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The photoprotein aequorin has been widely used as a bioluminescent label in immunoassays, for the determination of calcium concentrations in vivo, and as a reporter in cellular imaging. It is composed of apoaequorin (189 amino acid residues), the imidazopyrazine chromophore coelenterazine and molecular oxygen. The emission characteristics of aequorin can be changed by rational design of the protein to introduce mutations in its structure, as well as by substituting different coelenterazine analogues to yield semi-synthetic aequorins. Variants of aequorin were created by mutating residues His16, Met19, Tyr82, Trp86, Trp108, Phe113 and Tyr132. Forty-two aequorin mutants were prepared and combined with 10 different coelenterazine analogues in a search for proteins with different emission wavelengths, altered decay kinetics and improved stability. This spectral tuning strategy resulted in semi-synthetic photoprotein mutants with significantly altered bioluminescent properties.
Luminescent proteins are invaluable biochemical tools with applications in a variety of fields including gene expression analysis (Chalfie et al., 1994; Ward et al., 2000; Contag and Bachmann, 2002), drug discovery (Gonzalez and Negulescu, 1998; Kain, 1999), study of protein dynamics (DiPilato et al., 2004) and mapping signal transduction pathways (Hayes et al., 2004), to name a few. Extensive studies have been carried out to alter the spectral properties of fluorescent proteins such as green fluorescent protein (GFP) yielding mutants with different excitation/emission wavelengths (Heim and Tsien, 1996). This has further expanded the applications of GFP to multiplex and fluorescence resonance energy transfer studies (Ehrig et al., 1995; Pollok and Heim, 1999). Bioluminescent proteins are an alternative to the fluorescent proteins because of the low background signal associated with their detection yielding high sensitivity for biological sample analysis. Therefore, variants of a bioluminescent protein with different bioluminescence emission wavelengths can expand the applications of these types of proteins. The goal of this study was to produce variants of the bioluminescent photoprotein, aequorin, with different bioluminescence lifetimes and/or emission wavelengths.
Native aequorin is a stable complex of apoaequorin (189 amino acid residues), the imidazopyrazine chromophore coelenterazine and molecular oxygen (Prendergast, 2000). Aequorin contains three calcium-binding sites. Upon binding to calcium, the protein undergoes a conformational change that triggers the oxidation of the chromophore, resulting in the emission of light (λmax 469 nm). The emission wavelength largely depends on the electronic conjugation of the π-electrons of the aromatic rings of coelenterazine. Since the aromatic ring (1) can freely rotate along the σ bond that connects rings (1) and (2), the π–π interactions between these two rings changes according to the torsional angle between these planar rings (Fig. 1). The chromophore, coelenterazine, is stabilized by H-bonds, π–π interactions and hydrophobic interactions within the active site, and the torsional angle between the rings (1) and (2) is fixed by these interactions. Here we postulate that by changing the π–π interactions, H-bonding and bulkiness between the active site of the aequorin and coelenterazine, the emission maximum for the bioluminescence of aequorin can be changed.
Different methodologies, including site-specific (Ohmiya et al., 1992; Ohmiya and Tsuji, 1993; Stepanyuk et al., 2005) and random mutagenesis (Tsuzuki et al., 2005; Tricoire et al., 2006) for spectral tuning of aequorin have been used to produce aequorins with altered bioluminescent properties such as emission wavelengths, half-lives, stabilities and sensitivity to Ca2+ ions. In addition to modifications to the protein, synthetic analogues of the coelenterazine have been used to explore the effect of different coelenterazine structures on the bioluminescence of the protein. Pairing of coelenterazine analogues with native apoaequorin has been shown to result in altered bioluminescence of the protein (Shimomura et al., 1988; Shimomura et al., 1989; Shimomura et al., 1990; Shimomura et al., 1993; Inouye et al., 1997; Hirano et al., 1998; Zheng et al., 2000; Rowe et al., 2008). To design variants of aequorin with unique properties, we combined two strategies, rational site-directed mutagenesis and incorporation of coelenterazine analogues. This strategy rendered a series of mutants with varied characteristics. Notably, we observed a 74 nm shift in the emission maxima between aequorin mutant Y82F when paired with coelenterazine i and aequorin mutant W86F when paired with coelenterazine hcp.
