Crystal Structure of Human Gastric Lipase and Model of Lysosomal
Acid Lipase, Two Lipolytic Enzymes of Medical Interest*
Alain
Roussel
§,
Stéphane
Canaan§¶,
Marie-Pierre
Egloff
,
Mireille
Rivière¶,
Liliane
Dupuis¶,
Robert
Verger¶, and
Christian
Cambillau
From the
Architecture et Fonction des
Macromolécules Biologiques, CNRS-IFR1 UPR 9039 and
¶ Laboratoire de Lipolyse Enzymatique, CNRS-IFR1 UPR 9025, 31 chemin Joseph Aiguier, 13402 Marseille cedex 20, France
 |
ABSTRACT |
Fat digestion in humans requires not only the
classical pancreatic lipase but also gastric lipase, which is stable
and active despite the highly acidic stomach environment. We report
here the structure of recombinant human gastric lipase at 3.0-Å
resolution, the first structure to be described within the mammalian
acid lipase family. This globular enzyme (379 residues) consists of a
core domain belonging to the
/
hydrolase-fold family and a "cap" domain, which is analogous to that present in serine
carboxypeptidases. It possesses a classical catalytic triad (Ser-153,
His-353, Asp-324) and an oxyanion hole (NH groups of Gln-154 and
Leu-67). Four N-glycosylation sites were identified on the
electron density maps. The catalytic serine is deeply buried under a
segment consisting of 30 residues, which can be defined as a lid and
belonging to the cap domain. The displacement of the lid is necessary
for the substrates to have access to Ser-153. A phosphonate inhibitor
was positioned in the active site that clearly suggests the location of
the hydrophobic substrate binding site. The lysosomal acid lipase was
modeled by homology, and possible explanations for some previously
reported mutations leading to the cholesterol ester storage disease are given based on the present model.
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INTRODUCTION |
Since 1990, when the first three-dimensional structures of a
fungal (Rhizomucor miehei lipase) and a mammalian lipase
(human pancreatic lipase
(HPL))1 were published,
growing interest in lipolysis has led to the structural determination
of several lipases of various origins, including those present in
bacteria, fungi, and mammals. All the lipases investigated so far vary
considerably in size and in their amino acid sequences. However, they
are all serine esterases belonging to the
/
hydrolase superfamily
(1) in which the nucleophilic serine, part of a Ser-His-(Asp/Glu)
triad, is located in an extremely sharp turn (nucleophilic elbow).
Another feature that is common to all the members of the
/
hydrolase superfamily as well as to proteases is the occurrence of an
oxyanion hole, which stabilizes the transition state. Some
organophosphorous compounds inhibit lipases in a similar way to what
occurs in the case of serine proteases (2).
The three-dimensional structures of several complexes consisting of
lipases bound to covalent inhibitors have been solved: R. miehei lipase (3, 4) and Candida antartica B lipases bound to a C6-alkyl phosphonate (5), Candida rugosa lipase bound to long chain alkyl sulfonyl (6), Pseudomonas cepacia lipase (7) as well as cutinase (8) bound to a
dialkylcarbamoylglycerophosphonate, and human pancreatic
lipase-colipase complex bound to C11-alkyl phosphonate (9). It has been
established that the covalently inhibited lipases are in the so called
"open" conformation, i.e. that the lid has moved away to
give free access to the active-site serine. It has been suggested that
this mechanism may be instrumental in the binding of lipases to the
water-lipid interface and that the presence of a lid in the structure
of the enzyme may be involved in the interfacial activation process (4,
10).
Among the mammalian lipases, the acid lipases belong to a family of
enzymes that have the ability to withstand acidic conditions. This
family that includes the preduodenal lipases and human lysosomal lipase
shows no sequence homology with any other known lipase families (11).
The preduodenal lipases form a group of closely related enzymes
originating either from the stomach, the tongue, or the pharynx (12).
