PEDS Advance Access originally published online on October 21, 2005
Protein Engineering Design and Selection 2005 18(12):581-587; doi:10.1093/protein/gzi066
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A set of multicolored Photinus pyralis luciferase mutants for in vivo bioluminescence applications
Departments of 1Chemical Engineering and 2Bioengineering, University of Washington, Box 351750, Seattle, WA 98195-1750, USA
3 To whom correspondence should be addressed. E-mail: baneyx{at}u.washington.edu
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Abstract |
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Error-prone PCR was used to isolate Photinus pyralis luciferase mutants producing bright light in the red–orange region of the spectrum. All mutations were clustered in the ß5–
10–ß6 region of N-terminal subdomain B and appear to affect bioluminescence color by modulating the position of the Ser314–Leu319 mobile loop with respect to the putative active site. Two red variants (Q283R and S284G) and one orange mutant (S293P) contained a single substitution. Although the remaining orange variant contained two mutations, L287I mainly contributed to the color change. Emission spectra collected on whole cells at pH 7.0 revealed that while a single peak of 
max 
605 nm accounts for red light production by the Q283R and S284G variants, orange light results from the contribution of two peaks of max 560 and 600 nm. All spectra underwent a red-shift when cells were assayed under acidic conditions, whereas a blue-shift was observed at pH 8.0, indicating that the internal pH of Escherichia coli is close to the external pH shortly after imposition of acid or alkaline stress. In addition, changes in assay pH led to bimodal emission spectra, lending support to the idea that bioluminescence color is determined by the relative contribution of yellow–green and red–orange peaks. The set of multicolored luciferase mutants described here may prove useful for a variety of applications including biosensing, pH monitoring, and tissue and animal imaging.
Keywords: biosensor/firefly/imaging/luc/luminescence/lux
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Introduction |
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TopAbstractIntroductionMaterials and methodsResults and discussionConclusionsAcknowledgementsReferences |
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Bioluminescence has many uses in nature, ranging from the luring of prey to signaling for courtship and mating, to erasing the shadow of an organism, rendering it invisible from below (Visick and McFall-Ngai, 2000
; Greer and Szalay, 2002). Because light can be measured down to a few photons using photomultiplier tubes or charged-coupled devices, bio- and chemiluminescence have been exploited to detect chemicals present well below nanomolar levels in biological samples by coupling light emission with enzyme-catalyzed reactions involving oxidases, dehydrogenases and kinases. More recently, members of the luciferase family of bioluminescent proteins have found usefulness as reporters to monitor gene expression, protein localization, protein–protein interactions, analyte detection, and for the study of infection and cancer in whole animals (Doyle et al., 2004; Roda et al., 2004).
Luciferases oxidize chemically unrelated substrates in the presence of molecular oxygen to emit photons of visible light (Wilson and Hastings, 1998). The major advantage of these enzymes as reporters is the rapid (5–20 min) detection of metabolic events with high sensitivity and large dynamic range. Bacterial luciferase operons (e.g. the luxCDABE operons from Vibrio fischeri and Photorhabdus luminescens) which encode a heterodimeric luciferase (LuxAB) and all components necessary for light generation have been used in numerous biotechnology, environmental and genomic applications (Chatterjee and Meighen, 1995; Rhodius et al., 2002). Eukaryotic luciferases (e.g. the Photinus pyralis luc gene product) have the disadvantage of requiring exogenous addition of a luciferin substrate. However, they exhibit increased light production and higher efficiency than bacterial luciferases, exert a lower energy cost and metabolic burden on the host cell (Wilson and Hastings, 1998), and are widely used for in vivo luminescence monitoring and for a variety of medical and pharmaceutical screens and applications (Doyle et al., 2004; Roda et al., 2004).
Luciferase from the North American firefly P.pyralis (EC 1.13.12.7
[EC]
) is a monooxygenase that catalyzes the ATP-dependent conversion of firefly luciferin into a luciferyl-adenylate, which is oxidized to electronically excited oxyluciferin in a multistep reaction (Ugarova, 1989). Relaxation to the ground state results in the production of yellow–green light (max 560 nm) with a remarkable quantum yield of 0.9 (Seliger and McElroy, 1960; De Wet et al., 1985). The 61 kDa enzyme consists of a large N-terminal domain (residues 1–436) tethered to a smaller (residues 440–550) C-terminal domain via a flexible linker in an ‘anvil and hammer’ arrangement (Figure 1A) (Conti et al., 1996). Formation of the active site likely requires interactions between the two structural domains and 15 residues have been proposed to be involved in the architecture of the luciferyl-adenylate binding pocket (dark space-filled residues in Figure 1A) (Branchini et al., 1998, 2003).
