Original Paper
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Acta Biochim Biophys
Sin 2008, 40: 365-374
doi:10.1111/j.1745-7270.2008.00411.x
Spectroscopic and functional
characterization of Lampyris turkestanicus luciferase: a comparative
study
Mojtaba Mortazavi1,
Saman Hosseinkhani1*, Khosro Khajeh1,
Bijan Ranjbar2, and A. Rahman Emamzadeh1
1 Department of Biochemistry, Faculty of Basic Sciences,
Tarbiat Modares University, Tehran P. O. Box 14115-175, Iran
2 Department of Biophysics, Faculty of Basic
Sciences, Tarbiat Modares University, Tehran P. O. Box 14115-175, Iran
Received: November
26, 2007
Accepted: March 14,
2008
This work was
supported by a grant from the Ministry of Mines and Industries (Iran) and the
Research Council of Tarbiat Modares University (TMU 85-04-13)
Abbreviations: ANS,
8-anilino-1-naphthalenesulfonic acid; CD, circular dichroism; FT, Fourier transform;
IR, infrared; NTA, nitrilotriacetic acid; UV, ultraviolet
*Corresponding
author: Tel, 98-21-8800-4407; Fax, 98-21-8800-9730; E-mail,
Functional
expression and spectroscopic analysis of luciferases from Lampyris turkestanicus
and Photinus pyralis were carried out. cDNA encoding L. turkestanicus
luciferase was isolated by reverse transcription-polymerase chain reaction,
cloned, and functionally expressed in Escherichia coli. The
luciferases were purified to homogeneity using Ni-nitrilotriacetic acid
Sepharose, and kinetic properties of luciferase from L. turkestanicus
were compared with that from P. pyralis. Amino acid differences
in its primary structures in relation to P. pyralis luciferase
brought about changes in the kinetic properties of the enzyme as evidenced by
substantial lowering of Km for ATP, increased light decay time, and
decreased thermostability. Luciferase from L. turkestanicus was used to
carry out Michaelis-Menten kinetics with a Km of 95.5 mM for ATP and
20 mM for
luciferin. Maximum activity was recorded at pH 8.5, so it might be a suitable
reporter for microbial screening at alkaline pH. Tryptophan fluorescence for P.
pyralis luciferase was higher than L. turkestanicus luciferase.
Substitution of some residues in L. turkestanicus luciferase
appears to change the kinetic properties by inducing a substantial tertiary
structural change, without a large effect on secondary structural elements, as
revealed by intrinsic and extrinsic fluorescence, Fourier transform infrared
spectroscopy, and near-ultraviolet circular dichroism spectra.
Keywords firefly luciferase; enzyme activity; Lampyris
turkestanicus; emission spectrum; decay rate
Many living organisms are capable of producing visible light, a phenomenon
generally known as bioluminescence. The best-known bioluminescent organism is
the North American firefly, Photinus pyralis, with flash-type
luminescence reaction [1]. The enzyme responsible for the light emission by
firefly is called luciferase, a monomeric enzyme of 62 kDa [2]. Firefly
luciferase (EC 1.13.12.7) catalyzes the oxidation of substrate luciferin in the
presence of ATP, Mg2+, and molecular oxygen
[2,3]. The product, oxyluciferin, is generated in an excited state, then decays
to the ground state with the emission of light. The color of the light emitted
from luminous beetles ranges from green (540 nm) to red (620 nm), and is solely
determined by the active site residues [4,5]. The luciferases have similar
spectra to the color in situ. Each luminous beetle emits a distinctive
flashing pattern that is recognized by the opposite sex in the same species
[6]. The flash may last a few milliseconds, as with the adult Photinus,
or may be a glow lasting several hours, as in the glow worm Lampyris
[7].