They all have a low pH optimum, and none of them require any specific
protein cofactor. Human gastric lipase (HGL, EC 3.1.1.3) is secreted by
the chief cells located in the fundic part of the stomach (13), where
it initiates the digestion of triacylglycerols (14, 15). The maximum
specific activities of HGL are 1160 units/mg on TC4 (pH 6.0), 1110 units/mg on TC8 (pH 6.0), and 600 units/mg on IntralipidTM (pH 5.0)
(16). Native HGL has an apparent molecular mass of 50 kDa and is a
highly glycosylated molecule with 4 potential
N-glycosylation sites (17). The glycan moiety was estimated
to account for around 15% of its total protein mass (18).
This enzyme plays a crucial role in newborns, because pancreatic lipase
is not yet fully developed at this age (15). The physiological
importance of gastric lipase has been suspected for some time, based on
pathological situations involving pancreatic exocrine insufficiency,
such as the late stage of chronic pancreatitis or cystic fibrosis. In
these cases, even in the complete absence of pancreatic lipase, the
patients still absorb a high percentage of their ingested dietary fat
(19, 20). In substitutive enzymatic therapy, the use of
acidic-resistant lipases should in principle yield more satisfactory
results than the pancreatic preparations currently in use. The
co-administration of acidic lipases, which hydrolyze dietary lipids
under acidic conditions, should help to treat patients with various
forms of pancreatic deficiency. Physiological studies have shown that
preduodenal lipases are capable of acting not only in the stomach
but also in the duodenum in synergy with a pancreatic lipase (14).
Various clinical studies have been conducted on both animals and humans
to assess the efficacy of enzymatic replacement therapies using
acid-resistant lipases to treat exocrine pancreatic insufficiency (21).
This treatment significantly increased the weight and reduced the
steatorrhea in dogs.
Despite the close amino acid sequence similarities (59% of the amino
acids are identical) between HGL (17) and human lysosomal acid lipase
(HLAL, EC 3.1.1.3) (22, 23), HGL lacks the cholesteryl ester hydrolase
activity reported in HLAL. The latter enzyme hydrolyzes not only the
triglycerides that are delivered to the lysosomes by low density
lipoprotein receptor-mediated endocytosis but also cholesteryl esters
(24). The cholesterol released by this reaction plays an important
regulatory role in cellular sterol metabolism. Defective HLAL activity
has been found to be associated with two rare autosomal recessive
traits, Wolman disease and cholesteryl ester storage disease. In Wolman
disease (25), a lack of HLAL activity results in a pronounced
accumulation of cholesteryl esters and triacylglycerols in the
lysosomes in most of the body tissues. The patients usually succumb to
hepatic and adrenal failure within the first year of life. Cholesteryl
ester storage disease, the other clinically recognized phenotypic form
of HLAL deficiency, follows a more benign clinical course (26), and a
residual HLAL activity has been detected. Since the cloning of the
cDNA and determination of the genomic organization of the gene
(LIPA) located on chromosome 10, which encodes HLAL (22, 23,
27), some deleterious LIPA gene mutations have been
identified (28-34). Most of these mutations affect either the mRNA
splicing or the amino acid sequence of HLAL. The exact correlations
between these mutations and the biochemical and clinical phenotypes
still remains to be elucidated, however. In this study, the crystal
structure of recombinant HGL at 3.0-Å resolution is determined, and a
model of HLAL is discussed and used to possibly explain the previously
reported cholesteryl ester storage disease mutations.
 |
MATERIALS AND METHODS |
Recombinant HGL (rHGL) Expression and Purification--
rHGL was
expressed in the baculovirus/insect cell system (48). The active enzyme
was produced on a large scale (5-13 mg/liter) from recombinant
baculovirus-infected insect cells using a bioreactor and its specific
activity (µmole·min
1·mg
1) was around
700 units/mg (49). The amino acid sequence (KLHPG) of rHGL in the
purified protein starts at residue 4.