All restriction endonucleases, T4 DNA ligase, Luria Bertani broth, LB agar and DNA mass ladder, were purchased from Gibco-BRL (Gaithersburg, MD, USA). Bovine serum albumin (BSA) and Bradford Protein Assay were purchased from BioRad (Hercules, CA, USA). Tris(hydroxymethyl)amino methane (Tris), ethylenediaminetetraacetic acid (EDTA) sodium salt, agar, glucose, magnesium sulfate, magnesium chloride and all other reagents were from Sigma (St Louis, MO, USA). The primers were purchased from Operon Technologies Inc. (Alameda, CA, USA). QuikChange™ site-directed mutagenesis kit was obtained from Stratagene (La Jolla, CA, USA). All coelenterazine analogues were purchased from Biotium (Hayward, CA, USA). All solutions were prepared using deionized (Milli-Q Water Purification System, Millipore, Bedford, MA, USA) distilled water. All chemicals were reagent grade or better and were used as received.
The lyophilization of the solutions was performed with a Christ Alpha2-4 lyophilizer (Martin Christ Gefriertrocknungslangen GmbH, Osterode, Germany). Bioluminescent measurements were made on an Optocomp I luminometer from GEM biomedical (Carrborro, NC, USA). Half-life measurements for apoaequorin–coelenterazine i pairs were taken on a Polarstar Optima luminometer from BMG Labtech (Durham, NC, USA). Bioluminescence spectra were recorded on SpectroSystem instrument (Sciencewares Inc., Boston, MA, USA).
All molecular biology procedures were performed using standard protocols (Maniatis et al., 1982). Two expression systems were used for the production of the aequorin mutants. The first was the Bacillus system previously described (Lewis et al., 2000). The second system utilized Escherichia coli and is described below. The mutants H16A, H16F, H16I, H16G, H16C, W86S, W86C, W86F, Y82F, Y82W, Y82R, Y132C, Y132M and Y132S were expressed in E.coli and the remaining mutants were prepared in Bacillus. Site-directed mutagenesis was performed using the QuickChange site-directed mutagenesis kit from Stratagene. Primer sequences used to introduce the desired mutations are listed in Table I. Expression plasmids containing the gene of the cysteine-free mutant of apoaequorin (Lewis et al., 2000) were used as a template for the PCR. The cycling parameters for the PCR reactions were 95°C for 30 s, 55°C for 60 s and 68°C for 20 min. The plasmid DNA obtained by PCR reaction was digested with Dpn I to remove the non-mutated template DNA and used to transform E.coli XL-1 Blue. Site-directed mutagenesis was confirmed through DNA sequencing performed at the Macromolecular Center at the University of Kentucky.
A single colony of Bacillus subtilis was used to inoculate 10 ml of SP II solution [9 ml T-Base (2 g ammonium sulfate, 6 g potassium dihydrogen phosphate, 14 g potassium monohydrogen phosphate and 1 g sodium citrate in 1 l water), 100 µl 50% glucose, 100 µl 10% yeast extract, 100 µl 1% casamino acids, 100 µl 2% magnesium sulfate and 100 µl 5 mg/ml tryptophan, methionine and lysine mixture]. The cells were grown until the optical density at 600 nm reached 1.0–1.5 absorbance units. One hundred mircroliter of SP II containing 50 µl of 1 M CaCl2 was added to the culture and the cells were then incubated for 90 min at 37°C with shaking at 250 rpm followed by spinning for 5 min at 5000 rpm at room temperature. The medium was then decanted and saved, and the cell pellet was resuspended in 8 ml of the decanted medium. A volume of 2 ml of 50% glycerol was then added. One microliter aliquots of the cells were frozen in a dry ice/isopropanol bath.
