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(Received for publication, January 20, 1995; and in revised form, August 8,
1995) From the
Hepatic lipase (HL) and lipoprotein lipase (LPL) are key enzymes
that mediate the hydrolysis of triglycerides (TG) and phospholipids
(PL) present in circulating plasma lipoproteins. Relative to
triacylglycerol hydrolysis, HL displays higher phospholipase activity
than LPL. The structural basis for this difference in substrate
specificity has not been definitively established. We recently
demonstrated that the 22-amino acid loops (``lids'') covering
the catalytic sites of LPL and HL are critical for the interaction with
lipid substrate (Dugi, K. A., Dichek, H. L., Talley, G. D., Brewer, H.
B., Jr., and Santamarina-Fojo, S.(1992) J. Biol. Chem. 267,
25086-25091). To determine whether the lipase lid plays a role in
conferring the different substrate specificities of HL and LPL, we have
generated four chimeric lipases. Characterization of these chimeric
enzymes using TG (triolein and tributyrin) or PL
(dioleoylphosphatidylcholine (DOPC) vesicles, DOPC proteoliposomes, and
DOPC-mixed liposomes) substrates demonstrated marked differences
between their relative PL/TG hydrolyzing activities. Chimeric LPL
containing the lid of HL had reduced triolein hydrolyzing activity (49%
of the wild type), but increased phospholipase activity in DOPC
vesicle, DOPC proteoliposome, and DOPC-mixed liposome assay systems
(443, 628, and 327% of wild-type LPL, respectively). In contrast,
chimeric HL containing the LPL lid was more active against triolein
(123% of the wild type) and less active against DOPC (23, 0, and 30%,
respectively) than normal HL. Similar results were obtained when the
lipase lids were exchanged in chimeric enzymes containing the
NH In summary, the lid
covering the catalytic domains in LPL and HL plays a crucial role in
determining lipase substrate specificity. The lid of LPL confers
preferential triglyceride hydrolysis, whereas the lid of HL augments
phospholipase activity. This study provides new insight into the
structural basis for the observed in vivo differences in LPL
and HL function.
Hepatic lipase (HL) ( Together with
pancreatic lipase, HL and LPL are members of the human lipase family
and share a high degree of primary sequence homology(5) . Their
catalytic domain consists of a Ser-His-Asp
triad(6, 7, 8) . Conservation of disulfide
bonds suggests a similar folding pattern(9) , and by homology
to the reported three-dimensional structure of pancreatic
lipase(10) , LPL and HL may be organized into two distinct
amino- and carboxyl-terminal domains. Both lipases are anchored to the
capillary endothelium via glycosaminoglycans and can be released by
intravenous administration of heparin(1, 11) . There are, however, striking differences between LPL and HL. LPL is
primarily synthesized by adipocytes, muscle cells, and macrophages,
whereas HL mRNA is primarily detected in hepatocytes(12) . LPL
is inhibited by 1 M NaCl (11) and requires its
cofactor, apoC-II, for full activation(13, 14) . HL,
on the other hand, is fully active even in the presence of high salt
concentrations and in the absence of a cofactor, although its activity
may be modulated by apoA-II (15, 16) and
apoE(17, 18, 19) . The two enzymes also
differ in their substrate specificity, demonstrating preferences in
their fatty acid positional specificity (20, 21) ,
fatty acid chain length(22, 23) , and degree of fatty
acid saturation(24, 25) . In addition, LPL and HL
differ in their preferred lipoprotein substrates. Thus, patients with
LPL deficiency accumulate primarily chylomicrons as well as very low
density lipoproteins in plasma(11) , whereas HL deficiency
results in elevated plasma concentrations of intermediate density
lipoproteins and HDL(26, 27) . Characterization of
these patients demonstrates that in vivo, the preferred
substrate of LPL are the large triglyceride-rich lipoproteins, whereas
HL is more active in the hydrolysis of smaller lipoproteins such as
HDL(28, 29) , especially HDL Recent studies involving the analysis of chimeric
LPL-HL mutants (39, 40, 41) have suggested
that the COOH-terminal domains of LPL and HL (Fig. 1) may play a
role in determining the preferred substrate of the respective
lipase(42) . In addition, removal of the COOH-terminal 58 amino
acids of LPL by chymotryptic cleavage resulted in the inability of LPL
to bind to chylomicrons(43) , suggesting that this region of
the enzyme may mediate binding to lipoproteins. However, to date, the
structural basis for the differences in substrate specificity between
the two lipases remains poorly defined. We have recently demonstrated
that the lid covering the catalytic domain of the human lipases (Fig. 1) is critically involved in the interaction of the
lipases with their lipid substrate(42) . In this report, we
investigate the role of the lipase lid in determining the substrate
specificity of HL and LPL for phospholipids and triglycerides by
generating four different chimeric lipases in which the LPL and HL lids
have been exchanged. Our studies demonstrate that the LPL lid enhances
triglyceride hydrolysis, whereas the HL lid augments phospholipase
function, thus establishing the important role of the lid in mediating
lipase substrate specificity.