rHGL Crystallization and Enzymatic Activity in the
Crystal--
Crystallization experiments were performed using the
hanging-drop vapor diffusion method. Crystals of the rHGL were obtained by mixing 2 µl of a well solution (2 M ammonium sulfate,
1.4% tert-butanol, at pH 5.0) with 2 µl of a protein
solution at 5-6 mg/ml. The crystals are cubic, space group I
21 3 with cell dimensions a = b = c = 244.0 Å3. The protein mass, 47,673 Da, was determined from
collected crystals by matrix-assisted laser desorption ionization
time-of-flight spectroscopy. There are two molecules in the asymmetric
unit (see below), and the Vm was estimated to be
6.35 Å3/Da (81% water content). For the mercury
derivative preparation, the crystals were transferred in a synthetic
liquor corresponding to the well solution containing 23 mM
mercuric acetate.
To assess the catalytic activity of the crystallized enzyme, tests were
performed on both dissolved and intact crystals. Ten crystals were
washed three times in the crystallization buffer and were subsequently
dissolved in 50 µl of water, and the protein concentration was
estimated by absorbance at 230 nm. The lipase activity was measured
titrimetrically at 37 °C using a pH stat (metrohm) at pH 5.7 with a
tributyrin emulsion as the substrate: 0.5 ml of tributyrin added to
14.5 ml of 150 mM NaCl, 2 mM taurodeoxycholate, and 2 mM bovine serum albumin (16).
The specific activity
(µmole·min
1·mg
1) was calculated and
found to be around 580 units/mg. To test the catalytic activity
in situ, a single rHGL crystal was incubated in the
crystallization solution containing 0.1 mM Nitroblue
tetrazolium and 0.75 mM 5 bromo-chloro-3-indoyl butyrate as
the substrate. After 24 h of incubation, the crystal was intensely
colored in blue/gray.
Data Collection and Heavy Metal Derivative Search--
All data
sets were collected at 100 K using a cryo-stream cooler from Oxford
Cryosystems. A first native and a derivative data set were collected
in-house using a 300-mm MAR Research imaging plate detector mounted on
a RU200 rotating anode generator (Rigaku, Tokyo, Japan). The generator
was operated at 3.2 kW with a focal spot size of 0.3 × 0.3 mm2. A second native data set was collected at LURE (Orsay,
France) on DW32 beamline at 0.963 Å wavelength using a 345-mm MAR
Research imaging plate. All data were collected in frames of 1.0 degree and processed with DENZO. The scaling was performed with SCALA (CCP4
(50)), and the derivative was merged with FHSCAL. Data collection
statistics are given in Table I. The fact that most of the crystals
diffracted only very poorly made the heavy metal derivative search
laborious. In the end, it led to one mercury derivative diffracting to
3.6-Å resolution, isomorphous to the native crystal.
Phase Determination--
The position of the heavy metal was
determined using Patterson methods. The MLPHARE software program (51)
was then used to refine the heavy atom parameters and calculate phases
to 4.0 Å, taking into account both isomorphous and anomalous
differences. Resulting single isomorphous replacement with anomalous
scattering (SIRAS) phases were considerably improved after flattening
80% of the solvent using density modification. At this stage, the map
indicated clearly that the handedness of the helices was wrong. Repeating the previous steps with the heavy atom position giving the
negative coordinates yielded a suitable map for determining an envelope
for each of the two molecules in the asymmetric unit. The
noncrystallographic symmetry operators were determined using both the
GLRF program and the relation between the two molecular replacement
solutions (automated molecular replacement density modification (52))
obtained from a search using a bones model (MAPMAN (53)) calculated on
the basis of one molecule. The density modification program was then
used again to improve the initial phases (from MLPHARE) by performing
simultaneously solvent flattening, histogram matching, 2-fold
noncrystallographic symmetry averaging, and phase extension. The best
map was obtained when a suitable mask around one of the molecules was
added to the program. The resolution of the derivative data set was
actually better than that of the native one, and the map was therefore
calculated using the derivative structure factor amplitudes with the
phases extended to 3.6 Å.
