Directed evolution of bright mutants of an oxygen-independent flavin-binding fluorescent protein from Pseudomonas putida
© Mukherjee et al.; licensee BioMed Central Ltd. 2012
Received: 7 June 2012
Accepted: 19 October 2012
Published: 24 October 2012
Fluorescent reporter proteins have revolutionized our understanding of cellular bioprocesses by enabling live cell imaging with exquisite spatio-temporal resolution. Existing fluorescent proteins are predominantly based on the green fluorescent protein (GFP) and related analogs. However, GFP-family proteins strictly require molecular oxygen for maturation of fluorescence, which precludes their application for investigating biological processes in low-oxygen environments. A new class of oxygen-independent fluorescent reporter proteins was recently reported based on flavin-binding photosensors from Bacillus subtilis and Pseudomonas putida. However, flavin-binding fluorescent proteins show very limited brightness, which restricts their utility as biological imaging probes.
In this work, we report the discovery of bright mutants of a flavin-binding fluorescent protein from P. putida using directed evolution by site saturation mutagenesis. We discovered two mutations at a chromophore-proximal amino acid (F37S and F37T) that confer a twofold enhancement in brightness relative to the wild type fluorescent protein through improvements in quantum yield and holoprotein fraction. In addition, we observed that substitution with other aromatic amino acids at this residue (F37Y and F37W) severely diminishes fluorescence emission. Therefore, we identify F37 as a key amino acid residue in determining fluorescence.
To increase the scope and utility of flavin-binding fluorescent proteins as practical fluorescent reporters, there is a strong need for improved variants of the wild type protein. Our work reports on the application of site saturation mutagenesis to isolate brighter variants of a flavin-binding fluorescent protein, which is a first-of-its-kind approach. Overall, we anticipate that the improved variants will find pervasive use as fluorescent reporters for biological studies in low-oxygen environments.
KeywordsFlavin-binding fluorescent proteins Directed evolution Site saturation mutagenesis
Flavin-binding fluorescent protein
Green fluorescent protein
Light oxygen voltage
Sodium dodecyl sulfate polyacrylamide gel electrophoresis
Green fluorescent protein (GFP) and related analogs have been extensively engineered by directed evolution [1–4] to evolve fluorescent reporters with faster maturation times [5, 6], enhanced brightness [7–9], improved photostability , a wide range of emission wavelengths [11, 12], enhanced thermal tolerance , and improved efficiency of Förster resonance energy transfer (FRET) . However, the available palette of GFP-based fluorescent proteins is limited by a dependence on molecular oxygen, which mediates oxidation of a cyclic tripeptide chromophore that is strictly required for fluorescence [15, 16]. In this way, GFP-family reporters require oxygen for fluorescence and do not fluoresce in anaerobic environments [17–20].
Low oxygen environments are frequently encountered in a broad range of biomedical and industrial bioprocesses, including bioremediation and fermentation platforms for the production of high value reduced biomolecules (e.g., biofuels), hypoxic tissue environments that promote tumorigenesis, microbial pathogenesis and biofilm development, and in biotechnology applications based on obligate anaerobes. Consequently, there is a pressing need to develop a new class of reporter proteins that is capable of fluorescence in microaerobic and anaerobic environments. Recently, two flavin-binding photosensory proteins were isolated from P. putida and B. subtilis and were shown to fluoresce upon heterologous expression in E. coli. In a separate study, a flavin-binding photosensory protein based on a blue light photoreceptor (phototropin) from A. thaliana was isolated and used to track the development of viral lesions in tobacco mosaic virus-infected N. tabacum leaves . Importantly, flavin-binding fluorescent proteins (FbFPs) were observed to efficiently fluoresce when expressed in anaerobically cultured E. coli cells under identical conditions in which the GFP-family yellow fluorescent protein (YFP) was rendered nonfluorescent . FbFPs comprise a conserved light, oxygen, or voltage-sensing (LOV) domain core, which is widely employed by plant phototropins (blue light receptors) for phototaxis, stomatal opening of guard cells, chloroplast translocation, and a variety of light-driven regulatory responses [23–25]. FbFPs have a characteristic Per Arnt Sim (PAS) fold and employ a noncovalently bound flavin mononucleotide (FMN) cofactor as the light-sensing moiety . Flavin-binding photosensors have been implicated in mediating diverse functions in prokaryotes, including regulation of stress response and virulence and mediating cell adhesion [27, 28].
