(Received for publication, May 21, 1997, and in revised form, June 19, 1997)
From the Department of Chemistry and Biochemistry, University of California at San Diego, La Jolla, California 92093-0601
A 25-kDa murine lysophospholipase (LysoPLA I) has
been cloned and expressed, and Ser-119 has been shown to be essential
for the enzyme activity (Wang, A., Deems, R. A., and Dennis,
E. A. (1997) J. Biol. Chem. 272, 12723-12729).
In the present study, we show that LysoPLA I represents a new member of
the serine hydrolase family with Ser-119, Asp-174, and His-208
composing the catalytic triad. The Asp-174 and His-208 are conserved
among several esterases and are demonstrated herein to be essential for
LysoPLA I activity as the mutation of either residue to Ala abolished
LysoPLA I activity, whereas the global conformation of the mutants
remained unchanged. Furthermore, the predicted secondary structure of
LysoPLA I resembles that of the /
-hydrolase fold, with Ser-119,
Asp-174, and His-208 occupying the conserved topological location of
the catalytic triad in the
/
-hydrolases. Structural modeling of
LysoPLA I also indicates that the above three residues orient in such a manner that they would comprise a charge-relay network necessary for
catalysis. In addition, the regiospecificity of LysoPLA I was studied
using 31P NMR, and the result shows that LysoPLA I has
similar LysoPLA1 and LysoPLA2 activity. This
finding suggests that LysoPLA I may play an important role in removing
lysophospholipids produced by both phospholipase A1 and
A2 in vivo.
Lysophospholipids
(LysoPL)1 are the
detergent-like intermediates in phospholipid metabolism whose in
vivo levels must be strictly regulated for proper cell function
and survival. Accumulation of LysoPL can perturb the activities of many
membrane-bound signal-transducing enzymes (1-4), distort cell membrane
integrity, and even cause cell lysis (5, 6). Increased LysoPL levels
have also been detected in a variety of disease states including lethal
dysrhythmias in myocardial ischemia and segmental demyelination of
peripheral nerves (7-11). The increased LysoPL levels are believed to
be caused by the malfunction of LysoPL-regulating enzymes including lysophospholipases (LysoPLA), phospholipases A1 and
A2 (PLA1 and PLA2), transacylases,
and acyltransferases (Scheme I).
As shown in Scheme I, lysophospholipases (LysoPLA1 and LysoPLA2) regulate LysoPL levels by further hydrolyzing the LysoPL generated by PLA1 or PLA2. Over the past few years, PLA2 has attracted much attention due to its roles in signal transduction and in the release of arachidonic acid, an important precursor for other lipid messengers such as the prostaglandins and leukotrienes (12-16). Arachidonic acid that occurs predominantly in the sn-2 position of phospholipids, however, could also be released by the sequential actions of PLA1 and LysoPLA2. Therefore, LysoPLA2 may also contribute to arachidonic acid release in vivo.
LysoPLA has been identified in a variety of cells and tissues, and recently a rat and a mouse enzyme have been sequenced, cloned, and expressed in Escherichia coli cells (17, 18). These two enzymes (both of 25 kDa molecular mass) share very high sequence homology as well as similar properties and represent the first characterized mammalian lysophospholipid-specific LysoPLA (referred to as LysoPLA I) (18). Both the mouse and the rat enzymes contain a GXSXG motif, and the serine residue in the center of the motif was shown to be essential for enzymatic activity (18). In the present work, we have used site-directed mutagenesis and structural modeling to investigate the mechanism of action of LysoPLA I and to determine if a Ser/His/Asp catalytic triad is involved in catalysis. We have also investigated the substrate regiospecificity of LysoPLA I using 31P NMR spectroscopy.