Figure 1:
Schematic representation of the
structure of the
To evaluate a potential role of the lipase lid in modulating
the substrate specificities of LPL and HL, we first established the
importance of the lid region in the hydrolysis of different lipase
substrates. We had previously demonstrated that the integrity of the
amphipathic helices in the lipase lid is essential for the hydrolysis
of water-insoluble long chain fatty acid triglycerides(42) . In
a similar manner, we now investigated the role of the amphipathic
helices in the hydrolysis of water-insoluble phospholipid substrates. Table 1summarizes the concentration as well as activity in the
medium of cells transfected with wild-type LPL, wild-type HL, or
lipase-lid mutant plasmids. Disruption of the amphipathicity of helix 1
or helix 2 of the lid or deletion of the helices destroyed the ability
of the lipases to hydrolyze water-insoluble triglycerides, while the
esterase activity remained intact. Extensive rearrangement of both
helices in HL and LPL markedly reduced esterase activity, possibly by
reducing the ability of the lid to change to the open conformation upon
interfacial activation(42) . The tributyrin activity, however,
was still 5 times above background, whereas the more sensitive triolein
assay revealed absent activity. These results indicate that the
disruption of both helices in HL and LPL selectively prevented the
hydrolysis of liposoluble substrate. Similarly, disruption of the
amphipathic properties of the lipase lid abolished the ability of all
mutant enzymes to hydrolyze phospholipid substrate presented as a DOPC
vesicle (Table 1). Taken together, these studies demonstrate that
the lipase lid is essential for the hydrolysis of not only liposoluble
triglycerides, but also of liposoluble phospholipid substrates.
To
investigate a potential role of the lid in conferring substrate
specificity, we generated several mutant lipases. As illustrated in Fig. 2, in Mut I, the lid of LPL was replaced by the lid of HL,
and in the reciprocal mutant (Mut II), the lid of HL was replaced by
the lid of LPL. Mut III is a chimeric lipase in which the
carboxyl-terminal 134 amino acids of human LPL were replaced by the
carboxyl-terminal 146 amino acids of human HL. Mut IV is a chimeric
lipase that contains the NH
Figure 2:
Schematic representation of wild-type and
mutant LPL and HL constructs used for the analysis of substrate
specificity. The name as well as a general description of the
constructs utilized in this study are listed. LPL sequences are shown
in white, and HL sequences are shown in black. The
numbers indicate the amino acid residues of the particular lipase
comprising the NH
Table 2summarizes the tributyrin, triolein, and phospholipid
hydrolyzing activities in the medium of 293 cells transfected with the
different plasmids. Tributyrin is water-soluble at the concentrations
used in the assay and thus measures the esterase function of the
lipases. Table 2shows that substitution of the COOH-terminal
domain of LPL with that of HL (Mut III) leads to a parallel reduction
of tributyrin and triolein activities compared with LPL WT possibly due
to a destabilization of the active dimer(40) . Comparison of
Mut I with LPL WT, Mut II with HL WT, and Mut IV with Mut III indicates
that despite the exchange of the lids, the catalytic domain and
esterase function of the mutant lipases are preserved. Further analysis
reveals that the presence of the HL lid (Mut I versus LPL WT
and Mut IV versus Mut III) increased phospholipid and lowered
triolein hydrolysis, whereas the presence of the LPL lid (Mut II versus HL WT and Mut III versus Mut IV) had the
opposite effect.