Mutated plasmids isolated from E.coli XL-1 Blue cells were used to transform E.coli JM109 cells prior to transformation of B. subtilis cells. Plasmids isolated from JM109 were then used in the transformation of chemically competent B. subtilis cells as follows: a volume of 40 µl of filter sterilized fresh SPII solution was mixed with 12 µl miniprep DNA in a 15 ml disposable culture tube. The chemically competent B. subtilis cells were thawed rapidly in a 37°C water bath. The thawed cells were diluted with SP II solution in a 1:2 ratio. A volume of 300 µl of the competent cells was dispensed into each culture tube containing the plasmid DNA. The tubes were incubated at 37°C for 30 min. LB broth (500 µl) was then added into each tube and incubated at 37°C for 60 min with shaking at 250 rpm. The cell cultures were then centrifuged and 600 µl of the supernatant was discarded. The remaining supernatant was used to resuspend the cells. The cells were spread on to LB Agar plates containing 10 µg/ml kanamycin. The plates were incubated overnight in a 37°C incubator.
Bacillus was grown in 500 ml of LB broth containing 30 µg/ml kanamycin at 37°C with shaking (250 rpm) until the optical density reached OD600=1.5–2.0. The culture was centrifuged at 8000g at 4°C for 30 min to pellet the cells. The culture medium containing the secreted protein was filtered through a 0.2 µm cellulose acetate syringe filter to remove any remaining cell debris. To this solution, 1 ml of protease inhibitor cocktail (Sigma) was added. The pH of the solution was adjusted to 4.2 with glacial acetic acid to precipitate the protein. The supernatant was then allowed to stir at 4°C for 12 h. The precipitated apoaequorin was collected by centrifuging at 12 000g for 30 min at 4°C. The pellet containing aequorin was dissolved in 10 mM Tris buffer pH 8.0, containing 5 mM EDTA and 2 mM dithiothreitol (DTT). The pH of the solution was adjusted to 8.0 with NaOH and the clear solution was filtered through a 0.2 µm cellulose acetate syringe filter. The crude apoaequorin extract was then purified by perfusion chromatography using an HQH anion exchange column (4.6 mm × 100 mm). The column was pre-equilibrated with 10 mM Tris buffer pH 7.0, containing 5 mM EDTA and 2 mM DTT (Buffer A). A salt gradient from 0.0 to 1.0 M NaCl was employed to elute the protein. All apoaequorin mutants eluted between 0.10 and 0.20 M NaCl. The fractions containing the mutant protein were combined, and after addition of glucose to a final concentration of 30 mM, were lyophilized. The lyophilized mutant apoaequorin variants were then dissolved in a minimum amount of deionized water and the desired buffer exchange was achieved by dialysis. The purity of the mutant apoaequorins was verified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–PAGE). Polyacrylamide (12.5%) gels were developed by using the silver-staining method. Protein concentrations were estimated by using the Bradford Protein Assay, with BSA as the standard.
For expression in E.coli, the coding sequence for aequorin was ligated into a vector containing the lpp promoter and ompA leader sequence. Escherichia coli cells containing this plasmid were found to release apoaequorin into the culture medium. E.coli cells expressing apoaequorin were grown overnight in 1 l of Luria Bertani (LB)+ampicillin (100 µg/ml) incubated at 37°C at 250 rpm. The cells were pelleted at 12 000 g for 20 min and the supernatant was then transferred to a clean flask. The pH of the supernatant was adjusted to 4.2 by the slow dropwise addition of glacial acetic acid with stirring. The precipitated protein was then pelleted by centrifugation at 12 000 g for 30 min. The pellet was resuspended in 35 ml of 30 mM Tris/HCl, pH 7.5, 2 mM EDTA and the pH was adjusted to 7.5 by the dropwise addition of 1 N NaOH. The protein was filtered through a 0.8 µm filter and then a 0.2 µm filter before applying to a Poros HQ column (20 mm × 65 mm) which was equilibrated with Buffer C (30 mM Tris/HCl, pH 7.5, 2 mM EDTA). Apoaequorin was eluted with a gradient from 0 to 50% Buffer D (30 mM Tris/HCl, pH 7.5, 2 mM EDTA, 1 M NaCl) over 10 column volumes. Fractions were analyzed by SDS–polyacrylamide gels and by assaying for activity. The active fractions of the best purity were pooled together and ammonium sulfate was added to a 1 M final concentration. The protein was then applied to a Butyl Sepharose Fast Flow column (16 mm × 90 mm) which was equilibrated with Buffer E (20 mM Bis–Tris, pH 7.5, 2 mM EDTA, 1 M ammonium sulfate). Protein was eluted by first stepping to 80% Buffer F (20 mM Bis–Tris, pH 7.5, 2 mM EDTA) and was then with a gradient of 80 to 100% Buffer F over two column volumes. Fractions were analyzed by SDS–polyacrylamide gels and by assaying for activity. After this step aequorin was usually ≥95% pure. Purified apoaequorin was then sterilized by filtering through a 0.2 µM filter before storage.