Previously, we cloned the murine LysoPLA I gene into the pLEX vector (Invitrogen) at the multiple cloning sites of NdeI and EcoRI (18). To subclone LysoPLA I into the pET28a(+) vector (Novagen), both the pET28a(+) and the pLEX/LysoPLA I vectors were digested by the same two restriction enzymes, NdeI and EcoRI, and then separated by 1% agarose gel. The bands corresponding to LysoPLA I (~700 base pairs) and pET28a(+) (~5300 base pairs) were purified from the gel using WizardTM PCR Preps (Promega), and ligated together with T4 DNA ligase (Pharmacia Biotech Inc.). The ligation product was used to transform competent E. coli NovaBlue cells (Novagen), and the resulting colonies were screened by restriction enzyme analysis of the isolated plasmids. Finally, the entire LysoPLA I coding region in the pET28a(+) vector was checked by the automated DNA sequencer (Applied Biosystems 373 from Perkin-Elmer) at the University of California at San Diego Center for AIDS Research Molecular Biology Core. It should be noted that this cloned LysoPLA I has an extra 20 amino acids at the N-terminal of the protein, the sequence of which is shown in Sequence 1.
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Three pairs of mutagenic primers
with H28A, D174A, and H208A mutations were synthesized by Life
Technologies, Inc. (H28A, 5GCGGTTATTTTCCTTGCCGGATTGGGAGATACAGGGC 3
and
5
GCCCTGTATCTCCCAATCCGGCAAGGAAAATAACCGC 3
; D174A,
5
GCCATGGAGATTGTGCCCCTTTAGTTCCCC 3
and
5
GGGGAACTAAAGGGGCACAATCTCCATGGC 3
; H208A,
5
CTATGAAGGCATGATGGCCAGCTCATGTCAGCAGG 3
and
5
CCTGCTGACATGAGCTGGCCATCATGCCTTCATAG 3
). These primers were
used to generate the mutated proteins by PCR using QuickChange
Site-directed Mutagenesis kit from Stratagene. Here, the
pET28a(+)/LysoPLA I plasmid isolated from the NovaBlue cells was used
as the template for the Pfu DNA polymerase (Stratagene). After PCR, the wild-type parent plasmid remaining in the PCR product was selectively digested by the DpnI restriction enzyme
(Stratagene), and the resultant mixture was used to transform competent
E. coli NovaBlue cells. The desired mutations were selected
by restriction analysis and DNA sequencing and were confirmed to be the
only mutation introduced in each mutant.
The pET28a(+) vector harboring either the wild-type or
the mutant LysoPLA I insert was used to transform competent E. coli BL21(DE3) (Novagen), and a single colony was inoculated in an overnight culture in LB-kanamycin (50 µg/ml) medium. This overnight culture was then diluted 40-fold into Terrific Broth-kanamycin (50 µg/ml) medium and allowed to grow at 37 °C until the
A600 nm reached 1. Then IPTG (Fisher) was added
to a final concentration of 0.4 mM, and the cells were
grown at 22 °C for another 4 to 5 h to induce foreign protein
expression. Finally, the E. coli cells were centrifuged and
the pellet was stored at 20 °C.
The same purification scheme was used to purify both the wild-type and
mutant enzymes, and all procedures were carried out at 4 °C. The
purification was started by resuspending the E. coli pellet
in lysis buffer (25 mM Tris-HCl, pH 8.0, 500 mM
NaCl, 5 mM imidazole, and 10 mM
-mercaptoethanol). Lysozyme (Sigma) was added to a final
concentration of 0.5 mg/ml, and the mixture was stirred slowly for half
an hour. The mixture was then sonicated intermittently 6 times for
10 s each and then centrifuged at 100,000 × g for
45 min. The supernatant was collected and passed through a Ni-NTA
column (Qiagen). The column was then washed with 40 ml of the lysis
buffer, and the bound proteins were eluted with elution buffer (25 mM Tris-HCl, pH 8.0, 500 mM NaCl, 250 mM imidazole, and 10 mM
-mercaptoethanol).