Similar results were obtained when the
triacylglycerol hydrolase and phospholipase activities of the different
lipases were directly compared after normalization for tributyrin
(esterase) activity (Fig. 3). The ability of wild-type LPL and
HL as well as Mut I-IV to hydrolyze triglyceride (triolein) versus phospholipid (DOPC vesicles, DOPC proteoliposomes, and
DOPC-mixed liposomes) substrates is summarized in Fig. 3(A-E). Replacement of the LPL lid with the
lid of HL (Mut I) resulted in a 51% reduction in the ability of mutant
LPL to hydrolyze triolein as well as a 317-618% increase in the
phospholipid hydrolysis relative to that of wild-type LPL (Fig. 3A). Conversely, replacement of the HL lid with
that of LPL (Mut II) led to a 23% increase in triolein hydrolysis and a
reduction of phospholipase activity to <30% of wild-type HL (Fig. 3B).
Figure 3:
Triolein and DOPC hydrolyzing activities
of wild-type and mutant lipases. Activities are normalized for esterase
(tributyrin hydrolyzing) activity and presented as percent of wild-type
lipase activity. Wild-type lipase is shown as open bars, and
mutant lipase as closed bars. Standard deviations are
calculated from three transfection experiments. A, LPL WT and
Mut I; B, HL WT and Mut II; C, LPL WT and Mut III; D, LPL WT and Mut IV; E, Mut III and Mut IV. In A, D, and E, hatch marks on the x axis indicate the presence of separate scales for triolein
and DOPC activities.
Previous studies have proposed a role of
the COOH-terminal domain of the human lipases in mediating the
interaction with lipid substrates (40, 41) . We thus
analyzed a mutant LPL in which the COOH-terminal domain was replaced by
the COOH-terminal domain of HL (Mut III); the lipase lid in this
chimeric enzyme is from LPL. Fig. 3C shows that the
presence of the COOH-terminal domain of HL in LPL leads to only a
slight decrease in the ability to hydrolyze long chain fatty acid
triglycerides. The phospholipase activity of Mut III, however, was
significantly reduced using all three different phospholipid
substrates. To examine further the role of the lipase lid in modulating
lipase substrate specificity, we replaced the lid of LPL in Mut III
with the lid of HL to create a double mutant of LPL. In this case, both
the lid and the COOH-terminal domain of LPL are replaced by the
analogous structures of HL. Fig. 3D demonstrates that
compared with wild-type LPL, the double mutation leads to a 50%
reduction in the ability of the lipase to hydrolyze water-insoluble
triglyceride substrate and up to a >5-fold increase in phospholipase
activity, depending on the DOPC substrate used. When the ability of the
double mutant LPL Mut IV to hydrolyze different substrates was directly
compared with that of the chimera Mut III (Fig. 3E), a
42% reduction of triolein activity but up to a 13-fold increase in
phospholipase activity were evident. Thus, the additional presence of
the HL lid again resulted in marked enhancement of phospholipase
function, stressing the importance of the lipase lid in conferring
substrate specificity. Table 3summarizes the DOPC/triolein
ratios for wild-type LPL and HL as well as the mutant lipases using
either the DOPC vesicle or DOPC-mixed liposome substrate as a measure
of phospholipase activity. Compared with native LPL, the ratio for
native HL is 5-7-fold greater. To facilitate the evaluation of
the role the lipase lid plays in determining substrate specificity, the
data were separated into two groups. In the top half of Table 3,
all lipases containing the LPL lid are listed: wild-type LPL, HL with
the LPL lid (Mut II), and LPL with the COOH-terminal domain of HL (Mut
III). The constructs containing the lid of hepatic lipase are listed in
the bottom half of Table 3: wild-type HL, LPL with the HL lid
(Mut I), and LPL with the COOH-terminal domain and lid of HL (Mut IV).
When normalized for wild-type LPL, all constructs containing the lid of
LPL have a DOPC/triolein ratio of 1.4 or less, whereas the constructs
with the HL lid have a ratio of 3.5-11.5. Analysis of Mut I and
Mut II demonstrates that the exchange of the lipase lid leads to a
reversal of DOPC/triolein ratios of LPL and HL, indicating that the lid
may be the single most important region in conferring lipase substrate
specificity.