Model Building, Refinement, and Analysis--
The model was
built with the program TURBO-FRODO (54) using the recently developed
ab-initio building tools, which make it possible to build a
model from planar pseudo-residues in a very short time. When the
connectivity and the direction of the polypeptide chain have been
determined, the pseudo-residues are automatically replaced by the
actual residues in the sequence. Side-chain fitting is then performed
manually. When most of the model had been built, a 3.0-Å native data
set was collected on synchrotron radiation at the LURE. These better
data were then used to calculate a new map, with which the model was
completed (370 residues of 376 in the recombinant protein). The four
sugar glycosylation sites were already clearly identified in this
experimental electron density map. Refinement was carried out in X-PLOR
(55) against the 3.0-Å data. Five percent of the data were set aside for calculating and monitoring of the free R-factor (Rfree). Refinement involved cycles of simulated annealing using the slow-cool procedure interspersed with manual rebuilding. Noncrystallographic symmetry restraints were applied during the whole refinement procedure. Individual B-factor refinement was subsequently performed. The final
model consisted of 6194 atoms; the final R-factor was 22.5%, and the
R-free factor was 25.1%. The complete refinement statistics, given in
Table I, the quality of the Ramachandran
plot, and the electron density indicate that the model is better than
expected for a structure at 3.0-Å resolution. Maps calculated from the derivative structure factor amplitudes showed a clear electron density
for the mercury derivative bound to cysteine 244 but no significant
movements of any part of the model. Fig. 1 was drawn up with Molscript
(56) and Raster 3D (57), Fig. 2 with Clustal W (58) and Alscript (59),
Figs. 3, 4, and 6 with Turbo-Frodo (54), and Fig. 5 with GRASP (60).
The coordinates and structure factor amplitudes have been deposited in
the Protein Data Bank with entry code 1HLG.
 |
RESULTS AND DISCUSSION |
After numerous trials using weakly diffracting crystal forms
obtained from the native gastric lipases purified over the last 10 years from humans, rabbit, and dog (18), useful cubic crystals were
finally obtained from rHGL expressed in the insect cell/baculovirus expression system. The structure of rHGL has been solved by a combination of SIRAS, solvent flattening, and 2-fold averaging at 4.5 Å resolution. Phase extension procedures have made it possible to
extend the resolution to 3.6 Å. A preliminary model was constructed at
this resolution. The final structure was refined to 3.0-Å resolution using restrained noncrystallographic symmetry between the two molecules. The crystallographic R-factor based on all the data between
15.0 and 3.0 Å is 22.6%, and the corresponding R-free is 25.1% with
a model containing residues 9 to 53 and 57 to 379, 6 sugar residues
located on the 4 potential N-glycosylation sites, and 46 water molecules/monomer.
Overall Structure--
rHGL consists of one globular domain (Fig.
1) and belongs to the
/
hydrolase-fold family (1). The core domain, which is located between
residues 9-183 and 309-379 (Figs. 1 and 2) contains a central
sheet composed of 8 strands, 7 of which are parallel and 1 antiparallel
(strand 2) with 1(-2)435678 connectivity, and 6 helices, 3 on each side
of the
-sheet (Fig. 1C). Two segments are missing in the
electron density maps. The first of these was from residues 4 to 9, because the electron density starts abruptly at residue 9. In the
region of the second lacking segment (residues 54 to 56), some faint
electron density was observed, suggesting that the loop may be intact
but disordered (Fig. 1). The accessibility of this loop is consistent
with the previously reported preferential trypsin cleavage site
(Arg-55) of rabbit gastric lipase (35). In comparison with the
canonical
/
hydrolase fold, an extra helix (
1) is present at
the N terminus. Helix
A is shorter and has moved to the bottom of
the structure, helix
B is replaced by two helices (
B1 and
B2),
and helix
D is replaced by an extra domain (residues 184 to 308, Figs. 1 and 2) located between strands 6 and 7. Protrusions have been
observed in other lipases, generally constituting the device covering
the active site and called the lid. A "cap" domain occurs at the
same location in wheat serine carboxypeptidase II (residues 181 to 311, WCSII) (36) and in human protective protein (residues 181 to 347, HPP)
(37), two protease members of the
/
hydrolase-fold family. In
HPL, the lid (237 to 261) lies between strands 8 and 9, and a
-sandwich domain extends the enzyme at the C terminus (336 to 449).