Generally, LysoPLA I was eluted in the first 15 ml of the elution
buffer with some minor high molecular weight contaminants. It was then
loaded onto a gel filtration column, Sephadex G-75 (2.5 × 90 cm,
Pharmacia Biotech Inc.) equilibrated in buffer A (10 mM
Tris-HCl, pH 8.0, 2 mM EDTA, and 10 mM
-mercaptoethanol). The LysoPLA I eluted from the G-75 column was
essentially free of contaminants and was used for both CD measurements
and activity assays. To remove the His·Tag at the N-terminus of the
protein, the purified protein was digested with biotinylated thrombin
(Novagen) for more than 16 h at 4 °C, and the biotinylated
thrombin was removed at the end of the digestion by
streptavidin-agarose (Novagen). The His·Tag-removed protein was used
for both 31P NMR measurements and activity assays.
Lysophospholipase
activity was measured at 40 °C in 0.1 M Tris-HCl buffer,
pH 8.0, 125 µM
1-[14C]palmitoyl-sn-glycero-3-phosphorylcholine
(1.6 µCi/µmol) (obtained from Avanti and NEN Life Science Products)
in a total volume of 0.5 ml. The assay was initiated by adding an
aliquot of enzyme solution to the substrate mixture and incubating for
the desired time. The released fatty acid was extracted by the Dole
method and then quantified by scintillation counting (19). The protein concentration of the E. coli homogenate was quantified by
the Bio-Rad protein assay using bovine serum albumin as standard, and
the purified LysoPLA I was quantified by absorbance at 280 nm using an
extinction coefficient of 0.85 (mg/ml)1
cm
1. This coefficient was calculated based on the
absorbance (20) and the numbers of Trp and Tyr residues in the LysoPLA
I sequence and was found to give essentially the same result as that
obtained by Bio-Rad protein assay. Protein purity was examined using
12% SDS-polyacrylamide gel electrophoresis using the method of
Laemmli (21), and the protein bands were visualized by staining
with Coomassie Blue.
CD
spectra were measured using a modified Cary 61 spectropolarimeter (18).
CD spectra were collected at 7 °C using a cylindrical quartz cuvette
with path length of 0.5 mm. For each protein sample (purified wild-type
or mutant LysoPLA I) and blank solution (10 mM Tris-HCl, pH
8.0, 2 mM EDTA, and 10 mM -mercaptoethanol)
10 separate spectra were collected and averaged. The final protein spectra were obtained by subtracting the blank spectra from the sample
spectra and converting the difference to mean residue ellipticity.
Conversion of 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (Avanti Polar Lipids) to 2PGPC was achieved using Rhizopus lipase (Boehringer Mannheim) following published procedures (22). The enzymatically catalyzed reaction did not proceed to completion. However, 2PGPC could be separated from the starting material by chromatography on Sephadex LH-20 (Sigma). The 2PGPC prepared in this manner gave a single spot on analytical TLC but showed two peaks on 31P NMR corresponding to a 4:1 mixture of 2PGPC:1PGPC isomers. Similarly, commercial 1PGPC contains approximately 10% of the corresponding 2PGPC isomer. These cross-contaminations have been documented and are attributed to migration of the fatty acyl chain during the preparation and purification of lysophospholipids (22).
NMR MeasurementsNMR samples were made up in a final buffer
of 100 mM Tris-HCl, pH 8.0. Appropriate amounts of 1PGPC
(Avanti Polar Lipids) and 2PGPC (synthesized above) were combined to
give a ~1:1 mixture of isomers. As lysophospholipids were stored as
chloroform solutions at 20 °C, the chloroform was removed at the
beginning of the experiment by evaporation under a stream of nitrogen.
The resulting film was dissolved in the reaction buffer by vortexing
and bath sonication to give a clear and colorless solution (14.3 mM total lysophospholipid). From this lysophospholipid
solution, 350 µl was transferred to a 5-mm NMR tube. After a
reference spectrum was obtained, 150 µl of purified LysoPLA I was
added to initiate the reaction. The starting concentrations of reagents
were 10 mM substrate and 540 µg/ml enzyme in 500 µl of
buffer. The reaction was carried out at 20 °C. In addition, an
insert containing 10 mM pyrophosphate in D2O
(Cambridge Isotopes) was used as an external standard.