Generation of the mutant aequorin variants from their respective apoaequorins was achieved by mixing purified protein diluted in Tris–HCl buffer, pH 8.0, containing 10 mM EDTA in a glass test tube with a two to three molar excess of a coelenterazine analogue. The mixture was briefly vortexed and placed at 4°C for 16 h.
The bioluminescence activity of the mutant aequorins was measured by placing 10 µl of the regeneration mixture described above in a glass test tube, which was then placed in an Optocomp I luminometer. A volume of 50 µl of Buffer B (100 mM CaCl2, 100 mM Tris–HCl, pH 7.6) was injected into the sample solution in order to trigger the light emission. The bioluminescence signal was collected at 0.1 s intervals over a 6 s time period.
All half-life determinations, except when apoaequorin was paired with coelenterazine i, were performed using the Optocomp I luminometer. Apoequorin was diluted with buffer A until an activity between 5000 and 50 000 Relative Light Units (RLU) was reached. A volume of 50 µl of the diluted apoaequorin mutant was then incubated with 3 µl of the appropriate coelenterazine (100 µg/ml), in an air-tight, light-free environment at room temperature for 30 min. Then, 50 µl of buffer B was added to 10 µl of this solution, and light was collected over a 6 s time period, with real time measurements acquired every 100 millisecond. Half-life determinations were obtained in duplicates, and the mean of the duplicate blank was point-by-point subtracted from the mean of the sample. Blanks contained 50 µl of buffer A and 3 µl (100 µg/ml) of the relevant coelenterazine. Data were analyzed by employing the Prism 4.0 GraphPad software. Initial increasing bioluminescent data points were excluded from the half-life calculation in order to analyze only the decay of aequorin. The bioluminescent decay curve was fit into a nonlinear regression, one-phase exponential decay equation, with all curves having an R2 value greater than 0.90. The half-life was determined by fitting the data to a first-order kinetics equation.
Half-life determinations for apoaequorin paired with coelenterazine i were monitored on a Polarstar luminometer, in triplicate. This method was used because the Optocomp I luminometer had a maximum sampling time of 10 s using 0.1 s intervals, which was not sufficient for coelenterazine i. The mean of the sample was point by point corrected via subtraction of the mean of the blank. The blank contained 175 µl of buffer A incubated with 6 µl coelenterazine i then split into three wells of 50 µl each. Samples were diluted to a concentration that did not overload the photomultiplier tube detector in the luminometer, and then 175 µl aequorin were mixed with 6 µl coelenterazine and used, with 50 µl being pipetted into each well on a 96 well microtiter plate directly before analysis. A volume of 50 µl of buffer B was injected, and real time light readings were taken every 0.1 s for 100 s. The data were analyzed with the same software and equations as described above.
In order to determine the emission characteristics of the aequorin variants, a custom-designed instrument based on a Thermo-Labsystems Luminoskan Ascent luminometer capable of obtaining spectra from flash reactions of luminescent samples that emit in the 400–700 nm range was employed. The bioluminescence emission spectra of the mutant aequorins were obtained by placing 50 µl of the regeneration mixture described above in a 96-well microtiter plate, which was then placed in the SpectroSystem instrument. A volume of 100 µl of buffer B was injected into the sample solution in order to trigger the light emission. The luminescence signal was collected over a 10 s time period.