The enzyme converts chemical energy, efficiently, into light with a
quantum yield of 0.88 [8]. The sensitivity and convenience of the firefly
luciferase assay has created considerable interest in luciferase-based
biosensors, with a detection limit in the femtomole range. Light production of
firefly luciferase is one of the most sensitive analytical tools in the
ultrasensitive detection of ATP for measuring microbial contamination [9],
genetic reporter assays in molecular biology [10], detection of phosphatase
activity [11], use in DNA sequencing [12], and as a tool for monitoring in
vivo protein folding and chaperonin activity [13].
Assays based on luciferase are preferred due to its high
sensitivity, rapidity, and non-invasiveness. However, several factors limit
further application and development of this technology, including a low
turnover number, high Km for the substrate
ATP, and low stability [14]. For the practical use of luciferase like other
enzymes, both genetic engineering methods such as DNA shuffling and looking for
natural variants are important. Since the first cloning of P. pyralis
luciferase, the luciferase genes have been isolated from several species of
fireflies [15]. The cDNA encoding a glow worm luciferase from lantern mRNA of Lampyris
turkestanicus was cloned and functionally expressed in Escherichia coli
[16].The aim of this work was to characterize the luciferase from L.
turkestanicus and compare its properties with a flash-emitter luciferase
from P. pyralis. Both enzymes were purified to homogeneity; their
enzymatic and structural properties were compared. A further objective was to
determine whether the amino acid sequence or enzyme properties could account
for the characteristics of the glow worm’s glow, in contrast with the flashing
pattern of luciferase from P. pyralis.
Materials and Methods
Chemicals
Mg-ATP and D-luciferin (sodium salt), ATP, and isopropyl-b–D-thiogalactopyranoside
were purchased from Sigma (Poole, UK). Ni-nitrilotriacetic acid (NTA) Sepharose
was from Novagen (Madison, USA).
Plasmids and strains
The cDNA of L. turkestanicus luciferase was previously
cloned in pQE30 vector [16]. pET-16b (P. pyralis) was kindly provided by
Prof. Laurence Tisi (University of Cambridge, Cambridge, UK). Both
luciferase-containing vectors (pQE30-luc and pET-16b-luc) were transferred into
E. coli strains XL1-Blue and BL21, respectively. The pQE30-luc
and pET-16b-luc vectors used the T5 and T7 promoters, respectively, and encoded
six histidine residues on the amino terminus of the protein in the multiple
cloning sites that can be used for immobilized metal affinity chromatography.
Transformed colonies by pQE30-luc and pET-16b-uc were screened by X-ray films. Bacteria
containing the pQE30-luc and pET16b-luc were grown overnight at 37 ?C on
Luria-Bertani agar plates containing ampicillin. The master plates were sprayed
with 200 ml solution containing 0.2 mM D-luciferin (in 0.1 M Tris acetate, pH
7.8) for 5 min and exposed to X-ray film for 4 h. After film development, the
positive colonies (bioluminescent) were identified (they also could be observed
by eye after dark-adaptation).
Expression and purification of
recombinant luciferase
A 5 ml culture [2XYT (yeast extract, 10 g; tryptone, 16 g; NaCl, 5
g/L of medium) plus 50 mg/ml ampicillin] was inoculated with a single colony from a freshly
streaked plate of XL1-Blue. The culture was incubated at 37 ?C for 6–8 h then used to
inoculate a 200 ml fresh culture (2XYT plus 50 mg/ml ampicillin). The 200
ml culture was grown for approximately 5 h at 37 ?C (OD600=0.7). The temperature was then lowered to 22 ?C and the expression
was induced with 1 mM isopropyl-b–D-thiogalactopyranoside and 1 mM
lactose. Following overnight induction, the cells were harvested by
centrifugation (12,000 g for 15 min at 4 ?C). The cell pellets were
washed with 10 ml Tris buffer (50 mM, pH 7.8) and repelleted. Cells were
resuspended in 15 ml lysis buffer (10 mM imidazole, 50 mM NaH2PO4, 300 mM NaCl, pH 7.8)
containing 1 mM phenylmethylsulfonyl fluoride (Sigma). The cells were lysed by
ultrasonication. Sonication was carried out seven times with 10 s bursts with
the cells on ice to prevent excessive heating of the lysate. The cell lysate
was clarified by centrifugation (18,000 g for 25 min at 4 ?C). The clear
lysate was applied to the Ni-NTA Sepharose column then washed with 20 mM
imidazole in 50 mM NaH2PO4 and 300 mM NaCl (pH 7.8). The histidine-tagged L. turkestanicus
luciferase was eluted with 120–250 mM imidazole, desalted using dialysis, and placed into storage
buffer (20% glycerol). The desalted L. turkestanicus luciferase
was concentrated with a Centriprep-10 concentrator (Amicon, Beverly, USA). The
purity of the L. turkestanicus luciferase was more than 95% as
estimated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. This
process was also carried out to purify P. pyralis luciferase.