FbFPs have been utilized in only a limited set of biological studies such as plasmid conjugation in E. coli. Codon-optimized variants of the reporters have been shown to fluoresce when heterologously expressed in Saccharomyces cerevisiae, Candida albicans, Bacteroides fragilis, and Porphyromonas gingivalis in anaerobic conditions [36–38]. Under growth conditions that favor rapid bacterial growth, FbFPs exhibited superior performance compared to YFP as reporters of gene expression . In this work, fluctuations in oxygen tension occurred in the growth medium as cells transitioned from an actively respiring growth phase to the stationary phase, thereby resulting in time-dependent fluorescence of YFP due to fluctuations in oxygen, whereas the FbFPs were observed to be insensitive to variable oxygen tension. In a recent study, translational fusions between O2-independent FbFP and O2-sensitive YFP were constructed to generate FRET-based biosensors for real-time monitoring of cellular oxygen concentrations in E. coli.
FbFPs are promising candidates to serve as a new class of fluorescent reporters in low-oxygen conditions. In addition to oxygen-independent fluorescence emission, FbFPs exhibit a relatively small size (≈150 amino acids), which is a desirable feature for generating fluorescently labeled fusion proteins with small imprints. However, versatile application of FbFPs as robust imaging probes is currently hindered by their limited brightness. Indeed, fluorescence emission from FbFPs is weak unless FbFPs are expressed at elevated levels using strong promoters [35–40]. Based on the considerable promise of FbFPs as a new class of fluorescent reporter proteins, we aimed to evolve an FbFP for enhanced brightness and improved spectral properties. In this work, we evolved the FbFP isolated from P. putida, which has been shown to express well in both E. coli and the obligate anaerobe Rhodobacter capsulatus. We isolated two mutants involving an FMN-proximal amino acid (F37S and F37T) that confer a twofold enhancement in brightness of fluorescence emission relative to the wild type protein. Based on biochemical characterization, we conjecture that the F37S and F37T mutations improve brightness by relieving fluorescence quenching stacking interactions in the wild type protein, thereby increasing quantum yield, and by enriching the fraction of FMN-bound fluorescent holoprotein in the evolved mutants. Overall, we anticipate that these improved spectral variants will be valuable for investigating biological processes in low-oxygen conditions.
Results and discussion
Homology modeling and design of site saturation mutagenesis
FbFP fluorescence is mediated by a buried FMN chromophore and the fluorescence emission of FMN is known to be sensitive to its microenvironment . Therefore, we hypothesized that mutations within the chromophore-binding cavity could be used to enhance the fluorescence of wild type FbFPs by affecting the photophysical interactions between FMN and its neighboring amino acids. Therefore, we evolved FbFP using saturation mutagenesis of chromophore-proximal amino acids. Site saturation mutagenesis enables the incorporation all possible amino acid substitutions at selected target sites . Using this approach, degenerate oligonucleotides are used to engineer amino acid substitutions into proteins without the need for rigorously defined structure-function relationships. Previously, site saturation mutagenesis has been used to engineer key improvements in the fucose hydrolyzing activity and thermostability of an E. coli beta-galactosidase and a B. subtilis lipase respectively, by mutating amino acids in the active site [43–45]. In recent work, saturation mutagenesis of amino acids proximal to the pyrrole chromophore of a bacterial phytochrome was employed by the Tsien lab to evolve an infrared emitting fluorescent protein .