In this study, we have investigated a structural motif in LPL
and HL that mediates the different ability of the two enzymes to
hydrolyze triglyceride versus phospholipid substrates.
Relative to triolein hydrolyzing activity, HL has been shown to be a
more potent phospholipase than LPL in
vitro(1, 33, 34, 35) . This
difference in substrate specificity may play an important role in the
biological function of LPL and HL (35) and becomes apparent in
the markedly different phenotype observed in LPL (37, 38) and HL (36) deficiency states. We
have recently demonstrated that the 22-amino acid lid that covers the
catalytic site (Fig. 1) plays a crucial role in mediating the
interaction of the lipases with emulsified triglycerides. Since there
is little sequence homology between the LPL and HL lids(5) , we
postulated that the different primary structures of the lids of LPL and
HL may, in part, modulate lipase substrate specificity(42) . The data presented in Table 1demonstrate that the lids of LPL
and HL are important for phospholipid as well as triglyceride
hydrolysis. This is in contrast to guinea pig pancreatic lipase, which
contains only a ``minilid'' of 5 amino acids, but is still
able to hydrolyze phospholipid substrate(61) . Our mutant in
which the 22-amino acid lid was replaced by a 4-amino acid peptide had
absent phospholipase activity even though its esterase activity was
>2-fold higher than that of wild-type LPL. This, together with the
data from the other mutants listed in Table 1, indicates that in
human LPL and HL, the lid plays a crucial role in triglyceride as well
as phospholipid hydrolysis. Analysis of Mut I and Mut II (Fig. 3, A and B) demonstrates that the simple
exchange of the lids between LPL and HL markedly altered the ability of
the lipases to hydrolyze certain substrates. The lid of HL increased
the phospholipase and reduced the triglyceride hydrolase activity,
whereas the lid of LPL had the opposite effect. Similar findings were
obtained in lipase mutants in which the COOH termini alone or both the
COOH termini and the lids of LPL and HL were exchanged. Analysis of the
chimeric enzyme containing the NH Analysis of the DOPC/triolein ratios as an indicator of the relative
phospholipase/triglyceride hydrolase function of normal and mutant
lipases demonstrated that this ratio is higher for HL than for LPL (Table 3). Our studies clearly demonstrate that whenever the LPL
lid was present (Mut II and Mut III) (Table 3), the DOPC/triolein
ratio was similar to that of LPL ( The
mechanism by which the 22-amino acid lid mediates the different
substrate specificities of LPL and HL may be complex. Previous studies
have demonstrated that the lid must be repositioned to allow access of
substrate to the catalytic pocket(10) . We have suggested that,
upon opening of the lid, the hydrophobic sides of its amphipathic
helices are exposed(42) , thus creating a large hydrophobic
area, which may then function as a binding site for lipid substrates. A
similar mechanism has been previously demonstrated for an unrelated
fungal lipase(62, 63) . More recently, studies in
which the three-dimensional structure of pancreatic lipase was
cocrystallized with mixed micelles of phosphatidylcholine and bile salt (64) have demonstrated that, indeed, one of the two fatty acids
of phospholipid binds to the hydrophobic side chains of the amino acid
residues in the lipase lid. A molecular model of LPL based on the
pancreatic lipase x-ray structure (65) indicates that the lid
of LPL is indeed likely to be organized into amphipathic helices and
that it may play a role similar to that of the pancreatic lipase lid.