Luciferase assay
Luciferase assay
The activity levels were measured using a Sirius Single Tube
Luminometer (Berthold Detection Systems, Pforzheim, Germany) by integration of
total light emitted in 10 s. After the addition of 100 ml standard solution [1 mM
ATP, 0.5 mM D-luciferin, and 10 mM MgSO4 in 50 mM Tris-HCl buffer (pH 7.8)] to 100 ml luciferase-containing
extracts or purified luciferase at 25 ?C, total light units were measured in 10
s.
Optimum temperature
Thermal sensitivity of the luciferases from L.
turkestanicus and P. pyralis were compared. This was carried out
by incubating each luciferase at 10 ?C–45 ?C in 50 mM Tricine-NaOH buffer (pH 7.8)
containing 5% glycerol and 10 mM b-mercaptoethanol. Activity was measured at
different temperatures.
Optimum pH
The pH optimums of the luciferases were determined and compared by
preparation of a mix buffer (100 mM glycine, 100 mM succinic acid, and 50 mM
morpholinopropane sulfonic acid). The pH level of the mix buffer was set at 0.5
increments from 5.0 to 12.5. The reaction was initiated by injecting 15 ml diluted luciferase
into 170 ml mix buffer at each pH, then 15 ml substrate was immediately
added and initial activity was measured.
Thermal stability
Thermal stability of the luciferases from L. turkestanicus
and P. pyralis were compared. This was carried out by incubating
enzyme solutions in Eppendorf tubes in a circulating water bath at different
temperatures (20 ?C–45 ?C) for 5 min [in 50 mM Tricine-NaOH buffer (pH 7.8) containing
5% glycerol and 10 mM b-mercaptoethanol]. After heating at different temperatures, the
samples were cooled in ice water and remaining activities were assayed
immediately at 25 ?C.
Determination of kinetic
parameters
Luciferase activity was assayed by measuring the peak light emission
with a tube luminometer. The reaction was initiated by injecting 100 ml assay reagent
into 100 ml diluted luciferase. The assay reagent for estimation of Km and Vmax for ATP contained 10 mM MgSO4 and 1 mM luciferin plus 0.1, 0.2, 0.4, 0.8, 1.6, 2.5, 3.5, or 4.5
mM (final concentration) ATP in 50 mM Tricine-NaOH buffer (pH 7.8). The assay
reagent for luciferin Km and Vmax estimation contained 10 mM MgSO4 and 2 mM ATP plus 0.015, 0.025, 0.05, 0.1, 0.2, 0.5, 1.0, or 2.0 mM
luciferin in 50 mM Tricine-NaOH buffer (pH 7.8). The peak light emissions were
taken as a measure of the initial velocity and expressed as relative light
units per second. The data were collected and shown by Michaelis-Menten plots.
The Km and Vmax values for ATP and luciferin were calculated from Lineweaver-Burk
plots. Kinetic experiments were carried out three times and were reproducible
within 5%.