Oligonucleotides used in this work
Site saturation mutagenesis
Saturation mutagenesis at the nine amino acid targets was performed as described in Materials and Methods. For each target mutant, we screened up to 180 isolated colonies using fluorescence spectrophotometry, which corresponds to approximately sixfold coverage of the mutation space (4 × 4 × 2 = 32 possible mutations with NNK degenerate codon) and was deemed to be a statistically significant sample size by the CASTER program . CASTER is an open source worksheet that enables estimation of the number of colonies that are required to be screened in order to ensure a 95% coverage of all possible variants resulting from site saturation mutagenesis . FbFPs are significantly less bright compared to most proteins in the GFP-family (~13% and ~4% as bright as GFP and the spectrally equivalent mTFP1, respectively ). Therefore, we determined that at least a twofold enhancement in brightness or a substantial spectral shift would be useful criteria for obtaining fluorescent reporters of practical relevance. In this way, we defined beneficial mutants as cells displaying at least a twofold enhancement in fluorescence emission and/or a 10 nm or longer shift in emission wavelength. Mutants that scored in the 96-well plate assays and shake flask-based screens were isolated and purified, and the fluorescence spectra of the purified mutant proteins were compared against purified wild type FbFP. Direct comparison of fluorescence spectra between purified protein preparations ensures that improvements in spectral properties are the direct result of mutations in the open reading frame and do not arise due to improved cellular production of the mutant proteins (e.g., enhanced emission due to improved protein production by codon adaptation at a mutated site). Only mutants satisfying the improvement criteria in all three screens (96-well plate, shake flask, and purified protein) were classified as beneficial mutants that show significant improvement relative to the wild type protein.
Classification of mutants
Isolation of enhanced brightness F37S and F37T mutants
Characterization of bright mutants F37S and F37T
In order to elucidate a biochemical and biophysical basis for the improved fluorescence observed in the F37S and F37T mutants, we characterized these beneficial mutants with respect to their oligomeric structure, quantum yield, and the fraction of FMN-bound holoprotein in solution. Fluorescence properties of the beneficial mutants F37S and F37T can be affected by: 1) changes in the oligomeric state of the mutants, 2) changes in the dissociation constant of the FMN chromophore, which can affect the levels of fluorescent holoprotein (FMN chromophore + FbFP apoprotein) for mutants relative to the wild type protein, and/or 3) altered photophysical interactions between the FMN chromophore and the amino acid at the mutation site.
Characteristics of improved FbFP F37S and F37T mutants
0.30 ± 0.01
0.24 ± 0.01
Fraction of holoprotein
0.33 ± 0.01
0.49 ± 0.01
0.45 ± 0.03
In this work, we successfully employed site saturation mutagenesis to isolate two fluorescence enhanced mutants of an anaerobic fluorescent protein from Pseudomonas putida. Although directed evolution of a plant-based flavin binding fluorescent phototropin for enhanced fluorescence in plant cells has been described previously , our study utilizes laboratory evolution to develop flavin-based anaerobic fluorescent proteins for prokaryotic expression. We identified FMN-proximal aromatic amino acids as particularly important in affecting the spectral response of FbFPs by directly affecting the quantum yield of the bound FMN cofactor as well as the strength of FMN binding. Future efforts aimed at improving the brightness of FbFPs can be directed at enhancing both the quantum yield as well as the holoprotein fraction by engineering stronger binding mutants. Although we were unable to further enhance the brightness of the F37S mutant using error-prone PCR (Additional file 3), we speculate that additional improvements in FbFP fluorescence may be achieved by employing alternative evolution strategies such as family shuffling. In future work, we are investigating these options to generate improved variants of the F37S mutant. Furthermore, aromatic amino acids in other FbFPs (B. subtilis, A. thailana) could be considered as mutagenesis “hot spots” to engineer and fine-tune FbFP fluorescence. We anticipate that further engineering of anaerobic fluorescent proteins for spectral enhancements will enable new studies of real time gene expression, protein localization, and dynamic protein interactions in anaerobic bioprocesses with broad implications in systems and synthetic biology, biotechnology, and biological engineering. In this way, we anticipate that our study will furnish an experimental and theoretical basis to enable further engineering of FbFPs, which are currently at an early stage of development.
