorus luciferase also afforded the opportunity to design an improved luminogenic substrate.21
To achieve this, we combined structural optimization of the small catalytic subunit using various schemes of mutagenesis with the organic synthesis of a panel of novel substrate analogues. By folding these aspects together into an integrated process, we created a small luciferase called NanoLuc (Nluc; 19 kDa) capable of producing very bright and sustained luminescence. While elevated light intensity in short bursts has been possible with other luciferases, achieving sustained luminescence at these high levels has been elusive. Sustained luminescence greatly simplifies the instrumentation and processing requirements for sample quantification, and is essential for analysis using laboratory automation.
Nluc exhibits high physical stability despite being much smaller than Fluc, showing much greater tolerance to temperature, pH, and urea. In cells, Nluc is present as a single molecular species devoid of post-translational modifications and is uniformly distributed without apparent compartmental biases. The novel substrate, furimazine, yields light intensity higher than that of native coelenterazine and is more stable with lower background autoluminescence. The combination of these characteristics positions Nluc to be generally useful as a cellular reporter, generating a highly sensitive signal with a physical constitution that is robust to environmental influences. Nluc is particularly suited for high-throughput screening, where the bright luminescence facilitates measurements in very small sample volumes and the assay is resistant to interference from library compounds.
The structure of Nluc can be readily adapted to meet different experimental needs. It may be appended with a degradation signal (e.g., PEST) to allow expression levels to change rapidly in response to transcriptional dynamics or appended with a secretion sequence to allow export into the culture medium. The stability of Nluc does not rely on having disulfide bonds, so the enzyme may be efficiently expressed either inside or outside of cells. The small size of Nluc makes it well suited as a protein fusion tag, allowing luminescence to be associated with the physiological dynamics of specific intracellular proteins. For example, changes in luminescence can be correlated to the regulated degradation associated with many transcription factors. Luminescence from Nluc can be generated within living cells, providing sufficient signal intensity for imaging the subcellular location of fusion proteins. Further investigation will determine how broad the application space is for Nluc. The properties described here are expected to enable successful use in BRET, where the enhanced brightness over Rluc should improve assay sensitivity. Nluc may also provide unique opportunities for the development of protein complementation assays, where small size, brightness, and structural stability may offer advantages over existing approaches.
Variant Enzyme Screening
Random libraries were generated by error-prone PCR (average of 2–3 mutations per clone). Library 1 (phase 1; template = Oluc-N166R) was screened (4,400 variants) with coelenterazine. Library 2 (phase 2; template = C1A4E) was screened (4,400 variants) with 11 novel coelenterazine analogues: 3840, 3841, 3842, 3857, 3880, 3881, 3886, 3887, 3889, 3897, and 3900 (Supplementary Figure s4). The 11 analogues represented substitutions at positions 2, 6, and 8 and were considered to be representative of the entire set of 24 compounds; 2,200 variants were screened with compounds 3896 and 3894 (Supplementary Figure s4). All hits (improved luminescence) were screened again with the remaining coelenterazine analogues. Library 3 (phase 3; template = C1A4E + Q18L/K33N/F54I/F68Y/L72Q/M75K/I90V) was screened in the context of a mouse Id-X-HaloTag (where X = library) using coelenterazine and furimazine (Figure 1c). Library screens were performed on a Freedom robotic workstation (Tecan) as follows: induced bacterial cultures (in 96-well microtiter plates) were lysed with a buffer containing 300 mM HEPES pH 8, 200 mM thiourea, 0.3X Passive Lysis Buffer (PLB, Promega), 0.3 mg mL–1 lysozyme, and 0.002 units of RQ1 DNase (Promega). Assay reagent containing 1 mM CDTA, 150 mM KCl, 10 mM DTT, 0.5% (v/v) Tergitol, and 20 μM substrate was then added to equal volumes of lysate. Samples were measured on a GENios Pro luminometer (Tecan). Secondary screening to confirm hits (defined as those variants producing greater luminescence compared to that of the parental clone) and to test combination sequences was completed using a similar protocol but in manual fashion and in triplicate.
NanoLuc Assay buffer
The buffer for Nluc reactions consisted of 100 mM MES pH 6.0, 1 mM CDTA, 0.5% (v/v) Tergitol, 0.05% (v/v) Mazu DF 204, 150 mM KCl, 1 mM DTT, and 35 mM thiourea. Furimazine substrate was added to give a working reagent that was then added in equal volume directly to assay samples (final concentration of furimazine in the assay was commonly between 10 and 50 μM).
Complete methods and additional details can be found in the Supporting Information.
We thank M. Scurria, R. Arbit, H. Wang, L. Bernad, D. Simpson, R. Hurst, S. Saveliev, A. Niles, M. O’Brien, E. Strauss, J. Wilkinson, and T. Lubben for technical expertise and insightful discussions. We also thank G. Colwell at Gene Dynamics, LLC for help with vector constructions and J. Bujnicki at the IIMCB in Warsaw, Poland for assistance with fold-recognition analysis.
Supporting Information Available
This material is available free of charge via the Internet at http://pubs.acs.org
The authors declare no competing financial interest.
This paper was published ASAP on August 30, 2012. Additions were made to the Variant Enzyme Screening of the Methods section. The revised version was posted on October 24, 2012.
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