Decay rate
The time-course of the light emitted was measured with a
luminometer. The assay was carried out by addition of 50 ml substrate [1
mM luciferin, 50 mM Tris buffer, 2 mM ATP, and 10 mM MgSO4 (pH 7.8)] into a 50 ml purified luciferase solution from P.
pyralis or L. turkestanicus. The estimated mixing time was
1.0 s, then data was acquired.
Measurement of bioluminescence emission spectra
Bioluminescence emission spectra of the luciferases from P.
pyralis and L. turkestanicus were measured with an LS 50B
luminescence spectrophotometer (PerkinElmer Optoelectronics, Fremont, USA). A
volume of 2 ml substrate mixture consisting of 1 mM luciferin, 50 mM Tris
buffer (pH 7.8), 2 mM ATP, and 10 mM MgSO4 was added to 50 ml purified luciferase solution in a quartz cell. Data were collected
over the wavelength range 400–700 nm. The spectra were automatically corrected for the
photosensitivity of the equipment.
Fluorescence measurements
Tryptophan fluorescence was measured on the LS 50B luminescence
spectrophotometer. The excitation wavelength was set at 295 nm and the emission
spectra were obtained. Extrinsic fluorescence studies were carried out as
previously described [17] using 8-anilino-1-naphthalenesulfonic acid (ANS) as a
fluorescence probe. Measurements were taken on the same spectrofluorometer as
used for intrinsic fluorescence studies. All experiments were carried out at 25
?C with ANS and protein concentrations of 30 mM and 1 mM, respectively,
in 50 mM phosphate buffer. An excitation wavelength of 350 nm was used.
Infrared (IR) spectroscopy
Fourier transforms (FT)-IR spectra were recorded on a Nexus 870
FT-IR spectrophotometer (Thermo Fisher Scientific, Waltham, USA) with a
deuterated triglycine sulfate (DTGS) detector. The samples (1 mM) were placed
in a cell. Typically, 1 mM protein mixture was injected into the cell and a spectrum
recorded. The cell was then drained and the protein deposited on the crystal
was dried. Second-derivative spectra were calculated by a method reported
previously for bacterial luciferase [18].
Circular dichroism (CD)
measurements
CD spectra were recorded on a Jasco J-715 spectropolarimeter (Tokyo,
Japan) using solutions with protein concentrations of approximately 0.2 and 1.5
mg/ml for far- and near-ultraviolet (UV) regions, respectively. Results are expressed
as molar ellipticity, [q] (deg?cm2?mol–1), based on a mean amino
acid residue weight (MRW) of 115 for luciferase. The molar ellipticity was
determined as [q]=(q100MRW)/(cl), where c is the protein concentration in
mg/ml, l is the light path length in centimeters, and q is the measured
ellipticity in degrees at a wavelength q. The instrument was
calibrated with (+)-10-camphorsulfonic acid, assuming [q]291=7820 deg?cm2?mol–1, and with Jasco standard
non-hydroscopic ammonium (+)-10-camphorsulphonate, assuming [q]290.5=7910 deg?cm2?mol–1. The data were smoothed
using the fast FT noise reduction routine that allows enhancement of most noisy
spectra without distorting their peak shapes.
Calculation of essential
residues pKa
The data of crystal structure of Japanese firefly (Luciola
cruciata) luciferase were downloaded from Protein Data Bank (2D1Q; http://www.pdb.org/pdb/home/home.do).
The crystal structure of Japanese thermostable firefly luciferase was used as a
template to make a suitable model for L. turkestanicus and P.
pyralis. We used the software package MacroDox (Northrup, Cookeville, USA)
for our analysis [19]. The modeling data were then used to calculate pKa
of critical residues involved in enzyme catalysis or substrate binding.
Results
Purification of the
polyhistidine-tagged luciferases
The cDNA encoding both luciferases containing a polyhistidine tag (6?his) at the amino terminus of the protein were used to express
luciferases. Cells were lysed and the clarified cytoplasmic extracts applied on
a column. After washing with low imidazole (20 mM), unbound and weakly bound
proteins were removed. The polyhistidine-tagged luciferase was finally eluted
from the column by increasing the imidazole concentration gradient up to 250
mM. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis analysis and
Coomassie Brilliant Blue staining of the eluted fractions showed that the
polyhistidine-tagged enzymes were efficiently bound to the column and that the
corresponding fractions contained highly purified protein (more than 95%).