In order to study the stability of the apoaequorin mutants, the activity of a concentration of 1.0 × 10−7 M apoaequorin solution was monitored throughout a 6-month period. During this time, the solutions containing the apoaequorin mutants (1 × 10−5 M) were kept at 4°C in the Tris–HCl buffer, pH 8.0, containing 2 mM EDTA. At predetermined time periods, 100 µl of an aliquot were incubated with a molar excess of coelenterazine at room temperature for 30 min. A volume of 100 µl of Buffer B was injected into the sample solution in order to trigger the light emission.
The X-ray crystal structure of aequorin reveals a 600 Å3 hydrophobic core in which coelenterazine resides (Head et al., 2000). There are 21 hydrophobic residues in this core that stabilize the chromophore. Three sets of tyrosine, histidine and tryptophan residues form three triads that interact with the chromophore through hydrogen bonding and π–π interactions (Head et al., 2000). In our study, variants of aequorin were created by mutating residues His16, Met19, Tyr82, Trp86, Trp108, Phe113 and Tyr132. These residues were selected after careful examination of the active site of aequorin crystal structure and studying the previous works performed by other researchers. All of these residues are within the hydrophobic chromophore-binding pocket, and form H-bonds with the chromophore. The mutations were designed to alter π–π interactions, H-bonding and bulkiness. The gene of a cysteine-free mutant of aequorin previously constructed was employed as the template (Kurose et al., 1989; Lewis et al., 2000). The cysteine-free mutant was selected as the reference because in our previous work, it has been shown to have bioluminescence emission with higher intensity than native aequorin (Lewis et al., 2000). The properties of the mutants prepared in this study are summarized in Table I.
It has been demonstrated that the bioluminescence emission of aequorin is strongly dependent on the stability of coelenterazine within the pocket, which may explain why out of the 42 mutants, only seven of them showed activity greater than 20%.
Ring (1) of coelenterazine (Fig. 1) is fixed in its place with hydrogen bonds formed by the HIS16-TYR82-TRP86 catalytic triad. Any mutations involving these residues can result in emission shifts, as long as the changes do not destabilize the coelenterazine molecule to such extent that the activity is lost. This is best illustrated when TYR82 is mutated to phenylalanine producing a mutant with an emission peak at 508 nm, a shift of 37 nm from the peak of mutant S. The phenylalanine residue is very similar to tyrosine in size and the only difference is the OH bond. This OH bond, however, is one of the main hydrogen bonds that fixes the ring (1) of coelenterazine and responsible for the emission shift. In a previous study, a different research group (Stepanyuk et al., 2005) reported a shift of close to 45 nm. This difference is probably due to the different types of apoaequorins used in the study. They employed native aequorin whereas in our study we employed a cysteine-free aequorin mutant (mutant S) as the protein template for all of our mutagenesis studies.
The half-lives of the active mutants paired with native coelenterazine ranges from 0.58 to 3.72 s. Most of the mutants’ half-life is comparable with that of mutant S, and only 4 of the 11 mutants show a significant change. These mutants involve residues Tyr82 and Trp86. It is well known that these mutants are responsible in stabilizing the coelenterazine ring (1) and, thus, it can be inferred that they are responsible for the rate of bioluminescence emission.
The chromophore of aequorin is synthetic unlike the case of GFP in which the chromophore is internally formed. This allows the use of different analogues of the chromophore with apoaequorin to explore the effect of different coelenterazine structures on the bioluminescence of the protein. We selected apoaequorin mutants with luminescence activity and paired them with different chromophores. Pairing of coelenterazine analogues with native apoaequorin has been shown to result in altering the bioluminescence of the protein (Rowe et al., 2008). We observed that aequorin mutant Y82F paired with coelenterazine i resulted in a bioluminescence emission maximum of 519 nm (Table II), and that the aequorin mutant W86F when paired with coelenterazine hcp resulted in an emission maximum of 445 nm, yielding photoproteins with bioluminescence emission peaks easily separated by 74 nm (Fig. 2).