Thus, like in pancreatic lipase, the lids of LPL and HL may come into
intimate contact with lipase substrates and thus play a major role in
mediating lipase substrate specificity. In this context, the
primary, secondary, and tertiary structures of the lid may be
important. Although the secondary structural organization of the lid
into highly amphipathic helices appears to be very similar in LPL and
HL(42) , there is only a small degree of primary sequence
homology. In fact, 15 out of 22 amino acids of the lid are different
between LPL and HL. Further analysis of the lipase lids reveals that
the LPL lid contains four acidic and three basic amino acids, whereas
the lid of HL contains only one acidic but five basic residues. In the
investigation of substrate specificity, we controlled for different
fatty acid preferences between LPL and HL (22, 23, 24, 25) by using oleic acid
as the fatty acid in triglyceride (triolein) and phospholipid (DOPC)
substrates. Thus, the observed differences in substrate specificity
may, in part, be due to different abilities of the LPL and HL lids to
accommodate the polar head group of phospholipids. The differences in
the distribution of charged residues in the lid may thus be crucial
since a negatively charged lid could interfere with the entrance of a
phospholipid with its negatively charged polar head group into the
catalytic pocket. Therefore, the lid of HL may be more suited for
interaction with a phospholipid substrate than the LPL lid. A
molecular model of LPL based on the x-ray structure of pancreatic
lipase reveals another, smaller surface loop between residues 54 and 64
as well as three loops (residues 91-95, 157-160, and
187-196) that make up the hydrophobic groove surrounding the
active site(65) . These loops are more highly conserved between
LPL and HL, and the region contains two additional negative charges in
HL, making it less likely that this region confers preferential
phospholipid hydrolysis. While a contribution of these loops to the
determination of substrate specificity cannot be ruled out, the
reversal of phospholipid/triglyceride hydrolyzing ratios by the mere
exchange of the lids suggests that the lids of LPL and HL are the
single most important regions determining substrate specificity. In
conclusion, characterization of chimeric enzymes in which the LPL and
HL lids have been exchanged has demonstrated that the lipase lid plays
a crucial role in mediating the different substrate specificities of
LPL and HL. The lid of HL augments phospholipase activity, whereas the
lid of LPL increases triglyceride hydrolysis. This study provides new
insights into the structural basis for the observed in vivo differences in LPL and HL function.
Volume 270,
Number 43,
Issue of October 27, 1995 pp. 25396-25401
©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
-terminal end of LPL and the COOH-terminal domain of HL.
Exchange of the LPL and HL lids resulted in a reversal of the
phospholipase/neutral lipase ratio, establishing the important role of
this region in mediating substrate specificity.
)and lipoprotein lipase (LPL)
are critical enzymes that play a central role in lipoprotein
metabolism. Their main function is to hydrolyze triglycerides and
phospholipids present in circulating plasma lipoproteins, including
chylomicrons, very low and intermediate density lipoproteins, and
HDL(1) . In the process, they generate free fatty acids, which
can be utilized for storage or generation of energy. By influencing HDL
metabolism(2, 3, 4) , these two lipases may
also play a role in reverse cholesterol transport.(30) ,
and intermediate density lipoproteins (26, 31, 32) . Another physiologically
important difference in the function of HL and LPL is the relative
phospholipase versus triacylglycerol hydrolase activity of the
two enzymes. Several studies have demonstrated that, relative to
triglyceride hydrolysis, HL is a more active phospholipase than
LPL(1, 33, 34, 35) . In fact,
inhibition of HL by infusion of antisera in cynomolgus monkeys led to a
marked accumulation of not only triglycerides, but also of
phospholipids in different lipoprotein fractions, especially
HDL(36) . Similar changes in plasma phospholipid levels have
been observed in some patients with HL deficiency(32) , but not
with LPL deficiency(37, 38) . This difference in
relative phospholipase to triacylglycerol hydrolase activity of the two
enzymes may be important for modulating the function of LPL and HL in vivo.
-carbon backbone of horse pancreatic lipase as
reported by Bourne et al.(66) . The loop covering the
catalytic pocket and the COOH-terminal domain are highlighted.
Residues of the catalytic triad (corresponding to Ser-132, Asp-156, and
His-241 in human LPL) and the disulfide bridges are represented as ball-and-sticks. Solid arrows represent 
-sheet
structures. The region of highest homology between pancreatic lipase on
one hand and HL and LPL on the other hand is the
NH-terminal region, whereas the lid region and the
COOH-terminal domain are less strictly conserved. The drawing was
generated with the program
MOLSCRIPT(67) .
cDNA Expression Vector
The parent plasmid (pCMV)
used for site-directed mutagenesis and transfection is a pUC18-derived
vector containing the cytomegalovirus immediate early promoter and the
polyadenylation site of SV40 as described previously(44) . A
1473-base pair fragment of normal human LPL cDNA (pCMV-NLPL) (45) or a 1522-base pair fragment of normal human HL cDNA
(pCMV-NHL) (46, 47, 48) was cloned into the XbaI and HpaI restriction sites of pCMV. The DNA
sequence of each fragment, which spanned the signal peptide through the
termination codon, was confirmed by DNA sequence analysis using the
dideoxynucleotide chain termination method (49) and T7 DNA
polymerase (Sequenase, U. S. Biochemical Corp.).Synthesis of Mutant cDNA
The mutant HL and LPL
cDNAs were synthesized by the overlap extension polymerase chain
reaction (50) using either pCMV-NLPL or pCMV-NHL as template.