Determination of protein contents in each fraction was carried out using the
Bradford method. These results clearly showed that the enzymes were
expressed at relatively high levels (approximately 10 mg per liter culture),
and were efficiently purified, and that the enzymes had retained their
biological activity [22].
Decay rate
The time-course of the light emitted by both luciferases was also
measured for glow worm and firefly luciferases (Fig. 1). The decay in
light emission was different for each enzyme under the same conditions.
However, the rate of luminescence decay for P. pyralis firefly
luciferase was faster, suggesting lower stability of the intermediate structure
of the luciferase reaction (Table 1).
Bioluminescence emission
spectra
The color of the light emitted by the L. turkestanicus
luciferase was green as that of the P. pyralis luciferase. Fig.
2 compares the bioluminescence emission spectra of these luciferases in the
presence of luciferin and ATP at pH 7.8. Emission spectra of luciferases from L.
turkestanicus and P. pyralis showed a high similarity
without changing the lmax (Fig. 2) and only a minor broadening of the spectrum for P.
pyralis luciferase was observed.
Optimum temperature of
luciferase
The thermosensitivity of the P. pyralis and L.
turkestanicus luciferases was compared. The activity of both luciferases
reached a maximum at 25 ?C and started to decrease at higher temperatures (Fig.
3). When the temperature increased above the thermal transition temperature
(approximately 45 ?C), both luciferases lost almost all of their activity.
Thermostability of luciferase
The thermostability of the P. pyralis and L.
turkestanicus luciferases was also compared. The remaining activity of
luciferase from P. pyralis reached its maximum at 25 ?C, whereas
the activity of luciferase from L. turkestanicus started to decrease
at this temperature (Fig. 4). When the temperature increased above the
thermal transition temperature (approximately 45 ?C), both luciferases lost
almost all of their activity. The thermostability of these enzymes, however,
showed that luciferase from L. turkestanicus has lower
temperature stability than luciferase from P. pyralis.
pH optimums
The pH optimum of these luciferases was determined and
compared. The pH optimum for luciferase
from P. pyralis was 8.0, whereas the pH optimum for luciferase
from L. turkestanicus showed a maximum at 8.5 (Fig. 5).
Therefore, comparative study of these enzymes confirmed a higher optimum pH for
L. turkestanicus when compared to P. pyralis.
Kinetic properties of purified
luciferases
Table 1 compares some properties of L.
turkestanicus and P. pyralis luciferases. The bioluminescent
activity of luciferase is directly proportional to the concentration of ATP
present in the reaction mixture [9,20]. Therefore, extensive washing procedures
are necessary to eliminate ATP and other components originating from the host organism [21]. For L.
turkestanicus luciferase the Km for luciferin was 20 mM; for P.
pyralis luciferase the Km for [chshou1] luciferin was 30 mM. However, for L. turkestanicus luciferase the Km for ATP was 95.5 mM, whereas for P. pyralis luciferase it was 140 mM. The Vmax values for different concentrations of ATP were similar for both
luciferases. It should be noted that subcloning of the luciferases in
the same vector (pET-28a) did not change their kinetic properties.
Spectra of luciferases
CD spectra of both luciferases obtained in Tricine buffer (50 mM, pH
7.8) are shown in Fig. 6. As indicated in Fig. 6(A), the far-UV
CD spectra of luciferase showed a small decrease in negative ellipticity at 208
nm and 222 nm in L. turkestanicus luciferase. The near-UV CD
spectra indicated that the defined tertiary structure of the L.
turkestanicus luciferase was decreased compared to P. pyralis
luciferase [Fig. 6(B)].