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The availability of bright, red-shifted variants of P.pyralis luciferase is desirable for multiplex analysis in whole-cell biosensor applications and for minimizing light absorption and scattering by tissues in whole-animal bioluminescence imaging studies. Although wavelength-shifted P.pyralis luciferase mutants have been obtained by site-directed mutagenesis of putative active site residues, most of them exhibit reduced specific activity relative to the wild-type enzyme (Branchini et al., 1998
, 2001, 2003). Red-shifted mutants of the Japanese firefly (Luciola cruciata) luciferase have also been isolated following hydroxylamine mutagenesis but their suitability as reporters was not investigated (Kajiyama and Nakano, 1991). In this work, we describe and characterize a set of bright, multicolored P.pyralis luciferase variants that should prove useful for biosensing, imaging and pH monitoring applications. |
Materials and methods |
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TopAbstractIntroductionMaterials and methodsResults and discussionConclusionsAcknowledgementsReferences |
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Random PCR mutagenesis and screening
Plasmid pPluc*, which encodes the P.pyralis luc gene (Bonin et al., 1994), was obtained from the American Type Culture Collection (ATCC no. 87108). The random PCR mutagenesis protocol was adapted from Shafikhani et al. (1997). Primers 5'-AGGAAACAGACCATGGAAGACGCCA-3' and 5'-GTGGTGGTGGTGCTCGGTGCGGCCGCTCTAGAA-3' were used to amplify the 1652 bp luc gene from pPluc*, maintaining the NcoI site 2 bp upstream of the initiator codon and the NotI site downstream of the stop codon. PCRs consisted of 1x PCR buffer (Promega, Madison, WI, USA), 0.2 mM dATP, 0.2 mM dGTP, 1 mM dCTP, 1 mM dTTP, 1.25 mM MgCl2, 0.24 mM MnCl2, 0.1 µM of each primer, 1 µM (500 ng) of pPluc* template and 5 U of Taq DNA polymerase (Promega) in a 100 µl final volume. The thermocycler was programmed for 30 cycles of 40 s at 90°C, 40 s at 50°C and 2 min at 72°C. PCR products were digested overnight with NcoI and NotI, and were gel purified.
Plasmid pTac24 was constructed by ligating a 482 bp SphI–HindIII fragment from pKK233-2 (Amersham Pharmacia Biotech) into the same sites of pET24a (Novagen, Madison, WI, USA), thereby exchanging the T7 promoter for a trc promoter (Chow and Baneyx, 2005). This plasmid was digested with NcoI and NotI and the large DNA fragment was used as a backbone for ligation reactions with PCR-amplified products (typically 2 ng of backbone for 7 ng of amplified DNA in 10 µl final volume). Escherichia coli Top10 cells (Invitrogen, Carlsbad, CA, USA) were made electrocompetent and 50 µl were transformed with 3 µl of the ligation mixture. Aliquots (50 µl) were spread on Luria–Bertani (LB) agar plates supplemented with 50 µg/ml neomycin. After overnight growth at 37°C, colonies were transferred onto nitrocellulose filters and luciferase synthesis was induced by spotting 1 mM IPTG on individual clones. After 1.5 h of incubation at 37°C, the filters were soaked with 1 mM D-luciferin (Biotium, Hollywood, CA, USA) in 100 mM sodium citrate buffer (pH = 5.0) for 15 min and colonies were examined in the dark for a change in bioluminescence color. Color-shifted mutants were streaked twice on LB–neomycin plates and verified for altered luminescence before DNA sequencing of the full luc gene. Primers used for sequencing were 5'-CGTACCATGGAAGACGCCAAAAACATAAAG-3', which hybridizes at the start of luc; 5'-GATACTGCGATTTTAAGTGTTGTTCC-3', which hybridizes 700 bp downstream of the start of luc; 5'-CGGGCGTGGCAGGTCTTCCCGACGAT-3', which hybridizes 1400 bp downstream of the start of luc; and the reverse primer 5'-CGTATCTCTTCATAGCCTTATGCAG-3', which hybridizes 100 bp downstream of the start of luc. Approximately 10 000 colonies were screened. Plasmids encoding luciferases emitting orange and red light were named pTucO1, pTucO2, pTucR1 and pTucR2 (Table I).