In R. miehei lipase (4), a lid-containing extra domain
formed by an
-helix and two short strands (82-109) can be observed
between
4 and
B. The situation is more complex in C. rugosa lipase (6), where three protrusions have been observed that
together cover the central core of the enzyme: between strands 1 and 2 (residues 30 to 98), between
6 and
D (residues 241 to 331), and
between
7 and
E (residues 339 to 415). In the latter enzyme,
however, the lid covering the active site is composed of two helices
belonging to the first protrusion and comprising residues 66 to 92.

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Fig. 1.
Stereo ribbon representation of human gastric
lipase. A, view with the central -sheet parallel to
the paper plane. The cap domain on top of the / hydrolase core
domain is colored magenta and the putative lid,
green. For the / hydrolase fold, the color code is
helices (red), strands (blue), turns and random
coil (yellow). The catalytic triad, Ser-153, His-353, and
Asp-324, the disulfide bridge, and the Asn-attached sugars are shown in
ball and stick form. The disordered loop is
indicated by an arrow. The secondary structures were
calculated with Dictionary of Secondary Structure of Proteins.
B, view rotated about 90° from the previous one. Cys-244
is visible above the catalytic triad. C, schematic
representation of the rHGL catalytic domain topology with the secondary
structure elements identified (H, helix; B,
strand) as well as the catalytic triad.
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The amino acid sequence of the enzyme includes four consensus
N-glycosylation sites at Asn-15, -80, -252, and -308 (Fig.
2). Electron density patches attributable
to the first sugar attached to Asn have been observed at all four sites
(Fig. 1). GlcNAc residues were therefore introduced that nicely
sustained the refinement. No electron density was observed in the case
of the fucose residues attached at GlcNAc 1, in line with the fact that
the expression of the enzyme was carried out in a weakly glycosylating
cell type. The residual electron density made it possible to introduce
a second GlcNAc residue at sites 80 and 252. Sites 15, 80, and 252 are
located on one side of the molecule, whereas site 308 is on the other.
Site 80 is located on the core of the protein, whereas site 252, which
belongs to the cap domain, lies between helices
e5 and
e6. It is
worth noting that the glycan chains on the above-mentioned sites are in
close contact, which enhances the interactions between the core and the
cap.

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Fig. 2.
Sequence alignment of acid preduodenal
lipases and human lysosomal acid lipase. Note that there are no
insertions and only one deletion in the alignment. Identities are
displayed with yellow shading, and the values with respect
to HGL are: HLAL, 59%; rabbit gastric lipase, 85%; dog gastric
lipase, 85%; rat lingual lipase, 76%; calf pregastric esterase, 75%.
The catalytic triad, Ser-153, His-353, and Asp-324 is in
green, blue, and red, respectively.
The oxyanion hole residues are boxed. The secondary
structures have been calculated with DSSP.
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The amino acid sequence of HGL contains three cysteine residues, one of
which is free, whereas the other two are involved in a disulfide bridge
(35, 38-40). Based on the present three-dimensional structure, the
free cysteine can be unambiguously assigned to residue 244 (Fig.
3A), and the disulfide bridge,
to residues 227-236 (Fig. 1), as previously suggested by Lohse
et al. (40) using site-directed mutagenesis.

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Fig. 3.
The active site. A, stereo
view of the electron density map around the active site residues
Ser-153, His-353, and the single cysteine 244 (The density has been
contoured at 1 level). B, representation of the HGL
catalytic triad and of the oxyanion hole (Gln-154, Leu-67) superposed
to the same part of HPL (green, in C tracing).