Fluorescence measurements
Intrinsic and ANS fluorescence spectroscopy were used to
characterize the microenvironments of Trp and also hydrophobic clusters of
luciferases. As indicated in Fig. 7, an increase of fluorescence
intensity was observed for P. pyralis luciferase. Loss of
hydrophobic patches on the L. turkestanicus protein surface was
confirmed by fluorescence measurements using ANS (Fig. 8). ANS
fluorescence was clearly enhanced on interaction with native luciferase, as
observed earlier [22].
IR spectra
Fig. 9 compares the second-derivative
spectrum of luciferases at pH 7.8. In 1H2O, the amide I band
showed a maximum at 1652 cm–1 for both luciferases. The bands at 1658, 1666, and 1672 have been
assigned to a-structures, b-turns, and b-sheets, respectively. Changes in luciferase structure seemed to
cause a small modification in the amount of a-structures and b-turns. The band
at 1658 cm–1 had been previously
assigned to the C=O stretching vibration of the side chain amide groups. The
most obvious difference was the intensity of the band around 1658 cm–1, which was visibly a
shift to longer wave numbers in L. turkestanicus luciferase.
Discussion
The firefly luciferase gene, luc, is frequently used as a
reporter of genetic function, or for producing of recombinant luciferase [23].
To simplify the purification process, luciferases with six histidine residues
on their N-terminus were used. The luciferase bound to Ni-NTA Sepharose through
the strong chelating interaction between the His tag and Ni2+ directly from the cell lysate. A high purity recombinant luciferase
with a yield of 95% was achieved. The result reported earlier showed that the
luciferase responsible for the light emitting reaction in L.
turkestanicus, although three residues shorter, has 85% sequence
similarity with that of the firefly P. pyralis [16]. Both enzymes
have a similar C-terminus with the same key residues.
Kinetic differences
According to its kinetic properties, L. turkestanicus luciferase
could be considered a suitable indicator for ATP detection. In fact, the Km of L. turkestanicus luciferase for ATP was lower than
P. pyralis luciferase, whereas the Km towards luciferin was almost identical (Table 1). Using
ProSite [24], putative interacting residues with AMP in both luciferases have
been identified as 195-IMNSSGSTGLPK-206 [16]. The crystal structure of P.
pyralis luciferase shows that invariant residues Arg218, His245, Phe247,
Ala348, and Lys529 appear to interact with luciferin in luciferase binding site
[25]. Furthermore, the residues Asn197, Ser199, Thr343, Tyr340, Ala317, and
Gly339 in the P. pyralis luciferase sequence are believed to
interact with ATP [26]. Using homology modeling, the same residues have been
found to interact with ATP in L. turkestanicus luciferase (data
not shown). Therefore, it could be suggested that the higher affinity of L.
turkestanicus luciferase for ATP might be due to specific orientation of
these residues in the active site of the enzyme or conformational changes. The time-course of the light decay for both luciferases was also
compared (Fig. 1, Table 1). The rate of light decay for L.
turkestanicus, similar to another glow worm luciferase, was slower than P.
pyralis [27]. However, P. pyralis firefly luciferase produced
a reproducibly faster decay, suggesting faster decomposition of the
intermediate state of the luciferase reaction or more sensitivity to product
inhibition [27].
Similar light emission spectra of the L. turkestanicus
and P. pyralis luciferases (Fig. 2) also suggests
similarities in critical residues involved in the formation of substrate
intermediate structures. One of the main reasons for differences in the
luciferase bioluminescence color is the property of the emitter
microenvironment localized in the enzyme active site [28]. The variety of
bioluminescence color is also attributed to the luciferase structure [29]. However,
it seems the emitter environments of both luciferases are similar.Another intriguing result is that the shape of the bioluminescence
spectrum for L. turkestanicus luciferase under acidic conditions
was slightly changed [30]. In contrast to most firefly (Lampyridae)
luciferases, luciferase from L. turkestanicus showed a minor
shift in its emission peak under this condition, similar to that of
pH-insensitive luciferases (click beetles and railroad worms). Although firefly
luciferases have high amino acid similarity, multiple sequence alignment of L.
turkestanicus luciferase with other firefly luciferases revealed certain
amino acid changes in L. turkestanicus that are essentially
sufficient for it to have a pH-independent luminescence spectra profile. Among
them is Phe268, a conserved residue in firefly luciferases, which has been
substituted by Cys in L. turkestanicus [31].