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Site-directed mutagenesis
The wild-type luc gene isolated from Ppluc* by NcoI/NotI digestion was subcloned into the same sites of pTac24 to yield pTuc. Site-directed mutagenesis was conducted on this plasmid using the QuikChange system (Stratagene, La Jolla, CA, USA) and primer pairs 5'-TCTAATTTACGCGAAATTGCTTCTGGGG-3' and 5'-CCCCAGAAGCAATTTCGCGCTAAATTAGA-3' for the H310R mutation, and 5'-CAAAGTGCGTTGATTGTACCAACCC-3' and 5'-GGGTTGGTACAATCAACGCACTTTG-3' for the L287I mutation. The presence of the correct mutations was verified by sequencing. The plasmid encoding the L287I mutation (C859A, A861T) was named pTucO2N and that containing the H310R mutation (A929G) was named pTucH310R. The L287I mutation was also introduced by site-directed mutagenesis into plasmid pTucO1 to combine the S293P and L287I substitutions. The resulting plasmid was named pTucGS.
Fluorescence spectroscopy
Shake flasks (125 ml) containing 25 ml of LB and 50 µg/ml neomycin were inoculated at a 1:50 dilution using overnight cultures of E.coli Top10 [F' – endA1 recA1 hsdR17 (rK– mK+) supE44 thi1 gyrA96 relA1 
80
lacZM15 (lacZYA-argF) U169 deoR; Invitrogen] harboring pTuc, pTucO1, pTucO2, pTucR1, pTucR2, pTucO2N or pTucGS. Cells were grown at 30°C to A600 0.4, supplemented with 1 mM IPTG to induce luciferase expression and incubated for 1.5 h at the same temperature. Samples (1 ml) were collected and the A600 was adjusted to 0.9 using LB. Aliquots (1 ml) were centrifuged at 8000 g for 10 min, and cells were resuspended in 1 ml of 25 mM MES (pH = 5.5), 50 mM sodium phosphate (pH = 7.0) or 25 mM Tris–HCl (pH = 8.0) containing 0.5 mM D-luciferin. The suspension was immediately transferred to a 5 ml fluorescence cuvette. After 5 min incubation at room temperature, emission spectra were collected on a Hitachi F-4500 fluorescence spectrophotometer operated in the luminescence mode. Data shown correspond to the average of 10 consecutive spectra collected within 15 min of substrate addition.
Immunoblotting
Cultures were grown and induced as above. Samples (3 ml) were harvested 1.5 h post-induction and the A600 was recorded. Cells were centrifuged at 8000 g for 8 min, resuspended in 3 ml of 50 mM potassium phosphate monobasic, pH 6.5, and disrupted with a French press at 10 000 psi. Soluble fractions were clarified by centrifugation at 8000 g for 20 min. Whole-cell samples were also collected 1.5 h post-induction, recovered by centrifugation at 8000 g for 2 min, resuspended in SDS loading buffer and disrupted by boiling. Soluble and insoluble cellular fractions corresponding to identical absorbance units were prepared as described previously (Thomas and Baneyx, 1996). Aliquots were resolved on 12.5% SDS minigels and proteins were transferred onto nitrocellulose. Membranes were probed with polyclonal goat anti-luciferase antibodies (Promega) at a 1:1000 dilution followed by incubation with alkaline phosphatase-conjugated rabbit anti-goat IgG at 1:1000 dilution. Blots were developed using 5-bromo-4-chloro-3-indolyl phosphate and nitroblue tetrazolium.
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Results and discussion |
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TopAbstractIntroductionMaterials and methodsResults and discussionConclusionsAcknowledgementsReferences |
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Isolation of colored luciferase variants
In an effort to select luciferase variants displaying altered bioluminescence colors, we subjected the P.pyralis luc gene to random PCR mutagenesis as described in Materials and methods. Approximately 10 000 colonies were screened. Promising clones were identified by unaided visual examination to guarantee that light production would be sufficient for in vivo applications. Four bright variants emitting in the orange to red regions of the spectrum were selected for further analysis (Table I).