Polymerase chain reaction was performed in an automated DNA thermal
cycler (Perkin-Elmer) as described (51) utilizing DNA
polymerase from Pyrococcus furiosus (Stratagene Inc., La
Jolla, CA) and 30 cycles with 1-min denaturation at 95 °C, 1-min
annealing at 50 °C, and 2-min extension at 72 °C in 1 
buffer 2 (Stratagene Inc.), 200 µM each dATP, dCTP, dGTP,
and dTTP (Boehringer Mannheim), and 0.5 µM each primer.
The exchange of the lids was performed with partially complementary
oligonucleotide primers spanning the entire noncomplementary region of
the 22-amino acid lid. Generation of chimeric proteins by domain
exchange was performed as described previously(40) . The mutant
cDNAs were subcloned into the pCMV expression vector and amplified
using competent DH5 cells (Life Technologies Inc.). Clones
carrying the mutant cDNA were grown overnight at 37 °C in LB broth
(Biofluids, Inc., Rockville, MD), and DNA was isolated by one-tube
minipreparation(52) . All constructs were examined by sequence
analysis of the complete cDNA insert. Oligonucleotide primers for
overlap extension polymerase chain reaction and sequencing were
synthesized by the phosphoramidite method on a DNA synthesizer (Model
380B, Applied Biosystems, Inc., Foster City, CA).In Vitro Expression of cDNA in Human Embryonal Kidney 293
Cells
Plasmids used for transfection were purified by the cesium
chloride double-banding method(53) . Transfections were
performed using the calcium phosphate coprecipitation method (54) by adding 40 µg of plasmid DNA to each 100-mm plate of
subconfluent human embryonal kidney 293 cells (American Type Culture
Collection, Rockville, MD). Twenty-four hours after addition of DNA,
the cells were washed, and medium containing 10% (v/v) fetal calf serum
and 2 units/ml heparin sodium (Elkins-Sinn, Cherry Hill, NJ) was added.
Medium for activity determination was harvested 12-16 h after
washing and supplemented with glycerol to a final concentration of 30%
(v/v). Intracellular protein was harvested as described by Chait et
al.(55) . Aliquots of media and intracellular extracts
were kept at -70 °C until lipase assays were performed. Each
plasmid was transfected in triplicate. Wild-type HL and LPL were used
as positive controls, and the pCMV vector without insert was used as
negative control.Determination of HL and LPL Activities
Esterase
activity was quantitated in triplicate using
[
C]tributyrin(56) , and triglyceride
lipase activity was determined in triplicate using
[C]triolein as previously
published(57) . Phospholipase activities were measured in
triplicate utilizing three different substrates. Phospholipid vesicles
were generated by modifying the synthesis of triolein emulsion (57) as follows. Dioleoylphosphatidylcholine (DOPC) (1
mM; Sigma) was used instead of egg yolk extract. Labeled
triolein was substituted with [C]DOPC (Amersham
Corp.) at an activity of 0.1 µCi/ml of substrate. Proteoliposomes
were synthesized according to a previously published protocol (58) with the following modifications. DOPC was used as the
unlabeled phospholipid, and the proteoliposomes were labeled with 1
µCi of [C]DOPC. One-hundred microliters of
substrate were added to 200 µl of medium from transfected cells in
a total volume of 500 µl (0.15 M NaCl, 0.1 M Tris-HCl, pH 8.5, 2.5% bovine serum albumin, 25 µl of human
plasma (as source of apoC-II), 2 units/ml heparin). Samples were
incubated at 37 °C in a shaking water bath for 1-4 h, and
oleic acid was extracted by the method of Belfrage and Vaughan (59) . Mixed liposome substrate was prepared by a modification
of a previously published protocol (60) using unlabeled DOPC,
unlabeled triolein (Sigma), and 2 µCi of
[C]DOPC. Assay conditions used were the same as
in the proteoliposome protocol.Determination of LPL Mass
LPL mass was determined
six times by an enzyme-linked immunosorbent assay using the 5D2
monoclonal antibody (kindly provided by Dr. J. D. Brunzell, University
of Washington, Seattle) for capture and a chicken polyclonal antibody
(kindly provided by Dr. I. J. Goldberg, Columbia University, New York)
for measurement.