The optimum temperature of both luciferases was similar (Fig. 3).
However, the increased temperature sensitivity of the glow worm luciferase (Fig.
4) might make it a slightly better reporter for the study of gene control
elements for high turnover mRNAs and unstable proteins in live cells [32]. The
precise mechanism responsible for the low thermostability of luciferase from L.
turkestanicus is not fully understood. The lower thermostability of L.
turkestanicus luciferase compared to P. pyralis luciferase
was similar to another glow worm luciferase, as reported earlier [27]. Another property that should be considered in the application of L.
turkestanicus luciferase is its optimum pH. As indicated in Fig. 5,
its optimum pH is 0.5 higher than P. pyralis luciferase.
Calculation of the pKa of essential residues showed that most of the
critical residues involved in binding of substrates or catalysis had a higher pKa
in L. turkestanicus compared to P. pyralis (Table
2). Therefore, it could be suggested that the higher optimum pH of L.
turkestanicus luciferase could arise from the higher pKa of some
critical-reported residues.
Structural differences Characterization of L. turkestanicus luciferases
containing an N-terminal His-tag by spectroscopic devices revealed differences
with P. pyralis luciferase. It has been shown that the His-tag
has very little effect on the structure, activity, or stability of firefly
luciferases [33]. Differences in primary structure of L.
turkestanicus compared to P. pyralis luciferase have changed
the protein conformation, as confirmed by intrinsic fluorescence. For example, L.
turkestanicus luciferase has only one Trp at position 419, whereas P.
pyralis has two Trp residues. W419 can be used as a suitable reporter in
the study of L. turkestanicus luciferase conformational changes
under different conditions. Therefore, as shown in Fig. 7, differences in
primary structure and the presence of an additional Trp caused a clear increase
in intrinsic fluorescence of P. pyralis luciferase, suggesting
that Trp(s) are located in a more hydrophobic environment. A minor red shift in
the emission spectra was also seen, suggesting that other factors might be
involved in fluorescence emission. A similar result has been observed in
changes of fluorescence intensity in native and a mutant form of bacterial
luciferase [18,34]. The hydropathy profiles of these luciferases were similar,
however, there were some regions that showed major differences. In particular,
the regions from residue 140 to 200 and from 300 to 340, which indicates the
additional hydrophobic residues in P. pyralis luciferase, showed
major differences in hydrophobic pattern when compared to L.
turkestanicus luciferase. The presence of some substitution in the L.
turkestanicus luciferase primary structure is associated with decreased
surface hydrophobicity compared to P. pyralis luciferase, as was
also confirmed by ANS fluorescence spectroscopy (Fig. 8). Such a feature
could be potentially involved with considerable conformational changes among
these luciferases. It has been reported that such changes could affect the
bioluminescence colors and other kinetic properties of luciferases [35]. The
far-UV and near-UV CD spectra of both forms were taken, and they reflected
substantial changes in the tertiary structure of the protein with small changes
in its secondary structure (Fig. 6). The type of changes observed in the
CD and IR spectra of luciferases suggests conformational differences.In conclusion, results presented in this study show that L.
turkestanicus luciferase, similar to P. pyralis luciferase,
was successfully expressed in sufficient amounts in E. coli.
Moreover, the kinetic properties of L. turkestanicus, including
higher affinity to ATP, show it can be used as a suitable reporter in molecular
biology.
Acknowledgements
We thank
Ms. Z. Fazl-Zarandi (Department of Biochemistry, Tarbiat Modares University,
Tehran, Iran) for her helpful cooperation.
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