31P NMR spectra were obtained on a Brucker spectrometer operating at 243 MHz. A 90° pulse with a 2-s delay and a spectral width of 2671 Hz and 8 K data points was used. Broad band proton decoupling was utilized. Spectra were obtained at varying time intervals at 20 °C. The resulting free induction decay was apodized with an exponential multiplication with line broadening of 5 Hz. Peak intensities were taken to represent the relative concentrations of the phosphorus-containing species in solution. Under the same conditions as NMR experiments, nonenzymatic catalyzed hydrolysis was negligible as determined by the Dole assay.
Structure Prediction of LysoPLA IThe LysoPLA I sequence was aligned with that of the acetylcholinesterase (23) or dielelactone hydrolase (24) based on the predicted secondary structure of LysoPLA I (18) and the known crystal structures of the above two proteins. The modeling was performed on a Silicon Graphics workstation using Modeller 3, a program2 for automated comparative modeling based on the satisfaction of spatial restraints. The resulted protein structure was visualized by the RasMol and Insight II programs.
To optimize
protein expression and to simplify large scale protein purification,
the coding region of LysoPLA I in the pLEX vector was subcloned into
pET28a(+) at restriction sites of NdeI and EcoRI,
as described under "Experimental Procedures." This expression
system provides a His·Tag at the N-terminal of LysoPLA I, which can
be easily removed by thrombin cleavage after the fusion protein is
purified by the Ni-NTA column. As shown in Fig. 1, a protein band at an apparent
molecular mass of ~29 kDa was strongly induced by 0.4 mM
IPTG in E. coli (lane 3 versus lane 2). The
lysophospholipase activity in E. coli homogenate harboring pET28a(+)/LysoPLA I was also increased more than 35-fold compared with
the control, demonstrating that such a system expressed an active
LysoPLA I at a very high level. With the purification procedures described below, more than 20 mg of pure recombinant protein can be
obtained from a liter of E. coli culture.
To purify the expressed protein, the homogenate of induced E. coli cells was centrifuged at 100,000 × g for 45 min. The LysoPLA I in the supernatant fraction was then purified by a
Ni-NTA column, which yielded highly purified LysoPLA I (>96%) in a
relatively small volume (15 ml). To remove the minor high molecular
weight contaminations and to exchange the enzyme into a low salt buffer in which it is more stable, LysoPLA I isolated from the Ni-NTA column
was further purified using a gel filtration column. The LysoPLA I thus
obtained was essentially free of contamination on the
SDS-polyacrylamide gels and possessed a specific activity of 1.09 ± 0.02 µmol/min·mg (Fig. 2). After
the removal of the His·Tag at the N terminus of the protein by
thrombin, the specific activity of LysoPLA I remained the same.
Candidates for the Catalytic Triad
Previously, the Ser-119
residue in the conserved GXSXG motif of LysoPLA I
was found to be essential for protein function (18). This suggests that
LysoPLA I may be a new member of the serine hydrolase superfamily, the
catalytic mechanism of which generally involves a catalytic triad
composed of a nucleophile (Ser), an acid (Asp/Glu), and a base (His).
While the serine residue in the catalytic triad can often easily be
identified by the conserved GXGXG motif, the
sequences around the acid and the base are generally much less
conserved. Identification of the acid and the base residues is made
even more difficult by the fact that the three catalytic residues often
occur far apart in the amino acid sequence, and the order of their
appearance in the primary sequence also varies from enzyme to enzyme
(25-27). As a result, the candidates for the acid and the base
residues are often identified by comparison with other proteins based
on either amino acid sequences or secondary/tertiary structures (13,
25, 27, 28). As shown in Fig. 3, the
predicted secondary structure of LysoPLA I resembles those of the
/
-hydrolase fold, especially in the C-terminal half of the
sequence.
/
-Hydrolases constitute a family of enzymes with
different phylogenetic origins and catalytic functions but share a
common protein structure (termed
/
-hydrolase fold) and a
conserved catalytic mechanism (the catalytic triad) for activity (26).