Sequencing of the mutant genes revealed that a transition leading to a single amino acid substitution was responsible for the color change in variants LucO1, LucR1 and LucR2. Variant LucO2 contained two substitutions resulting from both a transition and a transversion. All mutations mapped to the ß-sheet B subdomain of the large N-terminal portion of luciferase, which involves residues 22–70 and 236–351 and consists of six ß-strands and six -helices (Conti et al., 1996). Remarkably, all substitutions were clustered in the region encompassing ß-strand B5, helix 10 and ß-strand B6, and none was located in the predicted luciferin-binding site, which involves 13 amino acids from the B subdomain (Figure 1A) (Branchini et al., 1998). Possible explanations for this result are that mutagenesis was not carried out to saturation and/or the fact that mutations in active site residues often decrease luciferase-specific activity (Branchini et al., 1998, 1999, 2003), which could preclude visual detection of color-shifted variants. Considering that the B5–10–B6 region directly connects to the Ser314–Leu319 loop that is a mobile component of the putative active site (Franks et al., 1998), the mutations described here may indirectly alter luminescence color by modulating loop location (Figure 1B). The fact that the G315A and G316A substitutions red-shift the maximum emission wavelength (max) of P.pyralis luciferase (Branchini et al., 2003) is consistent with this interpretation.
Expression, in vivo folding and proteolytic stability
To assess the impact of the mutations on luciferase structural integrity and in vivo folding, cultures expressing wild-type and mutant proteins were grown in LB medium at 30 or 37°C. Samples collected 90 min post-induction were separated into whole-cell, soluble and insoluble fractions and subjected to western analysis with anti-luciferase antiserum. Figure 2 shows that all strains produced comparable amounts of luciferase at both temperatures and that soluble cell fractions contained two immunoreactive bands corresponding to full-length luciferase and to a 59 kDa product (lanes s). Since the truncated fragment was not present in whole-cell fractions (lanes w), it probably results from a proteolytic event during sample processing. We surmised that the culprit was OmpT, an outer membrane protease specific for paired basic residues (Sugimura and Nishihara, 1988) that is well known for cleaving intracellular proteins following cell disruption (Baneyx and Mujacic, 2004). Indeed, when plasmid pLuc was introduced into the ompT deletion strain SF100 (Baneyx and Georgiou, 1990), the 59 kDa band disappeared, while it remained present in isogenic ompT+ cells (data not shown). The existence of a degradation fragment (which is consistent with OmpT-mediated cleavage of the K510–K511 bond) provided us with an opportunity to assess how our mutations affected the structural integrity of luciferase since destabilized mutants are usually more sensitive to proteolytic attack than properly folded proteins.
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At 37°C,
50% of the total luciferase produced accumulated as inclusion bodies and there were no major differences in solubility or proteolytic stability between native and mutant forms of the protein (Figure 2). Cultivation at 30°C improved in vivo folding as only about one-third of the total luciferase aggregated under these conditions. As expected from the fact that the L287I, H310R and Q283R substitutions are conservative, LucO2 and LucR2 behaved as the wild type. This was also the case for LucR1, which contains the less conservative S284G substitution at the beginning of strand B5 (Figure 1B). More surprisingly, neither the folding nor the stability of LucO1 was significantly different from that of the authentic enzyme despite the fact that it contains a presumably disruptive S293P substitution in the center of helix 10. Overall, the above results suggest that none of the mutations that affect bioluminescence color dramatically affects the structural integrity and/or folding of luciferase. Spectral characteristics
Because the intended applications of the luciferase mutants described herein are in the biosensing and imaging fields, we collected bioluminescence emission spectra using whole-cell samples from cultures grown at 30°C. At neutral pH, cells synthesizing the LucR1 or LucR2 variants exhibited a single emission peak with max shifted to the 600–610 nm range and showed no significant peak broadening relative to authentic luciferase (Figure 3 and Table II). However, the orange color of cultures expressing the LucO1 and LucO2 mutants was due to a major red-shifted peak with max of 600 nm and a minor peak with maximum emission wavelength near that of the native protein (Figure 3 and Table II).
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Figure 3 also shows that the maximum light emission intensities of cultures producing the LucR1 and LucR2 variants were 60–80% that of cells synthesizing wild-type luciferase. This decrease in light production may be related to slight differences in expression levels or to a direct effect of the mutations on enzyme activity. For comparison, light production by the two most active red-shifted variants isolated by Baranchini and co-workers (H245A,
max = 604 nm; A348V, max = 610 nm) was 25 and 40% that of purified luciferase at pH 7.8, respectively (Branchini et al., 2003). The fact that LucO1 and LucO2 cultures only produce 30–50% of the light generated by control cells (Figure 3) strongly suggests that the orange variants exhibit a lower quantum efficiency compared with authentic luciferase. Site-directed mutagenesis
To dissect the contribution of the two substitutions in the LucO2 variant, the L287I and H310R mutations were separately introduced in the native luciferase gene by site-directed mutagenesis, yielding LucO2N and LucH310R, respectively. Inspection of bioluminescence spectra collected as above revealed that the L287I substitution was predominantly responsible for the color change despite the fact that the side chain of Leu287 projects away from the putative active site (Figures 1B and 4; Table II). Thus, just like in the case of L.cruciata luciferase (Kajiyama and Nakano, 1991), a single amino acid change is sufficient to cause a pronounced alteration in the color of the light emitted by all our variants of P.pyralis luciferase.