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The Catalytic Machinery--
In lipases, as well as in serine
proteases, the catalytic machinery consists of a triad and an oxyanion
hole (2, 4). The present three-dimensional structure of rHGL is also
equipped with this machinery (Fig. 3B). The nucleophilic
serine belonging to the usual consensus sequence
GX1SX2G
(X1 and X2 being His and Gln, respectively, see Fig. 2) is located at position 153, between
5
and
C. This Ser-153 has an
conformation (41), which is a
characteristic feature of all the enzymes within the
/
hydrolase-fold family (1). His-353 (between
8 and
F) and Asp-324
(between
7 and
E) are the other two residues forming the
catalytic triad, as previously identified by site-directed mutagenesis
(42). The rHGL triad is almost superimposable on all the other lipase catalytic triads. The nucleophilic serine is covered by the cap (184-308) and is therefore not freely accessible to lipidic substrate molecules (Fig. 1). The environment of the catalytic triad seems not to
display any special features. However, one might expect His-353 to show
local pKa decrease to be able to explain the acidic
pH optimum of rHGL. There are no charged residues within a 10-Å sphere
centered on the Ser-153 O
atom. This absence makes it difficult to
suggest the mechanism possibly responsible for the local
pKa modulation.
Cys-244 was found to be very close to Ser-153 and His-353, and O
and
S
are at a distance of only 5.2 Å. This cysteine, which is
completely buried, has nevertheless permitted the binding of the
mercury acetate derivative. In the presence of specific cysteine reagents (C12-docecyldithio-5-2-nitrobenzoic acid or
5,5'-dithiobis(2-nitrobenzoic acid or 4,4'-dithiopyridine), it has been
established that a single cysteine can react stoichiometrically with a
concomitant loss of enzymatic activity (38). Another member of the acid
lipase family (calf pregastric esterose) has a threonine at this
position (Fig. 2). Furthermore, rHGL Cys-244 Thr mutant has been
constructed, and no loss of activity was observed in the purified
enzyme.2 In view of all the
above findings, the possibility that cysteine 244 may participate
directly in the catalytic mechanism can be ruled out. Based on the
three-dimensional structure, however, the specific inhibition observed
with sulfhydryl reagents was attributed to a steric hindrance
occurring at the level of the active site (see Fig. 3A).
The oxyanion hole is a device that stabilizes the oxyanion transition
state via hydrogen bonds with two main-chain nitrogens (2). It is often
evidenced by using organophosphate or organophosphonate complexes in
which the two main chain NH groups establish short hydrogen bonds with
an oxygen belonging to the phosphonate group. In rHGL, the oxyanion
hole has been identified on the basis of comparisons with other
esterases or lipases. Within the
/
hydrolase-fold family, one
mandatory component of the oxyanion hole is the NH group of the residue
following the nucleophile Ser (Gln-154 in rHGL). The second NH group
occupies a different position along the sequence but always originates
from the same spatial region. In some lipases, the oxyanion hole is not
preformed in the closed conformation but results from the concerted
movements during the opening process of the lid or of a small loop
bearing the nitrogen atom of the main chain. Comparisons between rHGL
and the open form of the C11-phosphonate HPL-colipase ternary complex
indicates that the second oxyanion NH group belongs to Leu 67 (Fig.
3B). The C
tracing of the loop bearing Leu-67 is fully
superimposable on that of the open HPL (Fig. 3B). It can
therefore be concluded that the oxyanion hole of rHGL is preformed as
in esterases, cutinase (43), and C. rugosa lipase (6). The
second oxyanion hole residue can be of very diverse nature in lipolytic
enzymes. R. miehei lipase (4) or cutinase (43) possess a
polar noncharged residue, Ser-82 and Ser-42, respectively. Furthermore,
the Ser-42 side chain in cutinase has been found to be a third
essential component of the oxyanion hole (44). It seems, however, that the Leu-67 side chain in rHGL may play a similar role to that of Phe-77
in the open form of HPL, leading to an exquisite interaction with the
alkyl group of the phosphonate inhibitor (see below).