Bacterial strains and growth media
E. coli BLR (DE3) expression strains (EMD Chemicals) were used for protein expression. E. coli DH5α cells were used for cloning and propagation of the wild type flavin-binding fluorescent protein (FbFP) gene from Pseudomonas putida. Cells were grown with vigorous shaking (200 r.p.m.) at 37°C in Lennox broth (10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl) supplemented with ampicillin at 100 μg/mL. For growth in solid phase, cells were grown on solid 1.5% agar plates containing LB medium, and plates were incubated at 37°C for 20–48 hours. Growth in high-brim 96-well plates (Axygen) was achieved using 800 μL cell culture volumes. The plates were sealed with oxygen permeable sealing mats (Axygen) to minimize evaporation and contamination. Antibiotics and salts were purchased from Sigma-Aldrich (St. Louis, MO) and were of the highest possible purity. Tryptone, agar, and yeast extract were purchased from Fisher Scientific (Pittsburgh, PA).
Cloning of FbFP gene from Pseudomonas putida
The Pseudomonas putida KT2440 gene coding for the FbFP [RefSeq ID: NP_744883.1] was synthesized by GenScript (Piscataway, NJ). The synthetic gene was cloned into the pQE80L expression vector (Qiagen) using BamHI and HindIII restriction enzymes, following standard protocols , and the construct was designated pQE80L-FbFP. DNA sequences for all primers used in this work are listed in Table 1. The pQE80L vector appends a hexahistidine tag to the N-terminus of the protein, thereby facilitating protein purification on nickel chelating columns. The expression constructs were transformed to E. coli DH5α cells by heat shock transformation at 42°C, and transformants were selected on LB plates solidified with agar at 1.5% and supplemented with ampicillin. Plasmids were isolated from E. coli DH5α transformants (Qiagen Miniprep kit) and were used to transform E. coli BLR (DE3) cells for protein expression. Restriction enzymes were purchased from New England Biolabs (Ipswich, MA), and primers were synthesized by Integrated DNA Technologies (Coralville, IA).
Directed evolution of FbFP from Pseudomonas putida
A schematic of the protocol for directed evolution is presented in Additional file 4. Mutations were engineered in wild type FbFP using site-saturation mutagenesis, followed by transformation into E. coli BLR (DE3) expression strains. Expression vectors and induction conditions were optimized for high transformation efficiencies and enhanced expression levels of the desired proteins. Specifically, we cloned the wild type FbFP gene in three different expression vectors: pQE80L (Qiagen), pET28a(+) (Novagen), and pET*28a(+), where the latter is an expression vector constructed in-house by mutating the promoter for the lac repressor in pET28a(+) to induce high levels of expression of the repressor protein . E. coli BLR cells were transformed with the aforementioned expression constructs, and the transformation efficiencies were assessed. Transformed cells were induced for FbFP expression using three different strategies: IPTG induction for 8 hours, 16 hours, and autoinduction . For these three plasmid constructs, fluorescence levels were measured and compared between induced and uninduced cells using these three different induction strategies. We observed that the pQE80L construct consistently yielded the highest transformation efficiencies as well as the largest normalized fluorescence levels in induced cells (Additional file 5). Therefore, the pQE80L vector was used to clone the mutant libraries. Screening was implemented in a 96-well format in liquid media and was performed approximately 8 to 12 hours after induction of protein expression with IPTG. Subsequent to the initial screen, improved mutants were carried forward and further assayed by fluorescence measurements in shake flask cultures and using purified protein preparations.
Site saturation mutagenesis
Site saturation mutagenesis was accomplished with the QuikChangeTM Multi Site Directed Mutagenesis kit (Agilent Technologies) using degenerate oligonucelotides that substitutes the targeted codon with an NNK triplet (N: any nucleotide, K: G or T). Briefly, 300 ng of the pQE80L-FbFP expression vector was used as a template in a 50 μL ligation-during-amplification PCR  comprising an initial denaturation at 95°C for 1 minute and 30 cycles of 95°C for 1 minute, 55°C for 1 minute, 65°C for 12 minutes and a final extension at 65°C for 15 minutes. The products of the reaction are single stranded circular plasmids harboring a degenerate NNK substitution at the desired codon. The reaction was digested with DpnI as before to eliminate plasmids bearing the native FbFP gene. Plasmids were transformed to E. coli BLR (DE3) cells by heat shock transformation at 42°C. Transformants were selected by ampicillin resistance. Primer sequences used to generate mutations for site saturation mutagenesis are listed in Table 1.