-terminal domain of LPL and the
COOH-terminal domain as well as the lid of HL. In addition to these
four chimeric constructs, plasmids containing wild-type LPL and HL
cDNAs and a negative control consisting of the parent vector were
transfected into 293 cells. Greater than 95% of the total triolein
hydrolyzing activity was found in the culture medium of all transfected
cells, indicating that the mutant lipases were secreted to a similar
extent as wild-type LPL and HL (data not shown).
-terminal domain of the respective
construct.
-terminal domain of human
LPL and the COOH-terminal domain of human HL (Mut III) demonstrated a
slight reduction of triolein as well as a significant decrease in DOPC
hydrolyzing activity compared with normal LPL (Fig. 3C). This finding is consistent with previous
studies (39, 40, 43) that suggest that the
COOH-terminal domain of LPL may play a role in mediating the initial
interaction of the lipase with lipoproteins and perhaps modulate the
major type of particle with which either lipase will
interact(42) . Davis et al.(41) conclude from
their analysis of similar chimeric enzymes in which the COOH-terminal
domains of LPL and HL are exchanged that the COOH-terminal domain of HL
augments phospholipase function. Based on the results presented here,
the lid appears to be the more important determinant of phospholipase
hydrolyzing activity. Analysis of the data reported by Davis et al.(41) indicates that replacement of the
NH-terminal domain of HL with the NH-terminal
domain and thus the lid of LPL resulted in a 2.5-fold reduction in the
dipalmitoylphosphatidylcholine/triolein ratio. Replacement of the
NH-terminal domain of LPL with that of HL, on the other
hand, led to a 20-fold increase in this ratio. In addition, direct
comparison of the activity of our Mut III with that of Mut IV, the
latter containing the COOH-terminal domain as well as the lid of HL in
the LPL backbone (Fig. 3E), demonstrates that the
presence of the HL lid in this chimera leads to a 40% reduction of
triolein activity and up to a 13-fold increase in DOPC hydrolyzing
activity. Thus, the presence of the HL lid markedly enhanced
phospholipase function in Mut IV, indicating that the lid may be more
important in determining phospholipase function than the COOH terminus. 
1.4). Conversely, whenever the HL
lid was present (Mut I and Mut IV) (Table 3), the ratio increased
to >3.5, similar to that of normal HL. Thus, regardless of the type
of COOH terminus or lipase backbone present, exchange of the lids
between the lipases resulted in a reversal of the
phospholipase/triglyceride hydrolase ratios of LPL and HL.
)
©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
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N. Griffon, E. C. Budreck, C. J. Long, U. C. Broedl, D. H. L. Marchadier, J. M. Glick, and D. J. Rader Substrate specificity of lipoprotein lipase and endothelial lipase: studies of lid chimeras J. Lipid Res., August 1, 2006; 47(8): 1803 - 1811. [Abstract] [Full Text] [PDF]
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M.-E. Paradis, P. Couture, Y. Bosse, J.-P. Despres, L. Perusse, C. Bouchard, M.-C. Vohl, and B. Lamarche The T111I mutation in the EL gene modulates the impact of dietary fat on the HDL profile in women J. Lipid Res., October 1, 2003; 44(10): 1902 - 1908. [Abstract] [Full Text] [PDF]
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R. J. Brown, J. R. Schultz, K. W. S. Ko, J. S. Hill, T. A. Ramsamy, A. L. White, D. L. Sparks, and Z. Yao The amino acid sequences of the carboxyl termini of human and mouse hepatic lipase influence cell surface association J. Lipid Res., July 1, 2003; 44(7): 1306 - 1314. [Abstract] [Full Text] [PDF]