The Cap and the Putative Lid--
The cap domain fold (residues
184-308) is an intricate mixture of 8 helices, turns, and random coils
(Figs. 1 and 2). When this domain is removed from rHGL on a display,
the active site becomes accessible to solvent. An apparently sufficient
degree of accessibility can be achieved, however, by removing a shorter stretch of residues between residues 215 and 244 (Fig.
4). The flanking residues of this stretch
are Pro-214 and Gly-245. It has been suggested that the
trans-cis-proline isomerization may be one of the
main factors involved in the C. rugosa lipase lid conformational changes (45). This, together with the flexibility of the
glycine residue, suggests that these residues might act as hinges in
the opening of the lid. It seems unlikely, however, that this lid
displacement might occur as a rigid body movement, as described
previously in R. miehei lipase (4). Given the complex
folding of this lid, the opening of rHGL is probably a more complex
process than those described previously. Cysteine 244 would therefore
be the last residue in the lid, before the hinge. The resulting
displacement of the chemically modified Cys-244, which occurs upon the
lid opening, might therefore be too limited to abolish the steric
hindrance during the formation of the lipase-substrate complex.

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Fig. 4.
Representations of human protective protein
(A) and rHGL (B) in the same
orientation with the catalytic triad in red. A,
Corey-Pauling-Koltun view of the core domain of HPP (yellow)
and of its cap domain (blue). The putative excision peptide
is given by a C trace (green). B,
Corey-Pauling-Koltun view of the core domain of rHGL
(yellow) and its cap domain (blue). The putative
lid is given by a C trace (green).
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Cap domains of this kind have been found to have the same topology in
WSCII (36), and in the related HPP, a serine carboxypeptidase zymogen
occurring in lysosomes (37). In HPP, the protruding domain is located,
as in rHGL, between strands 6 and 7, comprising residues 181-347. The
putative excision peptide is thought to be situated between residues
285 and 298. When this excision peptide is not removed, the active-site
serine remains covered (Fig. 4A), in the same way as the
putative lid covers the rHGL serine (Fig. 4B). In both
cases, the rest of the cap domain forms a ring around the active site,
ensuring the substrate specificity in the case of HPP and the lipid
affinity in the case of rHGL. In the framework of the above hypothesis,
the lid remains covalently attached to the body of rHGL after opening,
in contrast to HPP.
Active-site Accessibility and Model of a Phosphonate
Complex--
On the water-accessible surface of rHGL, a large number
of cavities that are accessible to water can be observed in the
structure (not shown). These are predominantly located at the interface between the core and the cap domains, which suggests that their packing
may be imperfect. Of particular interest is the small cavity (23 Å3) located above the active-site serine. Another cavity
of 113 Å3, lined by residues 274 to 278 and 284 to 287, makes it possible for the solvent to gain access to Ser-153, provided
that side-chain displacements can occur.
The question therefore arises as to whether or not the above channels
might make it possible for substrates or inhibitors to reach to the
active site. There exist several arguments against this hypothesis. It
is in fact impossible for a bulky long chain triglyceride to travel
through these channels to be accommodated in the closed active site.
When the putative lid domain was removed on the display, from residues
215 to 244, a large hydrophobic surface appears around the active site
and can act as a lipid binding site (Fig.
5). As observed in the three-dimensional
structures of open lipases, this hydrophobic surface may face the lipid
interface after the lid opening. Accordingly, the complementary surface part of the lid domain is also composed of hydrophobic residues facing
the putative lipid binding site in the closed form of rHGL.

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Fig. 5.
Molecular surface representations of
rHGL. A, the complete enzyme; B, same as in
A but with the putative lid depleted and the C11-phosphonate
enantiomers (red) of HPL represented in the active site (9).