Both the sequence (namely a nucleophile, followed by an acid and then a
base) and the topological position (all on loops formed between a
-strand and an
-helix) of the catalytic triad are highly
conserved in the
/
-hydrolase family.
In addition, when the amino acid sequence of LysoPLA I is compared with
several other esterases or putative esterases that share more than 25%
homology to LysoPLA I, only one acid residue, Asp-174, is conserved
among all of them (Fig. 3B). More importantly, this Asp-174
occurs on a loop between -strand 7 and helix E, the site conserved
for the acid residue in the
/
-hydrolases (Fig. 3A).
Three His residues, His-28, His-170, and His-208, are also conserved
among all the listed proteins (Fig. 3). However, only His-208 has the
features of the catalytic base residue in the
/
-hydrolases; it
occurs downstream of Asp-174 and is on a loop between
-strand 8 and
helix F. The previously identified Ser-119 also occupies the conserved
site termed the "nucleophile elbow" between
-strand 5 and helix
C (Fig. 3). Therefore, LysoPLA I appears to be a new member of the
/
-hydrolases with Ser-119, Asp-174, and His-208 composing its
catalytic triad.
To verify that the Asp-174 and His-208 in LysoPLA I are indeed the components of the catalytic triad, each residue was changed to an Ala by site-directed mutagenesis. In addition, His-28 was also mutated to an Ala to examine how important it is for LysoPLA I activity. E. coli cells transformed with the vectors harboring the mutant genes expressed the mutant proteins (D174A, H208A, and H28A) at about the same level as that of the wild-type protein (Fig. 1). However, lysophospholipase activity in the E. coli homogenate expressing either the D174A or the H208A mutant was at the same level as the control, more than 35-fold lower than that of the wild-type enzyme (Fig. 1). In contrast, the E. coli homogenate expressing the H28A mutant retained more than 40% activity of the wild-type enzyme. Similarly, when the mutant proteins were purified and assayed for activity, it was found that mutation at Asp-174 and His-208 abolished the activity of these two purified proteins (Fig. 2). The H28A mutant, on the other hand, had a specific activity of 0.500 ± 0.007 µmol/min·mg (~50% wild-type enzyme activity), indicating that His-28 is not absolutely required for LysoPLA I activity (Fig. 2).
CD Spectra of Wild-type LysoPLA I and H28A, D174A, and H208A MutantTo exclude the possibility that the loss of the enzyme
activity in the mutant proteins was due to conformational changes in the mutants, CD spectra were taken for each of the purified mutants as
well as the wild-type protein. As shown in Fig.
4, the CD spectra of all the proteins
were essentially identical, demonstrating that the decreased enzyme
activity, whether a 100% loss for D174A and H208A or a 50% loss for
H28A, is not the result of misfolding or global conformational changes
in the mutants.
Regiospecificity of LysoPLA I
To explore the regiospecificity
of LysoPLA I, 31P NMR was used to monitor the hydrolysis of
both natural regioisomers (1PGPC and 2PGPC) under conditions in which
acyl migration was minimized. As shown in Fig.
5A, LysoPLA I readily
processed both isomers at similar rates. As the substrate concentration
(10 mM) used in the NMR measurements was much higher than
the KM value (22 µM) reported
previously (19), a linear time course was expected if the reaction was
not complicated by substrate/product inhibition or activation. However,
examination of the time courses for the consumption of both isomers as
well as the production of glycerophosphocholine revealed that the
reaction had two zero-order phases (Fig. 5B), an early
slower phase for up to 25 min and a later faster phase for up to 40 min. The data points after 40 min became non-zero order, reflecting the
much smaller concentrations of substrates remaining. Similar reaction
profiles were observed with secretary PLA2s (29), and the
complex time courses were attributed to changes in the interface
resulting from the fatty acid produced initially in those reactions.