When we look at the emission wavelengths of various aequorin mutants, we observed that the smaller hydrophobic side chain on the coelenterazine is responsible for the higher energy blue-shift. It has been known that tight hydrophobic packing of the polypeptide core of a protein can increase its stability (Eriksson et al., 1993; Vieille and Zeikus, 2001). By decreasing the size of the hydrophobic side chain of the aequorin, the effectiveness of hydrophobic packing is reduced. This results in the slight destabilization of coelenterazine within the active site resulting in a rotational relaxation around the bond connecting the two ring systems (1) and (2). Since the planarity of the ring systems (1) and (2) is responsible for the shift in wavelength, this rotational relaxation is responsible for the shift in the emission maxima that is observed in the mutants with coelenterazines that contain smaller hydrophobic side chain.
To further characterize the bioluminescence properties of the aequorin mutants, we determined their bioluminescence half-life. The apoaequorin mutants paired with coelenterazine i showed notably longer half-life values, ranging between 11.8 and 50.1 s (Table III). The observed red-shift in the bioluminescence spectra and longer half-life when apoaequorin is paired with coelenterazine i can be explained on the basis of internal heavy atom effects (Nijegorodov and Mabbs, 2001). The presence of iodine can increase the probability of an excited state singlet to an excited state triplet transition. Forbidden triplet to singlet transition leads to increased residence time in the excited states resulting in longer half-life and shift in bioluminescence spectra. Even though these mutants have lower peak intensities, they would be suitable for imaging applications.
When we observe the data obtained from these mutations, we can see that by using two different mutants coupled with two different coelenterazine analogues—Y82F paired with coelenterazine i and F113W paired with coelenterazine hcp—we can achieve a wavelength difference of 73 nm. This difference is sufficient enough for us to detect two different analytes simultaneously in the same sample simply by measuring at two different wavelengths. Moreover, we can also use another pair of aequorin mutants with similar emission wavelengths with different decay kinetics such as cysteine-free mutant paired with native coelenterazine and W86F paired with coelenterazine i- to simultaneously detect two different analytes by integrating the emitted photons after 5 s followed by deconvolution of the contributions of photons emitted from the mutant with longer half-life. Therefore, when we combine this time-resolved approach with the spatial resolution, it is possible that we can detect up to four different analytes within a given sample (Fig. 3). Thus, given a wide-variety of aequorin mutants with sufficiently different wavelengths and half-lives, it is possible to achieve a true multi-analyte detection tool. Therefore, there is still a need for a larger variety of aequorins with broader spectral characteristics.
Finally, to determine the long-term stability, a stock solution of apoaequorin was stored at 4°C. At predetermined time intervals, an aliquot of this stock solution is charged with excess coelenterazine, and serially diluted to 1 × 10−7 and 1 × 10−8 M. The data show a similar trend in all aequorin variants, the activities decrease gradually as the time passes. The original cysteine-free mutant retains ~80% of its activity after 131 days and it is the most stable enzyme within the time frame. At the end of the 131 days, the activities of the mutants range from 5.5% (Y113H) to 64.9% (W86F). These results suggest that the mutations made within the active site of the aequorin can be detrimental to the long-term stability of aequorin (Fig. 4).
In conclusion, we have demonstrated the design and preparation of aequorin variants that have shifted-bioluminescence emission spectra, and longer bioluminescence half-life. Also, this study allowed us to investigate the relationship between the activities of aequorin with respect to the mutations in its active site. In order to achieve more subtle stability changes, one can propose to make mutations on the secondary layer of aequorin. This layer is defined as the amino acids in the immediate proximity of the residues that forms the active site. These amino acid residues stabilize the active site residues. Mutations done on these residues will affect the spatial positions of the active site residues without actually changing them. These subtle positional shifts can be effective enough to actually affect the stability of coelenterazine without the detrimental effects observed.
This work was supported by National Institutes of Health (CH 467917).
E. Dikici acknowledges the Research Challenge Trust Fund of Kentucky for predoctoral fellowship. S.D. is indebted to the Office of the Vice-President for Research at the University of Kentucky for a University Research Professorship. S.D. is also thankful for a Gill Eminent Professorship. L.R. acknowledges the support by Pre-doctoral Fellowships from the National Institutes of Health and the National Science Foundation-IGERT Program at the University of Kentucky.
Edited by Michael Hecht