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We also constructed LucGS, a double mutant combining the S293P mutation of LucO1 and the L287I substitution of LucO2N. To our surprise, the spectral characteristics of the double mutant—including maximum emission intensity—were much more similar to that of native luciferase than to either LucO variant (Figure 4 and Table II). Although this may be related to the fact that the two mutations compensate for each other, the experiment underscores the fact that attempts at rational design of bioluminescence color may yield quite unexpected results.
Influence of pH on light emission
It has long been known that the firefly luciferase emission spectrum becomes red-shifted when the enzyme is assayed at low pH (DeLuca, 1976). Because the internal pH of E.coli varies following acid or alkaline shock (Zilberstein et al., 1984; Richard and Foster, 2004), we next examined how basic and acidic environments affected the spectral characteristics of luciferase-producing cultures.
Results obtained with cells producing native luciferase mirrored those reported with the purified enzyme (Branchini et al., 1999). Although an increase in pH from 7 to 8 had little impact on spectral characteristics, acidification of medium led to a 50 nm red-shift in max (Figure 5 and Table II). In fact, the majority of our mutants exhibited narrow bandwith emission spectra with maximum emission wavelengths in the 610 nm range when cells were assayed at low pH. One exception was the LucGS variant, which displayed a bimodal spectrum at pH 5.5 with a minor peak of max = 556 nm about half as intense as the major red-shifted peak.
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However, when cultures synthesizing the LucR1 and LucR2 variants were assayed at pH 8.0, a new peak centered at
560 nm appeared. This peak was approximately as intense as the 590–600 nm peak that dominates under neutral conditions. Cells producing the LucO variants exhibited a similar behavior upon alkaline shock. However, in these strains, the 600 nm peak all but disappeared whereas the 560 nm peak became the main contributor to the spectrum (Figure 5 and Table II).
The fact that all mutants primarily luminesce in the red upon acid shock and in the yellow–green upon alkaline shock indicates that the internal pH of E.coli is very similar to the extracellular pH shortly after imposition of pH stress. It is also worth noting that, for all variants, including LucGS, we were able to identify pH conditions leading to a distinct bimodal emission spectrum. These results support the proposal that firefly luciferase bioluminescence color is determined by the relative contribution of short- and long-wavelength emission peaks, each of which corresponds to the preferential stabilization of a different resonance species of excited oxyluciferin that is ultimately controlled by the protein structure (Branchini et al., 2004).
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Conclusions |
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TopAbstractIntroductionMaterials and methodsResults and discussionConclusionsAcknowledgementsReferences |
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We have described a set of bright firefly luciferase mutants containing single amino substitutions that alter bioluminescence color from yellow–green to red–orange in the pH range 7–8. Because longer wavelength light more easily diffuses through tissues, LucR mutants may prove useful for tissue and whole-animal imaging. Owing to their rather wide dynamic response to changes in pH, LucO variants may find applications for non-invasing monitoring of intracellular pH. Such a system would be complementary to pH indicators based on green fluorescent protein in which changes in fluorescence emission intensity, rather than in emission wavelength, are used for in vivo pH determination (Kneen et al., 1998
). Finally, the availability of multicolored green fluorescent proteins and luciferases paves the way for the construction of complex whole-cell biosensors that will respond to a wide range of analytes and/or allow the monitoring of multiple metabolic events. |
Acknowledgements |
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TopAbstractIntroductionMaterials and methodsResults and discussionConclusionsAcknowledgementsReferences |
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E.S. gratefully acknowledges NSF for a graduate fellowship. This work was supported by the NSF award BES-0097430 and by a seed grant from the University of Washington Microscale Life Science Center.
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References |
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Received June 1, 2005; revised August 22, 2005; accepted September 12, 2005.
Edited by Ashutosh Chilkoti
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