Screening mutant libraries for improved variants
Transformants from the site saturation mutagenesis library were plated on LB-ampicillin plates without IPTG. Following incubation for 20 hours at 37°C, single colonies were picked from the plates with sterile toothpicks and inoculated in 800 μL LB-ampicillin media in high-brim 96-well plates and grown for 16 hours. Cells from the overnight cultures were diluted 100-fold in fresh LB media and grown for an additional 2 hours before inducing with 1 mM IPTG for 8–12 hours. Fluorescence measurements were then conducted in optically clear round bottom 96-well plates (BrandTech Scientific, Essex, CT) in a spectrofluorometer (Cary Eclipse Fluorescence Spectrophotometer, Varian). Fluorescence emission scans spanning 470 nm to 600 nm wavelength range (1 nm resolution) were recorded from each well at two excitation wavelengths (450 nm and 500 nm). Background fluorescence was measured in uninduced cells bearing wild type pQE80L-FbFP and was subtracted from all readings. The resulting spectra were smoothed by 3rd order Savitzky Golay filtering with a frame size of 5. Selected mutants were subject to further spectrofluorometric analyses using shake flask cultures and purified protein preparations. Induction of protein expression and subsequent fluorescence measurements were conducted as previously described. Analyses of fluorescence spectra were performed using custom codes written in MATLAB version 7.10 (MathWorks) and are available on request.
Homology Modeling of the FbFP from Pseudomonas putida
A predetermined structure of the Bacillus subtilis blue light photoreceptor, YTVA [PDB: 2PR5_A] was used as a template to model the structure of the Pseudomonas putida FbFP. Homology modeling was implemented in the Swiss PDB Viewer version 4.0 . An initial raw fit was first constructed by the iterative magic fit option. Amino acids showing steric clashes with the peptide backbone were corrected by iterative simulated annealing. The structure was energy minimized using the Gromos96 force field, and the final structure was validated by measuring the root mean-squared deviation of the Cα backbone and by inspecting the Ramachandran plot. Amino acids (except glycine and proline) lying in the prohibited areas of the Ramachandran plot (specifically, Y50, D107, H13, A21, I48, V119) were omitted from further consideration. As further validation, we compared amino acids located in a chromophore-proximal 0.4 nm cavity surrounding FMN in the modeled structure with chromophore-proximal amino acids in the known crystal structures of homologous LOV domain proteins from Chlamydomonas reinhardtii [PDB: 1N9L_A] and Arabidopsis thaliana [PDB: 2Z6D_A]. In all cases, an 80-85% agreement was obtained between the identities of the amino acids in the 0.4 nm cavity surrounding FMN in the modeled and the known X-ray diffraction structures (Additional file 6).