However, as fatty acids inhibit the LysoPLA I (19), it remains unclear
what causes this complex time course. It is apparent, however, that
LysoPLA I can function equally well as either a LysoPLA1 or
a LysoPLA2.
LysoPLAs have been identified in many mammalian tissues and cells and are considered to be the major route by which lysophospholipids are degraded. The substrate used in lyso-PLA I activity assays is known to exist as a 9:1 equilibrium mixture of 1PGPC and 2PGPC, with the fatty acid predominantly at the sn-1 position (22). The Dole extraction, however, is unable to distinguish which isomer is hydrolyzed. To examine whether LysoPLA I has a preference for one isomer over the other, we have followed the hydrolysis of both isomers by 31P NMR. Remarkably, the results show that LysoPLA I processes both regioisomers at almost identical rates. These findings were obtained at 20 °C, and similar results were also obtained at 40 °C (data not shown). These observations are in contrast to what was found for the lysophospholipase activity of the Group IV Ca2+-dependent cytosolic PLA2, which exhibited a strong preference for 1PGPC over 2PGPC (30). The ability of LysoPLA I to efficiently hydrolyze both regioisomers suggests that Lyso-PLA I may function as both a LysoPLA1 and a LysoPLA2 in vivo, controlling lysophospholipids produced by both PLA1 and PLA2 (Scheme I). In addition, LysoPLA I may also play a role in arachidonic acid release and signal transduction since the arachidonic acid that occurs predominantly in the sn-2 position of phospholipids could also be cleaved by the sequential actions of a PLA1 and LysoPLA I. Interestingly, the enzyme's activity does not appear to be affected by the aggregation state of the substrate. Its activity does not vary dramatically as the substrates' concentration increases above its critical micelle concentration suggesting that interfacial activation does not play the same role in lysophospholipase activity as it does in phospholipase A2 activity (31).
By structural comparison to other esterases and /
-hydrolase, we
have identified two putative catalytic residues Asp-174 and His-208
that, together with the previously identified Ser-119, may form the
catalytic triad in LysoPLA I. This hypothesis is supported by the
site-directed mutagenesis studies, which showed that mutations of
either residue rendered the enzyme completely inactive, although the
D174A and H208A mutants retained the same protein global conformation
as that of the wild-type enzyme. These results are in strong contrast
to that observed for another mutant, H28A. Even though His-28 is also
highly conserved, mutation of it to Ala retained 50% activity of the
wild-type enzyme. Taken together with the previous results, we have
herein demonstrated that Ser-119, Asp-174, and His-208 are essential
for LysoPLA I activity, and LysoPLA I appears to function by a
mechanism involving a catalytic triad.
Structural prediction for LysoPLA I was achieved based on the predicted
secondary structure of LysoPLA I and the known crystal structure of
acetylcholinesterase, a member of the /
-hydrolase family (23). As
shown in Fig. 6, the three catalytic
residues Ser-119, Asp-174, and His-208, although far apart in the
primary sequence, come together and orient in such a way that they
could form the charge-relay network. Similar results were obtained
using another
/
-hydrolase (dielelactone hydrolase) as template
and agree with the notion that through divergent evolution the
three-dimensional configuration of the triad and therefore the
catalytic mechanism are highly conserved in many hydrolases (26, 27).
The structural model obtained here provides a valuable tool for the
design of studies to determine the function and regulation of this
enzyme. The validity of this structural model, however, awaits x-ray
structural studies for the enzyme, which are currently underway.
In summary, we have established an improved protein expression/purification procedure to quickly purify a large amount of active enzyme and demonstrated that LysoPLA I represents a new member of the serine hydrolase family with the catalytic triad composed of Ser-119, Asp-174, and His-208. LysoPLA I hydrolyzes both 1PGPC and 2PGPC equally well and may play an important role in controlling the level of lysophospholipids produced by both PLA1 and PLA2.
We thank Dr. Joseph Noel at the Salk Institute for introducing us to the PET28a(+) expression system, Dr. Murray Goodman for use of the CD spectrometer, and Dr. Alan Deese for considerable help with the NMR experiments.