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 X.-Y. Wen, R. A. Hegele, J. Wang, D. Y. Wang, J. Cheung, M. Wilson, M. Yahyapour, Y. Bai, L. Zhuang, J. Skaug, et al. Identification of a novel lipase gene mutated in lpd mice with hypertriglyceridemia and associated with dyslipidemia in humans Hum. Mol. Genet., May 15, 2003; 12(10): 1131 - 1143. [Abstract] [Full Text] [PDF]
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 K. Winkler, B. Wetzka, M. M. Hoffmann, I. Friedrich, M. Kinner, M. W. Baumstark, H.-P. Zahradnik, H. Wieland, and W. Marz Triglyceride-Rich Lipoproteins Are Associated with Hypertension in Preeclampsia J. Clin. Endocrinol. Metab., March 1, 2003; 88(3): 1162 - 1166. [Abstract] [Full Text] [PDF]
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S. Y. Choi, K.-i. Hirata, T. Ishida, T. Quertermous, and A. D. Cooper Endothelial lipase: a new lipase on the block J. Lipid Res., November 1, 2002; 43(11): 1763 - 1769. [Abstract] [Full Text] [PDF]
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H. Wong and M. C. Schotz The lipase gene family J. Lipid Res., July 1, 2002; 43(7): 993 - 999. [Abstract] [Full Text] [PDF]
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M. G. McCoy, G.-S. Sun, D. Marchadier, C. Maugeais, J. M. Glick, and D. J. Rader Characterization of the lipolytic activity of endothelial lipase J. Lipid Res., June 1, 2002; 43(6): 921 - 929. [Abstract] [Full Text] [PDF]
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X.-P. Yang, L. A. Freeman, C. L. Knapper, M. J. A. Amar, A. Remaley, H. B. Brewer Jr., and S. Santamarina-Fojo The E-box motif in the proximal ABCA1 promoter mediates transcriptional repression of the ABCA1 gene J. Lipid Res., February 1, 2002; 43(2): 297 - 306. [Abstract] [Full Text] [PDF]
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T. Keiper, J. G. Schneider, and K. A. Dugi Novel site in lipoprotein lipase (LPL415;-438) essential for substrate interaction and dimer stability J. Lipid Res., August 1, 2001; 42(8): 1180 - 1186. [Abstract] [Full Text] [PDF]
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H. L. Dichek, S. M. Johnson, H. Akeefe, G. T. Lo, E. Sage, C. E. Yap, and R. W. Mahley Hepatic lipase overexpression lowers remnant and LDL levels by a noncatalytic mechanism in LDL receptor-deficient mice J. Lipid Res., February 1, 2001; 42(2): 201 - 210. [Abstract] [Full Text]
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 R. Abia, Y. M. Pacheco, J. S. Perona, E. Montero, F. J. G. Muriana, and V. Ruiz-Gutiérrez The Metabolic Availability of Dietary Triacylglycerols from Two High Oleic Oils during the Postprandial Period Does Not Depend on the Amount of Oleic Acid Ingested by Healthy Men J. Nutr., January 1, 2001; 131(1): 59 - 65. [Abstract] [Full Text]
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 K. A. Dugi, M. J. A. Amar, C. C. Haudenschild, R. D. Shamburek, A. Bensadoun, R. F. Hoyt Jr, J. Fruchart-Najib, Z. Madj, H. B. Brewer Jr, and S. Santamarina-Fojo In Vivo Evidence for Both Lipolytic and Nonlipolytic Function of Hepatic Lipase in the Metabolism of HDL Arterioscler. Thromb. Vasc. Biol., March 1, 2000; 20(3): 793 - 800. [Abstract] [Full Text] [PDF]
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Y. Yang and M. E. Lowe The open lid mediates pancreatic lipase function J. Lipid Res., January 1, 2000; 41(1): 48 - 57. [Abstract] [Full Text]
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 K.-i. Hirata, H. L. Dichek, J. A. Cioffi, S. Y. Choi, N. J. Leeper, L. Quintana, G. S. Kronmal, A. D. Cooper, and T. Quertermous Cloning of a Unique Lipase from Endothelial Cells Extends the Lipase Gene Family J. Biol. Chem., May 14, 1999; 274(20): 14170 - 14175. [Abstract] [Full Text] [PDF]
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Y. Nagai, J. Aoki, T. Sato, K. Amano, Y. Matsuda, H. Arai, and K. Inoue An Alternative Splicing Form of Phosphatidylserine-specific Phospholipase A1 That Exhibits Lysophosphatidylserine-specific Lysophospholipase Activity in Humans |
ABSTRACT