Protein expression and purification
An isolated E. coli BLR colony expressing the pQE80L-FbFP construct was inoculated in 5 mL LB-ampicillin medium and grown for 16 hours. Cells from the overnight culture were diluted 100-fold in 500 mL fresh LB-ampicillin medium in a 2 L shake flask and induced with IPTG at 1 mM concentration in the mid-exponential phase (A600 ≈ 0.5) of cell growth. Protein expression was continued for 5 hours at 37°C. Protein purification was achieved by immobilized nickel-affinity chromatography (Qiagen Ni-NTA resin) and anion-exchange chromatography (HiTrap Q Sepharose, GE Healthcare). Briefly, following IPTG induction for 5 hours, E. coli cells were centrifuged at 5000g for 15 minutes and resuspended in Buffer A (25 mM sodium phosphate, 500 mM sodium chloride, pH 7.4). The cells were incubated with lysozyme at 1 mg/mL at 4°C for 30 minutes and lysed with an ultrasonicator. Cell debris was removed by centrifugation at 10,000g for 30 minutes, and the supernatant was supplemented with imidazole at 10 mM and incubated with 4 mL nickel-chelating resin at 4°C for 1 hour. Non-specifically bound proteins were removed by washing the column with 50 mL Buffer B (25 mM sodium phosphate, 500 mM sodium chloride, 40 mM imidazole, pH 7.4) and the protein was eluted with 25 mL Buffer C (25 mM sodium phosphate, 500 mM sodium chloride, 500 mM imidazole, pH 7.4). Protein-containing fractions were visibly fluorescent and were pooled and diluted to reduce NaCl concentration to 200 mM and loaded onto a 5 mL HiTrap Q Sepharose anion exchange column (GE Healthcare). At this NaCl concentration, the protein was strongly bound the anion-exchange column. The bound protein was washed with 25 mL buffer D (25 mM sodium phosphate, 200 mM sodium chloride, pH 7.4), and eluted in 25 mL Buffer E (25 mM sodium phosphate, 1 M sodium chloride, pH 7.4. Protein fractions were assayed by denaturing polyacrylamide gel electrophoresis (SDS PAGE) (Additional file 7) and spectrofluorometry. Purified proteins were stored at 4°C and were stable for at least a week under these conditions. IPTG was purchased from Sigma-Aldrich (St. Louis, MO). Mutant FbFP proteins were purified exactly as described for the wild type FbFP.
Determination of oligomeric state of proteins
Oligomeric states of the wild type and mutant FbFP proteins were assessed using size exclusion chromatography using a Superdex 200 column in an AKTA FPLC system (GE Healthcare). The column was calibrated with globular protein standards of known molecular mass, including bovine thyroglobulin (670 kDa), bovine γ-globulin (158 kDa), chicken ovalbumin (44 kDa), and horse myoglobin (17 kDa) (Figure 7). Purified FbFP, F37S, and F37T mutant proteins were loaded onto a Superdex 200 column in 20 mM Tris, 1M NaCl buffer at a pH of 8.0, and elution volumes corresponding to the peaks in the 280 nm absorption chromatogram were recorded. Net molecular mass was then estimated and oligomeric state determined by dividing the net mass by the calculated mass of a monomer (16.3 kDa).
Determination of quantum yield
where QY is the fluorescence quantum yield, F is the fluorescence emission intensity, and A is the absorbance. Subscripts FMN and FbFP refer to the FMN standard and the FbFP samples of unknown quantum yields, respectively. Quantum yields were estimated using highly purified proteins, which contain negligible free FMN. Furthermore, we observed that the quantum yield in wild type FbFPs from P. putida and B. subtilis cannot be increased by addition of FMN to solutions of purified proteins (data not shown). Therefore, the concentration of free FMN in solutions of purified wild type and mutant proteins is assumed to be negligible.
Calculation of holoprotein fraction
where C holo is the concentration of the holoprotein, ε is the molar absorption coefficient of FMN (12500 M-1cm-1), and l is the cuvette path length. The holoprotein concentration was further divided by the total protein concentration (measured by the Bradford assay) to elucidate the fraction of fluorescent holoprotein in solution. Holoprotein fractions were calculated using highly purified proteins, which contain negligible FMN in solution. Moreover, we observed that the fluorescent holoprotein fraction in wild type FbFPs from P. putida and B. subtilis cannot be increased by addition of FMN to solutions of purified protein (data not shown). Therefore, the concentration of free FMN in solutions of purified wild type and mutant proteins is assumed to be negligible.
All sequencing reactions were performed at the Core DNA Sequencing Facility at the Roy J. Carver Biotechnology Center, University of Illinois at Urbana-Champaign.
We would like to acknowledge Prof(s) Isaac K.O. Cann and Roderick I. Mackie (University of Illinois) for helpful discussions on anaerobic microbiology. We acknowledge useful contributions from undergraduate researchers, Robert Beverly and Cassandra Schneider. We thank Dr. Melikhan Tanyeri and Dr. Amit Desai for insightful discussions and critical comments on the manuscript. We thank Sandy McMasters for preparing competent DH5α cells and ampicillin-supplemented LB-agar plates. Finally, we acknowledge our anonymous reviewers for their excellent feedback.
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