Hydrophobic residues are colored blue.
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When positioning the two enantiomers of the C11-phosphonate inhibitor,
keeping their spatial position as observed in the three-dimensional structure of the HPL-C11-phosphonate complex, it was noted that both
molecules fit well against the bottom of the active-site cleft (Figs.
5B and 6). The C11 enantiomer corresponding to the hydrolyzable triglyceride acyl chain fit nicely into the putative acyl
binding site and interacted only with hydrophobic residues (Fig.
6). The other phosphonate enantiomer is
surrounded by a more polar environment, as in the HPL-C11 phosphonate
complex.

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Fig. 6.
Stereo view of the interactions of the HPL
C11-alkyl phosphonate inhibitor (9) with rHGL. For the
C11-phosphonate enantiomer the identified hydrophobic cleft extends
beyond the C11 carbon atom by about 7 additional atoms (mimicking a
stearoyl chain) and involves on each side: F203, V198, L202 and F212,
F207, I206, F205, respectively.
|
|
The Lysosomal Acid Lipase--
HLAL displays 59% identity and
75% homology with HGL, without any insertion or deletion (Fig. 2). A
straightforward homology model of HLAL was built, and this model was
examined in the light of the mutations, resulting in a clinical
phenotype in humans. Besides mutants bearing large deletions, few
single mutations associated with cholesteryl ester storage disease have
been described. The deletion of fragment 205 to 253 and fragment 254 to
277 affect the cap domain (29, 31, 32, 46). It is possible that the residual activity reported in the latter study (5-10%) might be because of the fact that the
/
hydrolase fold of the catalytic domain has remained intact. The mutation Leu-273
Ser creates a new
potential glycosylation site (NMS), and this mutant expressed in HeLa
cells has been found to have a higher molecular mass than that of the
wild type HLAL expressed in the same system (29, 30). This residue is
water-exposed, and the bulky glycan moiety may prevent either the
substrate binding or the movement of the lid. The mutation Leu-336
Pro is located just inside helix
E and probably destabilizes its
structure, leading to an inactive enzyme as initially proposed by
Seedorf et al. (32). The mutation Gly-66
Val is easily
interpretable in structural terms, because the valine side chain
clashes with the active-site serine 153 and also partly blocks the
putative triglyceride binding site, on similar lines to what has been
found to occur with pancreatic lipase RP1 (47).
 |
ACKNOWLEDGEMENTS |
We are grateful to Frédéric
Carrière for helpful discussions and for his constant interest in
this topic. The contribution of C. Wicker-Planquart is particularly
acknowledged for setting up the expression of rHGL in a
baculovirus/insect cell system. R. Verger acknowledges the constant
help of the Institut de Recherche Jouveinal/Park Davis since 1989. We
thank C. Abergel and C. Jelsch for performing the preliminary
crystallization trials, J. Bonicel for performing mass
spectroscopy on the rHGL, and S. Diotallevi for technical assistance.
We thank J. Perez and A. Bentley for their assistance during the data
collection at the LURE. We also thank Dr. Jessica Blanc for correcting
the English.
 |
FOOTNOTES |
*
This research was carried out in the framework of the EU
Biotech- lipase project (BIO2-CT94-3041).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
The atomic coordinates and structure factors (code 1HLG) have
been deposited in the Protein Data Bank, Brookhaven National Laboratory, Upton, NY.
§
These authors have made equal contributions to the present study.
To whom correspondence should be addressed. Fax:
33-491-16-45-36; E-mail: cambillau{at}afmb.cnrs-mrs.fr.
2
Canaan, S. (1999) Biochem. Biophys. Res.
Commun., in press.
 |
ABBREVIATIONS |
The abbreviations used are:
HPL, human
pancreatic lipase;
HGL, human gastric lipase;
rHGL, recombinant HGL;
HLAL, human lysosomal acid lipase;
SIRAS, single isomorphous
replacement with anomalous scattering;
HPP, human protective
protein.
 |
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